DETECTION AND MOLECULAR CHARACTERIZATION OF PORCINE NOROVIRUSES AND SAPOVIRUSES

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in the Graduate

School of The Ohio State University

by

Qiuhong Wang, Bachelor of Medicine, M.S.

* * * * *

The Ohio State University

2005

Dissertation Committee: Approved by Distinguished University Professor Linda J. Saif, Adviser

Assistant Professor Julie A. Funk ______Adviser Professor John H. Hughes Graduate Program in Assistant Professor Jeffrey T. LeJeune Veterinary Preventive Medicine

ABSTRACT

Enteric caliciviruses are emerging pathogens that cause gastroenteritis in humans and animals. They comprise 2 genera, Norovirus and Sapovirus in the family Caliciviridae.

Human noroviruses (NoV) infect people of all ages. They cause more than 90% of nonbacterial gastroenteritis outbreaks worldwide and have been listed as class B biological pathogens by the National Institutes of Health/Biodefense Program. Human sapoviruses

(SaV) are associated with 1.8-9% of pediatric gastroenteritis worldwide. Whether NoVs and SaVs are zoonotic pathogens is unclear due to limited studies of animal NoVs and

SaVs. However, the classic calicivirus, vesiviruses cause cross-species infections and infect marine mammals, cats, dogs, cattle and pigs. Bovine NoVs may infect humans based on a bovine NoV seroprevalence study of veterinarians in contact with cattle. Compared to cattle, pigs have a gastrointestinal tract more similar to that of humans in anatomy and physiology. Compared to bovine NoVs, porcine NoVs are genetically and antigenically more closely related to human NoVs. These data raise concerns of whether pigs may be reservoirs for the emergence of human NoVs. For animal SaVs, only 2 genetically similar porcine SaV strains and one mink SaV strain have been reported. The porcine SaV prototype Cowden strain was isolated from a diarrheic pig. It causes diarrhea and intestinal lesions in gnotobiotic (Gn) pigs. To date, no prevalence study of porcine SaVs has been reported. The main objectives of this dissertation were: 1) to investigate the genetic

ii diversity of porcine NoVs and their relationship to human NoVs; 2) to investigate the

genetic diversity of porcine SaVs and their relationship to human SaVs; 3) to develop

sensitive and specific methods for detection of porcine NoVs and SaVs; and 4) to study the

prevalence of porcine NoVs and SaVs in US swine.

We screened for porcine NoVs by reverse transcription-PCR (RT-PCR) using

calicivirus universal primers on 275 fecal samples from normal US adult pigs. Six samples

were confirmed NoV positive by sequencing. Based on sequence analysis of the 3 kb of the

genomes of 5 porcine NoVs, 3 genotypes [2 new (GII-18 and 19) and 1 confirmed (GII-11)]

within GII and two potential recombinant strains were identified. One genotype (GII-18)

of porcine NoVs was genetically and antigenically related to human NoVs and replicated

in gnotobiotic pigs. By similar approaches, porcine SaVs were identified as genetically diverse viruses comprising at least 2 genogroups, one previously classified (GIII) and one potentially new genotype (GVI?). Two porcine SaVs were potential recombinants. One porcine SaV strain (Po/SaV/MI-QW19/2002/US) was genetically most closely related to human SaVs based on the partial RdRp sequence (286 nt).

Based on the obtained sequences of porcine NoVs and SaVs, several primer pairs

were designed and evaluated for the detection of porcine NoVs and SaVs, respectively. An

internal control RNA was developed and used in RT-PCR to monitor for RT-PCR

inhibition. A 10-fold dilution of sample RNA containing RT-PCR inhibitors usually no longer interfered in the RT-PCR reaction. Microwell hybridization assays were developed to confirm the porcine NoV- and SaV-specific amplicons.

A prevalence study of porcine NoVs and SaVs in US swine was performed by the

newly developed RT-PCR-hybridization assays. The porcine NoVs were detected

iii exclusively from finisher pigs in 4 of 7 farms and 1 slaughterhouse with an overall prevalence of 20% in finisher pigs (range of 3-40% in the positive farms). Porcine SaVs were detected from all ages of pigs. The prevalence of porcine SaVs was 62% overall, lowest in nursing pigs (21%) and highest in post-weaning pigs (83%), and varied from 37 to 100% depending on the farm. Mixed infections of NoVs and SaVs were common in finisher pigs with an overall prevalence of 27% among NoV- or SaV-positive pigs.

These findings have improved our understanding of the genetic diversity of porcine

NoVs and SaVs and their relationships to human strains. Certain porcine NoVs and SaVs are genetically or antigenically related to human strains. This finding and the high prevalence of NoVs and SaVs in subclinically infected swine including their detection from slaughterhouse pigs increase the risks that pigs may be reservoirs for human NoV and

SaV strains.

iv

Dedicated to my husband Hong Liu, son Hengyu,

my parents Xushong Wang and Jie Gao, and my parents-in-law Jingqin Liu and Yongbi Tang

v

ACKNOWLEDGMENTS

Grateful appreciation is due to my adviser, Dr Linda J. Saif, for her encouragement, challenge, guidance and support throughout my years of study. I have learned substantially from her unwavering emphasis on research quality and unlimited patience during my studies and during the preparation of manuscripts and this dissertation.

I would also like to thank my current committee members, Dr. Julie A. Funk, who collected most of the field fecal samples for my studies, Dr. John H. Hughes and Dr.

Jeffrey T. JeJeune, and my prior committee members (prior to their leaving OSU) Dr

Srinand Sreevatsan and Dr. Qijing Zhang, for their constructive comments, advice and encouragement throughout my studies.

Special thanks are due to all my colleagues working on enteric caliciviruses, Dr.

Keong-Ok Chang, Dr. Mingzhang Guo, Dr. Myung Guk Han, Menira Souza, Dr. Sonia

Cheetham, Veronica Costantini and Dr. Christopher Thomas for their sharing experience and knowledge, their technical assitance and their friendship throughout the years of my

study.

I also thank Peggy Lewis, Paul Nielsen, Wei Zhang and Ana Azevedo for their

technical assistance.

Many thanks are due to Dr. Marli Azevedo, Dr. Yuxin Tang, Dr. Lijuan Yuan, Dr.

Kwang-il Jeong, Trang Van Nguyen and Dr. Ana Gonzalez, whose friendship has helped me through difficult times.

vi I want to thank Dr. Julliete Hanson, Richard McCormick, Janette McCormick,

Todd Root, Greg Meyers and Don Westfall, for technical assistance and commitment.

Special thanks are to Hannah Gehman and Robin Weimer for indispensable support.

vii

VITA

September 3, 1969 Born – Beijing, China

1989 - 1994 Bachelor of Medicine Beijing Medical University, Beijing, China

1994 -1998 Research staff Institute of Virology Chinese Academy of Preventive Medicine, Beijing, China

1998 - 2000 Master of Health Sciences The University of Tokyo, Tokyo, Japan

2000 – present Graduate Research Associate Food Animal Health Research Program Ohio Agricultural Research and Development Center Department of Veterinary Preventive Medicine The Ohio State University Wooster, Ohio

PUBLICATIONS

Research Publication

1. Chang, K. O., S. S. Sosnovtsev, G. Belliot, Q. Wang, L. J. Saif, and K. Y. Green. 2005. Reverse genetics system for porcine enteric calicivirus, a prototype sapovirus in the Caliciviridae. J Virol 79:1409-16.

2. Han, M. G., Q. Wang, J. R. Smiley, K. O. Chang, and L. J. Saif. 2005. Self-assembly of the recombinant capsid protein of a bovine norovirus (BoNV) into virus-like particles and evaluation of cross-reactivity of BoNV with human noroviruses. J Clin Microbiol 43:778-85.

3. Tang, Y., Q. Wang, and Y. M. Saif. 2005. Development of a ssRNA internal control template reagent for a multiplex RT-PCR to detect turkey astroviruses. J Virol Methods 126:81-6. viii 4. Wang, Q. H., J. Kakizawa, L. Y. Wen, M. Shimizu, O. Nishio, Z. Y. Fang, and H. Ushijima. 2001. Genetic analysis of the capsid region of astroviruses. J Med Virol 64:245-55.

5. Sakamoto, T., H. Negishi, Q. H. Wang, S. Akihara, B. Kim, S. Nishimura, K. Kaneshi, S. Nakaya, Y. Ueda, K. Sugita, T. Motohiro, T. Nishimura, and H. Ushijima. 2000. Molecular epidemiology of astroviruses in Japan from 1995 to 1998 by reverse transcription-polymerase chain reaction with serotype-specific primers (1 to 8). J Med Virol 61:326-31.

6. Wang, Q., and Z. Y. Fang. 1997. Molecular biology of Norwalk-like viruses (review). Foreign Medical Sciences (field: Virology ) 3:27-31 (Chinese).

7. Wang, Q., Z. Y. Fang, Z. Lu, D. Zhou, J. Zhang, and D. Wu. 1996. Detection of hepatitis A virus contamination in shellfish by antigen capture/polymerase chain reaction. Chinese Journal of virology 12:123-128 (Chinese).

8. Pan, N., Z. Y. Fang, J. Yan, Q. Wang, Z. Gong, and J. Zhang. 1995. Molecular epidemiology of group A rotaviruses during 1989-1992 in Hubei province of China. Chinese Journal of Experimental and Clinical Virology 9:250-253 (Chinese).

9. Wang, Q., and Y. Chang. 1995. The effects of chemical toxins on secretion of luteinising hormone (LH) and follicle stimulating hormone (FSH) of male pituitary (review). Foreign Medical Sciences (field: Hygiene) 3:140-144 (Chinese).

10. Chang, Y., S. Lu, X. Jin, Q. Wang, and Z. Liu. 1994. Effects of HgCl2 on setolic cells of testis in rats. Journal of Health Toxicology 8:300 (Chinese).

FIELDS OF STUDY

Major Field: Veterinary Preventive Medicine

Studies in Molecular Virology

ix

TABLE OF CONTENTS

Page Abstract...... ii

Dedication...... v

Acknowledgments...... vi

Vita...... viii

List of Tables ...... xiii

List of Figures...... xv

List of Abbreviations ...... xvi

CHAPTER 1. Literature review: Noroviruses and sapoviruses 1.1 Introduction...... 1 1.2 Molecular biology and genetic classification ...... 3 1.2.1 Genome organization and classification ...... 3 1.2.2 Polyprotein processing and function of non-structural proteins...... 6 1.2.3 Capsid proteins and structure...... 9 1.2.4 Subgenomic RNA and recombination ...... 12 1.2.5 Evolution…………...... 14 1.3 General biology...... 15 1.3.1 General biologic characteristics...... 15 1.3.2 Cell culture and infectious clones...... 17 1.3.3 Molecular mechanisms of replication...... 18 1.3.4 Virus assembly and apoptosis...... 20 1.4 Pathogenesis...... 22 1.4.1 Studies in animal models ...... 23 1.4.2 Host receptors and viral ligands...... 25 1.4.3 Clinical features ...... 31 1.4.4 Site of primary replication ...... 33 1.4.5 Virulence...... 39 1.5 Laboratory diagnosis...... 39 1.5.1 Electron microscopy ...... 40 1.5.2 RT-PCR and real-time RT-PCR ...... 41 x 1.5.3 Immunoassays...... 46 1.5.4 Detection of serological responses...... 49 1.6 Epidemiology...... 50 1.6.1 Outbreaks, sporadic cases and asymptomatic infections ...... 50 1.6.2 Seroprevalence ...... 55 1.6.3 Zoonotic potential ...... 57 1.7 Immunity...... 59 1.8 Treatment ...... 63 1.8.1 Anti-gastroenteritis ...... 63 1.8.2 Anti-virus replication and RNAi...... 64 1.9 Control and Prevention...... 65 1.9.1 General approaches...... 65 1.9.2 Vaccines...... 66 1.10 References ...... 70

CHAPTER 2 Porcine noroviruses: genetic and antigenic relationships to human noroviruses 2.1 Summary...... 98 2.2 Introduction...... 99 2.3 Materials and Methods...... 101 2.4 Results ...... 106 2.5 Discussion...... 110 2.6 Acknowledgments...... 114 2.7 References...... 115

CHAPTER 3 Genetic diversity and recombination of porcine sapoviruses 3.1 Summary...... 130 3.2 Introduction...... 131 3.3 Materials and Methods...... 133 3.4 Results ...... 136 3.5 Discussion...... 141 3.6 Acknowledgments...... 146 3.7 References...... 147

CHAPTER 4 A new microwell hybridization assay for the detection of porcine noroviruses and sapoviruses by reverse transcription-PCR 4.1 Summary...... 161 4.2 Introduction...... 162 4.3 Materials and Methods...... 164 4.4 Results ...... 170 4.5 Discussion...... 176 4.6 Acknowledgments...... 181 4.7 References...... 181

xi CHAPTER 5 Prevalence of noroviruses and sapoviruses in swine determined by reverse transcription-PCR 5.1 Summary...... 194 5.2 Introduction...... 195 5.3 Materials and Methods...... 197 5.4 Results ...... 202 5.5 Discussion...... 205 5.6 Acknowledgments...... 211 5.7 References...... 211

Bibliography ...... 222

xii

LIST OF TABLES

Table Page 1.1 Representative strains of NoVs and SaVs...... 95

2.1 Classification and GenBank accession numbers of NoV strains and a SaV strain used for sequence analysis...... 120

2.2 The sizes of the deduced capsid protein VP1 and the minor capsid protein VP2, the overlap regions, and the 3’-UTR of porcine GII NoVs...... 121

2.3 Summary of the percent amino acid identities of NoVs within the capsid region ...... 122

2.4 Antigenic cross-reactivity between human GII NoV antigens (VLPs) and a pig convalescent antiserum against porcine GII NoVs as determined by ELISA ...... 123

3.1 Porcine SaV strains detected using primers p290/110 and sequenced in this study...... 151

3.2 Summary of sapovirus strains and representative strains for Lagovirus, Vesivirus, Norovirus genera and NB-like viruses used in sequence analysis… ...... 152

3.3 The sizes of the deduced capsid protein, the ORF1-ORF2 overlap region, the ORF2 protein, and the 3'-UTR of porcine SaVs...... 153

3.4 Summary of the amino acid (nucleotide) sequence identities (%) in the complete capsid region between the newly identified porcine SaVs and reference calicivirus strains ...... 154

4.1 Representative porcine NoV and SaV strains and positive fecal samples used in this study...... 184

4.2 RT-PCR primers used for the detection of porcine NoVs and SaVs...... 185

4.3 Hybridization DNA probes for the detection of porcine NoV- and SaV-specific amplicons ...... 186 xiii

4.4 Comparison of primer pairs p290/110, PEC66/65, PEC46/45, PEC68/67, PSV6/7 and PSV11/14 for the detection of porcine SaVs ...... 187

4.5 Detection spectrum of the NoV probes for PNV7-Bio/8 RT-PCR products…188

4.6 Detection spectrum of the SaV probes for the RT-PCR products of several primer pairs...... 189

4.7 The absorbance of porcine NoV and SaV positive, negative and non-specific amplicons in each RT-PCR-hybridization assay ...... 190

5.1 Number and time of fecal sample collection from nursing, post weaning, finisher pigs and sows from different farms ...... 216

5.2 The prevalence of porcine NoVs in pigs...... 217

5.3 The prevalence of porcine SaVs in pigs ...... 218

5.4 The prevalence of porcine SaVs in pigs of OH farm B at different times...... 219

5.5 The prevalence of porcine NoVs, SaVs and co-infection in finisher pigs...... 220

5.6 The prevalence of RT-PCR inhibitors in the pig fecal samples...... 221

xiv

LIST OF FIGURES

Figure Page

1.1 The genome organization of NoVs and SaVs...... 96

1.2 The proteolytic cleavage map of Hu/NoV/Southampton/91/UK and domains and subdomains of the capsid protein of Hu/NoV/Norwalk/68/US..... 97

2.1 Neighbor-Joining phylogenetic trees of genogroup II NoVs...... 124

2.2 Identification of QW170 and QW218 strains as potential recombinants ...... 126

2.3 Immune electron micrograph of porcine NoVs...... 128

2.4 Conserved amino acid motifs of the capsid proteins of NoVs ...... 129

3.1 Phylogenetic tree of the partial RdRp region (95 aa) of SaVs...... 155

3.2 Neighbor-Joining phylogenetic trees of SaVs ...... 156

3.3 Identification of QW270 and MM280 strains as potential recombinants...... 158

3.4 Sequence alignments of the RdRp-capsid junction region of SaVs...... 160

4.1 The complete RNA sequence of the internal control RNA (632 nt in length) for primer pairs PEC66/65 and PEC46/45...... 191

4.2 Agarose gel (1.5%) eletrophoresis of the RT-PCR products (ethidium bromide staining) of the Po/SaV/Cowden strain and the IC RNA ...... 192

4.3 Comparison of the detection limits of agarose gel electrophoresis and the microwell hybridization assay (probe PEC-P1) for detection of RT-PCR PEC68-Bio/67 products of Po/SaV/GVI?/JJ681 ...... 193

xv

LIST OF ABBREVIATIONS

Basic local alignment search tool (BLAST) Bovine coronavirus (BCV) Canine calicivirus (CaCV) Cell culture immunofluorescence (CCIF) Cesium chloride (CsCl) Cetyltrimethylammonium bromide (CTAB) Cholera toxin (CT) Dalton (Da) Day post-inoculation (DPI) Eagle’s minimal essential medium (EMEM) Enzyme-linked immunoassay (ELISA) Feline calicivirus (FCV) Genogroup (G) Gnotobiotic (Gn) Guanidinium thiocyanate (GTC) Histo-blood group antigens (HBGA) Horseradish peroxidase (HRP) Immune electron microscopy (IEM) Immunofluorescence (IF) Inducible nitric oxide synthase (iNOS) Interferon (IFN) Internal control (IC) International Committee on the Taxonomy of Viruses (ICTV) Internal ribosomal entry site (IRES) Intranasal (i.n.) Intracerebral (i.c.) Intraperitoneally (i.p.) Intestinal transplant (IT) 7-methylguanosine (m7G) Monoclonal antibody (MAb) Mutant Escherichia coli heat-labile toxin (mLT-R192G) Morpholino (PMO) Calicivirus Nebraska/80/US (NB) Norovirus (NoV) N-terminal protein (Nterm) Nucleotide triphosphatase (NTPase) xvi Nucleoside triphosphate (NTP) Open reading frames (ORF) Peripheral blood mononuclear cell (PBMC) Peroral (p.o.) Polyethylene glycol (PEG) Protein kinase A (PKA) Protein kinase RNA-activated (PKR) Protruding (P) domain Radio immunoassay (RIA) Rabbit hemorrhagic disease virus (RHDV) Recombinant Identification Program (RIP) Red blood cells (RBC) Reverse transcription (RT) RNA-dependent RNA polymerase (RdRp) RNA interference (RNAi) Secretor phenotype (Se+) Weak secretor phenotype (Sew) Nonsecretor phenotype (Se-) Sodium dodecyl sulfate (SDS) Solid phase IEM (SPIEM) Signal transducer and activator of transcription 1 (STAT1) San Miguel sea lion virus (SMSV) Sapoviruses (SaV) Shell domain (S domain) Small, round-structured viruses (SRSV) Mean tissue culture infective dose (TCID50) In vitro coupled transcription and translation (TNT) Untranslated region (UTR) Vesicular exanthema of swine-like virus (VESV) Virus-like particles (VLPs) Virus protein genome-linked (VPg)

xvii

CHAPTER 1

LITERATURE REVIEW

NOROVIRUSES AND SAPOVIRUSES

1.1 INTRODUCTION

Diarrhea, which is still one of the most common illnesses in humans, causes

nearly 2 million deaths in children in developing countries (248). The first virus that was

discovered to cause human diarrhea was Norwalk virus in stools of school children with diarrhea in Norwalk, Ohio (114). It was discovered in 1972 by Kapikian et al. (114) using immune electron microscopy (IEM) and was described as having an indistinctive morphology. Because Norwalk virus does not replicate in cell culture or animals except for primates, it was not classified until 1993 when molecular cloning and sequencing of the viral genome identified it as a member of the family Caliciviridae. However, the cup-

shaped surface depressions of Norwalk virus are not as clear as those of classical animal

caliciviruses (104). Norwalk virus is the prototype of human calicivirus. Subsequently,

many small, round-structured viruses (SRSV), which are morphologically similar to

Norwalk virus, were initially referred to as Norwalk-like viruses in the Caliciviridae (43).

Later, Norwalk-like viruses were assigned to the genus Norovirus approved by the

1 international Committee on the Taxonomy of Viruses (ICTV) in 2002 (160). Since the

discovery of Norwalk virus, noroviruses (NoV) have emerged as the most common

pathogens of food- and water-borne viral gastroenteritis in humans including both

outbreaks and sporadic cases worldwide (45, 46, 127, 161). The NoVs have been listed as

class B biological pathogens by the National Institutes of Health/Biodefense Program.

Members of the Norovirus genus, except for the newly identified murine NoV (117),

cause gastroenteritis in humans and calves (63, 71, 257). Norovirus RNA was also detected from adult pig fecal samples in Japan and Europe (232, 245), but their role in

diarrhea was undefined.

In addition to NoVs, a second distinct group of enteric caliciviruses, the

sapoviruses (SaV), belong to the genus Sapovirus and are associated with human and

animal gastroenteritis (26, 52, 73, 74). They have typical cup-shaped depressions on the

surface and were first found in stools of diarrheic children in 1976 by direct electron

microscopy (50, 148). The prototype Hu/SaV/Sapporo was first detected during a

gastroenteritis outbreak in an infant center in Sapporo, Japan in 1977 (26). Like NoVs,

SaVs were tentatively named Sapporo-like viruses and are now recognized as emerging

pathogens causing diarrhea in humans, pigs and mink (52, 63, 73, 74, 160).

During the past 30 years, significant progress has been made on characterization

of enteric caliciviruses. Major breakthroughs include the following. The genomes of

representative strains of NoVs and SaVs were characterized and the structure of Norwalk

virus was established by x-ray crystallography of baculovirus-expressed Norwalk virus-

like particles (VLPs) (103, 104, 138, 197). Reverse transcription (RT)-PCR was widely

used for diagnosis (6). Host histo-blood group antigens (HBGA) were discovered as host

2 cell receptor or co-receptors for NoV infection (95). A porcine SaV Cowden strain was shown to replicate in a continuous cell line (193). Murine NoVs were discovered in mice and mice were tested as an infection model to study NoVs (117). Pathogenesis studies of porcine SaV Cowden strain were performed in pigs (74). In spite of these breakthroughs, human NoVs and SaVs still do not grow in cell culture and there is still no animal enteric disease model available for NoVs. Knowledge of the pathogenesis of NoVs, immune responses and immunity to NoV and SaV infections are limited. This review focuses on the molecular biology, virus pathogenesis, diagnosis, epidemiology and immunity to

NoVs and SaVs. The zoonotic potential of animal NoVs is also discussed.

1.2 MOLECULAR BIOLOGY AND GENETIC CLASSIFICATION

1.2.1 Genome organization and classification

Viruses in the family Caliciviridae are nonenveloped, polyadenylated single-

stranded positive-sense RNA viruses (63). They are 27-35 nm in diameter and contain a

genome of 7 to 8 kb. Caliciviruses were classified into 4 genera by the ICTV in 2002:

Norovirus, Sapovirus, Vesivirus and Lagovirus (160). Later, a bovine enteric calicivirus

strain Bo/Nebraska/80/US was characterized as a potential new genus in Caliciviridae

but its classification has not yet been assigned by ICTV (219). The formal name of a

NoV or SaV strain is organized as follows: Host species from which the virus was

obtained/genus abbreviation/genogroup-genotype/strain name/year of identification/

country of origin (63). Researchers usually use the strain name directly after first giving

the formal name.

3 The first genome organization was determined for Norwalk virus in the early

1990s (104) (Fig. 1.1). Presently, a total of 23 genomes of human, 1 of bovine and 1 of

murine NoVs have been characterized. The NoV genome is 7.3-7.7 kb long and includes

3 open reading frames (ORF) that encode a non-structural polyprotein, a major capsid

protein (VP1 or capsid protein) and a minor capsid protein (VP2 or small basic protein).

The polyprotein undergoes protease processing to produce several nonstructural proteins

(Fig. 1.2A) (11, 63, 195). The capsid protein is composed of a conserved shell (S) domain and hypervariable protruding (P) domain, with the latter subdivided into P1A, P2 and

P1B subdomains based on x-ray crystallography of Norwalk virus VLPs (Fig. 1.2B)

(197).

The first genome of SaVs was determined for Hu/Manchester/93/UK (136, 138)

(Fig. 1.1). Since then, the full length genomes of 4 human and 1 porcine SaVs have been

characterized (72, 118, 204). The genome is 7.3-7.5 kb long and contains two main ORFs

based on sequence analysis. The ORF1 encodes a polyprotein that undergoes protease

processing to produce several non-structural proteins and a capsid protein. The capsid protein gene of SaV is contiguous with ORF1, whereas the capsid protein of NoV is encoded by a separate ORF2 (Fig. 1.1). This is the major difference in genome organization between NoVs and SaVs. The ORF2 of SaV encodes a small basic protein with unknown function. For GI, GIV and GV human SaVs, another ORF overlapping the

5’-end of the capsid gene was proposed because there is a conserved translation initiation motif GCAAUGG at the 5’-end of this ORF (49, 216).

Because there is no cell culture system or small animal model for NoVs and SaVs

except for the Po/SaV/GIII/Cowden/80/US strain (74, 193), antigenic classification of

4 these viruses by two-way cross-neutralization tests is not yet possible. Genetic classification of these viruses based on phylogenetic analysis of the complete capsid sequences is a generally accepted method (62, 119, 216). Katayama et al. (119) investigated which region of the genome was most suitable to classify the genotypes and genogroups of NoVs based on similarity plot analysis, bootstrap value and pairwise distances of the entire genomes of 18 strains, the complete sequences of ORF1, ORF2 and ORF3 and segments of ORF1 and ORF2. They found that the full-length genome, the complete ORF1 or ORF2 and the capsid N-terminal/S domain could segregate viruses into genus, genogroup and genotypes. However, the RNA-dependent RNA polymerase

(RdRp) could not be used to clearly identify genotypes; the P domain of the capsid protein could not be used to assign genogroups; and the VP2 could not distinguish genus.

There is no such detailed investigation of classification for SaVs. Sequence analysis of the complete capsid region is a generally accepted method for classification of SaVs into genogroups and genotypes.

The NoVs are genetically highly diverse viruses. Recently, they have been

divided into at least 27 genotypes within 5 genogroups: GI/1-8, GII/1-17, GIII/1-2, GIV and GV based on a Bayesian tree and maximum likelihood pairwise distance analysis of the capsid proteins of 164 strains (264). The pairwise amino acid distances of NoVs within genotypes, between genotypes and between genogroups are 0-14.1%, 14.3-43.8% and 44.9-61.4%, respectively.

Similar to NoVs, SaVs are genetically variable viruses and have been classified

into 5 genogroups (GI, II, III, IV and V) and at least 6 genotypes (GI/1-3, GII/1-3) based

on the complete capsid sequences (49, 216). The pairwise amino acid distances of SaVs

5 within genotypes, between genotypes and between genogroups are 1-8%, 9-29% and 51-

63%, respectively.

The classification and GenBank accession numbers of representative NoV and

SaV strains are summarized in Table 1.1.

1.2.2 Polyprotein processing and functions of non-structural proteins

The organization of caliciviruses includes a large nonstructural polyprotein gene that precedes the major capsid protein gene (Fig. 1.1). The strategy for defining NoV

ORF1 cleavage sites and nonstructural protein functions was the combination of site- directed mutagenesis, in vitro coupled transcription and translation (TNT) and N-terminal protein sequence analysis. There is no report on the polyprotein processing and functions of non-structural proteins of SaVs. However, conserved functional domains of each nonstructural protein have been also observed for SaV genomes based on comparative sequence analysis of SaVs, NoVs, lagoviruses and vesiviruses.

The polyprotein is about 200-kDa and contains at least 6 functional domains: N- terminal protein (Nterm, p48), 2C-like nucleotide triphosphatase (NTPase, p41), p20 or p22, virus protein genome-linked (VPg, p16), proteinase (protease or, Pro, p19) and polymerase (RdRp or Pol, p57) (11, 63, 86, 137, 140, 195, 218, 224). The functional regions of NoV polyprotein are shown in Fig 1.2A.

Proteolytic cleavage map. A proteolytic cleavage map based on the polyprotein amino acid sequence was determined for Hu/NoV/GI-2/Southampton strain by translation of the ORF1 polyprotein in reticulocyte lysate (137) and bacterial expression systems

(140): Q399/G, Q762/G, E961/G, E1099/A, E1280/G (Fig. 1.2A). The proteolytic

6 cleavage map for the GII NoV MD145 strain was similar to that of Southampton strain

using an in vitro TNT assay: Q330/G, Q696/G, E875/G, E1008/A, and E1189/G in the

ORF1 polyprotein (11). These cleavage sites are also conserved in the sequences of other

GI and GII NoVs. Some cleavage sites of these viruses were confirmed by other

experiments, e.g., the processing at EG and EA was confirmed for Hu/NoV/GII-

4/Camberwell strain in a eukaryotic expression system (COS cells) (218). Protein

precursors were also found during the processing including stable precursors p20VPg and

ProPol and less stable p20VPgProPol, p20VPgPro and VPgPro.

Non structural proteins. (i) The 3C-like proteinase. It is a 19 kDa viral protein and is responsible for cleavage of the polyprotein into several functional proteins during virus replication. The calicivirus 3C-like proteinase belongs to a family of viral proteinases that resemble the cellular chymotrypsin-like serine proteinase (17). The nucleophilic serine residue of the cellular enzymes is replaced by a cysteine in the viral proteinases. Activity of the 3C-like cysteine proteinase of human and animal caliciviruses has been analyzed by expression in bacteria, rabbit reticulocyte lysates, and most recently, in mammalian cells (137, 140, 153, 218). The active sites of the amino acid residues of the 3C-like proteinase of strain Hu/NoV/GI-4/Chiba407/87/JP were determined as His30 and Cys139 (based on the amino acid number of the proteinase), forming a catalytic dyad, by site-directed mutagenesis of the precursor VPgPro followed by expression in bacteria

(224). Site-directed mutagenesis of the proteinase of Hu/NoV/GI-2/Southampton confirmed residue C1238 as the nucleophilic residue (137). During polyprotein processing, a protein precursor ProPol is stable and possesses the proteinase and RdRp activities simultaneously, which is a bifunctional enzyme (10).

7 (ii) The NTPase. It is a 40-41 kDa viral protein depending on strains. The p41 of

Hu/NoV/GI-2/Southampton was expressed in bacteria and it exhibited NTPase activities: nucleoside triphosphate (NTP)-binding and NTP hydrolysis (195). This NTPase activity was not stimulated by single-stranded nucleic acids and it had no detectable helicase activity. Moreover, its function during virus replication is unclear. Stimulation of NTPase activity by RNA is a common property of helicases, whereas the NTPase activity of this p41 was inhibited by homopolymeric RNA, suggesting that the NTPase activity may need to be down regulated at some point during viral replication. Moreover, it did not unwind an RNA-DNA heteroduplex which was readily unwound by hepatitis C virus

RNA helicase NS3. Thus, although p41 of caliciviruses has adopted the NTP-binding and hydrolysis motifs of superfamily (SF3) helicases, they seem to use their ability to hydrolyze NTPs for an unknown function, but not nucleic acid unwinding.

(iii) Polymerase. The 3D-like polymerase itself is a mature protein with a size of

57-58 kDa depending on strains. It catalyzes the synthesis of viral genomic and subgenomic RNA by way of an RNA-dependent RNA polymerization during virus replication. The polymerase domain is also in a stable protein precursor ProPol. Recent studies demonstrated that the polymerase domain in both forms is active (10).

(iv) The VPg. It is a 16 kDa protein. Like the positive-stranded RNA viruses in

the families Picornaviridae, Potyviridae, Luteoviridae and Comoviridae, viruses in the

family Caliciviridae have a genome without the 5’ terminal 7-methylguanosine (m7G)

cap structure, which binds to the translation initiation factor eIF4F of eukaryotic cells

(209). Instead, a viral protein VPg is covalently linked at the 5' end of the genome.

Removal of VPg results in loss of infectivity and dramatically reduced translation of

8 feline calicivirus F9 strain (FCV) RNA in vitro (20, 90). However, a full-length cDNA

clone of FCV retained infectivity after capping with the m7G (225), indicating that m7G can substitute for VPg. Recent studies identified that the Norwalk virus VPg binds to eIF3d, subunit of translation initiation factor eIF3 by yeast two-hybrid analysis screening of an epithelial cell cDNA library (33). This binding pattern was confirmed using purified mammalian eIF3. Furthermore, the VPg inhibited the translation of reporter RNAs having the cap, encephalomyocarditis virus internal ribosomal entry site (IRES) or a cricket paralysis virus IRES structure in a dose-dependent manner using a cell-free translation system. These results indicate that VPg may function in translation initiation through protein-protein interactions with the translation machinery.

1.2.3 Capsid proteins and structure

Two capsid proteins are expressed for NoVs and SaVs. The major capsid protein

(capsid protein, VP1) is about 57-59 kDa for NoVs and 58-62 kDa for SaVs (26, 63, 75,

100, 103, 105, 130). It is encoded by ORF2 for NoVs and by the 3’-end of ORF1 for

SaVs. Another capsid protein (VP2) is about 22-29 kDa for NoVs and 18 kDa for SaVs and is encoded by the 3’-end ORF for both NoVs and SaVs (63). It is a basic protein and thus it is also named small basic protein. The VP2 is likely a genome-linked protein or functions during virus assembly. In fact, a subgenomic RNA is supposed to be the template to efficiently express both VP1 and VP2 (Fig. 1.1). Although the VP2 is not essential for the formation of Norwalk virus VLPs (13, 251), it in cis enhanced the expression of VP1 and the yields of VLPs examined in baculovirus-expression system by using different recombinant baculovirus constructs containing both ORF2 and ORF3 or

9 ORF2 alone (12). Elimination of the expression of VP2 by site-directed mutation resulted in reduced VP1 levels similar to those obtained with the construct containing ORF2 alone.

This in cis enhancement of VP1 expression by VP2 was also confirmed in mammalian cells transfected with wild-type or ORF3-inactivated subgenomic RNAs of Norwalk virus.

Furthermore, VP2 increased VP1 stability and protected VP1 from disassembly and protease degradation.

Structural studies of NoVs relied on the expression of virus capsid proteins in a baculovirus-expression system (197, 198) because of the lack of a cell culture system for enteric NoV propagation. The insect cell expressed Norwalk virus capsid protein can self-assemble into two kinds of VLPs, 38 nm and 23 nm in diameter. The 38 nm-VLPs, which are the predominant ones compared to the 23 nm-VLPs, contains 90 dimers (180 copies of VP1) of the capsid protein exhibiting T=3 icosahedral symmetry. The 23 nm

VLPs probably contains 60 copies of the capsid protein exhibiting T=1 symmetry (251).

The precise location of the primary amino acid sequence of the 38 nm-VLP has been revealed through x-ray crystallography (Fig. 1.2) (197). The S domain forms the inner part of the capsid and surrounds the RNA genome, and the P domain forms the protruding part of a virion. The entire S domain (amino acids 1 to 225) corresponds to the

N-terminal region of the capsid protein that is relatively conserved among NoVs. The P domain corresponds to the C-terminal region of the capsid protein, which is more variable compared to the S domain. The P domain is further divided into P1A (amino acids 226-278), P1B (amino acids 406-520) and P2 subdomains (amino acids 279-405).

The P1A and P1B subdomains form the sides of the arch of the capsomeres and position the P2 subdomain at the top of the arch. The P2 subdomain corresponds to a region of the

10 capsid protein that shows the highest sequence variability among NoVs. The exposure of the highly variable region on the surface of the virion coincides with the functions of formation of major antigenic site(s) and receptor binding ligand(s) conferring host specificity and immunity (87).

Comparative structure analysis of the recombinant capsids of NoV (GI-1/Norwalk virus and GII-4/Grimsby), SaV (GI-2/Parkville) and vesivirus [(San Miguel sea lion virus serotype 4 (SMSV4)] showed that all these capsids consist of 90 dimers of the capsid protein sharing a common T=3 icosahedral symmetry (25). They also had a similar subunit organization consisting of an S domain and a P domain. The highly conserved S domain provides an icosahedral skeleton; the P1 subdomain, located between S and P2, provides additional fine-tuning to position the P2 subdomain; the P2 subdomain is located at the most distal surface of the subunit. The P2 subdomain shows the most prominent differences both in shape and in size among caliciviruses, corresponding to the observed highest sequence variability in this region. In the relative orientation between the S and P domains, the SaVs and vesiviruses are similar, whereas the NoVs are quite different. This data suggests that SaVs are closer to animal vesiviruses than to NoVs in structure.

The conserved amino acid motifs of the capsid protein of NoVs (Fig. 2.4) include

IDPWI (amino acids 52-56, IXPWI for Mu/MNV-1/03/US), LAGNA (amino acids 113-

117) and LYTPLR (amino acids 187-192) motifs in the S domain, the QNGR (amino acids 267-270) motif in the P1A domain and the RGD(/K) (RGD-like, amino acids 292-

294) motif in the P2 domain (81, 186, 240, 262). Among these conserved motifs, the

11 IDPWI motif is recognized by a NoV-specific broadly reactive monoclonal antibody

(MAb) 1B4, which reacts with 18 of 23 genotypes of NoVs (262).

The NoVs bind to human HBGAs which were found to influence the genetic

susceptibility of individuals to NoV infections (88, 94, 96, 97, 135, 151). The RGD-like motif forms the bottom of a binding pocket by computational structure analysis and is critical for virus binding to HBGAs as determined by site-directed mutagenesis of 2 strains having different HBGA-binding patterns (240). The NGR motif is essential for the capsid assembly and/or capsid protein expression (240).

1.2.4 Subgenomic RNA and recombination

Human NoVs and SaVs can not be grown in cell culture, so it has not been possible to demonstrate the presence of a subgenomic RNA in infected cells. However, detection of an RNA molecule of ~2 kb in stool samples from a human volunteer infected with Norwalk virus and detection of a 2.3 kb RNA from murine NoV MNV-1-infected

RAW 264.7 cells strongly suggests that NoVs produce subgenomic RNAs (104, 255). A

2.2 kb RNA was detected from an in vitro replication assay with the replication complex of SaV Cowden strain extracted from virus-infected cells suggesting that SaVs generate subgenomic RNA during virus replication (24).

However, whether the genomic and subgenomic RNAs of NoVs are co- encapsidated into one virion is still unknown. The results of Meyers et al. (163) suggested that both the genomic and the subgenomic RNAs of lagovirus RHDV were encapsidated into virions. The RHDV genomic and subgenomic RNAs isolated from infected rabbit liver tissue were resistant to RNase degradation which may be due to RNA packaging

12 into viral particles. When they performed sucrose density gradient centrifugation of liver

homogenates to separate particles containing either genomic RNA or subgenomic RNA

followed by Northern blot, they found that virions in the bottom fraction of centrifugation contained only the genomic RNA, while virions in the top fraction contained only the subgenomic RNA, but the virions in the middle fraction contained both genomic and subgenomic RNAs. These results suggested that the genomic and subgenomic RNAs are encapsidized into separate virions. The middle fraction probably comprised two kinds of virions and thus showed both kinds of RNA in the Northern blot.

Co-infections with GI and GII NoVs and co-infections by multiple genotypes of

NoVs is common in NoV infections (5, 60, 159). The RdRp-capsid junction region of caliciviruses contains a highly conserved ~20 nt motif in genomic and subgenomic RNAs that is suggested to be a transcription start signal (63, 119). This conserved nucleotide motif is identical within each genogroup of NoVs and SaVs except for the

Hu/NoV/GII/J23, probably facilitating homologous recombination during co-infection of a cell by different genotypes of NoVs or SaVs within the same genogroup. In fact, recombinants were identified in some human and bovine NoVs at the RdRp-capsid junction region, and the recombination occurs exclusively between NoVs within the same genogroup (81, 84, 98, 119, 141, 247). In comparison, only one SaV recombinant strain

(Mc10 or C12) of human origin was reported (118) probably due to the less extensive studies of SaVs than of NoVs.

A recent study (207) revealed that NoV recombination probably also occurs in the capsid gene. Seven potential recombinants [GI/1 (n = 1), GII/1 (n = 1), GII/3 (n = 1),

GII/4 (n = 3), and GII/5 (n = 1)] were detected from 94 NoV strains examined. Further

13 analysis showed that recombination sites were mainly located between the putative P1A

and P2 domains and/or within the P2 domain. These putative recombination regions

showed common features, such as sequence length and composition (upstream and

downstream GC- and AU-rich sequences, respectively). Also, the predicted RNA

secondary structure of this region demonstrated characteristics of homologous

recombination activators.

1.2.5 Evolution

Epidemic strains of NoVs have emerged spreading quickly and widely and

causing increased outbreaks as well as sporadic cases. In the mid 1990s, a single NoV

GII-4 strain was associated with 55% (60/109) of unrelated outbreaks in the US and its

emergence coincided with an increase in the NoV-associated illness reported (179). This

strain was also identified in 7 other countries located on 5 different continents over the

same period. Evidence showed that this virus first emerged in the UK and then it spread

widely internationally. Recently, another new GII-4 variant circulating in Europe was

responsible for 50% of the outbreaks in the UK (143). Sequence analysis revealed that

this new strain had a mutation in the polymerase gene.

A chronic NoV infection was reported in a severe immunocompromised patient,

but with intact humoral immunity (176). A NoV GII-3 strain accumulated 32 aa changes

including 11 aa changes in the capsid (1, 2 and 8 aa in the S domain, P1 and P2

subdomains, respectively). The amino acid changes in the P2 subdomain resulted in

predicted structural changes including disappearance of one helix structure. Because the

P2 subdomain is on the surface of the virion which is consistent with its suspected role in

14 the formation of a major antigenic site and in receptor binding, the amino acid mutations

in the P2 subdomain were probably driven by the host humoral immune-response.

In summary, the mechanisms of generation and emergence of a new or a widely

spread variant is not completely understood. It probably includes virus mutation and

recombination, virus subjected to specific immune selection pressure and virus changes

in response to different host receptor profiles.

1.3 GENERAL BIOLOGY

1.3.1 General biologic characteristics

Morphology. NoVs and SaVs are small, round, nonenveloped and 27-40 nm

(NoVs 28-35 nm, SaVs 32-40 nm) in diameter (6). Their characteristic morphological

feature is the appearance of 32 cup-shaped depressions on the icosahedral virion surface.

The SaVs possess a typical “Star of David configuration” (a six-pointed star with a dark

hollow in the center) having a feathery edge and distinct surface hollows that are round or

oval (147), and are morphologically indistinguishable from the vesiviruses (26). However,

the NoVs, which include most of the SRSV, lack the defined surface substructure by IEM

(29, 43, 112).

Physicochemical properties. The NoVs and SaVs have a buoyant density of

1.33-1.41 g/cm3 and 1.37-1.38 g/cm3 in CsCl, respectively (63). From results based on volunteers, NoV particles are resistant to the following treatments: (1) pH 2.7 for 3 h at room temperature; (2) 20% ether at 4˚C for 18 h; (3) incubation at 60˚C for 30 min; and

(4) 3.75-6.25 mg/L (ppm) of chlorine, which is the normal concentration of chlorine in

15 drinking water. However, NoVs lost infectivity when they were treated with 10 mg/L of

chlorine (a concentration used to treat contaminated water) and at alkaline pH (63).

Because human NoVs and SaVs are still non-cultivable viruses, their

physicochemical properties have not yet been fully determined. Data from vesiviruses

and lagoviruses is available and it may be referenced to predict some of the human

calicivirus properties (213): (1) sedimentation coefficient is 170-183S; (2) resistant to

lipid solvents, nonionic and cationic detergents and most iodine-based disinfectants; (3) inactivated by ultraviolet irradiation, by phenolic disinfectants, hypochlorite, aldehydes;

(4) most strains inactivated by sodium dodecyl sulfate (SDS). A comparative study of calicivirus resistance was performed by using 2 cultivable vesiviruses [canine calicivirus no. 48 (CaCV) and FCV] and a human NoV strain GII-4/DenHaag22/02/NL (41). The infectivity of the vesiviruses was examined by mean tissue culture infective dose (TCID50) and vesiviruses and NoV RNA was tested by using quantitative conventional and real- time RT-PCR. Although the RT-PCR detection of viral RNA does not mean the presence of infectious virus particles, there is a good correlation between the decline in infectivity and the decline of RNA tested using vesiviruses. However, when the viral infectivity was lost, the viral RNA still remained detectable. Compared to the 2 vesiviruses, the NoVs showed similar resistance to 0-100˚C treatment, ultraviolet-B radiation, 70% ethanol and sodium hypochlorite solutions, but they were more resistant to low or high pH.

Inactivation of the vesiviruses was time-dependent for temperature treatment and was dose-dependent for ultraviolet-B treatment. The commonly used 70% ethanol and the high concentration (> 300 mg/L) of hypochlorite for cleaning environmental contamination did not completely inactivate their infectivity.

16

1.3.2 Cell culture and infectious clones

Cell culture. Since the discovery of Norwalk virus, extensive efforts have been

undertaken to adapt human NoVs to cell culture testing at least 27 cell lines (42).

However, at present, no human NoV strain replicates in cell culture. Recently, a murine

NoV strain, MNV-1, replicated in cultured primary dendritic cells and macrophages and the virus growth was inhibited by interferon (IFN) αβ receptor and signal transducer and activator of transcription 1 (STAT1). An amino acid substitution in the capsid protein of serially passaged MNV-1 was associated with virulence attenuation in STAT1 knockout

(STAT1-/-) mice (255). Examination of the ultrastructure of infected cells revealed that virus particles localized within or next to single- or double-membraned vesicles, like the animal caliciviruses (65, 145, 229) and other positive-stranded RNA viruses (30, 194), and were present in the cytoplasm by 12 h post inoculation. The vesiculate areas increased in size with time, and by 24 h post inoculation, large numbers of these vesicles and viral particles occupied most of the cytoplasm, displacing the nucleus. In addition, a complete rearrangement of intracellular membranes occurred, leading to a rearrangement of the endoplasmic reticulum and loss of an intact Golgi apparatus. Interestingly, these smooth-membraned vesicles were often surrounded by mitochondria.

Porcine SaV Cowden strain is the only enteric calicivirus that has been adapted to cell culture. It was first adapted to primary porcine kidney cells in 1988 and then to a continuous porcine kidney cell line, LLC-PK cells in 1991 (51, 193). Sequence comparison of the complete genomes of the cell culture adapted-Cowden and wild-type-

Cowden revealed that the tissue cultured-Cowden has one silent mutation in the predicted

17 proteinase, 2 aa changes and a silent mutation in the RdRp, and one distant and 3 clustered amino acid substitutions and a silent mutation in the capsid (72). These amino acid substitutions may be associated with the cell culture adaptation of tissue cultured-

Cowden and its attenuated virulence in pigs (74). The virus antigens were detected in the cytoplasm of infected LLC-PK cells by cell culture immunofluorescence (CCIF).

Cytopathic effects (CPE) of tissue cultured-Cowden SaV in the primary porcine kidney and LLC-PK cells appeared 2-3 days post inoculation (DPI) and included rounding, separation and detachment of the cells (22, 51). A detailed ultrastructural study of the infected cells has not been performed.

Infectious clones. Because of the lack of a cell culture system for human NoVs and SaVs, infectious viral clones of human NoVs and SaVs may mimic the natural viruses during replication in permissive cells, thus providing a system for investigation of the mechanisms of virus replication. Among enteric caliciviruses, the only infectious clone available is the Po/SaV/Cowden strain (23).

In summary, the cultivable NoV/MNV-1 and SaV/Cowden and the Cowden infectious clones are useful models for studies of the pathogenesis, replication and immunity to human NoVs and SaVs. The MNV-1 replication in macrophage-like cells has opened a window for trying to adapt human NoVs to the macrophage/dendritic cell lines or cells with IFNαβ/STAT1 defects.

1.3.3 Molecular mechanisms of replication

Different types of cells may contain the specific receptors for NoVs as shown by the binding and internalization of VLPs into cells, such as the differentiated Caco-2

18 (human colon) cells. Despite the use of many cell culture supplements in the cell culture

medium, such as insulin, DMSO and butyric acid, NoVs still did not replicate in these

cells (42). The porcine SaV Cowden strain grows in both primary and continuous cell

lines but only in the presence of supplements of diluted filtrates of porcine intestinal

contents from uninfected gnotobiotic (Gn) pigs in the cell culture medium (51, 193).

Studies using this model revealed several important molecular mechanisms during SaV

replication (22, 24).

Chang et al. (22) investigated the relationship between the intestinal content

medium supplements and the growth of Cowden strain. Pretreatment of cells or the virus

with intestinal contents or transfection of viral RNA into cells did not induce virus

growth unless the medium was supplemented with the intestinal contents. Among

modulators of cell signal transduction, the G protein uncoupler, suramin, adenylate

cyclase inhibitor, MDL-12330A, and the cAMP-dependent protein kinase (PKA)

inhibitor, N-(2-[bromocinnamulamino]ethyl)-5-isoquinolinesulfonamide (NBEI) inhibited the effect of intestinal contents on virus growth for up to 72 h. These data indicate that Cowden virus replication may be dependent on an initial cAMP signaling pathway induced by intestinal contents.

In a later study using this model (24), the bile acids in intestinal contents were

identified as active factors promoting the growth of Cowden SaVs. During virus

replication, they induced an increased cAMP concentration in LLC-PK cells that was

associated with down-regulation of IFN-mediated STAT1 phosphorylation, a key element in innate immunity. In addition, cAMP/PKA pathway inhibitors, suramin, MDL12330A, or H89 suppressed bile acid-mediated virus replication. Thus the mechanism proposed for

19 SaV growth is dependent on bile acids, which are ubiquitous molecules present in the

intestine at the primary site of the virus replication. Sapovirus replication involves the

PKA cell-signaling pathway and a possible down-regulation of innate immunity. The

latter correlates the findings of murine NoV MNV-1. The MNV-1 was more virulent in

IFNαβγR-/- mice than in wild-type mice and it replicated better in macrophage and

dendritic cells from IFNαβγR-/-, IFNαβR-/- and STAT1-/- mice than in the cells from wild- type or IFNγR-/- mice (117, 255). These results indicate that NoVs and SaVs are IFNαβ-

sensitive viruses.

1.3.4 Virus assembly and apoptosis

Little is known about the virus assembly and release of NoVs and SaVs from infected cells. Recent studies of murine NoV MNV-1 strain in macrophage-like cells revealed that virus replication was associated with intracellular membranes, like the animal caliciviruses (65, 145, 229) and other positive-stranded RNA viruses (30, 194).

Several studies have been performed using the FCV, which belongs to genus

Vesivirus, as an example for calicivirus replication. During virus infection of kitten

kidney cells, membrane bound vesicles and accumulated virus particles were observed in

the cytoplasm by EM (145). The ultrastructural changes of a feline embryo continuous

cell line infected with FCV were studied (229). Single virus particles, extensive and non-

regular accumulations of virions or paracrystalline arrays of virus particles associated with microfibrils were found exclusively in the cytoplasm. Other cellular changes included rounded cells and nucleus and loss of pseudopodia of cells. There was extensive production of smooth-membrane bound vesicles in the cytoplasm, which were frequently

20 associated with the virus accumulations and paracrystalline arrays. The cisternae of the

endoplasmic reticulum and the space between the two layers of the nuclear membrane

were enlarged. In addition, nuclear chromatin underwent condensation and usually

changed into a single, rounded, central mass by Feulgen staining and light microscopy, as

well as electron microscopy, which suggested that apoptosis may occur during FCV

replication.

The virus assembly process for FCV has been monitored by labeling the virus-

specific proteins with 3H-leucine (125). First, the capsid precursors rapidly aggregated

into 5S subunits, then formed the stable 15S subunit component. Finally, the 15S subunits

and the viral genomes formed the 170S mature infectious FCV particles. This process is

rapid and within 30 min of the initiation of protein synthesis, the mature FCV particles

appeared in the infected cells.

Animal caliciviruses induce cell apoptosis or programmed cell death, during

infection contributing to the pathogenesis of disease. The lagovirus rabbit hemorrhagic

disease virus (RHDV) causes apoptosis of infected cells that may be a determinant in the development of the acute fulminant hepatitis and lethal hemorrhagic fever in rabbits (4).

The RHDV infects the liver with viral antigens expressed in the hepatocytes, macrophages and endothelial cells as determined by immunohistochemistry. Apoptosis of these cells was observed by in situ DNA fragmentation in the nucleus. In addition, the virus-induced apoptosis correlated with the extensive parenchymal destruction of the liver causing a lethal, acute fulminant hepatitis. Apoptosis of intravascular monocytes and endothelial cells was observed together with fibrin thrombi in blood vessels. Because apoptotic cells are known sites of enhanced procoagulant activity, apoptosis of these cell

21 populations might be the trigger of disseminated intravascular coagulation (DIC) and hemorrhagic fevers.

The FCV also triggers apoptosis within infected cells (3, 226). Replication of

FCV in cells resulted in translocation of phosphatidyl serine to the cell outer membrane, chromatin condensation and oligonucleosomal DNA fragmentation, which are the characteristics of apoptosis (226). In addition, this apoptosis involved cleavage of the viral capsid protein (3). The FCV capsid protein is synthesized as a precursor (76 kDa) that is post-translationally processed into the mature 62 kDa capsid protein by removal of the N-terminal 124 amino acids. The caspases, which are important pathways of apoptosis, were activated during FCV infection and this coincided with cleavage of the capsid to a 40 kDa fragment, which is related to the FCV capsid protein. The cleavage of the FCV capsid protein was prevented by caspase inhibitors in cell culture. In addition, an in vitro cleavage assay revealed that caspase-2 and 6 cleaved the capsid protein to generate the 40 kDa fragment using recombinant human caspases and FCV capsid proteins.

One comparative histopathological study based on diarrheic children after intestinal transplantation with or without human calicivirus infection revealed that apoptosis may also occur during human calicivirus infection (167), which is discussed in detail subsequently. However, whether apoptosis occurs during NoV and SaV infections has not been investigated in normal volunteers, animals or cell culture systems.

1.4 PATHOGENESIS

22 1.4.1 Studies in animal models

Attempts to establish an animal infection or disease model for human NoVs have failed in many animals including mice, guinea pigs, rabbits, kittens, baboons and monkeys (63). Chimpanzees inoculated with Norwalk virus responded serologically and shed soluble viral antigens, which are Norwalk virus capsid proteins but these capsid proteins do not self-assemble into virions, in their stools, but they did not develop illness

(260). Another animal model Macaca nemestrina developed clinical signs including diarrhea, dehydration and vomiting and fecal virus shedding was detected by RT-PCR post NoV/GII-3/Toronto inoculation through a nasogastric tube (231). Recently, Norwalk virus antibody-free rhesus macaques were found to be susceptible to Norwalk virus infection as three animals inoculated with Norwalk virus intragastrically shed virus (206).

One of the 3 animals shed virus from 1-19 DPI and developed serum IgM and IgG antibody responses to Norwalk virus. However, no diarrhea or other clinical signs were observed for these 3 animals. Thus, the host range of human NoVs may be limited to humans and non-human primates. However, the existence of similar animal NoVs (63,

117, 139) and SaVs (72, 73) suggests that interspecies transmission among humans and animals might have occurred in the evolutionary history of these viruses although little information of such transmission of caliciviruses is available.

Murine NoV/MNV-1 replicated in STAT1-/-, IFNαβγR-/-, IFNαβR-/- and IFNγR-/- mice, mice lacking protein kinase RNA-activated (PKR) or inducible nitric oxide synthase (iNOS) and wild-type mice (117). Only the STAT1-/- and IFNαβγR-/- mice were

more susceptible to fatal disease after MNV-1 infection compared to the wild type mice.

The clinical signs included encephalitis, vasculitis of the cerebral vessels, meningitis,

23 hepatitis and pneumonia after inoculation through different routes: peroral (p.o.),

intranasal (i.n.) and intracerebral (i.c.). The p.o. route is the physiological route for

human and animal NoV infections. The MNV-1 induced systemic infections which

differred from the gastroenteritis associated with human NoV infections. The MNV-1

infected wild-type mice did not show any clinical signs although viral RNA was detected in the intestine, liver and spleen at 1 DPI. These results indicated that MNV-1 replication in STAT1-/- or IFNαβγR-/- mice provides a small animal model for systemic NoV

infection but this small animal model does not mimic the disease induced by human

NoVs. Further studies of MNV-1 in gene knockout mice will provide important

information on host susceptibility and immunity to systemic NoV infections. However,

MNV-1 is not a typical enteric NoV. Other animals, such as pigs and calves, are infected

naturally with enteric NoVs and they provide more similar infection or disease models

for enteric NoVs.

The bovine NoV/GIII-1/Jena strain was discovered by EM examination of diarrheic stools from newborn calves in Jena, Germany in 1980 (71). It caused diarrhea in all of the 11 orally inoculated calves used for serial passage of this strain. The Newbury-2 strain was isolated from diarrheic calf fecal samples in the UK (257). The detailed pathogenesis of the bovine NoVs is discussed subsequently in 1.4.4 section. Because of the genetic relationships and the similar disease spectrum of bovine and human NoV infections, replication of bovine NoVs in calves should be a useful infection and disease model for investigation of the pathogenesis and immune responses of NoVs.

24 The porcine SaV Cowden strain replicates in pigs and provides an important

enteric disease model for SaV infections (74). The detailed pathogenesis of the SaV

Cowden strain is discussed subsequently in 1.4.4 section.

1.4.2 Host receptors and viral ligands

Viral infection is generally initiated by the binding of virus particles (ligands) to

specific receptors on the host cell surface. In the early Norwalk virus challenge study

with volunteers (192), one subset of individuals was repeatedly infected with Norwalk

virus, whereas the other set of individuals was never infected even after repeated

inoculation. One hypothesis for this phenomenon is that host genetic factors may

determine the susceptibility and resistance to Norwalk virus infection. It has now been

proven that the Norwalk virus attaches to potential host cells in the gut only if the

individual expresses specific, genetically determined carbohydrates (88, 94, 96, 97, 135,

151).

Histo-blood group antigens. Human red blood cells (erythrocytes, RBC) contain

ABO(H) blood group determinants. The ABH antigens are produced by enzyme

catalyzed sequential addition of specific monosaccharides onto terminal saccharide

precursors linked to glycolipid or glycoprotein carrier molecules. The A and B alleles

encode an A transferase and B transferase, respectively, and O alleles are nonfunctional.

However, the ABH antigens are not restricted to RBCs, but are widely distributed among

tissues as HBGAs. The ABH antigens are found on epithelial cells of the intestinal tract

of all terrestrial vertebrates (188). However, their appearance on RBCs is a recent event

25 in phylogenetic terms, because expression on this cell type is restricted to anthropoid apes

(261).

Four main types of ABH antigens (H type 1 to 4) have been found on human

RBCs or in secretions (188). The RBCs express exclusively H type 2 antigens under the control of the H or FUT1 gene. People who do not express H2 antigens on their RBCs are referred to as the Bombay phenotype, which is found on less than 0.002% of the population worldwide.

The expression of H type 1 antigens is under the control of the FUT2 gene, which is located within a single 999-bp exon and encodes a α-1, 2-fucosyltransferase. Generally, individuals having an active FUT2 gene secrete H type 1 antigens into saliva and on the surface of mucosal epithelial cells (202), and are referred to as secretor phenotype (Se+); individuals carrying mutated FUT2 genes either secrete H type 1 antigens weakly or do not secrete and are referred to as weak secretor phenotype (Sew) or nonsecretor phenotype

(Se-), respectively. At present, several mutations have been identified as the molecular

basis of Sew and Se- phenotypes (187): the A385T missense mutation corresponds to the weak secretion and is found at high frequency in Asian populations; G428A nonsense mutation represents >95% of the inactive mutations found in individuals of European and

African descent; other inactive mutations include C571T, C628T, G849A and

685delTGG (235). About 20% of Europeans are homozygous recessive for the inactivating mutations and have a Se- phenotype.

H type 2 antigens are also expressed in glands of both secretors and non-secretors

(151, 188). In some areas of the digestive mucosa of an individual with Se+ phenotype,

26 both H type 1 and H type 2 are expressed but in different cells. The expression of H type

3 and 4 antigens is independent of the FUT2 gene.

Another group of HBGAs, the Lewis antigens, are expressed under the control of

FUT3 gene (89, 95). They exist independently from the ABH antigens and are associated with secretor phenotypes. They exist as free oligosaccharides in milk and urine, or may be protein and lipid bound as part of glycoprotein or glycolipid. The RBCs do not express

Lewis antigens by themselves but acquire these antigens from plasma. The digestive tract is a major site for synthesis of the Lewis plasma glycolipids. Like ABH antigens, they are made by enzyme catalyzed sequential addition of specific monosaccharides onto terminal saccharide precursors. About 80-90% of Northern Europeans and Caucasian Americans are Lewis antigen positive (Le+), independent of secretor status. The Lewisa (Lea) and

Lewisx (Lex) are associated with non-secretor phenotypes and Lewisb (Leb) and Lewisy

(Ley) are associated with secretor phenotypes. For European Caucasian adults, 3 main

and 1 minor RBC Lewis phenotypes and their relationships to ABH secretor phenotypes

are: 1) 72% Le(a-b+) is (Le+, Se+); 2) 22% Le(a+b-) is (Le+, Se-); 3) 6% Le(a-b-) is (Le-,

Se+ or Se-); and 4) Le(a+b+) is (Le+, Sew). An infant genetically carrying genes

responsible for Le(a-b+) phenotype shows Le(a+b-) RBC phenotype first, then Le(a+b+)

before 2 years of age due to the fact that Lewis activity increases before secretor activity

and the competition for common substrates.

Human genetic susceptibility and resistance to NoV infections. A recent

Norwalk virus challenge study of 77 volunteers showed that none of the Se- individuals

developed an infection after challenge, regardless of the dose (135). These results suggest

27 that one genetic factor, the Se- phenotype, is associated with lack of susceptibility to

Norwalk virus infection.

Among ABO blood groups, group O individuals are more susceptible than group

A individuals and group B individuals are least susceptible to Norwalk virus infection, but not enough to completely prevent infection. These results correspond to an earlier epidemiological study of Norwalk virus infection and in vitro binding assays (88, 96,

151).

However, it appears that these observations about Norwalk virus can not be extended to certain other NoVs. For instance, the infection of volunteers with GII-2/Snow

Mountain virus was not associated with blood type, secretor phenotype or Lewis phenotype investigated (134).

Histo-blood group antigens may act as host receptors to NoVs. The human

NoVs and SaVs remain refractory to cell culture. However, many cells contain the

specific receptors for Norwalk virus. White et al. (250) found that Norwalk virus VLPs

bound specifically to a variety of cell lines (including ones of human and animal origin).

Norwalk virus VLPs bound to differentiated Caco-2 cells in the highest efficiency

compared to their binding to other cell lines. However, only 1.4~6.8% of the total VLPs

bound to these Caco-2 cells were internalized. Also, the amount of bound VLPs among

the different cell lines did not correlate with the tissue or species of origin, suggesting

that host range specificity may not occur at the level of cell binding. They further

localized the viral ligand of the capsid protein by an immunoprecipitation assay using a

MAb and truncated and cleaved forms of the capsid protein. The viral ligand was

28 localized within amino acid residues 300~384 within the P2 subdomain (amino acid

residues 279-405) which is externally exposed on the virus surface (197).

Recently, the Norwalk virus VLPs have been found to attach to the surface of the epithelial cells of the gastroduodenal junction and to bind to saliva, but only from secretor donors (151). These attachments and bindings were abolished by α-1, 2- fucosidase and inhibited by the H types 1 and 3 trisaccharides or by anti-H type 1 and anti-H type 3/4 antibodies. Transfection of CHO (Chinese hamster ovary) and TS/A cells with a α-1, 2-fucosyltransferase cDNA also allowed attachment of VLPs. These transfectants and differentiated Caco-2 cells expressing H type 1 antigens internalized the

VLPs. These results suggest that H type 1 and/or H type 3/4 antigens on the gastroduodenal epithelial cells of S+ individuals may be receptors or co-receptors for

Norwalk virus.

The carbohydrate-binding patterns of 6 other NoV VLPs have also been characterized by ELISA assays using individual saliva samples (94). Interestingly, the binding patterns of different strains were highly variable. Four different binding patterns were observed: (1) A/O (GI-1/Norwalk); (2) A/B/O (GII-4/VA387) and (3) A/B (GII-

5/MOH) recognized saliva from Se+ individuals, but 1 binding pattern (GII-9/VA97207) recognized saliva from Lewis-positive Se- individuals (94). These bindings were blocked by the MAbs to human HBGAs. The binding of these VLPs to synthetic oligosaccharides containing the Lewis and ABH antigenic epitopes confirmed that different strains of

NoVs may recognize different human HBGAs. However, it remains to be determined if such binding patterns correlate with the genetic classification of NoVs or if they are more strain-specific.

29 Whether the HBGAs act as receptors alone or if other co-receptors are necessary

for NoV infection is still unclear. Tamura et al. (239) investigated the nature of the cell

receptors for NoVs by using the VLPs of NoV/GII-6/Ueno7k/94/JP strain and

mammalian cell lines. They found that the binding between viruses and Caco-2 cells is a

protein-protein interaction determined by enzymatic and chemical modifications of Caco-

2 cell surface molecules. Furthermore, a 105-kDa cellular protein was found to be a

candidate receptor by using a virus overlay protein-binding assay. The binding of Ueno7k

VLPs to the 105-kDa cellular protein was also observed in six other mammalian cell lines.

These data suggests that the attachment of Ueno7k virions to mammalian cells is

mediated by a cellular protein, which is ubiquitously expressed on the mammalian cells.

In a more recent study from the same lab (237), the VLPs of GII NoVs (Ueno7k

virus, Chitta 76 virus and Kashiwa virus) and GI NoVs (Seto 124 virus and Funabashi virus) were found to bind the cell surface molecule heparan sulfate, which is a sulfated polysaccharide on the surface of most cells and a part of the proteoglycans. The binding efficiency was 10-fold higher for GII VLPs than for GI VLPs. Digestion with heparinase

I resulted in a reduction of Ueno7k VLP binding of up to 90% and 50% in undifferentiated and differentiated Caco-2 cells, respectively. These data indicated that heparan sulfate accounts for 90% and 50% of the binding molecules for undifferentiated and differentiated Caco-2 cells, respectively. In this study, they also found that pre- incubation of Ueno7k VLPs and HBGA H type 1 antigens reduced more than 40% of the binding of the VLPs to differentiated Caco-2 cells. The α1, 2-fucosidase reduced about

40% of the binding in a dose-dependent manner. These data suggest that the Ueno7k

VLPs probably possess separate ligands for heparan sulfate and H type 1 antigens on cell

30 surfaces. Also, the previously suggested 105 kDa cellular protein as a candidate receptor

molecule in various mammalian cell did not correspond to cell heparan sulfate (237).

In summary, nonsecretor phenotype individuals lacking H type 1 antigens on the

surface of intestinal epithelial cells are genetically resistant to Norwalk virus infection;

different NoV strains have different binding patterns to the HBGAs; a 105-kDa cellular

protein interacts with NoV/GII-6/Ueno7k strain; GI and GII NoVs bind to cell surface

heparan sulfate. To sum up, NoVs may bind to more than one kind of molecule on the

cell surface and different strains may bind to different molecules in order to attach to

cells and initiate infection.

1.4.3 Clinical features

Noroviruses. NoVs generally cause acute gastroenteritis with an incubation time of 10-51 h. The clinical symptoms observed in 38 outbreaks associated with NoVs included nausea (79%), vomiting (69%), diarrhea (66%), abdominal cramps (30%), headache (22%), fever (37%), chills (32%), myalgias (26%) and sore throat (18%) (115).

The illness is generally mild, and self-limited, and usually lasts 12 to 60 h. The diarrheic stools are liquid, and do not contain mucus, blood or leukocytes (39). It has been estimated that about 50% of people exposed to NoVs become ill (43). Clinical features observed in 31 of 52 volunteers who developed illness following Norwalk virus challenge were similar to those observed in the outbreaks (259). The incubation period was 10 to 51 h and illness lasted 24-48 h. Virus shedding detected by IEM was maximal around the onset of illness, and persisted up to 72 h in most cases following onset. When tested by more sensitive diagnostic methods, such as ELISA and RT-PCR, virus shedding

31 continued up to 3 weeks after the onset of illness in individuals with subclinical infection

(59, 205). Illness caused by Hu/NoV/GII-1/Hawaii/71/US and Hu/NoV/GII-2/Snow

Mountain/76/US in volunteers was clinically indistinguishable from that observed with

the Hu/NoV/GI-1/Norwalk/68/US. Asymptomatic infections with Norwalk virus or

Hawaii virus were observed in infected volunteers and during natural outbreaks (63).

Other NoV strains have a similar clinical spectrum with asymptomatic infections

observed (54, 63, 201). Chronic NoV infections with prolonged and more severe diarrhea

were reported in immunocompromised patients (63, 120, 121, 176).

Sapoviruses. The clinical symptoms for human SaV-associated acute, gastroenteritis include vomiting, diarrhea, nausea, malaise, aching limbs and headache

(166, 205), which were indistinguishable from those recorded in NoV infected patients.

Generally, SaV-induced gastroenteritis is less severe than NoV-induced illness (211). The incubation period was 12-72 h and illness lasted 1-11 days. The duration of detectable viral shedding was up to 14 days by RT-PCR. Subclinical infection was common in children (26, 67, 157). One clinical difference found between NoV and SaV infections is that SaV infection was mainly in children < 5 years of age, whereas NoV infection was common in all age groups (205). There is no report about chronic infection by SaVs.

The “Kaplan criteria” (115, 126) is considered highly indicative for calicivirus- associated gastroenteritis outbreaks: an incubation period of 15-50 h, presence of acute symptoms (including vomiting) in more than one-half of cases and/or diarrhea, average duration of symptoms of 12-60 h, a high attack rate and stool samples that test negative for bacterial pathogens.

32 1.4.4 Site of primary replication

The NoVs and SaVs enter the host through the oral route or possibly via vomit-

contaminated aerosols (63). These virions are acid stable, so they can survive passage

through the stomach and maintain infectivity until they reach the sites of replication.

They replicate in the epithelial cells of the upper intestinal tract because virus antigens have been detected in these cells by immunofluorescence (IF) although they have not been detected directly by transmission EM in the enterocytes (2, 27, 38, 52, 74, 112, 214,

215).

Human NoV infection and lesions. In Norwalk virus or Hawaii virus volunteer studies, biopsies of the jejunum from diarrheic individuals exhibited broadening and blunting of the villi (2, 38, 214, 215) although the mucosa was intact. Infiltration with mononuclear cells and cytoplasmic vacuolization, swollen mitochondria were also observed. Shortening and distorting of the microvilli were viewed by transmission EM although the epithelial cells were intact. Histologic lesions were not observed in the gastric fundus, antrum, or rectal mucosa of volunteers with Norwalk virus induced illness

(2).

The NoV RNA (GII-4/Miami Beach) was also detected in the ileal allograft by

RT-PCR from an intestinal transplant (IT) infant who was under immunosuppressive therapy (combination of corticosteroids, tacrolimus, sirolimus and basiliximab within the first postoperative month, followed by prednisone and tacrolimus for maintenance) and developed severe secretory diarrhea 178 days after IT that persisted for more than 120 days (120). Following a reduction in immunosuppressive therapy, diarrhea and enteritis remitted in association with the disappearance of all NoV RNA.

33 The levels of digestive enzymes changed during NoV infection of volunteers.

The small intestinal brush-border enzymes trehalase and alkaline phosphatase were

significantly decreased, a transient malabsorption of fat, D-xylose, and lactose was

observed, whereas adenylate cyclase activity in the jejunum did not increase following

Norwalk or Hawaii virus induced illness (2, 129). The levels of hydrochloric acid, pepsin,

and intrinsic factor in gastric secretions were not altered during Norwalk virus illness

(162). Gastric emptying was profoundly delayed in diarrheic volunteers who had the

typical jejunal mucosal lesion (162). This is probably the reason that nausea and vomiting

occur during NoV infection. The studies also showed that NoVs are mainly released from

the feces of infected individuals.

Bovine NoV infection and lesions. The Bo/NoV/GIII-1/Jena/80/DE caused diarrhea in all the 11 orally inoculated calves used for serial passage of this strain (71).

The original fecal sample from which the Bo/NoV/GIII-2/Newbury-2/76/UK was isolated also contained "astrovirus-like" viruses, which were separated from the

Newbury-2 NoVs by Gn calf passages (257). The Newbury-2 caused diarrhea in gnotobiotic calves with villous atrophy and D-xylose malabsorption, whereas the astrovirus-like agent did not cause diarrhea in two gnotobiotic calves.

A distinct bovine enteric calicivirus Nebraska/80/US (NB) strain was originally identified with the presence of a bovine coronavirus (BCV) in a diarrheic stool sample collected from a dairy calf (219). The NB virus has morphology similar to other noroviruses by EM. However, the VLPs of NB strain showed slight morphological differences from the VLPs of Bo/NoV/GIII-2/CV186-OH strain (Han M.K. and Saif LJ, personal communication), such as a little larger in size and short surface projections on

34 the NB VLPs. The NB virus was separated from the BCV through filtration of the stool

suspensions through 0.22 µm filters and passage in gnotobiotic calves exposed to BCV

previously and immune to BCV. The NB strain induced diarrhea in orally or i.v.

inoculated gnotobiotic calves. The infected calves developed intestinal lesions that were

most severe in the upper small intestine (duodenum and jejunum). No lesions were

observed in the lung, liver, kidney and spleen by histologic examination. The complete

genome of NB strain has been identified and may represent a distinct genus (NB-like)

within the family Caliciviridae.

Another unclassified bovine enteric calicivirus strain, Newbury agent-1

(Newbury-1) was found together with astrovirus in one calf fecal sample from an outbreak of diarrhea in the UK (18, 79). Later, the Newbury-1 and the astrovirus were separated from each other by calf passage. The Newbury-1 caused anorexia, diarrhea, and xylose malabsorption in gnotobiotic calves, whereas the bovine astrovirus was nonpathogenic in the calves. A detailed pathogenesis study of Newbury-1 has been performed with fourteen, 21-day-old gnotobiotic calves inoculated orally with Newbury-

1 (79). These calves were killed at 0.5-10 DPI and the histological changes and enzyme activities were examined by light microscopy, scanning and transmission EM, enzymology and xylose absorption assays, respectively. First, the infected enterocytes detected by immunoperoxidase staining appeared on the sides of villi at the base and the small intestine lesions occurred as early as at 0.5 DPI. Then the villi became stunted resulting from the exfoliation of the degenerate enterocytes. The most severe lesions were seen at 1 DPI. The lamina propria was compressed slightly, appearing more cellular, and macrophages were seen in lacteals. These small intestinal damages were restricted to the

35 anterior half of the small intestine and were almost repaired by 10 DPI. In the distal small

intestine, there was no virus-induced damage, but the villi were lengthened possibly due

to increased mitosis of crypt cells stimulated by enteroglucagon release during virus

infection. No virus particles were observed by transmission EM in the degenerate

enterocytes exfoliating from the villi, nor in the macrophages in the lacteals. Mucosal

beta-galactosidase activity decreased and xylose malabsorption occurred.

The Newbury-1 strain has not been classified because no sequence data is

available for this strain. It is suspected to be different from any of the identified bovine

NoVs in the family Caliciviridae because of several reasons. First, the size of Newbury-1

(36 nm) is bigger than that of bovine NoV Newbury-2 (33 nm) and Jena (30 nm) strains

and other noroviruses (28-35 nm). Second, the Newbury-1 was antigenically distinct

from the Newbury-2 by two-way cross-protection experiments in calves and solid phase

IEM (SPIEM) (18, 32). Third, the same research group successfully sequenced the partial

RdRp, complete capsid and VP2 regions of the Newbury-2 strain, but failed to amplify

Newbury-1 although several primer pairs detecting both Newbury-2-like and Jena-like

NoVs were used (186). The Newbury-1 is probably a NB-like calicivirus (personal

communication, Dr. Janice Bridger).

Murine NoV infections and lesions. A study of the murine NoV MNV-1 strain

in STAT1-/- mice demonstrated that MNV-1 specific staining was observed in the spleen

and liver of the mice on 2 DPI and the virus staining pattern corresponded to the distribution of macrophages and dendritic cells in the liver and spleen. They further cultured the MNV-1 in primary macrophages and dendritic cells (255). Because the

MNV-1 does not induce typical intestinal infections, even in the immunocompromised

36 mice (STAT1-/- or IFNαβγR-/-), this model does not mimic the human NoV enteric

infection. However, the authors suggested that macrophage-like cells may play an

important role during NoV infections, as virus carriers that transmit NoV from the gut to

blood, or vice versa.

Porcine SaV infections and lesions. The pathogenesis of SaVs was determined

using Gn pigs inoculated with porcine SaV Cowden strain (52, 74). The Gn pigs

developed mild to moderate diarrhea, which lasted for 2 to 5 days, following oral

inoculation with the wild-type-Cowden. Fecal virus shedding persisted for at least 7 days

as detected by both RT-PCR and enzyme-linked immunoassay (ELISA). The Cowden

virus particles were also detected from fecal samples by IEM. Mild to severe (duodenum

and jejunum) villous atrophy and fusion were observed corresponding to the moderate to

severe villous shortening and blunting in the duodenum and jejunum observed using

scanning EM. Cowden virus antigens were observed by IF in the villous epithelial cells

of the proximal small intestine, but not in the colon or extraintestinal tissues of any

inoculated pigs. No lesions were observed in extraintestinal tissues. These results indicate

that porcine SaV Cowden strain replicates in the epithelial cells of the small intestine.

However, in one of the same studies, the researchers found that Gn pigs that were i.v. inoculated with the Cowden strain had similar histopathological lesions to those in pigs that were orally inoculated. Compared to oral inoculation, i.v. inoculation had an incubation period 1-2 days longer and induced more severe villous atrophy in the jejunum. More interestingly, viral RNA and antigens were also detected from the serum samples of 7 of 9 orally inoculated pigs. These virus-positive acute-phase serum samples were infectious as confirmed by oral or i.v. inoculation of 4 additional Gn pigs. These 4

37 pigs developed diarrhea, small intestinal lesions, virus shedding in the feces, and

seroconverted to Cowden strain. This data suggests that after the infectious virions enter the blood stream (viremia) during infection. They can enter the intestinal epithelial cells through both the apical surfaces and potentially through the basal epithelial surface via the blood stream. How and through which kind of cells the virions in the blood stream enter the enterocytes is still unknown because the crypt enterocytes were virus-negative by IF staining. How the virions in the intestine reach the blood stream is also unclear.

Whether the mononuclear cells, which infiltrate the small intestine during virus infection, are responsible for such viral transmission remains to be investigated.

Apoptosis occurs during the animal calicivirus RHDV and FCV infections contributing to the pathogenesis of diseases. One comparative histological study based on diarrheic IT children with or without human NoV infection revealed that apoptosis may also occur during human NoV infection (167). The intestinal biopsies of 13 IT recipients were collected. Five patients with high-volume diarrhea were diagnosed with human

NoV (GII-4/Miami Beach) enteritis by RT-PCR followed by sequencing of the RT-PCR products. Controls were 8 pediatric IT recipients with high-volume diarrhea but calicivirus negative by RT-PCR during the same time period. Compared to the control group, the disarray and apoptosis of the surface epithelial cells were characteristics of

NoV enteritis, whereas the glandular apoptosis was an overlap feature of NoV infection and mild acute cellular rejection.

In summary, enteric NoVs and SaVs replicate in the proximal intestinal tract causing histopathological changes in intestine epithelial cells, decreasing the level of digestive enzymes and prolonging gastric empting. Viremia occurs during SaV infection

38 and virions can enter the intestine through 2 different ways: the apical surface and the blood stream. Whether enteric NoVs replicate only in intestinal epithelial cells or also in marcrophage-like cells needs to be further investigated in human volunteers and animal models of enteric NoV infections.

1.4.5 Virulence

Little is known about human NoV virulence. The factors responsible for the

emergence of an “epidemic” strain are unknown. Recently, a new GII-4 variant

circulating in Europe was responsible for 50% of outbreaks in the UK (143). Sequence analysis revealed that this new strain had a mutation in the polymerase gene.

A comparative pathogenesis study was performed in Gn pigs with wild-type and

tissue cultured-Cowden SaV (74). Compared to the wild-type-Cowden that induced mild

to moderate diarrhea in pigs, the tissue cultured-Cowden induced subclinical infection.

Sequence comparisons of the complete genomes of the tissue cultured-Cowden and wild-

type-Cowden revealed that the former has 2 aa changes in the RdRp and one distant and 3

clustered amino acid substitutions in the capsid, except for silent mutations (72). These

substitutions are associated with attenuation in pigs (74). However, which mutation is

critical for virulence is still unclear. Hopefully use of site-directed mutagenesis of

infectious Cowden SaV clones will resolve this question in the future.

1.5 LABORATORY DIAGNOSIS

The etiological diagnosis of illness caused by enteric caliciviruses is based on laboratory diagnosis as well as clinical aspects. Laboratory diagnosis focuses on the

39 detection of viral antigen, viral RNA and serum antibody responses. The EM and

antigen-ELISA detect intact virions and viral antigens, respectively, RT-PCR and real- time PCR detect viral RNA, and recombinant VLPs have been widely used as antigens for antibody-ELISA detection of serum antibodies.

1.5.1 Electron microscopy

The NoV and SaV prototype strains, Norwalk virus and Sapporo virus, were first detected by EM or IEM (26, 114). The EM can detect new viruses and mixed infections by different viruses. Therefore, it is still used to screen fecal samples for enteric viruses in the public health laboratories worldwide (6, 55, 200). The EM performed using specific antiserum to aggregate virus particles is called IEM; otherwise it is called direct

EM.

The direct EM is not a very sensitive technique with a detection limit of 106 particles per ml of stool for enteric viruses (6). Compared to SaVs, NoVs do not have a typical calicivirus morphology and they are very hard to be differentiated from many other small round fecal viruses including enteroviruses, astroviruses and picornaviruses in stool specimens (43, 203). In many labs, fecal samples are prepared as 10 to 20% suspensions before being clarified by low-speed centrifugation. Afterwards, concentration and purification steps are usually necessary before EM, such as ultracentrifugation or semi-purification through sucrose cushions. Virus particles are negatively stained using electron dense stains, such as phosphortungstic acid and uranyl acetate.

40 The IEM can usually increase the sensitivity of EM up to 10 or more-fold (114,

132). One modified IEM procedure is called solid phase IEM (SPIEM) which is a capture

ELISA performed on an EM grid (133). These two methods are generally used to examine NoVs and SaVs in fecal samples because one can observe both the viral morphology and the antigen-antibody reaction. Compared to SPIEM and direct EM, one pitfall of IEM is that the antibodies can obscure the characteristic surface appearance of viral particles and mask virus particles if antibody is in great excess (31, 43). The specificity of IEM and SPIEM depends on the quality of the antiserum used in the experimental procedures. The disadvantages of EM include the need for highly skilled experts and expensive equipment as well as the time consuming process to examine individual specimens. Therefore, it is not a practical method for investigation of large numbers of samples.

1.5.2 RT-PCR and real-time RT-PCR

Nowadays, RT-PCR is the primary assay for detection of NoV and SaV nucleic acid because it is the most sensitive technique and a commercial broadly reactive ELISA is not available for detection of NoVs and SaVs. Several factors affect the sensitivity and specificity of RT-PCR assays, including the quality of the sample, the methods used for

RNA extraction and purification, the primers, RT-PCR conditions and the methods for detection of RT-PCR products. The primers mainly determine the specificity of the RT-

PCR reaction. Although nested RT-PCR can increase sensitivity up to 10-1,000 times, it is no longer suggested because of the high risk of cross-contamination (6). Real-time RT-

41 PCR is more sensitive and it has become an alternative method for detection of low

amounts of virus, especially from environmental samples.

Extraction of RNA. Two major aspects should be considered in the selection of an RNA extraction method: efficiency of RNA recovery and ability to remove RT-PCR inhibitors. Jiang et al. (102) evaluated a number of different methods for the removal of

RT-PCR inhibitors and they found that cetyltrimethylammonium bromide (CTAB) partially removed inhibitors. However, this method is time-consuming including concentration of samples by precipitation with polyethylene glycol (PEG) and proteinase digestion and thus it is not applicable for large-scale epidemiological studies. At present, the most accepted RNA extraction method for fecal samples is a mixture of guanidinium thiocyanate (GTC), phenol and chloroform extraction and precipitation of RNA by alcohol (6). The principal reagent is commercially available from several companies, such as TRIzol from Invitrogen (Carlsbad, CA). Modifications of the GTC method have

been reported using silica particles to absorb released RNA followed by washing and

elution of RNA. This method proved to be better than the PEG-CTAB method in the

extraction of NoV RNA (77).

Antibody capture of NoVs and then heating to release the viral RNA from

captured virus particles is a simple method adapted from that for detection of hepatitis A virus. It can concentrate and purify virus particles from large volumes of environmental samples containing low amount of viruses (56, 217). The limitation of this method is similar to that of ELISA: NoV hyperimmune serum is usually highly specific and only detects homologous strains. With NoV common epitope-specific MAbs available in the future, this method may be evaluated in more detail and applied to field studies.

42 Primer selection. The genetic diversity of NoVs and SaVs has made it very

difficult to select a single primer pair to detect all NoVs or SaVs. The majority of primers

have been designed based on the most conserved RdRp region of the genome (6).

However, sequence analysis of the polymerase region of a wide range of virus strains

indicated that this region is also variable with the nucleotide identity as low as 53%

between strains of different genogroups and 60-64% within genogroup (247).

Calicivirus “universal” primers based on the most conserved RdRp motifs have

been designed. The primer pair p290 and p289 (or p110) targets the conserved motifs

“DYSKWDST” and “YGDD” of the RdRp region of caliciviruses (99, 128). Compared

to p289, p110 is 2 nt shorter at the 5’-end and it is degenerate. The primer pair p290/289

amplifies human GI and GII NoVs (99), bovine GIII NoVs and NB-like enteric

caliciviruses (220), human, porcine and mink SaVs (49, 73, 99) and mink vesiviruses

(73) . New human SaVs have been identified by using this primer pair to screen for new

viruses from a large number of samples (49). Because these primers have nucleotide

mismatches for both NoVs and SaVs, RT-PCR is performed at relatively low stringency conditions, such as low annealing temperature (99). Although the detecting spectrums of these universal primers are wider, they are less sensitive for specific strains. For example,

Smiley et al. (220) found that the primer pair p290/289 was less sensitive than the bovine

GIII NoV-specific primers for detection of the bovine GIII NoVs. In addition, the specificity is lower because the reverse primer targets the YGDD motif, and especially the GDD motif is also conserved among other RNA viruses (16). A recent study reported that primer pair p290/289 also amplified rotaviruses (146). Therefore, sequencing of the

43 RT-PCR products of p290/289 (or p290/110) or performing hybridization with an

internal specific probe is necessary to prevent false-positive results.

A commonly accepted approach is to design and use several primer pairs for

detection of NoVs and SaVs. Generally, primers designed based on the sequence of

locally circulating strains have performed better than other primer pairs (210); primers

targeting nonpolymerase regions of the viral genome are less broadly reactive; very

specific primers are more sensitive than less specific primers for a specific strain. For example, Le Guyader et al. (128) reported 10- to 1000-fold differences in the quantities of several NoV strains could be detected by two primer pairs.

Internal control (IC) RNA. The RT-PCR inhibitors are common and sample- dependent in food, shellfish and fecal samples (6, 43) causing false-negative results.

These inhibitors may include body fluids, food and bacterial cell constituents (253). An

IC RNA co-amplified with the target viral RNA can be used to monitor for inhibitors as well as technical errors. Diluting the extracted RNA of such samples or further purifying the extracted RNA and performing RT-PCR again usually decreases the false negative results. Diluting the extracted RNA is simplest and most samples diluted 1:10 or 1:50 will no longer inhibit the RT-PCR reaction (43). Stool samples from Norwalk virus infected volunteers have been shown to be positive at dilutions ranging from 1:10 to 1:

10,000 (59). A drawback of this strategy is that dilution of weakly positive samples may exceed the detection limit of RT-PCR and therefore become negative.

There are two kinds of internal control RNAs: competitive and noncompetitive.

The competitive internal controls are suggested because the target virus RNA and internal control RNA are amplified in the same RT-PCR condition. The approaches of how to

44 generate such an internal control have been well reviewed elsewhere (93). When using

the competitive IC RNA, there are three possible outcomes of RT-PCR assays: (1) no

product (the assay has failed-inhibition may be present); (2) the IC RNA, but not the

virus fragment is obtained (the assay has worked, and the sample is most likely negative

or viral RNA is below the detection limit); (3) the virus fragment, with or without the IC

RNA fragment, is obtained (the assay has worked, and the sample is positive).

Confirmation of RT-PCR products. The simplest method for detection of RT-

PCR products is agarose gel electrophoresis. If a sample shows a band having a similar size to that of the positive control amplicons, it will be considered positive. However, it can produce false-positive bands and such nonspecific bands are frequently seen for NoV and SaV RT-PCR from fecal samples (6).

The use of a hybridization assay is probably the cheapest approach to interpret

and confirm RT-PCR products. The most commonly used hybridization assays include

dot or blot hybridization, liquid hybridization and Southern blot hybridization. Nowadays,

the time required to perform a hybridization assay can be shortened without loss of

sensitivity by using microwell hybridization assays (16, 159). Compared to sequencing of the amplicons, it is less time consuming and cheaper to confirm the specific amplicons, especially in epidemiological studies testing of large numbers of samples.

However, the limitation of hybridization assays for NoVs and SaVs is that the

genetic variability of NoVs and SaVs makes it difficult to select a single probe to detect

most if not all possible NoV and SaV strains (128). Although internal probes can detect strains with several nucleotide mismatches under hybridization conditions with lower stringency, strains with highly variable internal sequences may be missed. Consequently,

45 gel electrophoresis followed by sequencing of RT-PCR products is still the gold standard

to confirm NoV or SaV specific amplicons.

Real-time RT-PCR. In the past 2-3 years, real-time RT-PCR has been used for detection of NoVs from fecal and shellfish samples (69, 92, 109, 110, 142, 164, 177, 189).

Compared to conventional RT-PCR, the real-time RT-PCR is more sensitive, faster and quantitative if RT-PCR inhibitors are not present in the sample (142). It also avoids the

post RT-PCR confirmation steps, such as gel electrophoresis and hybridization. The

sensitivity of a LightCycler real-time RT-PCR has been reported to be 10,000 times more

sensitive than the conventional RT-PCR, and the detection limit is 5 to 5 × 106 copies of

RNA per reaction (189). The disadvantage of real-time RT-PCR is mainly the expense.

1.5.3 Immunoassays

Immunoassays detect both viral particles and soluble antigens. First radio

immunoassay (RIA) and subsequently ELISA were developed and used for large scale

epidemiological studies (66, 91, 149, 170). Initially human serum samples from

volunteers were used as capture and detection antibodies. Now hyperimmune antisera are

mainly produced using recombinant NoV VLPs. This has greatly improved the large

scale investigation of NoV and SaV associated gastroenteritis because the VLPs can be

produced in large quantity and in high purity. The ELISA has become the most popular

immunoassay to detect the viral antigens of NoVs and SaVs because it has less technical

requirements, it is fast and it is cheaper to perform.

46 The antigen ELISA for detection of NoVs has proven to be sensitive with a

detection limit of 0.025 ng of capsid protein (59, 101), with 1:10,000 dilutions of viral

antigens in the stools of volunteers still being detectable (59). The sensitivity of ELISA was similar to that of traditional RT-PCR probably due to the large amounts of NoV

soluble proteins in stools (6, 43).

The hyperimmune antiserum produced to VLPs is often highly specific to the

homologous strain or viruses within the same genotype (59, 76, 101, 111, 123, 182). For example, the ELISA using hyperimmune serum against NoV/GII-3/Mexico strain did not detect NoV/GII-4/Grimsby virus, and vice versa (76). The reason is probably that the most variable region, the P2 subdomain is on the surface of the VLPs and contains the immunodominant epitopes. The most conserved N-terminal/S domain is either buried inside and can not induce antibodies after inoculation of the animals with VLPs, or the S domain contains the common epitopes that are not immunodominant (43, 197). However, the high specificity of these ELISA assays becomes a disadvantage during screening of

NoV and SaV antigens in epidemiologic studies because the circulating strains are highly variable.

Search for common epitopes. If all NoVs or SaVs, or at least each genogroup of NoVs or SaVs contains one or more common epitopes, the development of a broadly reactive antigen-ELISA will be possible. Many researchers have worked on this aim and recent findings are very promising.

Broadly reactive MAbs against GI or GII NoVs have been generated. One MAb

was used as the capture antibody with a polyclonal antibody as detector antibody in an

antigen-ELISA and this assay detected 9 of 15 GI-positive (RT-PCR positive) fecal

47 samples representing 4 of 5 genotypes tested (78). Kitamoto et al. (122) generated 20

MAbs from mice immunized orally with either a single genotype of 4 NoV VLPs, one strain of SaV VLPs or with mixtures of two types of VLPs from different genogroups.

The MAbs were characterized by Western blotting and ELISA and were classified as strain-, genogroup-, or genus-specific MAbs to NoVs and SaVs. Of interest, the first reported NoV common MAbs were obtained from mice immunized with only one type of

NoV VLPs which means common epitopes exist among viruses within the Norovirus genus. These common epitopes can be used to develop a cross-reactive diagnostic assay for detection of NoVs. Two MAbs that reacted exclusively with SaV VLPs are SaV- specific and are the first reported MAbs for SaVs (122).

One rabbit hyperimmune sera produced by using the recombinant capsid proteins

of a GII NoV expressed in E. coli recognized both GI and GII viral recombinant capsid

proteins expressed in E. coli by Western Blot and ELISA, although the cross-reactivity

was weaker than the genogroup-specific reactivity (263). Whether these capsid proteins form VLPs is unknown. The N terminal/S domain of these capsid proteins may be exposed and induce antibodies against common epitopes.

Recently, two commercial ELISA kits for detection of NoVs in stool samples have been evaluated with a panel of 103 stool samples containing 4 genotypes of GI and

10 genotypes of GII NoVs and 39 stool samples containing other enteric viruses (21).

The sensitivities and specificities of the two kits were tested. One kit had a high sensitivity (>70% for 10 of the 14 subgroups) but a specificity of only 69%, and the other kit had a low sensitivity (<30% for 6 GII subgroups) but a high specificity of 100%.

48 Immunoassays for detection of SaVs have a similar problem. The SaV antigen-

ELISA was quite strain-specific using hyperimmune antiserum raised against the purified viruses from the stools of infected young children or VLPs produced by a baculovirus- expression system (85, 172). An antigen-ELISA for detection of porcine SaVs has been developed by using hyperimmune antiserum to the VLPs of the Cowden strain. It has been used successfully for detection of the Cowden strain from experimentally infected

Gn pigs or cell-culture supernatants (24, 74).

1.5.4 Detection of serological responses

About 50% or more of the individuals involved (or examined) in a NoV- associated outbreak demonstrate an antibody response which is strong evidence to indicate the cause of an outbreak (116). However, in volunteer studies of Norwalk virus, individual serological responses are not in 100% agreement with virus infection. For example, a number of volunteers who were clinically ill did not seroconvert to Norwalk virus (91).

In earlier serological studies, ELISA assays were developed by using virus-rich fecal extracts as antigens (6). Later, the detecting antigens were replaced by baculovirus expressed VLPs. Generally these antibody-ELISA assays are specific and sensitive for detection of the VLP-specific antibodies and thus they have been used in several large- scale epidemiologic studies (28, 35, 36, 48, 61, 106, 191). However, these antibodies are usually cross-reactive, detecting antibody responses to both GI and GII NoVs in volunteers (59, 165, 243), making it impossible to identify the antigenic type of the infecting strain (178).

49 Antibody-ELISA assays for detection of antibodies against bovine GIII-1 and

GIII-2 NoVs or porcine GII NoVs have been developed recently using the VLPs or

capsid proteins of bovine or porcine NoVs, respectively (35, 48, 82, 252). Similarly, these antibody-ELISA assays are usually cross-reactive. For example, the VLPs of

Po/NoV/GII/Sw918 strain cross-reacted with the antibodies against human GII (Hawaii and Mexico) (48).

1.6 EPIDEMIOLOGY

1.6.1 Outbreaks, sporadic cases and asymptomatic infections

NoVs and SaVs are emerging enteric pathogens that cause gastroenteritis in humans and animals (26, 63, 73, 75, 139, 143).

Human NoVs. Human NoVs are estimated to cause 23 million cases of illness annually in the US (161) and more than 90% of nonbacterial epidemic gastroenteritis worldwide (63). In the US, between 1996-1997 and between 1997-2000, NoVs were detected from fecal samples in 96% (86/90) and 93% (217/233) of the outbreaks of nonbacterial gastroenteritis, respectively, by RT-PCR or RT-PCR and direct EM (45, 46).

The GII NoVs were the predominant (73%) strains between 1997 and 2000. Certain GII clusters (GII/1, 4) were more commonly associated with outbreaks in nursing home settings than with outbreaks in other settings (45). Investigations performed in other countries also showed that NoVs are the primary cause of nonbacterial gastroenteritis outbreaks and GII NoVs were found more frequency in outbreaks than GI NoVs (47, 58,

143, 150, 154, 181, 190, 246). In communities, NoV infection accounts for 7-11% of

50 gastroenteritis in Europe and has become the most commonly diagnosed cause of

infectious gastroenteritis (34, 241, 249). In children, NoVs were detected from 20%

(230/832) of diarrheic fecal samples from Finish children from 2 months to 2 years old

by RT-PCR (190). The NoVs have emerged as the second viral cause, next to rotaviruses

(29%), of sporadic diarrhea in children. In hospitalized acute diarrheic children, whose

diarrhea was more severe, NoVs were detected from 7.1-7.6% cases (199, 265).

In certain years, increased outbreaks of NoVs are usually associated with the

emergence of genetically variable strains. For example, collaborative investigations

including ten European countries (143) showed that a striking increase and unusual

spring and summer peak of norovirus gastroenteritis in 2002 coincided with the

emergence of a novel GII-4 strain, which had a mutation in the polymerase gene.

Human SaVs. Compared to NoVs, the epidemiological studies of SaVs are limited due to a limited supply of available reagents for diagnosis and the fact that SaVs usually do not cause large scale food- and water-borne outbreaks. The SaV-associated gastroenteritis outbreaks and sporadic cases usually occurred in younger children although one outbreak and a few sporadic cases in adults have been described (49, 180).

After the discovery of the SaV prototype Sapporo virus in 1977 in an infant home in Sapporo, Japan, the antigenically and genetically identical Sapporo virus caused

3 more outbreaks in the same infant home during 1977 and 1982 (26). Between 1976 and

1995, SaVs accounted for 6 (16.7%) of 36 gastroenteritis outbreaks in this infant home, less than human group A rotavirus (10/36, 27.8%) and NoVs (8/36, 22.2%), but higher than astroviruses (2/36, 5.6%) and adenoviruses (3/36, 8.3%) (174).

51 Human SaVs are associated with 1.8-9% of the cases of pediatric acute

gastroenteritis worldwide (47, 124, 138, 156, 173, 175, 184, 190, 205, 256, 258). In the

community, SaVs were detected by RT-PCR from 6.2% (48/772) and 1.4% (11/765) of

diarrheic and normal control subjects, respectively, in a cohort study performed in the

Netherlands (205). The SaV infection was mainly restricted to children <5 years of age

(175, 205). A cohort study of children in Mexico reported that SaVs were detected from

7.6% and 2.8% of diarrheic and non-diarrheic samples, respectively, by RT-PCR followed by sequencing of RT-PCR products (47). In hospitalized acute diarrheic children, SaVs were detected from 0.2-1.4% of the cases (83, 265).

Mixed infections of NoVs and SaVs were reported in Japan and Thailand (26, 70).

Mixed infections of NoVs and other enteric viruses are common, particularly with rotaviruses and less frequently with adenoviruses and astroviruses (183, 196, 230).

Animal NoVs and SaVs. The prevalence of bovine GIII-1 (Jena-like) NoVs was

8.9% (34/381) of diarrheic samples collected from dairy herds in three areas of Germany during 1999-2002 by an antigen-ELISA using the VLPs of Jena strain (35). The GIII-2 bovine NoVs (Newbury-2-like) were endemic in cattle in The Netherlands by RT-PCR, with an overall prevalence of 31.6% (77/243) of pooled veal calf samples and 4.2% of the individual dairy cattle samples (13/312) (244). The GIII-1 and GIII-2 bovine NoVs have been detected from 53-62% (43/75) of veal calf fecal samples from 2 commercial Ohio farms by RT-PCR using one primer pair detecting both GIII-1 and GIII-2 bovine NoVs

(220). They occurred in 33-75% of diarrheic stool samples from calves on all 8 Michigan and 2 of 14 Wisconsin dairy farms tested by RT-PCR with another pair of GIII bovine

52 NoV-specific primers (254). No prevalence study has been done for porcine or murine

NoVs and porcine SaVs.

Seasonality. Although human NoV transmission occurred year-round in most surveys, generally a cold weather peak was found for both sporadic cases and outbreaks identified in The Netherlands, England and Wales, Japan, the US, Australia, Canada and

Denmark, and especially in Japan, Canada, and The Netherlands (144, 168). Seasonality did not appear to be related to the detection method used, age group, or suspected mode of transmission. Sometimes atypical seasonal peaks occurred probably due to the emergence of new NoV strains (15, 143).

A study of gastroenteritis outbreaks between 1976 and 1995 in the infant home

in Sapporo, Japan, from which the SaV prototype Sapporo virus was detected, showed

that NoV- and SaV-associated gastroenteritis outbreaks occurred almost every year, but

with no clear seasonality, which was in contrast to the outbreaks caused by group A

rotaviruses that occurred between October and April and peaked from December to

March (174).

One research group reported that oysters preferentially accumulated F(+)

coliphage, an enteric viral surrogate, to their greatest levels from late November through

January (19). This indicates that the winter seasonality of shellfish-related illnesses by

enteric viruses probably results from seasonal physiological changes of oysters that may

affect their ability to accumulate viral particles from waters.

Transmission and populations at risk. The human NoVs and SaVs are transmitted mainly by the fecal-oral route. Viruses can be spread through direct person-to person contact, contaminated food, water, or aerosol droplets (63). The distribution

53 pattern of 86 outbreaks caused by human NoVs in the US during one year was as follows:

43% occurred in nursing homes and hospitals; 26% in restaurants and catered meals; 11% in schools and day care centers; 11% in vacation settings (including cruise ships); 6% from oyster consumption and 3% from other settings (46). The percentage of NoV transmission models were: 21% foodborne, 11% person-to-person, 6% oyster consumption, 3% waterborne and 43% no data available. Mead et al. (161) estimated that as high as 40% (9.2/23 million) of illnesses caused by NoVs are foodborne. The SaVs cause outbreaks mainly in infants, especially in infant homes (26).

Airborne transmission of NoV in a hotel restaurant has been reported (152). The

low infectious dose, winter seasonality and rapid rate of spread of NoVs also supports

that aerosol transmission is important (168).

The NoV infection was common in all age groups (166, 205). However, the

elderly in nursing homes, children in day care centers and schools, customers in

restaurants, people in vacation settings, travelers, and soldiers are at a higher risk for

NoV infection (63). The SaV-associated diarrhea occurred more frequently in children

than in adults and was mainly restricted to children aged < 5 years (166, 205).

Nosocomial infections by NoVs and SaVs have been also reported. In a study of

children and adults hospitalized with gastroenteritis in a hospital in France during a 2-

year period, researchers found that 37.5% (9/24) of NoV infections were acquired in the

hospital based on the date of admission and the date of illness (242). A study in a

children’s hospital in Sweden from 1987-1992 revealed that 78% (25/32) of the episodes

of calicivirus infection were nosocomial (228). Other nosocomial infections have also

been reported (53, 227).

54 Recently investigators found that among ABO blood groups, group O individuals

are more susceptible than group A individuals, and group B individuals are least

susceptible to Norwalk virus infection but not completely resistant to prevent infection

(88, 96, 151).

1.6.2 Seroprevalence

Human NoVs. An early comparative seroprevalence study of Norwalk virus and rotavirus in infants, children and adults in the US (113) reported that antibodies to rotavirus are acquired rapidly during infancy and early childhood and by 36 months of age over 90% of individuals have rotavirus antibodies, whereas Norwalk virus antibodies are gradually acquired during childhood, then more quickly during the adult years and by

50 years of age about 50% of individuals have Norwalk virus antibodies. A study of a population in London, England using Norwalk virus VLPs showed similar results (191).

Cubitt et al. (28) compared the seroprevalence of IgG antibodies to recombinant

VLPs of Norwalk virus, Hawaii virus and Mexico virus in 338 children in London.

Infections with GII viruses (Hawaii virus and Mexico virus) were prevalent and occurred earlier in life than GI Norwalk virus infections. They also performed a serological study of 566 Canadians between 9-79 years of age using Norwalk virus and Hawaii virus VLPs, and found that the prevalence of antibodies increased with age from 53-100% for

Norwalk virus and from 65-100% to Hawaii virus.

However, seroprevalence studies in developing countries demonstrated a different pattern of NoV antibody acquisition. In a longitudinal study of young children in rural Bangladesh, Black RE et al. (14) found that the prevalence of antibody to

55 Norwalk virus was 7% in children younger than six months and increased to 80% in

children 2-5 years of age. A recent seroprevalence study for Norwalk virus and Mexico

virus in Beijing, China (106) using Norwalk virus and Mexico virus VLPs as antigens in

antibody-ELISA assays showed that the seroprevalence is high in neonates for both

Norwalk virus (99%) and Mexico virus (94%), then it decreased to 41% (Norwalk virus)

and 36% (Mexico virus) in infants of 7 to 11 months of age. This reflects maternal

antibodies present in infants that rapidly decreased. The seroprevalence increased sharply

from 1 to 3 years of age in young children and approached 100% at 8 to 9 years of age.

These results indicated that NoV infections are very common in this population and

children are infected with NoVs at an early age. In Australian aborigines NV infection

occurs early in life and by the age of 6 years over 90% of children were seropositive to

Norwalk virus (191). Another seroprevalence study using Norwalk and Mexico VLPs in

Kuwait populations showed similar results (36).

These results suggest that NoVs were not an important cause of illness in infants

and young children in developed countries. Although NoV infections are universal in

children less than 5 years of age in developing countries, childhood exposure does not

protect adults from disease. The reasons probably are: (1) NoVs have so many genetic

and antigenic types with little cross-protection that infection with one type can not protect from infection of other types; (2) Infection with NoVs does not induce a long-term protective immunity.

Animal NoVs. Recent seroprevalence studies of bovine and porcine NoVs have also been reported (35, 48). In Thuringia, Germany, serum samples from 99.1% of the cattle tested (n=824) were seropositive to the VLPs of bovine NoV/GIII-1/Jena/80/DE

56 strain. In the US, 71% of the pigs (n=110) were seropositive to the VLPs of the Japanese porcine NoV/GII/Sw918 strain, 63% were seropositive to human NoV/GI/Norwalk VLPs and 52% were seropositive for human NoV/GII/Hawaii VLPs. In Japan, the pig seroprevalence to the Sw918 strain was 36% (95/266). These results indicate that NoV infections are common among domestic cattle and pigs.

Human SaVs. In Southeast Asia, 70-94% of adults were seropositive to SaVs

tested by a radioimmunoassay blocking test (169). In Sapporo, Japan, where the SaV

prototype Sapporo strain was discovered, the seroprevalence of SaV-antibodies increased

greatly in 2-5-year-old children (65%, 13/20), and peaked at school age (90%, 18/20) and remained high (90%, 9/10) in adults by IEM using SaV antigens from fecal extracts (212).

Another SaV seroprevalence study performed in the US showed a similar pattern of acquisition of antibodies to SaVs, which began in infancy and became 100% by the age of 4 years (172).

No seroprevalence study has been reported for porcine and mink SaVs.

1.6.3 Zoonotic potential

Noroviruses have been identified in humans, cattle, pigs and mice. Murine

NoVs belong to a distinct GV within the genus Norovirus and the pathogenesis of murine

NoVs in mice differs from that of NoVs in humans (117). At present, no evidence of transmission of murine NoVs to humans exists. Bovine NoVs (GIII-1, 2) and NB-like enteric caliciviruses are genetically or antigenically different from human NoVs and

SaVs (81, 82, 139, 186). However, a recent seroprevalence study of antibodies against

Bo/GIII-2/Newbury-2 strain in veterinarians in The Netherlands demonstrated that there

57 is a possibility of cross-species transmission of bovine NoVs to humans (252). The

seroprevalence of Newbury-2 IgG antibody in veterinarians (58/210, 28%) was

significantly higher than in control populations [127/630, 20% (P = 0.03)]. Although

weak cross-reactivity may exist between the capsid proteins of Newbury-2 and antibodies

to human NoVs, 26% of the Newbury-2 antibody-positive samples showed high IgG

antibody titers to the Newbury-2 antigens but low titers to Norwalk VLPs, suggesting a

specific response to Newbury-2 antigens. In addition, young veterinarians who spent time

on farms [Odds Ratio (OR) = 1.8] and the veterinarians who were members of the Bovine

Practitioners Society were significantly more likely to be IgG seropositive to Newbury-2

(OR = 2.7). These results suggest that bovine GIII-2 NoVs may infect humans or cross-

reactive GIII strains exist in cattle and humans.

Porcine NoVs are genetically classified into the same genogroup, GII (233) as

the predominant human NoVs causing nonbacterial gastroenteritis worldwide. Also,

porcine NoVs are antigenically related to human NoVs (48). These results raise public

health concerns of the potential for zoonotic transmission of NoVs to humans.

Interspecies transmission of vesiviruses. Vesiviruses belong to the genus

Vesivirus within the family Caliciviridae. A bovine vesivirus, Tillamook strain, was

isolated originally in 1981 from 3 dairy calves in a herd in Oregon with persistent calf respiratory disease (222). This virus caused only minimal lesions in 2 experimentally inoculated calves, but established a persistent infection with virus shedding for 45 days after which time the calves were euthanized. Pigs also developed vesicular lesions after inoculation with this virus. Later, the neutralizing antibodies against Tillamook strain were found in two widely distributed species of sea lions suggesting the possibility of a

58 marine origin for this agent (9). A vesicular exanthema of swine-like virus (VESV) was

detected from an aborted bovine fetus (223). These results suggest that cross-species

transmission may occur for vesiviruses.

The RNA virus populations consist of closely related viral mutants and

recombinants, which are known as quasispecies (40). They are subjected to continuous

genetic variation, competition and selection during virus replication. When the virus

population size increases, the expected number of viral genomes with more mutation sites

also increases. Because of the high prevalence of human and animal NoVs, the risk of

emergence of “new” variants with changes in host cell tropism or the ability to replicate

in a different host species increases too. Further studies of NoVs and SaVs in animals and in humans with occupational animal exposure (e.g. farm workers, veterinarians and abattoir workers), but especially of these viruses in developing countries where animals and humans are in close contact are needed.

1.7 IMMUNITY

Human NoVs. The importance of immunity to NoVs infection is still in question because adults demonstrate a high degree of susceptibility to NoV infections although the seroprevalence among them is as high as 50-100% (28, 36, 106, 191). Little is known about the gastrointestinal secretory or serum neutralizing antibodies because of the absence of a cell-culture system to assay neutralizing antibody responses. The knowledge of immunity to NoVs is mainly from volunteer studies.

59 A recent Norwalk virus challenge study of 77 volunteers (135) showed that a

portion of the Se+ individuals were also resistant to infection. However, these Se+

resistant individuals showed earlier (≤ 2 days post challenge) saliva IgA antibody

responses compared to those (> 5 days post challenge) of the infected Se+ individuals.

The authors suggested that a memory immune response was responsible for the resistance

of these Se+ individuals. However, it is unclear whether protective immunity to Norwalk

virus is short-term or long-term and what frequency of repeated exposure is necessary to

induce protective immunity. Previous human volunteer studies of Norwalk virus also

demonstrated that short-term protective immunity to Norwalk virus exists (37, 135, 192)

and protective immunity is serotype specific. Volunteers who became ill following the

first Norwalk virus challenge were protected from illness when they were re-challenged

with the same virus 6 to 14 weeks later, but they were not protected when they were re-

challenged with Hawaii virus (259). During a rotavirus vaccine study of Finnish children

(131), the authors demonstrated that children with high Norwalk virus-specific serum IgG antibody titers were significantly less likely to acquire a Norwalk virus infection than children who had lower Norwalk virus serum IgG antibody titers (P < 0.05).

No long-term immunity exists even to the homologous strain in the volunteer

studies (192). Six of twelve volunteers developed illness following the initial challenge,

and they also developed symptomatic infection again following re-challenge 27-42

months later. In those individuals who did not become ill, the level of serum or intestinal

antibody before and after exposure was low, whereas individuals who became ill

developed a serological response after each challenge. The level of serum or intestinal

antibody before exposure seemed to be inversely proportional to protection (63). The

60 studies using recombinant VLPs also indicated that there is an inverse correlation

between the titer of prechallenge antibodies (fecal IgA or serum IgG ) and the

development of clinical illness in adult volunteers (59, 185). However, conflicting evidence also exists: in Bangladesh or Panama, the presence of serum antibodies to

Norwalk virus was found to correlate with resistance to virus infection (14, 208); volunteers who were clinically ill did not seroconvert to Norwalk virus (91). Several explanations for this paradox have been suggested. For example, repeated exposure to

Norwalk virus, which may influence long-term immunity, does induce protective natural immunity (44, 158). Similarly, the antibody levels of volunteers who were repetitively exposed to NV were associated with protection (107). In addition, virus receptors in the gastrointestinal tract are genetically variable and may be responsible for resistance to

NoV infections, providing variable genetic susceptibility to NoVs in different populations or age groups.

A recent study of murine MNV-1 strain demonstrated that the virus was more virulent in IFNαβγ-/- mice than wild-type mice and the MNV-1 replicated better in

macrophage and dendritic cells from IFNαβγ-/- and STAT1-/- mice (117). These data indicate that innate immunity is important during murine NoV infection.

Only one study of cellular and humoral immune responses to NoV infection has

been reported (134). Fifteen volunteers were challenged with the GII-2/Snow Mountain

virus. The stool, serum, saliva, and peripheral blood mononuclear cell (PBMC) immune

responses were analyzed pre- and post-challenge. Sixty percent (6/15) of the volunteers

were infected and showed an increased (≥ 4-fold) in serum IgG antibody, mainly IgG1

antibody responses post-challenge. The serum IgG cross-reacted with another GII-1

61 Hawaii virus, but salivary IgA was less cross-reactive with Hawaii strain. Neither serum

IgG nor salivary IgA cross-reacted with GI Norwalk virus. Serum cytokine levels were

examined by cytometric bead array assay. Serum IFN-γ and IL-2, but not IL-6 or IL-10, increased significantly at 2 days post challenge. For 91% (11/15) of volunteers including infected and uninfected, prechallenge PBMCs stimulated with Snow Mountain virus

VLPs secreted IFN-γ and other Th1 cytokines (TNFα and IL-2) suggesting previous NoV exposure and immunological memory in most volunteers. Like the IgG antibodies, the virus activated T cells cross-reacted with Hawaii virus, but not with Norwalk virus. The

IFN-γ production was dependent upon CD4+ T cells, consistent with a predominant, but

not exclusive, Th1 response.

Human SaVs. The SaV specific antibodies may be correlated with the

resistance to illness in infants and young children. One study analyzed paired serum

samples (pre- and post-outbreak) from sick infants and healthy contacts involved in a

Sapporo virus outbreak at an infant home by RIA (171). Of 41 infant residents, 18 had pre-existing antibodies to SaVs and 3 (17%) of them became ill, whereas 23 did not have pre-existing antibodies and 18 (78%) of them were sick. The percentages of sickness between infants with and without pre-existing antibodies differed significantly (P<0.01,

χ2 analysis).

Animal NoVs. Short-term protective immunity exists for bovine GIII-2/CV186-

OH strain and fecal IgA antibody responses may contribute to the protection. All Gn

calves recovered from virus infection had high titers of fecal IgA antibody responses and

were completely protected against challenge with the same strain (80).

62 1.8 TREATMENT

1.8.1 Anti-gastroenteritis agents

In adults, human NoV and SaV infections induced symptoms that are generally mild and self-limited. Supportive care is the rule to treat NoV or SaV associated gastroenteritis (63). For example, the oral rehydration solution (ORS) including electrolytes is usually sufficient to compensate for fluid loss with diarrhea and vomiting.

Severe dehydration, although rare, occurred with both NoV and SaV induced diarrhea and i.v. administration of solutions was needed. For intractable vomiting, administration of ORS in small amounts by a spoon, syringe, cup, or feeding bottle is recommended.

Continuous, slow nasogastric infusion of ORS via a feeding tube can be helpful for the child who is vomiting (1). For infected young children under 5 years of age, that show 2 or more of the following signs: (1) restless, irritable; (2) sunken eyes; (3) drinking

eagerly and thirsty; (4) skin pinch goes back slowly, indicate dehydration and care can be provided at home. If a child has 2 or more of these signs: (1) lethargic or unconscious; (2) sunken eyes; (3) not able to drink or drink poorly; (4) skin pinch goes back very slowly, it is considered severe dehydration and hospital emergency care is needed. If the child still

can drink, sips of ORS should be given on the way to the hospital [World Health

Organization regional office for South-East Asia. Guidelines for management of common

diseases in young children in emergencies (working document),

http://w3.whosea.org/LinkFiles/List_of_Guidelines_for_Health_Emergency_draft__docu

ment.pdf].

63 1.8.2 Anti-virus replication

Interference of virus binding. Tamura et al. (238) found that non-cytotoxic histone H1, was an extremely strong NoV-binding protein and was a potential anti-virus reagent. It interacted with NoV particles and also with the cell surface, and prevented the attachment of NoV but not of other viruses to intestinal cells. In a recent study, the same research group found that GII NoVs efficiently bind to the heparan sulfate on the cell surface (237). Because various interactions between histones and glycosaminoglycans in the nuclei and on the cell surface have been reported, the authors suggested that histone

H1 may block the binding of NoVs to cell surface by direct binding to heparan sulfate.

The Norwalk virus VLPs have been found to attach to the HBGAs on the surface of the epithelial cells of the gastroduodenal junction (151). These attachments were abolished by α-1, 2-fucosidase and inhibited by the H types 1 and 3 trisaccharides or by anti-H type

1 and anti-H type 3/4 antibodies. Because virus binding to specific cells is the first step during virus infection, the histone H1, α-1, 2-fucosidase and polysaccharides are potential anti-NoV replication reagents.

RNA interference. RNA interference (RNAi), or RNA silencing, is a sequence-

specific RNA degradation process in the cytoplasm of eukaryotic cells that is induced by

double-stranded RNA (108). It controls gene expression at transcriptional and post-

transcriptional levels. It is a natural defense mechanism against incoming viruses and the

expression of transposable elements. Based on the mechanism of RNAi, antisense oligos

were investigated for viral therapy. Morpholino (PMO) is one method to synthesize

antisense oligos (234). It starts with the introduction of an amine via a relatively simple ribose-to-morpholine transformation to ribonucleosides. The resulting PMO subunits can

64 be assembled into antisense oligos via simple and efficient coupling to the morpholine

nitrogen.

A PMO was designed from FCV consensus sequences. The PMO blocked a novel FCV, which caused fulminant hepatitis in cats, infection in cell culture in a dose- dependent manner (221). When this PMO was administrated intraperitoneally (i.p.) as a treatment for two typical FCV outbreaks in cats, which differed from the novel FCV infections, it significantly decreased the mortality. In the two outbreaks, the infected cats developed typical clinical signs of FCV infection: respiratory infection with mucosal and skin ulcers. The FCV was detected from lungs (7/9), but not from liver (0/9) by necropsies and cell culture isolation. In outbreak 1, the PMO treatment was the predictor of survival for all ill cats (n=28) and the PMO-treated cats (n=22) by multiple regression analysis. For the 22 PMO-treated cats, earlier administration of PMO (on the first disease day) significantly increased survival rate compared with first doses on disease days 2-5

(P=0.008). In outbreak 2, 80% (16/20) of PMO-treated cats survived, which was

significant higher than the survival rate (15%, 4/25) of cats without PMO-treatment.

These data suggests that the antisense nucleotide designed based on the conserved region

of FCVs inhibited the replication of two FCV strains in vitro and in vivo, respectively.

The RNAi mechanism should be tested with cell cultured adapted NoVs and

SaVs, such as murine MNV-1 strain and SaV Cowden strain in the future to investigate

whether RNAi is a way to prevent NoV and SaV infections.

1.9 CONTROL AND PREVENTION

1.9.1 General approaches

65 There are several features of NoV and SaVs that contribute to their highly contagious nature. First, these viruses are highly infectious. A few viral particles (<100 particles) permit droplet-transmission, person-to-person spread, and frequently secondary spread to family members and other close contacts (57). Second, they are stable in the environment. Third, it has been estimated that about 50% of people exposed to NoV become ill and that asymptomatic individuals can shed viruses longer than 1 week; thus these people could be important infectious sources, especially food handlers (43). Forth, shellfish can accumulate viruses, and NoV-contaminated shellfish can meet bacteriologic standards and be distributed to the market. Fifth, these viruses demonstrate great antigenic and genetic diversity, and induce little cross-protection. So people can be

infected serially by different strains. Sixth, people can be infected by the same strain after a period of time because long-term immunity may not exist for NoVs.

Prevention strategies should focus on the control of the infectious sources, block

the transmission routes, or use immunization if safe and effective vaccines become

available in the future. Investigation of the infection of food handlers, contamination of foods, oyster beds, drinking water and swimming pools and subsequent effective disposal or disinfection of contaminated material should decrease the frequency of outbreaks (63).

Personal hygiene including effective hand washing is the best way for individuals to prevent infection. The consumption of raw shellfish is also not recommended.

1.9.2 Vaccines

Virus vaccines. Natural infection and volunteer studies showed that repeated exposures to NoVs does induce protective immunity. Although some people still question

66 whether a human NoV vaccine is necessary because long-term immunity may not exist

and short-term immunity is not broadly cross-reactive for the genetically and

antigenically diverse NoVs, vaccines are required for those people at high risk. For

example, military soldiers, the elderly in nursing homes, travelers and younger children

in day-care centers may benefit from vaccines because short-term immunity does exist

(163, 170).

To develop a NoV vaccine, several challenges need to be overcome. The biggest difficulty is to overcome the genetic and antigenic diversity of these viruses and to produce a vaccine capable of preventing infection by most, if not all, of the important circulating strains.

The tissue cultured porcine SaV Cowden strain has been proven to be an

attenuated strain compared to the wild-type-Cowden in a pathogenesis study (178).

Gnotobiotic pigs inoculated with the tissue cultured Cowden strain did not developed diarrhea. Thus tissue cultured Cowden strain may be a potential candidate vaccine for swine against SaV infection.

VLP vaccines. Recombinant NoV VLPs produced in the baculovirus- expression system are similar to native virus particles in morphology and antigenicity (64,

103). The VLPs also are highly immunogenic when inoculated parenterally with adjuvant into experimental animals (103).

The immunogenicity of Norwalk virus VLPs were evaluated in mice following oral administration without or with the adjuvant cholera toxin (CT) (8). When the groups with CT and without CT were compared, Norwalk virus-specific serum IgG and intestinal IgA antibodies were detected in both groups, the number of responders was not

67 significantly different, but CT induced higher levels of serum IgG antibodies. In most

mice, serum IgG and intestinal IgA antibody responses were detected by 9 and 24 days

post immunization, respectively. In the absence of CT, IgG2b was the dominant IgG

subclass response reflecting a predominant Th1 response, which is often induced by virus

infections and viral antigens. These results show that non-replicating VLPs are

immunogenic when administered orally in the absence of a delivery vehicle or mucosal

adjuvant.

The Norwalk virus VLPs induced immune responses in mice when given

intranasally in the presence or absence of the mucosal adjuvant, mutant Escherichia coli

heat-labile toxin (mLT-R192G). Interestingly, the intranasal inoculation of low doses of

VLPs was more effective to induce an immune response than oral administration (68).

These preclinical studies in naïve mice demonstrated that VLPs are an excellent

immunogen when administered via both oral and nasal routes. Several unique properties

could contribute to the efficacious mucosal immunogenicity of VLPs: these VLPs are

stable at low pH, they can be lyophilized and reconstituted in water or buffer, and they

are particulate and therefore might be targeted to Peyer’s patches in the gastrointestinal

tract. The VLPs are a potential candidate vaccine for NV infection (44).

In a phase I trial (7), the safety and immunogenicity of recombinant Norwalk

virus VLPs as an oral immunogen have been evaluated in healthy adult volunteers who

had pre-existing antibodies. The VLPs boosted antibody responses. Oral administrations

of 100 µg and 250 µg doses of VLPs were safe. Serum total and subclass (IgG and IgA)

Norwalk virus-specific antibodies were monitored by antibody-ELISA. Upon administration of the non-replicating VLPs, the increases of antibody titers were

68 moderate, the frequency of antibody responses was similar to that induced by natural

infection, but the magnitude of the antibody responses was lower than that induced by

wild-type Norwalk virus. However, whether the VLPs can induce antibody responses in

immunologically naive subjects is unknown. Both Norwalk virus and its VLPs stimulated

a predominantly IgG1 subclass response (Th2 response) when administrated orally (7).

This predominant Th2 response differs from the predominant Th1 response of

Hu/NoV/GII-2/Snow Mountain virus infection in human volunteers (134). The

predominant Th2 response has also been seen previously in naïve BALB/c mice

immunized with Norwalk virus VLPs and adjuvant CT (8). Serum IgA antibodies were

detected but direct detection of fecal IgA antibodies was unsuccessful in most volunteers.

Norwalk virus capsids expressed in tobacco leaves and potatoes stimulated a

serum and mucosal immune response in most mice in the absence or presence of the

adjuvant CT. Potato tubers containing Norwalk virus VLPs were also immunogenic when

fed to mice and when given to volunteers (155, 236).

The VLP vaccine of Bo/NoV/GIII-2/CV186-OH strain was evaluated in Gn

calves (80). Calves were vaccinated with two or three doses (250 µg/dose) of the VLPs

with different adjuvants: oil, mLT-R192G or immunostimulating complexes (ISCOM).

Only calves vaccinated intranasally with VLP+mLT were fecal IgA antibody positive and partial protected after challenge, with delayed and shortened diarrhea (1-2 days).

Diarrhea was present for 2-6 days in calves vaccinated with VLP with oil or ISCOM and for 8-9 days in controls. Virus shedding was detected post-challenge by ELISA in all vaccinated calves. All calves recovered from bovine NoV infection had high titers of fecal IgA antibodies and were completely protected against re-challenge. The authors

69 concluded that only bovine NoV VLP+mLT given intranasally stimulated both serum and fecal IgA antibodies and partial protection.

Neutralizing antibodies are often critical for protective immunity and cannot be monitored because a cell culture system is not available. The main question remains is whether the immune responses induced by the VLPs are protective immune responses.

This can first be best evaluated by studies in animal models (calves and pigs) susceptible to enteric NoV infection.

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94 Genogroup- Genbank Genogroup- Genbank Genus Strain genotype accession # Genus Strain genotype accession # Norovirus Hu/Norwalk/68/US GI-1 M87661 Norovirus Po/Sw918/97/JP GII-11 AB074893 Hu/Southampton/91/UK GI-1 L07418 Hu/Gifu/96/JP GII-12 AB045603 Hu/Desert Shield/90/US GI-2 U04469 Hu/SaitamaU1/99/JP GII-12 AB039775 Hu/Chiba 407/87/JP GI-3 D38547 & AB022679 Hu/Fayetteville/98/US GII-13 AY113106 Hu/Musgrove/89/UK GI-4 AJ277614 Hu/M7/99/US GII-14 AY130761 Hu/SzUG1/99/JP GI-5 AB039774 Hu/J23/99/US GII-15 AY130762 Hu/BS5/DE GI-6 AF093797 Hu/Tiffin/99/US GII-16 AY502010 Hu/WUG1/00/JP GI-6 AB081723 Hu/Neustrelitz260/00/DE GII-16 AY772730 Hu/Boxer/01/US GI-7 AF538679 Hu/CS-E1/02/US GII-17 AY502009 Hu/Hawaii/71/US GII-1 U07611 Bo/Jena/80/DE GIII-1 AJ011099 Hu/Snow Mountain/76/US GII-2 AY134748 Bo/Newbury-2/76/UK GIII-2 AF097917 Hu/Mexico/89/MX GII-3 U22498 Bo/CV186-OH/00/US GIII-2 AF542084 Hu/Toronto/91/CA GII-3 U02030 Hu/Alphatron/98-2/98/NET GIV AF195847 Hu/SaitamaU18/97-99/JP GII-3 AB039781 Mu/MNV-1/03/US GV AY228235 Hu/SaitamaU201/98/JP GII-3 AB039782

95 Hu/Camberwell/94/AUS GII-4 AF145896 Sapovirus Hu/Sapporo/82/JP GI-1 U65427 Hu/Lordsdale/93/UK GII-4 X86557 Hu/Manchester/93/UK GI-1 X86560 Hu/Bristol/93/UK GII-4 X76716 Hu/Plymouth/92/UK GI-1 X86559 Hu/MD145-12/87/US GII-4 AY032605 Hu/Lyon30388/98/FR GI-1 AJ251991 Hu/Farmington Hills/02/US GII-4 AY502023 Hu/Houston/86/US GI-1 U95643 Hu/Langen1061/02/DE GII-4 AY485642 Hu/Parkville/94/US GI-2 U73124 Hu/Hillingdon/93/UK GII-5 AJ277607 Hu/Houston/90/US GI-2 U95644 Hu/New Orleans 306/94/US GII-5 AF414422 Hu/Stockholm/97/SE GI-3 AF194182 Hu/Baltimore/274/1993/US GII-6 AF414408 Hu/Mex14917/00/MX GI-3 AF435813 & AF435810 Hu/SaitamaU3/97/JP GII-6 AB039776 Hu/London/92/UK GII-1 U95645 Hu/SaitamaU4/97/JP GII-6 AB039777 Hu/Lyon598/97/FR GII-1 AJ271056 Hu/SaitamaU16/97/JP GII-6 AB039778 Hu/Bristol/98/UK GII-1 AJ249939 Hu/SaitamaU17/97/JP GII-6 AB039779 Hu/Mex340/90/MX GII-2 AF435812 & AF435809 Hu/Leeds/90/UK GII-7 AJ277608 Hu/Cruise ship/00/US GII-3 AY289804 & AY157863 Hu/Gwynedd/273/94/US GII-7 AF414409 Hu/Mc10/00/TH GII-4 AY237420 Hu/Amsterdam/98-18/98/NET GII-8 AF195848 Hu/C12/00/JP GII-5 AY603425 Hu/SaitamaU25/97-99/JP GII-8 AB039780 Po/Cowden/80/US GIII AF182760 Hu/VA97207/97/US GII-9 AY038599 Po/LL14/02/US GIII AY425671 Hu/NLV/Erfurt/546/00/DE GII-10 AF427118 Hu/Hou7-1181/90/US GIV AF435814 Hu/Mc37/00-01/THA GII-10 AY237415 Hu/Argentina39/Arg GV AY289803 & AF405715 Po/Sw43/97/JP GII-11 AB074892 Mink/MEC/1/1999/US G? AF338404 Table 1.1 Summary of norovirus and sapovirus representative strains. Strains that have a complete genome sequence are bolded. Norovirus ORF1 ORF2 ORF3 5 5374 6950 7588 5’ 1789 aa 212 aa 7654 (A)n Capsid 530 aa

5358 6950 Subgenomic RNA ~2 kb (A)n

Sapovirus 96 ORF1 ORF2 13 6855

5’ 1719 aa Capsid 561 aa 165 aa 7431 (A)n 161 aa 6852 7349 ORF3 Subgenomic RNA ~2 kb (A)n

Figure 1.1. The genome organization, open reading frames (ORF) and subgenomic RNA are shown for representative strains in the genus Norovirus (Hu/NoV/Norwalk/1968/US, GenBank accession number M87661) and genus Sapovirus (Hu/SaV/Manchester/1993/UK, GenBank accesion number X86560). The size of each ORF is indicated as total amino acids (aa). The numbers alone indicate nucleotide position. A poly(A) tail is shown as (A)n. A. Proteolytic cleavage map of polyprotein

399 762 961 1099 1280 Q/G Q/G E/G E/A E/G

p48 P41, NTPase p22 p16, VPg p19, Pro p57, Pol

B. Domains and subdomains of capsid protein 97

NS P1AP2P1B

10-49 50-225 226-278 279-405 406-520

Shell Protruding

Figure 1.2. A. The proteolytic cleavage map of Hu/NoV/GI/Southampton/91/UK (GenBank accession number L07418). Pro and Pol are abbreviations of proteinase and polymerase, respectively. The arrows indicate each cleavage site between amino acids Q/G, E/G or E/A. The numbers indicate the P1 positions of the cleavage sites of the polyprotein. B. Domain and subdomains of the capsid protein of Hu/NoV/GI/Norwalk/68/US (GenBank accession number M87661). The numbers indicate amino acid position of the capsid protein.

CHAPTER 2

PORCINE NOROVIRUSES: GENETIC AND ANTIGENIC RELATIONSHIPS TO

HUMAN NOROVIRUSES

2.1 SUMMARY

Detection of genogroup II (GII) norovirus RNA from adult pigs in Japan and

Europe and GII norovirus antibodies in US swine raises public health concerns about zoonotic transmission of porcine noroviruses to humans. No noroviruses have been detected in US swine. To detect porcine noroviruses and to investigate their genetic diversity and relatedness to human noroviruses, 275 fecal samples from normal US adult pigs were screened by reverse transcription-PCR using calicivirus universal primers. Six samples were confirmed norovirus positive by sequencing. Based on sequence analysis of the 3’-end 3 kb of 5 porcine noroviruses, 3 genotypes within GII and two potential recombinant strains were identified. One genotype of porcine noroviruses was genetically

98 and antigenically related to human noroviruses and replicated in gnotobiotic pigs. These

results raise concerns of whether subclinically infected adult swine may be reservoirs for

emergence of new human noroviruses or if porcine/human GII recombinants could

emerge.

2.2 INTRODUCTION

Noroviruses (NoV) are emerging enteric pathogens that cause diarrhea in humans

and animals (4, 19, 21). They belong to family Caliciviridae, genus Norovirus. (4). The

NoV genome is 7.3-7.7 kb long and includes 3 open reading frames (ORFs) encoding a

non-structural polyprotein, a major capsid protein (VP1, capsid) and a minor capsid protein (VP2, small basic protein). The polyprotein undergoes protease processing to produce several non-structural proteins and an RNA-dependent RNA polymerase (RdRp)

(1, 4, 27). The capsid protein is composed of a conserved shell (S) domain and hypervariable protruding (P) domain based on x-ray crystallography of virus-like particles (VLPs) of human Norwalk/68/US strain (28). Noroviruses are genetically diverse and comprise 27 genotypes within 5 genogroups: GI/1-8, GII/1-17, GIII/1-2, GIV and GV based on the capsid genes of 164 strains (39). Human NoVs are estimated to cause 23 million cases of illness annually in the US (24) and more than 90% of nonbacterial epidemic gastroenteritis worldwide (4). The low infectious dose, environmental resistance, strain diversity, shedding from asymptomatic individuals and

99 the varied transmission vehicles make human NoVs highly contagious.

Norovirus RNA was detected by reverse transcription-PCR (RT-PCR) in cecal

contents of 4 of 1017 normal slaughtered pigs in Japan (30) and in 2 of 100 pooled fecal

samples from 3-9-month old pigs in The Netherlands (32). These porcine NoV strains

(Sw43/97/JP, Sw918/97/JP and 34/98/NET) are genetically similar to each other and are

classified as NoV GII (30, 32) like most epidemic human NoVs (3, 22, 36). Also, VLPs of the Sw918 strain cross-react with antibodies against human GII NoVs (Hawaii/71/US and Mexico/89/MX), but not with antibodies against human GI NoVs (Norwalk and

VA115) (2). The close genetic and antigenic relationships between human and porcine

NoVs raise public health concerns of their potential for zoonotic transmission and reservoirs for emergence of new epidemic human strains.

Farkas et al. (2) reported that US swine sera react with the Po/NoV/GII/Sw918 strain, but no direct detection of NoVs from US swine has been reported. To detect porcine NoVs and assess their genetic diversity and relatedness to human NoVs, we screened 275 pig fecal samples from US swine by RT-PCR with a calicivirus universal primer pair p290/110 (14, 17) followed by sequencing of the RT-PCR products. To classify these porcine NoVs, we sequenced the 3’-end 3 kb of the genome of 5 strains.

Gnotobiotic (Gn) pigs were inoculated with porcine NoVs to examine their infectivity and to produce convalescent antiserum for antigenic analysis.

100 2.3 MATERIALS AND METHODS

Sampling. Fecal samples (n=275) were collected from December, 2002 to June,

2003 from finisher (10-24 weeks of age) pigs and gestating sows (≥1 years of age) from 3

Ohio swine farms (10, 60 and 32 samples from OH farms A, B and C, respectively), 1

Ohio slaughterhouse (83 samples) receiving pigs from multiple farms, 1 Michigan swine farm (61 samples), and 2 North Carolina swine farms (8 and 21 samples from NC farm A and B, respectively). OH farm B and MI farm A were typical US commercial swine farms housing about 50,000 pigs and the other farms housed about 1,000 pigs. Fresh fecal samples were collected from individual pigs, placed into sterile specimen containers and stored frozen.

RNA extraction. Sample RNA was extracted from 200 µl of centrifuged (1,200 × g, 30 min) 10% fecal suspensions in sterile Eagle’s minimal essential medium (EMEM,

Invitrogen, Carlsbad, CA) by using the Trizol LS (Invitrogen) procedure. The RNA pellet was resuspended in 40 µl RNase-free water and stored at -20 or -70ºC. For amplification of the 3’-end 3 kb fragment of porcine NoVs, the 10% fecal suspensions were filtered

(0.45 µm) and semi-purified through a 40% (w/v) sucrose-cushion by ultracentrifugation

(112,700 × g, 2 h). The pellets were resuspended in EMEM for RNA extraction. The

RNA was concentrated and purified by using QIAamp Viral RNA Mini kit (Qiagen,

Valencia, CA).

RT-PCR. (i) RT and PCR were performed separately using primer pair p290 (5’-

101 GAT TAC TCC AAG TGG GAC TCC AC -3’) (14) and p110 (5’-ACD ATY TCA TCA

TCA CCA TA-3’) (17) as previously reported (14) with minor modifications (48ºC for

annealing). The products were 317 bp for NoVs or 329 bp for sapovirus.

(ii) To amplify the 3’-end 3 kb fragment of the porcine NoVs, the cDNA was

synthesized by SuperScript III First-Strand cDNA synthesis kit (Invitrogen) according to

the manufacturer’s instructions with primer VN3T20 (5’-GAG TGA CCG CGG CCG

TM CT20-3’). Then PCR was performed using TaKaRa Ex Taq polymerase (TaKaRa Mirus

Bio, Madison, WI) with primers p290 and VN3T20. After initial denaturing at 94ºC for 3 min, the first 5 cycles were performed at 94ºC for 30 sec, 50ºC for 30 sec and 72ºC for 5 min, then another 30 cycles with a shorter elongation time of 3 min, and a final extension for 10 min at 72ºC. All RT-PCR products were analyzed by agarose gel electrophoresis stained with ethidium bromide and visualized using ultraviolet light.

Cloning and sequencing. The RT-PCR products were purified by QIAquick Gel

Extraction kit (Qiagen). Products of primer pair p290/110 were sequenced directly using primer p290 and p110 or after cloning into pCR2.1-TOPO (T/A) vector (Invitrogen). The

~3 kb RT-PCR products were cloned by using PCR XL cloning kit (Invitrogen). The insertion was confirmed by restriction enzyme digestion. Five clones of each sample were sequenced by a primer-walking strategy. DNA sequencing was done using BigDye

Terminator Cycle chemistry and 3700 or 3730 DNA Analyzer (Applied Biosystems,

Foster, CA).

102 Sequence analysis. Sequence editing was performed by Lasergene software package (version 5) (DNASTAR Inc., Madison, WI). Basic Local Alignment Search Tool

(BLAST, http://www.ncbi.nlm.nih.gov) was used to find homologous hits. Multiple

sequence alignment was done by ClustalW (v 1.83) at DDBJ (http://www.ddbj.nig.ac.jp).

Phylogenetic analysis (Neighbor-Joining) with bootstrap (1,000 replicates) and amino acid pairwise distance analysis (p-distance) were conducted using MEGA v 2.1 (16).

Recombination identification. Identification of recombinants was performed by comparing a query sequence to a set of background sequences by using Recombinant

Identification Program (29) (http://hivweb.lanl.gov/RIP/RIPsubmit.html) and creating separate phylogenetic trees for the segments resembling different genetic clusters of the query sequence.

Gnotobiotic (Gn) pigs. Two Gn pigs were inoculated with the new genotype

(GII-18) of porcine NoV (QW101-like). Because the QW101 and QW125 fecal samples

contained both porcine NoVs and sapoviruses (the latter detected by RT-PCR with primer

pair PEC66/65 (5) and confirmed by sequencing), we inoculated Gn pigs with two other

QW101-like porcine NoV strains, QW126 and QW144. The QW144 strain was obtained

by screening of other sapovirus-negative fecal samples collected from OH farm B by

RT-PCR with a newly designed porcine NoV primer pair PNV7 (5’- AGG TGG TGG

CCG AGG AYC TCC T-3’) and PNV8 (5’- TCA CCA TAG AAG GAR AAG CA-3’) targeting the RdRp of QW101 strain. These two strains were further confirmed as

103 QW101-like porcine NoVs in both the RdRp and capsid regions by sequence analysis of the RT-PCR products of the PNV7/8 and a primer pair PNV17 (5’- GGT CTC GTM CCA

GAG GTC A-3’) and PNV18A (5’- TTG TCT CAC ATC MAC TAT AA-3’) targeting the

S domain of the capsid of QW101 strain (211 bp for PNV7/8 and 402 bp for

PNV17/18A). They shared 99% and 100% amino acid identities to the QW101 strain for the PNV7/8 and PNV17/18A products, respectively.

Four Gn pigs were derived, maintained and euthanized as previously described (5,

25). The inoculum was composed of 5 ml of 0.2 µm-filtered 20% fecal suspensions in

EMEM of the QW126 or QW144 strains, or EMEM only (negative control). One Gn pig was inoculated with QW126 orally and intranasally at 9 days of age and maintained until post-inoculation day (PID) 26 to obtain convalescent antiserum, LL616. A second Gn pig was inoculated with QW144 orally at 35 days of age and euthanized at PID 5 when NoV shedding detected by RT-PCR with primer pair PNV7/8 was strongly positive. The other

2 Gn pigs were inoculated with EMEM orally as controls. Intestinal contents and acute serum were collected from the QW144-exposed and control pigs euthanized at PID 5.

Immune electron microscopy (IEM). The IEM was performed as described previously (12).

Enzyme linked immunoassay (ELISA). The recombinant baculovirus-expressed human NoV VLPs and as a negative control, a recombinant baculovirus-expressed rotavirus VP2 (bovine RF strain) and VP6 (human Wa strain) (2/6)-VLPs (38) were

104 purified through cesium chloride (CsCl)-gradients. Except for the VLPs of

GII-4/HS66/01/US (HS66, S. Cheetham and L. Saif, unpublished) produced in our lab,

the other human NoV VLPs were provided by Dr. Kim Green (NIAID, National Institutes

of Health). Briefly, 96-well microplates were coated with the human NoV VLPs (100-200

ng/well) (positive coating) or the rotavirus 2/6-VLPs (negative coating) in carbonate

buffer (pH 9.6) and incubated at 4ºC overnight. After washing and blocking with 5%

non-fat-dry milk in PBS-Tween 20 (0.05%) at 4ºC overnight, serially diluted Gn pig

convalescent antiserum LL616 was added to duplicate positive and negative coating

wells and the plates were incubated at room temperature for 2 h. Two pre-inoculation sera,

LL368 and MM982 from the 2 previously described Gn pigs exposed to porcine NoVs

and a human convalescent serum HS66CS to the HS66 strain served as negative and positive controls, respectively. After washing, horseradish peroxidase (HRP)-labeled goat anti-pig IgG (H+L) (KPL, Gaithersburg, MD) for pig sera or goat anti-human

IgG+IgA+IgM (H+L) (KPL) for the human serum were added. The plates were incubated at 37ºC for 1 h. After washing, the substrate 3,3’,5,5’-tetramethylbenzidine was added, the plates were incubated at room temperature for 20 min and the absorbance was measured at 650 nm. The cut-off value was determined as the mean absorbance of the negative coatings times 2.

Classification and GenBank accession numbers. The classification and

GenBank accession numbers of porcine NoVs identified in this study and other

105 representative strains for phylogenetic analysis are listed in Table 2.1.

2.4 RESULTS

The porcine NoVs were classified into 3 genotypes within GII including the previously recognized genotype containing the prototype Japanese strains Sw43 and

Sw918 and 2 new genotypes based on the complete capsid sequences. A total of 19 of

275 samples (7%) showed a potential positive band after agarose gel electrophoresis of the RT-PCR products of primer pair p290/110. Fourteen samples representative for each potential positive farm or the slaughterhouse were sequenced. After performing BLAST search, we identified 6 NoVs, 3 sapoviruses and 5 sequences that had no significant hit in the database. Porcine NoVs were from MI farm A (QW48), OH farm B (QW101, QW125 and QW126), and the OH slaughterhouse (QW170 and QW218). Because the QW126 strain shared 99% nucleotide identity with the QW101 and QW125 strains in the 274 nt

RdRp region (excluding the primer sequences), we choose QW48, QW101, QW125,

QW170 and QW218 strains for further sequence analysis.

We sequenced the 3’-end 3 kb of the genome containing the partial RdRp, VP1 and VP2 genes and the 3’-UTR of the 5 strains. The porcine NoVs separated into 3 distinct clusters: 1) Sw43, Sw918 and QW48; 2) QW101 and QW125; and 3) QW170 and QW218, based on the size of each gene and the ORF1-ORF2 overlap region (Table

2.2). Over the 3 kb, the QW101 and QW125 strains, and the QW170 and QW218 strains

106 shared 99% nucleotide identity, respectively.

In Table 2.3, the amino acid identity of the predicted complete and S and P

domains of the capsid protein of the 5 porcine NoVs, the previously reported porcine

NoVs (Sw43 and Sw918) and representative human, bovine and murine NoV strains are

summarized. In the complete capsid region, the QW48 strain was most closely related to

the porcine NoV prototype Sw43 strain with an amino acid identity of 98%; the QW170 and QW218 strains shared the highest amino acid identities (81%) to porcine Sw43 and

Sw918 strains; the QW101 and QW125 strains showed the highest amino acid identity to human GII-3/Mexico (71.4%), then to human GII-6/Baltimore (71.0%), porcine QW218

(71.0%) and porcine Sw43 (70.6%) strains. Analysis of the capsid S and P domains of these NoVs showed similar relationships. A Neighbor-Joining phylogenetic tree based on

the amino acid sequences of the complete capsids (Fig. 2.1A) revealed that QW48

grouped with the Japanese strains (Sw43 and Sw918) into GII-11 and QW170 and 218

formed a new genotype (GII-19) that was closer to the porcine than to the human strains.

However, QW101 and 125 formed a new genotype (GII-18) between human and porcine

GII NoVs.

Further analysis of the predicted C-terminal ~260 aa of the RdRp region (Fig.2.1B)

showed similar grouping results for QW48, QW101 and QW125 strains, but different for

QW170 and QW218 strains which were in the same cluster (GII-11) as Sw43, Sw918 and

QW48 in the RdRp region. This suggested that the QW170 and QW218 could be

107 potential recombinants between Sw43-like NoV (RdRp segment) and an unknown strain

with a capsid like QW170 and QW218 strains. The complete VP2 sequences of representative strains were also analyzed (data not shown). Results were similar to the

classification based on capsid sequences.

QW170 and QW218 represent recombinants between a Sw43-like strain

(RdRp) and a QW170/218-like strain (capsid). To determine where the recombination occurred for QW170 and QW218, we performed the Recombinant Identification Program by placing the 3’-end RdRp and the capsid sequence of QW170 or QW218 as a query sequence and the corresponding sequences of Sw43 and QW101 as background sequences. The VP2 gene was omitted because no sequence data for Sw43 or Sw918 is available. The resulting diagram (Fig. 2.2A) showed that QW170 had high similarity to

Sw43 strain in the RdRp region but not in the capsid region. This abrupt change

happened in the RdRp-capsid junction region. Therefore, we performed sequence

alignments of the RdRp-capsid junction of NoVs including the calicivirus

genomic-subgenomic conserved 18 nt motif (15) (Fig. 2.2B). Between Sw43, QW170

and QW218, all 18 nt were identical and identities decreased downstream of this motif.

The porcine NoVs replicated in Gn pigs. Two Gn pigs were inoculated with

GII-18 (QW101-like) porcine NoVs to verify if the NoVs replicate in pigs as confirmed

by IEM and to produce convalescent serum to examine the antigenic reactivity with

human NoVs. Porcine NoV shedding, assessed by RT-PCR, was detected at PID 2-5

108 (euthanized) post QW144 exposure coincident with mild diarrhea. Porcine NoV shedding

was detected only at PID 5 without diarrhea post QW126 exposure. Examination of the

intestinal contents of the pig inoculated with QW144 by IEM with pig convalescent

antiserum LL616 to QW126 revealed small clumps of typical ~32 nm diameter NoV

particles (Fig. 2.3). The 2 control Gn pigs had no virus shedding or diarrhea. These

results indicate that porcine NoVs replicated in the pigs. Detailed studies of the

pathogenesis of porcine NoVs in additional Gn pigs are in progress and will be described

in a separate report.

Antisera to QW101-like (QW126) porcine NoVs cross-reacts with VLPs of human GII NoVs in ELISA. An ELISA was performed to examine the one-way antigenic cross-reactivity between antiserum to porcine NoVs (QW101-like, QW126 strain) and human NoV VLPs of GI-3/Desert Shield, GII-1/Hawaii, GII-3/Toronto,

GII-4/MD145, GII-4/HS66 and GII-6/Florida strains (Table 2.4). The rotavirus 2/6 VLPs were used for negative coatings. The pig convalescent antiserum (LL616) to QW126, showed higher titers (400-800) to the GII strains: Toronto, MD145, HS66 and Florida; a lower titer (100) to the GII Hawaii strain; and the lowest titer (10) to the GI Desert Shield strain. The 2 pre-inoculation pig serum samples were negative for antibodies to each of these strains. This indicates that one-way antigenic cross-reactivity exists between the human NoV antigens and the porcine NoV (GII-18) antiserum, with moderate cross-reactivity to human GII-3, GII-4 and GII-6 NoVs, low to GII-1 and very low to

109 human GI-3 NoVs.

The NoV capsid conserved amino acid motifs in the QW101 strain were identified based on sequence alignments. Sequence alignments of the QW101 and 44

NoV strains including GII genotypes 1-19, GI/Norwalk, GIII/Newbury-2, GIV/Alphatron and GV/MNV-1 showed that the QW101 strain contains all the previously reported conserved motifs of NoVs in the capsid region including the IDPWI (amino acids 52-56,

IXPWI for MNV-1), LAGNA (amino acids 113-117) and LYTPLR (amino acids 187-192) motifs in the S domain, the QNGR (amino acids 267-270) motif in the P1A domain and the RGD(/K) (RGD-like, amino acids 292-294) motif in the P2 domain (6, 26, 31, 37).

The partial sequence alignments of 7 NoV strains are shown in Fig. 2.4. The QW101 strain also shared several newly identified motifs with other GII NoVs including the

NFVQAP (amino acids 59-64), GEFTVSPRNA(/S)PGE (amino acids 66-78) and

DVFTVSCRVLTRP (amino acids 199-211) motifs in the S domain, the NSRFP (amino acids 244-248, P1A domain) and LFFRS (amino acids 468-472, P1B domain) motifs.

2.5 DISCUSSION

In this study we choose the calicivirus universal primer pair p290/110 (14, 17) to screen for porcine NoVs because no sequence data for US porcine NoVs is available and

NoVs are highly variable genetically. From 275 pig fecal samples, we identified 6 genetically diverse NoVs. Although the primer pair p290/110 can detect human GI NoVs,

110 we did not find any GI NoV strains in pigs. This suggests that GII strains may be

predominant in pig NoV infections of the normal adult swine tested, which is very similar

to the current situation in humans (3, 21, 33, 35, 36). In addition, the calicivirus universal

primer pair p290/110 did not match well to every NoV strain probably resulting in low

sensitivity for certain strains. In this study, we found that at least 3 genotypes of porcine

NoVs, based on the complete capsid sequence, were circulating in US swine. The QW48

strain shared the highest amino acid identity (98%) in the capsid region with the porcine

NoV prototype Japanese Sw43 strain (GII-11) despite the geographic distance between

Japan and US. The QW170 and QW218 strains were potential recombinants containing

the RdRp from a Sw43-like strain and the capsid from another unknown but GII-19 strain.

More interestingly, the GII-18 porcine NoVs (QW101 and QW125) shared the highest

amino acid identity (71.4%) with the Mexico human strain in the capsid region.

Until now, only adult pigs were screened for NoVs and all porcine NoVs were

detected from pigs without clinical signs (30, 32). Subclinically infected pigs may be

natural reservoirs for NoVs and because porcine GII NoVs are genetically and

antigenically related to human NoVs, concerns exist about their zoonotic potential.

Whether human NoV strains similar to the QW101-like porcine NoVs circulate among

swine farm workers is unknown, but such studies could provide information on the zoonotic potential of these porcine NoVs.

The RdRp-capsid junction region contains a highly conserved 18 nt motif in

111 genomic and subgenomic RNA that is suggested to be a transcription start signal (4, 15).

All 18 nt were identical within each genogroup except for the Hu/GII/J23,

Po/GII/QW101 and Po/GII/QW125 strains (Fig. 2.2B, sequence alignments on other GI

and GIII strains are not shown). Four GII human strains (Fayetteville, Neustrelitz, J23 and M7) lack 4-5 nt information at the beginning. This information suggests that homologous recombination may occur within this motif between NoVs of different genotypes within the same genogroup. Recombinant human GII NoVs have been reported previously (7, 13, 15, 20, 34). To our knowledge, this study is the first identification of two potential GII NoV recombinants of pig origin, the QW170 and

QW218 strains. They are potential recombinants between the Sw43 (RdRp) strain and an unknown strain, which is genetically closer to the porcine than to the human GII NoVs in the capsid region. At present, NoV recombinants have been detected exclusively between viruses within the same genogroup and within the same host species origin, but few animal NoVs have been sequenced (RdRp and capsid) for comparative analysis, especially ones from animals in developing countries where humans and animals closely contact.

There are no previous reports of NoV replication in pigs or NoV particles detected from pig fecal samples. The QW101-like porcine NoVs replicated in Gn pigs with fecal shedding documented by RT-PCR and IEM. No cell culture system or animal disease models are available for human NoVs which greatly impedes the study of their

112 pathogenesis, replication strategies, host immune responses and preventive approaches.

The infection of pigs with porcine NoVs may provide a new infection or disease model for the study of NoV infections.

The NoV serotypes remain undefined due to the lack of cell culture system.

Evaluation of NoVs by cross-challenge studies in human volunteers, IEM or a solid phase

IEM, resulted in identification of 4 NoV antigenic types (Norwalk, Taunton, Hawaii, and

Snow Mountain) (4). By using ELISA based on hyperimmune antisera to VLPs,

GI-1/Norwalk, GI-3/Desert Shield, GII-1/Hawaii, GII-3/Mexico and GII-4/Grimsby strains were determined to represent distinct antigenic types. In this study one-way antigenic cross-reactivity was found between antiserum to QW101-like porcine NoVs and the VLPs of human NoVs, with cross-reactivity moderate to GII-3, GII-4 and GII-6, lower to GII-1 and the lowest to GI-3 NoVs. This coincides with the finding that the

QW101 strain shares high amino acid identity with GII-3 (71.4%), GII-6 (71.0%) and

GII-4 (63%) NoVs and more conserved amino acid motifs with GII NoVs than with any other genogroup of NoVs in the capsid region. The expression of VLPs of QW101 strain is in progress in our lab and will be used to examine the cross-reactivity between antiserum to human NoVs and the VLPs of QW101 strain. Among those conserved motifs, the IDPWI motif is recognized by a NoV-specific broadly reactive monoclonal antibodies 1B4, which reacts with 18 of 23 genotypes of NoVs (37). The NoVs bind to human histo-blood group antigens (HBGAs) which were found to influence the genetic

113 susceptibility of individuals to NoV infections (8-11, 18, 23). The RGD-like motif forms the bottom of a binding pocket by computational structure analysis and is critical for virus binding to HBGAs as determined by site-directed-mutagenesis of 2 strains having different HBGA-binding patterns (31). The NGR motif is essential for the capsid assembly and/or capsid protein expression (31).

In summary, 3 genotypes of porcine NoVs were detected from US swine. One

genotype (QW101-like) of porcine NoVs was genetically and antigenically most closely

related to human GII NoVs. Recombinant GII porcine NoV strains were identified and

characterized. The QW101-like NoVs infected Gn pigs with NoV particles evident in

intestinal contents. These results raise questions of whether pigs may be reservoirs for

emergence of new human NoVs or if porcine/human GII recombinants could emerge.

2.6 ACKNOWLEDGMENTS

This work was supported by grants from NIAID, National Institutes of Health

(Grant R01 AI 49742) and NRI, US Department of Agriculture (CGP Grant 1999 02009)

and by the Ohio Agricultural Research and Development Center (OARDC), The Ohio

State University (Graduate Student Research Enhancement Grant project 2002-114).

We thank Dr. Kim Green for providing the VLPs for the human NoV strains, Dr.

Duping Zheng (Centers for Disease Control and Prevention) for assistance in sequence

analysis, and Dr. S. Sreevatsan and Dr. J. LeJeune for critical review of the manuscript.

114 Sequencing was performed at the Plant-Microbe Genomics Facility of The Ohio State

University and the University of Wisconsin.

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119 Genus/Genogroup- Genbank Genus/Genogroup- Genbank Strain genotype b Abbreviation accession # Strain genotypeb Abbreviation accession # Hu/Norwalk/68/US Noro/GI-1 Norwalk M87661 Hu/Amsterdam/98-18/98/NET NoV/GII-8 Amsterdam AF195848 Hu/Hawaii/71/US Noro/GII-1 Hawaii U07611 Hu/SaitamaU25/97-99/JP NoV/GII-8 SaitamaU25 AB039780 Hu/Melksham/89/UK Noro/GII-2 Melksham X81879 Hu/VA97207/97/US NoV/GII-9§ VA97207 AY038599 Hu/Snow Mountain/76/US Noro/GII-2* SMV AY134748 Hu/NLV/Erfurt/546/00/DE NoV/GII-10 Erfurt AF427118 Hu/Mexico/89/MX Noro/GII-3 Mexico U22498 Hu/Mc37/00-01/THA NoV/GII-10* Mc37 AY237415 Hu/Toronto/91/CA Noro/GII-3 Toronto U02030 Po/Sw43/97/JP NoV/GII-11 Sw43 AB074892 Hu/SaitamaU18/97-99/JP Noro/GII-3 SaitamaU18 AB039781 Po/Sw918/97/JP NoV/GII-11 Sw918 AB074893 Hu/SaitamaU201/98/JP Noro/GII-3 SaitamaU201 AB039782 Po/MI-QW48/02/US NoV/GII-11 QW48 AY823303 Hu/Arg320/ARG Noro/GII-3* Arg320 AF190817 Hu/Gifu/96/JP NoV/GII-12§ Gifu AB045603 Hu/Camberwell/101922/94/AUS Noro/GII-4 Camberwell AF145896 HU/Wortley/90/UK NoV/GII-12* Wortley AJ277618 Hu/Lordsdale/93/UK Noro/GII-4 Lordsdale X86557 Hu/SaitamaU1/97-99/JP NoV/GII-12* SaitamaU1 AB039775

12 Hu/Bristol/93/UK Noro/GII-4 Bristol X76716 Hu/Fayetteville/98/US NoV/GII-13 Fayetteville AY113106

0 Hu/MD145-12/87/US Noro/GII-4 MD145 AY032605 Hu/M7/99/US NoV/GII-14 M7 AY130761 Hu/Farmington Hills/02/US Noro/GII-4 Farmington Hills AY502023 Hu/J23/99/US NoV/GII-15 J23 AY130762 Hu/Langen1061/02/DE Noro/GII-4 Langen AY485642 Hu/Tiffin/99/US NoV/GII-16 Tiffin AY502010 Hu/Hillingdon/93/UK Noro/GII-5 Hillingdon AJ277607 Hu/Neustrelitz260/00/DE NoV/GII-16 Neustrelitz AY772730 Hu/New Orleans 306/94/US Noro/GII-5 New Orleans AF414422 Hu/CS-E1/02/US NoV/GII-17 CS-E1 AY502009 Hu/Baltimore/274/1993/US Noro/GII-6 Baltimore AF414408 Po/OH-QW101/03/US NoV/GII-18 QW101 AY823304 Hu/SaitamaU3/97/JP Noro/GII-6 SaitamaU3 AB039776 Po/OH-QW125/03/US NoV/GII-18 QW125 AY823305 Hu/SaitamaU4/97/JP Noro/GII-6 SaitamaU4 AB039777 Po/OH-QW170/03/US NoV/GII-19§ QW170 AY823306 Hu/SaitamaU16/97/JP Noro/GII-6 SaitamaU16 AB039778 Po/OH-QW218/03/US NoV/GII-19§ QW218 AY823307 Hu/SaitamaU17/97/JP Noro/GII-6 SaitamaU17 AB039779 Bo/Newbury-2/76/UK NoV/GIII-2 Newbury-2 AF097917 Hu/Seacroft/90/UK Noro/GII-6* Seacroft AJ277620 Hu/Alphatron/98-2/98/NET NoV/GIV Alphatron AF195847 Hu/Leeds/90/UK Noro/GII-7 Leeds AJ277608 Mu/MNV-1/03/US NoV/GV MNV-1 AY228235 Hu/Gwynedd/273/94/US Noro/GII-7 Gwynedd AF414409 Po/Cowden/80/US sapovirus/GIII-1 Cowden AF182760 a Classification is based on the capsid gene sequences. The 5 porcine NoV sequenced in this study are bolded. b The * indicates previously reported recombinants (7, 9, 11, 15, 28) and the § indicates the potential recombinants found in this study.

a Table 2.1 Classification and GenBank accession numbers of NoV strains and a sapovirus strain used for sequence analysis .

Genotype/ ORF1-ORF2 VP1 ORF2-ORF3 VP2 3'-UTR Strain overlap (nt)a (aa)a overlap (nt) (aa) (nt) Genotype 11/Sw43b 17 547 NAc NA NA Genotype 11/Sw918b 17 547 NA NA NA Genotype 11/QW48 17 547 1 253 57

Genotype 18/QW101 20 557 1 275 48 Genotype 18/QW125 20 557 1 275 48

Genotype 19/QW170 17 548 1 254 51

12 Genotype 19/QW218 17 548 1 254 51 1 a nt=nucleotide, aa=amino acid. b See Table 2.1 for summary of these 2 reference porcine GII NoVs. cNA=Not available.

Table 2.2 The sizes of the deduced capsid protein VP1 and the minor capsid protein VP2, the overlap regions, and the 3’-UTR of porcine GII NoVs.

Complete capsid

(S domain, P domain) Hu/GI/ Bo/GIII/ Hu/GIV/ Mu/GV/ Strain Po/GIIa Hu/GIIb Norwalk Newbury-2 Alphatron MNV-1

96-98 63-71 43 45 53 39 QW48 (100, 94-97) (77-85, 53-63) (59, 36) (62, 36) (71, 42) (58, 29)

QW101 70-70.6 61-71.4 42 45 54 39 QW125 (83, 63) (77-86, 51-64) (59, 35) (62, 38) (71, 44) (58, 28) 12

2 QW170 81 62-69 43 45 53 39 QW218 (90, 74) (77-82, 52-62) (59, 36) (61, 37) (72, 40) (60, 27)

a Includes Sw43 and Sw918 strains. b Includes Hawaii, Snow Mountain, Mexico, MD145, New Orleans, Baltimore, Gwynedd, Amsterdam, VA97207, Erfurt, Gifu, Fayetteville, M7, J23, Neustrelitz strains.

Table 2.3 Summary of the percent amino acid identities of NoVs within the capsid region.

ELISA Antibody titer with each VLP antigen (genogroup-genotype) Hawaii Toronto MD145 HS66 Florida Desert Shield Antiserum (GII-1) (GII-3) (GII-4) (GII-4) (GII-6) (GI-3) HS66CS (positive control) Human convalescent 25,600 6,400 25,600 25,600 6,400 6,400 antiserum to human HS66 (GII-4)

LL616 Pig convalescent antiserum 100 800 400 400 400 10 to porcine QW126 a

12 (QW101-like, GII-18) 3 LL368 (negative control) <10 <10 <10 <10 <10 <10 Pre-inoculation serumb

MM982 (negative control) <10 <10 <10 <10 <10 <10 Pre-inoculation serumb

a The QW126 shared 99% and 100% amino acid identities to the QW101 strain (GII-18) for a 211 bp-segment in RdRp region and a 402 bp-segment in the capsid region, respectively. b LL368 and MM982 were sera from 2 gnotobiotic pigs before inoculation with porcine NoVs. Table 2.4 Antigenic cross-reactivity between human GII NoV antigens (VLPs) and a pig convalescent antiserum

against porcine GII NoVs as determined by ELISA.

69 Gifu96 100 A. Wortley GII-12 93 SaitamaU1 80 Hawaii GII-1 Neustrelitz 99 100 Tiffin GII-16 100 Erfurt Mc37 GII-10 100 Melksham 85 81 SMV GII-2 Hillingdon 58 GII-5 100 New Orleans 20 Fayetteville GII-13 98 CS-E1 GII-17 Arg320 100 Mexico 93 Toronto GII-3 84 SaitamaU18 41 98 SaitamaU201 68 Baltimore 100 Seacroft 100 SaitamaU3 99 SaitamaU4 GII-6 SaitamaU16 100 SaitamaU17 37 100 Leeds90 67 GII-7 Gwynedd 100 M7 GII-14 VA97207 GII-9 98 Amsterdam 91 100 SaitamaU25 GII-8 100 QW101 QW125 GII-18 100 100 QW170 80 QW218 GII-19 100 Sw918 100 Sw43 GII-11 80 QW48 99 100 Farmington Hills Langen 100 100 MD145 Camberwell GII-4 100 Bristol 100 Lordsdale J23 GII-15 Alphatron GIV Norwalk GI 0.05 (Continued) Fig. 2.1. Neighbor-Joining phylogenetic trees of genogroup II NoVs. The 5 newly identified porcine NoV strains are in bold. A. Tree was based on the complete capsid region. B. Tree was based on the partial RdRp region (C- terminal 260-266 aa). Genogroups (G) and genotypes (numbers followed G) are indicated. The human NoV GI-1/Norwalk and GIV/Alphatron strains were used as out-group controls.

124 Gifu Recombinant B. 84 60 SaitamaU1 Recombinant 24 Mc37 Recombinant Farmington Hills 31 GII-4 99 Langen Hawaii GII-1 41 64 Arg320 Recombinant MD145 64 37 Camberwell GII-4 92 Bristol 96 Lordsale 57 Snow Mountain Recombinant 49 Melksham GII-2 100 Neustrelitz GII-16 Mexico SaitamaU201 32 100 GII-3 SaitamaU18 98 Toronto QW101 GII-18 100 QW125 Recombinant 76 QW170 95 QW218 Recombinant 51 Sw43 100 Sw918 GII-11 QW48 GII-8 26 SaitamaU25 SaitamaU16 99 GII-6 100 SaitamaU17 100 VA97207 Recombinant SaitamaU3 63 GII-6 98 SaitamaU4 Norwalk GI

0.05 Fig. 2.1. Continued.

125 A.

3’-end RdRp S P w

o 100

d Sw43 n i to y 80 entit thin W d i w

e i 60 d

40 QW101

20 pe sequences y t o Percent nucleoti 0 gen 1000 2000 Nucleotide Position of center of window (Continued) Fig. 2.2. Identification of QW170 and QW218 strains as potential recombinants. A. Recombination Identification Program analysis of QW170 strain. At each position of the window, the query sequence (QW170) was compared to each of the background genotype representatives (GII-11/Sw43 and GII-18/QW101). When the query sequence is similar to the background sequences, the homologous regions are indicated as thick lines on the plot. Analysis parameters were Window size of 100 and Statistical significance of 90%. The nucleotide positions of the 3’-end RdRp and the S and P domains of the capsid protein are indicated. B. Sequence alignments of the RdRp-capsid junction region of NoVs. The genomic and subgenomic conserved 18 nucleotide motif was indicated by a horizontal line with 2 vertical bars. Asterisks indicate the identical residues to the sequence of the first line. Dashes represent gaps. The letter N indicates missing data on the residue. The start codon of ORF2 is underlined. Five NoV genogroups are indicated.

126 B. Genomic-subgenomic RNA conserved 18 nt motif

MD145 GTGAATGAAG ATGGCGTCGA GTGACG CCAACCCATC Sw43 ********** ********T* *C**** *TGCT***** Sw918 ********** ********T* *C**** **CCT***** QW48 ********** ********T* A***** *TGC****** QW170 ********** ********T* ****** *TGCT***** QW218 ********** ********T* ****** *TGCT***** QW101 ********T* *****A**** A***T* *T*C*G*TC* QW125 ********T* *****A**** A***T* *T*C*G*TC* VA97207 ********** ********** A***** *AGCT***** SaitamaU25 ********** ********** A***** *AGCT***** New Orleans ********** ********** A***** *T*CT***** SaitamaU16 ********** ********** A***** *TGCT***** Toronto ********** ********** A***** *TGCT***** Melksham ********** ********** A***** **GCT***** GII Snow Mountain ********** ********** A***** **GCT***** Mc37 ********** ********** A***** **GCT***** CS-E1 ********** ********** A***** **GCT***** Tiffin ********** ********** A***** **GCT***** Hawaii ********** ********** A***** **GC****** Gifu ********** ********** ****** **GCT***** SaitamaU1 ********** ********** ****** **GCT***** Fayetteville NNNN****** ********** A***** **GCT***** Neustrelitz NNNNN***** ********** A***** **GCT***** J23 NNNN****G* ********** ****** *GCC*GTT** M7 NNNN****** ********** A***** *T*CT***** Arg320 ********** ********** A***** ***CT***** SaitamaU3 ********** ********** A***** **GCT***** Gwynedd ********** ********** A***** *AGCT***** GIII Newbury-2 **A******* ***A*TGAC* *A***A TT*CTGAT** GI Norwalk **A*****T* ********T* AG**** *T*CAT**AG GIV Alphatron NNNN****** ********** ****** *TGCT***** GV MNV-1 ********G* ***AG---TG A**G** *AGCG***AA

Fig. 2.2. Continued.

127 100 nm

Fig. 2.3 Immune electron micrograph of porcine NoVs. The diluted intestinal contents of a Gn pig euthanized on PID 5 to QW101-like porcine NoVs (QW144) was incubated with convalescent serum LL616 from another Gn pig inoculated with QW101-like porcine NoVs (QW126) and visualized by negative staining with 3% phosphotungstic acid. The arrow bar indicates a small clump of NoV-like particles. The scale bar is 100 nm.

128 S domain (50-236 aa)

60 70 80 120 190 200 210 220 250 MD145 IIDPWIRNNF VQAPGGEFTV SPRNAPGEIL … VILAGNAFTA … LKLIAMLYTP LRANNAGDDV FTVSCRVLTR PSPDFDFIFL … EMSNSRFPIP … Hawaii V******M** ****N***** ****S***** … *L******** … MR*V****** **S*GS**** ********** *******NY* … *L******V* … Snow Mountain *******A** ****N***** ********V* … *M******** … MRIV****** **S*GS**** ********** *******TY* … *L******VS … QW101 V******G** ****N***** ****S***V* … *V******A* … *R*V****** ****SPT*** ********** *****E*T** … *L******A* … Toronto ******M*** ********** ****S***V* … *V******** … *R******** *****S**** ********** *****S*N** … ********V* … Baltimore *******E** ****Q***** ****S***M* … *V******** … MR*V****** ****S-*E** ********** *A***E*T** … *L******AA … Norwalk P*****I*** ****Q****I **N*T**DV* … IM******** … MR*VC***** **TGGGTG*S *V*AG**M*C *****N*L** … SL****A*L* …

P1A domain (237-279 aa) P2 domain (280-454 aa) P1B domain (455-576 aa) 12

9 270 280 290 300 400 420 430 470 480 MD145 AFVVQPQNGR CTTDGVLLGT TQLSAVNICN FRGDVTHIVG … HFTPKLGSVQ … TGQNTKFTPV GVIQDGD-HH … TFPGEQLLFF RSTMPGCSGY … Hawaii GVI******* S*L**E**** ***VPS***A L**RINAQ*P … K********I … LN*P*R**** *LFNT----- … L********* **HI*LKG*T … Snow Mountain VIS**C**** **L**E*Q** ***QVSG**A *K*E**AHLQ … QY*****Q** … VN*PV***** *LNDTEH--- … V****R**** **YL*LKG** … QW101 NI******** **L**E*Q** ***QPSL**S ***VTLNETS … RYA******E … LNDQ****** *LAHNRA--- … VY******** **NL*LAG*T … Toronto NI***C**** V*L**E*M** ***LPSQ**A ***TL*R--- … R*******LE … PN*P****** **GV*NE--- … N********* **QL*SSG*R … Baltimore SI******** **L**T*Q** ***VPTQ**A ***TLIS--- … KYA****TIL … *N*PIR**** *MGDN----- … S****RI*** **IV*SAG** … Norwalk VQS**F**** **L**R*V** *PV*LSHVAK I**TSNG--- … T*V*H***I* … GVLSWISP*S HPSGSQV--- … PGF**V*V** M*K***PGA* …

Fig. 2.4 Conserved amino acid motifs of the capsid proteins of NoVs. The S, P1A, P2 and P1B domains are divided by three single-arrow vertical bars after amino acid residue 236, 279 and 454 based on the aligned sequences of human GI-1/Norwalk, GII-1/Hawaii, GII-3/Toronto, GII-4/MD145, GII-6/Baltimore and porcine QW101 (bold) NoVs. Asterisks indicate the identical residues to the sequence of MD145 strain. Dashes represent gaps. The predicted conserved amino acid motifs of the capsid protein are underlined in the MD145 sequence. The motif IDPWI was confirmed by monoclonal antibody cross-reactivity as a common antigenic motif of NoVs.

CHAPTER 3

GENETIC DIVERSITY AND RECOMBINATION OF PORCINE SAPOVIRUSES

3.1 SUMMARY

Sapoviruses (SaV) are emerging enteric pathogens that cause diarrhea in humans and animals. To date, only two genetically similar porcine SaV strains have been reported that belong to genogroup III (GIII). Human SaVs are genetically variable and have been classified into 4 genogroups (GI, II, IV and V). To investigate the genetic diversity of porcine SaVs and their genetic relatedness to human strains, we sequenced 286 nt of the

RNA polymerase (RdRp) region of 9 porcine SaVs detected from field pig fecal samples collected in US swine farms during 1999-2003. One strain (Po/SaV/MI-QW19/2002/US) was most closely related to human GII SaVs. We also sequenced 3 kb of the viral genome including the partial RdRp (766-790 nt), the complete capsid, the ORF2 and the

3’-UTR of 4 strains representative for the positive farms or for the distinct genetic clusters. From the sequence analysis of the predicted complete capsid, we identified a

130 potential new genogroup of porcine SaVs with Po/SaV/OH-JJ681/00/US as the

representative strain. Further analysis of the predicted partial RdRp and the RdRp-capsid

junction regions also identified 2 strains, Po/SaV/NC-QW270/03/US and

Po/SaV/OH-MM280/03/US that were likely recombinants between 2 distinct porcine

SaV strains. To our knowledge this is the first report of a porcine SaV strain more closely

related genetically to human SaVs and the occurrence of porcine SaV recombinants. The

presence of porcine SaVs more similar to human SaVs is a significant finding because of

the potential for zoonotic infections or generation of porcine/human recombinants if

intra-genogroup human strains exist.

3.2 INTRODUCTION

Sapoviruses (SaV), previously referred to as Sapporo-like viruses, are emerging enteric pathogens that cause diarrhea in humans, pigs and mink (4, 5, 7, 8, 20). They are nonenveloped, polyadenylated single-stranded positive-sense RNA viruses and belong to the Sapovirus genus in the family Caliciviridae (5). The SaV genome is 7.3-7.5 kb long and contains two main ORFs based on sequence analysis (6, 13, 18, 24). The ORF1 encodes a polyprotein that undergoes protease processing to produce several non-structural proteins including a cysteine protease and an RNA-dependent RNA polymerase (RdRp), and a capsid protein. The ORF2 encodes a small basic protein with unknown function.

131 Human SaVs are primarily associated with 1.8-9% cases of pediatric acute gastroenteritis (2, 15, 22, 26), although SaV outbreaks in adults have been described (21).

Human SaVs are genetically variable viruses. They have been classified into 4 genogroups (GI, II, IV and V) and at least 8 genetic clusters or genotypes (GI/1-3,

GII/1-3, GIV/1 and GV/1) (2, 27). Porcine SaV Cowden strain was isolated from a

27-day-old diarrheic field pig (25). It causes diarrhea and intestinal lesions in gnotobiotic pigs (8). However, only two genetically closely related porcine SaV strains

(Po/SaV/Cowden/80/US and Po/SaV/LL14/02/US), sharing 96% nucleotide identity throughout the ORF1 and ORF2, were reported (1, 6). The complete genome of the

Cowden strain has been analyzed and it is classified as SaV GIII (6). It is the only cultivable SaV or enteric caliciviruses (1, 3, 23).

Recently, we identified porcine noroviruses, another genus of enteric caliciviruses causing diarrhea in humans and animals, that are genetically and antigenically similar to human noroviruses (Q. H. Wang, M. G. Han, S. Cheetham, M. Souza, J. Funk and L. J.

Saif, submitted for publication) raising questions of whether pigs may be reservoirs for emergence of human noroviruses. To investigate the genetic diversity of porcine SaVs and their relatedness to human SaVs, we chose a pair of calicivirus universal primers p290 and p110 targeting the conserved motifs “DYSKWDST” and “YGDD” of the RdRp region of caliciviruses (12, 17) to perform reverse transcription (RT)-PCR to screen for genetically variable SaVs in pigs. Nine SaVs were identified from field pig fecal samples

132 collected from US swine farms during 1999-2003. We further sequenced the 3’-end 3 kb including partial RdRp, the complete capsid and ORF2 regions of 4 strains representative for the positive farms or for the distinct genetic clusters. We then classified these newly identified porcine SaVs by phylogenetic analysis and a recombination identification program.

3.3 MATERIALS AND METHODS

Stool samples. A total of 377 fecal samples were collected from 8 swine farms

(OH farm A-E, MI farm A, NC farm A and B) and one OH slaughterhouse during April,

1999 to May, 2003 and used to survey for the presence of genetically diverse enteric caliciviruses. Nine SaV-positive pig fecal samples were identified by RT-PCR with a calicivirus universal primer pair p290/110 (12, 17) followed by sequencing of the

RT-PCR products. The age and diarrhea status of the pigs from which these 9 samples were obtained is summarized in Table 3.1. The MM280 strain (the large intestinal contents of a gnotobiotic pig) was the third passage of one field pig fecal sample performed using Gn pigs as previously reported (8). This sample was amplified in pigs because the SaV amount in the original specimen was inadequate for analysis. Fresh fecal samples or intestinal contents were placed into sterile specimen containers and stored frozen until tested.

RNA extraction. The RNA was extracted from the 10% (wt/vol) fecal

133 suspensions using the Trizol LS (Invitrogen, Corporation, Carlsbad, CA) procedure. For

amplification of the 3’-end 3 kb fragment of these samples, except for strain MM280, the

extracted RNA was further concentrated and purified by using the QIAamp Viral RNA

Mini kit (Qiagen Inc., Valencia, CA). For amplification of the 3’-end 3 kb fragment of

QW19, 60 ml of a 10% fecal suspension was filtered (0.45 µm) and semipurified through a 40% (w/v) sucrose-cushion by ultracentrifugation at 112,700 × g for 2 h. The pellets were resuspended in 1 ml of sterile Eagle’s minimal essential medium (EMEM,

Invitrogen Corporation, Carlsbad, CA) for RNA extraction and purification.

RT-PCR. (i) RT and PCR were performed separately using primer pair p290 (5’-

GAT TAC TCC AAG TGG GAC TCC AC -3’) (12) and p110 (5’-ACD ATY TCA TCA

TCA CCA TA-3’) (17) as previously reported (Q. H. Wang, M. G. Han, S. Cheetham, M.

Souza, J. Funk and L. J. Saif, submitted for publication). Products of primer pair p290/110 were 329 bp for SaVs.

(ii) To amplify the 3’-end, 3 kb fragment of the porcine SaVs, the cDNA was

synthesized by SuperScript III First-Strand cDNA synthesis kit (Invitrogen) according to

the manufacturer’s instructions with primer VN3T20 (5’-GAG TGA CCG CGG CCG

CT20-3’) (Q. H. Wang, M. G. Han, S. Cheetham, M. Souza, J. Funk and L. J. Saif, submitted for publication). Then the first PCR was performed using TaKaRa Ex TaqTM polymerase (TaKaRa Mirus Bio, Madison, WI) with primers p290 and VN3T20. For strains QW270 and JJ681, a second PCR was necessary to produce a visible band, using

134 an internal forward primer PSV4b (5’-GCA TGC CC(/G)T TCA CCA GTG T-3’) and

PEC68 (5’-CCG CTA TAA ATT TAT TGG GTG-3’) designed based on the obtained 286

nt RdRp region of the QW270 and JJ681, respectively, and the reverse primer VN3T20.

After initial denaturation at 94ºC for 3 min, the first 5 cycles were performed at 94ºC for

30 sec, 50ºC for 30 sec and 72ºC for 5 min, then another 30 cycles with a shorter elongation time of 3 min, and a final extension for 10 min at 72ºC. All RT-PCR products were analyzed by agarose gel electrophoresis and stained with ethidium bromide. The

RT-PCR products were visualized using ultraviolet light.

Cloning and sequencing. The RT-PCR products were purified by QIAquick Gel

Extraction kit (Qiagen Inc.). Products of primer pair p290/110 were sequenced directly using primers p290 and p110 or after cloning into pCR2.1 (T/A) vector (Invitrogen). The

~3 kb RT-PCR products were cloned by using PCR XL cloning kit (Invitrogen). Plasmid

DNA was extracted using an alkaline lysis method. The insertion was confirmed by restriction enzyme Eco RI digestion. Three to five clones of each sample were sequenced by a primer-walking strategy. DNA sequencing was done using BigDye Terminator Cycle chemistry and 3700 or 3730 DNA Analyzer (Applied Biosystems, Foster, CA).

Sequence analysis. Sequence editing was performed by Lasergene software package (version 5) (DNASTAR Inc., Madison, WI). Basic Local Alignment Search Tool

(BLAST, http://www.ncbi.nlm.nih.gov) was used to find homologous hits. Multiple

sequence alignment was done by ClustalW (version 1.83) at DDBJ

135 (http://www.ddbj.nig.ac.jp). Phylogenetic analysis (Neighbor-Joining) with bootstrap

(1,000 replicates) and amino acid pairwise distance analysis (p-distance) were conducted using MEGA v 2.1 (16).

Recombination identification. Identification of recombinants was performed by

comparing a query sequence to a set of background sequences by using Recombinant

Identification Program (28) (http://hivweb.lanl.gov/RIP/RIPsubmit.html) and creating separate phylogenetic trees for the segments resembling different genetic clusters of the query sequence.

Classification and GenBank accession numbers. The classification and

GenBank accession numbers of published representative strains for phylogenetic analysis

are listed in Table 3.2.

3.4 RESULTS

Genetically diverse porcine SaVs were detected. A total of 35 of 377 samples

(9%) were potentially positive by RT-PCR with the primer pair p290/110. We sequenced

the RT-PCR products (317 or 329 bp products) of 23 samples, at least 1 sample from

each positive farm at one sampling time. After performing BLAST search (blastn and

blastx), we identified 9 SaVs (Table 3.1) and 7 noroviruses. Seven sequences did not

have any significant hit in the database and were not analyzed further. All the SaVs and

noroviruses contained the conserved amino acid motif “GLPSG” of calicivirus RdRp.

136 The SaV-specific sequence was 286 nt encoding 95 aa of the RdRp region.

Neighbor-joining phylogenetic analysis was performed based on the deduced 95 aa RdRp sequences of porcine, human and mink SaVs. The

Ra/Lagovirus/RHDV/GH/1988/GE (RHDV), Fe/Vesivirus/FCV/F9/1958/US (FCV),

Hu/Norovirus/Norwalk/68/US (Norwalk) and Bo/Nebraska/80/US (NB, a potentially new genus) are representative strains of Lagovirus, Vesivirus and Norovirus genera and the NB-like viruses, respectively, within the Caliciviridae family (5, 29) (Fig. 3.1). We found that JJ259, II166, II176 and QW270 grouped with the porcine SaV prototype

Cowden strain and LL14 strain, sharing 91.6-98.9% amino acid identities. The QW152 and MM280 were similar to each other and shared 81.1% amino acid identities with the

Cowden strain, whereas the QW19, JJ681 and LL26 strains segregated into 3 new branches distant from the Cowden group. Interestingly, the QW19 strain shared the highest amino acid identity (66.3%) with human GII Mc10, C12 and Cruise ship strains.

In addition, the JJ681 strain shared the highest amino acid identity (63.2%) with the

LL26 strain, but both showed low amino acid identities to SaVs (25.3-47.4%), FCV

(31.6-33.7%), RHDV (33.0-34.0%), NB (32.6-38) and Norwalk (19.8-24.2%).

A potential new genogroup of porcine SaVs was found based on the complete capsid sequences. To genetically classify the newly identified porcine SaVs, we choose the QW270, MM280, JJ259, JJ681, QW19 and LL26 as representative strains for further analysis. Attempts to amplify the QW19 and LL26 strains, which included concentrating

137 and purifying the viral RNA and performing second PCR using internal forward primers, were unsuccessful. We also tried, but failed to detect another QW19-like strain from the

60 pig fecal samples collected at the same time from the same farm as the QW19 strain.

For the RT-PCR, we used a pair of QW19-specific primers PSV6 (5’-CGG TCA TTT

TGT GTG GAC TG-3’) and PSV7 (5’-ATT GCC CGT ATA AGG CAC A-3’) based on the obtained 286 nt of the RdRp region.

We successfully amplified and sequenced the 3’-end 3 kb of genome containing the partial RdRp (263 aa for the QW270, MM280 and JJ259 strains and 255 aa for the

JJ681 strain), capsid protein and ORF2 genes and the 3’-UTR of QW270, MM280, JJ259 and JJ681 strains. The sizes of the predicted capsid and the ORF2 proteins, the

ORF1-ORF2 overlap region and the 3’-UTR of each newly identified strain and the porcine SaV prototype Cowden strain are summarized in Table 3.3. The QW270 and

MM280 strains had exactly the same sizes for each region as the Cowden strain. The

JJ259 strain had the same size capsid, ORF1-ORF2 overlap and the 3’-UTR as those of the Cowden strain, but not the ORF2 protein. For the JJ681 strain, only the ORF1-ORF2 overlap region was identical in size and nucleotide sequence (ATGA) to that of the other

4 strains; the other targeted region sizes, especially for the 3’-UTR, were highly divergent from the other SaVs.

The amino acid and nucleotide percent identities of the predicted complete capsid protein sequences of the 4 newly identified porcine SaVs, the previously reported porcine

138 and human SaVs, and one representative strain of each of the other 4 genera within the

Caliciviridae family: RHDV, Norwalk, FCV, and NB (a potentially new genus) are

summarized in Table 3.4. In the complete capsid region, the QW270, MM280 and JJ259

strains were genetically closest to the porcine SaV prototype Cowden strain with an

amino acid identity of 88-95%. The JJ681 strain differed genetically from these strains,

but showed similar amino acid (32-36%) and nucleotide (45-48%) identities to those of

the different genogroups of SaVs. These percent identities were higher than those to

representative strains of the other four genera: NB (23% amino acid and 40% nucleotide

identities), RHDV (21% amino acid and 39% nucleotide identities), FCV (23% amino

acid and 39% nucleotide identities) and Norwalk strains (17% amino acid and 34% nucleotide identities). The previously reported SaV intra-genus sequence identity was somewhat higher than that noted for JJ681 strain, ranging from 37-49% for amino acids and 49-58% for nucleotides (2). The inter-genus amino acid identity of caliciviruses is

15-27% (5, 29). A Neighbor-Joining phylogenetic tree based on the predicted amino acid sequences of the complete capsid (Fig. 3.2A) revealed that the QW270, MM280 and

JJ259 strains grouped together with the Cowden strain into GIII. The JJ681 and other

SaVs segregated into different groups from a common ancestor with a high bootstrap value (99%). Based on this data, we tentatively classified the JJ681 into a new genogroup

(JJ681-like, GVI?) within the Sapovirus genus.

The QW270 and MM280 were identified as potential porcine SaV

139 recombinants. Further analysis of the predicted C-terminal ~260 aa of the RdRp region

showed that all strains contained another RdRp conserved motif “YGDD”. The

phylogenetic tree based on the nucleotide sequence of this partial RdRp region (Fig. 3.2B)

showed similar classification results to those based on the capsid sequence of the

corresponding strain (Fig. 3.2A) for the JJ259 and JJ681 strains, but the RdRp and capsid

assignments differed for the QW270 and MM280 strains. The QW270 strain shared only

83.4% nucleotide identity with the Cowden strain in the partial RdRp region, whereas it

shared much higher nucleotide identity (96.5%) to the JJ259 strain. The MM280 strain

shared lower nucleotide identity with the Cowden strain in this RdRp region (76.2%)

compared to that of the capsid protein (89.1%). Generally, the RdRp region is more

conserved than the capsid region. Therefore, these results indicated that the QW270 and

MM280 could be recombinants with RdRp and capsid genes from 2 different parent

strains.

The predicted ORF2 protein sequences of the 4 representative strains were also analyzed. Similar to the classification based on the capsid protein, QW270, MM280 and

JJ259 were grouped into the same cluster as the Cowden strain, whereas the JJ681 strain again formed a new branch distant from the other SaVs. The JJ259 had a 9 aa-insertion

(GVTTTPKPQ) compared to the other strains in this cluster (Table 3.3). The QW270 strain was most similar to the Cowden strain sharing 95% amino acid identity, but only sharing 87% amino acid identity with the JJ259 strain.

140 To determine where the recombination occurred for the QW270 and MM280

strains, we performed the Recombinant Identification Program by placing the 3’-end

3-kb, from the C-terminal RdRp (aligned 843 nt) to just before the polyA tail, of the

QW270 and MM280 strains as a query sequence, respectively, and the corresponding sequences of Cowden (and JJ259) as background sequence(s). The QW270 strain showed high homology to the JJ259 strain in the RdRp region, but it changed to shared high homology to Cowden strain in the capsid protein and ORF2 protein regions (Fig. 3.3A).

In Fig. 3.3B, the MM280 strain abruptly shared higher homology to Cowden strain after the RdRp region although there was low homology between these two strains in the partial RdRp region. The abrupt homology changes in both the QW270 and MM280

strains occurred in the RdRp-capsid junction region. The nucleotide alignments around this region (Fig. 3.4) of SaVs revealed that sequence conservation exists among strains within each genogroup. These results indicate that QW270 was a potential recombinant between a JJ259-like strain (RdRp region) and a Cowden-like strain (capsid, after RdRp), and the MM280 was a potential recombinant between an unknown strain (RdRp region), which may represent a new genotype within GIII SaVs, and a Cowden-like strain (capsid, after RdRp).

3.5 DISCUSSION

Since the discovery of porcine SaVs in the 1980s (25), few further studies have

141 been done to investigate their genetic diversity, the possibility of zoonotic transmission, and the disease spectrum of field strains. In this study, we investigated the genetic diversity of porcine SaVs and their genetic relationship to human and mink SaVs, and to the other four genera (Lagovirus, Vesivirus, Norovirus and NB-like viruses) in the

Caliciviridae.

To date, the complete genomes of only 5 SaV strains have been sequenced (6, 13,

18, 24) including 4 human SaV strains (Manchester, Bristol, Mc10 and C12) and the porcine Cowden strain. The classification of SaVs based on the complete capsid gene sequences is generally accepted although the consistency and reliability of SaV classification based on different regions, such as the complete and partial RdRp and the

ORF2 protein, of the genome remains to be investigated (2, 27). In this study, we found that the classification based on the complete capsid was consistent with that based on the larger partial RdRp (~260 aa) of most SaV strains except for the GIV/Hou7 strain and the potential recombinant strains (Mc10 or C12, QW270 and MM280) (Fig. 3.2). The short segment (95 aa) of the RdRp region could be used to identify genogroups but it did not reliably identify genotypes. For example using this fragment, the GI-1/Manchester strain was distinguished from the other three GI-1 strains (Sapporo, Houston86 and Lyon30388) and clustered with GI-2/Houston90 (Fig. 3.1). Based on the 95 aa RdRp segment, the

QW19 strain shares 66.3% amino acid identity to human GII Mc10, C12 and Cruise ship strains and the LL26 shares 63.2% amino acid identity to the JJ681 strain. The overall

142 intra-genogroup amino acid identity of the 95 aa RdRp region of SaVs is 80-100%. Thus

the QW19 and LL26 strains may belong to two new genogroups. However, no further

classification of the QW19 and LL26 strains was possible due to a lack of additional

sequence data.

In the predicted complete capsid region, the JJ681 strain showed similar amino acid identities (32-36%) to each SaV genogroup (GI-GV), which were higher than those

(17-23%) between JJ681 and representative strains (NB, RHDV, FCV and Norwalk) of the other four genera of Caliciviridae or the overall inter-genus amino acid identities

(15-27%) for caliciviruses (5, 29). The JJ681 strain also had the same size and identical nucleotides in the ORF1-ORF2 overlap region as the other porcine SaVs. However, in the RdRp-capsid junction region, which is conserved within each genogroup (Table 3.4), the JJ681 strain differed from all the other SaV genogroups (GI-V). Therefore, we tentatively classified the JJ681 strain as a new genogroup (JJ681-like, GVI?) within the

Sapovirus genus. Our results indicate that porcine SaVs circulating in US swine herds are genetically diverse and comprise at least 2 genogroups (GIII and JJ681-like) and may comprise 2 other potential new genogroups (QW19-like and LL26-like).

Among porcine SaVs, the QW19 strain is genetically closest related to human

GII SaVs. A similar sequence was found in GenBank (AY615810,

Po/Sapo/SWECII/VA14/NET, van der Heide et al., unpublished). These 2 strains share

100% amino acid and 84% nucleotide identities in the 286 nt (95 aa) RdRp region, the

143 region produced as the products of primer pair p290/110. This indicates that other

porcine SaVs closely related to human SaVs exist. Whether human SaV strains similar to

the QW19-like porcine SaVs circulate among swine farm workers is unknown, but such

studies could provide information on the zoonotic potential of these porcine SaVs.

We tried to passage the QW19 strain in 2 Gn pigs orally and intranasally

inoculated with QW19 fecal filtrates. No shedding of the QW19 SaV was detected in

feces by RT-PCR, or by immune electron microscopy using convalescent serum to this

strain from one of the above pigs. Only rotavirus particles were observed by immune

electron microscopy using the convalescent antiserum. The rotavirus infection may have

interfered with replication of the SaV QW19 strain. Although 60 other fecal samples

were collected and tested from the same farm (MI A) from which QW19 was identified,

no other QW19-like strains were detected. Also, no calicivirus-like particles were

observed in the original QW19 fecal specimen by electron microscopy (EM, data not

shown). Based on these limited results, possibly the QW19 strain was introduced only

occasionally into individual pigs. Potentially it did not replicate to high titer in the pig

host (EM negative, RT-PCR weakly positive) or spread efficiently in the herd, suggesting

a possible alternative host origin such as from humans based on the phylogenetic

analysis.

The SaV recombinant strain (Mc10 or C12) of human origin (13) and several norovirus recombinants of human, bovine and porcine origin have been reported (9-11,

144 14, 19, 30). All the recombination events occurred in the RdRp-capsid junction region

and exclusively within the genogroups of each genus, consistent with the nucleotide

conservation seen among strains within a genogroup in this junction region, facilitating

homologous recombination. The QW270 was a potential recombinant strain between a

JJ259-like strain (RdRp region) and a Cowden-like strain (capsid, after RdRp) of the

same genotype and the MM280 was a potential recombinant strain between 2 parent

strains ( an unknown strain for the RdRp region and a Cowden-like strain for the capsid region) belonging to different genotypes within the same genogroup. We did not detect a strain carrying an RdRp similar to that of MM280.

Interestingly for the human SaVs analyzed, the GIV/Hou7 strain is more closely related to GII SaVs in the partial RdRp region (~260 aa). It shares the highest nucleotide identity with the GII/Mc10 strain (78.4%) and C12 strain (78.3%), which is within the range of nucleotide identity (74.9-96.3%) among GII SaVs, whereas it shares lower but similar nucleotide identities (47.3-58.1%) to GI-III and GV SaVs. The GIV/Hou7 strain shares a 26 nt-motif with the GII SaVs in the RdRp-Capsid junction region (Fig. 3.4).

Therefore, the Hou7 strain may be a recombinant between an unknown GII strain (RdRp segment) and a GIV strain (after RdRp segment), although recombinants between two distinct genogroups of SaVs have not been described previously.

For GI, GIV and GV human SaVs, another ORF overlapping the 5’-end of the capsid gene was proposed because there is a conserved translation initiation motif

145 GCAAUGG at the 5’-end of this ORF (2, 27). We did not find this motif in the capsid

sequences of QW270, MM280, JJ259 and JJ681 strains. It will be of interest to determine

if QW19-like porcine SaVs which are more closely related to human SaVs have this

motif.

In summary, we choose the pair of calicivirus universal primers p290 and p110 targeting the conserved motifs “DYSKWDST” and “YGDD” of the RdRp region of caliciviruses (12, 17) to screen for genetically variable caliciviruses in pigs. In this study we identified genetically diverse porcine SaVs comprising at least 2 genogroups (GIII and JJ681-like GVI?) and possibly comprising 2 other potentially new genogroups

(QW19-like and LL26-like). One porcine SaV strain QW19 was genetically most similar to human SaVs suggesting the possibility of a pig reservoir for human strains or vice versa. We also identified 2 potential recombinant porcine SaV strains (QW270 and

MM280). To our knowledge, this is the first report of porcine SaV recombinants.

3.6 ACKNOWLEDGMENTS

This work was supported by grants from the National Institute of Allergy and

Infectious Diseases, National Institutes of Health (Grant R01 AI 49742) and NRI, US

Department of Agriculture (CGP Grant 1999 02009) and by the Ohio Agricultural

Research and Development Center (OARDC), The Ohio State University (Graduate

Student Research Enhancement Grant project 2002-114). Salaries and research support

146 were provided by state and federal funds provided to the OARDC.

We thank Dr. Duping Zheng (Centers for Disease Control and Prevention) for assistance in sequence analysis. We thank Dr. S. Sreevatsan and Dr. J. LeJeune for critical review of the manuscript. We also thank the Plant-Microbe Genomics Facility of

The Ohio State University and McGill University, Canada for DNA sequencing.

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150

Sample Date of Swine GenBank Strain Pig age Diarrhea number sampling farm accession # Po/SaV/OH-II166/99/US II166 Apr., 1999 OH D post-weaning No AY974193 Po/SaV/OH-II176/99/US II176 May, 1999 OH C post-weaning Yes AY974194 Po/SaV/OH-JJ259/00/US JJ259 June, 2000 OH C post-weaning Yes AY826423 Po/SaV/OH-JJ681/00/US JJ681 Dec., 2000 OH E post-weaning Yes AY974192 Po/SaV/OH-LL26/02/US LL26 Apr., 2002 OH C post-weaning No AY974195 Po/SaV/MI-QW19/02/US QW19 Dec., 2002 MI A Finisher No AY826424 Po/SaV/OH-QW152/03/US QW152 Mar., 2003 OH B Finisher No AY826425 Po/SaV/OH-QW270/03/US QW270 June, 2003 NC B Finisher No AY826426 Po/SaV/OH-MM280/03/US MM280 Mar., 2003 OH B Finisher No AY823308

Table 3.1 Porcine sapovirus strains detected using primers p290/110 and sequenced in this study.

151

Genus/Genogroup- Genbank Strain genotype Abbreviation accession # Hu/Sapporo/82/JP SaV/GI-1 Sapporo U65427 Hu/Manchester/93/UK SaV/GI-1 Manchester X86560 Hu/Plymouth/92/UK SaV/GI-1 Plymouth X86559 Hu/Lyon30388/98/FR SaV/GI-1 Lyon30388 AJ251991 Hu/Houston/86/US SaV/GI-1 Houston86 U95643 Hu/Parkville/94/US SaV/GI-2 Parkville U73124 Hu/Houston/90/US SaV/GI-2 Houston90 U95644 Hu/Stockholm/97/SE SaV/GI-3 Stockholm AF194182 Hu/Mex14917/00/MX SaV/GI-3 Mex14917 AF435813 & AF435810 Hu/London/92/UK SaV/GII-1 London92 U95645 Hu/Lyon598/97/FR SaV/GII-1 Lyon598 AJ271056 Hu/Bristol/98/UK SaV/GII-1 Bristol AJ249939 Hu/Mex340/90/MX SaV/GII-2 Mex340 AF435812 & AF435809 Hu/Cruise ship/00/US SaV/GII-3 Cruise ship AY289804 & AY157863 Hu/Mc10/00/TH SaV/GII-4 Mc10 AY237420 Hu/C12/00/JP SaV/GII-5 C12 AY603425 Po/Cowden/80/US SaV/GIII Cowden AF182760 Po/LL14/02/US SaV/GIII LL14 AY425671 Hu/Hou7-1181/90/US SaV/GIV Hou7 AF435814 Hu/Argentina39/Arg SaV/GV Arg39 AY289803 & AF405715 Mink/MEC/1/1999/US SaV/G? MEC AF338404 Ra/RHDV/GH/1988/GE Lagovirus RHDV M67473 Fe/FCV/F9/1958/US Vesivirus FCV M86379 Hu/Norwalk/68/US Norovirus Norwalk M87661 Bo/Nebraska/80/US NB-like viruses NB AY082891

Table 3.2 Summary of sapovirus strains and representative strains for Lagovirus, Vesivirus, Norovirus genera and NB-like viruses used in sequence analysis.

152

Genogroup/ Capsid ORF1-ORF2 ORF2 protein 3'-UTR strain (aa)a overlap (nt)b (aa) (nt)

GIII/Cowden 544 4 164 55 GIII/QW270 544 4 164 55 GIII/MM280 544 4 164 55 GIII/JJ259 544 4 173 55 GVI?/JJ681 554 4 168 28 a aa=amino acid. b nt=nucleotide.

Table 3.3 The sizes of the deduced capsid protein, the ORF1-ORF2 overlap region, the ORF2 protein, and the 3'-UTR of porcine sapoviruses.

153

Genus/Genogroup- Sapovirus NB-like Lagovirus Vesivirus Norovirus Strain genotype viruses RHDV FCV Norwalk GI/1a GI-2 b GI-3 c GII-1 d GII-2 e GII-3 f GII-4 g GIII h GIV i GV j

40 39-40 39-40 38-39 38 39 40 95 39 43 24 24 24 18 Sapovirus /GIII QW270 (50-51) (48) (48) (50) (49-50) (50) (49) (90) (47) (51) (41) (42) (39) (38)

40-41 40 39-41 39 39-40 40 40 91-93 40 43 24 24 24 19 Sapovirus /GIII MM280 (50) (47) (48) (49) (49-50) (49) (50) (88-89) (49) (51) (40) (40) (38) (37)

39-40 39-40 38-39 36-37 38 38 38 88-90 39 43 23 23 24 18 Sapovirus /GIII JJ259 (48- (50-51) (49) (47-48) (50) (49) (50) (85-86) (49) (51) (41) (41) (38) (37) 49) 15 4 33-34 32-33 34-35 33-34 34-35 35 34 36 34 35 23 21 23 17 Sapovirus /GVI? JJ681 (47) (45) (46) (46-47) (46-47) (47) (46) (47-48) (46) (45) (40) (39) (39) (34)

a GI/1 includes Sapporo, Houston86, Manchester, Plymouth and Lyon30388. b GI-2 includes Houston90 and Parkville. c GI-3 includes Stockholm and Mex14917. d GII-1 includes London92, Bristol and Lyon598. e GII-2 includes Mes340 and Mc10. f GII-3 includes Cruise ship. g GII-4 includes C12. h GIII includes Cowden and LL14. i GIV includes Hou7. j GV includes Arg39.

Table 3.4 Summary of the amino acid (nucleotide) sequence identities (%) in the complete capsid region between the newly identified porcine sapoviruses and reference calicivirus strains.

50 Lyon598 85 Bristol 51 GII London92 100 Mex340 Hou7 GIV Cruise ship 33 Mc10 72 73 GII 74 C12 76 Manchester 100 Houston90 Parkville 52 80 Mex14917 GI Lyon30388 75 Sapporo 100 21 Houston86 QW19 MEC 100 MM280 47 QW152 89 98 Cowden 99 LL14 GIII 97 JJ259 II166 65 78 78 II176 68 QW270 73 Arg39 GV FCV JJ681 GVI? 100 LL26 Norwalk RHDV 96 NB

0.1

Fig. 3.1. Neighbor-Joining phylogenetic tree of the partial RdRp region (95 aa) of SaVs and RHDV, NB, Norwalk and FCV strains representing Lagovirus, NB-like virus, Norovirus and Vesivirus genera, respectively, in Caliciviridae. The 9 newly identified porcine SaV strains are in bold.

155 A. 63 Sapporo 58 Manchester Plymouth 100 GI-1 84Lyon30388 96 Houston86 Parkville 100 GI-2 100 Houston90 76 Stockholm 100 Mex14917 GI-3 63 Hou7 GIV Arg39 GV 100 C12 GII-4 100 Lyon598 100 100 Bristol GII-I London92 9 Cruise ship 8 53 GII-3 Mex340 82 GII-2 100 Mc10 JJ259 99 MM280 100 QW270 93 GIII LL14 99 44 48 Cowden 99 JJ681 GVI? NB 78 RHDV FCV NV

0.1 (Continued) Fig. 3.2. Neighbor-Joining phylogenetic trees of SaVs. The RHDV, NB, Norwalk and FCV strains represent Lagovirus, NB-like virus, Norovirus and Vesivirus genera, respectively, in Caliciviridae. A. Tree based on the predicted complete capsid amino acid sequences. B. Tree based on the partial RdRp region (C-terminal aligned 843 nt). The SaV genogroups (G plus roman numerals) and genotypes (arabic numbers following genogroup numbers) are indicated. The 4 newly sequenced porcine SaV strains are in bold.

156 B.

75 Lyon598 100 Bristol GII-1

100 London92 Hou7 GIV Mc10 99 Recombinant/parent 100 100 C12 Recombinant/parent

100 Houston90 GI-2 Parkville Manchester 100 100 Lyon30388 100 GI-1 Sapporo 86 87 Houston86 MM280 Recombinant

66 100 Cowden 100 LL14 GIII 100 JJ259 100 QW270 Recombinant JJ681 GVI? RHDV 43 71 NB FCV Norwalk

0.05

Fig. 3.2. Continued.

157 A.

3’-end RdRp capsid ORF2 3’-UTR thin Window i entity to d strains w d Cowden Nucleotide i JJ259 backgroun

Position of center of window (nucleotide)

(Continued)

Fig. 3.3. Identification of QW270 and MM280 strains as potential recombinants. Recombination Identification Program analysis (RIP) was performed based on the 3’-end 3 kb sequences including the partial RdRp, the capsid, the ORF2 and the 3’-UTR. At each position of the window, the query sequence was compared to the background sequence(s). When the query sequence is similar to the background sequence(s), the homologous regions are indicated as thick dots on the plot. Analysis parameters were Window size of 100 and Statistical significance of 90%. The nucleotide positions of the 3’-end RdRp, the predicted capsid, ORF2 and the 3’-end UTR are indicated. A. RIP of QW270 strain. The QW270 strain was a query sequence compared to the background Cowden and JJ259 strains. B. RIP of MM280 strain. The MM280 strain was a query sequence and only the Cowden strain was the background.

158 Nucleotide identity to B. Cowden strain within Window Position ofcenterw Fig. 3.3.Continued. 15 9 Cowden i ndow (n ucleotide) Genomic-subgenomic RNA conserved nucleotide motif

Cowden AAGAGCCAGA AGTGTTCGTG ATG GAGGCGCCTG CCCCAACCCG LL14 ********** ********** *** ********** *****G**** GIII QW270 ********** ********** *** *****A**** ********** JJ259 ********** ********** *** *****T**** ********** MM280 ****A*A*** ********** *** ********** **T******* Sapporo CCC*CAA*AT ******T*A* *** ****GCAA** G*T*C*A*TC Manchester CCC*CAA*AT ******T*A* *** ****GCAA** G*T*C*A**C GI-1 Lyon30388 CCC*CAA*AT ******T*A* *** ****GCAA** G*T*C*A**C Plymouth CCC*CAA*AT ******T*A* *** ****GCAA** G*T*C*A**C Houston86 CCC*CAA*AT ******T*A* *** ****GCAA** G*T*C*A**C Houston90 CACCCAA*TT ******T*A* *** ****GCAA** G*T*C*AATT GI-2 Parkville C*CCCAA*TT ******T*A* *** ****GCAA** G*T*C*AATT London92 CTACCAA*TT ******T*AA *** ****GC-T** G**AGC*A*A GII-I Lyon598 CTACCAA*TT ******T*AA *** ****GC-T** GG***G**AC Bristol CTACCAA*TT ******T*AA *** ****GC*TG* G**AGC*A*A GII-4 C12 CCACCAA*TT ******T*AA *** ****GTGTAC ***GCC*AGA GII-2 Mc10 CCACCAA*TT ******T*AA *** ****GC*TA* G**A*C*A*A GIV Hou7 CCACCAA*TT ******T*AA *** ****GCAA** G**T*C***A GVI? JJ681 GC*G*G*T*T T****A*ACA *** ****G***AA AG**CT*T**

Fig. 3.4. Sequence alignments of the RdRp-capsid junction region of SaVs. The predicted genomic and subgenomic conserved nucleotide motif of Cowden strain is indicated by a horizontal line with 2 vertical bars. Asterisks indicate the identical residues to the sequence of the first line (Cowden). The start codon for the capsid protein is underlined. Six SaV genogroups (GI-VI) are indicated. The 4 newly sequenced porcine SaV strains are in bold.

160

CHAPTER 4

A NEW MICROWELL HYBRIDIZATION ASSAY FOR THE DETECTION OF

PORCINE NOROVIRUSES AND SAPOVIRUSES BY REVERSE

TRANSCRIPTION-PCR

4.1 SUMMARY

Recently, genetically diverse porcine noroviruses (NoV) and sapoviruses (SaV) were identified from field pig fecal samples. Reverse transcription-PCR is the primary method used for detection of human NoVs and SaVs. However, RT-PCR inhibitors frequently cause false-negative results. In this study, a competitive internal control (IC) RNA to monitor for inhibition of RT-PCR was developed; primers for detection of genetically diverse porcine NoVs and SaVs were designed; and microwell hybridization assays to confirm the specific RT-PCR products were developed. The primer pairs and the RT-PCR-hybridization combinations were compared using representative porcine NoV and SaV strains, positive pig fecal samples and a panel of 30 field pig fecal samples. Extracted RNA from 3 of 30 samples failed to amplify the IC RNA. However, this inhibition was not present after

161 10-fold dilution of the extracted RNA and one of these 3 samples became positive for

NoV and SaV after dilution. The five different RT-PCR-hybridization combinations

developed specifically detected all 3 genotypes of porcine NoVs, all GIII porcine

SaVs, unclassified JJ681-like, QW19 and LL26-like porcine SaVs, respectively.

These RT-PCR-hybridization assays are specific, less time consuming and economical

and particularly applicable to testing large number of samples for porcine NoVs and

SaVs.

4.2 INTRODUCTION

Porcine noroviruses (NoV) and sapoviruses (SaV) are members of the family

Caliciviridae, comprising 2 genera: Norovirus and Sapovirus, respectively. The porcine NoVs replicate in pigs with NoV particles observed by immune electron microscopy (IEM) in intestinal contents of experimentally infected gnotobiotic pigs

(Wang et al., submitted), but their pathogenesis is undefined. The porcine SaVs cause diarrhea and intestinal lesions in pigs (4, 7). In addition, some porcine NoVs and

SaVs are genetically or antigenically related to human NoVs and SaVs (Wang et al., submitted), respectively, raising questions of whether pigs are reservoirs for emergence of human strains or if recombinant human/porcine NoVs and SaVs could emerge.

Recently, porcine NoVs and SaVs have been recognized as genetically diverse viruses. The porcine NoVs are classified genetically into 3 genotypes within genogroup II (GII) (14, 16). The porcine SaVs comprise at least 2 genogroups (5);

162 Wang et al., submitted). This genetic diversity increases the difficulty of detection of all strains of different genogroups using a single test.

Reverse transcription PCR (RT-PCR) has been used widely for the detection of human NoVs and SaVs from fecal samples (1). However, there are few reports of

RT-PCR detection of porcine NoVs and SaVs. Two primer pairs, PEC66/65 and

PEC46/45 based on the porcine SaV Cowden strain targeting two overlapping

RNA-dependent RNA polymerase (RdRp) regions have been used for detection of

Cowden SaV RNA from experimentally infected gnotobiotic pigs (7). A pair of calicivirus universal primers (p290 and p110 or p289) (9, 10) has been used widely to screen for genetically variable caliciviruses in humans and animals (3, 6). However, these primers have not been evaluated for detection of genetically variable NoVs and

SaVs from field fecal samples of pigs and new strain-specific primers are needed.

The sensitivity of RT-PCR may be much lower than expected because of the presence of RT-PCR inhibitors in a sample. Use of an internal amplification control in a RT-PCR or PCR assay is strongly recommended if not required (2, 8). Alternatively, an RT-PCR assay using template RNA directly extracted from fecal samples often produces non-specific amplifications which can result in false-positive results (1).

Sequencing of the RT-PCR products or performing hybridization assays is necessary to confirm the positive results with the latter being the simplest approach.

The objectives of this study were to produce a competitive internal control (IC)

RNA to monitor the presence of RT-PCR inhibitors, to design and compare primers for the detection of porcine NoVs and SaVs by RT-PCR and to develop microwell

163 hybridization assays to confirm the positive amplicons.

4.3 MATERIALS AND METHODS

Porcine NoV and SaV strains. The representative porcine NoV and SaV strains and positive fecal samples used in this study are summarized in Table 4.1.

There are 3 genotypes of porcine NoVs based on sequence analysis of the complete capsid and partial RdRp (~260 aa): GII-11 including the prototype porcine NoV

Sw43/97/JP, GII-18 (QW101-like) and GII-19 (QW170-like) (15). Cowden, QW270 and JJ259 are GIII-1 SaVs. The MM280 is a potential SaV GIII recombinant between an unknown SaV strain (RdRp region) and a Cowden-like SaV (capsid region). Its partial RdRp region (aligned 843 nt encoding 263 aa) shares 76% identity with the

Cowden strain and may represent another genotype (GIII-2?) in the RdRp region

(Wang et al., submitted). The JJ681, QW19 and LL26 may represent 3 different new genogroups of SaVs referred to as JJ681-like, QW19-like and LL26-like SaVs based on sequence analysis of the complete capsid and partial RdRp (255 aa) of JJ681 and the partial RdRp (95 aa) of QW19 and LL26 (Wang et al., submitted). Other positive field pig fecal samples used to assess the primers and probes included: (1) QW126,

QW133 and QW144, belonging to Po/NoV/GII-18; (2) II166, II175, II176 and JJ46, belonging to Po/SaV/GIII-1 and QW152 belonging to Po/SaV/GIII-2; (3) JJ672,

JJ674, JJ675 and JJ680, belonging to Po/SaV/JJ681-like and (4) LL27, LL28 and

LL31, belonging to Po/SaV/LL26-like. They were amplified by strain-specific primers (see below) and confirmed by sequencing of the RT-PCR products.

164 A panel of 30 fecal samples [nursing (1-3 weeks, n=2), post-weaning (3-10

weeks, n=2), finisher (10-24 weeks, n=21), sow (1-2 years, n=5)] collected from 6

swine farms [Michigan A (n=6), Ohio A (n=2), Ohio B (n=10), Ohio C (n=3), North

Carolina A (n=1), North Carolina B (n=2)] and one Ohio slaughterhouse (n=6) were

used to test the competitive IC RNA and the specific SaV and NoV

RT-PCR-hybridization combinations.

RNA extraction. Sample RNA was extracted from 200 µl of centrifuged

(1,200×g, 30 min) 10% fecal suspensions in sterile Eagle’s minimal essential medium

(EMEM, Invitrogen, Carlsbad, CA) by using the Trizol LS (Invitrogen) procedure.

The RNA pellet was resuspended in 40 µl RNase-free water and stored at -20 or

-70ºC.

A competitive IC RNA for the primer pairs PEC66/65 and PEC46/45. We

produced an IC RNA using a pair of composite primers to amplify PCR products,

which differ in size from our target viral amplicons (8). The forward primer Int1

(5’-GTG CTC TAT TGC CTG GAC TAC AGC AAG TGG GAT TCC ATA GTG

GAC TCT TGT TCC A-3’) contained the sequences of the primers PEC46 (bolded),

PEC66 (underlined) and the vector pCR2.1-TOPO nucleotides 841-859 (Invitrogen) in the 5’→3’ sequence (Fig. 4.1). The reverse primer Int2 (5’-ATA CAC ACA ATC

ATC CCC GTA TCT GTG GTG CGG TTA GCC TTA GAA AGC CAT CCA GTT

TAC-3’) contained the sequences of the primers PEC65 (bolded), PEC45 (underlined) and the vector pCR2.1-TOPO nucleotides 1250-1232 in the 5’→3’ sequence. The Int1 and Int2 were purified by High Performance Liquid Chromatography (Integrated

165 DNA Technologies, Coralville, IA). The vector pCR2.1-TOPO was amplified by PCR with these primers to produce a 487 bp DNA fragment. The 487 bp amplicons were purified by QIAquick Gel Extraction kit (Qiagen Inc. Valencia, CA) and then were cloned into the vector pCR2.1-TOPO. The direction of insertion was confirmed by

DNA sequencing. Thereafter, the circular recombinant plasmid was digested completely by the restriction enzyme Hind III and purified by phenol

(pH8.0)-chloroform extraction. Then the recombinant RNA was synthesized in vitro by T7 RNA polymerase using Riboprobe system-T7 (Promega, Madison, WI).

Plasmid DNA was removed completely by digestion with RNase-free DNase I for 15 min at 37˚C. Clearance of the plasmid DNA was confirmed by PCR without RT. The

IC RNA was then extracted using phenol (pH5.0)-chloroform. To avoid RNA degradation by trace amounts of RNase, RNase inhibitor RNasin (Promega) was added to the IC RNA. The concentration of the IC RNA was spectrophotometrically measured at 260 nm and the IC RNA aliquots were stored at –70˚C. The amount of IC

RNA used in RT-PCR with primer pair PEC66/65 or PEC46/45 was optimized by testing 10-fold serially diluted IC RNA. With primer pair PEC66/65, the RT-PCR product of IC RNA was 472 bp, whereas that of the SaV Cowden strain was 330 bp.

With primer pair PEC46/45, the IC RNA product was 466 bp whereas that of the SaV

Cowden strain was 572 bp.

Using this competitive IC RNA, there were three possible outcomes of

RT-PCR assays (8): (1) no product (the assay has failed-inhibition may be present); (2) the IC RNA, but not the virus fragment is obtained (the assay has worked, and the

166 sample is most likely negative or viral RNA below detection limit); (3) the virus

fragment, with or without the IC RNA fragment, is obtained (the assay has worked,

and the sample is positive).

Design of primers for detection of porcine NoVs and SaVs by RT-PCR.

Primers compared in this study and their RT and PCR annealing temperatures are summarized in Table 4.2. These primers target the RdRp region of porcine NoVs and

SaVs. The primer pairs PNV7/8 and PNV15/16 were designed based on conserved nucleotides of all 3 genogroups of porcine NoVs (sequence alignments performed by

ClustalW (version 1.83) at DDBJ, http://www.ddbj.nig.ac.jp) and should detect all 3 genogroups of porcine NoVs. The primer pairs PEC68/67, PSV6/7 and PSV11/14 were designed based on the JJ681, QW19 and LL26 sequence data using PrimerQuest software provided on line (http://www.idtdna.com) by Integrated DNA Technologies.

All primers were synthesized by Integrated DNA Technologies.

One-step RT-PCR assays were carried out using the NoV primer pair

PNV15/16 and the SaV primer pairs PEC66/65, PEC46/45, PEC68/67, PSV6/7 and

PSV11/14. Briefly, a final volume of 50 µl reaction mixture (46 µl of master mixture

plus 4 µl of sample RNA) contained 1 × PCR buffer (50 mM KCl, 10 mM Tris-HCl

[pH9.0], and 0.1% Triton X-100), 2.5 mM MgCl2, 200 µM of dNTPs, 0.5 µM of each primer, 10 U of RNase inhibitor RNasin (Promega, Madison, WI), 4 U of avian myeloblastosis virus reverse transcriptase (AMV RT, Promega), 2 U of Taq DNA polymerase (Promega). The RT reaction was conducted at 42-45ºC for 60 min followed by heat inactivation at 94ºC for 3 min. The PCR reaction was performed for

167 40 cycles with each cycle at 94ºC for 30 sec, 50-55ºC for 30 sec (PEC66/65,

PEC68/67, PSV6/7 and PSV11/14) or 1 min (PEC46/45 and PNV15/16), and 72ºC

for 30 sec; and a final extension for 10 min at 72ºC.

Two-step RT-PCR assays were performed using the primer pairs p290/110 and

PNV7/8 as previously reported with minor modifications (9). For RT, 25 µl of

reaction mixture (22 µl of master mixture plus 3µl of sample RNA) containing 1 ×

PCR buffer, 2.5 mM MgCl2, 400 µM of dNTPs, 1 µM of reverse primer p110 or

PNV8, 10 U RNasin and 2 U AMV RT was incubated at 42-45 ºC for 60 min

followed by heat inactivation at 94ºC for 3 min then chilling to 4ºC. Afterwards,

another 25 µl of reaction mixture containing 1 X PCR buffer, 2.5 mM MgCl2, 1 µM of forward primer p290 or PNV7, and 1 U Taq DNA polymerase was added into the above RT mixture to a final volume of 50µl to perform PCR reaction for 40 cycles with each cycle at 94ºC for 30 sec, 48 (for p290/110) or 50ºC (for PNV7/8) for 30 sec and 72ºC for 30 sec, and a final extension for 10 min at 72ºC.

The RT-PCR products were analyzed by agarose gel electrophoresis stained with ethidium bromide and visualized using ultraviolet light and microwell hybridization assays (see below).

Microwell hybridization assays for confirmation of the positive RT-PCR products. The 5’-end of the forward NoV primer PNV7 and SaV primers PEC66,

PEC68, PSV6, and PSV11 was Biotin-labeled (Integrated DNA Technologies) and referred to as PNV7-Bio, PEC66-Bio, PEC68-Bio, PSV6-Bio, and PSV11-Bio, respectively, in this manuscript. The amplicons of the primer pairs PNV7-Bio/8,

168 PEC66-Bio/65, PEC68-Bio/67, PSV6-Bio/7 and PSV11-Bio/14 were detected by a

microwell hybridization assay modified from that described previously for detection of Brucella abortus and/or Mycobacterium bovis (13). The internal DNA probes were

designed using PrimerQuest software to hybridize to the forward strands of the

corresponding RT-PCR products (Table 4.3). The probes PoNoroP1A, PoNoroP1B

and PoNoroP1C targeted the porcine NoV GII-11 (QW48-like), GII-18 (QW101-like)

and GII-19 (QW170-like), respectively. Probes PoSapoP1A, PoSapoP1B, PoSapoP1C,

PEC-P1 and PEC-P2 targeted the porcine GIII-1 SaV, MM280, QW19, JJ681 and

LL26, respectively. They were synthesized by Integrated DNA Technologies.

The EIA/RIA 8-well strips (Corning Inc., Corning, NY) were coated with 100

ng/well of each probe in 1 M ammonium acetate solution (pH 7.5) at 37ºC overnight.

After washing, the wells were blocked with 1% BSA in PBS-Tween 20 (PBS-T,

0.05%) at 37ºC for 2 hours. Then the plates were used immediately or were air-dried

and stored at 4ºC for up to 3 months with the presence of dehumidizer. The RT-PCR

products were denatured by adding an equal volume of denaturing buffer (0.4 M

NaOH, 80 mM disodium EDTA, 0.005% thymol blue). Next 25µl of the denaturing

solution containing 12.5µl of RT-PCR product was added to duplicate wells

containing 100µl of neutralization-hybridization buffer (1 M sodium thiocyanate, 80

mM NaH2PO4; 10mM Na2HPO4, pH5.0 ±0.2). After incubating at 37ºC for 1 h, the

wells were washed with the PBS-T. Then 100 µl of 0.4 µg/ml NeutravidinTM

Horseradish Peroxidase conjugate (Pierce Biotechnology, Rockford, IL) in 1% BSA

buffer was added to each well and incubated at 37ºC for 15 min. The wells were

169 washed, the substrate tetramethylbenzidine (KPL, Gaithersburg, MD) was added and

strips were incubated in the dark for 10 min. The reaction was stopped by adding 100

µl of 1.8 % HCl. After 5 min equilibration, the absorbance (A) was measured by a spectrophotometer at 450 nm. For each hybridization assay, RT-PCR positive and negative controls and water as a blank were included. The cut-off value for each primer pair-probe combination was determined based on the highest A450 of the

non-specific amplicons multiplied by 2.

Sequencing and sequence analysis. The RT-PCR products were purified by

QIAquick Gel Extraction kit (Qiagen Inc.). The purified DNA was sequenced directly using forward and reverse primers or after cloning into pCR2.1-TOPO (T/A) vector

(Invitrogen). The DNA sequencing was done using BigDye Terminator Cycle chemistry and 3700 or 3730 DNA Analyzer (Applied Biosystems, Foster, CA).

Sequence editing was performed by Lasergene software package (version 5)

(DNASTAR Inc., Madison, WI). Basic Local Alignment Search Tool (BLAST,

http://www.ncbi.nlm.nih.gov) was used to find homologous hits.

4.4 RESULTS

Design and application of the IC RNA. A single-stranded (SS) IC RNA for

RT-PCR including the sequences of the SaV forward primers PEC66, PEC46 and the reverse complementary sequences of the SaV reverse primers PEC65 and PEC45 was produced by in vitro transcription (Fig. 4.1). It was 632 nt long from the T7 promoter to the restriction enzyme Hind III site. Based on the molecular weight of a

170 1kb-ssRNA of about 3.4 × 105 Dalton, the estimated molecular weight of the IC RNA

5 is about 2.16 × 10 Dalton. The 100-fold diluted IC RNA had an A260 of 0.21.

Therefore the concentration of the undiluted IC RNA was 0.084 µg/µl, equal to 2.34

× 1011 molecules/µl.

To determine how much of the IC RNA to use in a 50 µl RT-PCR with the primer pair PEC66/65, 1 µl of 10-fold serially diluted IC RNA was tested. We amplified a 472 bp-band with the 10-2 to 10-6 dilutions, but no visible band was observed at the 10-7 dilution by agarose gel electrophoresis. We co-amplified this 10-6

dilution of IC RNA with the 5-fold serially diluted viral RNA (50 to 56-fold) of the

SaV Cowden strain, representing from high to low amounts of viral RNA in the fecal specimens (Fig. 4.2). The Cowden-specific band (330 bp) was readily differentiated from that of the IC RNA band (472 bp) by agarose gel electrophoresis. No IC RNA band was seen when the viral RNA was tested undiluted, but both the IC RNA and the

Cowden-specific bands appeared when the viral RNA level was moderate (5 to

125-fold dilution), and only the IC RNA band was evident when the viral RNA level was low (≥ 625-fold dilution). We also amplified each dilution of the viral RNA without the IC RNA to investigate whether the IC RNA interferes with amplification of viral RNA. We found that with or without the IC RNA, the highest dilution of the viral RNA for a visible band was 1:125, and the intensity of the bands at each dilution was similar. These results indicated that 1 µl of 106 diluted IC RNA did not interfere

with the amplification of diluted viral RNA at high to low levels.

We also tested the IC RNA co-amplified with the primer pair PEC46/45 and

171 we found that 2 µl of the IC RNA were needed for a reproducible visible band (data not shown) by agarose gel electrophoresis. The Cowden-specific band (572 bp) and

the IC RNA-specific band (466 bp) were also well differentiated.

We tested a panel of 30 field pig fecal samples and found that 3 (10%) showed

RT-PCR inhibitors. After 10-fold dilution of the extracted RNA, we performed

RT-PCR again and found that the inhibitors no longer interfered with RT-PCR of any

of these 3 samples and one of the 3 samples became porcine NoV and SaV positive.

Comparison of the primer pairs used for detection of porcine NoVs.

Primer pairs PNV7/8 and PNV15/16 produced only weak bands for the porcine

NoV-positive samples when one-step RT-PCR was performed. Next we tried two-step

RT-PCR for these two primer pairs and we found that the intensity of PNV7/8

amplicons was increased and PNV7/8 detected all 3 genogroups of NoVs, whereas

NoV amplification with PNV15/16 was not greatly improved. Therefore, we

performed two-step RT-PCR using PNV7/8 and one-step RT-PCR using PNV15/16.

The PNV7/8 detected all 8 porcine NoVs tested, whereas PNV15/16 detected

only GII-18, QW101-like NoVs (QW101, QW125, QW126, QW133 and QW144)

and p290/110 did not detect 2 of the QW101-like NoVs (QW133 and QW144). Next

we compared the detection limit of primer pairs PNV7/8 and p290/110 by using

serially diluted QW101 RNA (5, 50, 500, 2,500, 12,500, 62,500-fold) as template. We

found that PNV7/8 amplified a visible band by agarose gel electrophoresis up to a

12,500-fold dilution of viral RNA whereas the p290/110 primers detected viral RNA

up to only a 50-fold dilution. The primer pair PNV7/8 was 250-fold more sensitive

172 than the p290/110 primer pair for detection of QW101.

After the 30 pig fecal samples were tested using primer pair PNV7/8, we

sequenced representative RT-PCR products to examine the specificity. We found that

non-specific amplicons with a similar size to that of porcine NoVs (211 bp) included

the following: uncultured Cytophagales MB11E04 16S ribosomal RNA gene [blastn,

E value = 9e-24, Identities = 124/145 (85%)] and a sequence that had no significant hit in the database (blastn and blastx) (Table 4.7).

Comparison of the primer pairs used for detection of porcine SaVs. We

compared primer pairs PEC66/65, PEC46/45, PEC68/67, PSV6/7, PSV11/14 and

p290/110 for the detection of different genogroups of SaVs (Table 4.4). We found that

the PEC66/65 detected all GIII and weakly detected 3 of 4 LL26-like porcine SaVs,

but not the JJ681-like and QW19 porcine SaVs. The PEC46/45 detected only 4 of 9

GIII porcine SaVs. The PEC68/67, PSV6/7 and PSV11/14 exclusively detected

JJ681-like, QW19, and LL26-like strains, respectively. Thus primer pairs PEC66/65,

PEC68/67, PSV6/7 and PSV11/14 were the best for detection of GIII, JJ681-like,

QW19-like and LL26-like porcine SaVs, respectively.

The specificity of primer pairs PEC66/65, PEC68/67, PSV6/7 and PSV11/14

was tested by sequencing of representative RT-PCR products of the 30 pig fecal

samples followed by BLAST search for significant hits in the database. One

non-specific amplicon of primer pair PEC66/65 with a similar size to that of porcine

SaVs (330 bp) had a significant hit [blastn, E value = 3e-08, identity = 66/77 (85%)]

to Treponema denticola ATCC 35405, section 5 of 10 of the complete genome (Table

173 4.7). The primer pair PSV6/7 amplified a non-specific band with a similar size to that of porcine QW19-like SaVs (219 bp), which was Oceanobacillus iheyensis HTE831

DNA [blastn, E value = 1e-35, identity = 124/139 (89%)]. We did not find non-specific bands with a similar size to the SaV-specific products for primer pairs

PEC68/67 (219 bp) and PSV11/14 (231 bp), respectively.

Confirmation of porcine SaV- and NoV-specific amplicons by microwell hybridization assays. Biotin-labeling of the forward primers did not change the corresponding RT-PCR efficiency tested by using serially diluted positive RNA as template. For detection of porcine NoVs, we tested probes PoNoroP1A, PoNoroP1B and PoNoroP1C separately with the PNV7-Bio/8 RT-PCR products of positive samples (Table 4.5). We found that every probe detected all genogroups of porcine

NoVs. However, the PoNoroP1A, PoNoroP1B and PoNoroP1C showed higher absorbance for QW48, QW101-like and QW170-like strains, respectively. Then we mixed the 3 probes together for coating to perform the hybridization assay. For the

QW48, QW101-like and QW170-like strains, the absorbance was similar to that for the hybridization with a single probe. This RT-PCR PNV7-Bio/8-hybridization

(PoNoroP1A+ PoNoroP1B+PoNoroP1C) did not detect porcine SaVs.

Similarly, we tested the SaV probes for detection of SaV amplicons with different pairs of primers (Table 4.6). The PoSapoP1A and PoSapoP1B detected the

PEC66-Bio/65 products of all GIII SaVs and also detected weakly LL26-like SaVs.

When we mixed these 2 probes together for a single hybridization, they also weakly detected JJ681-like SaVs. The RT-PCR PSV6-Bio/7-hybridization (PoSapoP1C),

174 RT-PCR PEC68-Bio/67-hybridization (PEC-P1) and RT-PCR

PSV11-Bio/14-hybridization (PEC-P2) detected exclusively QW19, JJ681-like and

LL26-like porcine SaVs, respectively. None of the porcine SaV RT-PCR and hybridization combinations detected porcine NoVs.

The specificity of the combination of each primer pair and probe in the

RT-PCR-hybridization assay was examined in two ways (Table 4.7). First, we tested

the non-specific bands with similar sizes to the virus-specific products. Second, we

examined the non-specific bands that had different sizes to the virus-specific products because these non-specific products could be differentiated on agarose gel electrophoresis, but they may cause non-specific binding to the probes in the microwell hybridization assays. Based on these results, we determined the cut-off value as 2 × A450 of the highest value of the non-specific products, which was 0.30 for

RT-PCR PEC66-Bio/65-hybridization (PoSapoP1A+PoSapoP1B) and was 0.16-0.20 for the other RT-PCR–hybridization assays. Consequently the cut-off value used was

0.30 for RT-PCR PEC66-Bio/65-hybridization (PoSapoP1A+PoSapoP1B) and was

0.20 for the other RT-PCR–hybridization assays (Table 4.7).

The sensitivity of the hybridization assays for detection of RT-PCR products was compared to that of agarose gel electrophoresis by using serially diluted RT-PCR

products of porcine NoV and SaV representative strains. We found that the

hybridization assay had similar or higher sensitivity (1-8 fold, Table 4.7) than agarose

gel electrophoresis depending on the probes and the cut-off value of each

hybridization assay. For example, in the hybridization assay detecting PEC68-Bio/67

175 RT-PCR products with probe PEC-P1 (Fig. 4.3), the highest dilution (1:128) of the

RT-PCR products showing a visible band in the agarose gel produced an A450 of 1.40, whereas the microwell hybridization results were positive (> cut-off value 0.20) through the dilution of 1:1024. This means that the sensitivity of the microwell hybridization assay was increased 8-fold compared to agarose gel electrophoresis for the detection of RT-PCR PEC68-Bio/67 products. The increased sensitivity of other

RT-PCR-hybridization combinations is summarized in Table 4.7.

4.5. DISCUSSION

The RT-PCR inhibitors are common and sample-dependent in field fecal samples (1, 2) causing false-negative results. They may include pig body fluids, food and bacterial cell constituents (17). An IC RNA co-amplified with the target viral

RNA can be used to monitor for RT-PCR inhibition as well as technical errors.

Diluting the extracted RNA of such samples, further purifying the extracted RNA or using alternative RNA extraction methods and performing RT-PCR again usually decreases the false negative results. Diluting the extracted RNA is simplest and most samples diluted 1:10 or 1:50 will no longer inhibit RT-PCR (2). A drawback of this strategy is that the dilution of weakly positive samples may exceed the detection limit of RT-PCR and therefore become negative.

In this study, we produced a competitive IC RNA originally for two pair of primers, PEC66/65 and PEC46/45. We found that twice the amount of IC RNA was needed for PEC46/45 compared to PEC66/65 to produce a reproducible visible band

176 by agarose gel electrophoresis. This means that the RT-PCR efficiency is lower with

primer pair PEC46/45 than with PEC66/65. Later we also found that the PEC46/45

was less sensitive than the PEC66/65 for detection of GIII SaVs. Thus we choose the

primer pair PEC66/65 to amplify GIII SaV RNA and the IC RNA simultaneously.

When we used the IC RNA at a minimum level (1µl of 10-6 dilution) for a reproducible visible band with the primer pair PEC66/65, no obvious interference with the SaV RNA amplification was observed (Fig. 4.2). This is probably due to the smaller size of the viral RNA amplicon (330 bp) compared to that of the IC RNA amplicon (472 bp) because the PCR amplifies smaller products more efficiently (8).

The RT-PCR inhibitors mainly interfere with activities of the reverse transcriptase and

DNA polymerase which catalyze the RT and PCR reactions, respectively. When a sample tests RT-PCR inhibitor positive by primer pair PEC66/65 with the IC RNA, this sample likely also contains RT-PCR inhibitors for other primer pairs. This sample

RNA should then be diluted or purified until it is no longer inhibited in the RT-PCR with PEC66/65 and the IC RNA before performing RT-PCR with other primers.

When RT-PCR with primer pair PEC66/65 is not used for a study, a new IC RNA can be produced using our approach for other primer pairs. We tested 30 fecal samples and found that 3 samples contained RT-PCR inhibitors. After the RNA was diluted

1:10, the RT-PCR was no longer inhibited for all 3 samples and one of them became

NoV and SaV positive.

From a previous review (8) and our experience, there are several ways to

avoid IC RNA degradation: storing the IC RNA at high concentration with RNase

177 inhibitors at -70˚C and storing in aliquots to avoid repeated freezing and thawing. The working dilution of IC RNA should be defined after storage. Also, the IC RNA should be added in the master mixture, instead of being added to each tube for the RT-PCR because the IC RNA molecules have a Poisson’s distribution (12). Finally, the IC

RNA amplicons can be detected by either agarose gel electrophoresis or microwell hybridization. However, for the latter, a cut-off value should be determined without ignoring weak inhibition.

Although the RT-PCR primers used for detection of the genetically diverse

NoVs and SaVs are usually designed based on the most conserved RdRp region,

several primer pairs are usually necessary to detect the genetically variable strains (1).

In this study, we designed 5 pairs of primers (PNV7/8 and PNV15/16 for porcine

NoVs and PEC68/67, PSV6/7 and PSV11/14 for porcine SaVs) targeting the RdRp

region. Comparison of the detection spectrum and detection limits of these new

primer pairs and the previously reported 3 primer pairs (p290/110, PEC66/65 and

PEC46/45) revealed that p290/110 is the most broadly reactive primer pair for

detecting genetically variable porcine NoVs and SaVs, but its sensitivity is 250-fold

lower for the QW101 strain than the NoV-specific primer pair PNV7/8. In fact, most

porcine NoV and SaV strains were identified in our lab by performing RT-PCR with

the p290/110 followed by sequencing of the amplicons (Wang and Saif, submitted).

Therefore, RT-PCR with this primer pair combined with sequencing of the amplicons

remains a good way to screen for genetically new caliciviruses, but is likely to miss

the porcine NoVs and SaVs at lower RNA amounts in the samples, especially

178 important for screening subclinically infected animals. The detection spectrum of primer pair PNV7/8 is greater than that of primer pair PNV15/16 for detection of all three genogroups of porcine NoVs. For detection of porcine GIII SaVs, the primer pair PEC66/65 was more sensitive than PEC46/45. The PEC68/67, PSV6/7 and

PSV11/14 were more restricted and detected only the JJ681-like, QW19-like and

LL26-like porcine SaVs, respectively.

However, after sequencing the amplicons of selected samples, we found that primer pairs p290/110, PNV7/8, PEC66/65 and PSV6/7 amplified non-specific products of a similar size to those of the virus-specific products. These included bacterial genes and sequences with no known homology, likely because fecal samples contain mixtures including sloughed epithelial cells, undigested food components and the normal intestinal bacterial flora. The RT-PCR followed by hybridization increases the specificity by two step-specific selections: primer- and probe-selection. The

RT-PCR PNV7-Bio/8-hybridization (PoNoroP1A+PoNoroP1B+PoNoroP1C) detected all 3 genogroups of porcine NoVs. The RT-PCR PEC66-Bio/65-hybridization

(PoSapoP1A+PoSapoP1B) detected all GIII SaVs and weakly detected JJ681-like and

LL26-like SaVs. The RT-PCR PEC68-Bio/67-hybridization (PEC-P1), RT-PCR

PSV6-Bio/7-hybridization (PoSapoP1C) and RT-PCR PSV11-Bio/14-hybridization

(PEC-P2) only detected JJ681-like, QW19-like and LL26-like porcine SaVs, respectively. Compared to agarose gel electrophoresis, the microwell hybridization assay is similar or more sensitive (2-8 fold), depending on the probes and the cut-off value for each assay. The assay is also quick (2 h for one plate detecting 48 RT-PCR

179 products and controls) and economical (about $10/plate). Compared to a previously

reported microwell hybridization assay using biotinylated primers and

digoxigenin-labeled probes (11), our approach is simpler. Compared to sequencing of

the amplicons, it is less time consuming and cheaper to confirm the specificity of the

amplicons, especially in epidemiological studies testing large number of samples.

Although under conditions of low stringency (e.g., incubation at 37ºC), internal

probes can hybridize with amplicons with several nucleotide mismatches, strains with

highly variable internal sequences may be missed. Consequently, in some

circumstances sequencing of the amplicons from the calicivirus universal primer pair

p290/110 may be a better strategy than performing hybridization with a single probe,

which may miss the genetically highly variable or new calicivirus strains.

Alternatively, hybridizations with an array of strain-specific probes may also be

applied for efficient identification and differentiation.

In summary, we generated an IC RNA for SaV primers to monitor the

existence of RT-PCR inhibitors in the fecal samples to decrease false negative results.

New primer pairs were designed and compared with published primers for detection

of porcine NoVs and SaVs. We also developed microwell hybridization assays for

specific detection and confirmation of porcine NoV and SaV amplicons. These

RT-PCR-hybridization assays can be used for future epidemiological studies of

porcine NoVs and SaVs. The microwell hybridization assays, with appropriate modifications for other NoV and SaV genogroups/genotypes, should be applicable for

widespread use for detection of NoVs and SaVs.

180

4.6 ACKNOWLEDGMENTS

This work was supported by grants from the National Institute of Allergy and

Infectious Diseases, National Institutes of Health (Grant R01 AI 49742) and NRI, US

Department of Agriculture (CGP Grant 1999 02009) and by the Ohio Agricultural

Research and Development Center (OARDC), The Ohio State University (Graduate

Student Research Enhancement Grant project 2002-114). Salaries and research support were provided by state and federal funds provided to the OARDC.

We thank Dr. Norma Ramirez and Mr. Mike Kauffman for technical assistance.

We thank Dr. Jeffrey LeJeune and Dr. Ken Theil for critical review of the manuscript.

We also thank the Plant-Microbe Genomics Facility of The Ohio State University for

DNA sequencing.

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15. Sugieda, M., and S. Nakajima. 2002. Viruses detected in the caecum contents of healthy pigs representing a new genetic cluster in genogroup II of the genus "Norwalk-like viruses". Virus Res 87:165-72.

16. Van Der Poel, W. H., J. Vinje, R. van Der Heide, M. I. Herrera, A. Vivo, and M. P. Koopmans. 2000. Norwalk-like calicivirus genes in farm animals. Emerg Infect Dis 6:36-41.

17. Wilson, I. G. 1997. Inhibition and facilitation of nucleic acid amplification. Appl Environ Microbiol 63:3741-51.

183 Representative strain Genbank Positive fecal sample (Genus/Genogroup-genotype) Abbreviation accession # NoV/GII-11/MI-QW48/02/US QW48 AY823303 NAb

NoV/GII-18/OH-QW101/03/US QW101 AY823304 QW126, QW133, QW144 NoV/GII-18/OH-QW125/03/US QW125 AY823305

NoV/GII-19/OH-QW170/03/US QW170 AY823306 NAb NoV/GII-19/OH-QW218/03/US QW218 AY823307

SaV/GIII-1/Cowden/80/US Cowden AF182760 II166, II175, II176, JJ46, 184 SaV/GIII-1/NC-QW270/03/US QW270 AY826426 SaV/GIII-1/OH-JJ259/00/US JJ259 AY826423

SaV/GIII-2?/MM280/03/USa MM280 AY823308 QW152

SaV/GVI?/OH-JJ681/00/US JJ681 AY974192 JJ672, JJ674, JJ675, JJ680

SaV/G?/MI-QW19/02/US QW19 AY826424 NAb

SaV/G?/OH-LL26/02/US LL26 AY974195 LL27, LL28, LL31 a: The classification (GIII-2?) based on the partial RdRp region (aligned 843 nt encoding 263 aa). b: NA = not available Table 4.1 Representative porcine NoV and SaV strains and positive fecal samples used in this study.

Primer Target Target RdRp ProductTm RT Annealing pair Sense Sequence (5'→3') virus(es) (genome position) size (bp)(℃) T (℃) T (℃) Reference a 4568-4590 , p290 +GAT TAC TCC AAG TGG GAC TCC AC Caliciviruses 4327-4349b NoVs: 317 58 42 48 Jiang et al., 1999 a 4884-4865 , Le Guyader et al., b p110 - ACD ATY TCA TCA TCA CCA TA Caliciviruses4655-4636 SaVs: 32949 1996

PNV7 + AGG TGG TGG CCG AGG AYC TCC T NoV/GII-11, 18, 19 4422-4443 a 211 66 45 50 This study PNV8 - TCA CCA TAG AAG GAR AAG CA NoV/GII-11, 18, 19 4632-4613a 56 This study

a PNV15 + TCC TTY TAT GGT GAT GAT GA NoV/GII-11, 18, 19 4619-4638 416 49 45 50 This study PNV16 - CTG GGC ACG TAA AAC TCC A NoV/GII-11, 18, 19 5034-5016a 55 This study

185 b PEC66 + GAC TAC AGC AAG TGG GAT TCC SaV/GIII 4327-4347 330 55 45 50 Guo et al., 2001 PEC65 - ATA CAC ACA ATC ATC CCC GTA SaV/GIII 4656-4636b 55 Guo et al., 2001

b PEC46 + GTG CTC TAT TGC CTG GAC TA SaV/GIII 4312-4331 572 54 42 50 Guo et al., 2001 PEC45 - TCT GTG GTG CGG TTA GCC TT SaV/GIII 4883-4864b 63 Guo et al., 2001

b PEC68 + CCG CTA TAA ATT TAT TGG GTG SaV/GVI?/JJ681 4374-4394 225 50 45 50 This study PEC67 - ACG GGA CCC CAT ATT TTT GG SaV/GVI?/JJ681 4598-4579b 56 This study

b PSV6 + CGG TCA TTT TGT GTG GAC TG SaV/G?/QW19 4391-4410 219 54 45 50 This study PSV7 - ATT GCC CGT ATA AGG CAC A SaV/G?/QW19 4609-4591b 54 This study

b PSV11 + CAC CCA GAG GTG ATT TCA ACAC SaV/G?/LL26G 4355-4378 231 59 45 55 This study PSV14 - TTC TGC GTA ACA CTG GAG CAC CASaV/G?/LL26 4585-4562 b 63 This study a Based on Hu/NoV/GI-1/Norwalk/68/US (Genebank accession #: M87661). b Based on Po/SaV/GIII/Cowden/80/US (Genebank accession # AF182760). Table 4.2 RT-PCR primers used for the detection of porcine NoVs and SaVs.

Probe Sequence (5'-3') For primer pair(s) Target virus(es) AGC CAG TGG GCG AAG GAG PoNoroP1A TTC CAC TGT GAT GTG CA PNV7/8 NoV/GII-11/MI-QW48

AGC CAA TGC GCT ATG GAG NoV/GII-18/OH-QW101 PoNoroP1B TTC CAC TGT GAT GTG CA PNV7/8 & NoV/GII-18/OH-QW125

AGC CAA TGG GCA AAG GAA NoV/GII-19/OH-QW170 PoNoroP1C TTC CAT TGT GAT GTG CA PNV7/8 & NoV/GII-19/OH-QW218

SaV/GIII/Cowden, SaV/GIII/LL14 186 ATR ACA CTG GTG AAG GGC PEC66/65, SaV/GIII/NC-QW270, PoSapoP1A ATG CCA GAG GGG AG PEC46/45 SaV/GIII/JJ259

ATC ACA CTG GTG AAG GGC PEC66/65, PoSapoP1B ATG CCT GAT GGT AA PEC46/45 SaV/GIII/MM280

ATG ACG CTG GTG AAT GGC PoSapoP1C ATG CCA GAG GGA AG PSV6/7 SaV/G?/MI-QW19

AAC TGA GTT GAT CAC ACT PEC-P1 GGT GCC TGG CAT TCC AGA PEC68/67 SaV/GVI?/OH-JJ681

GTA AGC CCT TCT GTG TGG PEC-P2 GCA ATG AGA TGT CAA ACA PSV11/14 SaV/G?/OH-LL26 Table 4.3 Hybridization DNA probes for the detection of porcine NoV- and SaV-specific amplicons.

Classification Strain p290/110 PEC66/65 PEC46/45 PEC68/67 PSV6/7 PSV11/14 GIII Cowden + + + - - - II166 + + - - - - II175 - + + - - - II176 + + + - - - JJ46 - + - - - - JJ259 + + + - - - QW270 + + - - - - QW152 + + - - - - MM280 + + - - - - 187 GVI?, JJ681-like JJ681 + - - + - - JJ672 - - - + - - JJ674 + - NT + - - JJ675 - - NT + - - JJ680 - - NT + - - QW19-like QW19 + - - - + - LL26-like LL26 + + - - - + LL27 + - - - - + LL28 + + - - - + LL31 - + - - - + a:"+" = positive, "-" = negative, NT = not tested. Table 4.4 Comparison of primer pairs p290/110, PEC66/65, PEC46/45, PEC68/67, PSV6/7 and PSV11/14 for the

detection of porcine SaVsa.

Strain PoNoroP1A PoNoroP1B PoNoroP1C Mix of 3 probes Po/NoV/GII-11/MI-QW48 ++++ +++ ++++ ++++ Po/NoV/GII-18/OH-QW101 +++ ++++ +++ ++++ Po/NoV/GII-18/OH-QW125 +++ ++++ +++ ++++ Po/NoV/GII-19/OH-QW170 +++ +++ ++++ ++++ Po/NoV/GII-19/OH-QW218 +++ +++ ++++ ++++

188 Po/SaV/GIII/Cowden NT NT NT - Po/SaV/GIII/MM280 NT NT NT - SaV/G?/MI-QW19 NT NT NT - SaV/GVI?/OH-JJ681 NT NT NT - SaV/G?/OH-LL26 NT NT NT -

a: "++++" = A450 ≥ 3.00, "+++" = 3.0 > A450 ≥ 2.00, NT = not tested, "-" = negative.

Table 4.5 Detection spectrum of the NoV probes for PNV7-Bio/8 RT-PCR productsa.

PEC66-Bio/65- PEC66-Bio/65- PEC66-Bio/65- PSV6-Bio/7- PEC68-Bio/67- PSV11-Bio/14- Strain PoSapoP1A PoSapoP1B PoSapoP1A+PoSapoP1B PoSapoP1C PEC-P1 PEC-P2 SaV/GIII/Cowden ++++ +++ ++++ - - - SaV/GIII/NC-QW270 ++++ ++ ++++ - - - SaV/GIII/OH-JJ259 ++++ ++++ ++++ - - -

SaV/GIII/MM280 ++ ++++ ++++ - - - SaV/GIII/OH-QW152 +++ ++++ ++++ - - -

SaV/G?/MI-QW19 - - - ++++ - -

189 SaV/GVI?/OH-JJ681 - - + - ++++ -

SaV/G?/OH-LL26 ++ + ++ - - ++++

Po/NoV/GII-11/MI-QW48 NT NT - --- Po/NoV/GII-18/OH-QW101 NT NT - --- Po/NoV/GII-19/OH-QW170 NT NT - - - -

a: "++++" = A 450 ≥ 3.00, "+++" = 3.0 > A 450 ≥ 2.00, "++" = 2.0 > A 450 ≥ 1.00, "+" = d: 1.0 > A 450 ≥ cut-off value, NT = not tested, "-" = negativ

Table 4.6 Detection spectrum of the SaV probes for the RT-PCR products of several primer pairs (primer pair-probe)a

Positive, negative and non-specific RT-PCR products Hybridization Cut-off Increased Primer pair Hybridization probe Target virus d A 450 value A 450 sensitivity Positive 211 bp 3.80 0.20 2-fold PNV7-Bio/8 Negative no 0.05 Similar sizea Bacteria ribosomal RNA, unknown se quencesc PoNoroP1A+PoNoroP1B Po/NoV/GII 0.06-0.09 b +PoNoroP1C -11,18,19 Different size 100 bp, 250 bp, 350 bp,450 bp, 0.06-0.10 Positive 330 bp 4.00 0.30 similar Negative no 0.05 PEC66-Bio/65 Similar sizea Bacteria DNA PoSapoP1A+PoSapoP1B 0.05 Different sizeb 100 bp, 200 bp, 300bp, 400 bp, 600bpPo/SaV/GIII 0.05-0.15 IC RNA 472 bp 0.05 Positive 225 bp 4.00 0.20 8-fold PEC68-Bio/67 Negative no 0.05 a Similar size no PEC-P1 Po/SaV/ 190 Different sizeb 150 bp, 350 bp, 420 bp, 690 bpJJ681-like 0.05-0.09 Positive 219 bp 4.00 0.20 4-fold Negative no 0.05 PSV6-Bio/7 Similar sizea Bacterial DNA PoSapoP1CPo/SaV/ 0.06 Different sizeb 150bp, 350 bpQW19-like 0.05-0.08 Positive 231 bp 4.00 0.20 Negative no 0.05 PSV11-Bio/14 Similar sizea no PEC-P2Po/SaV/ 4-fold Different sizeb 150 bp, 250 bpLL26-like 0.05-0.08 a: Non-specific RT-PCR products with a similar size to the virus-specific products detected by agarose gel electrophoresis. b: Non-specific RT-PCR products with a different size to the virus-specific products detected by agarose gel electrophoresis. c: These sequences had no significant hit in the database by BLAST (blastn and blastx) search. d: The hybridization increased sensitivity for detection of RT-PCR products compared to agarose gel electrophoresis.

Table 4.7 The absorbance of porcine NoV and SaV positive, negative and non-specific amplicons in each RT-PCR-hybridization assay.

Nucleotide sequence of the IC RNA (632 nt) 1 taatacgact cactataggg cgaattgggc cctctagatg catgctcgag cggccgccag tgtgatggat atctgcagaa 81 ttcgcccttg tgctctattg cctggactac agcaagtggg attccatagt ggactcttgt tccaaactgg aacaacactc 161 aaccctatct cggtctattc ttttgattta taagggattt tgccgatttc ggcctattgg ttaaaaaatg agctgattta 241 acaaaaattt aacgcgaatt ttaacaaaat tcagggcgca agggctgcta aaggaagcgg aacacgtaga aagccagtcc 321 gcagaaacgg tgctgacccc ggatgaatgt cagctactgg gctatctgga caagggaaaa cgcaagcgca aagagaaagc 401 aggtagcttg cagtgggctt acatggcgat agctagactg ggcggtttta tggacagcaa gcgaaccgga attgccagct 481 ggggcgccct ctggtaaggt tgggaagccc tgcaaagtaa actggatggc tttctaaggc taaccgcacc acagatacgg 561 ggatgattgt gtgtataagg gcgaattcca gcacactggc ggccgttact agtggatccg agctcggtac ca

1 200 400 600 nucleotide 191

Int1 Int2 PEC46 PEC45 PEC66 PEC65

pCR2.1-TOPO nucleotides 841-1250

Fig. 4.1 The complete RNA sequence of the internal control RNA (632 nt in length) for primer pairs PEC66/65 and PEC46/45. It starts with the T7 promoter (nucleotides 1-20) and ends before the restriction enzyme Hind III cutting site (right after nucleotide 632). The primer Int1 (nucleotides 90-144) includes the sequences of the porcine SaV Cowden-specific primers PEC46 (nucleotides 90-109) and PEC66 (nucleotides 105-125) and the pCR2.1-TOPO vector nucleotides 841-859 (nucleotides 126-144). The reverse complementary sequence of the primer Int2 (nucleotides 517-576) includes the reverse complementary sequences of the pCR2.1-TOPO vector nucleotides 1250-1232 (nucleotides 517-535), the porcine SaV Cowden-specific primers PEC45 (nucleotides 536- 555) and PEC65 (nucleotides 556-576) Viral RNA dilution 1 5-1 5-2 5-3 5-4 5-5 5-6 Neg 1 5-1 5-2 5-3 5-4 5-5 5-6 Neg +/- IC RNA (10-6)++++++++ ------192

472 bp 330 bp

123456 78 M 9 1011121314 1516

Fig. 4.2 Agarose gel (1.5%) eletrophoresis of the RT-PCR products (ethidium bromide staining) of the Po/SaV/Cowden strain and IC RNA. Viral RNA was serially diluted and amplified with or without the presence of IC RNA with the primer pair PEC66/65. Lane M = 100 bp DNA ladder. Lanes 1-7, the undiluted RNA and 5-fold serially diluted viral RNA was co-amplified with the IC RNA, lane 8 was an RNA negative control with IC RNA; lanes 9-15, the undiluted RNA and 5-fold serially diluted viral RNA was amplified without IC RNA, lane 16 was an RNA negative control. The 330 bp bands are the virus-specific amplicons and the 472 bp bands are the IC RNA amplicons. 4 8 6 4 9 Dilution of RT- 8 0 : 16 32 :6 128 256 512 102 204 4 eg M 1 : N M PCR products 1: 1: 1 1: 1: 1: 1: 1: 1 193

225 bp

Hybridization 1 7 0 0 0 8 4 2 7 0 5 .2 .0 .6 .1 .4 .8 .5 .3 .1 .1 .0 3 3 2 2 1 0 0 0 0 0 0 A450 nm

Cut-off=0.20 Fig. 4.3 Comparison of the detection limits of agarose gel electrophoresis and the microwell hybridization assay (probe PEC-P1) for detection of RT-PCR PEC68-Bio/67 products of Po/SaV/GVI?/JJ681. The positive viral RNA amplicons were 2-fold serially diluted from 1:8 to 1:4096 and detected by agarose gel electrophoresis (10 µl/well) and the hybridization assay (12.5 µl/well), which are shown as A450. M = 100 bp DNA ladder. Neg = negative control for RT-PCR. Agarose gel electrophoresis was positive at a 1:128 dilution of the RT-PCR products, whereas the microwell hybridization results were positive through the dilution of 1:1024 (8-fold increased sensitivity).

CHAPTER 5

PREVALENCE OF NOROVIRUSES AND SAPOVIRUSES IN SWINE

DETERMINED BY REVERSE TRANSCRIPTION-PCR

5.1 SUMMARY

Noroviruses (NoV) and sapoviruses (SaV) are emerging enteric pathogens that cause diarrhea in humans and animals. Porcine NoVs replicate in pigs but their pathogenesis is undefined. Porcine SaV Cowden strain causes diarrhea and intestinal lesions in pigs. Recently, porcine NoVs and SaVs genetically or antigenically related to human strains were identified from normal pig fecal samples, raising questions of whether pigs are reservoirs for human strains. Only one seroprevalence study of one genotype of porcine NoV has been reported and no prevalence study has been reported for porcine SaVs. To investigate the prevalence of porcine NoVs and SaVs, six hundred and twenty-one fecal samples were collected from swine of various ages originating from

194 8 swine farms and 1 slaughterhouse in 3 states in the US. Except for 11 samples from

diarrheic pigs, all swine were clinically normal at the time of sampling. Fecal samples

were tested by reverse transcription (RT)-PCR with porcine NoV- and SaV-specific

primers. Surprisingly, porcine NoVs were detected exclusively from finisher pigs (10-24 wks of age) with an overall prevalence of 20% (range of 0-40%). Porcine SaVs were detected in 62% (range of 37-100%) of pigs with the prevalence highest in post-weaning pigs and lowest in nursing pigs. Because some porcine NoVs and SaVs are genetically or antigenically related to human strains, the high prevalence and subclinical infection rate of these viruses in pigs raise the question of whether pigs may be reservoirs for human strains.

5.2 INTRODUCTION

Members of the Norovirus genus within the family Caliciviridae, except for the newly identified murine norovirus (NoV) (19), cause gastroenteritis in humans and calves

(8, 21). Norovirus RNA has been detected from field adult pig fecal samples in Japan and

Europe (31, 33). Human noroviruses (NoV) are estimated to cause 23 million cases (66%) of food-borne human illness annually in the US (24) and more than 90% of human

nonbacterial epidemic gastroenteritis worldwide (5, 8, 22). Porcine NoVs have been

demonstrated to replicate in gnotobiotic (Gn) pigs but their pathogenesis is yet to be

defined (Q. H. Wang, M. G. Han, S. Cheetham, M. Souza, J. A. Funk and L. J. Saif,

195 submitted for publication). Sapoviruses (SaV) belong to another genus, Sapovirus within the family Caliciviridae. They cause sporadic cases and outbreaks of gastroenteritis mainly in young children < 5 years of age (27). Porcine SaV prototype Cowden strain causes diarrhea and intestinal lesions in gnotobiotic (Gn) pigs (7, 9).

Recently, genetically diverse porcine NoVs and SaVs were identified (32) (Q. H.

Wang, M. G. Han, S. Cheetham, M. Souza, J. A. Funk and L. J. Saif, submitted for publication). Based on phylogenetic analysis of the predicted complete capsid and partial

(~260 aa) RNA-dependent RNA polymerase (RdRp) regions, porcine NoVs were classified into 3 genotypes within genogroup II (GII) [GII-11 including the porcine NoV prototype Sw43/97/JP; GII/18 (QW101-like) and GII/19 (QW170-like)], which include the predominant human NoV strains (5). Porcine SaVs comprise at least 2 genogroups

[GIII including the porcine SaV prototype Cowden strain and GVI? (JJ681-like)]. More interestingly, the porcine NoVs were genetically and antigenically related to human GII

NoVs (6). Recently, a high prevalence (71%) of GII NoV antibodies in US swine was reported (6). These results raise questions of whether pigs are reservoirs for emergence of human NoVs.

At present, no prevalence study has been reported for porcine NoVs and SaVs in pigs or in humans with occupational exposure to pigs (e.g., farm workers, swine veterinarians and abattoir workers). Such studies would provide important information as to whether NoV and SaV gastroenteritis is a zoonotic disease. In this study, we collected

196 621 pig fecal samples from 7 swine farms and 1 slaughterhouse in 3 states. They were

tested by reverse transcription (RT)-PCR with porcine NoV-specific primer pair PNV7/8

(Q.H. Wang, K.O. Chang, M.G. Han, S. Sreevatsan and L.J. Saif, submitted for

publication) and porcine SaV-specific primer pair PEC66/65 (9), respectively. The primer

pair PNV7/8 targets the conserved RdRp region of all 3 genotypes of porcine NoVs and

PEC66/65 detected all GIII porcine SaVs and weakly detected LL26-like porcine SaVs

(unclassified, Q.H. Wang, M.G. Han, J. Funk, G. Bowman and L.J. Saif, submitted for publication). A competitive internal control (IC) RNA was used with primer pair

PEC66/65 to monitor for RT-PCR inhibition. Microwell hybridization assays using NoV- and SaV-specific DNA probes were performed to confirm the positive RT-PCR products.

5.3 MATERIALS AND METHODS

Sampling. Fecal samples (n=621) were collected from December, 2002 to March,

2005 from 7 swine farms and 1 slaughterhouse in 3 states [Ohio (OH), Michigan (MI)

and North Carolina (NC)] in the US, referred to as OH farm A, B, C and D, MI farm A,

NC farm A and B and OH slaughterhouse A (Table 5.1). Four age-groups of pigs were

included: nursing pigs [1-3 wks]; post-weaning (nursery) pigs (3-10 wks); finisher pigs

(10-24 wks) and sows (> 1 year of age). Except for 4 nursing pigs (1, 2 and 1 pigs from

OH farms A, C and D, respectively) and 7 post-weaning pigs (1, 5 and 1 pigs from OH farms A, B and C) that had diarrhea at sampling, all others were clinically normal at the

197 time of sample collection. Most samples were convenience samples from another

on-going project. Later, we sampled OH farm B for 5 more times from different ages of

pigs and in different seasons to investigate whether the prevalence of porcine NoV and

SaV differed among ages and seasons. The OH farm B and MI farm A had total pig

inventories of approximately 50,000 pigs and the other farms had total farm inventories

of approximately 1,000 pigs. The Ohio slaughterhouse received pigs from multiple farms

and the farms were not identified during collection. Fresh fecal samples were collected

from individual pigs, placed into sterile specimen containers and stored frozen at -20ºC.

RNA extraction. Sample RNA was extracted from 200 µl of centrifuged (1,200 × g, 30 min) 10% fecal suspensions in sterile Eagle’s minimal essential medium (EMEM,

Invitrogen, Carlsbad, CA) by using the Trizol LS (Invitrogen) procedure. The RNA pellet was resuspended in 40 µl RNase-free water and stored at -20 or -70ºC.

RT-PCR to detect porcine GIII SaVs and to monitor for RT-PCR inhibitors.

One-step RT-PCR was performed first with primer pair PEC66 (5’-GAC TAC AGC AAG

TGG GAT TCC-3’) and PEC65 (5’-ATA CAC ACA ATC ATC CCC GTA-3’) (9) and a competitive IC RNA (Q.H. Wang, K.O. Chang, M.G. Han, S. Sreevatsan and L.J. Saif, submitted for publication) to detect GIII SaVs and to monitor for RT-PCR inhibition simultaneously. Briefly, a final volume of 50 µl reaction mixture (45 µl of master mixture plus 4 µl of sample RNA and 1µl of 10-6 dilution of IC RNA) contained 1 × PCR buffer

(50 mM KCl, 10 mM Tris-HCl [pH9.0], and 0.1% Triton X-100), 2.5 mM MgCl2, 200

198 µM of dNTPs, 0.5 µM of each primer, 10 U of RNase inhibitor RNasin (Promega,

Madison, WI), 4 U of avian myeloblastosis virus reverse transcriptase (AMV RT,

Promega), 2 U of Taq DNA polymerase (Promega). The RT reaction was conducted at

45ºC for 60 min followed by heat inactivation at 94ºC for 3 min. The PCR reaction was performed for 40 cycles with each cycle at 94ºC for 30 sec, 50ºC for 30 sec and 72ºC for

30 sec, and a final extension for 10 min at 72ºC.

The RT-PCR products were tested by microwell hybridization assays with a mixture of two GIII SaV-specific probes (PoSapoP1A and PoSapoP1B) or with one IC

RNA-specific probe ICRNA-P1 (see below). Alternatively, the IC RNA amplicons were detected by agarose gel electrophoresis stained with ethidium bromide and visualized using ultraviolet light. The IC RNA amplicons are 472 bp, which were readily differentiated from the porcine GIII SaV-specific amplicons (330 bp).

There were three possible outcomes of the RT-PCR assays (12): (1) no RT-PCR

products (the assay has failed and inhibition may be present); (2) the IC RNA, but not the

virus amplicons are obtained (the assay has worked, and the sample is most likely to be

negative or viral RNA below detection limit); (3) the virus amplicons, with or without the

IC RNA amplicons, are obtained (the assay has worked, and the sample is positive).

When a sample contained RT-PCR inhibitors tested with primer pair PEC66/65, this

sample likely also contains RT-PCR inhibitors for primer pair PEC7/8 because the

RT-PCR inhibitors mainly interfere with activities of the reverse transcriptase and DNA

199 polymerase which catalyze the RT and PCR reactions, respectively. Then, this sample

RNA was diluted 1:10 or more in RNase-free water until it no longer inhibited the

RT-PCR reaction. This freshly diluted RNA was also used for RT-PCR with primer pair

PNV7/8.

RT-PCR to detect porcine NoVs. RT and PCR were performed separately using primer pair PNV7 (5’-AGG TGG TGG CCG AGG AYC TCC T-3’) and PNV8 (5’-TCA

CCA TAG AAG GAR AAG CA-3’) as previously reported with minor modifications (17).

For RT, 25 µl of reaction mixture (22 µl of master mixture plus 3µl of sample RNA) containing 1 X PCR buffer, 2.5 mM MgCl2, 400 µM of dNTPs, 1 µM of reverse primer

PNV8, 10 U RNasin and 2 U AMV RT was incubated at 45 ºC for 60 min followed by

heat inactivation at 94ºC for 3 min then chilling to 4ºC. Afterwards, another 25 µl of reaction mixture containing 1 × PCR buffer, 2.5 mM MgCl2, 1 µM of forward primer

PNV7, and 1 U Taq DNA polymerase was added into the above RT mixture to a final volume of 50µl to perform PCR reaction for 40 cycles with each cycle at 94ºC for 30 sec,

50ºC for 30 sec and 72ºC for 30 sec, and a final extension for 10 min at 72ºC. The porcine NoV-specific amplicons (211 bp) were confirmed by a microwell hybridization assay (see below).

Microwell hybridization assays. Microwell hybridization assays for detection of porcine NoV, SaV and IC RNA amplicons were performed as previously described for detection of Brucella abortus and/or Mycobacterium bovis (29). Forward primer PNV7

200 and PEC66 were biotinylated at the 5’-end (Integrated DNA Technologies, Coralville, IA).

For detection of the porcine NoV amplicons of RT-PCR with primer pair PNV7/8, the

EIA/RIA 8-well strips (Corning Inc., Corning, NY) were coated with a mixture of 3

NoV-specific probes PoNoroP1A (5’-AGC CAG TGG GCG AAG GAG TTC CAC TGT

GAT GTG CA-3’), PoNoroP1B (5’-AGC CAA TGC GCT ATG GAG TTC CAC TGT

GAT GTG CA-3’) and PoNoroP1C (5’-AGC CAA TGG GCA AAG GAA TTC CAT

TGT GAT GTG CA-3’) (100 ng/well of each probe). The mixture of SaV-specific probes

PoSapoP1A (5’-ATR ACA CTG GTG AAG GGC ATG CCA GAG GGG AG-3’) and

PoSapoP1B (5’-ATC ACA CTG GTG AAG GGC ATG CCT GAT GGT AA-3’) (100 ng/well of each probe) were used to coat the EIA/RIA strips for detection of porcine

SaV-specific amplicons by RT-PCR with primer pair PEC66/65. Probe ICRNA-P1

(5’-AGA ATA GAC CGA GAT AGG GTT GAG TGT TGT TCC AGT-3’) (100 ng/well) was used to coat the strips to detect IC RNA amplicons. After coating at 37ºC overnight, the wells were blocked with 1% BSA in PBS-Tween 20 (PBS-T, 0.05%) at 37ºC for 2 hours. The RT-PCR products were denatured by adding an equal volume of denaturing buffer (0.4 M NaOH, 80 mM disodium EDTA, 0.005% thymol blue). Next 25µl of the denaturing solution containing 12.5µl of RT-PCR product was added to duplicate wells containing 100µl of neutralization-hybridization buffer (1 M sodium thiocyanate, 80 mM

NaH2PO4; 10mM Na2HPO4, pH5.0 ±0.2). After incubating at 37ºC for 1 h, the wells were washed with the PBS-T. Then 100 µl of 0.4 µg/ml NeutravidinTM Horseradish Peroxidase

201 conjugate (Pierce Biotechnology, Rockford, IL) in 1% BSA buffer was added to each

well and incubated at 37ºC for 15 min. The wells were washed, the substrate

tetranethylbenzidine (KPL, Gaithersburg, MD) was added and strips were incubated in

the dark for 10 min. The reaction was stopped by adding 100 µl of 1.8 % HCl. After 5

min equilibration, the absorbance (A) was read by a spectrophotometer at 450 nm. For

each hybridization assay, RT-PCR positive and negative controls and water as a blank

were included. The cut-off value for each primer pair-probe combination was determined

based on the highest A450 of the non-specific amplicons multiplied by 2. The cut-off

values were A450 = 0.20 for porcine NoVs, A450 = 0.30 for porcine SaVs and A450 = 0.20 for the IC RNA. These microwell hybridization assays were similar (for SaVs and IC

RNA) or 2-fold more sensitive (for NoVs) than agarose gel electrophoresis for detection of the RT-PCR products.

Statistical analysis. Prevalence differences among farms, pig age groups and

collection time points were analyzed by Fisher’s exact test (Statistical Analysis System,

SAS Institute). Statistical significance was assessed throughout at P < 0.05.

5.4 RESULTS

A total of 621 pig fecal samples were tested for porcine NoVs and SaVs by

RT-PCR. These samples were collected during a 4 year-period from 7 US swine farms and 1 slaughterhouse.

202 Porcine NoVs were most prevalent among subclinically infected finisher pigs.

We used primer pair PNV7/8 to detect all 3 genotypes of porcine NoVs and the

NoV-specific amplicons were confirmed by a microwell hybridization assay. Surprisingly,

NoVs were detected exclusively from finisher pigs although other ages of pigs were also

tested (Table 5.2). In finisher pigs, the overall NoV prevalence was 20%; 4 of 6 swine

farms with samples from finisher pigs and the slaughterhouse were NoV positive; the

NoV prevalence ranged from 3-40% among NoV-positive farms.

Next, we examined whether the NoV prevalence differed among the different collection times. We took OH farm B as an example and collected 105 fecal samples from finisher pigs at 3 times including spring (Mar., 2003, n=60; Mar., 2005, n=30), and summer (June, 2004, n=15) (Table 5.1). The prevalence of NoVs was up to 70% (42/60) among samples collected in March, 2003, whereas only 1 of 15 samples collected in June,

2004 was NoV positive and no samples (n=30) collected in March, 2005 were NoV positive.

Porcine SaVs were most prevalent among post-weaning swine. Primer pair

PEC66/65 targeting the most conserved RdRp region of SaVs was used to detect mainly

GIII porcine SaVs and weakly detected unclassified LL26-like porcine SaVs. The SaVs were detected from all farms and from all ages of pigs (Table 5.3). The overall prevalence of porcine SaVs was 62% and ranged from 37-100% among farms. More interestingly, the prevalence of SaV in post-weaning pigs (83%) was significant higher than in sows

203 (71%) and finisher pigs (64%) and the prevalence of SaV in nursing pigs (21%) was significantly lower than in any other age group.

We then examined whether the SaV prevalence in pigs was different among different collection times using OH farm B as an example (Table 5.4). Samples were collected at 6 time points (Table 5.4) including winter (Dec.), spring (Mar. and May) and summer (June and July). The SaVs were detected at every time tested from this farm. In post-weaning pigs and sows, samples collected in June and July showed lower prevalence than the samples collected at other times. One unexpected observation was that all 30 finisher pigs were SaV negative whereas 100% (30/30) of post-weaning pigs were

SaV-positive in March, 2005, which differed from the May, 2003 and June, 2004 results when both groups had SaV positive pigs.

We also compared the SoV prevalence in normal and diarrheic pigs. Only 4 nursing and 7 post-weaning pigs had diarrhea when sampled. The SaV prevalence in normal nursing pigs (21/94, 22%) was higher than that (0/4, 0%) of diarrheic nursing pigs, whereas for post-weaning pigs, the SaV prevalence of normal pigs (97/117, 83%) was lower than that (7/7, 100%) of diarrheic pigs. However, because of the low diarrheic sample numbers, such differences were not significant.

Co-infection with NoVs and SaVs occurred in finisher pigs. The co-infection of pigs by porcine NoVs and SaVs was found in finisher pigs from 3 of 4 NoV positive farms and the slaughterhouse (Table 5.5). The co-infection rate [number of

204 co-infection/number of SaV and (or) NoV positive] was 27% overall ranging from 0-62% among farms.

Prevalence of RT-PCR inhibitors in field pig fecal samples. We used a competitive IC RNA control to monitor for RT-PCR inhibitors. We found that overall 7% of samples contained RT-PCR inhibitors (Table 5.6). After 10-fold dilution of extracted

RNA, those samples were no longer inhibited in RT-PCR reactions and 44% of them became NoV or/and SaV positive.

5.5 DISCUSSION

In this study, we investigated the prevalence of porcine NoVs and SaVs in US swine by RT-PCR with NoV- and SaV-specific primers targeting the viral RdRp region, respectively. We analyzed 621 pig fecal samples collected from 7 swine farms and one slaughterhouse from 3 states.

Surprisingly, NoVs were detected only from finisher pigs although pigs of other ages were tested. We sampled 98 nursing pigs and 124 post-weaning pigs at 12 different times, but still failed to detect porcine NoVs from these two age groups. In human NoV infections, NoVs have been detected in young children between 2 months and 2 years of age (26), but no NoV infection was reported in infants less than 2 months of age. In a comparative seroprevalence study of Norwalk virus and rotavirus in infants, children and adults in the US (18), investigators found that antibodies to rotavirus are acquired rapidly

205 during infancy and early childhood and by 36 months of age over 90% of individuals have rotavirus antibodies, whereas Norwalk virus antibodies are acquired gradually in childhood, then more quickly during the adult years and by 50 years of age about 50% of individuals have Norwalk virus antibodies. These results suggest that Norwalk virus was not important cause of illness in infants and young children, and in adults probably only a

portion are susceptible to Norwalk virus infection. Recently, researchers found that the

human NoVs bind to human histo-blood group antigens (HBGA) which were found to

influence the genetic susceptibility of individuals to NoV infections (10, 13-15, 20, 23).

Because the HBGAs are developmental antigens (11, 35), they may not be expressed in

high levels in the gut of the younger (nursing and post-weaning pigs). In this case, perhaps only adult pigs can be most readily infected with NoVs. However, why the adult sows were NoV negative in our study is unclear. Possibly the sows (> 1 year) were infected with porcine NoVs when they were younger and had acquired immunity to NoV infection. Sampling and testing first litter swine (gilts) of a similar age to the finisher pigs

(< 24 wks) may help to answer this question. Alternatively the greater concentration of swine in the finisher facilities as well as differences in housing and management as compared to sow facilities may have influenced NoV shedding. The finisher barn environment may be more analogous to the crowded, confined human populations,

typical on cruise ships (16).

In OH farm B, the prevalence of NoVs in Mar., 2003 (42/60, 70%) was much

206 higher than at the 2 other times: July, 2004 (1/15, 7%) and Mar., 2005 (0/30). Likely a

NoV outbreak occurred at this time in the finisher pigs in OH farm B. Excluding these possible outbreak samples, NoV sporadic infection was 10% in the finisher pigs.

On the other hand, the SaVs were detected from all ages of pigs with an overall prevalence of 62%. The SaV prevalence in nursing pigs (21%) was significantly lower than in any other age group, possibly due to protection of nursing pigs by maternal antibodies to SaVs in the milk. On the other hand, the prevalence of SaV shedding in sows was high (71%). It is not surprising that the prevalence of SaVs is the highest in post-weaning pigs (83%) because during this period pigs stop receiving maternal antibodies through milk and are under social (mixing) and environmental stresses. When we compared the SaV prevalence in normal or diarrheic pigs, we found that in nursing pigs, the prevalence in normal nursing pigs (21/94, 22%) was higher than that in diarrheic pigs (0/4, 0%). However, in post-weaning pigs the SaV prevalence in normal pigs

(97/117, 83%) was lower than that (7/7, 100%) in diarrheic pigs. These results suggest that SaVs may be involved in causing diarrhea in post-weaning pigs. However, there was no statistical significance due to the too small of a number of diarrheic pigs. The ability of porcine SaVs to cause intestinal lesions and diarrhea has been confirmed using the

SaV Cowden strain in studies of SaV seronegative Gn pigs (7, 9).

No obvious clinical signs were observed for the NoV or SaV fecal positive adult pigs (sows and finisher pigs) and only a small portion of SaV-positive post-weaning pigs

207 (7/104, 7%) had diarrhea at sampling. These results suggest that porcine NoVs may not

cause clinical signs in older pigs. However, SaVs may induce diarrhea in post-weaning

pigs but may not cause clinical signs in adult pigs. Further studies should be done to

detect NoV and SaV infection in pregnant sows versus gilts and to investigate whether

NoV and SaV are also common in pigs with diarrhea as few diarrheic pigs were sampled

and tested.

On the other hand, the high subclinical infection rate of NoVs and SaVs in pigs is important for the persistence of these viruses in nature and raises the risks of emergence of strains that may transmit to humans. Instead of being homogeneous, RNA virus populations consist of closely related viral mutants and recombinants, which are known as quasispecies (3). They are subjected to continuous genetic variation, competition and selection during virus replication. When the virus population size increases, the expected

number of viral genomes with more mutation sites also increases. Subsequently, this

raises the risk of emergence of “new” variants with changes in host cell tropism or the

ability to replicate in a different host species. For example, hantaviruses are apathogenic

in several rodent species. However, they infected humans causing severe pulmonary

syndrome in 1993 because of the increased numbers of infected mice during the

unusually mild and wet spring season (28).

We have inoculated 2 Gn pigs with the filtrates of 2 pig fecal samples collected

from OH farm B that tested positive for porcine GII-18 NoVs (Q. H. Wang, M. G. Han, S.

208 Cheetham, M. Souza, J. Funk and L. J. Saif, submitted for publication). Virus shedding was detected in 2 pigs by RT-PCR with primer pair PNV7/8. The NoV particles were observed in intestinal contents of one pig by immune electron microscopy. These results indicate that the subclinically NoV infected finisher pigs shed infectious virus. Because certain porcine NoVs are genetically and antigenically related to human GII strains, such a high prevalence of infectious viruses in subclinically infected swine including their detection from slaughterhouse pigs raises concerns of whether pigs may be reservoirs for human strains.

Co-infection of NoVs and SaVs was detected in 27% of 188 finisher pigs. In

humans, mixed infections of NoVs and SaVs were reported in 1 of 36 outbreaks (2) and

mixed infections of NoVs and other enteric viruses, particularly rotaviruses and less

frequently adenoviruses and astroviruses, are also common (25, 30). Whether NoVs and

SaVs are present as co-infections with known pig pathogens, such as porcine rotaviruses,

and whether such mixed infections exacerbate clinical conditions of pigs needs to be

investigated.

RT-PCR is the primary assay for detection of NoVs and SaVs because it is the

most sensitive technique. Primers are key factors for the sensitivity and specificity of

RT-PCR assays. Primer pair PNV7/8 and PEC66/65 were the most sensitive primers for

detection of porcine NoVs and GIII SaVs, respectively, compared to several other NoV

and SaV primer pairs tested (Q.H. Wang, K.O. Chang, M.G. Han, S. Sreevatsan and L.J.

209 Saif, submitted for publication). In addition, fecal samples often contain RT-PCR

inhibitors decreasing the sensitivity of reactions (1). These inhibitors are

sample-dependent and may include body fluids, food and bacterial cell constituents (34).

An IC RNA co-amplified with the target viral RNA can be used to monitor for inhibitors

as well as technical errors. Diluting the extracted RNA is simplest and most fecal samples

diluted 1:10 or 1:50 will no longer inhibit RT-PCR (4). In this study, we found that 7% of

pig fecal samples contained RT-PCR-inhibitors and a 1:10 dilution of sample RNA no

longer inhibited RT-PCR. A pitfall of this strategy is that the dilution of weakly positive

samples may exceed the detection limit of RT-PCR and therefore such samples may

become negative. On the other hand, false-positive results may be produced because fecal

samples contain mixtures including sloughed epithelial cells, undigested food

components and the normal intestinal bacterial flora (Q.H. Wang, K.O. Chang, M.G. Han,

S. Sreevatsan and L.J. Saif, submitted for publication). Compared to sequencing of the

amplicons, the use of a microwell hybridization assay probably is the cheapest, least time-consuming approach to interpret and confirm RT-PCR products, particularly for a

large epidemiological study.

To our knowledge, this is the first prevalence study of the shedding of NoVs and

SaVs in pigs. Porcine NoVs were detected exclusively from normal adult finisher pigs.

Porcine SaVs infected all ages of pigs with the shedding highest in post-weaning pigs and lowest in nursing pigs. Mixed infections of NoVs and SaVs were also found in finisher

210 pigs. The high prevalence of porcine NoVs and SaVs in pigs raises public health concerns

of their potential for zoonotic transmission.

5.6 ACKNOWLEDGMENTS

This work was supported by grants from the National Institute of Allergy and

Infectious Diseases, National Institutes of Health (Grant R01 AI 49742) and NRI, US

Department of Agriculture (CGP Grant 1999 02009) and by the Ohio Agricultural

Research and Development Center (OARDC), The Ohio State University (Graduate

Student Research Enhancement Grant project 2002-114). Salaries and research support were provided by state and federal funds provided to the OARDC.

Special thanks to Mr. Terry Workhemp, Mr. Carl Link, Mr. Alan Evers, Dr. Gary

Bowman and Dr. Myung Guk Han for help in sample collection. We thank Ana Azevedo for technical assistance. We also thank Dr. Jeffrey LeJeune (Food Animal Health

Research Program, The Ohio State University) for critical review of the manuscript and help in sample collection.

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215

Swine farm or Season/date collected Nursing Post-weaning Finisher Sow slaughterhouse Spring Summer Winter (1-3 w) (3-10 w) (10-24 w) (≧1 y) Total MI farm A Dec., 02 NA NA 61 NA 61 NC farm A May, 03 NA NA 8 NA 8 NC farm B June, 03 NA NA 21 NA 21 OH farm A Apr., 04 14 12 12 13 51 OH farm A Mar., 03 NA NA 10 NA 10 OH farm B Mar., 03 NA 30 60 NA 90 OH farm B May, 04 NA 15 NA NA 15 OH farm B June, 04 NA 15 15 NA 30 OH farm B July, 04 31 NA NA 15 46

21 OH farm B Dec., 04 30 NA NA 15 45 6 OH farm B Mar., 05 NA 30 30 NA 60 OH farm C May, 03 NA NA NA 32 32 OH farm C May, 04 15 12 12 12 51 OH farm D May, 04 8 10 NA NA 18 OH slaughterhouse Apr., 03 NA NA 83 NA 83 Total 98 124 312 87 621 a: NA = not available.

Table 5.1 Number and time of fecal sample collection from nursing, post weaning, finisher pigs and sows from different farmsa.

Swine farm or slaughterhouse Nursing Post-weaning Finisher Sow Total MI farm A NA NA 2/61(3) NA 2/61(3) NC farm A NA NA 2/8(25) NA 2/8(25) NC farm B NA NA 0/21(0) NA 0/21(0) OH farm A 0/14(0) 0/12(0) 5/22(22) 0/13(0) 5/61(8) OH farm B 0/61(0) 0/90(0) 43/105(40) 0/30(0) 43/286(15) OH farm C 0/15(0) 0/12(0) 0/12(0) 0/44(0) 0/83(0) 21

7 OH farm D 0/8(0) 0/10(0) NA NA 0/18(0) OH slaughterhouse NA NA 12/83(14) NA 12/83(14) Total 0/98(0) 0/124(0) 64/312(20) 0/87(0) 64/621(10) a: NA = not available. a Table 5.2 The prevalence of porcine NoVs in pigs [No. positive/total (%)] .

Swine farm or slaughterhouse Nursing Post-weaning Finisher Sow Total MI farm A NA NA 23/61(37) NA 23/61(37) NC farm A NA NA 8/8(100) NA 8/8(100) NC farm B NA NA 16/21(76) NA 16/21(76) OH farm A 4/14(28) 12/12(100) 20/22(90) 13/13(100) 49/61(80) OH farm B 7/61(11) 71/90(78) 58/105(55) 23/30(76) 159/286(55) OH farm C 7/15(46) 12/12(100) 12/12(100) 26/44(59) 57/83(68) OH farm D 3/8(37) 9/10(90) NA NA 12/18(66) 21 OH slaughterhouse NA NA 65/83(78) NA 65/83(78) 8 Total 21/98(21)C b 104/124(83)A 202/312(64)B 62/87(71)B 389/621(62) a: NA = not available. b: Numbers with different superscript letters (A, B and C) differ significantly (P<0.05).

a Table 5.3 The prevalence of porcine SaVs in pigs [No. positive/total (%)] .

Pig age May, 2003 May, 2004 June, 2004 July, 2004 Dec., 2004 Mar., 2005 Total Nursing NA NA NA 4/31 (13) 3/30 (10) NA 7/61 (11) Post-weaning 22/30(73) 11/15 (73) 8/15 (53) NA NA 30/30 (100) 71/90 (79) Finisher 53/60 (88) NA 5/15 (33) NA NA 0/30 (0) 58/105 (55) Sow NA NA NA 9/15 (60) 14/15 (93) NA 23/30 (77)

21 Total 75/90 (83) 11/15 (73) 13/30 (43) 13/46 (28) 17/45 (38) 30/60 (50) 159/286 (56) 9 a: NA = not available.

Table 5.4 The prevalence of porcine SaVs in pigs of OH farm B at different times [No. positive/total (%)]a.

Swine farm or slaughterhouse NoVs SaVs Co-infection/positivea MI farm A 2/61(3) 23/61(37) 0/25(0) NC farm A 2/8(25) 8/8(100) 2/8(25) OH farm A 5/22(22) 20/22(90) 3/22(13) OH farm B 43/105(40) 58/105(55) 39/62(62) OH slaughterhouse 12/83(14) 65/83(78) 6/71(8)

22 Total 64/279 (23) 175/279 (63) 50/188 (27) 0 a: NoV or SaV or both positive.

Table 5.5 The prevalence of porcine NoVs, SaVs and co-infection in finisher pigs [No. positive/total (%)].

Swine farm or NoV or SaV postitive/ slaughterhouse Nursing Post-weaning Finisher Sow Total total inhibited (%)c

MI farm A NAb NA 0/61(0) NA 0/61(0) 0 NC farm A NA NA 0/8(0) NA 0/8(0) 0 NC farm B NA NA 1/21(4) NA 1/21(4) 1/1(100) OH farm A 0/14(0) 0/12(0) 0/22(0) 0/13(0) 0/61(0) 0 OH farm B 4/61(6) 10/90(11) 29/105(27) 0/30(0) 43/286(15) 20/43(46) OH farm C 0/15(0) 0/12(0) 0/12(0) 2/44(4) 2/83(2) 0/2(0) OH farm D 0/8(0) 0/10(0) NA NA 0/18(0) 0 22

1 OH slaughterhouse NA NA 3/83(3) NA 3/83(3) 1/3(33) Total 4/98(4) 10/124(8) 33/312(10) 2/87(2) 49/621(7) 22/49(44) a: RT-PCR inhibitors were tested by RT-PCR with primer pair PEC66/65 and a competitive IC RNA. b: NA = not available. c: After 1:10 dilution of extracted RNA and performed RT-PCR again.

Table 5.6 The prevalence of RT-PCR inhibitors in the pig fecal samples [No. positive/total (%)]a.

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