Avian Bornavirus Infection in Waterfowl

Home , Avian bornavirus, Borna disease, Borna disease virus

Pauline G. Delnatte

A Thesis presented to The University of Guelph

In partial fulfilment of requirements for the degree of Doctor of Veterinary Science in Pathobiology

Guelph, Ontario, Canada

©Pauline G. Delnatte, August, 2013

ABSTRACT

AVIAN BORNAVIRUS INFECTION IN WATERFOWL

Pauline G. Delnatte Advisor: University of Guelph, 2013 Dr. Dale A. Smith

Avian bornavirus (ABV) is a newly recognized cause of neurological disease and mortality in free-ranging geese and swans in Ontario. To determine the correlation between clinical signs, pathological lesions and presence of ABV in tissues of wild waterfowl, 955 pathology cases from Canada geese (Branta canadensis), trumpeter swans (Cygnus buccinator) and mute swans (Cygnus olor) were reviewed, and 51 cases selected based on the presence of pathology or clinical history suggestive of ABV infection. The presence of virus in brains, assessed by immunohistochemistry and reverse transcriptase-polymerase chain reaction (RT-PCR) was highly correlated with the presence of non-suppurative inflammation in the central, peripheral and autonomic nervous systems. Partial sequencing of the ABV-nucleocapsid gene from infected geese indicated a unique waterfowl genotype.

To estimate the prevalence of ABV infection in southern Ontario, cloacal swabs and blood samples were collected from 624 asymptomatic free-ranging waterfowl and evaluated using RT-PCR and an enzyme-linked immunosorbent assay, respectively.

Thirteen percent of Canada geese caught on the Toronto Zoo site shed ABV in urofeces iii

compared to none of the geese sampled at three other locations. The prevalences of ABV shedding in mute swans, trumpeter swans and mallard ducks (Anas platyrhynchos) were

9.3%, 0% and 0%, respectively. Serum antibodies were present, often at high prevalence, in birds from all four species and at each sampling site.

To investigate the possibility of vertical transmission of ABV in wild Canada geese, 53 eggs were collected from an infected flock. ABV was detected in the yolk of one infertile egg.

To determine whether poultry species were susceptible to infection with the waterfowl strain of ABV, domestic ducks, chickens and geese were inoculated with brain homogenate from ABV-infected Canada geese. ABV was not detected using immunohistochemistry and RT-PCR in any inoculated or control bird at 45 or 90 days post inoculation. No histological lesions consistent with ABV infection were found in any ducks and chickens; however, non-suppurative inflammation was present in nervous tissues of 5/13 inoculated geese, 5/13 control geese, and 3/8 geese euthanized prior to inoculation, suggesting the presence of a pre-existing, non-ABV neurotropic virus. iv

ACKNOWLEDGEMENTS

I would like to thank, most sincerely, my advisor Dale Smith for her support and mentorship throughout this program, and for offering me this project in the first place. Thank you so much for your patience, your generosity and your dedication. Thanks for always taking the time to go over any “problem”, whether it was looking at slides at the last minute or helping me to complete a tax return… I could not wish for a better advisor.

I am also extremely grateful to the other members of my advisory committee, Éva Nagy, Davor Ojkic, Josepha DeLay and Simon Hollamby, for their invaluable insights in this research, their enthusiasm and for always having their doors open for me. It has been a pleasure working with you.

I would like to thank all the pathologists of the department that have taught me the basics of pathology at the beginning of this program. Thanks also to Doug Campbell, Marina Brash and Claire Jardine for their contributions and for being happy to answer any of my questions. Special thanks go to Ian Barker. Ian, I feel extremely lucky to have made my first steps in wildlife pathology with you. I greatfully acknowledge Ian Tizard and Susan Payne of the Schubot Exotic Bird Health Centre (Texas A&M University) for their help and collaboration.

This work would have not been completed without the help of many other dedicated people. I have been impressed by the expertise and kindness of so many of you:

- I am indebted to David Leishman who ran an impressive number of ELISAs in a very timely manner. Thanks also to Betty-Anne McBey who provided essential technical assistance in the early stage of the ELISA design and for the sample collection of the experimental birds.

- I am very appreciative of the staff at the Animal Health Laboratory, especially Susan Lapos, Ana Rita Rebelo, Sarah Hoyland, Jane Coventry and Elizabeth Hillyer, for their immense help with this project. v

- I am grateful to Lenny Shirose, Carol Lee Ernst and Dave Cristo (CCWHC) for making graphs, findings slides and reports, and sorting out samples for me and always smiling.

- I would like to thank the Ontario Trumpeter Swan Re-introduction Program (Harry Lumsden, Bev and Ray Kingdon, Julie Kee and Kyna Intini), the Canadian Wildlife Service (James Vanos [CWS London & Guelph] and Christopher Sharp [CWS Ottawa]), the Ministry of Natural Resources (Rob Brook [MNR Peterborough]), the Toronto Region Conservation Authority (Danny Moro) and the Golf Glen Veterinary Clinic (Aurora, Ontario) for their assistance in collecting and processing samples for the prevalence part of this study. Field work has always been wonderful and I am very glad I had the chance to meet each of you.

- Thanks to the Animal Care staff of the OMAFRA Animal Isolation Facility for the daily care of my experimental birds and for having coped so well with the unexpected!

- Thanks to all the previous Toronto Zoo residents for their goose and swan histology reports that were critical for the retrospective part of this study. Special thanks go to Maya Kummrow and Charlene Berkvens for their key contribution early in this project.

- Thanks to all students that helped me immensely in this project, particularly Monika Janssen, Veronica Kay, Elizabeth Beck, Nicole Zaranek, Matthew Mak and Kyle Elias.

- I gratefully acknowledge Amy Kistler and Joseph deRisi (University of California - San Francisco) for providing the rabbit polyclonal antiserum used for the IHC; Susan Payne (Texas A&M) for providing the antigen used for the ELISA; and Pierre Yves Daoust for providing the samples from CCWHC - Atlantic Region used in the retrospective part of this study.

- Thanks to the Toronto Zoological Foundation, the Ontario Ministry of Agriculture, Food and Rural Affairs (OMAFRA), the CCWHC-Ontario, the OVC Pet Trust, the Animal Health Laboratory and the Schubot Exotic Bird Health Centre for their vi

generous financial help for this project. The stipend of my DVSc degree was provided by the Toronto Zoological Foundation.

I would like to take this opportunity to express my deep gratitude to the other people who supported me during this program and made these three years truly fantastic. I would like to dedicate this work to the staff of the Wildlife Health Center of the Toronto Zoo.

This residency would have been completely different without Graham Crawshaw and Chris Dutton. Thanks for trusting me, supporting me and teaching me so many things. You will always be the ones who made me the zoo veterinarian I am today… and it means a lot to me.

Graham, your passion for zoo and wild animals is beautiful to watch and inspirational. Your dedication and professionalism will always be a role model. I guess I will have to agree with this little kid… “You rock Dr Graham!”.

Chris, I’m not sure that I can thank you enough for all these unforgettable days spent together, whether we were driving around the zoo or spotlighting ferrets. I loved every single one and I miss them already. You are a brilliant veterinarian, a wonderful teacher and an incredible person. You have been my day-to-day support for these last three years. Thanks for everything…

A very special thank-you goes to Tasha Long, Michelle Lovering and Dawn Mihailovic for being the best technicians in the world! Thanks so much for your constant support, your kindness and your friendship. Thanks for always being so knowledgeable and resourceful. I wish I could “steal” you for wherever I’m going to work in the future!

I am still amazed every day by the expertise and dedication of the keepers in this beautiful zoo. I would especially like to thank all of the Health Center keepers, in particular Charles Guthrie, Andrew Lentini, Lydia Attard, Mark Bongelli, Margaret Kolakowski, Paula Roberts and Andrea Dada (the budgie keeper!). Thanks for making this hospital such a great place to work. I will miss you deeply. Thanks to all the other vii

keepers, curators and staff from the nutrition and reproduction departments. I learnt so much from all of you. It was an honour to be part of your team. And thanks to Iga Stasiak, Adriana Nielsen and Mélanie Ammersbach for all the fun we shared at the zoo or at Guelph.

I am also very grateful to Stéphane Lair, Guy Fitzgerald and Marion Desmarchelier for affording me the incredible opportunity to partake in this residency three years ago.

I obviously cannot forget my family and friends from France and elsewhere who have loved and supported me from a distance. It has always been such a good feeling to hear your voices on the phone or to see you very occasionally and realize that nothing had changed! A special thank to Yann for being the best best-friend one can hope for.

I would like to deeply thank my wonderful parents and brothers for their unconditional love. I know you think I am too far away, but I’m happy and it is worth it. I will always love you.

Pauline Delnatte,

Toronto, August 2013 viii

DECLARATION OF WORK PERFORMED

I declare that, with the exception of the technical analyses listed below, all the work reported in this thesis was performed by me.

All RT-PCRs were performed by the virology division of the Animal Health Laboratory (University of Guelph, Ontario). The histological staining and immunostaining of tissue sections were performed by the histotechnology division of the Animal Health Laboratory (University of Guelph, Ontario). All ELISAs were performed by Kyle Elias, Bethy-Anne McBey, and David Leishman at the Ontario Veterinary College (Dr Éva Nagy’s laboratory). The purification of the ABV nucleocapsid protein used for the ELISA was performed by David Leishman at the Ontario Veterinary College (Dr Éva Nagy’s laboratory). All slides of the retrospective and experimental studies were reviewed in consultation with Dr Dale Smith (histology) and Dr Josepha DeLay (IHC). ix

TABLE OF CONTENTS

ABSTRACT ...... ii

ACKNOWLEDGEMENTS...... iv

DECLARATION OF WORK PERFORMED ...... viii

TABLE OF CONTENTS ...... ix

LIST OF TABLES ...... xiv

LIST OF FIGURES ...... xvi

LIST OF ABBREVIATIONS ...... xvii

CHAPTER 1: LITERATURE REVIEW...... 1

PROVENTRICULAR DILATATION DISEASE IN AVIAN SPECIES ...... 1

Descriptive epidemiology of proventricular dilatation disease ...... 1

Clinical signs of proventricular dilatation disease...... 3

Pathological lesions of proventricular dilatation disease ...... 5

Diagnosis of proventricular dilatation disease: identification of characteristic lesions...... 6

Etiology of proventricular dilatation disease ...... 7

Clinical management of proventricular dilatation disease ...... 8

BORNAVIRIDAE VIRUSES ...... 11

Members of family Bornaviridae ...... 12

Genomic organization of bornaviruses ...... 13

Pathogenesis of bornavirus infections ...... 14

Epidemiology of Borna disease virus ...... 17

PATHOGENESIS OF AVIAN BORNAVIRUS INFECTION ...... 20

Experimental inoculation...... 20 x

Viral distribution in tissues ...... 21

Shedding and transmission ...... 22

Seroconversion ...... 23

DIAGNOSIS OF AVIAN BORNAVIRUS INFECTION ...... 24

Virus culture ...... 24

Reverse Transcriptase Polymerase Chain Reaction ...... 26

Sequence analysis ...... 28

Immunohistochemistry ...... 29

Serology ...... 30

Summary statement: diagnosis of proventricular dilatation disease versus avian bornavirus infection ...... 31

AVIAN BORNAVIRUS INFECTION IN WATERFOWL SPECIES ...... 32

First identification of avian bornavirus in wild waterfowl ...... 33

Subsequent identification of avian bornavirus in waterfowl species ...... 34

Molecular characterization of waterfowl avian bornavirus ...... 35

Significance of ABV infection in waterfowl ...... 36

RESEARCH GOALS AND OBJECTIVES ...... 39

CHAPTER 2: RETROSPECTIVE STUDY - Pathology and diagnosis of avian bornavirus infection in wild Canada geese, trumpeter swans and mute swans in Canada ...... 41

Abstract ...... 42

Introduction ...... 43

Materials and Methods ...... 45

Case selection ...... 45

Histopathology...... 45 xi

Immunohistochemistry ...... 46

Nucleic acid isolation and RT-PCR...... 46

Sequencing and sequence analysis ...... 50

Statistical Evaluation ...... 50

Results ...... 51

Cases meeting inclusion criteria ...... 51

Diagnostic findings ...... 56

Discussion ...... 71

CHAPTER 3: PROSPECTIVE STUDY – Seroprevalence and prevalence of cloacal shedding of avian bornavirus in free-ranging waterfowl ...... 77

Abstract ...... 78

Introduction ...... 79

Material and methods ...... 81

Study area and sample collection ...... 81

Antigen preparation ...... 84

Serology ...... 85

Controls and determination of cut-off for seropositivity ...... 86

Nucleic acid isolation and RT-PCR...... 86

Statistical analysis...... 87

Results ...... 88

Seroprevalence...... 88

Prevalence of viral shedding...... 94

Correlation between cloacal shedding and seropositivity ...... 95

Discussion ...... 99 xii

CHAPTER 4: Investigation of the possibility of vertical transmission of avian bornavirus in free-ranging Canada geese ...... 105

Abstract ...... 106

Introduction ...... 107

Materials and Methods ...... 108

Egg collection ...... 108

Artificial incubation...... 109

Necropsy and sample collection ...... 109

Nucleic acid isolation and RT-PCR...... 110

Serology ...... 110

Results and discussion ...... 111

CHAPTER 5: EXPERIMENTAL TRIAL - Susceptibility of domestic poultry to infection with the waterfowl genotype of avian bornavirus ...... 115

Abstract ...... 116

Introduction ...... 117

Materials and Methods ...... 118

Bird acquisition and housing ...... 118

Study Design...... 118

Preparation of the inoculum and inoculation...... 121

Animal Use ...... 122

Post-mortem examination ...... 122

Histopathology...... 122

Immunohistochemistry ...... 123

Nucleic acid isolation and RT-PCR...... 123

Antigen preparation and serology...... 124 xiii

Statistical analysis...... 126

Results ...... 126

Clinical observations and post-mortem macroscopic findings ...... 126

Histopathology...... 127

Immunohistochemistry and RT-PCR results ...... 132

Serology ...... 132

Discussion ...... 133

CONCLUSIONS – FUTURE DIRECTIONS...... 139

REFERENCES ...... 141

Appendix A: Birds and bornaviruses – Review...... 160

Appendix B. Sampling protocol for trumpeter and mute swans provided to the biologists and volunteers of the Trumpeter Swan Reintroduction Program ...... 173

Appendix C. Avian bornavirus nucleocapsid protein gene cloning, recombinant expression, and purification – Protocol...... 175

Appendix D. Avian bornavirus infection trial - Group assessment sheet ...... 177

xiv

LIST OF TABLES

Table 2.1. Real-time RT-PCR primer and probe sequences (a) and cycling parameters (b) used to identify avian bornavirus in frozen and fixed paraffin-embedded tissues from free-ranging waterfowl in Ontario ...... 49 Table 2.2. Number of Canada geese, trumpeter swans and mute swans meeting initial case inclusion criteria in a retrospective survey of necropsy databases from Toronto Zoo and CCWHC-Ontario from 1992 to 2011 ...... 52 Table 2.3. Summary of clinical history, gross necropsy findings, histopathology, IHC and RT-PCR results in 41 wild Canada geese evaluated for the presence of ABV ...... 54 Table 2.4. Summary of clinical history, gross necropsy findings, histopathology, IHC and RT-PCR results in 8 wild trumpeter swans and 2 wild mute swans evaluated for the presence of ABV ...... 55 Table 2.5. Tissue distribution of non-suppurative inflammatory lesions, immunohistochemical staining and RT-PCR results in 35 wild Canada geese evaluated for the presence of ABV ...... 57 Table 2.6. Tissue distribution of non-suppurative inflammatory lesions, immunohistochemical staining and RT-PCR results in eight wild trumpeter swans and one wild mute swan evaluated for the presence of ABV ...... 59 Table 2.7. Summary of overall, positive and negative percent agreements among diagnostic tests in Canada geese, trumpeter swans and mute swans evaluated for the presence of avian bornavirus...... 67 Table 3.1. Capture dates and locations and demographic information for Canada geese, mallards, mute swans and trumpeter swans in southern Ontario from which blood samples and cloacal swabs were collected between October 2010 and May 2012 to determine the prevalence of ABV infection ...... 83 Table 3.2. Prevalence of cloacal shedding of ABV determined by RT-PCR and prevalence of antibodies against ABV determined by ELISA in 206 Canada geese, 208 mallards, 75 mute swans and 135 trumpeter swans caught in southern Ontario ...... 89 xv

Table 3.3. Results of statistical analysis, where significant, using Fisher’s exact tests to compare the seroprevalences among species, locations, shedders versus non-shedders, age class and sexes as part of a prospective survey of ABV infection in free-ranging waterfowl in southern Ontario ...... 93 Table 3.4. Results of statistical analysis, where significant, using Fisher’s exact tests to compare the prevalences of cloacal shedding among species and locations as part of a prospective survey of avian bornavirus infection in free-ranging waterfowl in southern Ontario, Canada ...... 95 Table 5.1. Summary of diagnostic tests performed for each group of birds in a trial carried out to evaluate the susceptibility of domestic poultry species to experimental infection with ABV ...... 120 Table 5.2. Summary of the severity and distribution of central nervous system histological lesions observed in domestic geese used in a trial carried out to evaluate the susceptibility of domestic poultry species to experimental infection with ABV...... 129 xvi

LIST OF FIGURES

Figure 2.1. Histological lesions in waterfowl infected with avian bornavirus...... 62 Figure 2.2. Immunohistochemical staining for avian bornavirus ...... 64 Figure 2.3. Phylogenetic tree illustrating the genetic relationships between the waterfowl bornavirus and other bornaviruses ...... 70 Figure 3.1. Study area in southern Ontario, Canada, showing sampling locations for Canada geese, mute swans, trumpeter swans and mallards from which cloacal swabs and blood samples were analyzed to determine the prevalence of ABV infection ...... 82 Figure 3.2. Sample-to-positive ratio of serum samples assessed for the presence of antibodies against ABV using an ELISA for 203 Canada geese, 75 mute swans, 130 trumpeter swans and 92 mallards in southern Ontario...... 97 Figure 3.3. Sample-to-positive ratio of serum samples assessed for the presence of antibodies against ABV using an ELISA for 203 Canada geese at four different locations in southern Ontario...... 97 Figure 3.4. Sample to positive ratio of serum samples assessed for the presence of antibodies against ABV using an ELISA for 203 Canada geese that were or were not shedding ABV in cloacal swabs at the time of blood collection ...... 97 Figure 3.5. Sample to positive ratio of serum samples assessed for the presence of antibodies against ABV using an ELISA for 75 mute swans that were or were not shedding ABV in cloacal swabs at the time of blood collection ...... 97 Figure 5.1. Histological lesions observed in control and experimental domestic geese in a trial carried out to evaluate the susceptibility of domestic poultry species to experimental infection with ABV ...... 131 xvii

LIST OF ABBREVIATIONS

ABV avian bornavirus ABV-CG avian bornavirus Canada goose genotype AHL Animal Health Laboratory AHY after hatch year BDV Borna disease virus BLAST basic local alignment search tool bp base pair BSA bovine serum albumin CAGO Canada goose CCWHC-Atl Canadian Cooperative Wildlife Health Center - Atlantic region CCWHC-Ont Canadian Cooperative Wildlife Health Center - Ontario region CD8-T cluster of differentiation 8 T-cell CI confidence interval Cp crossing point value CWS Canadian Wildlife Service DNA deoxyribonucleic acid EBLN endogenous bornavirus-like nucleoprotein ELISA enzyme-linked immunosorbent assay FFPE formalin-fixed paraffin-embedded GPS global positioning system H&E hematoxylin and eosin HY hatching year IHC immunohistochemistry MALL mallard duck mRNA meesenger ribonucleic acid MNR Ministry of Natural Resources MUSW mute swan NPA negative percent agreement NSAID nonsteroidal anti-inflammatory drug nt nucleotide OD optical density OD 405 optical density read at 405 nm OPA overall percent agreement OR odd ratio ORF open reading frame PAGE polyacrylamide gel electrophoresis PBS phosphate buffered saline PCR polymerase chain reaction PDD proventricular dilation disease PPA positive percent agreement RBV reptile bornavirus RNA ribonucleic acid xviii

RT-PCR reverse transcriptase polymerase chain reaction rtRT-PCR real-time reverse transcriptase polymerase chain reaction SD standard deviation SDS-PAGE sodium dodecyl sulfate S/P ratio sample to positive ratio SST serum separator tube TRSW trumpeter swan U unknown uORF upstream open reading frame USDA United States Department of Agriculture

CHAPTER 1: LITERATURE REVIEW

PROVENTRICULAR DILATATION DISEASE IN AVIAN SPECIES

Proventricular dilatation disease (PDD) is a devastating disease affecting primarily psittacine birds (parrots) and has been seen as a clinical syndrome in North America since the late 1970’s. It is considered by many to be the greatest threat to the aviculture of psittacine birds including several highly endangered species, such as the Spix’s macaw, that depend on captive breeding for their survival (Wyss et al., 2009; Enderlein et al., 2011).

The name PDD refers to the clinical syndrome of the disease and is derived from the predominant feature of the disease in parrots: dilatation of the proventriculus as a result of localized atony due to virus-induced immune damage to the autonomic nerves affecting the upper and middle gastrointestinal tract. Other synonyms used in the literature reflect the underlying clinico-pathological features of the disease and include proventricular dilatation syndrome, macaw wasting/fading disease/syndrome, neuropathic gastric dilatation of Psittaciformes, psittacine encephalomyelitis, myenteric ganglioneuritis and infiltrative splanchnic neuropathy (Gancz et al., 2010; Staeheli et al., 2010). In 2008, a major advance in PDD research was made with the identification of a novel avian bornavirus (ABV) as a plausible etiological agent of PDD (Honkavuori et al., 2008; Kistler et al., 2008).

Descriptive epidemiology of proventricular dilatation disease

PDD was first described as a clinical syndrome in psittacine birds in the United States in 1978 (Ridgway et al., 1983; Mannl et al., 1987; Gregory et al., 1994) and in Canada in 1982 (Berhane et al., 2004). It was hypothesized that the disease was introduced to North America via an outbreak that occurred in young macaws (Ara sp.)

newly imported from Bolivia and released from United States Department of Agriculture (USDA) quarantine (Woerpel et al., 1984). PDD has since been observed in more than 80 species of psittacine birds (Clark et al., 1984; Mannl et al., 1987; Weissenböck et al., 2009a; Shivaprasad et al., 2010). Certain species, such as African grey parrots (Psittacus erithacus), blue and gold macaws (Ara ararauna), cockatoos, and Amazon parrots, seem most frequently affected (Schmidt et al., 2003) whereas other species, such as budgerigars (Melopsittacus undulatus), appear to be more resistant to the disease (Graham et al., 1991; Reavill et al., 2007). Whether this reflects a true difference in susceptibility or other factors that influence exposure or diagnosis is unclear (Gregory et al., 1994). Both sexes appear equally affected and the disease can affect birds as young as 5 weeks of age (Kistler et al., 2010), although PDD is more generally identified in adults (Graham et al., 1991; Gregory et al., 1994). Under most circumstances the disease seems to spread slowly; however, acute outbreaks with high mortality have been described in psittacine aviaries (Lublin et al., 2006; Kistler et al., 2010). Crowded indoor aviaries and nurseries seem to favour the occurrence of PDD outbreaks (Gancz et al., 2010).

Cases of PDD have been diagnosed in psittacine birds from most parts of the world. The majority of cases have been described in Europe and North America (Clark et al., 1984; Graham et al., 1984; Turner et al., 1984; Woerpel et al., 1984; Gerlach et al., 1991; Gregory et al., 1994; Gregory et al., 2000) but the disease has also been reported in Australia (Sullivan et al., 1997; Doneley et al., 2007), the Middle East (Lublin et al., 2006; Kistler et al., 2008; Wyss et al., 2009), South America (Marietto-Goncalves et al., 2009), South Africa (Gancz et al., 2010; Last et al., 2012) and Japan (Ogawa et al., 2011). All psittacine cases have been diagnosed in captive birds and it is assumed that intensive trading and bird smuggling have contributed to spread the disease among continents (Gregory et al., 1997; Doneley et al., 2007; Staheli et al., 2010). No PDD cases have been reported, to date, in free-ranging parrots on any continent (Villanueva et al., 2010); however, large-scale active surveillance has not been carried out in the wild. The possibility of transmission of PDD from captive parrots to wild birds or the possibility of the spreading of PDD to parrots by wild birds remains open.

Although the vast majority of cases of PDD occur in psittacine birds, clinical and pathological features consistent with PDD have been described in several other avian

orders, including canaries (Serinus canaria) (Perpiñán et al., 2007; Weissenböck et al., 2009a; Rinder et al., 2012; Rubbenstroth et al., 2013), a greenfinch (Carduelis chloris), a long-wattled umbrella bird (Cephalopterus penduliger), a bearded barbet (Lybius dubius) (Perpiñán et al., 2007), Canada geese (Branta canadensis) (Daoust et al., 1991; Smith et al., 2010), trumpeter swans (Cygnus buccinator) (Smith et al., 2010), a peregrine falcon (Falco peregrinus) (Shivaprasad et al., 2005), a bald eagle (Haliaeetus leucocephalus) (Hoppes et al., 2013), a roseate spoonbill (Ajaja ajaja), a toucan (Ramphastos sp.), a honeycreeper (Cyanerpes sp.) and a weaver finch (Gregory et al., 2000).

Clinical signs of proventricular dilatation disease

The clinical signs of PDD generally reflect malfunction of the gastrointestinal and / or nervous system, and vary in nature, severity and duration (Lumeij et al., 1994; Rosskopf et al., 2003). Non-specific signs include depression, lethargy, weight loss associated with reduced appetite or polyphagia, muscle atrophy, abdominal enlargement, polyuria and polydipsia (Doneley et al., 2007), as well as sudden death (Shivaprasad et al., 2010). Commonly reported gastrointestinal signs include dysphagia, crop stasis, regurgitation, impaction, maldigestion (passage of undigested seeds), and progressive loss of body condition leading eventually to emaciation and death (Gregory et al., 1994; Hoppes et al., 2010). Central nervous system signs include changes in awareness and demeanor, tremors, seizures, erratic head movements, torticollis, head-pressing, opisthotonus, abnormal gait and posture, inability to perch, proprioceptive and motor deficits, ataxia, paralysis and status epilepticus (Gregory et al., 1994). Recent papers have reported that ophthalmologic abnormalities, especially lesions of the fundus, are an important part of the clinical picture of PDD and could be early signs of the disease. Lesions include dilated pupils, transient anisocoria, chorioretinitis, loss of retinal pigmented epithelium and bilateral retinal degeneration; and can result in blindness (Steinmetz et al., 2008; Korbel et al., 2011).

The incubation period of PDD seems to be extremely variable. Under experimental conditions, a minimum of 11 days has been reported (Gregory et al., 1997) and some unweaned chicks developed clinical disease within 2 to 4 weeks of exposure to

PDD (Kistler et al., 2010), whereas in other studies the incubation periods were months or even years (Gregory et al., 1995, Gancz et al., 2009, Gray et al., 2010).

Parrots affected with PDD sometimes exhibit secondary or concurrent conditions which may significantly increase mortality rates. Secondary opportunistic infections, such as fungal pneumonia, clostridial enteritis and bacterial septicemia, are frequently described in affected birds, likely resulting from immunosuppression or stress (Doneley et al., 2007; Perpiñán et al., 2007). Recent reports describe concurrent congestive heart failure in an African grey parrot (Vice et al., 1992; Juan-Salles et al., 2011) and a concurrent feather picking disorder in an Eclectus parrot (Eclectus roratus) (Orosz et al., 2010; Horie et al., 2012). A clear association between PDD and either of these two conditions has not been established; these isolated reports are unlikely of significance in understanding the overall pathogenesis of the disease.

Proventricular enlargement and passage of undigested food in the feces are usually considered as good indicators of PDD, but are not pathognomonic for the condition. Differential diagnoses include megabacteriosis (infection with Ornithogaster macrorhabdos), clostridial enteritis, fungal and yeast infections, heavy metal toxicosis, intestinal obstruction, neoplasia and parasitism, as well as pyloric dysfunction and pancreatitis (Bond et al., 1993; Taylor et al., 1997; Antinoff et al., 2001; Hoppes et al., 2010). A variety of other nervous system diseases can mimic the central and peripheral neurologic signs seen in PDD.

It is generally accepted that PDD is a progressive disease which becomes fatal once marked clinical signs, especially neurological signs, develop (Graham et al., 1984). However, some reports suggest the existence of an acute form, with birds dying within days or weeks after acute onset of symptoms; and of a persistent form, with birds able to live for years without significant clinical impairment and with the possibility of recovery (Phalen et al., 1986).

Pathological lesions of proventricular dilatation disease

The macroscopic and microscopic lesions in birds with PDD have been extensively studied (Graham et al., 1991; Shivaprasad et al., 1995; Reavill et al., 2007). Gross lesions include mild to severe emaciation, atrophy of the pectoral, proventricular and ventricular muscles, proventricular dilatation (Berhane et al., 2001; Gancz et al., 2010) and duodenal distension. Proventricular rupture and resulting peritonitis have been reported rarely. Accumulation of a transparent fluid in the subarachnoid space occurs in some birds with neurological signs (Berhane et al., 2001). Occasionally, no gross lesions are observed.

Microscopically, PDD is characterized by non-suppurative inflammation in peripheral, central and autonomic nervous tissues (Shivaprasad et al., 1995; Berhane et al., 2001; Schmidt et al., 2003). Lymphoplasmacytic infiltrates within myenteric ganglia and nerves of the proventriculus and ventriculus, and less frequently of the crop, esophagus and duodenum, are the histological hallmarks of PDD (Ritchie et al., 1998) and are considered pathognomonic for the disease by some authors (Schmidt et al., 2003; Berhane et al., 2004). Similar infiltrates may also be present in the brain and spinal cord in the form of perivascular cuffing, and in peripheral nerves (Shivaprasad et al., 1995; Berhane et al., 2001), adrenal glands, myocardium, conductive tissue of the heart (Vice et al., 1992) and nerves or ganglia adjacent to various tissues including adrenals, epicardium and testes (Doneley et al., 2007). Non-suppurative leiomyositis, polyserositis (Doneley et al., 2007), retinal degeneration, chorioretinitis (Steinmetz et al., 2008; Korbel et al., 2011) and perivascular dermatitis have also been described (Mannl et al., 1987; Shivaprasad et al., 2010). Cerebellar Purkinje cell necrosis, neuronophagia, myelin degeneration, gliosis and axonal swelling sometimes accompany the inflammatory neural lesions (Berhane et al., 2001).

The frequency and extent of inflammation in various organs and in the different parts of the brain and spinal cord have been described by several authors (Hugues et al., 1984; Gerlach et al., 1991; Graham et al., 1991; Gregory et al., 1994; Shivaprasad et al., 1995; Berhane et al., 2001; Berhane et al., 2004). These vary greatly, are not consistent among cases, and often do not reflect the type or severity of clinical signs or the gross

lesions (Berhane et al., 2001). Whether there is a relationship between lesions and species of bird affected or stage of the disease is not known (Berhane et al., 2001; Doneley et al., 2007).

Diagnosis of proventricular dilatation disease: identification of characteristic lesions

Post-mortem examination and identification of characteristic lymphoplasmacytic infiltration in nerves and ganglia of the upper and middle digestive tract has been, and will likely remain, the gold standard for diagnosing PDD as a clinical entity (Schmidt et al., 2003). Whereas necropsy has provided a clear and concise diagnosis, ante-mortem diagnosis has proven to be more challenging.

Thorough physical examination of the bird, including ophthalmologic examination and cardiac auscultation, should always be performed. The presence of any of the digestive and/or neurologic symptoms mentioned earlier should suggest PDD; however, most of these are non-specific (Gregory et al., 1994). Diagnostic imaging, including survey radiography of the coelomic cavity, measurement of the proventriculus/keel ratio (Dennison et al., 2009), contrast radiography and gastrointestinal fluoroscopy, may indicate proventricular enlargement and reduced gastro-intestinal motility (Gancz et al., 2010). However, suboptimal positioning, interspecific differences and normal physiologic variations usually make diagnostic images challenging to interpret and unreliable to definitively confirm or rule out PDD (Dennison et al., 2010; Gancz et al., 2010; Hoppes et al., 2010; Villanueva et al., 2010). Clinical pathology findings are not specific and may reflect functional disturbances in multiple organs, including gastro-intestinal malabsorption, or secondary infections (Suedmeyer et al., 1992; Keller et al., 2010). Other tools that can help in diagnosing PDD include electrocardiography (Vice et al., 1992), ophthalmoscopy and electroretinography (Steinmetz et al., 2008; Korbel et al., 2011). All the previously mentioned tests will allow the clinician to increase his/her index of suspicion but are not considered confirmatory for the disease (Bond et al., 1993; Taylor et al., 1997; Antinoff et al., 2001; Hoppes et al., 2010).

A more specific ante-mortem test to confirm the diagnosis of PDD is crop biopsy and histopathology to identify the presence of non-suppurative ganglioneuritis. Although crop biopsy has been the standard recommended method of diagnosing PDD ante- mortem for decades, it is less than ideal. The inconsistent presence of inflammatory cells in the nerves of the crop (Doolen et al., 1994; Berhane et al., 2004) and the numerous technical factors that can affect the quality of samples obtained (Gregory et al., 1996; Ritchie et al., 2004) explain the variable and low sensitivity reported for this diagnostic tool, which ranges from 22% to 76% depending on the author (Graham et al., 1991; Doolen et al., 1994; Shivaprasad et al., 1995; Gregory et al., 1996; Berhane et al., 2001; Deb et al., 2008).

The discovery of the etiologic agent of PDD in 2008 (Honkavuori et al., 2008; Kistler et al., 2008) has led to the development of new diagnostic tests to confirm ABV infection. These tests are described in detail further (in the diagnosis section).

Etiology of proventricular dilatation disease

A viral etiology for PDD has long been suspected based on epidemiologic observations, the apparent infectious nature of the disease, the typical microscopic lesions and efforts to rule out other possible causes (Gerlach et al., 1991; Graham et al., 1991; Shivaprasad et al., 1995). A variety of viruses have been discussed as candidate causative agents, including paramyxovirus (Grund et al., 1999; Grund et al., 2002), eastern equine encephalitis virus (Gaskin et al., 1991), coronavirus (Gough et al., 1996; Gough et al., 2006), avian herpesvirus, togavirus, polyomavirus, adeno-like virus, enterovirus, reovirus, and avian encephalitis virus (Ritchie et al., 1995). Numerous attempts were made to identify a putative virus using viral culture in various cell lines, electron microscopy and serological studies, and infectious trials failed to confirm any of these agents as causative (Gerlach et al., 1991; Gregory et al., 1994; Gregory et al., 1995; Shivaprasad et al., 1995; Gregory et al., 1997; Gregory et al., 2000; Grund et al., 2005; de Kloet et al., 2009). Using electron microscopy, virus-like particles of 30 to 250 nm were visualized in tissues and feces of affected birds (Mannl et al., 1987), and an enveloped virus of about 80 nm in diameter was demonstrated in feces of affected birds

(Gregory et al., 1997; Hartcourt-Brown et al., 1997; Gregory et al., 2000; Berhane et al., 2004). An enveloped virus of approximately 80 nm in diameter was also isolated from the tissues of affected birds using macaw embryonic cell culture (Gough et al., 1996). Experimental inoculation of tissue homogenates containing virus-like particles from birds with PDD into healthy birds resulted in clinical symptoms and histopathological lesions consistent with PDD and confirmed the transmissibility of PDD (Gregory et al., 1997; Gregory et al., 2000; Ritchie et al., 2003). However, the nature of this virus could not be elucidated at that time and the etiology of PDD remained open.

In 2008, two research teams independently identified a novel viral agent from birds with PDD using advanced molecular technologies (Honkavuori et al., 2008; Kistler et al., 2008). Kistler et al. used a panviral DNA microarray containing representatives of all viral groups to investigate tissues from birds with PDD which had been collected in Israel and in the United States. They detected a bornavirus hybridization signature in 62.5% of the PDD cases and in none of the controls, and subsequently used ultra high- throughput sequencing combined with conventional PCR-based cloning to recover a complete viral genome sequence (Kistler et al., 2008). Pair-wise comparison of sequences showed that these newly identified avian strains shared only 64% nucleic acid homology with the Borna disease virus (BDV) of mammals (Kistler et al., 2008). Honkavuori et al. used unbiased pyrosequencing on tissues of three birds affected with PDD, and compared sequences obtained with known viral sequences using BLAST search and GenBank database. Viral sequences were not found in PDD-negative control birds. This group identified two distinct strains of a new virus sharing genetic similarity with the family Bornaviridae. The work of these two independent groups resulted in the identification of a novel species of bornaviruses, which was named avian bornavirus.

Clinical management of proventricular dilatation disease

The diagnosis of PDD in an individual bird or group of birds has implications for the flock and for any in-contact birds. Management strategies must be put into place to limit the spread of the disease to other birds, to determine the disease status of all birds in the collection and, if possible, to treat the affected birds. Assessment of the disease status

of in contact birds is problematic as there is no gold standard clinical diagnostic test. Prior to the identification of ABV, radiographic and fluoroscopic evaluation for gastric motility abnormalities and histopathological assessment of crop biopsies were the only methods of assessing live birds. Molecular screening tests for ABV are now available for clinical samples, and serologic screening tests are just entering the market. Interpretation of these tests can be challenging with the existence of intermittent shedders and asymptomatic carriers, and with minimal information available on test sensitivity and specificity. Infected birds should be isolated and treated (Clubb et al., 2009; Gancz et al., 2010).

Husbandry measures

Preventive measures currently aim to avoid introducing the pathogen into new flocks and thus rely on excellent husbandry practices and strict quarantine protocols including testing of all newly introduced birds (Gancz et al., 2010). Measures include strict isolation of all affected and in-contact birds, practicing good hygiene via thorough cleaning and disinfection and following strict biosafety rules, such as traffic control (Gancz et al., 2010; Hoppes et al., 2010). Although there are still no data on survival of ABV in the environment or sensitivity to disinfectants, it may be assumed that it has much the same stability as other enveloped RNA viruses of similar size and structure, and thus disinfection with phenols, formaldehyde or hypochlorites is recommended (Hoppes et al., 2013). Whereas horizontal transmission may be controlled by appropriate quarantine, housing and hygienic procedures, the possibility of vertical transmission of ABV (see later) adds difficulties to the management of ABV-infected aviaries. Pairing ABV-positive birds, incubating their eggs artificially, and hand-raising the chicks separately until they show negative test results might be a viable option, particularly for breeding projects involving rare species where every individual is genetically important and cannot be eliminated from the program (Lierz et al., 2011; Kerski et al., 2012).

Therapeutic measures

There is no specific antiviral treatment for birds infected with ABV (Staeheli et al., 2010). However, with supportive and symptomatic therapy, affected birds can sometimes survive for months to years (Suedmeyer et al., 1992; Gerlach et al., 1994; Gregory et al., 1997). Very few case / control studies on the efficacy of therapeutic drugs have been published (Gancz et al., 2010; Hoppes et al., 2010; Hoppes et al., 2012) and thus most recommendations are based on the personal experience of various authors.

As non-suppurative inflammation is a histological hallmark of PDD, therapeutic trials have included the use of various anti-inflammatory drugs. The treatment of affected birds with non-steroidal anti-inflammatory drugs (NSAIDs), such as celecoxib, tepoxalin and meloxicam, has been commonly recommended but results have been inconsistent. Celecoxib has been reported as successful in treating PDD, with a slow and gradual clinical response (Dahlhausen et al., 2002; Clubb et al., 2009); however, there was no untreated control group for comparison in this study. A recent study in experimentally infected cockatiels suggested that the use of meloxicam would actually be detrimental to birds affected with PDD as they observed the development of more severe lesions in treated birds in comparison with the non-treated group (Hoppes et al., 2012).

Because selective removal of T cells with immunosuppressive drugs significantly enhances survival of BDV-infected rats (Stitz et al., 1989), it is anticipated that immunosuppressive protocols, especially selective T-cell elimination by drugs such as cyclosporine, may be of therapeutic benefit (Stitz et al., 1989). Research on the efficacy of cyclosporine as a treatment for birds with PDD is currently underway at Texas A&M.

Antiviral drugs, such as amantadine hydrochloride, have been described as being beneficial in birds exhibiting severe gastro-intestinal and neurological signs by some authors (Clubb et al., 2009; Gancz et al., 2010), but have been reported as having no apparent effect on fecal viral shedding by others (Hoppes et al., 2010). Ribavirin readily kills ABV in tissue culture (Hoppes et al., 2013) but does not appear to have a measurable effect on viral shedding in vivo.

Finally, an immuno-modulating therapeutic approach is recommended by some authors (Rossi et al., 2012). Recent trials used an anti-COX2 NSAID (Robenacoxib)

combined with Mycobacterium bovis extracts, intended to redirect the activated antigen- specific T cells to local inflammatory sites, and showed some promise; however, this study was again not case-controlled and thus the results are inconclusive.

In addition to the above anti-inflammatory and antiviral protocols, numerous general palliative therapeutic measures have been described for the long term management of birds affected with PDD (Clubb et al., 2009; Gancz et al., 2010). Response to therapy can be assessed by regular physical examinations, repeated radiographs, serial crop biopsies, and monitoring of fecal shedding of the virus (Clubb et al., 2009; Gancz et al., 2010).

Prophylactic measures

The development of an effective vaccine for PDD has been the object of great hopes among aviculturists since the discovery of ABV (Staeheli et al., 2010); however, there has been little work as yet in this area. Recent studies revealed that seropositivity to ABV is not protective (Payne et al., 2011b) and that ABV escapes recognition by the innate immune system (Reuter et al., 2010); PDD may thus belong to that group of infections whereby immune responses increase severity of disease and vaccination may be contraindicated (Hoppes et al., 2010).

BORNAVIRIDAE VIRUSES

Viruses in the family Bornaviridae are enveloped RNA viruses belonging to the order Mononegavirales. Borna disease virus (BDV) and avian bornaviruses (Hoppes et al., 2010) are the main members of the family. BDV causes epidemics of a unique neurologic syndrome, called Borna disease, which affects primarily horses and sheep in central Europe (Richt et al., 2000; Tomonaga et al., 2002; Gray et al., 2010). ABV was identified in 2008 as the etiologic agent of PDD in psittacine birds (Honkavuori et al.,

2008; Kistler et al., 2008). Both Borna disease and PDD share many attributes, including dysfunction of the central, peripheral and autonomic nervous systems (Stitz et al., 1993; Briese et al., 1999).

Members of family Bornaviridae

Until 2008, the only recognized member of the family Bornaviridae was BDV. Sequence analysis of isolates of BDV obtained from various species over several decades has shown remarkable sequence conservation with only two distinct subtypes described (Dürrwald et al., 2007; Kistler et al., 2008; Nowotny et al., 2000). This is an uncommon feature for RNA viruses. In contrast, twelve avian bornaviruses have been identified to date based on nucleotide and amino acid sequence identity. These include ABV genotypes 1 to 7 (ABV-1 to ABV-7) isolated from psittacine birds (Honkavuori et al., 2008, Kistler et al., 2008, Rinder et al., 2009, Weissenböck et al., 2009a; Rubbenstroth et al., 2012), ABV-canary genotypes 1 to 3 (ABV-C1, ABV-C2 and ABV-C3) isolated from canaries (Serinus canaria) (Weissenböck et al., 2009b; Rinder et al., 2012; Rubbenstroth et al., 2013) and ABV-CG isolated from free-ranging waterfowl (Delnatte et al., 2011). ABV is widespread in the captive psittacine population; it has been described in at least 33 different genera among the order Psittaciformes (Heffels- Redmann et al., 2012). ABV genotypes 2 and 4 are the most predominant psittacine strains circulating worldwide, with ABV-4 being the most common genotype in North America (Rinder et al., 2009; Weissenböck et al., 2009b; Hoppes et al., 2010; Nedorost et al., 2012). Co-infections with several ABV strains (ABV 4 and 7; ABV 2 and 4; ABV 2 and 6) have been occasionally reported and disease severity and lesions did not appear to differ from cases of mono-infections (Weissenböck et al., 2010; Nedorost et al., 2012; Rubbenstroth et al., 2012).

In addition, a putative additional (fourth) genotype of ABV-canary (ABV-C4) originating from a Bengalese finch (Hura striata domestica) was identified through BLAST search (Kato et al., 2010); and a bornavirus signature has also been recently detected from a cDNA library derived from a Gaboon viper (Bitis gabonica) venom gland and was named reptile bornavirus (RBV) (Horie et al., 2010; Fujino et al., 2012).

Further work on the molecular characterization of the bornaviruses is necessary and would help in understanding the evolutionary relationships among them. However, one must be aware of the presence of bornavirus sequences integrated into the genomes of a variety of mammals (Belyi et al., 2010; Horie et al., 2010; Katzourakis et al., 2010), which further complicates the determination of the relationships of the viruses in this group. At the present time, no endogenous bornavirus-like nucleoprotein (EBLN) elements have been found in species of birds whose genomes have been described, the chicken, zebra finch and scarlet macaw.

To illustrate the genetic relationships among bornaviruses, a phylogenic tree using partial N gene sequences is presented as part of Appendix A. Whether these different strains are actually different species, genotypes or genogroups is a matter for debate.

Genomic organization of bornaviruses

Bornaviruses are enveloped spherical viruses measuring 70 to 130 nm in diameter and containing a non-segmented negative-sense, single-stranded RNA genome of approximately 8,900 bases (de la Torre et al., 2006). Unlike other viruses in the order Mononegavirales such as paramyxoviruses and rhabdoviruses, transcription and replication of bornaviral genome occur inside the host-cell nucleus (Cubitt et al., 2001), allowing alternative splicing of the viral mRNA and the use of unusual initiation and termination signals (Briese et al., 1992). This highly uncommon and complicated replication mechanism represents a smart viral evasion strategy that allows the virus to avoid activation of innate immune response receptors in infected cells (Schneider et al., 2007; Habjan et al., 2008). A recent study showed that ABV and BDV use similar strategies to evade the host immune response (Reuter et al., 2010).

The BDV genome encodes six major proteins, including an RNA L-polymerase (L, 190 kDa), a membrane-bound glycoprotein (G, 57 kDa), two isoforms of a nucleoprotein (N, 38 kDa and 40 kDa), a regulatory phosphoprotein (P, 23 kDa), a matrix protein (M, 16 kDa), and a more recently characterized X virus protein (X, 10 kDa) (Hornig et al., 2003; de la Torre et al., 2006; Watanabe et al., 2009). The nucleoprotein and the phosphoprotein together form the viral nucleocapsid and protect the genomic

RNA (Kobayashi et al., 2001, Schneider et al., 2004). Thus the nucleoprotein, and to a lesser extent the phosphoprotein, is the main target of both the humoral and cellular immune response (Tomonaga et al., 2002). A review of BDV genomic organization detailing the function of each protein has been published (Villanueva et al., 2010). Despite the genetic variability among ABV strains, preliminary work on ABV revealed a similar overall organization of the genome (Honkavuori et al., 2008; Kistler et al., 2008; Horie et al., 2012) with the same number and order of genes and the same structure of transcription initiation and termination sites as those observed in BDV.

Minor structural differences have; however, been documented among the bornavirus genomes and could explain the overall lack of antigenic cross reactivity between avian and mammalian bornaviruses (Honkavuori et al., 2008; Villanueva et al., 2010) or among avian bornaviruses (Horie et al., 2012). A notable difference between the various viral genomes appears to affect a regulatory upstream open reading frame (uORF) in the N/X inter-region that serves a critical regulatory function for the genes coding for viral proteins X and P (Kobayashi et al., 2003; Poenisch et al., 2008; Rinder et al., 2009; Wanatabe et al., 2009; Horie et al., 2012; Mirhosseini et al., 2012). Whereas ABV-5 and ABV-CG contain a slightly shorter fragment of this uORF compared to BDV (Payne et al., 2011a; Horie et al., 2012), ABV genotype 1, 2, 3, 4 and 7, and RBV lack the entire regulatory uORF, having a 21-22 nt deletion in this region (Kistler et al., 2008; Rinder et al., 2009; Fujino et al., 2012; Mirhosseini et al., 2012; Rubbenstroth et al., 2012), and ABVs-Canary lacks this uORF but does not have a nucleotide gap at this position (Rubbenstroth et al., 2013). This suggests that different species and genotypes of bornaviruses have different strategies for regulation of expression of X and P proteins.

Pathogenesis of bornavirus infections

Most of the fundamental virology research on Bornaviridae has been done on BDV using rodents as experimental models. A review of the life cycle of BDV, based on experimental studies with BDV on various animal models, has been published (Villanueva et al., 2010). Epidemiological data on ABV and preliminary virology work including viral cultivation and experimental inoculations suggest that ABV shares many

characteristics with BDV and that the pathogenesis of the disease is likely similar (Rinder et al., 2009; Staeheli et al., 2010). However, the understanding of both mammalian and avian bornavirus-related disease remains limited.

BDV exhibits a high tropism for the central nervous system in both natural and experimental hosts. It establishes persistent, non-cytolytic infections of neurons and astrocytes with the functionality of infected cells being not or only marginally impaired (Stitz et al., 1995). Manifestations of natural or experimentally induced disease range from fatal meningo-encephalitis to subtle behavioral changes or asymptomatic persistent infection (Ludwig et al., 2000).

Bornaviruses are thought to spread mainly by cell-to-cell contact in a manner similar to that of the rabies virus; the ribonucleoproteic complexes spread along the neuronal networks inside axons (Carbone et al., 1987; Gosztonyi et al., 1993; Lipkin et al., 2006; Ackermann et al., 2010; Villanueva et al., 2010). In both BDV and ABV- infected cells, viral antigen is found in both the cytoplasm and the nucleus and shows a characteristic speckled immunofluorescence pattern in the nucleus (Herzog et al., 1980; Schwemmle et al., 1998; Rinder et al., 2009; Gray et al., 2010). The presence of abundant viral products in infected cells suggests that the virus must actively suppress apoptosis, possibly via the viral accessory protein X (Poenisch et al., 2009). However, cells persistently infected with BDV or ABV release very few infectious viral particles (Sauder et al., 2003; Rinder et al., 2009; Staeheli et al., 2010), raising questions regarding their mode of transmission in nature. Efficient release of virions may be restricted to some specialized cell types, likely present in the kidney or urinary tract (Sauder et al., 2003; Heatley et al., 2012).

Serologic evidence of widespread exposure to the virus, yet little clinical disease are a common feature of BDV and ABV. While up to 12% of the horses within the affected region in central Europe are seropositive, only a small fraction of these animals actually develop clinical disease and many animals become persistently infected but fail to develop disease (Payne et al., 2012). Similarly, asymptomatic ABV infection has been identified in both experimentally and naturally infected birds and likely plays an important role in the epidemiology of PDD (de Kloet et al., 2009; Gancz et al., 2009;

Kistler et al., 2010; Lierz et al., 2009, Lierz et al., 2010; Villanueva et al., 2010; Heffels- Redmann et al., 2012). Many apparently healthy psittacines carry ABV for prolonged periods (Hoppes et al., 2013). In a recent study, ABV was detected in 40% (12/30) of sampled canary captive flocks in Germany and most of these birds were apparently healthy (Rubbenstroth et al., 2013).

The exact factors that trigger the development of lesions or clinical signs are unknown. Experimentally, Borna disease development varies with host species, age and immune function (Lipkin et al., 2011). Age at infection may determine the rate of progression of disease in ABV-infected birds. Younger birds appear to have a more rapid course of disease progression than older birds (Kistler et al., 2010). Vertical transmission of BDV has been demonstrated in mammals (Okamoto et al., 2003) and is believed to occur in birds with ABV (Lierz et al., 2011; Kerski et al., 2012; Monaco et al., 2012; Rubbenstroth et al., 2013); this route of transmission may also play an important role in the outcome of infection. It is also possible that the ABV genotype affects the form of clinical disease. Although a direct relationship between genotype and virulence has not been formally demonstrated (de Kloet et al., 2009; Gancz et al., 2009; Payne et al., 2011b), a recent study suggests that there may be differences in pathogenicity among ABV isolates with cockatiels infected with ABV-2 showing earlier and more severe clinical symptoms compared to birds infected with ABV-4 (Lierz et al., 2012).

The lesions and symptoms seen with BDV in experimentally-infected rodents are the result of neural invasion by CD8 T-lymphocytes and subsequent cytotoxicity, rather than of virus-inflicted cellular damage (Rott et al., 1988; Stitz et al., 1989; Hallensleben et al., 1998; Matsumoto et al., 2012). Thus, it is generally believed that bornavirus infection related lesions are in large part immunologically mediated (Stitz et al., 1995; Stitz et al., 2002; Schwemmle et al., 2004; Baur et al., 2008). The mechanism by which ABV causes the lesions and clinical syndrome of PDD remains unclear; however, a similar immune-mediated pathogenesis is likely (Gancz et al., 2009).

Several recent communications (Rossi et al., 2008; Pesaro et al., 2011; Rossi et al., 2011; Rossi et al., 2012) have suggested that an autoimmune mechanism (production of auto-reactive anti-ganglioside antibodies), similar to the one observed in Guillain-

Barré syndrome in humans, is involved in the pathogenesis of PDD. The detection of anti-ganglioside antibodies in the sera of PDD affected birds and the successful reproduction of PDD via inoculation of purified gangliosides into healthy cockatiels support this theory. However, it is unclear how significant these antibodies are, whether they develop secondary to tissue damage, and how they contribute to the disease process. Although PDD is very unlikely to be an autoimmune disease per se, transient autoimmune responses occur in some PDD cases and may contribute to the complex pathogenesis of this disease (Hoppes et al., 2010).

Epidemiology of Borna disease virus

Host range

Although usually described in horses and sheep, natural BDV infection or exposure has been reported in a large variety of hosts, including humans, sheep, horses, cattle, cat, raccoons, deer, primates, and zoo animals such as sloths, alpacas, llamas and hippopotami (Carbone et al., 2001; Ikuta et al., 2002; Tomonaga et al., 2002). In addition to this extremely broad natural host range, experimental infections have been described in a wide variety of vertebrates, including chickens, quail, rats, rabbits, cats, shrews, and nonhuman primates (Zwick et al., 1927; Ludwig et al., 1973; Rott et al., 1995). One-day old chicks inoculated intra-cerebrally with brain homogenates from rabbits with Borna disease developed paralysis of legs and wings, had intranuclear Joest-Degen inclusion- bodies in their brains and anti-BDV antibodies in their sera (Ludwig et al., 1973).

The only report of naturally occurring disease attributed to BDV in an avian species was in farmed ostriches in Israel in the early 1990s (Malkinson et al., 1993; Weisman et al., 1994a; Malkinson et al., 1995). A paretic syndrome associated with high morbidity and mortality was attributed to BDV based on the presence of BDV specific antigen in the brains of affected birds as detected by ELISA. The syndrome could be transferred to naïve birds by intramuscular injection or oral application of brain homogenates derived from diseased animals (Ashash et al., 1996). Although the pathologic lesions in these ostriches were quite different from those seen in ABV- infected parrots or waterfowl, the possibility of ABV rather than BDV as being the cause

of the disease must remain open. Unfortunately, no tissues from affected animals are available for testing, so the actual etiology remains unclear (Hoppes et al., 2010).

BDV sequences were recovered from feces of wild asymptomatic mallards (Anas platyrhynchos) and jackdaws (Corvus monedula) in Sweden (Berg et al., 2001). Brain homogenates from several species of wild aquatic birds in Syria and Egypt collected during an outbreak of encephalitis in horses in the 1950's were experimentally inoculated into rabbits and kittens and encephalitis resulted. It was later hypothesized that these birds may have been infected with BDV (Daubney et al., 1967). BDV may be the etiological agent of a paralytic lethal syndrome in cats in central Europe and Scandinavia (Lundgren et al., 1992) and some authors hypothesized that cats were infected by predating BDV-infected birds (Berg et al., 2001).

The broad host spectrum of BDV has raised the question of whether this agent may also affect humans. Several publications suggested this (Rott et al., 1985; Weisman et al., 1994b; Bode et al., 1995) as well as an association between BDV and various neuropsychiatric disorders (Bode et al., 2003). However, the results of a recent blinded case–control study (Hornig et al., 2012), concern over the specificity in the serological tests used (Wolff et al., 2006) and the possibility of laboratory contamination of samples (Dürrwald et al., 2007) make these hypotheses very debatable.

Origin and possible reservoirs

Borna disease was first described in the late 18th century in Borna, Germany (Richt et al., 2000; Tomonaga et al., 2002; Villanueva et al., 2010). Although endemic to a restricted region of central Europe, BDV may have a broader geographic range than initially thought with sporadic cases reported in South Africa, Israel, Australia and Japan (Richt et al., 1997; Villanueva et al., 2010).

Several unique epidemiologic features point towards the existence of a natural reservoir population for BDV that is different than the final hosts, and free-living rodents have been the prime suspects (Jordan et al., 2001; Hornig et al., 2003; Dürrwald et al., 2006). In addition, clear clustering of different genetic strains of BDV within the endemic

area (Kolodziejek et al., 2005) and apparent lack of direct spread among livestock (Staeheli et al., 2000) suggest that the occurrence of multiple introductions of genetically stable bornaviruses is a more likely scenario than the spread of a single virus from a single point of origin.

Recent publications suggest that the natural reservoir host of BDV is the bicoloured white-toothed shrew (Crocidura leucodon), a small insectivore. This shrew was found in an area where BDV was endemic, harbored the virus in many tissues without histopathological signs of infection, and virus from the shrew was almost identical to that from diseased horses. Thus, it is possible that the horses acquired the infection by ingesting or inhaling virus from pastures contaminated via saliva, feces or urine from persistently infected possibly immunotolerant shrews (Hilbe et al., 2006; Puorger et al., 2010). Seropositive bank voles (Myodes glareolus) were found in Finland, where the bicoloured white-toothed shrew is not present (Kinnunen et al., 2007). Although no other reservoirs have been formally demonstrated, the broad host range of BDV and the restricted geographical distribution of these insectivores suggest that multiple genetically distinct strains of BDV might exist in other species of wild animals (Staeheli et al., 2010).

Although the role of wild birds in the maintenance and transmission of BDV in Europe remains unknown, there is evidence that avian vectors may play a role in maintaining and disseminating this disease (Payne et al., 2012). The epidemiology of Borna disease shows a certain seasonality (occurring mainly during spring and summer) and disease tends to occur at low altitudes, and preferentially along coast or river valleys that correspond to migratory flyways or sites preferred by birds (Dürrwald et al., 1993; Richt et al., 2000; Teplitsky et al., 2003). The presence of BDV sequences from the feces of wild birds in Sweden, as described above, may also support the possibility of wild birds as a natural reservoir for BDV (Berg et al., 2001).

PATHOGENESIS OF AVIAN BORNAVIRUS INFECTION

Since the identification of ABV in 2008, numerous publications have reported the presence of ABV in PDD-positive birds or in birds exposed to PDD cases (Gancz et al., 2010; Heffels-Redmann et al., 2011). Molecular techniques and, to a lesser extent, cultivation of the virus, have started to open the way for more sophisticated investigations into infection with and the pathogenesis of disease associated with this virus. Large scale infection trials, as would be carried out in species such as domestic poultry, have not yet been undertaken.

Experimental inoculation

Although the first identifications of ABV in PDD-affected parrots suggested that ABV was a plausible cause of PDD, full proof of a causal relationship using Koch’s postulates required isolation of the agent from infected birds; its propagation in culture; and manifestation of the disease after reintroduction of the isolate into a susceptible host. These postulates were fulfilled when cockatiels (Nymphicus hollandicus) and Patagonian conures (Cyanoliseus patagonus) were inoculated with cultured ABV genotype 4 via intramuscular (IM), intracerebral and intravenous (IV) routes and disease resulted (Gray et al., 2010; Payne et al., 2011b; Lierz et al., 2012; Piepenbring et al., 2012). PDD has also been reproduced in cockatiels inoculated with brain homogenate containing ABV genotype 4 via multiple routes (intramuscular, intraocular, intranasal, and oral) (Gancz et al., 2009) and with cultured ABV genotype 2 via oral, IM, IV and intracerebral routes (Mirhosseini et al., 2011; Lierz et al., 2012). The results of these experiments provide overwhelming support for the causal relationship between ABV and PDD in psittacine birds.

More recently, successful inoculation of canaries with cultured ABV-C2 via IM, SC, oral and oculo-nasal routes has been documented: shedding of virus, seroconversion and re-isolation of ABV-C2 from the brains of inoculated birds occurred but no clinical signs or macroscopic lesions attributable to ABV infection were observed in any of the

inoculated birds and histopathological changes consistent with ABV were minimal (Rubbenstroth et al., 2013).

Specific pathogen free (SPF) mallard ducklings inoculated with cultured psittacine ABV genotype 4 shed virus in feces and seroconverted within 3 weeks, but inoculation did not result in disease, based on a lack of clinical signs and of histopathological lesions on necropsy 8 months after inoculation (Gray et al., 2009; Hoppes et al., 2010).

Rats and mice were challenged with ABV-4, but no clinical signs or lesions resulted (Hoppes et al., 2013). Currently, there is no evidence that ABV can infect or cause disease in mammals.

Viral distribution in tissues

The distribution of ABV in the tissues and organs of PDD-positive birds has been studied using immunohistochemistry (IHC), Western blot, and reverse transcription- polymerase chain reaction (RT-PCR) (Honkavuori et al., 2008; Gancz et al., 2009; Kistler et al., 2010; Lierz et al., 2009; Ouyang et al., 2009; Rinder et al., 2009; Weissenböck et al., 2009a; Weissenböck et al., 2009b; Lierz et al., 2010; Raghav et al., 2010; Reßmeyer et al., 2010; Wunschmann et al., 2011; Rubbenstroth et al., 2013). ABV exhibits a high tropism for neuroectodermal cells including neurons, astroglia, and ependymal cells of the central nervous system; neurons of the peripheral nervous system; and adrenal cells. Although necrosis of Purkinje cells has been described in cases of PDD (Shivaprasad et al., 1995), IHC consistently fails to demonstrate that Purkinje cells themselves are infected. Nearby Bergmann glial cells are; however, heavily infected and it is likely that these cells are required to support Purkinje cell function and viability (Ouyang et al., 2009). In most affected birds, ABV is also present in many other organs, suggesting the possibility of transient viremia. A pancytotropic viral distribution has been described in acute symptomatic infections involving ABV genotypes 2 and 4 (Kistler et al. 2010; Payne et al. 2011b; Wünschmann et al. 2011).This broad tissue and cell tropism of ABV is strikingly different from classical BDV, where the virus shows a marked and almost exclusive neurotropism (Hoppes et al., 2013).

Whether the tissue distribution of ABV infection differs among birds with and without clinical evidence of disease (PDD) is not clear. While some authors report that RNA detection appears restricted to nervous system tissue in asymptomatic birds (Lierz et al., 2010), others suggest that ABV is widely distributed regardless of clinical presentation (Wunschmann et al., 2011).

Shedding and transmission

It has been demonstrated that birds shed ABV in their urofeces and that this shedding is intermittent and unpredictable for experimentally and naturally infected birds (Raghav et al., 2010; Villanueva et al., 2010; Payne et al., 2011a; Payne et al., 2011b). Psittacine birds may shed detectable (by RT-PCR) amounts of ABV on a single occasion and then test negative for months. The viral shedding is not restricted to the urofeces and ABV RNA has also been detected in swabs from the nares, the choana and feathers; and in the air of infected aviaries (Hoppes et al., 2010; de Kloet et al., 2011). Recent research has in fact shown that the greatest amount of virus may actually be excreted in urine (Heatley et al., 2012). This may explain why a fecal sample can be negative whereas a cloacal swab taken at the same time is positive (Dorrestein et al., 2010).

Kistler et al. detailed the spread of ABV and PDD through an aviary after introduction of an adult bird with (eventually) fatal PDD (Kistler et al., 2010). Epidemiological investigations of outbreaks of proventricular dilatation disease in psittacine aviaries and the presence of virus in feces and urine suggest urofecal-oral transmission of ABV as the primary method of spread (Hoppes et al., 2010). This was confirmed in an unintentional study, where mallards were infected with ABV by eating infected cockatiel droppings (Hoppes et al., 2010).

In addition, there is increasing evidence for in ovo infection of ABV. Vertical transmission of BDV has been demonstrated in mice (Okamoto et al., 2003) and is strongly suspected in horses (Hagiwara et al., 2000). ABV antigens are present in the testes and ovaries of infected parrots (Raghav et al., 2010; Payne et al., 2011b), infected domestic duck eggs used for cell culture have been identified (Hoppes et al., 2013) and ABV RNA was detected in eggs, embryos, and hatchlings of various psittacine species

and canaries (Lierz et al., 2011; Kerski et al., 2012; Monaco et al., 2012; Rubbenstroth et al., 2013). There are three possible routes of infection of eggs: penetration of the shell by virus after uro-fecal contamination; contamination of the albumin by infected oviductal secretions during oviposition or, more likely, ABV-infected ova or sperm (Monaco et al., 2012). ABV-positive samples included albumin, yolk, and embryonic brain, liver and eye tissue from fertilized eggs containing either non-viable or developing early- and late- stage embryos; and brain, feather and blood from hatchlings raised by infected parents. Whether ABV infection was the cause of the embryonic and hatchling deaths observed in these studies remains unclear; and whether the hatchlings were truly infected in ovo or were infected by exposure after hatch is uncertain.

Seroconversion

Antibodies to ABV have been identified in the serum of PDD-affected birds as well as in asymptomatic birds (de Kloet et al., 2009; Herzog et al., 2010; Villanueva et al., 2010; de Kloet et al., 2011; Rinder et al., 2011; Heffels-Redmann et al., 2012). Seroconversion occurs as early as 6 days post infection in experimentally infected cockatiels (Lierz et al., 2012). As in mammals, most of the antibodies detected in ABV- infected or exposed birds seem to be non-neutralizing and directed toward the non- structural phosphoprotein, nucleocapsid protein, matrix protein and X virus protein (de Kloet et al., 2009; de Kloet et al., 2011; Heffels-Redmann et al., 2011; Villanueva et al., 2010). The relative concentrations of these antibodies can vary widely in the individual sera of infected psittacine birds (Kerski et al., 2012). Antibodies against the viral glycoprotein, which are considered neutralizing in mammals (Stitz et al., 1998), and against ABV RNA polymerase have not been detected even in long term infected birds (Kerski et al., 2012). Antibodies were also found in egg yolks and serum of late-stage embryos originating from ABV infected sun conures, suggesting the possibility of maternal antibody transfer. The IgY composition of egg yolk in these embryos reflected the IgY composition of the maternal serum (Kerski et al., 2012).

The detection of ABV-antibodies in serum, the presence of ABV-RNA in feces and the distribution of ABV in tissues of the same animal do not always correlate

(Enderlein et al., 2011; Heffels-Redmann et al., 2011; Rubbenstroth et al., 2013). Some ABV-infected birds do not develop a detectable antibody response (Herzog et al., 2010). Sudden seroconversion sometimes immediately precedes the onset of clinical disease (Hoppes et al., 2013). However, for some authors, there seems to be a positive correlation between the level of antibodies produced and the development of disease (Hoppes et al., 2013).

The presence of antibodies to ABV does not appear to prevent the development of clinical disease and pathological lesions. Heffels-Redmann et al. (2012) reported that various psittacine birds shed the virus, developed PDD and died despite the presence of anti-ABV antibodies, sometimes in very high titer, suggesting that a humoral antibody response is not protective. This is similar to the results of BDV studies in immunocompetent rats (Narayan et al., 1983). In cockatiels, Payne et al. demonstrated that seropositive birds naturally infected with ABV-4 and showing no clinical evidence of disease developed unusually severe lesions of PDD subsequent to inoculation with a related virulent strain of ABV-4 (Payne et al., 2011b).

DIAGNOSIS OF AVIAN BORNAVIRUS INFECTION

Until the discovery of ABV as the etiologic agent of PDD, the definitive diagnosis of PDD relied on the identification of histological lesions in biopsy or post-mortem samples. More recent approaches to diagnostic testing have focused on identifying virus in secretions, excretions, or tissue; and in assessing exposure to the virus using serology.

Virus culture

Viral isolation and propagation remain the gold standard in virology and the only definitive method of demonstrating the presence of a viable infectious agent in tissues or excretions from a host.

Prior to the identification of ABV, numerous researchers tried to identify a causal agent for PDD through tissue cultures using various cell lines with no success (Clark et al., 1984; Daoust et al., 1991; Berhane et al., 2004). The absence of cytopathic effects was considered to indicate a lack of virus but, in retrospect, these attempts to propagate the virus may have been successful but were hampered by a lack of detection methods as ABV is very difficult to visualize by electron microscopy in cell cultures (Gray et al., 2010; I Tizard, personal communication). In 1996, Gough et al. did demonstrate cytopathic effects in macaw embryonic cell lines inoculated with virus-like particles derived from the feces of parrots affected with PDD (Gough et al., 1996). Whether these virus-like particles were ABV was never proven.

The identification of ABV using cutting-edge molecular technologies has provided a means of reliably identifying the virus in cell culture, and several authors describe its propagation (Gray et al., 2010; Hoppes et al., 2010; Rubbenstroth et al., 2012). Various genotypes of ABV originating from psittacine, canary and waterfowl samples have been successfully grown in duck cell lines (duck embryo fibroblasts (DEF) (Gray et al., 2010; Hoppes et al., 2010; Rubbenstroth et al., 2013) and quail cell lines (quail fibroblast cell line CEC32 and quail skeletal muscle cell line QM7) (Rinder et al., 2009; Herzog et al., 2010; Rubbenstroth et al., 2012; Rubbenstroth et al., 2013). Although cytopathic effects were not detected, progressive increases in viral antigen were demonstrated by either Western blot (Gray et al., 2010), IHC or indirect immunofluorescence assays (Rinder et al., 2009; Herzog et al., 2010; Rubbenstroth et al., 2013). Characteristic speckled intranuclear fluorescence similar to that considered diagnostic for mammalian BDV, so called Joest-Degen inclusion bodies (Herzog et al., 1980; Schwemmle et al., 1998), has been observed with ABV in culture (Rinder et al., 2009; Gray et al., 2010).

Virus titration has also been used to demonstrate and quantify viral replication and compare the efficiency of different cell lines (Rubbenstroth et al., 2012; Rubbenstroth et al., 2013). In a recent comparative study (Rubbenstroth et al., 2012; Rubbenstroth et al., 2013), the quail fibroblast CEC32 appears to outperform other avian cell lines, with isolation of ABV from organs, cloacal and pharyngeal swabs, and the buffy coat of blood samples. Cell culture characteristics vary between ABV strains and

genotypes (Rubbenstroth et al., 2012), and viral culture works best on freshly obtained tissues (Hoppes et al., 2013). Given that domestic duck eggs have been shown to be naturally infected with ABV on several occasions (Hoppes et al., 2013), all cell lines, especially DEF cell lines, should be screened by RT-PCR prior to use for viral isolation.

The use of chicken cell lines (chicken fibroblasts DF-1, and chicken hepatocytes LMH) has resulted in inconsistent results (Hoppes et al., 2010; Rinder et al., 2009; Rubbenstroth et al., 2012). Only one research group (Rubbenstroth et al., 2013) has reported growing ABV in a cell line of mammalian origin, the VERO cell. This supports the suggestion that ABV may be unable, or poorly able, to infect mammals but the topic requires further investigation (Rubbenstroth et al., 2013).

Viral isolation is not currently used as a screening tool for ABV infection, but rather as a source of virus for research activities such as sequencing of the viral genome (Mirhosseini et al., 2012), experimental animal inoculation studies and development of new molecular techniques and serologic assays. However, because of the extensive genetic heterogeneity of ABV, classical virus isolation may represent a useful diagnostic option, particularly for the detection of new bornavirus genotypes (Rubbenstroth et al., 2012; Rubbenstroth et al., 2013).

Reverse Transcriptase Polymerase Chain Reaction

Both gel-based conventional RT-PCR and real time RT-PCR have been developed for the identification of ABV. Various degenerate and non-degenerate primers have been used that target the nucleocapsid N gene, the matrix M gene, the phosphoprotein P gene, the polymerase L gene and sometimes ABV segments not entirely within a single protein coding gene (Honkavuori et al., 2008; Kistler et al., 2008; Gancz et al., 2009; Lierz et al., 2009; Rinder et al., 2009; Dorrestein et al., 2010; Kistler et al., 2010; Horie et al., 2012; Kerski et al., 2012). In several studies, primers were specifically designed to better target certain ABV genotypes, including ABV-canary (Rubbenstroth et al., 2013) and ABV-CG (Delnatte et al., 2013). In psittacine birds, assays for the M and N genes appear to have a similar high sensitivity, whereas RT-PCR for L and P genes seems generally less accurate (Gancz et al., 2010; Raghav et al., 2010).

Designing primers able to detect a broad range of ABV genotypic variants has been challenging (Rinder et al., 2011; Rubbenstroth et al., 2013). As mentioned earlier, whereas BDV sequences show a remarkable homogeneity, ABV displays an extensive genetic heterogeneity with at least 12 different genotypes (Kistler et al., 2008; Rinder et al., 2009; Weissenböck et al., 2009a; Weissenböck et al., 2009b; Rubbenstroth et al., 2012; Rubbenstroth et al., 2013), making molecular diagnostic tests more complex to design and their results more difficult to interpret. For example, in one study, RT-PCR using primers targeting the N gene amplified the canary strain of ABV, whereas repeated amplification attempts using primers targeting the M gene remained negative (Weissenböck et al., 2009b; Rubbenstroth et al., 2013). It is possible and even likely that currently used primers are not able to detect distantly related variants or as yet unknown genotypes.

As described previously, the tissue distribution of ABV is broad and viral particles are shed via various routes. RT-PCR can thus theoretically be used on a variety of clinical samples and necropsy materials. The most consistently infected tissues, and thus the most reliable samples for post-mortem diagnosis, include brain, proventriculus, ventriculus, adrenals and vitreous of the eye (Honkavuori et al., 2008; Gancz et al., 2009; Lierz et al., 2009; Rinder et al., 2009; Weissenböck et al., 2009b; Raghav et al., 2010; Weissenböck et al., 2010; Wunschmann et al., 2011; Hoppes et al., 2013). Specimens that are commonly recommended for ante-mortem detection of ABV RNA include crop tissue, blood, choanal and cloacal swabs, feces (Gancz et al., 2010) and, more recently, calami of plucked chest contour feathers (de Kloet et al., 2011). It is interesting that recommendations vary greatly among laboratories (de Kloet et al., 2011, Avian Biotech website). RT-PCR can also be used to identify ABV in formalin-fixed paraffin embedded (FFPE) tissues (Weissenböck et al., 2009b; Last et al., 2012; Nedorost et al., 2012; Delnatte et al., 2013) allowing the testing of birds for which fresh-frozen tissues are not available.

Sample selection, collection method and transport medium used can influence the results of RT-PCR assays (Dorrestein et al., 2010; Hoppes et al., 2013). Systematic comparisons among samples and techniques have not been conducted. ABV detection appeared to be less sensitive from blood samples as compared to cloacal swabs (de Kloet

et al., 2009; Gancz et al., 2009; Lierz et al., 2009; de Kloet et al., 2011; Rubbenstroth et al., 2012). Whereas some authors report no apparent difference in results between swabs of fresh droppings or of the cloaca (Lierz et al., 2009; Hoppes et al., 2010), others report that cloacal swabbing is the more sensitive technique (Dorrestein et al., 2010).

Despite being a highly sensitive technique per se, RT-PCR can produce false negative results if primers are not appropriate, if the virus was not present in the specific tissue at the time of sample collection, or if the viral copy number is extremely low (Kistler et al., 2008; Gancz et al., 2010). Intermittent shedding of virus in urine and feces is perhaps the most significant situation where false negatives may occur. The testing of a single dropping from a single bird is of limited usefulness; pooling multiple droppings from a single bird over several days or samples from multiple birds in an aviary increases test sensitivity. Until we better understand the pathogenesis of ABV, simply the presence of virus should not be used to diagnose the clinical/pathological syndrome of PDD. It is likely that birds may carry and perhaps shed the virus without ever developing any form of clinical disease (Hoppes et al., 2013).

Sequence analysis

The recognition of an increasing number of ABV genotypes makes genome sequencing a critical component in the diagnosis and management of ABV infection and of PDD, as well as in our understanding of ecology, evolution and pathogenesis of these diverse bornaviruses. Genome sequencing is particularly important when studying the epidemiology of the spread of ABV among collections and/or geographically, and to identify new circulating strains of the virus (Weissenböck et al., 2009a) or mixed infection with several ABV strains (Weissenböck et al., 2010; Nedorost et al., 2012; Rubbenstroth et al., 2012). Whether there is a correlation between ABV genotype and pathogenicity has not been determined. As we learn more, specific identification of the ABV strain affecting a patient may have clinical implications (Lierz et al., 2012).

Ideally, the entire genome of each variant should be sequenced, as was done recently for ABV-1 (Mirhosseini et al., 2012). However, because of financial considerations, the recovery of partial nucleotide sequences (usually RT-PCR products)

is most commonly achieved. After sequence recovery, a BLAST search is generally performed allowing the comparison of nucleotide sequences with those available in international databases, such as GenBank, and the determination of the ABV genotype. Specialized computer programs are then able to calculate probable phylogenetic relationships if necessary.

Immunohistochemistry

Immunohistochemistry has been used primarily to describe the tissue distribution of ABV antigen in necropsy tissues as part of research investigations (Ouyang et al., 2009; Rinder et al., 2009; Weissenböck et al., 2009a; Weissenböck et al., 2009b; Raghav et al., 2010; Reßmeyer et al., 2010; Wunschmann et al., 2011). However, IHC can also be used as a diagnostic tool to identify ABV in tissues, especially when histological lesions are mild or equivocal, and as a post-mortem confirmatory test.

Reagents used for IHC include antibodies directed against recombinant ABV nucleocapsid protein (Ouyang et al., 2009), polyclonal antiserum raised against ABV nucleocapsid protein (Raghav et al., 2010; Wunschmann et al., 2011), as well as polyclonal antiserum raised against the BDV phosphoprotein (Rinder et al., 2009; Weissenböck et al., 2009a; Weissenböck et al., 2009b; Reßmeyer et al., 2010). While some authors mention an overall lack of cross-reactivity between mammalian and avian sequences (Villanueva et al., 2010, Honkavuori et al., 2008), others report a high degree of cross-reactivity among ABV variants and between ABVs and BDV (Rinder et al., 2009; Weissenböck et al., 2009a; Weissenböck et al., 2009b; Herzog et al., 2010; Reuter et al., 2010; Payne et al., 2011a; Rubbenstroth et al., 2012; Rubbenstroth et al., 2013).

Immunohistochemical staining can be present in both the nuclear and cytoplasmic compartments of the cell, and has been demonstrated in neural and extraneural tissues. Cross-reactivity of the primary antibody with cellular proteins can complicate the interpretation of the immunostaining (Weissenböck et al., 2009b; Raghav et al., 2010). Raghav et al. defined the presence of a moderately intense, diffuse, intracytoplasmic staining accompanied by intranuclear staining in neurons and certain epithelial cells as specific staining for ABV antigen (Raghav et al., 2010; Delnatte et al., 2013).

Serology

Serological tests aim to detect the presence of specific antibodies to ABV in the bloodstream indicating previous exposure to the virus. The two major immunogenic proteins of bornaviruses are the nucleoprotein N and the phosphoprotein P (Tomonaga et al., 2002; Gancz et al., 2010). Historically, serological assays have not been very convincing tests for diagnosing BDV infections of mammals (Staeheli et al., 2010). However, unlike BDV, ABV shows a broad tissue tropism and ABV antigens are abundantly present in affected organs (Gancz et al., 2009; Rinder et al., 2009; Weissenböck et al., 2009a; Weissenböck et al., 2009b; Raghav et al., 2010), which suggest that the immune response to ABV may be stronger than the one to BDV.

To date, serologic assays described for the identification of antibodies to ABV include indirect immunofluorescence (Herzog et al., 2010; Heffels-Redmann et al., 2012), Western blot (de Kloet et al., 2009; Hoppes et al., 2010; Villanueva et al., 2010; de Kloet et al., 2011) and indirect ELISA (de Kloet et al., 2009; Dorrestein et al., 2010; de Kloet et al., 2011; Rinder et al., 2011; Rubbenstroth et al., 2013). Various primary sources of antigen have been used to design serological tests for ABV including BDV infected cell culture (Herzog et al., 2010; Heffels-Redmann et al., 2012), recombinant BDV N protein (de Kloet et al., 2009; Lierz et al., 2009; Dorrestein et al., 2010) and BDV P protein (Lierz et al., 2009), ABV-infected brain homogenates (Hoppes et al., 2010), ABV-infected cell culture (Villanueva et al., 2010; Herzog et al., 2010) and recombinant ABV N protein (Lierz et al., 2009; de Kloet et al., 2011; Rinder et al., 2011; Kerski et al., 2012; Rubbenstroth et al., 2013), ABV P protein (Lierz et al., 2009; de Kloet et al., 2011; Kerski et al., 2012), ABV M protein (de Kloet et al., 2011) and ABV X protein (Kerski et al., 2012). Secondary antibodies used for ELISA work include goat anti-bird IgG (de Kloet et al., 2009; Dorrestein et al., 2010; de Kloet et al., 2011) and rabbit anti-canary IgG (Rubbenstroth et al., 2013) conjugated to either horseradish peroxidase or alkaline phosphatase.

Determining a cut-off value for seropositivity is particularly challenging given the difficulty in identifying true control groups of uninfected and infected birds. A cut-off value of 3 standard deviations above the mean absorbance value (optical density (OD)

readings) of sera from a negative control group is a common practice (Classen et al., 1987; Ojkic et al., 2003) and was chosen for ABV specific ELISA designed for psittacine birds and canaries (de Kloet et al., 2011; Rubbenstroth et al., 2013). Assessment and comparison of the specificity and sensitivity of these different serological tests are very difficult, especially due to the complex and poorly understood epidemiology of the disease, the differences in methodology, the absence of a gold standard for identifying infection in live birds, and the relatively poor correlation between the presence of antibodies, fecal shedding of ABV, pathology and clinical disease. These parameters have been published for several assays: a Western blot using a lysate of ABV-infected cultured cells as the primary antigen showed a sensitivity of 90% and a specificity of 82% (Hoppes et al., 2010; Villanueva et al., 2010) and an ELISA using a recombinant ABV N protein as antigen showed a 75% sensitivity and 75% specificity for the diagnosis of PDD in affected birds (de Kloet et al., 2011).

Due to the presence of anti-ABV antibodies in asymptomatic ABV-positive macaws (de Kloet et al., 2009; Lierz et al., 2009) and to the occurrence of maternal transfer of antibodies (Kerski et al., 2012), serology can lead to false positive results in diagnosing PDD. Whether a bird can "clear" the infection, and thus be seropositive but not infectious has been hypothesized (Dorrestein et al., 2010) but has never been formally proven. In addition, some birds without detectable serum antibodies have been shown to shed the virus in feces (Villanueva et al., 2010; Lierz et al., 2010; Heffels- Redmann et al., 2011) (see previous paragraph on seroconversion).

Summary statement: diagnosis of proventricular dilatation disease versus avian bornavirus infection

With the rapid development of new diagnostic tests for ABV infection, it is important to remember that diagnosing PDD and diagnosing ABV infection are different. A diagnosis of PDD requires the presence of clinical signs and/or somewhat specific pathological lesions. A diagnosis of ABV infection requires identification of virus in excretions, secretions, or tissue samples. The significance of anti-ABV antibodies to the diagnosis of either current PDD or ABV infection remains to be clarified.

Each of the tests described above has its advantages and its shortcomings. Intermittent shedding, subclinical infection, asymptomatic carriers, inconsistent seroconversion and technical limitations make the interpretation of these tests challenging (Heffels-Redmann et al., 2012). Histopathology remains the gold standard to diagnose PDD, whereas serological tests and RT-PCR are useful non-invasive tools for the diagnosis of ABV infection in the live bird. Detection of viral RNA or antibodies to ABV provides evidence of ABV infection or exposure but may not necessarily correlate with clinical disease and thus does not differentiate between patients with PDD, asymptomatic shedders and previously exposed birds. A recent study showed that in general, birds with a high ABV load in their crop and cloaca combined with the presence of high levels of antibodies had the highest risk of developing disease (Heffels-Redmann et al., 2012). The optimal screening protocol for PDD and ABV in psittacine flocks is yet to be determined. Finally, with an ever-increasing number of ABV strains identified, tests using more highly conserved primers should be preferred in order to encompass a maximum of genotypes and avoid underestimation of the prevalence of infection.

In conclusion, the recent development of specific molecular and serologic assays for detection of ABV offers promise to avian veterinarians dealing with clinical situations. However, test results should be interpreted cautiously given our still-limited knowledge of this novel virus and the inherent limitations of the diagnostic techniques available.

AVIAN BORNAVIRUS INFECTION IN WATERFOWL SPECIES

Reports of PDD have largely been restricted to captive psittacines, but prior to the identification of ABV there were descriptions of a “PDD-like syndrome” and “BDV-like particles” from several other avian orders (see first section), including waterfowl species.

In 1991, Daoust et al. (1991) described two free-ranging Canada geese showing a PDD-like syndrome, including emaciation, severe proventricular dilatation associated

with food impaction and moderate cloacal distension. Common causes for proventricular dilatation in waterfowl, such as chronic lead poisoning and mechanical obstruction (e.g., soybean ingestion), were ruled out. Histology revealed a lymphoplasmacytic encephalomyelitis and ganglioneuritis involving the submucosal and myenteric plexuses and their autonomic ganglia along the upper gastrointestinal tract. Attempts to identify a causative agent, including viral culture in different cell lines and inoculation of young mice and chicken embryos, were not successful (Daoust et al., 1991).

Berg et al. (2001) amplified fragments of the BDV p24 and p40 genes from feces originating from two mallard ducks (Anas platyrhynchos) and one jackdaw (Corvus monedula) at a bird pond, suggesting that wild birds could be a potential natural reservoir for BDV (Berg et al., 2001). Sequencing identified three strains of BDV exhibiting 97- 99% identity with BDV horse isolates at both nucleic acid and amino acid levels. These sequences were distinct from all ABV genotypes identified in psittacines and canaries. The genetic similarities between these strains and the mammalian virus, and the fact that these samples were collected from the ground, raised the possibility that these putative avian sequences have resulted from environmental contamination (Kistler et al., 2008).

First identification of avian bornavirus in wild waterfowl

ABV was first identified in free-ranging waterfowl in 2009 at the University of Guelph, Canada. The discovery of ABV as a candidate etiologic agent of PDD in 2008 (Honkavuori et al., 2008; Kistler et al., 2008) and the fact that several waterfowl cases from the Toronto Zoo had histological lesions resembling PDD (DA Smith, personal communication) led to the hypothesis that ABV might be associated with similar pathologic changes in wild waterfowl. A retrospective evaluation of necropsy reports from wild Canada geese and trumpeter swans that were euthanized or found dead in southern Ontario between 1992 and 2011 was carried out to select cases with a clinical history, gross lesions or neuropathology consistent with PDD and without a specific etiologic diagnosis (Smith et al., 2010; Delnatte et al., 2011).

Immunohistochemistry using rabbit polyclonal antiserum raised against ABV nucleocapsid protein and/or conventional gel based RT-PCR using primers targeting the

ABV nucleocapsid protein gene (originally designed for ABV identification in parrots) revealed the presence of ABV in the brain of more than 50% of the selected cases. The majority of the infected birds had shown clinical signs including esophageal dysfunction, emaciation and neurological abnormalities prior to death or euthanasia, and all infected cases showed non-suppurative inflammation in the central, peripheral and autonomic nervous systems in various tissues on histology. These findings confirmed the association of ABV infection with clinical and histopathological lesions consistent with PDD in free- ranging waterfowl in Canada (Smith et al., 2010; Delnatte et al., 2011). The details of this retrospective study are covered in the chapter 2 of this manuscript.

Subsequent identification of avian bornavirus in waterfowl species

After the first identification of ABV in waterfowl (Smith et al., 2010; Delnatte et al., 2011), several researchers subsequently reported the presence of the virus in asymptomatic or diseased waterfowl.

We demonstrated that ABV was present in other Canadian provinces: archived tissues from several geese originating from Prince Edward Island (Daoust et al., 1991) and from a goose found dead in Québec – all of them showing consistent neuropathology - were positive for ABV on RT-PCR and IHC (Delnatte et al., 2013). More recently, ABV has been detected in a Canada goose flock in California with many birds dying as a result of encephalitis (HL Shivaprasad, personal communication).

RT-PCR using primers targeting the matrix protein gene was used to screen oropharyngeal / cloacal swabs and brain samples from wild Canada geese and mute swans (Cygnus olor) in the United States. Brain samples were collected by either dissection of the calvarium or syringe aspiration from the foramen magnum with no apparent difference of sensitivity between the two techniques (Guo et al., 2012). ABV sequences were detected in 2.9% (12 out of 409) goose swabs collected from various U.S. states and in 44% (11 out of 25) of the goose brains collected from two urban flocks in New Jersey (Payne et al., 2011a); and in 6.4% (14 out of 219) swabs from the mute swans and in 23% (45 out of 197) of the swan brains (Guo et al., 2012). All sequences were similar or closely related to the waterfowl ABV described previously (Smith et al.,

2010; Delnatte et al., 2011; Payne et al., 2012a; Guo et al., 2012). The vast majority of these samples were obtained from birds described as healthy; however, no clinical examination or necropsies were performed.

The difference between the detection rate in brain and swabs can be explained by the fact that urofecal shedding is likely intermittent in waterfowl as it is in parrots (Villanueva et al., 2010; Payne et al., 2011a; Payne et al., 2011b; Guo et al., 2012). These results also showed that viral prevalence was not uniform across the continent, and that infections could be clustered within flocks. For example, 20–42% of birds within selected flocks in Kansas, New Hampshire and New York were RT-PCR-positive (Payne et al., 2012).

ABV infection has been subsequently shown to be widespread in wild birds across the United States where the virus has been identified in apparently healthy wild snow geese (Chen caerulescens), Ross’s geese (Chen rossii), northern shovelers (Anas clypeata), northern pintails (Anas acuta), gadwalls (Anas strepera), mallards (Anas platyrhynchos), American widgeons (Anas americana), redheads (Aythya americana), herring gulls (Larus argentatus), ring-billed gulls (Larus delawarensis), and laughing gulls (Larus atricilla) using oropharyngeal / cloacal swabs and brain samples (Payne et al., 2011a; Guo et al., 2012, Payne et al., 2012). During the winter of 2011-2012, ABV was found in approximately 10% of 212 hunter-killed ducks of various species in Texas, with the highest prevalence in gadwalls and widgeons, and the lowest is pintails (Hoppes et al., 2013). The apparent prevalence in these waterfowl varied between 6.5% and 50% depending on the species and location sampled. In addition, several domestic duck eggs from two different commercial hatcheries in Texas have been shown to be infected with ABV-CG (Hoppes et al., 2013).

Molecular characterization of waterfowl avian bornavirus

Smith et al. (2010) reported that partial nucleotide sequences of ABV-N genes isolated from several Canada geese in Ontario showed 100% identity on nucleotide and amino acid levels among themselves, but the N gene sequences clustered separately from those found in psittacines and passerines, and from BDV (Delnatte et al., 2011). This

separate genotype was named ABV-CG. This is in agreement with the recent papers describing ABV in the brains of wild waterfowl from various locations in the USA (Payne et al., 2011a; Guo et al., 2012). Brain isolates from wild Canada geese and wild mute swans were cultured in duck embryo fibroblasts and pair-wise comparisons of partial N, M, X, P, and G sequences showed 95.5 to 100% nucleotide identity for all regions among the waterfowl isolates, 68% to 73% homology with parrot ABVs, and 67% to 69% homology with BDVs (Payne et al., 2011a; Guo et al., 2012). Interestingly, an alignment of the N/X intergenic region of two Canada goose bornavirus isolates revealed that the waterfowl bornaviruses were configured more like BDV, sharing important regulatory sequences with BDV (Payne et al., 2011a).

These data confirm that the waterfowl bornaviruses circulating in North America represent a separate, independent cluster and a new species of bornavirus, distinct from psittacine ABVs, BDVs and from the putative avian BDV sequence isolated from mallards and jackdaws in Sweden (Berg et al., 2001). The very close similarity of swan and geese isolates and the fact that these species occupy similar habitats, suggests that ABV likely circulate between various waterfowl species in North America.

Significance of ABV infection in waterfowl

The presence of ABV in tissue has been associated with clinical disease and pathologic lesions in Canada geese and trumpeter swans in Canada (Smith et al., 2010; Delnatte et al., 2011), and in California (HL Shivaprasad, personal communication) suggesting that ABV infection can cause disease in waterfowl. While some species of waterfowl, such as Canada geese and mallards, are numerous, other species are rarer. Trumpeter swans were extirpated from eastern Canada as a result of overhunting by the early 20th century and have been the subject of a successful captive breeding and reintroduction program since the 1980s. The Ontario population is now estimated at approximately 1000 birds (Lumsden et al., 2009), and trauma and lead poisoning were historically the main causes of mortality. Whether mortality associated with ABV infection plays a significant role in this small population of relatively high genetic relatedness remains uncertain.

ABV was also detected in many apparently healthy Canada geese and other waterfowl species (Payne et al., 2011a; Guo et al., 2012; Payne et al., 2012). The pathogenesis of ABV infection is not well understood and the exact factors that trigger or influence the onset of clinical disease remain uncertain. Whether this can be attributed to a difference in pathogenicity among strains, a difference of susceptibility among species and individuals or other environmental factors has yet to be determined.

The frequency of transmission of ABV from waterfowl to other species of birds and its significance are unknown. Waterfowl produce large wet droppings that can heavily contaminate the environment and can be a source of virus for other birds. Predation on waterfowl may be another method of transmission of ABV. A bald eagle, a species known to predate on geese, recently died in Texas from an acute encephalitis and ABV-CG strain was detected in its brain (Payne et al., 2012). Other species of raptors have been reported to be affected with a PDD-like syndrome (Shivaprasad et al., 1995).

It is not known whether ABV can spread between waterfowl and parrots. Experimental inoculation of ABV-CG into cockatiels is currently underway at Texas A&M (I Tizard, personal communication). Based on the high lethality rate of ABV among large psittacines, ABV likely originated in other species. The presence of ABV infection in apparently healthy wild waterfowl (Payne et al., 2011a; Guo et al., 2012) suggest that waterfowl species may function as reservoir hosts. Some authors hypothesized that macaws were originally infected by contact with ABV-infected waterfowl while in quarantine (Payne et al., 2012).

It is not also known whether ABV can spread between waterfowl and poultry species. Domestic duck eggs from commercial hatcheries have been shown to be infected with ABV-CG, raising the concern that the waterfowl genotype of ABV may be a new threat or a previously unrecognized pathogen of domestic ducks. (Hoppes et al., 2013). The risk for the commercial poultry industry has not been determined.

Recent communication with colleagues in Britain indicates that encephalitis similar to that seen in Canada geese with ABV infection has been seen in wild geese in the UK (JP Duff, personal communication). Specific identification of ABV infection in waterfowl outside of North America has not been made as far as we are aware. However,

given the migratory nature of the host species, ABV-CG infection is unlikely to be restricted to North America.

The exact role of wild birds in the epidemiology of ABV and BDV is unknown and the natural reservoirs of these viruses remain unidentified. In order to assess the overall impact of this virus on wildlife, a more detailed knowledge of the susceptibility of wild avian species and of the geographic distribution of the virus will need to be determined (Smith et al., 2010).

RESEARCH GOALS AND OBJECTIVES

The recent identification of a novel genotype of avian bornavirus in wild Canada geese and trumpeter swans in Ontario motivated this graduate research project.

Subsequent to the discovery of ABV in waterfowl, various papers, including several arising from the work described in this thesis, have been published (Delnatte et al., 2011; Payne et al., 2011a; Guo et al., 2012; Payne et al., 2012; Delnatte et al., 2013). The design and the objectives of the research project presented in this manuscript actually predate these publications. A review of bornaviruses in birds, with a specific emphasis on waterfowl, summarizes most of the information currently available (Payne et al., 2012). This review was the result of a collaborative effort between researchers from the University of Guelph and Texas A&M University, and is presented in the Appendix A.

The overall objectives of this study were to acquire further knowledge about the pathology and epidemiology of avian bornavirus in free-ranging waterfowl in Ontario using retrospective and prospective methods, and to assess the susceptibility of domestic poultry species to this virus.

The present study was driven by several hypotheses:

- Hypothesis 1: Avian bornavirus causes disease (i.e., clinical signs and pathological lesions) in free-ranging Canada geese and trumpeter swans, and thus the virus is present in the tissues of birds showing disease consistent with ABV infection in other avian species (psittacines).

- Hypothesis 2: Avian bornavirus is present in Ontario waterfowl at a prevalence of 1% to 4% and waterfowl species shed the virus in their feces and develop serum antibodies as a result of infection.

- Hypothesis 3: Avian bornavirus can be transmitted vertically in wild waterfowl species.

- Hypothesis 4: Domestic fowl are susceptible to infection with avian bornavirus.

Each of these hypotheses was addressed by the following objectives:

- Objective 1: To determine the prevalence of and correlation among clinical signs, pathological lesions and presence of ABV in tissues from wild waterfowl in Ontario (retrospective study).

- Objective 2: To determine the prevalence of ABV in wild waterfowl in southern Ontario (prevalence survey) and assess the correlation between the presence of virus in feces (shedding) and the presence of antibodies in serum.

- Objective 3: To demonstrate the presence of ABV in eggs, developing embryos, and newly-hatched Canada geese originating from a flock infected with ABV.

- Objective 4: To infect domestic chickens, ducks and geese, and to characterize the viral distribution in tissues, seroconversion and viral shedding in these species (experimental infection trial).

CHAPTER 2: RETROSPECTIVE STUDY

Pathology and diagnosis of avian bornavirus infection in wild Canada

geese (Branta canadensis), trumpeter swans (Cygnus buccinator) and

mute swans (Cygnus olor) in Canada: a retrospective study

Pauline Delnatte1,2, Davor Ojkic3, Josepha DeLay3, Doug Campbell4, Graham Crawshaw2 and Dale A Smith1*

1 Ontario Veterinary College, University of Guelph, Guelph, Ontario, Canada, N1G 2W1

2 Toronto Zoo, Toronto, Ontario, Canada, M1B 5K7

3 Animal Health Laboratory, University of Guelph, Guelph, Ontario, Canada, N1G 2W1

4 Canadian Cooperative Wildlife Health Centre, Guelph, Ontario, Canada, N1G 2W1

Keywords: Avian bornavirus, Bornaviridae, Branta canadensis, Canada goose, Cygnus buccinator, encephalitis, ganglioneuritis, immunohistochemistry, polymerase chain reaction, trumpeter swan.

Published in Avian Pathology. 2013. 42(2):114–128.

ABSTRACT

Nine hundred fifty-five pathology cases collected in Ontario between 1992 and 2011 from wild free-ranging Canada geese, trumpeter swans and mute swans were retrospectively evaluated for the pathology associated with avian bornavirus (ABV) infection. Cases were selected based on the presence of upper gastrointestinal impaction, central nervous system histopathology or clinical history suggestive of ABV infection. The proportion of birds meeting at least one of these criteria was significantly higher at the Toronto Zoo (30/132) than elsewhere in Ontario (21/823). Central, peripheral and autonomic nervous tissues were examined for the presence of lymphocytes and plasma cells on histopathology. The presence of virus was assessed by immunohistochemistry (IHC) and RT-PCR on frozen brains and on formalin-fixed paraffin-embedded (FFPE) tissues. Among selected cases, 86.3% (44/51) were considered positive on histopathology, 56.8% (29/51) were positive by IHC, and RT-PCR was positive on 88.2% (15/17) of the frozen brains and 78.4% (40/51) of the FFPE samples. Histopathological lesions included gliosis and lymphoplasmacytic perivascular cuffing in brain (97.7%), spinal cord (50%), peripheral nerves (55.5%), and myenteric ganglia or nerves (62.8%), resembling lesions described in parrots affected with proventricular dilation disease. Partial amino acid sequences of the nucleocapsid gene from seven geese were 100% identical amongst themselves and 98.1 to 100% identical to the waterfowl sequences recently described in the United States. Although ABV has been identified in apparently healthy geese, our study confirmed that ABV can also be associated with significant disease in wild waterfowl species.

INTRODUCTION

In 2009, we identified a novel genotype of avian bornavirus (ABV) in association with non-suppurative inflammation in the central, peripheral and autonomic nervous systems of several Canada geese (Branta canadensis) and trumpeter swans (Cygnus buccinator) in southern Ontario as part of our surveillance program for diseases in free- ranging wildlife (Smith et al., 2010; Delnatte et al., 2011). ABV was first identified in 2008 and proposed as the etiology of proventricular dilation disease (PDD), a significant pathological syndrome that has been reported worldwide in more than 80 species of psittacine birds (Honkavuori et al., 2008; Kistler et al., 2008). Subsequent research, including inoculation studies in birds (Gancz et al., 2009; Gray et al., 2010; Mirhosseini et al., 2011; Payne et al., 2011b; Piepenbring et al., 2012) and investigations of outbreaks (Gancz et al., 2010; Kistler et al., 2010; Wünschmann et al., 2011; Heffels-Redmann et al., 2012) provided strong supporting evidence for a causal relationship. Psittacine birds affected with PDD exhibit various non-specific gastrointestinal and neurological signs with a high case fatality rate once clinical signs develop. Characteristic gross pathological findings of PDD include poor body condition, proventricular and ventricular dilation, and duodenal distension (Shivaprasad et al., 1995; Berhane et al., 2001; Gancz et al., 2010). Lymphoplasmacytic infiltrates within myenteric ganglia and nerves of the proventriculus and ventriculus are considered the histological hallmarks of PDD (Ritchie et al., 1998) and are considered by some authors to be pathognomonic for the disease (Berhane et al., 2001; Schmidt et al., 2003). Similar infiltrates are frequently present in the brain, spinal cord, peripheral nerves (Shivaprasad et al., 1995; Berhane et al., 2004), adrenal glands, heart (Vice et al., 1992), eye (Steinmetz et al., 2008; Korbel et al., 2011) and autonomic nerves and ganglia adjacent to various tissues (Doneley et al., 2007). In infected parrots, ABV exhibits a high tropism for neuroectodermal cells including neurons, astroglia, and ependymal cells of the central nervous system, neurons of the peripheral nervous system, and adrenal cells but is also present in many other tissues, as demonstrated by immunohistochemistry (IHC), Western blot analysis and reverse transcription polymerase chain reaction (RT-PCR) (Honkavuori et al., 2008; Gancz et al., 2009; Lierz et al., 2009; Ouyang et al., 2009; Rinder et al., 2009; Weissenböck et al., 2009a; Weissenböck et al.,

2009b; Kistler et al., 2010; Lierz et al., 2010; Raghav et al., 2010; Wünschmann et al., 2011).

Although PDD had been considered a disease of captive psittacine birds, consistent histopathological lesions have been identified occasionally in a variety of other avian species, including passerines, falconiformes, piciformes and anseriformes (Daoust et al., 1991; Gregory et al., 2000; Perpiñán et al., 2007; Weissenböck et al., 2009b; Delnatte et al., 2011). In 1991, Daoust et al. described non-suppurative encephalitis and ganglioneuritis in two emaciated free-ranging Canada geese from eastern Canada that showed proventricular dilatation (Daoust et al., 1991). The similarity to PDD was highlighted in that publication, particularly as common causes of proventricular dilatation in waterfowl, such as chronic lead poisoning and mechanical obstruction, had been excluded. Attempts to identify a causative agent, including viral culture in several cell lines and inoculation of young mice and chicken embryos, were not successful.

Since our initial report of ABV in waterfowl, others have identified the presence of the virus in apparently healthy wild Canada geese and mute swans in the United States using oropharyngeal / cloacal swabs and brain samples. No clinical examination or necropsies were performed on these birds (Payne et al., 2011a; Guo et al., 2012). Partial nucleotide sequences of ABV identified in Canadian and American waterfowl are very similar, sharing 95.8% to 100% nucleotide identity as determined using pairwise comparison, with sequences clustering separately from those previously described in psittacines and other avian species, and from Borna disease virus (BDV) (Delnatte et al., 2011; Payne et al., 2011a). It thus appears that the ABV identified in waterfowl is a separate genotype within the family Bornaviridae, and that the virus may be present in birds that do and do not appear to suffer from clinical disease.

The objectives of this study were to describe the results of a detailed retrospective investigation evaluating the presence of ABV in wild Canada geese and swans in southern Ontario and to assess the correlation among clinical signs, histopathological lesions and presence of ABV in tissues.

MATERIALS AND METHODS

Case selection

Nine hundred and fifty five necropsy reports from the Toronto Zoo (n=132) and the Canadian Cooperative Wildlife Health Centre – Ontario Region (CCWHC-Ont) (n=823) databases from 1992 to 2011 were reviewed to identify Canada geese or swans with upper gastrointestinal impaction (esophagus, proventriculus or ventriculus); non- suppurative encephalitis, myelitis, neuritis, or ganglioneuritis; or having a clinical history suggestive of ABV infection (e.g. weakness, neurological signs). Cases were excluded if a specific diagnosis accounted for the clinical signs and pathological lesions; e.g., soybean impaction, lead poisoning or West Nile virus infection. Six additional cases (Canada geese) with a histological diagnosis of non-suppurative encephalitis were submitted from the CCWHC-Atlantic Region (CCWHC-Atl), including the two birds reported in Daoust’s paper in 1991 (Daoust et al., 1991). Nine negative control birds were selected from the CCWHC-Ont database based on the absence of these inclusion criteria.

The following necropsy information was reviewed as available: species, geographic location where found, age class, sex, clinical history, gross necropsy findings including body condition, histopathology, and the results of any additional diagnostic tests such as toxicologic and microbiologic testing.

Histopathology

Slides from all selected cases were reviewed by one pathologist, blinded by case, for the presence of histological lesions. Tissue samples had been formalin-fixed, paraffin- embedded, sectioned at 4-μm and stained with haematoxylin and eosin. Nerves and ganglia in all sections of all tissues (central, peripheral and autonomic nervous system) were specifically examined for the presence of lymphocytes and plasma cells. A case was considered to be positive for ABV-consistent histopathology if non-suppurative inflammation was present in any nervous tissue.

Immunohistochemistry

Immunohistochemistry (IHC) was performed on one or more sections of formalin-fixed paraffin-embedded (FFPE) brain from each case as available. When brain was not available, spinal cord or peripheral nerves were assessed. Briefly, IHC was performed on 4-μm tissue sections mounted on charged slides and using an automated staining instrument (Dako Autostainer, Dako Canada Inc., Mississauga, Ontario, Canada) with rabbit polyclonal antiserum raised against the ABV nucleocapsid protein (ABV-N) (provided by the deRisi Laboratory, UC-SF, San Francisco, California, USA). Detailed description of the cloning, recombinant expression, purification, and generation of polyclonal antisera against the ABV nucleocapsid (ABV2-N) protein has been previously published (Gancz et al., 2009; Raghav et al., 2010). Following manual deparaffinization and rehydration, tissue sections were treated with 3% hydrogen peroxide to quench endogenous peroxidase activity. Antigen retrieval was accomplished by incubation with Proteinase K (Dako) for 6 min. Sections were then incubated with rabbit anti-ABV-N antiserum (1:2,000) for 30 min, followed by 30-min incubation with a goat antimouse/antirabbit polymer visualization system (UltraVision ONE, Lab Vision Corp., Fremont, California, USA). All incubations were completed at room temperature. Nova Red was used as the chromogen (Vector Laboratories, Burlington, Ontario, Canada). Positive ABV-positive control brain sections included with each run were from a psittacine previously tested by both IHC and PCR. For negative reagent controls, duplicate sections of each control and test tissue were subjected to the same immunohistochemical procedure with substitution of pre-immune rabbit serum at a similar protein concentration to the anti-ABV-N antisera. All slides were interpreted by one author (Dr Josepha Delay) who was blinded to case information. A case was considered IHC positive when strong, diffuse intranuclear staining was present in neurons or glial cells.

Nucleic acid isolation and RT-PCR

Formalin-fixed paraffin-embedded (FFPE) tissues and thawed frozen or fresh tissue or cloacal swabs collected at necropsy (when available) were evaluated for the

presence of ABV by reverse transcriptase polymerase chain reaction (RT-PCR). The paraffin blocks were the same as those used for immunohistochemistry, and thus included one or more sections of brain in most cases, or spinal cord or peripheral nerves when brain was not available.

RNA extraction was performed on FFPE sections of brain using the RNeasy® FFPE Kit RNeasy® (FFPE Kit, Qiagen Inc., Mississauga, Ontario, Canada) according to manufacturer's instructions. Total nucleic acids were extracted from thawed frozen or fresh brain samples and cloacal swabs using MagMAX-96 Viral RNA Isolation Kit in a MagMAX Express-96 Magnetic Particle Processor (Applied Biosystems Inc., Foster City, California, USA) from 50 µl aliquots of 5-10% tissue suspensions or, for cloacal swabs, from 50 µl aliquots of virus transport media (Starplex Scientific Inc., Etobicoke, Ontario, Canada).

Thawed frozen or fresh brain samples and cloacal swabs collected at necropsy were evaluated for the presence of ABV by conventional gel-based RT-PCR detection of the ABV nucleocapsid (ABV-N) gene and by real-time RT-PCR (rtRT-PCR) detection of the ABV matrix (ABV-M) gene. FFPE tissues were evaluated for the presence of ABV by rtRT-PCR detection of the ABV matrix (ABV-M) gene.

For the conventional gel-based RT-PCR, the primers used were directed against ABV-N and have previously been used for ABV identification in psittacine birds (Kistler et al., 2008). The RT-PCR was carried out in 25 µl reactions using One-Step RT-PCR Kit (Qiagen Inc., Mississauga, Ontario, Canada).

The rtRT-PCR assay was a duplex test with two sets of primers and TaqMan probes: (1) ABV_M_120201 – targeting psittacine ABV matrix gene sequences; and (2) ABVG_M_111029 – targeting geese ABV matrix gene sequences (Table 2.1a). The amplification was carried out in 25 µl reactions in a LightCycler 480 Real-Time PCR System (Roche, Laval, Quebec, Canada) using AgPath-ID One-Step RT-PCR Kit (Applied Biosystems, Foster City, California, USA) under cycling parameters described in Table 2.1b. A rtRT-PCR crossing point value (Cp) less than 37.00 was considered positive, between 37.00 and 39.99 inconclusive, and equal or greater than 40.00 negative. A case was classified as ABV-PCR positive if one or more frozen, fresh or FFPE samples

tested were positive on at least one RT-PCR (either conventional or rtRT-PCR), and negative if all samples tested were found to be negative.

Table 2.1a.

NAME FUNCTION SEQUENCE ABV_M_F_120201 Primer, Forward 5'-CAAGGTAATYGTYCCTGGATGGCC-3' ABV_M_Pr_120201 Probe, TaqMan 5'-TAATGTTGGARATAGACTTTGTTGG-3' ABV_M_R_120201 Primer, Reverse 5'-TCACTGAAAGAAANGGTATRTTGAT-3' ABVG_M_F_111029 Primer, Forward 5'-CGAGGGAGAAGAGACTGGTTGATT-3' ABVG_M_Pr_111029 Probe, TaqMan 5'-ATGTGGAACCTGCTGGTCACTCA-3' ABVG_R_111029 Primer, Reverse 5'-ACTGCCAAAGAGTTGAGCGT-3'

Table 2.1b.

PARAMETER CYCLES TIME TEMPERATURE RT 1 600 seconds 45°C ACTIVATION 1 600 seconds 95°C DENATURATION 5 seconds 94°C 45 ANEALING/EXTENSION 60 seconds 60°C

Sequencing and sequence analysis

Partial nucleotide sequences of ABV-N genes were determined at the Laboratory Services, University of Guelph, Ontario. Sequences were assembled with the SeqMan module of Lasergene 10.0 software package (DNASTAR, Inc., Madison, Wyoming, USA). The sequences reported in the present study have been deposited in GenBank under accession numbers JX258867, JX258868, JX258869, JX258870, JX258871, JX258872, and JX258873. Multiple alignments were carried out with the MegAlign module of Lasergene 10.0 software package using the default settings of its ClustalW algorithm. Sequence comparisons were carried out using a 313-base pair (bp) fragment from nt 692 to nt 1004, relative to complete genomic sequence of ABV isolate bil, GenBank accession no. EU781967. Phylogenetic trees were also generated with the MegAlign module of Lasergene software using its default settings.

Statistical Evaluation

Statistical analyses were performed using GraphPad QuickCalcs software (http://www.graphpad.com/quickcalcs/). Fisher’s exact test was used for comparisons among species (overall and within each location) and between locations (overall and for each species) for percentage of birds meeting inclusion criteria; results of histopathology, IHC, RT-PCR on frozen brain, and rtRT-PCR on FFPE; and distribution of lesions (histopathology and IHC). When the difference was significant (p<0.05), the odd ratio (OR) was calculated and reported with a 95% confidence interval (CI). For rtRT-PCR, the mean Cp values were compared between type of tissues (frozen brain vs FFPE) using paired t-tests, and between location and species using unpaired t-tests. Two-tailed p values less than 0.05 were considered significant.

Test agreement was estimated using overall percent agreement (OPA), positive percent agreement (PPA) and negative percent agreement (NPA) (Feinstein et al., 1990) for all combinations of histopathology, IHC, RT-PCR on fresh/frozen tissues, and RT-PCR on FFPE. Test agreement was carried out for each species and for all species combined.

RESULTS

Cases meeting inclusion criteria

Forty-one of 555 Canada geese (7.4%), 8/321 trumpeter swans (2.5%) and 2/79 mute swans (2.5%) met at least one of the inclusion criteria and were included in the study (Table 2.2). The proportion of Canada geese selected was significantly higher than the proportion of swans (p=0.0007; OR = 3.11; 95% CI [1.54; 6.29]), and was significantly higher at the Toronto Zoo (27/88) than elsewhere in Ontario (14/467) (p<0.0001; OR = 14.32; 95% CI [7.12; 28.8]). There was no difference between the two sites for trumpeter swans (p = 0.07) or mute swans (p=1.00).

Table 2.2. Number of Canada geese, trumpeter swans and mute swans meeting initial case inclusion criteria in a retrospective survey of necropsy databases from Toronto Zoo and CCWHC-Ontario from 1992 to 2011.

Species Canada goose trumpeter swan mute swan Toronto CCWHC - Toronto CCWHC- Toronto CCWHC- Inclusion Zoo Ont Zoo Ont Zoo Ont criteria** (n=88) (n=467) (n=41) (n=280) (n=3) (n=76)

Clinical history^ 21 12 3 5 0 2

Upper gastrointestinal impaction 10 2 0 0 0 1

Non-suppurative inflammation within nervous system^^ 22 12 3 5 0 1

Total cases 27 14 3 5 0 2 included (30.6%) (3%) (7.3%) (1.8%) (0%) (2.6%)

** Cases were excluded if a specific diagnosis explained the clinical signs or pathological lesions identified. ^ Clinical history was not available for all cases within the database. ^^ Brain was not available for all cases within the database. CCWHC- Ont = Canadian Cooperative Wildlife Health Center - Ontario region CNS = central nervous system

Summaries of clinical history and gross necropsy findings in all selected geese and swans are presented in Tables 2.3 and 2.4. Clinical signs from the necropsy histories which met the clinical inclusion criteria included: somnolence, weakness, and lethargy; and suspected blindness, lameness, ataxia, hypermetria, inability to stand or to fly, torticollis, head tremors, stargazing, and opisthotonos. Of the selected cases, 30/41 (73.2%) Canada geese, 5/8 (62.5%) trumpeter swans and 1/2 (50.0%) mute swans were in poor body condition.

Six additional Canada geese from CCWHC-Atl database with nervous system histopathology were also included in this study; 3/6 showed upper gastrointestinal impaction; 5/6 were in poor body condition and 3/3 that were not found dead showed neurological sings prior to death.

Table 2.3. Summary of clinical history, gross necropsy findings, histopathology, IHC and RT-PCR results in 41 wild Canada geese evaluated for the presence of avian bornavirus.

Clinical history Gross necropsy Histopathology and testing

Neuro- Poor Upper Histo- RT-PCR logical Weak- body GI im- patholo- RT-PCR (FFPE Bird ID Origin Sex Age signs ness condition paction gy (H&E) IHC (brain) tissue)

CAGO-Z1 TZ F AHY N Y Y Y + + + (33.15)* + (24.64)

CAGO-Z2 TZ M AHY Y Y N N + + + (19.24)* + (20.33) CAGO-Z3 TZ F HY Y N Y Y + + + (27.21)* + (12.9) CAGO-Z4 TZ F AHY Y Y N Y + + + (25.6)* + (19.74)

CAGO-Z5 TZ M HY Y Y Y N + + + (21.71)* + (19.9) CAGO-Z6 TZ M AHY Y N Y N + + + (17.0) + (13.25) CAGO-Z7 TZ M AHY Y Y Y N + + + (17.92) + (16.37)

CAGO-Z8 TZ M AHY U U Y N + + + (22.88) + (30.83) CAGO-Z9 TZ M AHY Y N Y N + - + (25.32)* + (21.32) CAGO-Z10 TZ M AHY Y N Y N + + + (29.62) - ^^

CAGO-Z11 TZ M AHY Y Y Y N + + NA + (20.72) CAGO-Z12 TZ F U Y Y Y N + + NA + (31.95) CAGO-Z13 TZ F U Y N N N + + NA + (28.94)

CAGO-Z14 TZ M U Y N N N + + NA + (29.3) CAGO-Z15 TZ M HY Y N N N + + NA + (33.88)

CAGO-Z16 TZ M U Y Y N N + + NA + (15.79) CAGO-Z17 TZ M AHY N Y Y Y + + NA + (29.36) CAGO-Z18 TZ F AHY N Y Y N + + NA + (26.59) CAGO-Z19 TZ F AHY U U Y Y + + NA + (12.49) CAGO-Z20 TZ M U U U N N + - NA - CAGO-Z21 TZ M U N Y Y Y + - NA -

CAGO-Z22 TZ M U Y Y Y N + - NA + (28.66) CAGO-Z23 TZ M HY U U Y Y + - NA + (28.35) CAGO-Z24 TZ M HY Y Y Y N - - NA -

CAGO-Z25 TZ M U N Y Y Y - - NA - CAGO-Z26 TZ M U N N Y Y - - NA -

CAGO-Z27 TZ F AHY U U Y Y - - NA -

CAGO-W1 Burlington M AHY Y N Y N + + + (18.6)* + (11.81)

CAGO-W2 Mississauga M AHY Y Y Y N + + + (15.99) + (14.36)

CAGO-W3 Toronto M AHY Y Y Y N + + + (12.78) + (18.91)

CAGO-W4 Burlington F AHY Y N Y N + + + (14.48) + (19.94)

CAGO-W5 Etobicoke F AHY Y Y Y Y + - + (24.76)^ + (27.93)

CAGO-W6 Burlington M AHY Y N Y N + + - ** + (31.27)

CAGO-W7 Guelph M HY Y N Y N + - NA + (24.61)

CAGO-W8 Guelph M AHY Y Y N N + - NA + (22.95)

CAGO-W9 Mississauga U AHY N N N N + - NA + (31.64)

CAGO-W10 Winona F AHY Y N N N + - NA + (33.71)

CAGO-W11 Markham F AHY U U N N + - NA + (21.22)

CAGO-W12 Toronto M AHY Y N Y N + - NA + (20.59)

CAGO-W13 King City U AHY Y Y Y Y - - NA -

CAGO-W14 Fergus M HY Y Y Y N - - NA -

Table 2.4. Summary of clinical history, gross necropsy findings, histopathology, IHC and RT-PCR results in 8 wild trumpeter swans and 2 wild mute swans evaluated for the presence of avian bornavirus.

Clinical history Gross necropsy Histopathology and testing Neuro- Poor Upper Histo- RT-PCR logical Weak- body GI im- patholo- RT-PCR (FFPE Bird ID Origin Sex Age signs ness condition paction gy (H&E) IHC (brain) tissue)

TRSW-Z1 TZ M AHY Y Y N N + + NA + (29.18) TRSW-Z2 TZ M AHY N Y Y N + + NA + (25.26) TRSW-Z3 TZ F HY Y N Y N + - NA -

TRSW-W1 Toronto M AHY Y N Y N + + NA + (21.1) TRSW-W2 Oshawa M AHY Y Y N N + + NA + (23.81) TRSW-W3 Dundas F AHY Y Y N N + + NA + (26.31) TRSW-W4 Aurora F AHY Y Y Y N + + NA + (22.84) TRSW-W5 Burlington F AHY Y Y Y N + - NA + (20.52)

Frenchmans MUSW-W1 Bay M AHY Y Y N N + - - + (36.41) Humberbay MUSW-W2 Park M AHY N Y Y Y - - NA -

Legend for the tables 2.3 and 2.4:

Location (all in Ontario): TZ = Toronto Zoo Sex: F = female; M = male; U = unknown Age: HY = hatching year; AHY = after hatching year; U = unknown Clinical history and gross necropsy: Y = yes; N = no; U = unknown Testing: IHC = immunohistochemistry; RT-PCR = reverse transcriptase polymerase chain reaction; NA = tissues not available for testing; RT-PCR (brain): Unless mentioned, brains were tested for the presence of ABV by both gel -based RT-PCR detection of the ABV nucleocapsid gene and by quantitative RT-PCR detection of the ABV matrix gene; the crossing point values (Cp) are indicated in brackets. RT-PCR (FFPE tissue): Formalin-fixed paraffin-embedded (FFPE) tissues were tested for the presence of ABV by quantitative RT-PCR detection of the ABV matrix gene. The crossing point values (Cp) are indicated between brackets.

Diagnostic findings A summary of the histopathology findings and IHC and RT-PCR results for all selected geese and swans are presented in Tables 2.3 and 2.4. Among selected cases, the proportion of cases identified as positive for each diagnostic test (histopathology, IHC, RT-PCR on frozen brain and RT-PCR on FFPE) did not differ significantly among species or between locations.

Histopathology

Of the 41 selected Canada geese, 35 (85.4%) were considered positive on histopathology, including 23 of the 27 cases from the Toronto Zoo and 12 out of 14 cases found elsewhere in Ontario. All selected trumpeter swans and one of the two selected mute swans were considered positive on histopathology. The tissue distribution of non- suppurative inflammatory lesions is shown in Tables 2.5 and 2.6, and some characteristic histological lesions are illustrated in Figure 2.1.

Table 2.5. Tissue distribution of non-suppurative inflammatory lesions, immunohistochemical staining and RT-PCR results in 35 wild Canada geese evaluated for the presence of avian bornavirus.

Histopathology

RT-PCR Central nervous system PNS Gastro-inte stinal tract Adrenal Kidney Other

CAGO-Z1 + + + - - - + + NA NA NA NA - NA + + - NA NA NA - -

CAGO-Z2 + + + + - + + + + + NA NA NA ------+ thyroid thyroid, CAGO-Z3 + + + + + + + + + + NA NA NA + - NA + + NA NA + - heart

CAGO-Z4 + + + - + + + NA + + + NA + NA - - - - + + - +

CAGO-Z5 + + + + + NA + NA NA NA + NA NA - - - - NA - - - -

CAGO-Z6 + + + + + NA - NA NA NA NA NA NA + + - - - + - - +

CAGO-Z7 + + + + - - + + + + NA NA + - + + - NA - - - +

CAGO-Z8 + + + + + + + + + + - NA + + + + + - + + - -

CAGO-Z9 + + + - + NA + NA NA NA NA NA - NA - NA - NA - - - -

CAGO-Z10 + - ^^ + + + - + + + + NA NA NA - - - - NA + + + -

CAGO-Z11 NA + + + + + + + NA NA NA NA NA - - - + NA + + - +

CAGO-Z12 NA + + + + - + + + + NA NA - + + NA + - + - - - heart

CAGO-Z13 NA + NA NA + + + + NA NA NA NA NA + - - + + NA NA - +

CAGO-Z14 NA + + + + + + + NA NA NA NA - - + NA + - NA NA - -

CAGO-Z15 NA + + + + + + + + + NA NA NA + + - + + NA NA - +

CAGO-Z16 NA + + + - + + + + NA NA NA + + + - + NA NA NA - +

CAGO-Z17 NA + + + + + + NA + NA NA NA NA + + + - NA + - - + thyroid CAGO-Z18 NA + + + - NA + NA NA NA NA NA ------NA NA - +

CAGO-Z19 NA + + + + NA + NA + NA NA NA + + + + + - + + - + thyroid CAGO-Z20 NA - +^ - +^ - + - - - NA NA NA - - - - NA + - - +

CAGO-Z21 NA - - - - - NA NA - NA NA NA - + - - - - - + - +

CAGO-Z22 NA + + - + NA + - NA NA NA NA NA + - - + NA NA NA - +

CAGO-Z23 NA + + NA + - NA NA NA NA NA NA NA + + + + - + + - + thyroid

Table 2.5 (Cont.).

Histopathology

RT-PCR Central nervous system PNS Gastro-inte stinal tract Adrenal Kidney Other

Brain Brain Cerebrum Cerebellum Brainstem Optic lobes Spinal cord Eso- AN inter- AN inter- Bird ID (fro - (FF pha- Proven- Ventri- Intes- Cloa S stitium S stitium ANS . zen) PE) H&E IHC H&E IHC H&E IHC H&E IHC H&E IHC gus triculus culus tines ca * ** * ** *

CAGO-W1 + + + + + + + + - + - + - NA NA NA + NA - - - - Pan - CAGO-W2 + + - + - + NA NA + + NA NA - NA + NA + NA NA NA - - creas CAGO-W3 + + NA NA - + - + NA NA - NA + - + - - - + - - +

CAGO-W4 + + + + - + + + - + NA NA + + + + + NA - + + +

CAGO-W5 + + +^ - - - + - + - + NA + + + + + + + + - - ovary CAGO-W6 - + + + - + - + NA NA NA NA NA - - NA - NA - - - +

CAGO-W7 NA + - - + - NA NA NA NA NA NA - NA NA NA NA NA - - - - Thy - CAGO-W8 NA + + - + - + - - NA NA NA + NA NA NA + NA - + - + roid CAGO-W9 NA + + - NA NA NA NA - - NA NA NA NA NA NA - NA NA NA - +

CAGO-W10 NA + +^ ------NA NA - - - - - NA - - - + heart, CAGO-W11 NA + + - + - + - NA NA NA NA + + + NA + NA + + - - lung CAGO-W12 NA + + - + NA + - + - NA NA + - NA - - NA + - - -

Total 19/ 15/ 16/ 11/ 11/ 14/ 3/3 positive 15/16 32/35 30/33 32 22/34 27 26/30 23 14/21 16 3/6 1/1 21 15/27 16/30 8/24 17/34 4/15 25 11/25 5 21/35

For histopathology: + = presence of lesions consistent with avian bornavirus infection; - = absence of lesions consistent with avian bornavirus infection; NA = tissue not examined. For IHC: + = presence of specific staining for avian bornavirus antigen; - = absence of specific staining for avian bornavirus antigen; NA = tissue not examined. +^ = Inflammation present only in the meninges ^^ RT-PCR inconclusive * ANS = autonomic nervous system: considered positive if lymphocytes or plasma cells present within nerves or ganglia adjacent to or within tissue of interest ** Interstitium: considered positive if lymphocytes or plasma cells present within the interstitium of tissue of interest PNS = peripheral nervous system: vagus, brachial or/and sciatic nerves were assessed depending on cases as available H&E = haematoxylin and eosin staining IHC = immunohistochemistry RT-PCR = reverse transcriptase polymerase chain reaction FFPE = Formalin-fixed paraffin-embedded tissue CAGO = Canada goose TRSW = trumpeter swan MUSW = mute swan 58

Table 2.6. Tissue distribution of non-suppurative inflammatory lesions, immunohistochemical staining and RT-PCR results in eight wild trumpeter swans and one wild mute swan evaluated for the presence of avian bornavirus.

Histopathology

RT-PCR Central nervous system PNS^ Gastro-inte stinal tract Adrenal Kidney Other

Cerebrum Cerebellum Brainstem Optic lobes Spinal cord inter- inter- Bird ID Brain Brain Eso- Proven- Ventri- Intes- ANS stitium ANS stitium ANS . (frozen) (FFPE) H&E IHC H&E IHC H&E IHC H&E IHC H&E IHC phagus triculus culus tines Cloaca * ** * ** * TRSW-Z1 NA + + + + NA + NA NA NA + NA + - - - + NA + + - - - TRSW-Z2 NA + + + - NA + NA + + NA NA + NA - - - - + - - - +(testes) TRSW-Z3 NA - - - + - - - NA NA ------NA NA NA - - -

+ TRSW-W1 NA + + + + NA + NA NA NA + NA NA - - - - NA - - - + (thyroid) TRSW-W2 NA + + NA + - + + NA NA NA NA + NA - NA + NA - + - - - TRSW-W3 NA + + + + + + + + + NA NA + NA NA NA + NA NA NA - - - TRSW-W4 NA + + NA + + + + NA NA NA NA NA NA NA NA + NA NA NA - + - TRSW-W5 NA + + - NA NA + - NA NA NA NA NA NA NA NA - NA NA NA - - + (heart)

MUSW-W1 - + +^ - + - + - NA NA - NA ------+ - - + -

Tot al positive 0/1 8/9 8/9 4/7 7/8 2/5 8/9 3/6 2/2 2/2 2/4 0/1 4/6 0/4 0/6 0/5 4/9 0/2 3/5 2/5 0/9 3/9

For histopathology: + = presence of lesions consistent with avian bornavirus infection; - = absence of lesions consistent with avian bornavirus infection; NA = tissue not examined. For IHC: + = presence of specific staining for avian bornavirus antigen; - = absence of specific staining for avian bornavirus antigen; NA = tissue not examined. +^ = Inflammation present only in the meninges * ANS = autonomic nervous system: considered positive if lymphocytes or plasma cells prese nt within nerves or ganglia adjacent to or within tissue of interest ** Interstitium: considered positive if lymphocytes or plasma cells present within the interstitium of tissue of interest PNS = peripheral nervous system (vagus, brachial or/and sciatic nerves as available) H&E = haematoxylin and eosin staining IHC = immunohistochemistry RT-PCR = reverse transcriptase polymerase chain reaction FFPE = Formalin-fixed paraffin-embedded tissue CAGO = Canada goose TRSW = trumpeter swan MUSW = mute swan

All selected cases except one had histological lesions in the brain, which included perivascular cuffing by lymphocytes, plasma cells and, less frequently, macrophages; and focal and diffuse gliosis. Less common additional neuropathology included non- suppurative inflammation of the cerebral meninges; Wallerian degeneration with swollen axons, dilated axonal sheaths and gitter cells; and rare malacia and cerebral edema. The most commonly affected parts of the brain were the cerebrum (30/33 geese and 8/9 swans; overall 90.4%) and the brainstem (26/30 Canada geese and 8/9 swans; overall 87.2%) following by the optic lobes (14/21 Canada geese and 2/2 swans; overall 69.6%) and the cerebellum (22/34 Canada geese and 7/8 swans; overall 69.0%). Lesions were noticed in both grey and white matter, in cerebellar peduncles, in Purkinje layers, in choroid plexi and in the pituitary (pars nervosa). Lymphoplasmacytic perivascular infiltrates and Wallerian degeneration were also observed in peripheral nerves (11/21 Canada geese and 4/6 swans; overall 55.5%), including the brachial plexi, sciatic nerves and vagus nerves, and in spinal cord (3/6 Canada geese and 2/4 swans; overall 50.0%).

Among the histopathology-positive cases, 23/34 geese and 4/9 swans had lymphocytes or plasma cells within myenteric nerves or ganglia; with the intensity of the infiltrate varying greatly from nerve to nerve along the gastro-intestinal tract of an individual animal. Lesions were present in the esophagus (15/27 geese, 0/4 swans; overall 48.4%), proventriculus (16/30 geese, 0/6 swans; overall 44.4%), ventriculus (8/24 geese and 0/5 swans; overall 27.6%), intestines (17/34 geese and 4/9 swans; overall 48.8%), and cloaca (4/15 geese and 0/2 swans; overall 23.5%). Among histopathology positive cases, the proportions of Canada geese showing histological lesions in proventriculus and ventriculus were significantly higher when compared to combined values for trumpeter and mute swans (p=0.024, and p=0.0146, respectively). The distribution of histological lesions in other tissues did not differ significantly among species. All histopathology positive cases with a history of upper gastro-intestinal impaction had characteristic lesions in the esophagus, proventriculus and/or ventriculus. Four Canada geese and one mute swan were included in this study based on the presence of upper gastro-intestinal dilation but did not show any histological lesions in any nervous tissues, and thus were considered as histopathology negative cases.

Most cases with ganglioneuritis of the gastrointestinal tract also showed perivascular, perineural or periganglional non-suppurative infiltrates in the serosa, and/or tracking through the muscularis. However, as outlined in the material and methods, this alone was insufficient for the classification of a case as “histopathology positive”.

Lymphocytes and/or plasma cells were present within autonomic nerves or ganglia adjacent to the adrenal (14/25 geese and 3/5 swans; overall 56.7%), thyroid (n=7), heart (n=4), kidney (n=3), testes (n=1), ovary (n=1), lung (n=1), and pancreas (n=1). Non-suppurative interstitial adrenalitis and nephritis were noted, respectively, in 43.3% (11/25 geese and 2/5 swans) and 54.5% (21/35 geese and 3/9 swans) of histopathology-positive cases.

Other recurrent histological findings that were considered incidental included hepatic and splenic hemosiderosis and amyloidosis, pulmonary anthracosis, renal tubular mineralization, skeletal muscle degeneration, perivascular hepatitis and the presence of parasites in various tissues (renal and intestinal coccidia, vascular schistosomes, and gastro-intestinal nematodes and cestodes).

Figure 2.1. Histological lesions in waterfowl infected with avian bornavirus (ABV). Haematoxylin and eosin staining. A, Trumpeter swan, cerebrum. Marked and extensive perivascular lymphoplasmacytic cuffing. B, Canada goose, cerebrum. Large perivascular lymphoplasmacytic cuff. C, Canada goose, cerebrum. Well-defined glial nodule. D, Canada goose, vagus nerve. Diffuse and perivascular lymphoplasmacytic infiltration. E, Canada goose, adrenal. Interstitial infiltration by lymphocytes and plasma cells primarily around chromaffin cell islets. F, Canada goose, proventriculus. Lymphoplasmacytic infiltration within a myenteric ganglion.

Immunohistochemistry Out of the 41 selected Canada geese, 23 were positive by immunohistochemistry including 18/27 cases from the Toronto Zoo and 5/14 cases from the CCWHC-Ont. Six of eight trumpeter swans were positive by IHC, including 2/3 cases from the Toronto Zoo and 4/ 5 cases from the CCWHC-Ont. Neither mute swan was positive on IHC. Immunohistochemical staining was present consistently in the nucleus and occasionally in the cytoplasm of both neurons and glial cells (Figure 2.2). Staining intensity varied from mild to intense among cases. Staining that was considered non-specific cross- reactivity with cellular proteins was sometimes observed in circulating leukocytes.

The tissue distribution of IHC staining is presented in Tables 2.5 and 2.6. The ABV antigen was most commonly detected in the optic lobes (11/16 Canada geese and 2/2 swans; overall 72.2%) and in the brainstem (16/23 Canada geese and 3/6 swans; overall 65.5%) followed by the cerebrum (19/32 Canada geese and 4/7 swans; overall 59.0%) and the cerebellum (15/27 Canada geese and 2/5 swans; overall 51.5%). The tissue distribution of immunohistochemical staining did not differ significantly among species.

Figure 2.2. Immunohistochemical staining for avian bornavirus (ABV) antigen using rabbit polyclonal antiserum raised against ABV nucleocapsid protein. Canada goose, cerebrum. Note the intense intranuclear staining in neurons and glial cells, and the less frequent intracytoplasmic staining in neurons.

RT-PCR results

Fresh / frozen brains were available for only 16 Canada geese and one mute swan. Fifteen of the 16 Canada geese (93.7%) tested positive by at least one RT-PCR method (gel-based N gene RT-PCR or M gene rtRT-PCR), and one was negative by gel- based RT-PCR but no brain was available for subsequent rtRT-PCR. The mute swan tested negative by both methods. For the rtRT-PCR, the Cp values of the positive birds ranged from 12.78 to 33.15 with a mean and standard deviation of 21.75 (+/-5.85). The mean Cp for frozen brain samples was significantly lower in Canada geese that did not originate from the Toronto Zoo (p = 0.032).

Four cloacal swabs were tested by RT-PCR; three were taken from Canada geese for which frozen brain was positive by RT-PCR and one was collected from a mute swan for which the brain was negative. Only one of 3 goose swabs was positive, with a Cp of 16.95, and the swan swab was negative.

Out of the 51 selected birds, 40 were positive by rtRT-PCR on formalin fixed paraffin embedded (FFPE) tissues, including 32/41 Canada geese, 7/8 trumpeter swans and 1/2 mute swans. The result for one Canada goose was inconclusive (Cp = 37.4). The Cp values of the positive birds ranged from 11.81 to 36.41 with a mean and standard deviation of 24.22 (+/- 6.47). The mean Cp for FFPE samples did not differ significantly between locations or among species, and did not differ significantly from the mean value obtained from frozen brain in animals which were tested by both methods.

Agreement among tests

A summary of the overall (OPA), positive (PPA) and negative (NPA) percent agreements among diagnostic tests for all species combined and for OPA for each species alone, is presented in Table 2.7. Test agreement was not examined by location. For the 51 birds selected in this study, the overall percent agreements among tests (histopathology, immunohistochemistry, rtRT-PCR on frozen brain and rtRT-PCR on FFPE tissue) ranged from 70.6% to 92.2%, with the highest overall percent agreement being between

histopathology and rtRT-PCR on FFPE samples (all species considered together). PPA ranged from 65.9% to 96.6%, and was above 90% for rtRT-PCR (FFPE) and histopathology, rtRT-PCR (FFPE) and rtRT-PCR (frozen brain), IHC and rtRT-PCR (FFPE), and IHC and rtRT-PCR (frozen brain). NPA ranged from 0 to 100%, with 100% agreement between histopathology and IHC, and histopathology and rtRT-PCR (FFPE).

Table 2.7. Summary of overall, positive and negative percent agreements among diagnostic tests in Canada geese, trumpeter swans and mute swans evaluated for the presence of avian bornavirus.

CAGO TRSW MUSW All species combined (n*=51) (n*=41) (n*=8) (n*=2)

PPA NPA OPA OPA OPA OPA

histopathology vs IHC 65.9 (29/44) 100 (7/7) 70.6 (36/51) 70.7 (29/41) 75.0 (6/8) 50.0 (1/2)

histopathology vs RT-PCR (frozen brain) 88.2 (15/17) NA (0/0) 88.2 (15/17) 93.7 (15/16) NA 0 (0/1)

histopathology vs RT-PCR (FFPE) 90.9 (40/44) 100 (7/7) 92.2 (47/51) 92.7 (38/41) 87.5 (7/8) 100 (2/2)

IHC vs RT-PCR (frozen brain) 92.9 (13/14) 33.3 (1/3) 82.4 (14/17) 81.2 (13/16) NA 100 (1/1)

IHC vs RT-PCR (FFPE) 96.6 (28/29) 45.5 (10/22) 74.5 (38/51) 73.2 (30/41) 87.5 (7/8) 50.0 (1/2)

RT-PCR (frozen brain) vs RT-PCR (FFPE) 93.3 (14/15) 0 (0/2) 82.4 (14/17) 87.5 (14/16) NA 0 (0/1)

*n represents the total of birds selected in this study; CAGO: Canada goose; TRSW: trumpeter swan; MUSW: mute swan Agreement tests: PPA: positive percent agreement; NPA: negative percent agreement; OPA: overall percent agreement IHC: immunohistochemistry; RT-PCR: reverse transcription polymerase chain reaction; FFPE: formalin-fixed paraffin-embedded tissue NA: not able to calculate the percentage (denominator equal to 0) or no frozen brain available (for TRSW)

Cases submitted from the CCWHC-Atl

All six Canada geese cases received from CCWHC-Atl had histological lesions consistent with ABV based on their initial histopathology report and reassessment of tissues that were available at the time of this review. Blocks of paraffin embedded brain were not available from the two cases included in Daoust et al., 1991; testing was thus carried out on sections of spinal cord showing inflammatory lesions from one bird (A) and peripheral nerve without lesions in the second (B). The results of testing for these two birds are as follows: A - rtRT-PCR positive, IHC negative; B - rtRT-PCR inconclusive, IHC negative. Blocks of paraffin embedded brain were available from three additional cases, all three were positive by rtRT-PCR and two were also positive by IHC. A block of paraffin embedded spinal cord was available from one additional case, and was positive by rtRT-PCR and negative by IHC.

Birds negative for inclusion criteria

At the commencement of the study, six Canada geese, two trumpeter swans and one mute swan were selected from the CCWHC-Ont database as negative controls, based on the absence of the previously defined inclusion criteria. All birds were negative on histopathology and IHC. Frozen brain was available from four birds (one Canada goose, two trumpeter swans, one mute swan) and all four samples were negative for ABV on conventional RT-PCR. A sample from only one of these cases was available for re-testing with rtRT-PCR, and was again negative. Formalin-fixed paraffin embedded tissues were negative for ABV by rtRT-PCR in 5 Canada geese, and were positive in one Canada goose (Cp = 36.09), two trumpeter swans (Cp = 27.33 and 32.20) and one mute swan (Cp = 36.25).

Sequence analysis

Seven RT-PCR products from positive Canada geese were sequenced and their nucleotide sequences were 100% identical amongst themselves. Using pairwise sequence comparison based on amino acid sequence, these N gene sequences were compared with 68

the available ABV sequences in GenBank and a phylogenetic tree based on the amino acid sequences generated (Figure 2.3). The N gene sequences from Ontario geese were 100% identical to a Canada goose sequence from the United States (US) (Gen Bank # HQ123585.1) and 100% and 98.1% identical to two mute swan sequences from the US (Gen Bank # JQ687271.1 and JQ687270.1, respectively). Our Ontario geese N gene sequences were 84.6% (GenBank # FJ002315.1-ABV genotype 3) to 87.5% (GenBank # FJ002320.1, HC356270.1, HC356265.1, all ABV-genotype 2) identical to sequences from parrots; 93.1% identical to sequence from a canary (GenBank # GQ161095.1); and 85.6% identical to various mammalian Borna virus sequences available in the GenBank (GenBank # EU095836-S05-1178-sheep, EU095835-MP68-shrew, AY11416-CRNP5 - human, AY374525-H3452-horse).

Figure 2.3. Phylogenetic tree illustrating the genetic relationships between the waterfowl bornavirus and other bornaviruses, based on a partial amino acid sequence of the nucleocapsid protein gene. Representatives of the five known avian bornavirus (ABV) genotypes, the canary genotype and several representatives of Borna disease virus (BDV) are included. Sequences are identified with GenBank accession number followed by species (if non-psittacine). The seven Canada goose sequences originating in this study are highlighted in grey. The scale bar indicates the genetic distance among clusters. CAGO: Canada goose; MUSW: mute swan.

DISCUSSION

The objective of this study was to retrospectively identify cases of ABV infection in wild waterfowl species and to characterize associated pathology in cases with the presence of clinical signs or pathological lesions that would suggest ABV infection in psittacines. ABV was detected by RT-PCR in 78.4% (40/51) of the selected cases, indicating the effectiveness of our criteria. Birds infected with ABV but not showing the specified inclusion criteria would not have been chosen, thus our method was not appropriate to estimate the prevalence of infected birds within the databases. Other authors have looked for infection in asymptomatic waterfowl species and reported prevalences from 5 to 50% depending on species, location and type of sample (Payne et al., 2011a; Guo et al., 2012). The percentage of ABV infected birds selected from our pathology databases was thus significantly higher than the background level observed in apparently healthy flocks.

Pathological findings in ABV infected birds were very similar to those typically found in parrots affected with PDD (Shivaprasad et al., 1995; Berhane et al., 2001; Gancz et al., 2010). Although both neurological signs and gastrointestinal impaction were seen associated with ABV infection, the former was more consistently identified within this group of cases. Non-suppurative perivascular cuffing in the central nervous system was dramatic and widespread in the majority of birds; however, lesions in peripheral nerves and in the autonomic system could be very subtle and had often been missed on initial histopathologic assessment. Many cases showed a marked degree of autolysis, making evaluation of the gastrointestinal tract difficult, and inconsistent sample collection also affected assessment of lesion distribution. Lesions in the nerves and ganglia of the proventriculus and ventriculus were; however, present in a significantly greater proportion of Canada geese as compared to trumpeter and mute swans, perhaps indicating a difference in pathology among species.

Overall, the results of the diagnostic tests used in this study (histology, IHC, RT- PCR on tissues, RT-PCR on FFPE) appeared well-correlated. Actual assessment and comparison of specificity and sensitivity is difficult due to the absence of a recognized

gold standard for ABV infection, and to the identification of a true control group of uninfected birds. Test agreement was thus estimated using overall, positive, and negative percent agreement (Feinstein et al., 1990). In this study, histopathological lesions, with an emphasis on careful examination of a broad selection of nerves and ganglia, correlated strongly with the presence of virus in brain. Using histopathology alone would likely result in false positives, particularly if only brain is examined, as non-suppurative encephalitis is common to several neurotropic viral diseases. Lesions in the autonomic nerves and ganglia and the adrenal do appear to be more specific to ABV infection. Histopathology would not be a sensitive method of detecting virus infected birds as, in selecting negative controls, we identified several birds that were ABV positive but without clinical or pathological abnormalities. IHC and RT-PCR would be more appropriate for this purpose.

The results of this study also demonstrate that RT-PCR and IHC designed to identify psittacine ABV also recognize the strain of ABV found in waterfowl. The positive percent agreement between IHC and rtRT-PCR, performed on FFPE samples from the same block, was excellent. However, IHC did appear less sensitive than rtRT- PCR, and thus false negative results could occur with IHC if the viral load in the tissue is very low or as the result of antigen degradation. Non-specific immunohistochemical staining was observed in several cases and can be a source of false positive IHC results. In this study, we used strict criteria (the presence of strong and diffuse intranuclear staining in neurons or glial cells) to define IHC positivity (Raghav et al., 2010).

Both gel-based conventional RT-PCR (ABV-N gene) and rtRT-PCR (ABV-M gene) were used as a result of changes in diagnostic laboratory protocols during the study. This provided us with the opportunity to compare the results of the two testing methods. Results disagreed in only one case (of the 17 frozen brains tested), which was positive on rtRT-PCR but inconclusive on conventional RT-PCR. In parrots, assays for the M and N genes appear to have a similar high sensitivity (Gancz et al., 2010; Raghav et al., 2010); while the nucleoprotein N gene is the more highly expressed region of the genome, the matrix M gene is the more highly conserved region (Kistler et al., 2008).

In our laboratory, conventional RT-PCR using the N gene was not successful using FFPE tissues, whereas rtRT-PCR for the M gene was suitable for use with FFPE samples. Extraction of RNA from FFPE can result in short fragments of nucleic acids that may not be suitable for amplification in PCR assays that produce large amplicons. The use of different primers to amplify a shorter fragment in our rtRT-PCR likely explains the success of this method. The ability to perform rtRT-PCR on FFPE samples is crucial in the retrospective evaluation of disease, where frozen samples are rarely available.

IHC and PCR were carried out on brain, or spinal cord or peripheral nerve in the few cases where brain was not available. The decision to test only one tissue was a pragmatic one. Brain was selected due to the frequency of lesions in that tissue as well as the results of a study in psittacine birds that evaluated the presence of ABV in multiple tissues (Raghav et al., 2010). A comparison of the tissue distribution of ABV in waterfowl with that in parrots would be interesting; based on histopathologic lesions we predict that it would be equally widespread.

The proportion of ABV infected diseased birds was not equal in the two databases searched. The proportion of Canada geese was significantly higher at the Toronto Zoo when compared to goose cases arising elsewhere in Ontario. Recent publications about the prevalence of ABV in Canada geese and mute swans in the United States also showed that infections were often clustered within flocks and the prevalence of ABV appears to vary greatly among locations (Payne et al., 2011a; Guo et al., 2012). The Toronto Zoo goose population of approximately 150 free-flying individuals is composed of resident birds remaining on the zoo grounds year-round and migrating birds coming from the Arctic or leaving for the Great Lakes for the winter. Determining whether there is a truly higher prevalence of disease at the zoo site is made difficult by enhanced surveillance at the zoo. Numerous members of staff and the public are available to observe ill or dead birds, and a thorough necropsy is performed on every wild bird euthanized or found dead. Through the rest of Ontario, passive surveillance is carried out at a much lower rate and in a much more random manner and more often involves the submission of birds found dead without any clinical history. The prevalence of other diseases commonly found in Ontario waterfowl, such as lead poisoning or botulism, are very rarely encountered on the

zoo site but constitute a significant percentage of death elsewhere in Ontario, thus diluting the importance of ABV as a cause of mortality in the greater population. Finally, the Toronto Zoo environment may provide opportunities for ABV to transmit and propagate among birds allowing virus loads to increase and permit a disease cluster to develop. Greater food availability may also allow a sick bird to survive longer than it would in the wild, increasing the opportunity for viral shedding and environmental contamination, as well as the likelihood of being observed and identified as ill. These hypotheses are supported by that fact that the proportion of infected diseased trumpeter swans was not different between the birds found at Toronto Zoo and found elsewhere; this species is only transient on the zoo site, thus all birds should be considered as the same population. The relatively high prevalence of ABV-infected Canada geese at the zoo is of some concern for the zoo’s captive collection; however, there has been no report of PDD-lesions or ABV infection in any bird in the Toronto Zoo collection over the past 20 years. The possibility of transmission of ABV to other waterfowl species, to different avian families housed in outside aviaries and possibly to some mammals at the zoo remains uncertain.

As mentioned previously, ABV may be present in waterfowl that do not appear to suffer from clinical disease (Payne et al., 2011a; Guo et al., 2012). However, no necropsies were performed on any of the birds in these studies so it is possible that subclinical nervous system disease may have been present. Widespread asymptomatic infection with less common clinical disease is a consistent feature of bornavirus infections, and may be attributed to the non-cytopathic effect of Bornaviridae. This pattern has been documented with BDV in horses and sheep (Lipkin et al., 2006) and is now well-recognized in parrots (de Kloet et al., 2009; Lierz et al., 2009; Kistler et al., 2010; Villanueva et al., 2010; Heffels-Redmann et al., 2012). The exact circumstances under which disease develops in waterfowl species is unknown but influences are likely multifactorial. The existence of frequent asymptomatic birds may also explain our difficulty in identifying negative controls in this study. A small number of birds was selected based on the absence of inclusion criteria to act as negative controls for the IHC and PCR testing. However; one Canada goose, two trumpeter swans and one mute swan

within this group were positive for ABV by RT-PCR on FFPE tissues. It is of interest to note that their mean Cp (33.0) was significantly higher (ie. lower viral load) than the mean Cp of diseased birds (24.2). Further detailed evaluation of asymptomatic birds is necessary to estimate the true prevalence of ABV infection and its relationship to the presence of clinical disease in these species.

A non-lethal method of reliably testing for ABV would assist in identifying the prevalence of infection in populations, particularly those such as the trumpeter swan that are the subject of conservation programs. PCR testing of feces and cloacal swabs is used in psittacine medicine, but appears to be relatively insensitive based on studies of experimentally and naturally infected parrots (Villanueva et al., 2010; Payne et al., 2011b). In our study, only one out of three cloacal swabs collected from ABV-positive geese was positive by RT-PCR, supporting the presence of intermittent fecal shedding.

The partial nucleotide sequences of the ABV-N gene recovered from seven Canada geese in this study are highly conserved among individuals and closely match recently published sequences from Canada geese and mute swans in the United States (Payne et al., 2011a; Guo et al., 2012). This unique group of ABV sequences significantly diverges from the six genotypes of ABV previously found in psittacines (Kistler et al., 2008; Rubbenstroth et al., 2012), from the one described in a canary (Weissenböck et al., 2009b), and from mammalian BDVs. This confirms that the waterfowl bornaviruses represent a separate, independent cluster within the family Bornaviridae, now referred as ABV-CG (Delnatte et al., 2011; Payne et al., 2011a). Payne et al. recently showed that the genome organization of ABV-CG is in fact more closely related to mammalian BDV that to other ABVs, with the presence of a regulatory open reading frame in the N/X intergenic region that is lacking in psittacine ABVs (Kistler et al., 2008; Rinder et al., 2009; Payne et al., 2011a; Guo et al., 2012). This retrospective evaluation only looked at waterfowl entered into the databases since 1992; however, we found ABV in the tissues of a bird described by Daoust and submitted to the CCWHC-Atl in 1988 (Daoust et al., 1991). The finding of ABV in older archival material would support the premise that this virus has been long present in North American waterfowl populations and is not a recent introduction, as the psittacine strains

appear to be. As BDV sequences obtained from various species over several decades have shown remarkable sequence conservation (Durrwald et al., 2006; Kistler et al., 2008), the recognition of an increasing number of avian bornaviruses is surprising and makes molecular characterization a critical component in the understanding of the epidemiology of ABV infection.

In conclusion, this study confirmed that ABV is present in wild Canada geese, trumpeter swans and mute swans in Canada and showed evidence that it can be associated with significant disease (Smith et al., 2010; Delnatte et al., 2011). Although our findings strongly suggest that ABV is the cause of the clinical and pathological changes observed, formal proof of a causal relationship typically requires fulfilment of Koch's postulates. This was demonstrated for ABV / PDD in several species of parrots, but has yet to be carried out in waterfowl species (Gray et al., 2010; Mirhosseini et al., 2011; Payne et al., 2011b; Piepenbring et al., 2012). ABV infection should, nonetheless, be included in the differential diagnosis for any Anseriform exhibiting upper gastrointestinal impaction or non-specific neurological signs and having non-suppurative inflammation of the central, peripheral, or autonomic nervous systems. The overall impact of this disease on the health and size of populations of wild waterfowl in North America is yet to be determined.

CHAPTER 3: PROSPECTIVE STUDY

Seroprevalence and prevalence of cloacal shedding of avian bornavirus

in free-ranging waterfowl

Pauline Delnatte1,2, Éva Nagy1, Davor Ojkic3, David Leishman1, Graham Crawshaw2,

Kyle Elias1, and Dale A Smith1*

1 Ontario Veterinary College, University of Guelph, Guelph, Ontario, Canada, N1G 2W1

2 Toronto Zoo, Toronto, Ontario, Canada, M1B 5K7

3 Animal Health Laboratory, University of Guelph, Guelph, Ontario, Canada, N1G 2W1

Keywords: avian bornavirus, Canada goose, mallard, mute swan, serology, shedding, trumpeter swan, waterfowl

Submitted to Journal of Wildlife Diseases.

ABSTRACT

The objectives of this study were to survey wild Canada geese (Branta canadensis), trumpeter swans (Cygnus buccinator), mute swans (Cygnus olor) and mallards (Anas platyrhynchos) to estimate the seroprevalence and the prevalence of cloacal shedding of avian bornavirus (ABV) in southern Ontario, Canada. Blood samples and cloacal swabs were collected from 206 free-ranging Canada geese, 135 trumpeter swans, 75 mute swans and 208 mallards at different locations between October 2010 and May 2012. Sera were assessed for the presence of antibodies against ABV by an enzyme- linked immunosorbent assay and swabs were evaluated for the presence of virus using real-time reverse transcription polymerase chain reaction. Serum antibodies were present in birds from all four species and at each sampling site. Thirteen percent of the geese caught on the Toronto Zoo site shed ABV in feces compared to none of the geese sampled at three other locations in Ontario. The prevalences of shedding of ABV in mute swans, trumpeter swans and mallards were 9.3%, 0% and 0%, respectively. Birds that were shedding the virus were more likely to have antibodies against ABV and to have higher antibody levels than those that were not, although many birds with antibodies were not shedding the virus. This study confirmed that exposure to or infection with ABV is widespread in asymptomatic free-ranging waterfowl in Canada, however the correlation between cloacal shedding, presence of antibodies and presence of disease is not fully understood.

INTRODUCTION

The waterfowl genotype of avian bornavirus was first identified in several free- ranging Canada geese (Branta canadensis) and trumpeter swans (Cygnus buccinator) in 2009 in southern Ontario, Canada, where it was associated with non-suppurative encephalitis and mortality (Smith et al., 2010; Delnatte et al., 2011). This was also the first identification of ABV in any species of wild bird. A retrospective evaluation of 955 necropsy cases from free-ranging Canada geese, trumpeter swans and mute swans (Cygnus olor) in Ontario confirmed that the presence of ABV in brain, as assessed by immunohistochemistry (IHC) and reverse transcription polymerase chain reaction (RT- PCR), is strongly correlated with the presence of lymphoplasmacytic inflammation in the central, peripheral and autonomic nervous tissues (Delnatte et al., 2013). These histopathologic lesions are very similar to those seen in parrots affected with proventricular dilation disease (PDD), which is caused by a number of psittacine- associated genotypes of ABV (Gancz et al., 2010). More recently, ABV infection has been shown to be widespread in wild birds across the United States. The virus has now been identified in a variety of waterfowl and gull species using oropharyngeal/cloacal swabs and brain samples (Payne et al., 2011a; Guo et al., 2012, Payne et al., 2012). Partial nucleotide sequences of ABV identified in Canadian and US waterfowl are very similar, sharing 96 to 100% nucleotide identity, and cluster separately from those described in psittacine birds and canaries and from mammalian Borna disease virus (BDV) (Payne et al., 2011a; Guo et al., 2012; Payne et al., 2012; Delnatte et al., 2013). The bornaviruses circulating among North American free-living waterfowl thus appear to form a unique genotype (ABV-CG genotype).

The prevalence and distribution of ABV infection in wild waterfowl in southern Ontario are not known. In our retrospective necropsy study (Delnatte et al., 2013), the proportion of wild Canada geese affected by ABV-related neurological disease was significantly higher in birds originating from the Toronto Zoo site as compared to birds found elsewhere in the province. The apparent prevalence also differed among species examined. The prevalence of ABV infected birds in the United States ranged from 0 to

50% depending on the species and sampling sites, also supporting uneven geographic and species distribution of the virus (Guo et al., 2012; Payne et al., 2012).

The impact of ABV infection on waterfowl populations is unknown. While significant pathology was described in our retrospective necropsy study (chapter 2), ABV positive birds randomly sampled in the United States were described as healthy (Payne et al., 2011; Guo et al., 2012). As no clinical examinations or necropsies were performed on these birds, it remains possible that subclinical nervous system disease may have been present.

Avian bornavirus infection has been studied best in psittacine birds where investigations of PDD outbreaks have shown that ABV is often detectable in cloacal swabs and in the urofeces of affected birds using RT-PCR. Estimating the infection rate of ABV in an avian population is problematic as shedding can be intermittent and somewhat unpredictable (Raghav et al., 2010; Villanueva et al., 2010; Payne et al., 2011b). Recent research has shown that the greatest amount of virus may actually be excreted in urine (Heatley et al., 2012).

Various RT-PCR primers have been used to target different ABV genes. While the nucleoprotein (N) gene is the most highly expressed region of the genome, the matrix (M) gene appears to be the most highly conserved region (Kistler et al., 2008). Given the genetic heterogeneity among ABVs, designing primers able to detect a broad range of ABV variants can be challenging (Rinder et al., 2011). The results of our retrospective study demonstrated that RT-PCR primarily designed to identify psittacine ABV and targeting either the N or the M gene also recognize the waterfowl genotype of ABV (Delnatte et al., 2013).

Identification of serum antibodies (Ab) can be used to estimate prevalence of exposure to ABV within a population. Antibodies to ABV have been detected in experimentally-infected parrots and Khaki Campbell ducks (Gray et al., 2010; Villanueva et al., 2010; Hoppes et al., 2013). To date, serologic assays described for the identification of ABV-specific Ab in psittacine birds include indirect

immunofluorescence assay, Western blot analysis and indirect enzyme-linked immunosorbent assay (ELISA) (Hoppes et al., 2013).

In order to detect exposure to or infection with ABV in a population of wild birds, we chose to use a combination of serology and RT-PCR on cloacal swabs. Our objectives were to survey wild Canada geese, trumpeter swans, mute swans and mallards (Anas platyrhynchos) to estimate the seroprevalence of ABV and the prevalence of cloacal shedding in southern Ontario.

MATERIAL AND METHODS

Study area and sample collection

Cloacal swabs were collected from 206 Canada geese, 208 mallards, 75 mute swans and 135 trumpeter swans in southern Ontario between October 2010 and May 2012. Exact sampling locations are mapped in Figure 3.1, and global positioning system (GPS) coordinates of the main capture sites are presented in Table 3.1. For each of the Ottawa (mallard) and Toronto and Peterborough (Canada goose) locations, two to three sites were used within a 15 km radius and were considered as a single location. Geese and ducks were captured and handled by the Canadian Wildlife Service (CWS) or by the Ontario Ministry of Natural Resources (OMNR) as part of their annual banding exercises, except for the Canada geese at the Toronto Zoo that were specifically captured for sampling. Goose and duck capture sites were pre-defined in order to assess the possibility of an uneven geographic distribution of the virus among locations. In contrast, swan samples were collected by biologists and volunteers from the Trumpeter Swan Reintroduction Program at locations within the swans’ winter range at time points when birds could be baited into catching areas, and when members of the management team were available.

Figure 3.1. Study area in southern Ontario, Canada, showing sampling locations for Canada geese (Branta canadensis), mute swans (Cygnus olor), trumpeter swans (Cygnus buccinator) and mallards (Anas platyrhynchos) from which cloacal swabs and blood samples were analyzed to determine the prevalence of avian bornavirus infection, 2010-2012. 82

Table 3.1. Capture dates and locations and demographic information for Canada geese (Branta canadensis), mallards (Anas platyrhynchos), mute swans (Cygnus olor) and trumpeter swans (Cygnus buccinator) in southern Ontario, Canada from which blood samples and cloacal swabs were collected between October 2010 and May 2012 to determine the prevalence of avian bornavirus infection.

a b Sampling location Number Age class Sex Species Sampling period Area i GPS coordinates of birds HY AHY U Male Female U

Canada Guelph N43° 33', W80° 13' June 2011 50 7 43 0 29 21 0 goose Torontoc ~ N43° 39', W79° 22' June 2011 48 12 36 0 18 27 3 June 2011 (50) / Spring 2012 Toronto Zoo N43° 49', W79° 11' 56 0 56 0 17 37 2 (6) Peterboroughc ~ N44° 29', W78° 29' June 2011 52 20 32 0 32 20 0 206 39 167 0 96 105 5

Mallard Komoka N42° 57', W81° 26' Sept 2011 104 83 21 0 67 37 0 Ottawac ~ N45° 12', W75° 51' Sept 2011 104d 20 37 47 71 23 10 208 103 58 47 138 60 10

Mute LaSalle Park N43° 18', W79° 51' Winter 2011-2012 74 0 74 0 48 26 0 swan Hamilton N43° 18', W79° 48' Winter 2011-2012 1 0 1 0 1 0 0 75 0 75 0 49 26 0 Winter Winter

2010-2011 2011-2012 Trumpeter LaSalle Park N43° 18', W79° 51' 22 72 94 26 68 0 59 35 0 swan Bluffers Park N43° 42', W79° 14' 12 3 15 7 8 0 7 8 0 Aurora N44° 00', W79° 29' 7 0 7 2 5 0 2 5 0 Others various 4 15 19 7 12 0 10 7 2 45 90 135 42 93 0 78 55 2

GPS = global positioning system; a HY = hatching year; AHY = after hatching year; U = unknown; b U = unknown; c 2-3 locations sampled within a 15km radius - GPS coordinates indicate the average coordinates; d only cloacal swabs collected (no blood). 83

The birds were captured using different techniques. Canada geese were caught by baiting flocks into capture areas and then herding molting, flightless birds into drive traps; mallards were caught in baited swim-in traps; and mute and trumpeter swans were caught by baiting and then manual capture at winter feeding stations. Capture methods and sample collection were approved by the University of Guelph Animal Care Committee in accordance with the requirements of the Animals for Research Act of Ontario and the recommendations of the Canadian Council for Animal Care. A CWS Migratory Bird Research Permit was also obtained.

For each bird captured, the species, bird’s identification (band and/or wing tag numbers), sex and age class (when available), and capture date and location were recorded. A summary is presented in Table 3.1. Sex and age class were determined according to the visualization of a phallus when the cloaca was everted and/or based on plumage. Most trumpeter swans are banded during their first year and thus exact age was often available.

Blood samples were collected from the medial metatarsal vein from all mute swans (n=75), from the majority of trumpeter swans (n=130) and Canada geese (n=203), and from 92 mallards. Sera were separated and stored at -20oC. Cloacal swabs were collected by opening the cloacal orifice, inserting a sterile tipped applicator swab 1-2 cm into the cloaca and swabbing the cloacal mucosa two to four times using gentle pressure in a circular motion. Any large (more than 0.5 cm) pieces of feces were shaken off and the swab was placed in a virus transport medium tube (Multitrans System, Starplex Scientific Inc., Etobicoke, Ontario, Canada). Swabs were frozen at -20oC until being shipped to Guelph and then held at -80oC until testing (up to 1.5 years).

Antigen preparation

A pET21a plasmid based expression cassette for the M24N protein was kindly provided by Dr. Susan Payne, Schubot Exotic Bird Health Centre, Texas A&M University. The isolated plasmid DNA was transformed into BL21 cells (DE3) (Novagen pET system manual, 10th Ed., 2003). Protein purification was performed with the Qiagen 84

Ni-NTA purification system according to the manual (Qiagen, Dusseldorf, Germany). The purified protein was analyzed by SDS-PAGE and protein concentration was determined by the Bio-Rad Protein Assay (Bio-Rad, Hercules, California, USA). The detailed protocol of the protein purification is provided in the Appendix C.

Serology

An ELISA was developed to estimate ABV specific Ab in serum samples. ABV- N protein was purified as described above, and diluted in a carbonate buffer (35 mM

NaHCO3, 15 mM Na2CO3 [pH 9.6]). Immulon 2HB 96-well microtiter plates (Dynex Technologies Inc., Chantilly, Virginia, USA) were coated with antigen at a concentration of 100 ng/well and incubated at 4oC for 16 h. The plates were then blocked with 3% bovine serum albumin at 37oC for 75 minutes. The serum samples were heat-treated for 30 minutes at 56oC. Serum samples were diluted to 1:20 in a wash buffer (0.05% Tween 20 in PBS) and 100 µl were added to each well and incubated for 1 hour at 37oC. Each reaction was performed in duplicate wells. The plates were washed four times with wash buffer between each step. One hundred µl of horseradish peroxidase-labelled goat anti- bird IgG heavy and light chain antibody conjugate (Bethyl Laboratories Inc., Montgomery, Texas, USA) at a dilution of 1:5000 was added and incubated for 75 minutes at 37oC. The colour was developed with ABTS peroxidase substrate system (Kirkegaard & Perry Laboratories, Gaithersburg, Maryland, USA) until the average optical density (OD) of the two positive control wells reached 0.8, read in a BIO-Tek ELISA microplate reader (Bio-Tek Instruments, Winooski, Vermont, USA) at 405nm.

When the OD405 between two duplicates differed more than 15%, the sample was retested.

The ABV specific antibody responses were determined by calculating the sample- to- positive (S/P) ratio. Sample-to-positive ratio was calculated as: [sample mean – negative control mean] / [positive control mean – negative control mean]. Mean group S/P values (per species or per location) were obtained from individual S/P ratios. Serum

from a domestic goose (Anser anser domesticus, see below) and from a Canada goose that was shedding ABV were the negative and positive controls, respectively.

Controls and determination of cut-off for seropositivity

Setting cut-offs for determination of seropositive and seronegative birds was hindered by a lack of gold standard for determining ABV infection and a lack of known positive and negative birds for each species studied. Serum samples were obtained from 13 domestic geese and 13 domestic ducks (Anas platyrhynchos domesticus) that were part of a larger experimental infection trial (See chapter 5). The goslings and ducklings were obtained at one day of age from a commercial supplier and maintained in an isolation facility. Blood collected at 40, 55 and 70 days (39 samples total for each of the two species) was evaluated for Ab to ABV by ELISA as described above. These groups of birds were determined to be free of ABV infection using histology, IHC and RT-PCR on necropsy samples, and thus used as negative controls. The mean S/P ratio for the 39 goose samples was -0.525 with a standard deviation (SD) of 0.470; the mean S/P ratio for the 39 duck samples was 2.154 with a SD of 0.731. Field samples were considered positive if the S/P ratio of the sample was higher than the mean S/P ratio for the negative control group used plus 2 SD (Ojkic et al., 2003). Calculations of sample positivity were also performed using 3 SD in order to provide a second estimate of the seroprevalence with a lower sensitivity but higher specificity. Negative control domestic geese were used for the calculation of seropositivity cut-offs for the goose and swan samples; samples were considered positive if the S/P ratio was higher than 0.416 (2SD) or 0.886 (3SD). Negative control domestic ducks were used for the calculation of the seropositivity cut- offs for the mallard samples; samples were considered positive if the S/P ratio was higher than 3.616 (2SD) or 4.346 (3SD).

Nucleic acid isolation and RT-PCR

Cloacal swabs were evaluated for the presence of ABV by real time RT-PCR detection of the ABV-M gene. Total nucleic acids were extracted from 50-µl aliquots of 86

inoculated virus transport medium using MagMAX-96 Viral RNA Isolation Kit in a MagMAX Express-96 Magnetic Particle Processor (Applied Biosystems Inc., Foster City, California, USA) according to manufacturer's instructions. The RT-PCR assay was a duplex test with two sets of primers and TaqMan probes: (1) ABV_M_120201 – targeting psittacine ABV-M gene sequences; and (2) ABVG_M_111029 – targeting geese ABV-M gene sequences. The amplification was carried out in 25 µl reactions in a LightCycler 480 Real-Time PCR System (Roche, Laval, Quebec, Canada) using AgPath- ID One-Step RT-PCR Kit (Applied Biosystems, Foster City, California, USA). Primers and cycling parameters for ABV RT-PCR were those previously described (Delnatte et al., 2013). A RT-PCR crossing point value (Cp) less than 37.00 was considered positive, between 37.00 and 39.99 inconclusive, and equal or greater than 40.00 negative.

Statistical analysis

Statistical analyses were performed using Graph-Pad QuickCalcs software (http://www.graphpad.com/quickcalcs/). We strived to achieve a sample size of 200 birds of each species based on a 95% confidence interval for detection of a disease at an estimated prevalence of 0 to 4% (Dohoo et al., 2009). The prevalence of cloacal shedding, seroprevalence and their 95% confidence intervals (CI) were calculated separately for each species and at each location. Prevalence of shedding was defined as the percentage of birds sampled for which the cloacal swab tested positive by RT-PCR. For statistical analysis, inconclusive RT-PCR results were considered negative. Fisher’s exact test was used for comparisons of prevalences among species, among different locations, between shedders versus non-shedders, between age classes and between sexes. When the difference in prevalence was significant (P<0.05), the p values, odds ratio and 95% CI were calculated. Unpaired t tests were used to compare the mean Cp values (RT-PCR) of shedding birds between species and to compare the mean S/P ratios (ELISA) among different locations, between shedders and non-shedders, between age classes and between sexes. Two-tailed P<0.05 was considered significant.

RESULTS

Seroprevalence

The ELISA results for Ab against ABV in serum samples are summarized in Table 3.2 through presentation of mean S/P ratios and seroprevalences using the 2SD and 3SD seropositivity cut-offs. A total of 500 serum samples were analysed. The overall seroprevalence of ABV infection was 58% (117/203) in Canada geese, 71% (53/75) in mute swans, 62% (81/130) in trumpeter swans and 23% (21/92) in mallards using the 2SD seropositivity cut-off and was 14% (29/203) in Canada geese, 23% (17/75) in mute swans, 35% (46/130) in trumpeter swans and 6.5% (6/92) in mallards using the 3SD seropositivity cut-off.

Table 3.2. Prevalence of cloacal shedding of avian bornavirus determined by real time reverse transcription polymerase chain reaction and prevalence of antibodies against avian bornavirus determined by an enzyme-linked immunosorbent assay in 206 Canada geese (Branta canadensis), 208 mallards (Anas platyrhynchos), 75 mute swans (Cygnus olor) and 135 trumpeter swans (Cygnus buccinator) caught in southern Ontario, Canada from October 2010 to May 2012.

Seroprevalence Prevalence of Sampling Species cloacal mean Seropre- Seropre- location a shedding N S/P valence valence ratiob (2SD)c (3SD)d

Canada Guelph 0% 0/50 50 0.370 50% 2.0% goose Toronto 0% 0/48 48 0.519 65% 10%

Toronto Zoo 13% 7/56 53 0.557 51% 17%

Peterborough 0% 0/52 52 0.632 65% 27%

Overall 3.4% 7/206 203 0.521 58% 14%

Mallard Overall 0% 0/208 92 2.613 23% 6.5%

Mute swan Overall 9.3% 7/75 75 0.636 71% 23 %

Trumpeter Overall 0% 0/135 130 0.645 62% 35% swan

a N = number of birds from which a blood sample was available; b S/P ratio = sample-to-positive ratio calculated as: [sample mean of optical absorbance – negative control mean] / [positive control mean – negative control mean]; c Sample considered positive if its S/P ratio was higher than the mean S/P ratio of the negative control group plus 2 standard deviations; d Sample considered positive if its S/P ratio was higher than the mean S/P ratio of the negative control group plus 3 standard deviations.

The S/P ratio of individual samples by species (Fig. 3.2) and by location for Canada geese (Fig. 3.3) are presented in Figures 3.2 and 3.3 with both 2SD and 3SD seropositivity cut-offs indicated for goose/swan samples and for duck samples. Sample- to-positive ratios (all species considered together) ranged from -1.024 to 5.288 with a mean (± standard deviation) of 0.521 (± 0.474) for Canada geese, 0.636 (± 0.478) for mute swans, 0.645 (± 0.626) for trumpeter swans, and 2.613 (± 1.09) for mallards. Antibodies were found in all four species and at each location.

Using the 2SD cut-off values, the seroprevalence in mallards was significantly lower than that for the other three species of birds examined (Table 3.3). Using the 3SD cut-off values, the seroprevalence in trumpeter swans was significantly higher than in Canada geese and in mallards, and the seroprevalence in mute swans was significantly higher than in mallards (Table 3.3).

There were no significant differences in seroprevalence among Canada geese from the four sampling locations using the 2SD seropositivity cut-off. Using the 3SD cut- off, birds sampled at Peterborough had a significantly higher seroprevalence than those caught at Toronto and at Guelph, and Canada geese sampled at Toronto Zoo had a significantly higher seroprevalence than geese caught at Guelph (Table 3.3).

Female Canada geese sampled at Toronto Zoo had a significantly higher seroprevalence than males using the 3 SD cut-off (8/34 vs 0/17) (Table 3.3) and had a significantly higher mean S/P ratio (0.663 vs 0.256) (P=0.025). Male mallards had a significantly higher seroprevalence than females using the 2SD cut-off (18/60 vs 3/31) (Table 3.3). There were no effects of sex on seroprevalence or mean S/P ratio for any other species or at any other location.

When all species were considered together, adult birds (after hatch year) had a significantly higher seroprevalence than young birds (hatch year) using the 2SD cut-off (Table 3.3). There were no effects of age class on seroprevalence or mean S/P ratio if each species was considered separately.

Figure 3.2. Sample-to-positive ratio of serum samples assessed for the presence of antibodies against avian bornavirus using an enzyme-linked immunosorbent assay for 203 Canada geese (Branta canadensis), 75 mute swans (Cygnus olor) and 130 trumpeter swans (Cygnus buccinator) and 92 mallards (Anas platyrhynchos) in southern Ontario, Canada.

SD: standard deviation; a S/P ratio: sample to positive ratio calculated as: [sample mean of optical absorbance – negative control mean] / [positive control mean – negative control mean]; b Calculations of sample seropositivity for geese and swans were performed using the mean S/P ratio of the goose negative control group plus 2 and 3 standard deviations in order to provide two levels of sensitivity and specificity; c Calculations of sample seropositivity for ducks were performed using the mean S/P ratio of the duck negative control group plus 2 and 3 standard deviations in order to provide two levels of sensitivity and specificity ; d Sample considered positive if its S/P ratio is higher than the mean S/P ratio of the negative control group plus 2 standard deviations; e Sample considered positive if its S/P ratio is higher than the mean S/P ratio of the negative control group plus 3 standard deviations.

Figure 3.3. Sample-to-positive ratio of serum samples assessed for the presence of antibodies against avian bornavirus using an enzyme-linked immunosorbent assay for 203 Canada geese (Branta canadensis) at four different locations in southern Ontario, Canada.

CAGO: Canada goose; SD: standard deviation a S/P ratio: sample to positive ratio calculated as: [sample mean of optical absorbance – negative control mean] / [positive control mean – negative control mean]; b Calculations of sample seropositivity were performed using the mean S/P ratio of the negative control group plus 2 and 3 standard deviations in order to provide two levels of sensitivity and specificity; c Sample considered positive if its S/P ratio is higher than the mean S/P ratio of the negative control group plus 2 standard deviations; d Sample considered positive if its S/P ratio is higher than the mean S/P ratio of the negative c ontrol group plus 3 standard deviations.

odds Seroprevalence [cut-off used] P value 95% CI ratio CAGO vs MALL [2SD] <0.0001 4.6 2.62; 8.06

TRSW vs MALL [2SD] <0.0001 5.59 3.06; 10.2

MUSW vs MALL [2SD] <0.0001 8.15 4.06; 16.3

TRSW vs CAGO [3SD] <0.0001 3.29 1.93; 5.60

TRSW vs MALL [3SD] <0.0001 7.85 3.18; 19.4

MUSW vs MALL [3SD] 0.0032 4.2 1.56; 11.3

CAGO: Peterborough vs Guelph [3SD] 0.0004 18.1 2.27; 143

CAGO: Peterborough vs Toronto [3SD] 0.0432 3.17 1.04; 9.62

CAGO: Zoo vs Guelph [3SD] 0.016 10 1.22; 82.3

CAGO Zoo: female vs male [3SD] 0.04 11.2 0.610; 207

MALL: male vs female [2SD] 0.0356 4.14 1.12; 15.4

All species: adult vs young [2SD] 0.0003 2.04 1.39; 2.99

All species: shedder vs non-shedder [2SD] 0.0045 11.4 1.48; 87.8

All species: shedder vs non-shedder [3SD] <0.0001 11.3 3.47; 36.9

CAGO: shedder vs non-shedder [2SD] 0.022 11.7 0.66; 208

CAGO: shedder vs non-shedder [3SD] <0.0001 116 6.43; 2106

CAGO Zoo: shedder vs non-shedder [2SD] 0.01 19.4 1.05; 360

CAGO Zoo: shedder vs non-shedder [3SD] <0.0001 267 11.6; 6128

CAGO = Canada goose; CI = confidence interval; MALL = mallard; MUSW = mute swan; SD = standard deviation; TRSW = trumpeter swan.

Prevalence of viral shedding

The RT-PCR results for ABV in cloacal swabs are presented in Table 3.2. The overall prevalence (all locations considered together) of shedding was 3.4% (7/206) in Canada geese and 9.3% (7/75) in mute swans. Cloacal swabs from two geese had Cp values in the inconclusive range (Toronto Zoo site). None of the cloacal swabs from the 208 mallards or 135 trumpeter swans were positive.

The prevalences of shedding in Canada geese and in mute swans were significantly higher than in mallards and trumpeter swans (Table 3.4). The prevalence of cloacal shedding was not significantly different between Canada geese and mute swans (P=0.06). The Cp values of the 14 positive birds ranged from 24.61 to 36.89 with a mean and standard deviation of 31.95 (± 3.53). The mean Cp for the seven positive goose samples (29.67) was significantly lower than the mean Cp for the seven positive mute swan samples (34.23) (P=0.0087), indicating shedding of greater amounts of virus in the geese. The prevalence of ABV shedding in Canada geese caught on the Toronto Zoo site was 13% (7/56) and was significantly higher than the 0% prevalence found in geese sampled at Guelph, Toronto and Peterborough (Table 3.4).

Table 3.4. Results of statistical analysis, where significant, using Fisher’s exact tests to compare the prevalences of cloacal shedding among species and locations as part of a prospective survey of avian bornavirus infection in free-ranging waterfowl in southern Ontario, Canada.

Prevalence of shedding P value odds ratio 95% CI

CAGO vs MALL 0.0072 15.7 0.89; 276

MUSW vs MALL <0.0001 45.7 2.57; 810

CAGO vs TRSW 0.045 10.2 0.58; 180

MUSW vs TRSW 0.0006 29.7 1.67; 527

CAGO: Zoo vs Guelph 0.0136 15.3 0.85; 275

CAGO: Zoo vs Toronto 0.0144 14.7 0.82; 265

CAGO: Zoo vs Peterborough 0.0131 15.9 0.89; 286

CAGO = Canada goose; CI = confidence interval; MALL = mallard; MUSW = mute swan; TRSW = trumpeter swan

Correlation between cloacal shedding and seropositivity

The S/P ratio of each serum sample was plotted against the shedding status of the Canada goose (Fig. 3.4) or mute swan (Fig. 3.5) from which it was obtained (Figures 3.4 and 3.5). All Canada geese that were shedding ABV in cloacal swabs were seropositive using both seropositivity cut-offs. Six of seven mute swans that were shedding the virus were seropositive using the 2SD cut-off; however, only 3/7 (43%) were seropositive using the 3SD cut-off.

Amongst shedders, there was no direct correlation between individual Cp values and S/P ratio and no significant difference in mean S/P ratios between shedding Canada geese and shedding mute swans (1.686 vs 1.059) (P=0.11).

Figure 3.4. Sample to positive ratio of serum samples assessed for the presence of antibodies against avian bornavirus (ABV) using an enzyme-linked immunosorbent assay for 203 Canada geese (Branta canadensis) that were or were not shedding ABV in cloacal swabs at the time of blood collection.

CAGO: Canada goose; SD: standard deviation a S/P ratio: sample to positive ratio calculated as: [sample mean of optical absorbance – negative control mean] / [positive control mean – negative control mean]; b Calculations of sample seropositivity were performed using the mean S/P ratio of the negative control group plus 2 and 3 standard deviations in order to provide two levels of sensitivity and specificity; c Sample considered positive if its S/P ratio is higher than the mean S/P ratio of the negative control grou p plus 2 standard deviations; d Sample considered positive if its S/P ratio is higher than the mean S/P ratio of the negative control group plus 3 standard deviations.

Figure 3.5. Sample to positive ratio of serum samples assessed for the presence of antibodies against avian bornavirus (ABV) using an enzyme-linked immunosorbent assay for 75 mute swans (Cygnus olor) that were or were not shedding ABV in cloacal swabs at the time of blood collection.

MUSW: mute swan; SD: standard deviation a S/P ratio: sample to positive ratio calculated as: [sample mean of optical absorbance – negative control mean] / [positive control mean – negative control mean]; b Calculations of sample seropositivity were performed using the mean S/P ratio of the negative control group plus 2 and 3 standard deviations in order to provide two levels of sensitivity and specificity; c Sample considered positive if its S/P ratio is higher than the mean S/P ratio of the negative control group plus 2 standard deviations; d Sample considered positive if its S/P ratio is higher than the mean S/P ratio of the negative control group plus 3 standard deviations.

Birds that were shedding ABV (all species considered together) had a significantly higher seroprevalence than birds that were not shedding the virus (13/14 vs 259/486 [2SD cut-off]; 0/14 vs 88/486 [3SD cut-off]) (Table 3.4). This difference of seroprevalence between shedders versus non-shedders was also significant considering only the Canada geese (7/7 vs 110/196 [2SD cut-off]; 7/7 vs 22/196 [3SD cut-off]); and only the Canada geese sampled at Toronto Zoo (7/7 vs 20/46 [2SD cut-off]; 7/7 vs 2/46 [3SD cut-off]) (Table 3.4). However, this difference (shedders vs non-shedders) was not significant for mute swans considered separately.

Birds that were shedding ABV had a significantly higher mean S/P ratio (1.372 vs 0.553 (P<0.0001) for swans and geese considered together; 1.686 vs 0.480 (P<0.0001) for Canada geese only; 1.686 vs 0.385 (P<0.0001) for Canada geese from Toronto Zoo only; 1.058 vs 0.592 (P=0.0013) for mute swans only).

DISCUSSION

The results of this study provide the first estimates of the prevalence of cloacal shedding of ABV and of presence of antibodies against ABV in four species of free- ranging waterfowl in southern Ontario, Canada. Previous research studies on ABV in wild waterfowl populations have used RT-PCR, IHC and histology to describe the pathology that can be associated with ABV in geese and swans (Delnatte et al., 2013) and RT-PCR on brains and oropharyngeal / cloacal swabs to document the widespread presence of ABV in various species of Anseriformes (Payne et al., 2011a; Guo et al., 2012, Payne et al., 2012).

Our results suggest that exposure to ABV is widespread in southern Ontario in the species tested, but that infection rates and/or the biology of the disease vary among species and locations. Antibodies were found in all species and at all locations sampled with seroprevalences ranging from 6.5% to 71%, depending on species and location and on how seropositivity was determined. A higher seroprevalence in adult birds suggests that exposure to ABV increases with age. Widespread asymptomatic infection with less 99

common clinical disease is a consistent feature of bornavirus infections, and may be attributed to the non-cytopathic effect of these viruses. Anti-ABV Ab can be present in asymptomatic ABV-positive (shedding) macaws (de Kloet et al., 2009; Lierz et al., 2009; Kistler et al., 2010; Villanueva et al., 2010; Heffels-Redmann et al., 2012). Although ABV infection has been associated with significant neuropathology in waterfowl in Ontario (Delnatte et al., 2013), poor body condition or neurologic deficits were not noted in any of the birds captured and sampled in this study.

While antibodies to ABV were widespread in all four species of birds, only 9.3% of mute swans, 3.4% of Canada geese, and none of the trumpeter swans or mallards were shedding ABV in cloacal swabs at the time of sampling. These shedding rates are consistent with the ones recently reported in American studies, where 2.9% (12/409) of Canada geese (Payne et al., 2011b) and 6% (14/219) of mute swans (Payne et al., 2012) were positive using RT-PCR on oropharyngeal / cloacal swabs. However, the difference in shedding rates among species was surprising given the fact that these four species commingle and that that ABV-diseased trumpeter swans had previously been identified in the sampled flock. A species based difference in the biology of ABV infection is suggested. While the trumpeter swan is a native North American species, mute swans were introduced in the late 19th century from Eurasia and thus may have a different evolutionary exposure to ABV (Lumsden et al., 2009). Further studies are required to verify whether a true difference in shedding frequency exists in natural and, ideally, experimental infections.

Assessment of cloacal shedding of virus likely underestimates the prevalence of ABV infection. Rates of viral shedding might be expected to vary with stage of infection, being higher soon after initial infection or in association with viral reactivation and active replication. In addition, intermittent urofecal shedding of ABV is described in both experimentally and naturally infected parrots (Villanueva et al., 2010; Payne et al., 2011b; Heatley et al., 2012), and a similar pattern is suspected in waterfowl (Payne et al., 2012; Delnatte et al., 2013). Although the testing of a single cloacal swab is thus of limited usefulness in determining the ABV status of an individual bird, this method remains a useful tool to screen for the presence of active ABV infection on a flock scale.

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Pooling multiple droppings collected over several days, the current recommendation for investigation of ABV status in individual psittacine birds (Hoppes et al., 2013), is impractical and unrealistic for studies of free-ranging wild birds.

The results of serologic and cloacal swab evaluation were compared on a group, species and an individual bird basis. Birds that were shedding ABV in cloacal swabs had higher mean S/P ratios compared to non-shedding birds; however, not all birds shedding virus were seropositive. Possible explanations for this include the presence of acutely- infected birds that had not yet seroconverted, that some individuals simply do not seroconvert, or that, less likely and contrary to our underlying assumption, the virus in feces was ingested on contaminated material and was simply passing through the digestive tract. This discordance between shedding and seropositivity was also seen in several psittacine studies; the detection of ABV-Abs in serum and ABV RNA in feces from an individual bird did not always correlate (Enderlein et al., 2010) and birds without detectable Abs could shed the virus in feces (Villanueva et al., 2010).

The research presented here supports previous indications that infection rates vary according to geographic location (Payne et al., 2011a; Guo et al., 2012; Payne et al., 2012; Delnatte et al., 2013). The prevalence of shedding in Canada geese caught on the Toronto Zoo site was 13% compared to a 0% prevalence in geese sampled at the three other locations. Although, in general, Canada geese in Ontario are considered to belong to one migratory metapopulation, local factors might influence the presence and expression of disease. Our retrospective study showed that the proportion of birds with a clinical history or pathological lesions suggestive of ABV infection was significantly higher at the Toronto Zoo (30/132) than elsewhere in Ontario (21/823) (Delnatte et al., 2013). This appears to correlate with the higher prevalence of cloacal shedding at the zoo site and supports the premise that environmental or ecological factors may affect the shedding of ABV and possibly the onset of disease. The persistent high density of Canada geese on the zoo site, and the fact that the majority of these birds remain year- around in this attractive geographically restricted area, very likely enhance the transmission of ABV leading to the development of a disease cluster.

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Although we generated what appear to be valid serological data, the assessment of the presence of Ab in these birds was hindered by several methodological and technical difficulties. Historically, serological assays have not been very convincing tests for diagnosing BDV infections of mammals (Staeheli et al., 2010). However, unlike BDV, ABV in affected parrots shows a broad tissue tropism and ABV antigens are abundantly present in affected organs (Gancz et al., 2009; Rinder et al., 2009; Raghav et al., 2010). This suggests that the immune response to ABV may be stronger than the one observed to BDV and has encouraged researchers to develop serological tests for avian species. To date, serologic assays described for the identification of Ab to ABV in psittacine birds include indirect immunofluorescence assay, Western blot assay and indirect ELISA tests with a variety of sources of primary antigen used (Hoppes et al., 2013). The N-protein appears immunodominant (Lierz et al., 2009; Villanueva et al., 2010), thus our choice to use a purified M24N protein as the source of antigen in our ELISA. Secondary antibodies previously used for ABV ELISA include goat anti-bird IgG (de Kloet et al., 2011) and rabbit anti-canary IgG (Rubbenstroth et al., 2013) conjugated to either horseradish peroxidase or alkaline phosphatase.

Actual assessment and comparison of the specificity and sensitivity among various serological tests are very difficult due to differences in methodology, and perhaps more importantly, poor understanding of epidemiology of the disease, the absence of a gold standard for identifying infection in live birds, the difficulty in identifying a true control group of uninfected birds, the variability among species, and the relatively poor correlation between shedding, pathology, and seroconversion. We encountered these same difficulties in developing and interpreting our ELISA test, in addition to those related to working with free-ranging, non- domestic species.

In order to design an assay independent of the species tested, we used an anti-bird IgG antibody conjugate designed for diverse avian species and used previously for the detection of antibodies to ABV in avian sera (de Kloet et al., 2011). The reactivity of this conjugate towards immunoglobulins for each of the species studied was initially verified using dot blots (P Delnatte, Unpublished data). The intensity of OD405 readings in our

ELISA varied greatly among species, with OD405 values being much higher for ducks

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(including the negative control domestic ducks) than for the geese and swans. This is most likely due to non-specific binding of the secondary Abs, the intensity of which appeared to be species specific. These species variations precluded any direct comparison of S/P ratios among species and highlighted the need for species-specific secondary antibodies and seropositivity cut-offs.

A common practice with ELISA testing is to report a sample as positive if the S/P ratio is higher than the mean S/P ratio for the negative control group plus 2 and/or 3 SD (Ojkic et al., 2003). A cut-off value of 3 SD above the mean OD value of a negative group was previously used in ELISA studies to detect anti-ABV antibodies in psittacine birds and canaries (de Kloet et al., 2011; Rubbenstroth et al., 2013). Calculations of sample seropositivity in this study were performed using both 2 and 3 SD in order to provide two levels of sensitivity and specificity of the assay. Negative control domestic geese were used for the calculation of positivity cut-offs for the Canada geese samples; and negative control domestic ducks were used for the calculation of the positivity cut- offs for the mallard samples. As we did not have any known negative swans, we chose to use the goose positivity cut off based on the fact that swans are more closely related phylogenetically to geese than to mallards and that the range of swan OD405 readings was more similar to the range of OD405 observed in Canada geese that in mallards.

An additional concern relates to the fact that OD405 readings higher than 1, which occurred in many duck samples, are usually considered to be less reliable and more difficult to interpret than OD405 readings in the 0 to 1 range. Using a more species- specific secondary antibody, such as an anti-duck IgG, or using different serum or antigen dilutions may have helped overcome these high OD reading in duck samples and should be considered in future studies. Despite the above limitations, our ELISA test allowed us to detect ABV exposure and estimate the prevalence of ABV in several free- ranging waterfowl species.

Statistical comparisons between groups were made using Fisher’s exact test. Although several comparisons were statistically significant, the validity of the conclusions obtained could be questioned given that the confidence intervals sometimes

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included the null value of one (Dohoo et al., 2009). Using logistic regression would be an alternative approach.

In conclusion, this prospective study demonstrated for the first time that antibodies directed against ABV are widespread in wild, apparently healthy, Canada geese, trumpeter swans, mute swans and mallards in southern Ontario. This study also confirmed that virus can be shed in the urofeces of asymptomatic mute swans and Canada geese. Despite their respective limitations, serologic assays and RT-PCR testing of cloacal swabs can be used to estimate the prevalence of ABV at a flock level. Because ABV can also be associated with significant pathology in these species, the overall impact of the waterfowl genotype of ABV on waterfowl populations, as well as on other wild or domestic species whose environment may be contaminated by this virus, remains uncertain. Avian bornavirus infection is likely under-recognized in wild waterfowl and is unlikely to be restricted to North America given the migratory nature of the host species, thus free-ranging waterfowl from other continents should be evaluated for the presence of ABV.

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CHAPTER 4:

Investigation of the possibility of vertical transmission of avian

bornavirus in free-ranging Canada geese

Pauline Delnatte1,2, Éva Nagy1, Davor Ojkic3, Graham Crawshaw2, and Dale A Smith1*

1 Ontario Veterinary College, University of Guelph, Guelph, Ontario, Canada, N1G 2W1

2 Toronto Zoo, Toronto, Ontario, Canada, M1B 5K7

3 Animal Health Laboratory, University of Guelph, Guelph, Ontario, Canada, N1G 2W1

Keywords: avian bornavirus, Branta canadensis, Canada goose, egg, in ovo infection, vertical transmission, waterfowl

Submitted to Avian Pathology.

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ABSTRACT

In order to investigate the possibility of in ovo infection with avian bornavirus (ABV) in wild Canada geese, 53 eggs were opportunistically collected at various stages of embryonic development from the 16 nests of free-ranging Canada geese at a site where ABV is known to be endemic. ABV RNA was detected in the yolk of one of three unembryonated eggs using real time reverse transcription polymerase chain reaction. ABV RNA was not identified in the brains from 23 new hatched goslings or 19 embryos, and from 3 early whole embryos. Antibodies against ABV were not detected in the plasma of any of the hatched goslings using an enzyme-linked immunosorbent assay. Possible reasons for the failure to detect ABV RNA in hatchlings or embryos include low sample size, eggs deriving from parents possibly not actively infected with ABV, the testing of only brain tissue, and possible failure of the virus to replicate in Canada goose embryos. In conclusion, the presence of ABV in the yolk of a Canada goose egg provides the first evidence for the potential for vertical transmission of ABV in waterfowl.

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INTRODUCTION

A novel genotype of avian bornavirus (ABV) was recently identified in association with significant neuropathology and mortality in free-ranging Canada geese (Branta canadensis), trumpeter swans (Cygnus buccinator) and mute swans (Cygnus olor) in southern Ontario, Canada (Smith et al., 2010; Delnatte et al., 2013). More recently, ABV infection has been shown to be common and widespread in a variety of apparently healthy wild waterfowl species across North America (Payne et al., 2011a; Guo et al., 2012; Payne et al., 2012). The bornaviruses circulating among free-living waterfowl species appear to form a unique genotype (ABV-CG genotype) whose impact on waterfowl populations remains uncertain (Payne et al., 2011a; Guo et al., 2012; Payne et al., 2012; Delnatte et al., 2013).

The prevalence of ABV infected birds in the United States and in Canada shows an uneven geographic distribution of the virus (Guo et al., 2012; Payne et al., 2012; Delnatte et al., 2013). In a survey carried out at the Toronto Zoo in June 2011, the seroprevalence of ABV in Canada geese was 51% and the prevalence of shedding was 13%; significantly higher than the prevalence found in geese sampled at other locations in the province (see chapter 3). In addition, between 1992 and 2011, at least 23 free- ranging Canada geese originating from the Toronto Zoo died or were euthanized with ABV-related neurological disease that was confirmed at necropsy by histology, immunohistochemistry and reverse transcription polymerase chain reaction (RT-PCR) (Delnatte et al., 2013). The persistent high density of Canada geese at the Toronto Zoo, and the fact that the majority of these birds remain year-around in this attractive geographically restricted area, may play a role in enhanced transmission of ABV at this particular location.

Transmission of ABV is thought to occur primarily via ingestion of contaminated droppings. Avian bornavirus is present in the feces and urine of infected parrots and waterfowl species (Raghav et al., 2010; Heatley et al., 2012); urofecal-oral transmission of ABV was confirmed in an unintentional study where mallards acquired the infection by ingesting droppings from infected cockatiels (Hoppes et al., 2010). Epidemiological investigation of outbreaks of proventricular dilation disease in psittacine aviaries also 107

suggests urofecal-oral transmission of ABV as the primary method of spread. Respiratory tract secretions can also be a source of virus; and social interactions such as mutual preening would enhance virus transmission (Hoppes et al., 2010; de Kloet et al., 2011). More recently, ABV RNA was detected in eggs, embryos, and hatchlings of various psittacine and passerine species suggesting the possibility of vertical transmission of ABV (Lierz et al., 2011; Kerski et al., 2012; Monaco et al., 2012; Rubbenstroth et al., 2013). In addition, vertical transmission of Borna disease virus has been demonstrated in mice (Okamoto et al., 2003) and is strongly suspected to occur in horses (Hagiwara et al., 2000). Vertical transmission of a variety of viral infections is well recognized in poultry (Brentano et al., 2005; Grgic et al., 2006; Pillai et al., 2010).

Our hypothesis was that in ovo infection with ABV occurs in Canada geese and thus that evidence of infection will be found in some eggs, embryos and/or hatchlings collected from an infected flock. The objective of this work was to opportunistically collect Canada goose eggs from a known ABV-infected flock to investigate the possibility of in ovo transmission of ABV infection in a wild waterfowl species.

MATERIALS AND METHODS

Egg collection

Eggs were collected from Canada goose nests on the Toronto Zoo site (N43° 49', W79° 11') between April and June 2012 as part of the annual spring goose population control program held under a Canadian Wildlife Service (CWS) permit (#DA 0326). The zoo property was searched for Canada goose nests twice weekly during this period. Depending on the number of eggs present, the protocol for each nest was as follows: one egg was oiled and left in place to prevent the birds from laying a second clutch unless the parents were aggressive towards the public, in which case no eggs were left in the nest and the nest was destroyed; two eggs were collected for incubation; any remaining eggs were opened and any embryos present necropsied. For each nest records were made of the location, the date of egg collection, the total number of eggs present, whether an oiled 108

egg was left in the nest, the number of eggs incubated and the number of eggs necropsied. Whenever possible, the parents were captured, manually restrained and a blood sample and a cloacal swab were collected.

Artificial incubation

After collection, the eggs that were selected to be incubated (n = 28) were cleaned, labeled and transported to the laboratory in an insulated cooler containing a warm towel. Incubation was carried in "Octagon 20 Advance" incubators (Brinsea Products Inc., Titusville, Florida, USA) with automatic control of temperature and humidity, and an automatic egg turning cradle. The temperature and humidity were maintained at 38.3oC and 60% throughout incubation; at external pipping humidity was increased to 70%. Eggs were positioned on their sides and manually rotated half a turn on the long axis once daily in addition to the automatic movement of the egg cradle. Eggs were checked for viability once weekly using a digital egg monitor (“Buddy v2”; Animal Genetics Inc., Tallahassee, Florida, USA). Goslings that hatched (n=23) were, as soon as possible, deeply anesthetized using isoflurane administered via a small induction chamber and then euthanized via cervical dislocation.

Egg collection, incubation and euthanasia were approved by the University of Guelph and Toronto Zoo Animal Care Committees (Animal Use Protocol Guelph # 10R058; TZ #2012-03-01) in accordance with the requirements of the Animals for Research Act of Ontario and the recommendations of the Canadian Council for Animal Care. A CWS Migratory Bird Research Permit (#CA 0291) was also obtained.

Necropsy and sample collection

Eggs/Embryos

Eggs were disinfected with alcohol, opened with sterile equipment, and their contents were placed into a sterile Petrie dish. The contents of the egg were photographed and when the egg was not embryonated (n = 3), samples of albumin and of yolk were

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frozen. Early embryos (less than 4 cm crown-rump length) (n = 3) were frozen and embryos greater than 4 cm in length (n = 19) were decapitated and then necropsied; samples of brain and proventriculus/ventriculus were collected and frozen.

Hatched Goslings

Blood samples were collected immediately post-mortem via cardiocentesis. Blood samples were centrifuged for 5-10 minutes at 2500 rpm, and plasma was transferred into a cryovial tube (Sarstedt Inc., Nümbrecht, Germany). The skull was opened aseptically and half the brain was collected and frozen. All plasma and tissue samples were frozen at -80oC until testing (up to 1 year).

Nucleic acid isolation and RT-PCR

Brain samples from the 23 hatched gosling and from the 19 embryos greater than 4 cm, whole embryos (for the 3 early stage embryos), yolk samples from the 3 non- embryonated eggs and cloacal swabs from the parents (when available) were evaluated for the presence of ABV by real time RT-PCR detection of the ABV matrix gene. Primers and cycling parameters were as previously described (Delnatte et al., 2013). A RT-PCR crossing point value less than 37.00 was considered positive, between 37.00 and 39.99 inconclusive, and equal to or greater than 40.00 negative.

Serology

Plasma samples from each one-day old gosling and from the parents (when available) were assessed for the presence of ABV specific antibodies using an enzyme- linked immunosorbent assay (ELISA) (see chapter 3). ABV nucleocapsid purified protein (provided by Susan Payne, Texas A&M) was the antigen and horseradish peroxidase- labelled goat anti-bird IgG (Bethyl Laboratories Inc., Montgomery, Texas, USA) was used as the secondary antibody. The optical density (OD) was read at 405 nm and the ABV specific antibody responses were determined by calculating the sample-to-positive

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(S/P) ratio using mean OD405 readings. Sample to positive ratio was calculated as: [sample mean – negative control mean] / [positive control mean – negative control mean]. Plasma samples were considered positive if the S/P ratio of the sample was higher than the mean S/P ratio for the negative control plus 2 standard deviations. The methods for determining positive and negative control values and the seropositivity cut-off have been previously described (see chapter 3).

RESULTS AND DISCUSSION

A total of 53 Canada goose eggs were collected from 16 nests. Twenty-eight eggs were incubated and 23 of them hatched within 5 to 26 days; the remaining 5 were not embryonated. Twenty five eggs were opened immediately after collection; 22 contained embryos.

ABV RNA was detected by RT-PCR in one yolk sample from a non-embryonated egg (Cp of 28.45). This egg was laid in the morning and abandoned a few hours later. When collected, the egg was cold and thus was not incubated. No other eggs were present in this nest. The remaining samples were negative for ABV RNA, including brain samples from 23 hatched goslings and 19 embryos, 3 whole early-stage embryos, and yolk samples from 2 non-embryonated eggs. Antibodies against ABV were not detected in the plasma of the hatched goslings. S/P ratios ranged from -0.915 to -0.278, with a mean and a standard deviation of -0.748 (±0.180).

This is the first report of the detection of ABV RNA in the egg yolk of a wild Canada goose. This finding is not surprising given previously described evidence for vertical transmission of ABV in psittacine birds, the demonstration of ABV antigen in testes and ovaries of infected parrots (Raghav et al., 2010; Payne et al., 2011b), the presence of lymphoplasmacytic inflammation in the gonads of infected Canada geese (Delnatte et al., 2013), and the identification of ABV in infected domestic duck eggs used for cell culture (Hoppes et al., 2013).

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We were not able to identify evidence of ABV infected embryos or hatchlings in this flock of Canada geese, and there are a number of reasons why this may have occurred. Firstly, it is quite likely that our sample size was not sufficient. In a previous study on the prevalence of infection in adult birds, it was calculated that a sample size of approximately 200 was necessary to identify an event present at an estimated prevalence of 0-4% with a 95% confidence interval (Dohoo et al., 2009). There is no data to provide guidance on estimating the rate of vertical transmission should it occur, but we expect the rate would be very low.

Secondly, it is possible that all fertile eggs collected were from uninfected parents. Ideally, eggs would only have been collected from nests whose parents were ABV positive. We anticipated being able to assess parental status using cloacal swabs and serology; however, catching free-flying parents during breeding season proved to be much more challenging than expected. The adult geese were not very protective of their nests and most flew away quickly when the nests were approached. Only five parent geese (3 females and 2 males) were captured: cloacal swabs were collected from each of them, and a blood sample was collected from only one male. These five cloacal swabs were negative for the presence of ABV by RT-PCR; and the only plasma sample available was negative for anti-ABV antibodies (S/P ratio of -0.478). The parents from the egg containing the positive yolk sample were unfortunately not caught. Given a prevalence of ABV shedding of 13% and seroprevalence of 51%, as assessed in this flock one year previously, it would be surprising if none of the 32 parents assessed showed evidence of ABV exposure or infection (see chapter 3). However, the 2011 survey included Canada geese of all ages and breeding status, and thus these figures do not represent the actual prevalence within the adult breeding population. It is also possible that ABV infected birds may be less likely to pair up and breed than are non-infected birds.

The lack of antibodies to ABV in the plasma of newly hatched goslings also supports the possibility of our having inadvertently sampled only the nests of uninfected parents. In birds, the IgY composition of egg yolk commonly reflects the IgY composition in serum of the female parent (Hamal et al., 2005). ABV specific antibodies

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were found in egg yolk and serum of late-stage embryos of sun conures (Kerski et al., 2012), and the ratio of antibodies against the ABV nucleocapsid protein, viral X protein, phosphoprotein and matrix protein reflected the antibody ratio in the maternal serum (Kerski et al., 2012). Because the ABV nucleocapsid protein appears to be immunodominant (Lierz et al., 2009; Villanueva et al., 2010), we used a purified nucleocapsid protein as antigen in our ELISA. However, as the relative levels of the antibodies against the different ABV proteins can vary widely among individual birds in psittacine species (Kerski et al., 2012), it is not impossible that although we did not detect any antibodies against the nucleocapsid protein in these 23 goslings, antibodies against other ABV proteins may have been present.

Inappropriate sample collection might also have prevented our identifying evidence of in ovo disease transmission in embryos or hatchlings. The decision to test the brains of the hatchling geese and of the embryos (more than 4 cm) was based on the frequency of brain lesions and the presence of high viral load in the brains of infected diseased adult Canada geese (Delnatte et al., 2013), as well as in infected parrots (Raghav et al., 2010). The pathogenesis of in ovo infection with ABV has not been studied; hence the distribution of virus in infected embryos or hatchlings is unknown. However, ABV was detected in the brain tissue of a nonviable embryo from an Indian ring-necked parakeet (Psittacula krameri), two late-stage (22 day-old) nonviable embryos from sun conures, and in the brain tissue of viable embryos from a lovebird (Agapornis roseicollis) and from a cockatiel (Nymphicus hollandicus) (Monaco et al., 2012; Kerski et al., 2012). These results suggest that ABV appears to reach the brain during in ovo infection in psittacine species.

And finally, it is also possible that in ovo infection of embryos may simply not occur in Canada geese. It seems; however, unlikely that the routes of transmission of the waterfowl genotype of ABV are significantly different from those of other bornaviruses and that the psittacine and passerine strains are vertically transmitted whereas the Canada goose strain is not (Okamoto et al., 2003; Lierz et al., 2011; Kerski et al., 2012; Monaco et al., 2012; Rubbenstroth et al., 2013). Given the presence of ABV RNA in the yolk of one infertile egg, the possibility of transmission to a developing embryo remains open.

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In conclusion, we were able to demonstrate the presence of ABV RNA in the yolk of an infertile egg from a wild Canada goose by opportunistically collecting eggs from an infected flock. We were not able to demonstrate the entire cycle of in ovo transmission that would have required the isolation of ABV in a developing embryo and in a newly hatched bird; however there were several obvious limitations to this project. Additional studies using a larger sample size of eggs derived from parents that are known to be positive for ABV infection, perhaps in a captive or better controlled situation, and evaluation of a greater variety of tissues of embryos and hatchlings of a range of ages are required to formally demonstrate the occurrence of in ovo infection and the potential for vertical transmission of ABV in waterfowl.

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CHAPTER 5: EXPERIMENTAL TRIAL

Susceptibility of domestic poultry to experimental infection with the

waterfowl genotype of avian bornavirus

Pauline Delnatte1,2, Éva Nagy1, Josepha DeLay3, Davor Ojkic3, and Dale A Smith1*

1 Ontario Veterinary College, University of Guelph, Guelph, Ontario, Canada, N1G 2W1

2 Toronto Zoo, Toronto, Ontario, Canada, M1B 5K7

3 Animal Health Laboratory, University of Guelph, Guelph, Ontario, Canada, N1G 2W1

Keywords: avian bornavirus, ABV-CG, chicken, duck, experimental infection, goose, waterfowl

To be submitted to Avian Diseases.

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ABSTRACT

The objective of this experimental trial was to determine whether domestic chickens (Gallus gallus domesticus), Pekin ducks (Anas platyrhynchos domesticus) and Embden geese (Anser anser domesticus) were susceptible to infection with the waterfowl strain of avian bornavirus (ABV). For each species, eight birds were euthanized prior to inoculation, 13 birds were inoculated intramuscularly with brain homogenates from ABV-infected Canada geese and 13 birds were used as sham inoculated controls. Birds were observed for the presence of clinical signs and samples were collected biweekly to assess viral shedding and seroconversion using real time reverse transcriptase – polymerase chain reaction (RT-PCR) and an enzyme-linked immunosorbent assay, respectively. Necropsies were performed at day 45 or day 90 post-inoculation to look for evidence of infection using histology, immuno-histochemistry (IHC) and RT-PCR. ABV was not detected using IHC and RT-PCR in any of the experimental or control birds of the three species, and none of the birds seroconverted. No histological lesions consistent with ABV infection were found in any ducks or chickens. However, mild to moderate non-suppurative inflammatory infiltrates and glial nodules were present in the central, peripheral and autonomic nervous tissue of 5/13 experimental geese, 5/13 control geese and 3/8 geese euthanized prior to the commencement of the trial, which were also ABV negative by IHC and RT-PCR. Given the similarity in prevalence and morphologic appearance of lesions in the three groups of geese, the presence of a pre-existing, non- ABV neurotropic virus was suspected.

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INTRODUCTION

Avian bornavirus (ABV) is a newly discovered viral agent that affects a variety of avian species. The virus was first identified in 2008 and is recognized as the cause of proventricular dilatation disease (PDD) in psittacine birds, a devastating condition for which there is no effective vaccine or treatment (Honkavuori et al., 2008; Kistler et al., 2008). Characteristic pathologic findings include emaciation, proventricular and ventricular dilatation, and non-suppurative encephalitis, myelitis, peripheral neuritis, and ganglioneuritis (Shivaprasad et al., 1995; Berhane et al., 2001; Schmidt et al., 2003). Histopathologic lesions consistent with PDD have been identified occasionally in a variety of other avian species, including passerines, falconiformes, piciformes and anseriformes (Daoust et al., 1991; Gregory et al., 2000; Perpiňán et al., 2007; Weissenböck et al., 2009b; Delnatte et al., 2011).

A genotype of ABV different from those identified in psittacine birds, named ABV-CG, was identified in 2009 as being associated with significant neuropathology and disease in free-ranging Canada geese (Branta canadensis) and trumpeter swans (Cygnus buccinator) (Smith et al., 2010; Delnatte et al., 2013). This virus has since been found in a variety of wild waterfowl and gull species across the United States and Canada, where it has been identified in oropharyngeal/cloacal swabs and brain samples from birds that appeared clinically unaffected (Payne et al., 2011a; Guo et al., 2012; Payne et al., 2012; chapter 3, Delnatte et al., 2013).

There has been no evaluation as to whether the presence of free-ranging waterfowl species shedding ABV in their feces poses a risk to the poultry industry. The susceptibility of domestic poultry species to ABV is currently unknown, and the transmission of ABV-CG from wild waterfowl to poultry species has never been documented. However, ABV appears to have a broad avian species tropism, and several domestic duck eggs from two different commercial hatcheries in Texas were found to be infected with ABV-CG (Hoppes et al., 2013). Thus concern has been raised that the waterfowl genotype of ABV may be a threat to the commercial poultry industry, especially to backyard, open range, and organic producers.

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The histopathologic lesions of ABV infection in psittacine birds and waterfowl are similar to those caused by a variety of neurotropic viruses, including reportable paramyxoviruses (Woolcock et al., 2008). If infection of domestic poultry occurs and results in similar pathologic changes, ABV infection could be an important cause of misdiagnosis and should be recognized as a differential diagnosis for clinical neurologic disease and neuropathology. Other clinical signs of ABV infection, including weight loss and failure to thrive, would also have economic consequences.

MATERIALS AND METHODS

Bird acquisition and housing

Thirty-four domestic white Leghorn chickens, 34 Pekin ducks and 34 Embden geese were purchased at one day of age from a research station (Arkell Poultry Research Centre, University of Guelph, Guelph, Ontario, Canada for chickens) or from Ontarian commercial suppliers.

Birds were housed within dedicated rooms in the Isolation Unit – Central Animal Facility (University of Guelph, Ontario). All birds of each species were housed together in one room until the start of the trial, after which point experimental and control groups were held in separate rooms. Feed and water were provided ad libitum and heat lamps hung as needed. Housing was appropriate to the species and altered as necessary as the birds aged.

Study Design

The same protocol was used for all species. For each species, birds (n = 34) were randomized into three groups. On day 0, one group (experimental group; n = 13) was 118

inoculated intramuscularly with 0.5 ml of ABV-infected brain homogenate (see below for the preparation of the inoculum), the second group (control group, n = 13) was sham inoculated with the same volume of phosphate buffered saline (PBS), and the third group (pre-inoculation control group; n =8) was euthanized. Geese were inoculated at 6 weeks of age, ducks at 8 weeks of age and chickens at 10 weeks of age.

Twice daily throughout the study, each room was assessed for food and water intake and amount of feces, and each bird was assessed for general attitude, activity, respiratory pattern, and presence of neurological signs. The spreadsheet used for daily group assessment is provided in Appendix D.

Blood samples and cloacal swabs were collected from all birds in the control and experimental groups immediately before inoculation (day 0), and then every two weeks (day-15, 30, 45, 60 and 90) until the end of the study. As described in Chapter 3, blood samples were collected from the medial metatarsal vein and sera were separated and stored at -80oC; and cloacal swabs were taken as previously described and placed in a virus transport medium tube (Multitrans System, Starplex Scientific Inc., Etobicoke, Ontario, Canada) and stored at -80oC.

Birds from each of the control and experimental groups were randomly selected for euthanasia at 45 days post-inoculation (dpi) (n=6) and 90 dpi (n=7). All birds in the study were euthanized by placement in a confined chamber that was subsequently filled with CO2.

Diagnostic tests performed for each group of birds are summarized in Table 5.1.

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Table 5.1. Summary of diagnostic tests performed for each group of birds in a trial carried out to evaluate the susceptibility of domestic poultry species to experimental infection with avian bornavirus.

Post- Histo- Status / Number Species mortem patholog IHC RT-PCR ELISA Group of birds (dpi) y Pre- D0 8 none none none none inoculation D45 6 Experi- brain brain bi- mental spinal spinal Chicken D90 7 weekly cord cord brain serum adrenal adrenal (frozen) samples D45 6 kidney kidney

Control pancreas pancreas D90 7 Pre- D0 8 none none none none inoculation D45 6 Experi- brain brain bi- Duck mental spinal spinal weekly D90 7 cord cord brain serum adrenal adrenal (frozen) samples D45 6 kidney kidney Control pancreas pancreas D90 7 Pre- D0 8 brain none none none inoculation brain brain D45 6 spinal (frozen & Experi- cord brain FFPE) bi- mental Goose adrenal spinal adrenal weekly D90 7 kidney cord (frozen) serum pancreas adrenal samples D45 6 (+ full set kidney brain Control of tissues pancreas (frozen) D90 7 for four birds) dpi = day post-inoculation; ELISA = enzyme linked immunosorbent assay; FFPE = formalin -fixed paraffin-embedded; IHC = immunohistochemistry; RT-PCR = reverse transcriptase polymerase chain reaction.

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Preparation of the inoculum and inoculation

The inoculum was prepared from brain tissue harvested from two free-ranging Canada geese that were part of a retrospective study describing the pathology associated with the waterfowl genotype of ABV (ABV-CG) in wild geese and swans (CAGO-Z-7 and CAGO-Z-8) (see chapter 2). These geese were in poor body condition and had moderate to severe neuropathology consistent with ABV infection, including marked non-suppurative encephalitis. Their brains were positive for ABV on gel-based RT-PCR (N gene), real time RT-PCR (M gene) and IHC. RT-PCR products from these two brains were sequenced and their nucleotide sequences were 100% identical. Using pairwise comparison based on nucleotide sequence, the N gene sequences were compared with the available ABV sequences in GenBank and were found to be 100% identical to the Canada goose sequences that we deposited in GenBank for the retrospective study (Gen Bank # JX258867 and JX258868) (see Chapter 2). These brains were thus confirmed to contain the waterfowl genotype of ABV (ABV-CG).

The brains from these two geese were evaluated for the presence of other pathologic agents that could be transmitted to the experimental birds. Routine microbial culture was performed on blood-agar and McConkey's agar media and no bacterial growth was identified. IHC for WNV was performed on sections of formalin-fixed paraffin-embedded (FFPE) brain and was negative. To test for the presence of other viruses, brain homogenates were inoculated onto four different cell lines [chicken hepatoma (CH-SAH), chicken fibroblast (DF-1), duck embryo fibroblast (DEF), and quail cell (QT-35)]. No cytopathic effect was observed.

Thawed brains were macerated by two passages through a 3 ml syringe and then diluted with PBS. The preparation was homogenized for 15 seconds at 6000 rpm in a tube containing sterile grinding beads. The homogenates were then centrifuged for 5 minutes at 4000 rpm, and the supernatants were diluted with PBS for a final dilution of approximately 1:3 (weight per volume), equivalent to 340 mg of brain per ml of inoculum.

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The inoculum was prepared immediately before administration to experimental birds and was kept on ice until use. A total of 0.5 ml (equivalent to 170 mg of brain) of the inoculum was divided in 2 and administered by deep intramuscular injection into the cranial one-third of the right and left pectoralis muscles, approximately 1 cm lateral to midline, using a 23-gauge, 25.4 mm long needle. The inoculum prepared from the CAGO-Z-7 brain was used for the inoculation of the ducks and geese, and the inoculum prepared from CAGO-Z-8 brain was used for the inoculation of the chickens.

Animal Use

All animal use and procedures were approved by the University of Guelph’s Animal Care Committee, and by the Toronto Zoo’s Animal Care Research and Acquisition Committee, according to the regulations of the Canadian Council of Animal Care.

Post-mortem examination

Immediately after euthanasia, complete necropsies were performed on all birds. Tissue samples from brain, spinal cord (cervical and thoracic), peripheral nerves (vagus, sciatic and brachial nerves), eye, lungs, syrinx, trachea, heart, liver, spleen, bursa of Fabricius, thymus, pancreas, adrenal glands, kidneys, pectoral and thigh muscles, skin, growing feather, bone marrow, gonad, thyroid and parathyroid glands, esophagus, crop, proventriculus, ventriculus, duodenum, jejunum, ileo-cecal colic junction, cecum and cloaca were preserved in 10% buffered formalin. Additionally, samples of brain, cervical spinal cord, lung, liver, kidney, growing feather, and adrenal were frozen at -80°C.

Histopathology

Multiple sections of brain and a section of cervical spinal cord, adrenal, kidney and pancreas from all experimental and control birds and multiple sections of brains from the 8 pre-inoculation geese were reviewed for the presence of histological lesions. For

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cases where significant lesions were seen in the central nervous system, the entire set of tissues (listed above) was reviewed. Tissue samples had been formalin-fixed and paraffin-embedded (FFPE), sectioned at 4 mm and stained with haematoxylin and eosin. Central, peripheral and autonomic nervous tissues were specifically examined for the presence of lymphocytes and plasma cells.

Immunohistochemistry

IHC was performed on all experimental and control birds using sections from the paraffin blocks used for the initial histopathologic evaluation (brain, spinal cord, adrenal, kidney, pancreas). As described in chapter 2, 4 mm tissue sections mounted on charged slides were stained with rabbit polyclonal antiserum raised against the ABV nucleocapsid (ABV-N) protein (provided by the deRisi Laboratory, UC-SF, San Francisco, California, USA) using an automated staining instrument (Dako Autostainer; Dako Canada Inc., Mississauga, Ontario, Canada). A detailed description of the cloning, expression, purification, and generation of polyclonal antisera against the ABV nucleocapsid (ABV2- N) protein has been published previously (Gancz et al., 2009; Raghav et al., 2010). ABV- positive control brain sections included with each run were from a psittacine bird previously identified as ABV positive by both IHC and RT-PCR. For negative reagent controls, duplicate sections of each control and test tissue were subjected to the same immunohistochemical procedure with substitution of pre-immune rabbit serum at a similar protein concentration to the anti-ABV-N antisera. All slides were interpreted by one author, who was blinded to case information. A case was considered IHC-positive when strong, diffuse intranuclear staining was present in neurons or glial cells.

Nucleic acid isolation and RT-PCR

FFPE tissues and thawed frozen tissue (brain and adrenal) collected at necropsy were evaluated for the presence of ABV by real time RT-PCR detection of the ABV-M gene. Total nucleic acids from FFPE brain sections were extracted using the RNeasy® FFPE Kit (Qiagen Inc., Mississauga, Ontario, Canada) according to the manufacturer’s 123

instructions. Total nucleic acids from thawed frozen brain samples were extracted from 50 µl aliquots of 5 to 10% tissue suspensions using MagMAX-96 Viral RNA Isolation Kit in a MagMAX Express-96 Magnetic Particle Processor (Applied Biosystems Inc., Foster City, California, USA) according to the manufacturer's instructions.

The RT-PCR assay was a duplex test with two sets of primers and TaqMan probes: (1) ABV_M_120201 – targeting psittacine ABV-M gene sequences; and (2) ABVG_M_111029 – targeting geese ABV-M gene sequences. The amplification was carried out in 25 µl reactions in a LightCycler 480 Real-Time PCR System (Roche, Laval, Quebec, Canada) using AgPath-ID One-Step RT-PCR Kit (Applied Biosystems, Foster City, California, USA). Primers and cycling parameters for ABV RT-PCR were those previously described in chapter 2. A RT-PCR crossing point value (Cp) less than 37.00 was considered positive.

Antigen preparation and serology

A pET21a plasmid based expression cassette for the M24N protein was kindly provided by Dr. Susan Payne, Schubot Exotic Bird Health Centre, Texas A&M University. As described in Chapter 3, the isolated plasmid DNA was transformed into BL21 cells (DE3) (Novagen pET system manual, 10th Ed., 2003). Protein purification was performed with the Qiagen Ni-NTA purification system according to the manual (Qiagen, Dusseldorf, Germany). The purified protein was analyzed by SDS-PAGE and protein concentration was determined by the Bio-Rad Protein Assay (Bio-Rad, Hercules, California, USA).

Serum samples were assessed for the presence of antibodies using an ELISA that was specifically designed for this project as described in Chapter 3. Prior to performing the ELISAs, optimum dilutions of serum and secondary antibodies and optimum concentration of the antigen were determined using standard checkerboard analyses. The same investigator performed all ELISA tests for all serum samples.

ABV-N protein was purified as described above, and diluted in a carbonate buffer

(35 mM NaHCO3, 15 mM Na2CO3 [pH 9.6]). Immulon 2HB 96-well microtiter plates 124

(Dynex Technologies Inc., Chantilly, Virginia, USA) were coated with the antigen at a concentration of 100 ng/well and incubated at 4oC for 16h. The plates were then blocked with 3% bovine serum albumin at 37oC for 75 minutes. The serum samples were heat- treated 30 min at 56oC. Serum samples were diluted to 1:20 (goose samples) or 1:200 (duck and chicken samples) in a wash buffer (0.05% Tween 20 in PBS) and 200 µl of diluted serum were added to each well and incubated for 1 hour at 37oC. Each reaction was performed in duplicate wells. The plates were washed four times with wash buffer between each step.

For the chicken samples, 100 µl of alkaline phosphatase-labelled goat anti- chicken IgG heavy and light chain antibody conjugate (Kirkegaard & Perry Laboratories, Gaithersburg, Maryland, USA) at a dilution of 1:1000 was added and incubated for 75 minutes at 37oC. The colour was developed with p-Nitrophenyl phosphate system (Sigma Inc., Saint Louis, Missouri, USA). For the duck and goose samples, 100 µl of horseradish peroxidase-labelled goat anti-bird IgG heavy and light chain antibody conjugate (Bethyl Laboratories Inc., Montgomery, Texas, USA) at a dilution of 1:5000 was added and incubated for 75 minutes at 37oC. The colour was developed with ABTS peroxidase substrate system (Kirkegaard & Perry Laboratories, Gaithersburg, Maryland, USA). For all samples, the colour was developed until the average optical density (OD) of the two positive control wells reached an OD reading of 0.8 when read in a BIO-Tek ELISA microplate reader at 405 nm. When the OD405 between two duplicates of any sample differed more than 15%, the sample was retested.

The ABV specific antibody responses were determined by calculating the sample- to- positive (S/P) ratio. Sample-to-positive ratio was calculated as: [sample mean – negative control mean] / [positive control mean – negative control mean]. Sera from one of the control domestic geese (Anser anser domesticus) that tested negative for ABV using IHC and RT-PCR, and from a Canada goose that was shedding ABV were the negative and positive controls, respectively (see chapter 4).

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Statistical analysis

Statistical analyses were performed using Graph-Pad QuickCalcs software (http://www.graphpad.com/quickcalcs/). Unpaired t tests were used to compare the weight of the birds between the experimental versus control groups for each species at necropsy and to compare the mean S/P ratios (ELISA) between the experimental versus control groups for each species and at each sampling date. Two-tailed P<0.05 was considered significant.

RESULTS

Clinical observations and post-mortem macroscopic findings

All birds were determined to be in good health based on physical examination immediately prior the inoculation (day 0). Food and water intake, general attitude, activity, respiratory pattern and appearance of feces were normal for all birds throughout the study period. None of the birds showed neurological signs. At necropsy, none of the birds had macroscopic lesions consistent with ABV infection (crop stasis, dilatation of proventriculus or emaciation).

Chickens

Two experimental chickens and one control chicken had multiple superficial skin wounds, due to intraspecific aggression. At necropsy, all chickens were in good body condition, except for one experimental male that was in poor body condition (1.13 kg); this male had a string constricting its lower beak, likely impeding its ability to feed. Weight at necropsy varied from 1.13 kg to 2.48 kg, with an average of 1.74 kg and astandard deviation of 0.42 kg for the experimental birds and 1.80 kg (±0.34) for the control birds. This difference in body weight was not statistically significant (p=0.645).

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Ducks

The majority of ducks (both experimental and control) had mild to severe lesions of pododermatitis and mild conjunctivitis. At necropsy, all ducks were in good to excellent body condition with abundant fat stores. Weight at necropsy varied from 3.01 kg to 4.93 kg, with an average of 3.95 kg and a standard deviation of 0.35 kg for the experimental birds and 3.59 kg (±0.38) for the control birds. This difference in body weight was statistically significant (p=0.019), but was not considered biologically important.

Geese

The majority of geese (both experimental and control) had mild lesions of pododermatitis and two geese (one experimental and one control) had their carpi rotated outward ("angel wings"). One experimental goose (euthanized at day 90) had a large perirenal abscess and severe lesions of pododermatitis. At necropsy, all geese were in good to excellent body condition with abundant fat stores. Weight at necropsy varied from 3.5 kg to 7.15 kg, with an average of 4.61 kg and a standard deviation of 0.82 kg for the experimental birds and 5.59 kg (±1.09) for the control birds. This difference in body weight was not statistically significant (p=0.980).

Histopathology

Chickens

Three experimental chickens and two control chickens had one single, less than 5 cell-thick, perivascular lymphocytic cuff in the section of the cerebrum or of the brainstem. One experimental chicken had a single small glial nodule in its cerebrum. There was lymphocytic infiltration of one villus of the choroid plexus in two control chickens. There were no lesions in the brain or cervical spinal cord of any other birds. Most birds (both experimental and control) had a few aggregates of lymphocytes in the interstitium of the adrenal and pancreas.

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Ducks

There were no lesions in the brain or cervical spinal cord of any bird. Most birds (both experimental and control) had a few aggregates of lymphocytes in the interstitium of the adrenal and pancreas.

Geese

Three geese euthanized before inoculation (day 0), two experimental geese, and two control geese euthanized at day 45 and one experimental goose, and one control goose euthanized at day 90 had mild focal lesions in their brains. These lesions included 1 to 5 small (less than 5 cell-thick) lymphoplasmacytic perivascular cuffs or glial nodules restricted to one region of the brain (cerebrum or optic lobe).

Two experimental geese and two control geese euthanized at 90 dpi, had moderate widespread lesions in their brains. These lesions include numerous scattered, mostly small but up to 10 cell-thick, perivascular cuffs composed of mostly lymphocytes and fewer plasma cells, and diffuse and focal gliosis. Lesions were found in the cerebrum, the white and grey matters of the cerebellum, the cerebellar peduncles, the brainstem, the optic lobes, the meninges and the spinal cord. The status and time of necropsy of the geese having central nervous system histopathology is summarized in Table 5.2.

Most birds (both experimental and control) had a few aggregates of lymphocytes in the interstitium of the adrenal and pancreas.

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Table 5.2. Summary of the severity and distribution of central nervous system histological lesions observed in domestic geese used in a trial carried out to evaluate the susceptibility of domestic poultry species to experimental infection with avian bornavirus.

Histopathology Post- (central nervous system) Number mortem Status / Group Moderate of birds No Mild focal (dpi) widespread lesions lesions lesions Day 0 pre-inoculation 8 5 3 0 control 6 4 2 0 Day 45 experimental 6 4 2 0 control 7 4 1 2 Day 90 experimental 7 4 1 2

dpi = day post-inoculation.

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For the two experimental and the two control geese that had the most significant lesions in their brains, histopathological evaluation was performed on the entire set of tissues collected at necropsy. This revealed the presence of mild to moderate lymphoplasmacytic infiltration or perivascular cuffing in cervical and thoracic spinal cord, in peripheral nerves (sciatic and vagus nerves), and in the autonomic nervous system associated with the gastro-intestinal tract (myenteric ganglia of the esophagus, proventriculus, ventriculus and cloaca), the heart and the adrenal. The central, peripheral and autonomic nervous system histological lesions observed in these four cases are illustrated in Figure 5.1.

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Figure 5.1. Histological lesions observed in control (left column) and experimental (right column) domestic geese of a trial carried out to evaluate the susceptibility of domestic poultry species to experimental infection with avian bornavirus. Hematoxylin and eosin staining. A, B. Cerebrum, x40. Perivascular lymphoplasmacytic cuffing. C, D. Cerebrum, x60. Well-defined glial nodule. E, F. Sciatic nerve, x40. Perivascular lymphoplasmacytic cuffing. G, H. Proventriculus, x100. Lymphoplasmacytic infiltration or cuffing within a myenteric ganglion.

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Immunohistochemistry and RT-PCR results

The brain, cervical spinal cord, adrenal, kidney, and pancreas of all experimental and control chickens, ducks and geese were negative for ABV by IHC.

The frozen brains from all experimental and control chickens, ducks and geese and from the geese that were euthanized before inoculation were negative for ABV on RT-PCR. The FFPE brain and the frozen adrenal of the 13 experimental geese were also negative for ABV by RT-PCR.

Serology

Chickens

The mean S/P ratio and its standard deviation were -0.259 (±1.182) for the experimental birds and -0.122 (±0.752) for the control birds; this difference was not statistically significant (p=0.405).

Ducks

The mean S/P ratio and its standard deviation were 0.132 (±0.344) for the experimental birds and 0.271 (±0.517) for the control birds; this difference was not statistically significant (p=0.058).

Geese

The mean S/P ratio and its standard deviation were 0.510 (±0.477) for the experimental birds and -0.037 (±1.063) for the control birds; this difference was considered statistically significant (p<0.0001). When comparing S/P ratios between experimental and control birds at each sampling date, the difference was significant at 0 dpi (p=0.0009), 15 dpi (p<0.0001), 30 dpi (p=0.0005) and 45 dpi (p<0.0001), but not at 60 dpi (p=0.234), 75 dpi (p=0.7034) and 90 dpi (p=0.3208).

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DISCUSSION

The results of this study suggest that domestic chickens, Pekin ducks and Embden geese are not susceptible to experimental infection with the waterfowl strain of ABV. The intramuscular inoculation of 0.5 ml of brain homogenate containing ABV-CG did not result in disease in any of the experimental birds, and ABV was not detected in any brain by RT-PCR and IHC. There has been no previous experimental work using the waterfowl strain of ABV in any commercial poultry species, and this experiment was designed based on previous inoculation studies performed on psittacine birds that were published before the beginning of this trial (summer 2011). Further experiments using cultured titrated virus as the inoculum, other routes of inoculation and other species or strains of poultry are required to confirm the resistance of domestic poultry species to ABV-CG. The major weakness in the study was the failure to demonstrate viable ABV in the brains used for experimental challenge.

The only publication describing the inoculation of a poultry species with ABV was an attempt to experimentally infect ducklings with a psittacine strain of virus in 2009. This was done in an effort to minimize the experimental sacrifice of psittacine species. Twenty-two specific pathogen free (SPF) Khaki Campbell mallards were inoculated at 5 days of age or 8 weeks of age by either oral, intraocular, or intramuscular routes with DEF-cultured psittacine ABV-4. As was the case for our experiment, the inoculation did not result in disease, based on a lack of clinical signs and of macroscopic or microscopic pathological lesions on necropsies performed up to eight months after inoculation. However, several birds seroconverted and ABV was identified in the brains of two of the infected birds (Gray et al., 2009; Hoppes et al., 2010; S Payne, personal communication). It is possible that the ABV-CG used in our trial was less pathogenic than the psittacine ABV-4 used in the Khaki Campbell duck experiment. A recent study suggests that there may be differences in pathogenicity among ABV isolates with cockatiels infected with ABV-2 showing earlier and more severe clinical symptoms compared to birds infected with ABV-4 (Lierz et al., 2012).

The brain homogenates we used for inoculation tested positive by RT-PCR several times with consistently low Cp suggesting that these brains contained high viral 133

load. Attempts to grow ABV-CG were made at the time of the trial as our original intention was to use cultured ABV-CG for inoculation rather than brain homogenates; however, these attempts were unsuccessful using four different cell lines (CH-SAH, DF- 1, DEF, QT-35) and IHC and RT-PCR to detect viral particles in cell cultures. No viral titration of the inoculum was thus available. Brain homogenates containing psittacine ABV-4 were successfully used to inoculate cockatiels (Nymphicus hollandicus) via combined intramuscular, intraocular, intranasal, and oral routes and resulted in clinical signs and pathological lesions consistent with PDD (Gancz et al., 2009). Cell culture propagated virus was administered via the intramuscular route in several successful infection trials in cockatiels and Patagonian conures (Cyanoliseus patagonus) (Gray et al., 2010; Mirhosseini et al., 2011; Payne et al., 2011b). However, it is possible that, in our study, the quantity of virus inoculated was insufficient, the brain homogenates used were not infective or the intramuscular route was not appropriate.

The age of the birds at inoculation (6, 8 and 10 week-old) could also have contributed to the lack of observable disease. Most experimental trials in parrots were carried out in adult birds. We specifically chose not to inoculate the experimental birds at a too early an age as it is generally believed that lesions related to bornavirus infection are in large part immunologically mediated (Stitz et al., 1995; Stitz et al., 2002; Schwemmle et al., 2004; Baur et al., 2008) and thus that immuno-competent birds would be more likely to develop lesions than would very young birds with an immature immune system.

While the lack of disease development was not completely surprising, the fact that ABV was not found in any of the brains of the experimental birds was less expected. It is possible that the duration of our experiment was insufficient. However, in the experiment using Khaki Campbell ducklings mentioned earlier, ABV was identified in the brains of two infected birds on necropsy 6 weeks after inoculation (S Payne, personal communication). As well, in cockatiels, clinical signs and pathological lesions were found as early as 33 days post-inoculation (Mirhosseini et al., 2011). If it takes more than 90 days for disease to develop in poultry species, the relevance for the poultry industry

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with respect to meat-producing chickens, ducks or geese is less given their average life span.

Seroconversion in inoculated birds should have occurred within the time frame of the experiment: experimentally infected cockatiels seroconverted as rapidly as 6 days post infection (Lierz et al., 2012) and Khaki Campbell ducks as early as 3 weeks after inoculation (Gray et al., 2009; Hoppes et al., 2010). However, although there were significant differences in S/P ratios between the experimental and control geese at the early time points, there are several reasons why we do not believe this represents real seroconversion of the experimental birds. Differences between the two groups were significant at 0 dpi, i.e., prior to the experimental group being inoculated with ABV. The samples collected at times 0, 15, 30 and 45 dpi from the control and experimental birds were run on two different plates. The OD for the negative control was substantially higher on the control bird plate (0.605) than the same sample run on the experimental bird plate (0.498), while the OD readings for the positive control were similar (0.888, 0.892). This difference affects the S/P ratios calculated for the control geese run on this plate (0 dpi to 45 dpi) and thus the significant difference when compared to the S/P ratios of the experimental geese.

As discussed in Chapter 3, the assessment of the presence of antibodies in the experimental infections we performed was hindered by several methodological and technical difficulties. For the chicken samples, the secondary antibody was specific “anti- chicken” whereas a generic “anti-bird” IgG antibody conjugate was used for the duck and geese samples. This latter conjugate was used previously for the detection of antibodies to ABV in avian sera (de Kloet et al., 2011), and its reactivity towards immunoglobulins for both ducks and geese was initially verified by dot blots (P Delnatte, Unpublished data). The intensity of OD405 readings in our ELISA varied greatly among the 3 species, with OD405 values being much higher for the chickens and ducks than for the geese.

Given the fact that OD405 readings higher than 1 are usually considered to be less reliable and more difficult to interpret than OD405 readings in the 0 to 1 range, optimum serum dilution for each species was determined using standard checkerboard analyses prior to performing the ELISAs. Thus the duck and chicken sera were diluted 1:200 compared to

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a 1:20 serum dilution for the geese. These species variations precluded any direct comparison of S/P ratios among species. Despite our efforts to standardize the ELISA technique, several seemingly random, individual OD readings (and thus S/P ratios) in both control and experimental groups were much higher than the rest of the values, including values from the same birds at previous or later sampling dates. These S/P ratios were considered outliers. In order to prevent over-interpretation of these individual, potentially erroneous, results, we compared the S/P ratios for each species at a group level (experimental versus control) rather than on an individual basis.

The cloacal swabs collected at regular interval during this study were not evaluated for the presence of ABV, and thus the possibility of viral shedding could not be definitively ruled out. In order to comply with time and budgetary constraints, diagnostic tests were performed following a pre-defined logical order. Our first objective was to evaluate whether the inoculation of these birds with brain homogenates containing ABV- CG resulted in ABV-infection. Because ABV antigen has been shown to be most consistently detected in brain, spinal cord, adrenal gland, pancreas, and kidney using IHC in ABV-infected parrots (Raghav et al., 2010), we initially submitted these tissues for histopathology and IHC for all control and experimental birds. Similarly, because the brain is one of the most reliable tissues to test for ABV infection in psittacine birds by RT-PCR, the brains of all control and experimental birds were submitted for RT-PCR. Given that none of these samples tested positive for the presence of ABV on IHC and RT-PCR, further investigation was carried out on only the domestic geese as histological lesions were present in the brains of several birds. Frozen adrenals and FFPE brains of all experimental geese were subsequently submitted for RT-PCR and also tested negative. To better characterize the type and distribution of lesions in the four geese where significant lesions were seen in the central nervous system, the entire set of tissues collected from each bird was submitted for histopathological evaluation.

In this study, multiple histological sections of the central nervous system were meticulously reviewed to detect subtle lesions. The very mild focal lesions observed in a few chickens (both control and experimental) were considered incidental and would have likely been discounted during a routine histological screening (Gimeno et al., 2005).

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However, the changes observed in the nervous system tissues of several control and experimental geese were more compelling. Brains from the eight birds euthanized prior to inoculation were examined and found to contain similar lesions.

The presence of lymphoplasmacytic infiltrates within the central, peripheral and autonomic nervous system, and particularly within the myenteric ganglia of the proventriculus and ventriculus, are often considered pathognomonic for PDD in psittacine birds (Schmidt et al., 2003; Berhane et al., 2004). In a retrospective study of ABV infection of Canada geese and trumpeter and mute swans, correlation between the presence of ABV and lesions in these locations was also very high (Delnatte et al., 2013; Chapter 2) Although we cannot definitively rule out the possibility of the occurrence of a different genotype of ABV that remained undetected by our RT-PCR and IHC, this hypothesis is very unlikely. The duplex RT-PCR and the IHC used in this trial detect various psittacine strains of ABV as well as ABV-CG, which share 85% amino-acid sequence identity (Delnatte et al., 2013).

The goslings were purchased at one day of age after hatching at a non-SPF standard farm. A brief retrospective search for pathology cases originating from this facility during the time period when the goslings were obtained (summer 2011) revealed one 8 week-old gosling with numerous small to medium-sized glial nodules in the cerebellum and cerebrum. These lesions were considered incidental; the cause of death of the gosling was attributed to urate nephrosis and visceral and articular gout. Evidence thus points to the presence of a non- or poorly-pathogenic neurotropic virus that was present at the original facility and infected the birds prior to their delivery to the Isolation Facility.

Neurotropic viruses described in waterfowl include, but are not limited to, West Nile virus (WNV), avian paramyxoviruses (APMV), influenza viruses (AIV), eastern equine encephalitis virus (EEEV) and Marek’s disease virus (MDV). The brains of the four geese exhibiting the most severe brain lesions were negative for WNV, APMV, AIV and EEEV by RT-PCR. WNV virus has been associated with severe clinical neurological signs, meningo-encephalitis and myocarditis with high fatality in both naturally and experimentally infected young domestic geese (OIE, 1999; Swayne et al., 2001; Austin et 137

al., 2004; Meece et al., 2006). The geese in this trial were clinically normal and there was no evidence of myocarditis or myocardial necrosis in the four cases for which the full set of tissues was reviewed histologically. APMV-1 has been described in geese (Jinding et al., 2005) but is considered very rare in waterfowl (Woolcock et al., 2008). AIV has been isolated from numerous species of Anseriformes; however mildly pathogenic AIV have a marked tropism for the respiratory tract, disease of which was not observed in these cases; and highly pathogenic AIV has never been documented in Ontario. MDV has a rather narrow host range within the Galliformes, being primarily a disease of chickens and to a lesser extent quails, turkeys and pheasants (Schat et al., 2008). There are very few reported cases of Marek's disease in waterfowl. MDV genome has been detected from feather tips of healthy wild geese in Japan and Russia but no histological examination was performed and thus this finding may not be relevant in our case (Murata et al., 2007). The existence of a yet to be discovered, non- to low- pathogenic neurotropic virus in domestic geese is possible.

In conclusion, the inoculation of brain homogenates containing ABV-CG did not result in disease or in ABV infection in domestic chickens, Pekin ducks and Embden geese. Further experiments are required to definitely confirm a lack susceptibility to ABV-CG infection and/or resistance to the development of ABV-CG-related disease, especially given our failure to prove the presence of viable virus in our experimental inoculum. The presence of pre-existing lesions in the brain of several control and experimental geese with no evidence of ABV infection highlighted the complexities of experimental infection trials in non-SPF animals, and the difficulties in the diagnosis of neurotropic viral infection based on histological evaluation alone.

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CONCLUSIONS – FUTURE DIRECTIONS

The overall objectives of this study were to acquire further knowledge about the pathology and epidemiology of avian bornavirus in free-ranging waterfowl, and to assess the susceptibility of domestic poultry species to this virus.

We showed that free-ranging waterfowl in Ontario are infected with a unique genotype of ABV that can be associated with significant neurological disease (clinical signs and pathological lesions) in Canada geese, trumpeter swans and mute swans. ABV should thus be considered as a potential pathogen in these species and should be included in the differential diagnosis for any Anseriforme exhibiting upper gastrointestinal impaction or non-specific neurological signs, and having non-suppurative inflammation of the central, peripheral or autonomic nervous systems.

Our prevalence survey revealed that ABV exposure or infection is widespread in wild, apparently healthy Canada geese, trumpeter swans, mute swans and mallards in southern Ontario, and provided the first estimates of the prevalence of cloacal shedding of ABV and of presence of antibodies against ABV in free-ranging Anseriformes.

We were not able to infect domestic poultry species with the waterfowl strain of ABV, which suggests that the risk of spread of ABV from free-ranging waterfowl to the poultry industry is low, if it exists. There were; however, several limitations in our experimental trial and thus further investigation, including using other routes of inoculation and other sources of virus and animals, would be required to definitively rule out the possibility of transmission of ABV-CG to domestic poultry.

Since the first description of ABV in 2008 in psittacine birds, an impressive amount of data, mostly on parrots, has been gathered through research worldwide. However, many questions about bornaviruses, and more specifically about ABV in waterfowl, remain open. The exact pathogenesis of ABV-related disease still remains to

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be elucidated. Future research should be directed at determining the factors that trigger the development of disease in infected birds.

The remarkable genetic variability among ABVs suggests that additional bornaviruses will likely be discovered in other avian and non-avian species and new geographic locations in the near future. Further work on the molecular characterization of the bornaviruses is necessary and would help understanding the evolutionary relationship among them.

Avian bornavirus infection is likely under-recognized in wild waterfowl and it is likely that waterfowl constitute the largest pool of bornaviruses. Free-ranging waterfowl from other continents should be evaluated for the presence of ABV. The exact role of wild birds in the epidemiology of ABV and BDV is unknown and the natural reservoirs of these viruses remain unidentified.

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APPENDIX A: BIRDS AND BORNAVIRUSES – REVIEW

Payne SL, Delnatte P, Guo J, Heatley JJ, Tizard I, Smith DA. 2012. Birds and bornaviruses. Anim Health Res Rev 13:145–156. doi: 10.1017/S1466252312000205.

“Reprinted with permission”

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*c Cambridge University Press 2012 Animal Health Research Reviews 13(2); 145–156 ISSN 1466-2523 doi:10.1017/S1466252312000205

Birds and bornaviruses

Susan L. Payne1*, Pauline Delnatte2, Jianhua Guo1, J. Jill Heatley3, Ian Tizard1 and Dale A. Smith2 1Department of Veterinary Pathobiology and the Schubot Exotic Bird Health Center, Texas A&M University, College of Veterinary Medicine and Biomedical Sciences, College Station, TX 77843, USA 2Department of Pathobiology, Ontario Veterinary College, University of Guelph, Guelph, Ontario, Canada 3Department of Small Animal Clinical Sciences, Texas A&M University, College of Veterinary Medicine and Biomedical Sciences, College Station, Texas, USA

Received 18 September 2012; Accepted 1 October 2012

Abstract In 2008, avian bornaviruses (ABV) were identified as the cause of proventricular dilatation disease (PDD). PDD is a significant condition of captive parrots first identified in the late 1970s. ABV infection has subsequently been shown to be widespread in wild waterfowl across the United States and Canada where the virus infects 10–20% of some populations of ducks, geese and swans. In most cases birds appear to be healthy and unaffected by the presence of the virus; however, infection can also result in severe non-suppurative encephalitis and lesions similar to those seen in parrots with PDD. ABVs are genetically diverse with seven identified genotypes in parrots and one in canaries. A unique goose genotype (ABV-CG) predominates in waterfowl in Canada and the northern United States. ABV appears to be endemic in North American waterfowl, in comparison to what appears to be an emerging disease in parrots. It is not known whether ABV can spread between waterfowl and parrots. The discovery of ABV infection in North American waterfowl suggests that European waterfowl should be evaluated for the presence of ABV, and also as a possible reservoir species for Borna disease virus (BDV), a related neurotropic virus affecting horses and sheep in central Europe. Although investigations have suggested that BDV is likely derived from a wildlife reservoir, for which the shrew and water vole are currently prime candidates, we suggest that the existence of other mammalian and avian reservoirs should not be discounted.

Keywords: bornavirus, goose, swan, duck, canary, gull, phylogeny, proventricular dilatation disease, avian

Introduction the associations between members of the family Borna- viridae and birds, especially North American waterfowl. Until 2008, the family Bornaviridae consisted of one virus – Borna disease virus (BDV), known primarily as a cause of neurological disease in horses and ruminants in a geographically restricted area of Central Europe. The 2008 Borna disease virus identification of multiple genotypes of a novel avian bornavirus (ABV) opened the door to investigations on History the role of these viruses in causing disease in captive and free-ranging birds. The purpose of this article is to review The town of Borna lies in the eastern part of Germany near Leipzig, in the state of Saxony. The region has long been associated with epidemics of a unique neurologic *Corresponding author. E-mail: [email protected] disease of horses, especially a devastating epidemic 146 S. L. Payne et al. among cavalry horses between 1894 and 1896 (Richt association between BDV proteins and cellular chromatin et al., 2000). Affected animals show a diverse array of (Matsumoto et al., 2012). This association probably changes in behavior including ataxia, head tilt, muscle ensures that viral genomes are effectively delivered to fasciculation, hind-limb paresis, localized hypo- or hyper- daughter cells during mitosis. aesthesia, disturbances in chewing and swallowing, and aggression. A similar disease occurs in sheep. The pathology of the disease consists of a virally induced Borna disease pathology/immunopathology progressive, non-suppurative encephalomyelitis characterized by lymphocytic infiltrates affecting the gray matter BDV can infect a variety of mammals and birds, including (Lipkin and Briese, 2006). This disease is restricted to a rodents, non-human primates, chickens and ostriches fairly small geographic area encompassing central and (Carbone, 2001; Tomonaga and Carbone, 2002). In many southern Germany and neighboring countries, and was cases, animals become persistently infected but fail to named Borna disease after its region of origin. Serologic develop disease. This is not surprising as, in tissue culture, studies have shown that while BDV infection is wide- the virus appears to have minimal adverse effects on cell spread in horses and sheep within the affected region, function or survival. A small proportion of infected only a small fraction of these infected animals actually animals develop a progressive neurologic disease. develop clinical disease. Once disease develops, Experimentally, disease development varies with host however, Borna disease mortality may reach 100% in species and age (reviewed in Lipkin et al., 2011). For horses and 50% in sheep. example, 4–5-week old Lewis rats respond to intracranial infection with transient behavioral abnormalities and develop meningoencephalitis and retinitis characterized Characteristics of the virus by perivascular mononuclear cell infiltrations (Narayan et al., 1983). In contrast, newborn rats can be chronically BDV is an enveloped, non-segmented negative strand infected and develop subtle behavioral abnormalities RNA virus with a genome size of approximately 8.9 kb. but the infection is non-fatal (Lipkin et al., 2011). The The bornaviral genome encodes six proteins: nucleocap- differences in these scenarios appear to result from sid (N), X protein (X), phosphoprotein (P), matrix (M), differences in host immune function. Specifically, brain envelope glycoprotein (G) and the RNA-dependent RNA damage results from the activities of host T cells (Rott polymerase (L). Its overall genomic organization is similar et al., 1988). The presence of CD8+ T cells in the brain to that of other viruses in the order Mononegavirales parallels the onset of neurologic dysfunction. Conversely, including the paramyxoviruses and rhabdoviruses. suppression of T cell responses reduces the severity of However, BDV is the only non-segmented, negative experimental BDV infections (Narayan et al., 1983). BDV strand RNA virus to replicate within the nuclei of infected spreads throughout the host largely through neural cells and hence was placed into its own family, the networks (Carbone et al., 1987; Ackermann et al., 2010) Bornaviridae. Bornaviruses also have a unique genome and any route of infection that allows virus access to the replication strategy that involves trimming of 50 -terminal nervous system will eventually lead to CNS infection and nucleotides (Schneider et al., 2005). disease in susceptible adult rats (Lipkin and Briese, 2006). Virions appear to be enveloped particles of 80–100 nm, with 50–60 nm electron dense cores, but are difficult to visualize (Lipkin et al., 2011). In tissue culture, BDV is Borna disease epidemiology highly cell-associated, non-cytopathic, and found in only small amounts in cell culture supernatants (Schneider, Serologic studies based on ELISA assays suggest that up to Schwemmle and Staeheli, 2005; Tomonaga et al., 2002). 12% of horses in central and southern Germany may have Many mammalian cell types are permissive for BDV been exposed to BDV, implying that inapparent infec- replication. Cells in which BDV can be cultured include, tions are not uncommon (Lipkin and Briese, 2006). The but are not limited to human oligodendrocyte (OL), status of Borna disease in hoofstock, outside its core African green monkey kidney (Vero), Madin–Darby range, is unclear but it appears to be present in Israel, canine kidney (MDCK), rat glial cells (C6,) human Japan and other Asian countries (Richt et al., 1997). embryonic kidney cells (HEK) and guinea pig 1505 cells Studies within the core range have long suggested that the (Herzog and Rott, 1980; Staeheli et al., 2000; Schneider, virus is maintained within a natural reservoir, and rodents Schwemmle and Staeheli, 2005). have been the prime suspects. BDV has been found in the BDV spreads predominantly by direct cell-to-cell bicolored white-toothed shrew (Crocidura leucodon)in contact. Indirect immunofluorescence assays using anti- Central Europe (Hilbe et al., 2006), but the shrew cannot bodies against the viral N or P proteins reveal a be held wholly responsible for the transmission of Borna characteristic speckled pattern in the nucleus. These disease, as it is not found everywhere that disease intranuclear inclusions (karyosphaeridia) are known as occurs. In Finland, seropositive bank voles (Myodes Joest–Degen bodies and form as a result of the close glareolus) were identified by Kinnunen et al. (2007). Birds and bornaviruses 147

Domestic mammals are believed to acquire infection as a BDV, sharing 95.9–99% nucleotide sequence identity with result of eating or inhaling on pastures contaminated with BDV sequences from mammals. The jackdaw and mallard shrew urine. While other reservoirs have not been sequences were not identical to one another, sharing formally demonstrated, there is evidence that birds may 98–98.5% nucleotide sequence identity (Berg et al., 2001). play a role in maintaining and disseminating this disease. The publications cited above predate the discovery of ABV and involve little or no genetic characterization of the putative bornaviruses. The role of avian species in the BDV and birds epidemiology of BDV thus remains unclear, and there is as yet no evidence for the presence of any other members A few reports have noted the presence of BDV in birds, of the family Bornaviridae in wild birds in Europe. either as a cause of clinical disease or as inapparent Additional evidence also suggests that links may exist infections playing a part in transmission to more between BDV and an avian vector. For example, Borna susceptible species. A review by Rott and Becht (1995) disease recurs in specific areas or on individual farms reported that chickens are ‘frequently susceptible’ to BDV during spring and summer months (April, May and June) and cited previous studies, including those of Zwick at several year intervals (Du¨rrwald, 1993; Richt et al., et al. (1927). Experimental infections of chickens were 2000). The prevalence of disease drops significantly in the also described by Ludwig et al. (1973). In that study, 1-day autumn and winter months. BD tends to occur at low old chicks were inoculated intracerebrally with brain altitudes and there may be an association with river homogenates from rabbits with Borna disease; at 5–8 valleys – sites where waterfowl occur. Staeheli et al. weeks post-inoculation, 9 of 13 chicks developed (2000) pointed out the remarkable stability of the paralysis of legs and wings. After sacrifice the affected endemic area in central Europe over many years, despite birds were found to have intranuclear Joest-Degen the widespread movement of animals into and out of the inclusion-bodies in brain sections and anti-BDV anti- area. Clusters of BDV appear to have different origins, bodies were demonstrated by immune-diffusion tests which tends to exclude the spread of a single virus from a using sera from recovered chickens (Ludwig et al., 1973). single point of origin. This is somewhat surprising given BDV was identified as the cause of an outbreak of the widespread movement of livestock, especially horses, neurologic disease in ostriches (Struthio camelus)in between these areas, but is consistent with the apparent Israel (Malkinson et al., 1993a, b). Between 1989 and lack of direct spread among farm animals (Staeheli et al., 1992, 7–26% of all hatched chicks on affected farms died 2000). More recently, Kolodziejek and her colleagues from paresis (Asash et al., 1996). In one group of ostriches have also demonstrated, by genetic analysis, that within BDV antigen was detected, using an ELISA, in 7 of 13 the Borna disease endemic area there is clear clustering of brain homogenates from paralyzed birds, whereas it was different genetic strains of the virus (Kolodziejek et al., found in only 1 of 10 brains from healthy birds. Brain 2005). Thus specific genetic sequences are associated extracts from paralyzed ostriches, given orally or intra- with certain German, Austrian and Swiss regions, which muscularly to 5-week-old ostrich chicks, reproduced the have no obvious geographic barriers between them. clinical signs and microscopic lesions of naturally infected Studies such as those described above support multiple birds. Whether this disease outbreak was in fact caused by introductions of genetically stable bornaviruses into an avian bornavirus is unknown; the described pathologic livestock. Small mammal reservoirs of BDV have been lesions were not at all similar to those seen in parrots or identified, but there is no evidence of widespread BDV waterfowl. Unfortunately these tissues are not available infection in the Borna disease endemic area or elsewhere. for additional testing. An avian vector of BDV was also suggested by studies Beginning in the late 1980s, a neurologic disease on the epidemiology of the outbreak of Borna disease in characterized by stiffness leading to paralysis and death ostriches in Israel. Teplitsky et al. (2003) showed that was observed in cats in Central Europe and Scandinavia horses in the region were seropositive for BDV as (Lundgren, 1992). BDV was subsequently isolated from determined by an ELISA. Interestingly, the positive some of these cats and challenge of pathogen-free cats samples came either from the coast or the Jordan valley with BDV induced a neurologic disease resembling the rather than from the highlands between these two natural disease. The source of this cat infection was regions. The authors pointed out that the coast and valley unclear but Berg and his colleagues in Sweden (Berg are major flyways for migrating birds and went on to et al., 2001) speculated that the cats might have acquired suggest the possibility that birds may serve as vectors of this infection from wild birds. They investigated this BDV in Israel. possibility by using a nested RT-PCR assay to test for the presence of BDV in bird droppings and in fact detected BDV sequences in the droppings of a mallard (Anas Avian bornaviruses platyrhyncos) and a jackdaw (Corvus monedula) from an urban pond in Uppsala, Sweden. The partial genome Work thus far suggests that ABV behave much like BDV in sequences of these Swedish bird isolates were clearly culture. Infections are non-cytopathic and there is no 148 S. L. Payne et al.

Canada geese mute swans trumpeter swans ABV 4

ABV canary

ABV 2 Psitacine ABV 5 Psittacine

ABV3

ABV 1 Borna disease ABV 7 Mallard (Sweden) viruses Endogenous N Spermophilus 0.05 tridecemlineatus Nucleotide substitutions per site

Fig. 1. A phylogeny was generated using partial N-gene sequences of bornaviruses from birds and mammals. A consensus tree was generated using a Neighbor-Joining algorithm, with no outgroup assigned. To generate the consensus tree 1000 bootstrap replicates were generated. evidence that a significant amount of virus is released genotypes in captive parrots worldwide (Ogawa et al., (Rinder et al., 2009; Staeheli et al., 2010). In ABV-infected 2011; Rubbensroth et al., 2012). cells, viral antigen is found in the nucleus and shows the Two additional ABV genotypes have recently been same speckled immunofluorescence pattern as BDV identified. One genotype was recovered from a canary (Rinder et al., 2009; Gray et al., 2010). Not surprisingly, (Serinus canaria) and is identified as ABV-canary bird cells are more permissive for ABV replication than (Weissenbock et al., 2009b; Rinder et al., 2012). The are mammalian cells. Cells used successfully for ABV second non-psittacine ABV genotype was recovered from culture include primary duck embryo fibroblasts (DEF), a wild Canada goose (Branta canadensis) (Delnatte et al. the quail fibroblast cell line CEC32, the quail skeletal cell 2011), and was named ABV-CG. This was the first ABV line (QM7) and a chicken hepatoma cell line (Rinder, identified from wild birds. As described below, ABV-CG is et al. 2009; Gray et al., 2010; Villanueva et al., 2010). common across North America and has been isolated Attempts to grow ABV in mammalian cells (Vero, MDCK from trumpeter swans (Cygnus buccinator) and mute or C6 cell lines) have been unsuccessful (Rinder, et al., swans (Cygnus olor) (Delnatte et al., 2011; Guo et al., 2009). In vivo, ABV appears to be widely disseminated. 2012). Thus, to date nine ABV genotypes have been Viral antigen can be found in a broad spectrum of organs identified. and cell types in diseased birds (Rinder et al., 2009; To illustrate the genetic relationships among the Raghav et al., 2010). This is in sharp contrast to BDV, bornaviruses, the phylogeny (shown in Fig. 1) was which has a preference for cells of the central and generated using a short fragment of the nucleocapsid peripheral nervous systems. (N) gene. The phylogeny shows the psittacine ABVs on multiple branches of the tree. As ABV genotypes 1–7 and ABV-canary were recovered from captive birds it is impossible to meaningfully correlate virus genotype to ABV genotypes the region of origin of the host. It also appears that there is no obvious correlation between psittacine species and In the two studies that first identified ABV in parrots, five infecting genotype. For example, ABV4 and ABV2 are genotypes (ABV1–5) were recognized on the basis of found in captive birds of South American, African and nucleotide and amino acid sequence identity (Honka- Australian species. In contrast, the bornaviruses identified vuori et al., 2008; Kistler et al., 2008). The ABV genotypes, in wild Canada geese, mute swans and trumpeter swans all with <70% nucleotide sequence identity with BDV, form a tight cluster with over 90% nucleotide sequence were sufficiently different as to comprise a new species identity among them (Payne et al., 2011a, b; Guo et al., (Honkavuori et al., 2008; Kistler et al., 2008). Two 2012). Thus, it appears that, at least in North America, additional psittacine bornavirus genotypes (ABV 6 and there may be a predominant genotype circulating among 7) were subsequently identified (Weissenbock et al., free-living waterfowls. 2009a; Rubbensroth et al., 2012). Among these seven Natural cases of Borna disease are most prevalent in genotypes, ABV4 and ABV2 are by far the most common Central Europe, and the BDVs from that region are for the Birds and bornaviruses 149 most part closely related, forming their own cluster. BDV 2012). International trade in captive parrots subsequently was also found in ducks in Sweden, and that finding resulted in the disease being widely spread geographi- suggests a possible correlation between genotype and cally. PDD has yet to be identified in a wild parrot geographic location. population. In 2008, the intensive search for an etiology A particularly intriguing question is the nature of the ended when a novel bornavirus was identified and relationship between ABV infection in waterfowl and in subsequently shown to be the causative agent of PDD parrots. As described earlier, PDD first appeared in (Honkavuori et al., 2008; Kistler et al., 2008; Gancz et al., captive parrots in the USA in the mid-1970s at a time 2009; Gray et al., 2010). when many species of birds were being captured in the PDD is characterized histopathologically by non- wild and imported into Europe and the USA. These birds suppurative inflammation in the central, peripheral and were subjected to quarantine while being tested for autonomic nervous systems (Berhane et al., 2001; Raghav Newcastle disease, and facilities were not designed to et al., 2010). Clinical signs reflect the wide distribution of prevent cross species contact. Based on the lack of lesions, but are generally classified as neurologic or recognition of PDD in psittacine birds prior to this date, gastrointestinal. They include weakness, ataxia, proprio- on the rapid spread of the disease globally, and the lack ceptive deficits, seizures and blindness; weight loss, of reports of PDD (or ABV infection) in wild psittacine passage of undigested food, regurgitation and delayed populations, it is tempting to hypothesize that macaws crop emptying, respectively. Gastrointestinal malfunction were infected by contact with ABV-infected waterfowl results in the most common gross pathologic lesions – while in quarantine. Mixing of species and high stress dilation and thinning of the walls of the proventriculus levels would have enhanced viral transfer and perhaps and ventriculus, and in maldigestion, emaciation and facilitated development of disease in the new host. The death by starvation (Hoppes et al., 2010). Birds may presence of multiple discrete ABV genotypes in psittacine present with any combination of these signs, and the birds suggests multiple introductions, perhaps from severity and progress of the disease can vary tremen- different sources. Evaluation of waterfowl (or other wild dously. Subclinically affected birds may show very few if birds) for ABV genotypes has hardly begun, and has thus any clinical signs over long periods of time (Raghav et al., far largely been restricted to examining birds within 2010; Villanueva et al., 2010). continental North America. There is no information ABV infects primarily the central and enteric nervous currently available regarding the presence of ABV in wild systems of birds. Although the virus is non-cytopathic it birds in South America, Asia, Africa or Australasia; thus can induce inflammation and a selective loss of glial cells the potential for discovering other genotypes in other and neurons. Based on the known pathogenesis of BDV, species and geographic locations is high. it is believed that this cell loss is secondary to T cell A recent observation that further complicates our cytotoxicity although this has not yet been formally understanding of the relationships among bornaviruses demonstrated for ABV. ABV can be detected throughout is the identification of bornavirus sequences integrated the CNS and enteric nervous systems but is also found in into the genomes of a variety of mammals (Belyi et al., the kidneys, adrenals and gonads and in cell types other 2010; Horie et al., 2010; Katzourakis and Gifford, 2010). It than neurons (Rinder et al., 2009; Raghav et al., 2010; appears that bornavirus genes found their way into Weissenbock et al., 2010; Payne et al., 2011b). primate genomes over 40 million years ago, and that integration events occurred more than once. One North Identification of ABV in birds American mammal, the 13-lined ground squirrel (Spermo- philus tridecemlineatus) has an intact bornavirus N gene Psittacines in its genome. The endogenized N gene is more closely related to currently circulating ABV than to the endogen- ABV is readily detected by both RT-PCR and immunohis- ous bornavirus-like elements (EBLN) in human and tochemistry (IHC) in necropsy tissues from parrots with non-human primates. clinical and histopathological evidence of PDD (Raghav et al., 2010). It is present in all major organs, especially the central nervous system. In live birds, the identification of Proventricular dilatation disease (PDD) virus and the determination of its significance are less straightforward. ABV is often detectable in cloacal swabs PDD is a fatal neurologic condition of captive parrots and the urofeces of birds affected with PDD and this has (Hoppes et al., 2010). The disease was first identified in been recommended as the primary antemortem method North America and Europe during the mid-1970s at a time of identifying birds affected with the disease. However, when extensive trading in wild parrots was permitted. Its surveys have shown that ABV can be found in droppings source or sources are unknown (Graham, 1984), but from apparently healthy captive psittacines, and shedding South America was suspected to be the site of origin as of virus from the ABV-positive birds is often intermittent the disease was often observed in macaws imported from and unpredictable (Raghav et al., 2010; Villanueva et al., Bolivia (S. Clubb, personal communication, October, 2010). For example, ABV was detected in droppings from 150 S. L. Payne et al. apparently healthy parrots of several species, including seen in psittacine birds with PDD. ABV was identified in multiple cockatiels (Nymphicus hollandicus) purchased 11/12 goose brains and 2/2 swan brains by IHC, using from several local aviaries without a history of PDD rabbit polyclonal antiserum against ABV N-protein, as (Villanueva et al., 2010). Some clinically unaffected birds well as by RT-PCR testing for N-protein genes. Sequence shed virus for at least 5 years. analysis of the amplified gene products confirmed the Recently, serologic tests appear to be of limited presence of ABV in Canada geese and identified it as a usefulness in disease diagnosis. Indirect fluorescent unique new genotype (Delnatte et al., 2011). antibody testing (Lierz et al., 2012) and Western blot Subsequently, Delnatte and her colleagues selected 51 assays (Villanueva et al., 2010) appear to be the most necropsy reports from Canada geese, trumpeter swans sensitive and specific tests. Both tests appear to be and mute swans, euthanized or found dead in southern 90–95% sensitive and specific. They cannot how- Ontario, based on the presence of upper gastrointestinal ever differentiate between diseased birds and healthy impaction, central nervous system histopathology or a carriers. clinical history suggestive of ABV infection (Delnatte et al., 2012a). IHC and conventional RT-PCR for the N-protein gene and quantitative RT-PCR (qPCR) for Passerines the M-gene on fresh and formalin fixed paraffinized tissue again revealed the presence of ABV in birds Since the first appearance of PDD in large psittacines, with widespread non-suppurative encephalomyelitis avian pathologists have, from time to time, noted and ganglioneuritis. As with psittacine PDD, the clinical neurologic lesions in other bird species that ‘look like’ history and gross necropsy findings in these birds PDD (Daoust et al., 1991; Perpinan et al., 2007). However, generally reflected the occurrence of nervous system this was impossible to prove without the ability to detect lesions and included neurologic disease, weakness or the presence of ABV in the lesions. Since the discovery inability to fly, and signs suggesting possible defects in of ABV, infections have been detected in canaries gastrointestinal motility such as upper gastrointestinal (S. canaria). Weissenbock et al., (2009b) examined a tract impaction. In 1991, Daoust and colleagues described bird that had died after a few days of ‘apathy’. On two cases of PDD-like disease in Canada geese from necropsy the bird had a severely dilated proventriculus. Prince Edward Island (Eastern Canada) (Daoust el al., On histopathology the bird had a non-suppurative 1991), but this was before the identification of ABV. encephalitis and ganglioneuritis in the proventriculus and Recently, archived tissues from these birds were found to ventriculus. Bornaviral antigen was detected in multiple be positive for ABV based on RT-PCR (Delnatte et al., tissues and confirmed by RT-PCR. Sequence analysis 2012a), and one Canada goose from the province of demonstrated that this was a unique genotype of ABV. Quebec showing consistent neuropathology has also Subsequently, Rinder et al. (2012) described two canaries been shown to be ABV-positive (D.A. Smith, unpublished with neurologic disease associated with the presence of work). The conclusion of this work was that ABV ABV. One bird showed ‘apathy’ and sudden death; the infection can be associated with significant neuropathol- other showed prolonged depression, neurologic disease ogy in waterfowl species, and that the resulting disease is (head tilting and inability to fly), and visual impairment very similar to PDD in psittacine birds. with chorioretinitis. Necropsy showed a dilated proven- Comparisons of the site of origin of 40 Canada geese triculus with ganglioneuritis and non-suppurative ence- affected by ABV-related neurological disease suggested phalitis. Sequencing identified variants of the previously an uneven geographic distribution. Using retrospective reported canary genotype. Canaries are archetypical necropsy data, the proportion of affected wild geese passerines and the ability of ABV to cause disease in this found dead or euthanized at a large urban zoo was species suggests the possibility that all passerines may be significantly higher than that found elsewhere in the susceptible to this virus. province. Unequal surveillance intensity made estimation of the true prevalence of the disease in these two locations impossible to determine. Geese Cloacal swabs were subsequently collected from 200 free ranging Canada geese in four different locations, Smith and colleagues were the first to identify the including the zoo site, to better estimate the prevalence presence of ABV in waterfowl, specifically Canada geese of ABV infection in Ontario (Delnatte et al., 2012b). The and trumpeter swans during a retrospective survey of prevalence of fecal shedding at the zoo site was diseases of wild birds in Southern Ontario (Smith et al., significantly higher than at the other three collection sites 2010). These samples had been obtained from birds (7/50 versus 0/150), supporting their premise that suffering from neurologic disease of unknown origin but environmental or ecological factors can affect the with consistent neuropathology. Careful examination of prevalence of ABV infection and subsequent disease. central, peripheral and autonomic nervous tissue revealed Canada geese are often considered to be nuisance birds non-suppurative inflammatory lesions similar to those when they gather in large permanent flocks in public Birds and bornaviruses 151

a single nuisance ﬂock while four were from hunter-killed birds. RT-PCR testing also has been carried out on the brain materials of hunter-killed geese in Texas and Kansas. These include 115 snow geese (Chen caerulescens), 58 Ross’s geese (Chen rossii) and 10 greater white-fronted geese (Anser albifrons). None of the samples from greater white-fronted geese were ABV-positive, but 10% of Ross’s geese and 19% of snow geese samples were positive.

Swans Swans Ducks 1–5 5–10 > 10 Geese Trumpeter swans were native to Eastern Canada until extirpated as a result of overhunting by the early 20th Fig. 2. Sampling locations and the number of RT-PCR- Century. Beginning in the 1980s, a restoration project has positive samples at each site. The smallest circles represent succeeded in reintroducing this bird to its former range. 1–5 positive samples, the medium circles show 5–10 positive There are now more than 1000 trumpeter swans in samples and the largest circles represent states or provinces from which >10 positive samples were obtained. The Ontario and more than 100 breeding pairs established numbers do not reflect prevalence at any site. Of the states (Moser, 2006; Lumsden, 2009). Close monitoring of this where Canada geese were tested, only Ohio had no positive population and an appreciation of the importance of lead samples detected. Likewise, of the states where mute swans poisoning as a cause of morbidity and mortality, result in were tested, only New York provided no positive samples. the early recognition and capture of clinically abnormal birds, and an enhanced likelihood of the submission for spaces such as parks and playgrounds. In the USA, these necropsy of birds that die. In the retrospective study flocks are culled by the Wildlife Services Agency of the US performed by Delnatte et al. (2012a), ABV-associated Department of Agriculture. These culled birds are neuropathology was present in six of eight trumpeter routinely swabbed in the oropharynx and the cloaca to swans that met the study inclusion criteria intended to determine the presence of avian influenza viruses. Payne select ABV affected birds from a large database. Among and her colleagues extracted RNA from influenza- the birds presented for necropsy, the proportion of negative swabs and subjected them to RT-PCR analysis trumpeter swans meeting the inclusion criteria was lower for ABV M-genes (Payne et al., 2011a, b). In their first than that for Canada geese but did not differ between the survey, 300 samples consisting of 25 samples from each of two sites of sample origin. There was no significant 12 states across the USA were obtained. Subsequently, an difference in virus detection frequency between the two additional 109 swab samples were surveyed from the sites of origin, which is not surprising as all swans in three western coastal states. Of the 409 goose swab Southern Ontario belong to a single mobile population. samples tested, 24 from 5 states were positive for ABV In contrast to trumpeter swans, mute swans are an M-genes (Fig. 2). Twelve of these products (2.9%) were introduced species in North America and are found subsequently sequenced and confirmed to be of the ABV predominantly in the northeast. They are often consid- goose strain genotype. Of the states where Canada geese ered nuisance birds because of their aggression and their were tested, only Ohio had no positive samples detected. destructive effects on the environment. As a result, some These results also showed that viral prevalence was not US states permit their capture and euthanasia. Captured uniform across the continent, and that infections could be birds were therefore available to permit estimation of the clustered within flocks. For example, 20–42% of birds prevalence of ABV in this species. Combined orophar- within selected flocks in Kansas, New Hampshire and yngeal and cloacal swabs were collected from 219 mute New York were RT-PCR-positive. There were no apparent swans. Fourteen of these samples gave a product of gender differences in infection prevalence in either this appropriate size on gel electrophoresis (Guo et al., 2012). study or the Canadian studies. Four of these positive samples were selected for sequen- While cloacal/oropharyngeal swabs provided a conve- cing and were confirmed to be ABV M-gene sequences. nient method of testing wild goose flocks, experience Phylogenetic analysis of these amplified sequences based on psittacine testing suggests that this is a relatively indicated that the genes clustered in a single group insensitive method of detecting ABV infection (Villanueva closely related to the M-genes of the previously described et al., 2010). For this reason 25 freshly frozen Canada goose isolates (Payne et al., 2011a, b). goose heads were obtained from sites in New Jersey. Brain samples taken from 197 mute swan were tested Brain tissue from these geese was extracted and tested by by RT-PCR by Guo et al. and ABV sequences were RT-PCR. Of these, 11 (44%) were RT-PCR-positive (Payne detected in 45 (23%) (Guo et al., 2012). Two mute swan et al., 2011a, b). Seven of the positive samples were from isolates were cultured, their M, N and X–P sequences 152 S. L. Payne et al. were analyzed, and they were found to closely match the large numbers. ABV was detected in 7 of 84 redheads Canada goose genotype. Positive mute swan samples (8.3%) tested. were obtained from Michigan, Rhode Island and New Jersey (Fig. 2). Further analysis of brain samples obtained from selected counties in Michigan suggested that ABV Domestic ducks prevalence could be as high as 50% in some populations. The ‘health’ of these mute swans was assessed at the While growing primary duck fibroblasts (DEF) from time of capture and five were determined to be ‘sick’. commercially available fertile Pekin duck eggs, Payne However, at the time, their ill health was believed to be a et al. (2012) found that these cells were infected with a result of either parasitic worms or of lead poisoning. No strain of ABV whose M-protein gene sequence was necropsies were performed on any of these birds and it is different from those of previous laboratory isolates. entirely possible that some may have suffered from Eighteen fertile eggs were subsequently purchased from subclinical nervous system disease. Of the states where each of four commercial duck hatcheries. Upon receipt, mute swans were tested, only New York provided no eggs were incubated for 8–14 days before the embryo was positive samples (Guo et al., 2012). ABV-associated removed and processed for DEF production. The cultured neuropathology has also been described in one DEFs were subsequently tested by RT-PCR for ABV-M mute swan in the retrospective study by Delnatte et al. protein. Two eggs from two of the four sources were (2012a). ABV-positive. Sequencing of these PCR products indi- It is of interest to note the great difference between cated that the viruses belonged to the ABV-CG genotype. apparent viral prevalence in mute swan cloacal swabs Minor differences in sequence from ABV genotypes (6%) and brains (23%). This disparity is similar to that previously cultured in the laboratory supported the observed in ABV-infected parrots and almost certainly premise that these viruses were not laboratory contami- reflects the intermittent nature of viral shedding. The nants. This result also suggests that ABV may be vertically presence of ribonucleases in the feces may also contribute transmitted in domestic ducks. This has implications for to this disparity. In addition, Delnatte et al. (2012b) transmission, diagnosis and immunopathology. Thus collected cloacal swabs from 67 mute swans and 132 free birds infected in-ovo may be immunologically tolerant ranging trumpeter swans, and found six positive mute of ABV. If, as is believed, disease develops as a result of swan samples (qRT-PCR analysis for ABV M-gene) T cell responses to the virus, this tolerance may ensure compared with no positive samples in trumpeter swans. that ABV infection is inapparent in many birds. Whether This was despite the fact that these species commingle in-ovo infection occurs in Canada geese is currently under and that ABV-diseased trumpeter swans had previously investigation by Delnatte and colleagues. been identified in the sampled flock. The reason for this difference is unclear, but a species effect on fecal shedding of ABV is one possibility. Gulls

Wild ducks Brains from several species of gull were tested for the presence of ABV M-gene RNA by RT-PCR (Payne et al., Given the high prevalence of ABV in Canada geese and 2012). Birds had been culled as a result of collision mute swans, it was logical to test for the presence of virus avoidance strategies at airports (New Jersey, New York in ducks. Thus 212 fresh duck heads were obtained from and New Hampshire) or euthanized after submission to processors of hunter-killed ducks in Texas and brain rehabilitation centers (Texas). Three of 26 herring gulls samples were extracted (Payne et al., 2012). These (Larus argentatus), 1/5 ring-billed gulls (L. delawarensis), samples reflected the species composition in the areas 3/13 laughing gulls (L. atricilla) and 0/4 great black- hunted. One set of samples was harvested near Houston, backed gulls (L. marinus) were RT-PCR-positive; none of Texas and consisted of a mixture of the predominant these birds were from Texas. wintering dabbling ducks. Most were northern pintails (Anas acuta) and gadwalls (Anas strepera). Other species tested included northern shovelers (Anas clypeata), Raptors mallards (Anas platyrhynchos) and American widgeons (Anas americana). ABV was detected by RT-PCR in 4 of A bald eagle (Haliaeetus leucocephalus) that was unable 61 northern pintails (6.5%), 6 of 25 gadwalls (24%), 1 of 10 to fly was submitted to an avian rehabilitator in East Texas mallards (10%) and 4 of 12 American widgeons (33%). It in February, 2011 and died soon thereafter. Necropsy was not detected in either of the two wood ducks (Aix showed the presence of acute encephalitis and ABV-CG sponsa) tested. Diving ducks, represented by redheads strain was present in the brain as demonstrated by (Aythya americana), were obtained from a hunting camp RT-PCR. Bald eagles are known to predate on waterfowl on the central Texas coast, where this species winters in flocks in Texas, killing and eating sick birds. Birds and bornaviruses 153

Thus transmission of ABV from geese to eagles is not persistently infected but the clinical disease patterns that unexpected. developed varied among individuals. Five birds developed clinical signs of PDD, while on necropsy 7 of the 18 had a dilated proventriculus. All infected birds did, Experimental infection with ABV however, show mononuclear cell infiltrates characteristic of PDD in a wide range of organs. ABV-induced disease in psittacines Mirhosseini et al. (2011) isolated ABV genotype 2 from a cockatiel and used infected DEF to inoculate two adult The causal association between experimental ABV infec- cockatiels by the oral and intramuscular routes. One bird tion of psittacine birds and the development of PDD has developed clinical signs on day 33 and the second on day been demonstrated by multiple investigators. Gancz et al. 41. While both challenged birds had slightly enlarged (2009) were the first to demonstrate that PDD could be proventriculi, histopathology showed typical PDD transmitted to healthy birds by the use of infected-brain lesions in the brain, spinal cord, heart, adrenal gland tissue. They inoculated cockatiels by multiple routes with and intestine. A control, uninoculated cockatiel was a brain homogenate from either an ABV4-positive bird or apparently healthy when euthanized on day 50 and at from a PDD-/ABV-control bird. The birds inoculated with necropsy, no gross abnormalities were observed. On healthy control bird homogenate remained healthy, histopathologic examination, the liver, pancreas and whereas all three birds inoculated with brain homogenate spleen had mild-to-moderate infiltration of lymphocytes, from ABV-infected birds developed both gross and some of which were forming lymphoid nodules microscopic lesions typical of PDD. Two of these had (Mirhosseini et al., 2011). However, all tissues from this also exhibited clinical signs. These investigators went on bird were negative for ABV by RT-PCR. Lierz and his to demonstrate the presence of ABV, with a sequence colleagues have subsequently conducted a similar chal- nearly identical to that of the challenge strain, in the lenge study with ABV2 and suggested that this genotype brains of the challenged birds. High throughput pyrose- is more pathogenic in cockatiels than is genotype 4 (Lierz quencing of the inoculum suggested however, that other et al., 2012). viruses may have been present in the inoculums although The results of all these experiments provide over- they were not identified in the challenged birds. While whelming support for the proposition that ABV is the sole persuasive, these results could not prove that ABV alone cause of PDD in psittacines. That is not to say that PDD was responsible for the development of PDD. inevitably develops in ABV-infected parrots. As with both In a formal attempt to prove Koch’s postulates, Gray BDV and the Canada goose strains of ABV, it is also clear et al. (2010) isolated ABV in cultured DEF. After six that many, apparently healthy psittacines may carry ABV passages, these infected cells were injected intramuscu- for prolonged periods. larly into two Patagonian conures (Cyanoliseus patago- nis). Clinical signs of PDD developed within 66 days post-infection in both challenged birds. The presence of ABV infection of ducks typical PDD was demonstrated on necropsy and histopathology. RT-PCR demonstrated the presence of the Gray et al. (2009) inoculated specific-pathogen-free inoculated strain in the brains of the challenged birds. A domestic mallard ducklings with ABV4 (psittacine origin) third, uninoculated control bird remained healthy. These cultured in DEF. Although virus could be identified in the conures, although apparently healthy, had previously brains of two infected birds and most seroconverted, been shown to be carriers of psittacine herpesvirus and as clinical disease and pathologic lesions did not result. a result, some uncertainty persisted regarding the etiology Delnatte and colleagues inoculated domestic Embden of PDD. geese intramuscularly with brain homogenates from ABV- Four apparently healthy cockatiels from a flock known infected Canada geese (P. Delnatte, unpublished work). to be shedding ABV4 were challenged with a known Inoculated birds showed mild non-specific clinical signs, virulent strain of ABV4 (strain M24) (Payne et al., 2011b). and mild-to-moderate non-suppurative encephalitis was The challenged birds either died or were euthanized for present in 4/13 experimental birds. ABV was identified by humane reasons between days 92 and 110. Typical PDD qRT-PCR in the brain of one of these birds; further was apparent on necropsy but the histopathologic lesions RT-PCR testing on tissue samples, characterization of the were reported to be unusually severe. Control birds fecal shedding of ABV and the development of any inoculated with uninfected tissue culture cells remained serologic response are ongoing. healthy until euthanized on day 150, and no histopathologic lesions of PDD were found at necropsy. Recently, Piepenbring et al. (2012) inoculated 18 ABV in North America cockatiels by both the intracerebral and intravenous routes with an isolate of ABV4 cultured for six passages Within North America, ABV are associated with two very in a quail cell line (CEC-32). All challenged birds became different patterns of disease. In captive psittacine birds, 154 S. L. Payne et al. infection is often confined to particular flocks and is cause of a geographically restricted disease of mammalian considered to have a high morbidity and mortality hosts, has been joined by a number of avian bornavirus although, as described earlier, subclinical or chronically genotypes. Bornaviruses infecting psittacine birds show infected birds have been identified and are likely under genetic diversity and proventricular disease of parrots fits recognized. In wild birds, particularly waterfowl, infec- the epidemiologic profile of an emerging disease. tion appears to be present in populations across Bornaviruses infecting waterfowl, in contrast, show a enormous geographic ranges, sometimes at very high remarkable lack of genotypic diversity and appear to be prevalence, and frequently associated with a lack of stable and endemic in North America, although they are observable clinical disease. also capable of producing significant pathology and While the initial identification of ABV in free-ranging mortality. Indeed, based on current information, wild waterfowl was surprising, the broad distribution of the waterfowl appear to constitute the largest pool of virus is not. North American waterfowl often occur in bornaviruses. Given the migratory nature of these birds, large flocks that can migrate long distances, and mix it is unlikely that waterfowl bornaviruses are restricted to together and separate into subgroups through the course a single continent. We anticipate that they will eventually of a year. The waterfowl are widely distributed and some be found among Central/South American and Eurasian species, for example, Canada geese and mute swans waterfowl as well. We suggest, therefore, that the occur in both migratory and resident populations. Close Bornaviridae, like avian influenza, are predominantly mixing of waterfowl species occurs frequently, particu- waterfowl viruses. However, based on their possible larly in areas with limited appropriate habitat, enhancing ‘jump’ into parrots, their identification in canaries, and the transmission of the virus among, as well as within, sporadic reports of ABV identification and PDD-like species. disease in a wide range of avian species, we have likely Density-dependent transmission of disease in water- only started to recognize the broad range of avian species fowl has been shown for a number of different avian that may be infected, and affected by this virus. pathogens and ABV infection is likely similarly influ- Many questions remain to be answered. For example, enced. The predominant means of spread of ABV is likely our understanding of the pathogenesis of ABV-related fecal-oral, and waterfowl graze and defecate both on land disease is limited. Much has been assumed on the basis of and in water. Viral infection probably occurs through rodent studies using BDV. On a deeper level, what is the ingestion, but intranasal, inhalation and even trans-cloacal relationship among the avian bornaviruses, and what is routes of spread are all possible. Viral survival and spread the origin of the psittacine ABV genotypes? EBLN are may be enhanced by the propensity of waterfowl to present in the genome of a variety of mammals, especially spend time in moist areas, and by the abundant, large, African ones, implying a very ancient origin and a much wet droppings with which they heavily contaminate their wider distribution of the Bornaviridae in the past. The environment. Persistent high bird density is seen with presence of EBLN in ground squirrels implies that ABV is some species, such as Canada geese, which remain in not new to North America and that a mammalian reservoir geographically restricted areas while raising their young may remain undetected in this region. There are no each year, and in locations that are particularly attractive. significant EBLN in the genomes of chickens, zebra finch Based on studies in parrots that show persistent infection and scarlet macaw, implying that, at least in the past, and shedding (in our aviary we have detected intermittent bornaviruses were predominantly found in mammals. ABV shedding by healthy birds for at least 5 years), The topic of birds and bornaviruses is one that will seemingly healthy waterfowl could transmit virus for long continue to occupy researchers for decades to come. periods of time and migrating birds could transmit ABV over long distances. The presence of virus in the brains and feces of large numbers of apparently healthy birds suggests that subclinical disease, or even more benign References forms of viral carriage, may be the normal state. The conditions that result in the development of the impress- Ackermann A, Guelzow T, Staeheli P, Schneider U and Heimrich ive pathologic lesions described by Delnatte et al. (2012a) B (2010). Visualizing viral dissemination in the mouse are yet to be elucidated. As pointed out above, vertical nervous system using a green fluorescent protein- transmission may also occur and this may play a role in expressing Borna disease viral vector. Journal of Virology 84: 5438–5442. determining the outcome of ABV infections. Asash E, Malkinson M, Meir R, Perl S and Weisman Y (1996). Causes of losses including a Borna disease paralytic syndrome affecting young ostriches of one breeding Conclusions organization over a five-year period (1989–1993). Avian Diseases 40: 240–245. Belyi VA, Levine AJ and Skalka AM (2010). Unexpected Since 2008 our knowledge of the family Bornaviridae has inheritance: multiple integrations of ancient bornavirus and undergone a remarkable transformation. Borna disease ebolavirus/marburgvirus sequences in vertebrate genomes. virus, the sole member of the family and known as the PLoS Pathogens 6: e1001030. Birds and bornaviruses 155

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APPENDIX B. SAMPLING PROTOCOL FOR TRUMPETER AND MUTE SWANS PROVIDED TO THE BIOLOGISTS AND VOLUNTEERS OF THE TRUMPETER SWAN REINTRODUCTION PROGRAM

Purpose of the project: Survey wild waterfowl for infection by avian bornavirus (ABV)

Sequence of events: - “Sampling kits” will be sent to you. - Blood sample and cloacal swabs are collected in the field. - The samples are then delivered to a local veterinary clinic as soon as possible in order to centrifuge blood and to freeze the swabs. - The samples are finally sent to OVC Guelph to detect antibodies in serum and ABV genome in - cloacal swab.

Equipment list for sample collection: - Sampling kits (labelled Ziploc bag with one syringe, one needle, one tube, one swab, one blue cap viral transport media tube) - Cotton balls soaked in alcohol - Large Ziploc bags - Permanent markers - One shipping cooler and ice packs - Biohazard bag for garbage - Sharps container - Extra syringes, needles, tubes, labelled Ziploc bags, swabs, viral transport media tubes

Sampling kits: To make easier the sampling process in the field, one kit will be one Ziploc bag and should be used for one bird. One bird = One kit =One Ziploc bag

Each “Ziploc bag” we will send to you will contain: - 1 printed label (already stuck on the bag) - 1 syringe + 1 needle - 1 SST gel separator tube (yellow cap) - 1 cloacal swab - 1 blue cap viral transport media tube

Procedure (for each bird): - Take a sampling kit (everything you need for sampling should be inside). - Label the bag with the bird’s identification, minimally wing tag number and date. - Label each tube (1 yellow cap and 1 blue cap with transport media) with the wing tag number and date (keep tubes closed and open only before collecting bird). - BLOOD SAMPLING:

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- Blood is collected from the medial metatarsal vein (preferred site in waterfowl).The vein is located on the medial (inner) side of the lower leg below the hock. - Wet the venipuncture area lightly with alcohol to better observe the vein and to clean the skin. - Collect blood using a 22 G needle and a 3 mL syringe. The minimum quantity required is 2 mL (but 3 mL would be better and still safe for the bird). - Apply digital pressure until bleeding stops (30-60 seconds). - Immediately transfer blood slowly from the syringe to the pre-labelled SST gel separator tube directly through the rubber cap (no need to remove the cap). - Put this tube (yellow cap) back in the Ziploc bag.

- CLOACAL SWAB: - Open the swab envelope on the “stick” end, remove the swab. - With one hand, gently open the cloacal orifice, and with the other hand, gently insert the swab into the cloaca of the bird (1-2 cm deep). - Use gentle pressure and in a circular motion, swab the inside of cloaca 2-4 times. - Shake off any large (>0.5 cm) pieces of feces. - Open the tube containing viral transport media (blue cap). - Place the swab tip in the transport media approximately ¾ of the way towards the bottom of the tube. Snap the stem of the swab against the top of the tube so that the entire swab end and a portion of the stem should be left in the tube (do not use scissors or other cutting instrument to break the swab shaft) and replace the cap. - Put this tube (blue cap) back in the same Ziploc bag. - Be sure to complete the printed labels (stamped on the bags) with bird’s ID (wing tag number and band number), sex and age (when available), date, time and location of sampling and person who collected samples. Ensure that labels are marked with pencil or permanent ink, which will not dissolve when they get wet or frozen.

Recheck: After sampling, each Ziploc bag should contain: - 1 fully-completed label (stamped on the bag) - 1 blood sample (in a labelled yellow cap tube) - 1cloacalswab in a labelled blue cap viral transport media tube - Place the Ziploc bag in the cooler with an ice-pack until delivery to the local veterinary clinic.

Shipping to the local veterinary clinic: Place all the Ziploc bags together in a larger Ziploc bag in the provided cooler with ice packs and deliver to the local veterinary clinic as soon as possible. If you are not able to deliver the samples to the clinic the same day, Ziploc bags must be stored in fridge (DON’T FREEZE) until submitted.

THANK YOU VERY MUCH FOR YOUR TIME If you have any questions, please feel free to contact Pauline Delnatte (226-820-0419, [email protected]) or Dale Smith (519-824-4120 x54622, [email protected]) 174

APPENDIX C. AVIAN BORNAVIRUS NUCLEOCAPSID PROTEIN GENE CLONING, RECOMBINANT EXPRESSION, AND PURIFICATION – PROTOCOL

The expression cassette for the M24N protein carried by the pET21a plasmid was obtained with thanks from Dr. Susan Payne, Schubot Exotic Bird Health Centre, Texas A&M University. The plasmid was isolated using the Life Technologies High Pure DNA miniprep kit (Life Technologies, Carlsbad, California, USA). It was then transformed into BL21 (DE3) pLysS to be expressed under the T7 polymerase promoter (Novagen pET system manual, 10th Ed., 2003). A volume of LB media was inoculated from an overnight culture and grown to OD600 of 0.600. Expression of the N protein was induced by addition of IPTG to 1 mM final concentration and the culture was grown for an additional 4 hours. The E.coli cells containing the expressed protein were pelleted and washed once with cold 20 mM Tris-HCl pH 8.0. The pelleted cells were resuspended in a buffer compatible with the Qiagen Ni-NTA purification system (Qiagen, Dusseldorf, Germany) and lysed with the addition of lysozyme, PMSF and Benzonase endonuclease (EMD Millipore, Billerica, Massachusetts, USA). Buffers for the purification were based on the Protein Purification under Native conditions protocol found in the Qiagen Ni-NTA Spin Kit Handbook (2008). The cell lysate was spun as described in the handbook. The soluble fraction was collected the 6x his-tagged N protein was purified from this fraction. Three milliliters of Qiagen Ni-NTA agarose beads were dispensed into a 10 cm column (Bio-Rad, Hercules, California, USA) and washed with several column volumes of NPI buffer containing 10 mM imidazole. The soluble lysate was applied to the column and allowed to run through. The column was washed twice with two column volumes of buffer containing first 20 mM imidazole and then 50 mM imidazole. The 6x his-tagged protein was eluted from the column using first one column volume of buffer containing 250 mM imidazole and a second column volume with buffer containing 500 mM imidazole. The purified protein was analyzed by gel electrophoresis (SDS-PAGE) and the concentration was determined by the Bio-Rad Protein Assay.

For the SDS-PAGE, 50 µl of each fraction was collected at each step of the purification, mixed with 10 µl of loading buffer and heated at 95oC for 10 minutes. 10 µl of each 175

treated sample was loaded onto a 10% SDS-PAGE gel and run at 100 V until the dye front approached the bottom of the gel. The acrylamide gel was treated with Coomassie Brilliant Blue dye to stain the separated protein bands and an image was taken using a Bio-Rad Chemi-Doc XRS (marker is Bio-Rad # 161-0374 Precision Plus Protein™ Dual Color Standards).

Coomassie blue stained polyacrylamide gel showing the purified ABV M24N protein (arrow)

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APPENDIX D. AVIAN BORNAVIRUS INFECTION TRIAL - GROUP ASSESSMENT SHEET

*Shaded areas are for additional observations per day if requested by researchers based on changes in scoring as noted below.* Usual monitoring: Observations by researcher or student will be performed twice daily in the absence of any clinical signs. Findings/scorings are recorded on a "per room" basis. Enhanced monitoring: If any bird(s) show scoring greater than 0, researchers will be contacted to assess subject(s) and determine whether an increase in daily observations is appropriate / necessary. Scoring Key: 1Drinking – 0 – normal; 1 – reduced; 2Feeding – 0 – normal; 1 – reduced; 3Breathing – 0 – normal; 1 – laboured; 4Activity – 0 – normal; 1 – reduced; 2 – recumbent; 5Feces – 0 – normal; 1 – diarrhea or undigested food in feces 6Neurologic – 0 – normal coordination and behaviour; 1 – mild ataxia or abnormal head position; 2 – moderate ataxia or abnormal head position; 3 – severe (unable to stand or ambulate)

NOTE: It is not known whether birds will exhibit any clinical signs of disease following experimental infection. ENDPOINT DECISION: If any bird(s) show an Activity score of 2 or a Neurologic score of 2 or 3 a researcher listed below should be contacted to determine whether a further period of observation, or euthanasia and necropsy should be performed. EMERGENCY CONTACTS: Dr. Dale Smith ([email protected]) (Ext: 54622) Dr. Éva Nagy ([email protected]) (Ext 54783)

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Room # Infected Embden geese Week # Date of hatch: XX Date of inoculation: XX Drinking1 Feeding2 Breathing Activity4 Feces5 Neurologi Comments Initials 3 c6 Morning *Noon End of Day *Night Drinking1 Feeding2 Breathing3 Activity4 Feces5 Neurologic Comments Initials 5 Morning *Noon End of Day *Night Drinking1 Feeding2 Breathing3 Activity4 Feces5 Neurologic Comments Initials 5 Morning *Noon End of Day *Night Drinking1 Feeding2 Breathing3 Activity4 Feces5 Neurologic Comments Initials 5 Morning *Noon End of Day *Night

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