AUISCHHOLZHAUSEN ERMANY · Turkey coronavirus – molecular biology and pathogenicity of a 225...

World Veterinary Poultry Association and Clinic for Birds, Reptiles, Amphibians and Fish Justus Liebig University Giessen, Germany

VI. IN T E R N A T I O N A L SY M P O S I U M O N A V I A N CO R O N A - A N D PN E U M O V I R U S E S A N D

CO M P L I C A T I N G PA T H O G E N S

RA U I S C H H O L Z H A U S E N , GE R M A N Y , 14-17 JU N E 2009

PROCEEDINGS

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EDITORS DR. URSULA HEFFELS-REDMANN DAGMAR SOMMER PROF. DR. ERHARDT F. KALETA Clinic for Birds, Reptiles, Amphibians and Fish Justus Liebig University Frankfurter Strasse 91-93 D-35392 Giessen, Germany Phone: +49(0)641 99 384 52 Fax: +49(0)641 99 384 39 Email: [email protected] PUBLISHERS VVB LAUFERSWEILER VERLAG édition scientifique Staufenbergring 15 D-35396 Giessen, Germany Phone: +49(0)641 55 99 888 Fax: +49(0)641 55 99 890 Email: [email protected] ISBN / EAN – BOOK and CD ISBN-10: 3-8359-5484-9 ISBN-13: 978-3-8359-5484-7 EAN code: 9783835954847 © 2009 by VVB LAUFERSWEILER VERLAG, Giessen Printed in Germany

mailto:[email protected]


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SPONSORS The organizers gratefully acknowledge the financial support of: INTERVET INTERNATIONAL, BOXMEER, THE NETHERLANDS

MERIAL, LYON, FRANCE

FORT DODGE VETERINÄR, WÜRSELEN, GERMANY

LOHMANN TIERZUCHT, CUXHAVEN, GERMANY

IDEXX, LUDWIGSBURG, GERMANY

LOHMANN ANIMAL HEALTH, CUXHAVEN, GERMANY

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CONTENTS

Page PREFACE 1 AVIAN CORONAVIRUS – IBV – EPIDEMIOLOGY Observations on global and local epidemiology of avian coronaviruses 2 JONES RC, SAVAGE CE, WORTHINGTON KJ and HUGHES LA Sequence analysis of IBV from outbreaks in Denmark 2006-2009 7 HANDBERG KJ, KABELL S, OLESEN L and JØRGENSEN PH Molecular epidemiology of slovene infectious bronchitis virus strains 13 isolated between 1990 and 2006 KRAPEŽ U, SLAVEC B, BARLIČ-MAGANJA D and ZORMAN ROJS O

First report of IBV QX-like strains in Spain 27 DOLZ R, BERTRAN K and MAJÓ N

Molecular survey of IBV in Europe in 2008 34 MONNE I, DRAGO A, FASOLATO M, CAPUA I and CATTOLI G False layers: IB case report in broiler breeders in Germany 38 BLOCK H IBV in Jordan: Molecular subtype 40 ALROUSSAN DA, TOTANJI WS and KHAWALDEH GY

Recombination, point mutations and positive selection on the basis of the 47 molecular diversity of Brazilian strains of avian IBV BRANDAO PE, SANDRI TL, SOUZA SP, KUANA SL, RICHTZENHAIN LJ and VILLARREAL LYB

Genotyping and serotyping of Guangxi IBV isolates during 1985~2008 59 WEI P, LI M, WEI ZJ, WANG XY, MO MJ and CHEN QY Avian infectious bronchitis viruses from slaughtered chickens exhibit 67 wide varieties CHEN HW, HUANG YP and WANG CH

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AVIAN CORONAVIRUS – IBV – VIRUS PROPERTIES AND VIRUS-HOST INTERACTIONS A polymorphism study on IBV genes S1 and 3 using RT-PCR and RFLP 78 MAJDANI R; ALIABAD FN, MARANDI MV, MORSHEDI A and MARDANI K IBV induces NF-KB signalling 87 MCCRORY SA, MACDONALD A and HISCOX JA

Genetic and phenotypic variation of IBV within the host 88 GALLARDO R, VAN SANTEN VL and TORO H

Significance of minor viral subpopulations within Ark-type IB vaccines 92 VAN SANTEN VL, NDEGWA EN, JOINER KS, TORO H and VAN GINKEL FW Are the structural and accessory genes of IBV responsible for pathogenesis? 96 ARMESTO M, BRITTON P and CAVANAGH D

Importance of sialic acid for the infection of the tracheal epithelium by 100 different strains of IBV ABD EL RAHMAN S, NEUMANN U, HERRLER G and WINTER C

High throughput proteome screening of IBV infected cells reveals novel 108 host-cell interactions EMMOTT E and HISCOX JA

AVIAN CORONAVIRUS – IBV – DIAGNOSIS The use of FTA® cards to transport samples for diagnosis of IBV 109 and Avian Metapneumovirus by RT-PCR SAVAGE CE, COWLEY K and JONES RC Molecular detection and typing of IBV 114 LÜSCHOW D, DE QUADROS VL and HAFEZ HM

Test for extraneous agents in avian inactivated vaccines using PCR: 118 Detection of IBV MOTITSCHKE A and JUNGBÄCK C

The avian coronavirus IBV as a model for an in-house virus detection 125 microarray ABU-MEDIAN AA and BRITTON P

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Rapid detection and objective characterisation of IBV isolates 130 using high resolution melt curve analysis and a mathematical model HEWSON KA, NOORMOHAMMADI AH, DEVLIN JM, MARDANI K and IGNJATOVIC J

AVIAN CORONAVIRUS – IBV – PATHOGENESIS AND IMMUNOLOGY Pathogenic patterns in chicken challenged with variant strains of IBV 138 isolated from chicken flocks with different clinical manifestations CHACÓN JL, ASSAYAG MS, REVOLLEDO L, IVO MA, VEJARANO MP, PEDROSO AC and FERREIRA A

IBV induces acute interferon-gamma production through polyclonal 142 stimulation of chicken leukocytes JANSEN C, ARIAANS M, VAN HAARLEM D, VAN DE HAAR P, DE WIT JJ and VERVELDE L AVIAN CORONAVIRUS – IBV – VACCINATION The experimental production of bivalent IB/ND vaccine (793/B IBV 151 serotype / Clone 30 NDV) KHALESI B, MASOUDEI S and MOGHADDAMPOUR M Enhanced efficacy of the use of a monovalent Infectious Bronchitis Virus 158 inactivated vaccine in layers primed with H120 and 793B live IBV vaccines to increase the protection against challenge with 3 European serotypes of IBV DE WIT JJ, VAN DE SANDE H and PRANDINI F

Development of in ovo vaccine against IB in poultry flocks 167 KHALESI B, MASOUDEI S and MOGHADAMPOUR M Efficacy of combined vaccines at day of hatch against D388 challenge 176 in SPF and commercial chickens DE WIT JJ and VAN DE SANDE H

Immune responses in chicks after single or dual vaccination with live 183 IB Massachusetts (H120) and variant vaccines: some preliminary findings GANAPATHY K, ROTHWELL L, LEMIERE S, KAISER P and JONES RC

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Predictive value of the results of an α-IBV IgM ELISA for the efficacy 188 of IBV vaccinations in the field DE WIT JJ, VAN DER MEULEN A and FABRI THF

Immune response in chickens to recombinant S1 and N proteins of IBV 193 MEIR R, MAHARAT O, KRISPEL S and KATZ E

Manipulation of the IBV genome for vaccine development 198 BRITTON P, ARMESTO M, CASAIS R and CAVANAGH D AVIAN CORONAVIRUS – TURKEY CORONAVIRUS Molecular identification and characterization of a turkey coronavirus 209 in France MAUREL S, TOQUIN D, QUEGUINER M, LE MEN M, ALLEE C, LAMANDE J, BERTIN J, RAVILLION L, RETAUX C,.TURBLIN V, MORVAN H, PICAULT JP and ETERRADOSSI N Subgenomic RNA transcription of turkey coronavirus 219 CAO JZ, WU CC and LIN TL Comparative genomics on avian coronaviruses; Origin and divergence 220 associated with host and pathogenic shifts JACKWOOD MW, PATERSON AH, KISSINGER JC, HILT DA, MCCALL AW, MCKINLEY ET and BOYNTON TO

Turkey coronavirus – molecular biology and pathogenicity of a 225 growth-robbing pathogen GOMAA MH, YOO D, OJKIC D and BARTA JR

Pathology and virus tissue distribution of turkey coronavirus (TCOV) in 235 experimentally infected chicks and turkey poults GOMES DE, HIRATA KY, TEIXEIRA MCB, FERRARI HF, VICENTE RM, LUVIZOTTO MCR, GAMEIRO R and CARDOSO TC Matrix metalloproteinases expression in experimentally infected 245 poults with turkey coronavirus CARDOSO TC, GOMES DE, ASTOLPHI RD, NOVAIS JB, GUEDES ACR, FERRARI HF, SILVA-FRADE C and LUVIZOTTO MCR

DNA-mediated vaccination against challenge infection by turkey 250 coronavirus LIN TL, ABABNEH M, HSIEH MK, CHEN YN and WU CC

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AVIAN METAPNEUMOVIRUS – EPIDEMIOLOGY Subtypes of AMPV circulating in Brazilian commercial flocks 251 CHACÓN JL, PEDROSO AC, TOQUIN D, ETERRADOSSI N, PATNAYAK D, GOYAL S and FERREIRA AJP

Field observations after natural infection of Brazilian layer chickens 255 with a phylogenetically divergent lineage of subtype B AMPV VILLARREAL LYB, SANDRI TL, ASSAYAG MS, RICHTZENHAIN LJ, MALO A and BRANDÃO PE

Turkey rhinotracheitis outbreak in 7 week old turkeys caused by a 260 vaccine derived AMPV RICCHIZZI E, CATELLI E, CECCHINATO M, LUPINI C, BROWN P and NAYLOR CJ

Human metapneumovirus infection in turkeys 265 NAGARAJA KV, VELAYUDHAN BT, HALVORSON, DA and GRAY GC

AVIAN METAPNEUMOVIRUS – VIRUS PROPERTIES AND DIAGNOSIS Identification of two regions within the subtype A AMPV fusion protein 269 (amino acids 211 – 310 and 336 – 479) recognized by neutralizing antibodies BROWN PA, BONCI M, RICCHIZZI E, JONES RC and NAYLOR CJ

Avian Metapneumoviruses in Italy: Evidence of attachment protein 278 evolution coincident with mass live vaccine introduction CECCHINATO M, CATELLI E, LUPINI C, RICCHIZZI E, CLUBBE J and NAYLOR CJ

An investigation into molecular differences between Avian 285 Metapneumoviruses (AMPVS) of chicken and turkey origin CLUBBE J, JONES RC, GALLUDEC HL and NAYLOR CJ

Use of reverse genetics to develop a positive control virus for RT 294 nested PCR detection of subtype A and B AMPV FALCHIERI M, BROWN PA, CATELLI E and NAYLOR CJ

The effect of SH gene modifications on cytopathic effects seen in Vero cells 299 BROWN PA and NAYLOR CJ

Construction of GFP AMPV recombinant lacking the small hydrophobic 304 protein gene LUPINI C, CATELLI E, CECCHINATO M and NAYLOR CJ

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Growth in vitro and in vivo of a recombinant AMPV expressing eGFP 307 EDWORTHY N, TOQUIN D, ZWINGELSTEIN F, ETERRADOSSI N and EASTON AJ

Generation and biological assessment of recombinant avian 315 metapneumovirus subgroup C (AMPV-C) viruses containing different length of the G gene YU Q, ESTEVEZ C, KAPCZYNSKI D and ZSAK L AVIAN METAPNEUMOVIRUS – VACCINATION AND IMMUNITY AMPV – nearly 30 years of vaccination 326 COOK JKA

Field AMPV evolution avoiding vaccine induced immunity 334 CATELLI E, CECCHINATO M, LUPINI C, RICCHIZZI E and NAYLOR CJ

Low AMPV vaccine performance due to turkey astrovirus (TAstV-2) 338 persistence infection: Field study, Brazil CARDOSO TC, FERREIRA HL, DA SILVA SEL, FERRARI HF, TEIXEIRA MCB and LUVIZOTTO MCR

The role of humoral and cell-mediated immunity in the control of 344 avian metapneumovirus infection in turkeys RAUTENSCHLEIN S and RUBBENSTROTH D

Activity and efficacy of an experimental inactivated oil-adjuvanted 355 AMPV-C antigen preparation in white Pekin and specific pathogen free Muscovy ducklings TOQUIN D, ALLÉE C, LE BRAS MO, AMELOT M and ETERRADOSSI N COMPLICATING PATHOGENS Peritonitis due to Ornithobacterium rhinotracheale co-infection with IBV 363 and Escherichia coli in laying chickens THACHIL AJ, VELAYUDHAN B, SHAW DP, HALVORSON DA and NAGARAJA KV Role of unusual mycoplasmas in poultry – Mycoplasma lipofaciens as 371 an example LIERZ M

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IBD efficacy data in presence of maternal antibodies in broilers and 376 layers with a novel vector HVT-IBD vaccine LE-GROS FX

Novel HVT/ILTV-based recombinant vaccine to simultaneously 385 control infectious laryngotracheitis and Marek’s Disease in chickens (INNOVAX®-ILT) HEIN R SUMMARIES OF THE SESSIONS AND PANEL DISCUSSION 387 ADDRESS LIST OF THE PARTICIPANTS 391 AUTHOR INDEX 398

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VI. INT. SYMPOSIUM ON AVIAN CORONA- AND PNEUMOVIRUSES, RAUISCHHOLZHAUSEN, GERMANY, 2009

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PREFACE

In 1988 Prof. Kaleta realized the idea for a joint meeting of scientists and veterinarians from diagnostic laboratories, from the field and from vaccine producing companies, all working on the avian coronavirus causing infectious bronchitis which was and still is one of the major problems in poultry industry. He organized the I. International Sympsoim on Infectious Bronchitis in the Castle of Rauischholzhausen, the conference centre of the Justus Liebig University Giessen, which is located in a little village between the cities of Giessen and Marburg in Germany. The programme comprised all aspects of this virus disease, and the beautiful castle and its parc created a pleasant atmosphere for intense exchange of information and discussion. During the three days of the symposium at this separated place the attendees from all over the world came closer together not only with regard to social but also to scientific aspects. This had the positive effect of the formation of new working groups. Since then, eleven symposia on special avian virus infections had been organized. This year already the VI. Symposium on Avian Corona- and also Pneumoviruses took place in the Castle with the for non-Germans so difficult to pronouncing name indicating that the idea of Prof. Kaleta has become a known event. But unfortunately this symposium was the last one under his superintendence. For this reason, Prof. Hafez as President of the WVPA took the chance to recognize Prof. Kaleta’s long lasting efforts for the promotion of exchange of information and knowledge on avian medicine. Dr. Ursula Heffels-Redmann as his assistant and co-organizer since the beginning in 1988 thanked him in the name of all staff members from the Clinic for Birds, Reptiles, Amphibians and Fish, Justus Liebig University Giessen for the good and trustful cooperation. To make his retirement easier Prof. Lierz as his successor as head of the clinic promised to continue the work he started and expressed the hope that he furthermore can account on Prof. Kaleta`s support in this intension.

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OBSERVATIONS ON GLOBAL AND LOCAL EPIDEMIOLOGY OF AVIAN CORONAVIRUSES

JONES RC, SAVAGE CE, WORTHINGTON KJ and HUGHES LA

Department of Veterinary Pathology University of Liverpool, Leahurst Campus

Neston, South Wirral, CH662SY SUMMARY The global distribution of different serotypes/genotypes of infectious bronchitis virus (IBV) is discussed in the light of our relative ignorance as to how the viruses are transmitted between regions. Molecular techniques have enabled tracking of similar or identical viruses in different parts of the world, although the means of spread is seldom understood. In the light of their perceived importance in the spread of avian influenza viruses, it is tempting to suggest that they might also transmit IBV. Some examples of recent detections of avian coronaviruses in wild species are given but to date there is little evidence that they are of importance in IBV transmission. INTRODUCTION Avian coronaviruses are the cause of infectious bronchitis (IB), one of the most economically important endemic diseases in chickens worldwide and turkey and pheasant coronavirus infection. In recent years we have been investigating the prevalence of IBV genotypes in Western Europe and Latin America and also the presence of coronaviruses in wild birds. This paper discusses the distribution of different IBV genotypes globally and draws attention the lack of knowledge relating to virus spread and the occasional detection of avian coronaviruses in wild birds. IBV GENOTYPES: GLOBAL DISTRIBUTION Many genotypes are recognised worldwide. They were originally classified according to cross-neutralisation tests in eggs or tracheal organ cultures but more recently, genotyping has been used, based on the nucleotide sequencing of the S1 spike gene,


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which in most cases, correlates well with serotyping. Genotypes differ remarkably in their distribution worldwide. Massachusetts IBV for example is worldwide, in part due to the universal use of Mass-type vaccines. Arkansas and Connecticut are sometimes found outside the USA. The 793B group is present in many countries but not in the USA. Asia, Africa and Latin America have their own indigenous types and some established ‘international’ ones too. Australia, with its very strict import controls has only indigenous types. Some viruses, like the nephropathogenic B1640 appear to have spread very little from the country of origin. While the latter may relate to a different mode of pathogenesis, the ways in which different IBVs spread across continents is poorly understood. IBV WESTERN EUROPEAN SURVEY In a recent RT-PCR survey of IBVs in certain Western European countries between 2002-2006, we showed that 793B and Mass types were predominant, about 50% of each being considered vaccine virus (Worthington et al., 2008). Two novel genotypes appeared, Italy 02, of unknown origin and QX, identical with a virus first described in China in 1998. The appearance of the latter in Europe from its far Eastern origin resembled the transition of highly pathogenic H54N1 avian influenza from South East Asia to Europe and in a not dissimilar time span. However, in the case of H5N1, though wild migratory waterfowl are generally considered to have been instrumental in spread, there is no evidence to date that wild species promote the movement of important IBVs. A report by Bochkov et al. (2006) on strain in Russia, described QX detection in 2001 very close to the China border, then one year later in Volgograd, more than 6000 km away. It is possible that virus could have been transported perhaps by humans carrying livestock or meat products on the Trans-Siberian railway or by air travel. RECENT DIAGNOSTICS: IBVS IN LATIN AMERICA Since that study, we have been examining samples from Latin America. Apart from Massachusetts and 793 genotypes, among those encountered in South America was a QX-like virus resembling one we originally detected in France and a Chinese type which was not QX. How these viruses traversed the Pacific remains unknown. In some Latin American countries, are H120 vaccine is the only one licensed against IBV, when it is well recognised that other potentially important variants are prevalent. Control of IB in these cases represents a real enigma. GLOBAL SPREAD OF IBV: CONCLUSIONS Modern molecular techniques allow the precise fingerprinting of IBVs so molecular epidemiology can be used to trace the origin of novel viruses in new regions. However, even when the disease/new virus is recognised early and full investigations are undertaken, it is rarely possible to determine how the virus enters the country. With notifiable avian influenza virus (AI), there is limited capacity of some countries to


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investigate disease, there are delays in reporting outbreak s when they first occur and there is a scarcity of information on illegal movements of poultry or poultry products. The involvement of migratory wild birds in the transmission of highly pathogenic AI has prompted the search for IBV-like coronaviruses in wild avian species with the possibility of explaining intercontinental movement of these viruses. CORONAVIRUSES IN WILD BIRDS Originally, searches for coronaviruses in wild birds were hampered by their fastidious growth requirements and few viruses could be cultivated in chicken eggs. However, more recently, RT-PCR technology has been used widely to detect a diversity of coronavirus in several wild species. The use of microarrays will in future provide a further useful tool. In a study in Norway, type 3 coronaviruses were detected by Jonassen et al. (2005) in teal, mallard, greylag goose, wigeon and pigeon. Comparison of nucleotide sequences in the replicase gene showed that all were avian viruses but unrelated to IBV. Gough et al. (2006) described the first coronavirus from a psittacine species, an Amazon green-cheeked parrot. Other reports describe coronaviruses from the bulbul, thrush and munia (Woo et al., 2009) and mallard faecal pools (Muradrasoli et al., 2009). None of these appeared to be closely related to IBV. In contrast, two Chinese papers describe IBV like coronaviruses from a pigeon (Qian, 2006) and a peafowl (Sun et al., (2007). The pigeon isolate caused pancreatitis in chicks and was closely related to several IBV genotypes on the basis of S1 spike gene sequences. The peafowl virus was shown to be a Massachusetts type and was also pathogenic for chickens. In both these instances it appears that the viruses might have originated in poultry. THE LIVERPOOL STUDY The presence and prevalence of coronaviruses in wild bird populations in Northern England was investigated between 2004 and 2007 (Hughes et al., 2009). Faecal samples and sometimes oropharyngeal swabs were taken from more than 400 birds in 42 species, including both resident and migratory types. They were from live birds caught fro ringing and from dead wildfowl and corvids provided by local shooters. After extraction of the viral RNA, primers were used which targeted the 3’ untranslated region of the coronavirus genome. Coronavirus RNA was detected in seven faecal sample pools of which four were from ducks, one from a whooper swan, one from red knot and one from Eurasian oystercatchers. The pools represented estuarine, salt march or standing water habitats. All birds from which viruses were detected appeared to be healthy. Phylogenetic analyses were based on a final usable sequence of 146 nucleotides after removal of primer sites (Figure 1.). Sequences detected in three pooled duck samples and the sequence from the whooper swan clustered with a sequence from an IBV H120 (Massachusetts) vaccine. Sequences within this cluster were relatively homogeneous, with low within-group distance values (0.0%-2.8%) and bootstrap support for the individual nodes was relatively low. Coronavirus sequences in red knots clustered with


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a previously described goose coronavirus and divergence at the nucleotide divergence was 2.0%. The virus from oystercatchers clustered with a previously established duck coronavirus. Although some 42 species of wild birds from numerous were sampled, coronaviruses were detected only in wildfowl (Anseriformes) and waders (Chadriiformes). All the detections were from faecal samples, which were simpler to obtain than oropharyngeal ones. Although primarily a respiratory virus, many strains of IBV have been shown to replicate in the chicken intestine, usually without enteric disease. Further work is needed to compare the sequences of the S1 spike protein gene which is the commonest way to compare IBV genotypes. CONCLUSIONS Many different serotypes/genotypes of IBV have been described and due to the innate variability of the virus, will continue to be described. Molecular analysis and detailed sequencing allows the interrelationships of these viruses to be explored although in most instances, how viruses are transported between continents can be speculated on but is largely unknown. Drawing from the experiences of the role of wild birds in highly pathogenic avian influenza viruses, it is tempting to suggest that wild species could also have a role in dissemination of IBV genotypes. Although coronaviruses have been detected in several diverse species of wild birds and are likely to continue to be found in further investigations, to date there is little evidence that wild birds play an important part in long distance transmission of IBV. REFERENCES Bochkov YA, Batchenko GV, Shcherbakova LO, Borisov AV, Drygin VV. (2006).

Molecular epizootiology of avian infectious bronchitis in Russia. Avian Pathology. 35, 379-93.

Jonassen, CM, Kofstad, T, Larsen IL, Løvland, A, Handeland, K, Follestad, A, Lillehaug, (2005). A Molecular identification and characterization of novel coronaviruses infecting graylag geese (Anser anser), feral pigeons (Columbia livia) and mallards (Anas platyrhynchos). Journal of General Virology. 86, 1597-607.

Hughes, LA, Savage, CE, Naylor, CJ, Bennett, M, Chantrey, J and Jones, RC (2009). Genetically diverse coronaviruses in wild bird populations of Northern England. Emerging Infectious Diseases. 15, 1091-1094

Muradrasoli S, Mohamed N, Hornyák A, Fohlman J, Olsen B, Belák S, Blomberg J. (2009). Broadly targeted multiprobe QPCR for detection of coronaviruses: coronavirus is common among mallard ducks (Anas platyrhynchos). Journal of Virological Methods. 159, 277-87.

Qian DH, Zhu GJ, Wu LZ, Hua GX. (2006). Isolation and characterization of a coronavirus from pigeons with pancreatitis. American Journal of Veterinary Research. 67, 1575-9.


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Sun L, Zhang GH, Jiang JW, Fu JD, Ren T, Cao WS, Xin CA, Liao M, Liu WJ.A (2007) Massachusetts prototype like coronavirus isolated from wild peafowls is pathogenic to chickens. Virus Research. 130:121-8.

Worthington KJ, Currie, RE and Jones, RC (2008). A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. Avian Pathology, 37, 247-257.

Woo PC, Lau SK, Lam CS, Lai KK, Huang Y, Lee P, Luk GS, Dyrting KC, Chan KH, Yuen KY. (2009). Comparative analysis of complete genome sequences of three avian coronaviruses reveals a novel group 3c coronavirus. Journal of Virology. 83:908-17.

Figure 1. Minimum-evolution tree of avian coronaviruses based on a 146bp fragment of the 3’ UTR region of IBV. Coronaviruses detected in wild birds in the Northern England study (Hughes et al., 2009) are denoted with an asterisk. Genbank accession numbers are shown in brackets.


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SEQUENCE ANALYSIS OF IBV FROM OUTBREAKS IN DENMARK 2006-2009

HANDBERG KJ1a, KABELL S1b, OLESEN L2 and JØRGENSEN PH1c

1National Veterinary Institute, Technical University of Denmark, 2 Hangoevej, DK-8200, Aarhus N, Denmark. a [email protected], b [email protected], c [email protected].

2Danish Agricultural Advisory Service, National Centre, Udkaersvej 15, DK-8200 Aarhus N. [email protected].

SUMMARY IBV is still a significant problem for the poultry industry worldwide and new types continue to emerge in an arms race with the development of new vaccines. We reported in this study the detection of the IBV type QX D388 in Danish poultry and that this type has been the most prevalent type in the period 2006-2009. Typing was based on sequence analysis of a 260 bp fragment of the S1 gene. Further analysis of the sequences revealed silent variations, which potential could be used as epidemiological markers. INTRODUCTION Infectious bronchitis virus (IBV) is belongs to the family Coronaviridae, genus coronavirus, group III which are positive single stranded RNA viruses. The main impact of IBV is respiratory disease in chickens. In the majority of infections mortality is absent or moderate while the infection often causes production losses and compromised animal welfare due to secondary infections. Several vaccines against IBV have been developed and are widely used. Because of IBVs nature as a positive single-stranded RNA virus a high mutation and heterologues recombination rates of the viral genome occurs. By these mechanisms new IBV subtypes often emerge. Immunity against such new variants may be sub-optimal (Cavanagh. (2007)) In Denmark, IBV vaccines based on the Massachusetts (Mass) virus subtype have been used for decades. In the nineties the virulent variant strain 793B was introduced in the Danish poultry population. The negative impact of this strain was efficiently controlled by use of a homologues vaccine. The occurrence of IBV was managed satisfactorily until 2006, when the variant strain QX D388 was introduced in Danish commercial poultry.



mailto:[email protected].

mailto:[email protected].


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MATERIALS and METHODS Sampling and RNA extraction was done according to Handberg et al. 1999. RT-PCR protocols were according to Handberg et al. (1999) (IBVN primers) and Cavanagh et al. (1999) (XCE primers). All RT-PCR analyses were done using both assays and only considered positive if both were positive. Sequencing was performed by DNA Technology (www.dna-technology.dk). For sequence analysis we used CLC-DNA-Workbench (www.clcbio.com) for assembly, alignment and tree-building (NJ, bootstraps 1000). The sequence between position 755 to 1015 (260 bp) of the S1 gene was used for the analysis. +1 was defined as the A in the initial ATG. Data concerning abbatoir rejects were supplied by the Danish Poultry Council. RESULTS Since 2006 sequence fragments of the S1 gene have been derived from 174 cases of IBV infection detected in diagnostic samples from Danish poultry (Table 1). The sequence identifies the subtypes of IBV, and the results show that the QX D388 had the highest prevalence with detection in 133 samples. The subtypes Massachusetts and 793B were detected in 13 and 17 samples, respectively. D8880/B1648 and D274 were detected in 1 sample each. Phylogenetic analysis shows that the types were clearly separated (Fig 1). A phylogenetic analysis of the QX D388 type shows very similar sequences, however differences could be observed (Fig 2). The most significant differences with bootstraps value at 60 % were found in a group of samples from late 2008 and all samples from 2009. The QX D388 alignment (Fig 3) showed a unique variation at position 786 (CGC->CGT) for the sequences from late 2008 and all 2009. This variation correlated in time with increase of lesions for airsacculitis at the abattoirs (data not shown). A variation at position 987 (CAC->CAT) included the sequences above in addition to sequences from the middle of 2008. None of these variations could be found in any sequence deposited in Genbank. A variation at position 888 (TAC->TAT) was detected in all QX D388 from 2008-2009 and in 2007 sequences from the Danish island Bornholm. No other obvious connections to farm locations were observed among the sequences. The 888 variation was present in sequences deposited in Genbank. DISCUSSION This is the first report on detection of QX D388 in Danish poultry. The QX D388 sequences were very similar to the QX D388 deposited in Genbank, in agreement with the observations of Monne et al. (2008) and Worthington et al. (2008). QX D388 was the most prevalent type, but Massachusetts and 793B were detected as well, probably due to the vaccines. Other field type as D8880/B1648 and D274 were also detected, showing that theses types were circulating in Danish poultry. Internally the QX D388 type showed high degree of similarity. Only sequences from viruses sampled in late 2008 and in 2009 were significantly grouped. The time period

www.dna-technology.dk

www.clcbio.com


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when these viruses were sampled coincided with the period in which the Danish poultry industry reported increasing airsacculitis problems. Examination of the sequences revealed a silent variation at position 786 as the most significant change. However, two other silent variations were also observed in these sequences, but these groups included more sequences than the sequences possessing the 786 variation. Interestingly these variations could also be associated to a time period. The early sequences including the 888 variation were limited to the Danish island Bornholm. Bornholm is remote from the rest of Denmark, and was at that time a non-IBV-vaccinating area. These variations would be interesting for use as potential makers in epidemiological analyses. However, this will depend on improved validation including full-length S1 sequencing of selected viruses. This would also reveal if non-silent variations were associated with the pathological lesions, in accordance with observations in a study of Ammayappan et al. (2009). We therefore concluded that sequencing, even of a limited fragment of the IBV S1 gene could be used for typing and the variation observed could have potential as epidemiological markers. REFERENCES Arun Ammayappan, Chitra Upadhyay, Jack Gelb Jr and Vikram N.Vakharia. 2009.

Identification of sequence changes responsible for the attenuation of avian infectious bronchitis virus strain Arkansas DPI. Arch. Virol. 154: 495-499.

Cavanagh, D. 2007. Coronavirus avian infectious bronchitis virus. Vet. Res. 38: 281-297.

Cavanagh, D., Mawditt, K., Britton, P. & Naylor, C.J. 1999. Longitudinal field studies of infectious bronchitis virus and avian pneumovirus in broilers using type-specific polymerase chain reactions. Avian Pathology, 28, 593– 605.

Handberg, K.J., Nielsen, O.L., Pedersen, M.W. and Jørgensen, P.H. 1999. Detection and strain differentiation of infectious bronchtis virus in tracheal tissues from experimentally infected chickens by reverse transcription-polymersea chain reaction. Comparison with an immunohistochemical technique. Avian Pathology. 28: 327-335.

Monne I, Cattoli G, Jones R, Worthington K, Wijmenga W. 2008. QX genotypes of infectious bronchitis virus circulating in Europe. Vet. Rec. 163: 606-607.

Worthington K.J., Currie R.J.W., Jones R.C.. 2008. A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. 37: 247-257.


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Table 1: Detection of IBV in Danish poultry during 2006-2009

2006 2007 2008 2009 Total D388 2 37 67 27 133 793B 2 8 6 1 17 Mass 0 3 6 4 13 D274 1 0 0 0 1 B1648 0 0 1 0 1 D1466 0 0 0 0 0 Unknown 5 1 1 2 9 Total 10 49 81 34 174

Figure 1. Phylogentic tree of the Danish IBV sequences.

QXD388

D8880

793B

Mass

D274 Ref.


11

Figure 2. Part of the phylogenteic tree of the Danish QX D388 sequences

Late 2008 and most 2009


12

Figure 3. Part of the alignment of Danish QX D388 sequences The box indicating the variation CGC->CGT at position 786


13

MOLECULAR EPIDEMIOLOGY OF SLOVENE INFECTIOUS BRONCHITIS VIRUS STRAINS ISOLATED BETWEEN 1990 AND 2005

KRAPEŽ U1, SLAVEC B1, BARLIČ-MAGANJA D2 and ZORMAN ROJS O1

1 Institute for Poultry Health, Veterinary Faculty, University of Ljubljana, Ljubljana, Slovenia

2 College of Health Care Izola, University of Primorska, Izola, Slovenia

SUMMARY Molecular epidemiology of fifteen Slovenian infectious bronchitis virus (IBV) strains isolated between years 1990 and 2005 was studied. Fifteen IBV strains were divided into four genotypes by the molecular analysis of the S1 gene region. Four strains belonged to the Massachusetts genotype, one strain was placed into the genotype QX, one strain belonged to the B1648 genotype and nine strains were classified into the 624/I genotype. Nine Slovenian IBV strains of 624/I genotype formed two subgroups independently of the time of isolation and the geographical origin. Phylogenetic analysis of the partial N gene sequences revealed lower sequence variability and different clustering of the Slovenian IBV strains. Fourteen strains were grouped together with the reference strains from Massachusetts genotype. One strain formed a genetic group with the strain 793/B. Molecular analysis of the partial sequences of S1 and N gene showed that mutations together with intra and intergenic recombination events could contribute to the genetic diversity of the Slovenian isolates. INTRODUCTION Infectious bronchitis virus (IBV) is the etiological agent of infectious bronchitis (IB), which is an acute and highly contagious disease of the respiratory and sometimes the urogenital tract of chickens causing tracheal rales, sneezing, coughing, a poor weight gain and reduced feed efficiency in broilers and a decline in egg production and egg shell quality in layers (Cavanagh and Naqi, 2003a). Since IBV is endemic on all commercial sites, most flocks are vaccinated with live attenuated vaccines. Despite vaccination, IBV continues to be severe economic problem in commercial chickens


14

because many different serotypes of the virus exist and do not cross protect (Gelb et al., 1997; Farsang et al., 2002; Cavanag et al., 2003). IBV belongs to the family Coronaviridae. Coronaviruses are devided into three antigenic groups based primarily on their structural proteins. IBV is placed in antigenic group three (Enjuanes et al., 2000). Characteristic of this group are a cleaved spike (S) glycoprotein, an N-glycosilated membrane (M) protein, and no hemagglutininin/esteraze protein (Sidell SG, 1995). Virus genome is a linear, non-segmented, positive sense, single stranded RNA of approximately 27-kilo bases (kb) in length. The first 20 kb encode the viral RNA-dependent RNA polymerase and proteases. The reminder of the genome encodes five structural proteins, the spike (S) consisting of S1 and S2, envelope (E), membrane (M) and nucleocapsid (N) proteins, four small non-structural proteins, 3a, 3b, 5a and 5b, and a 3’ untranslated region (UTR) (Lai and Cavanagh, 1997). The S1 subunit contains epitopes that induce virus-neutralizing antibodies and serotype-specific antibodies (Cavanagh et al., 1992; Jia et al., 1996; Cavanagh et al., 1997). The S1 is also responsible for cell attachment and determining the tissue tropism (Cavanagh et al., 1983). Different serotypes and variants of IBV are thought to be generated by amino acid changes in the N-terminal part of the S1 protein resulting from nucleotide deletions, insertions, or point mutations made by the viral polymerase (Kusters et al., 1990; Cavanagh et al., 1992; Kant et al., 1992; Jia et al., 1996). Variations in the S1 and N genes, in particular, are believed to be of circuital importance for emergence of variants because of their role in virus replication and immunity, and hence S1 and N have been used most frequently to determine the relatedness of emerging IBV (Cavanagh et al., 1992; Tseng et al., 1996; Cavanagh et al., 1997; Lee and Jackwood, 2000). The N protein, located in the capsid of the virion is involved in RNA replication and carries group-specific antigenic determinants (Ignajatovic et al., 1995). The purpose of the present study was to genetically characterize fifteen IBV strains isolated in Slovenia between 1990 and 2005 by sequencing the region encompassing hyper variable region (HVR) 1 and HVR2 of S1 gene and region on the N gene. Sequences were compared to other published IBV sequences to establish genetic features, origin and evolution of Slovenian strains. MATERIAL and METHODS Virus strains Virus isolations were performed at the Institute for Poultry Health, Veterinary Faculty, University of Ljubljana, by inoculation of different tissue suspensions, obtained from dead chickens, into the allantoic cavity of 9 to 10-day-old embryonated specific-pathogen-free (SPF) chicken eggs as described by Gelb, Jr. and Jackwood (Gelb, Jr. and Jackwood, 1998).


15

RNA extraction, reverse transcription, polymerase chain reaction and nucleotide sequencing RNA was extracted from infectious allantoic fluids obtained after first passage by QIAamp Viral RNA Mini Kit (QIAGEN, USA), according to the manufacturer’s instructions. A region of 573 to 601-bp of the S1 gene, including HVR1 and HVR2, was amplified by oligonucleotide primer pair: CK4: 5'-TCA AAG CTT CAN GGN GGN GCN TA-3' and CK2: 5'-CTC GAA TTC CNG TRT TRT AYT GRC A-3' (Keeler et al., 1998). Oligonucleotide primer pair NP1: 5'-GGT AGY GGY GTT CCT GAT AA-3' and NP2: 5'-TCA TCT TGT CRT CAC CAA AA-3' was used for the amplification of 618-bp region of the N gene (Tseng et al., 1996). A single tube RT-PCR system (OneStep RT-PCR Kit, QIAGEN, USA) was used for the genomic RNA amplification. The RT-PCR was performed by uninterrupted thermal cycling with the following program: 30 min. at 50oC for reverse transcription, reverse transcriptase inactivation at 95oC for 15 min. was followed by 40 cycles of denaturation at 94oC for 30 sec., annealing at 50oC for 1 min. (with primers CK4/CK2) or 47oC for 1 min. (with primers NP1/NP2), extension at 72oC for 1 min. and final extension at 72oC for 10 min. The reaction’s products were analyzed by electrophoresis on a 1,8% agarose gel stained with ethidium bromide. The DNA fragments were excised from gel and purified with a Wizard PCR Preps DNA Purification System (Promega Corp, USA). PCR products were cloned by TOPO TA Cloning kit (Invirogen, USA). Double-stranded nucleotide sequencing was completed by using the ABI PRISM Big Dye Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems, USA) and oligonucleotide primers used for RT-PCR. Reactions were analyzed with an ABI3730xl Genetic Analyzer (Applied Biosystems, USA). Sequence analysis The MEGA 3.1 software (Kumar et al., 2004) was used for editing nucleotide sequences and deducing amino acid sequences. Nucleotide and deduced amino acid sequences were aligned with ClustalW software (Thompson et al., 1994). Phylogenetic analyses were constructed with MEGA 3.1 software using the neighbor-joining method with the Kimura-2 parameter substitution model and 1000 bootstrap replicates to assign confidence level to branches. Virus isolates used for pairwise comparisons on S1 gene B1648, (X87238); N1/62, (U29522); D274, (X15832); Gray, (L14069); Ark99, (M99482); 4/91, (AF093794); Connecticut, (L18990); Beaudette, (X02342); Mass41, (X04722); H120, (M21970); QXIBV, (AF193423); LX4, (AY189157); L-1148, (DQ431199); FR/L-1450T/05, (EF079118); IS/1201, (DQ400359); K1255/3, (AY790364); N1/88, (U29450); DE/072/92, (U77298); D1466, (X58001).


16

Virus isolates used for pairwise comparisons on N gene N1/62, (U52596); 793/B, (DQ294723); Ark99, (M85245); Gray, (S48137); Connecticut, (AY942746); Beaudette, (M28565); Mass41, (AY851295); H120, (AY028296); QXIBV, (AF199412); LX4, (AY338732); K1255/3, (AY790353); N1/88, (U52599); DE/072/92, (AF203001); D1466, (AF203006). RESULTS Phylogenetic analysis of partial S1 genes indicated that the Slovene IBV strains do not cluster together, but instead are split between the four different genotypes. Four strains (SLO/54/90, SLO/632/92, SLO/244/93 and SLO/202/94) were placed in the Massachusetts genotype, strain SLO/99/05 clustered together with the strains from Chinese QX genotype, strain SLO/266/05 form a cluster together with the strain B1648 and nine strains (SLO/682/91, SLO/2/96, SLO/31/96, SLO/136/96, SLO/186/96, SLO/809/97, SLO/263/98, SLO/276/98 and SLO/267/99) were classified in the Italian 624/I genotype. Slovenian strains of 624/I genotype formed two clusters independently of year and geographical origin of isolations. Strains SLO/682/91, SLO/31/96, SLO/136/96, SLO/186/96 and SLO/809/97 were grouped together with the Italian strain 624/I isolated in 1996 (Figure 2). Partial S1 gene nt sequences identities between the fifteen Slovenian IBV strains were 67.6% to 99.8%. Identities between aa sequences were 63.6% to 100%. Strains SLO/54/90, SLO/632/92, SLO/244/93 and SLO/202/94 placed in Massachusetts genotype by phylogenetical analysis (Figure 2) had nt and aa sequences the most similar (98.4% to 99% and 97.6% to 98.2%, respectively) to the strain H120. Alignment of the aa sequences of strains SLO/54/90, SLO/632/92, SLO/244/93 and SLO/202/94 with strain H120, showed following aa replacements at the position 121 within HVR2: Val to Tyr (SLO/632/92 and SLO/54/90), Val to Gly (SLO/244/93) and Val to Asp (SLO/202/94) (Figure 1). Strain SLO/99/05 placed in Chinese QX genotype (Figure 2) had nt and aa sequences the most similar (97.3% and 94.7%, respectively) to the strain FR/L-1450T/05. Alignment of the aa sequences of strain SLO/99/05 with the other strains from that genotype (FR/L-1450T/05, QXIBV, LX4, L-1148, IS/1201 and K1255/03) revealed that five out of six aa differences between SLO/99/05 and other five strains are at the positions within HVR1 and HVR2: 61 Ser to Asp, 67 Val to Ile, 119 Ser to Ile, 130 Arg to Ser, 142 Ser to Phe and 204 Glu to Gln (Figure 1). Strain SLO/266/05 classified together with the strain B1648 by phylogenetical analysis (Figure 2) had nt and aa sequences most identical (86.3% and 80.6%, respectively) to the same strain. Comparison of the protein S1 5’ region comprising HVR1 (region of protein S1 comprising 48 aa) showed that strain SLO/266/05 has less aa changes (seven, 14.5%) when compared with the strain 624/I, then when compared to the strain B1648 (sixteen, 33.3%). However, the other investigated part of the protein S1 of SLO/266/05 (130 amino acids) comprising HVR2, had less amino acids changes when compared to strain B1648. In that region eighteen aa substitutions (13.8%) between


17

SLO/266/05 and B1648, and twenty-seven (20%) aa substitutions between SLO/266/05 and 624/I were found (Figure 1). Strains SLO/682/91, SLO/2/96, SLO/31/96, SLO/136/96, SLO/186/96, SLO/809/97, SLO/263/98, SLO/276/98 and SLO/267/99 of Italian 624/I genotype (Figure 2) had nt and aa sequences the most similar (86.3% to 96.9% and 80.8% to 97.1%, respectively) to the strain 624/I. Aligned aa sequences of the strains SLO/682/91, SLO/2/96, SLO/31/96, SLO/136/96, SLO/186/96, SLO/809/97, SLO/263/98, SLO/276/98 and SLO/267/99 were compared with each other and with the strain 624/I. Strains from first group: SLO/682/91, SLO/31/96, SLO/136/96, SLO/186/96 and SLO/809/97 had aa sequences more similar to reference strain 624/I than strains from second group: SLO/2/96, SLO/263/98, SLO/276/98 and SLO/267/99. Alignment of the aa sequences of strains from the first group with the strain 624/I revealed aa changes at following positions: 117 Ser to His (except SLO/682/91), 121 Gln to Asn (SLO/31/96, SLO/136/96 and SLO/186/96) or Gln to Val (SLO809/97) (Figure 1). Alignment of the aa sequences of strains from second group with the strain 624/I revealed aa changes at following positions: 54 Leu to Thr, 79 Ser to Tyr, 91 Gln to Gly, 94 Thr to Val, 95 Asn to Ser, 107 Phe to Ile, 119 Ala to Gln, 121 Gln to Glu, 126 Gln to Leu, 128 Leu to Ile, 129 Pro to Gln, 136 Ser to Ala, 143 Lys to Asn, 145 Asn to Pro 146 Ser to Thr, 147 Ser to Asp, 157 Thr to Ala, 161 Thr to Lys, 163 Lys to Met, 169 Asn to Ser, 171 His to Tyr, 189 Ser to Hist, 190 Ile to Val, 192 Ala to Gly (Figure 1). Slovenian strains were classified into two genetic groups by the phylogenetic analysis of partial N protein genes. Fourteen strains were grouped together with strains H120, D1466 and 624/I. Strain SLO/682/91 formed a separate cluster together with the strain 739/B (Figure 3). Partial N gene nucleotide sequences identities between the fifteen Slovenian IBV strains were 92% to 100%. Identities between aa sequences were 82.5% to 100%. Strain SLO/682/91 had nt and ak sequences most identical (92.1% and 85.2%, respectively) to strain 793/B. Fourteen other Slovenian strains had nt and aa sequences the most similar (97.1% to 99.6% and 97.2% to 99.4%, respectively) to the strain H120. DISCUSSION Fifteen Slovenian IBV strains were classified into four different genotypes based on analysis of S1 gene region (Figure 2). Four strains: SLO/54/90, SLO/632/92, SLO/244/93 and SLO/202/94 that shared high nucleotide and amino acid identities, notwithstanding that they were isolated over a period of four years in different geographical origins, were placed in the Massachusetts genotype (Figure 2). Four strains were the most similar (98.4% to 99%) to the attenuated vaccine strain H120. Strains SLO/54/90, SLO/632/92, SLO/244/93 and SLO/202/94 were isolated from the broilers flocks that showed clinical signs of infectious bronchitis but have not been vaccinated against IBV. Based on these


18

findings, the question arises whether the strains derived from attenuated vaccine strain H120. On the basis of molecular analysis of the S1 and N protein gene could be assumed that the strains SLO/54/90, SLO/632/92, SLO/244/93 and SLO/202/94 developed from the vaccine strain H120. The data that support this assumption are: a) Strains derived from an unvaccinated broiler flocks. Vaccination of broilers against IBV has been at that time period largely dependent on the epidemiological situation (Zorman-Rojs, 1993), b) The pathogenicity of the strain SLO/244/93 for the SPF chicken embryos as well as for the day-old SPF chickens was examined. It was found that the ELD50 increased with passages on the embryos, and that after inoculation, the strain caused respiratory disease and death in SPF chickens (Zorman-Rojs, 1993). Vaccines with attenuated viruses are prepared by passages of field strains in the chicken embryos. This is a selection process in which subpopulations of lower virulence prevails, but the vaccine still contains subpopulations of viruses, much smaller fractions, with virulence similar to that of the original strain for the preparation of the vaccine. Subpopulations have also slightly different antigens, thus providing a wider immunity. The possibility of re-expression of pathogenic subpopulations is more likely in an environment where the use of the vaccine have been stopped, and implemented bio-security measures were poor (Nix et al., 2000). In such an environment cyclical infections of animals with vaccine strains may lead to increase in virulence of attenuated strains or re-expression of pathogenic subpopulations (Hopkins and Yoder, 1986). The four Slovenian strains that were isolated from broilers with clinical signs of IB have very likely arisen from vaccine strain H120. The differences between them could be explained by different subpopulations of viruses in a vaccine and/or accumulation of point mutations during the passages of these strains in non-immune broiler flocks. Strain SLO/99/05 was classified by molecular analysis of a region on the S1 gene into the Chinese QX genotype of nephropathogenic strains that was proposed by Liu and Kong (Liu and Kong, 2004) (Figure 2). IBV strains from Chinese QX genotype have been demonstrated also in Italy (Beato et al., 2005), France (Worthington & Jones, 2005), Holland (Worthington et al., 2004), Poland (Domanska-Blicharz et al., 2006), Russia (Bochkov et al., 2006), Israel (Meir and Manarat, 2004) and Korea (Jang and Kwon, 2004). Strain SLO/99/05 shared high partial S1 gene identity with the strains from QX genotype, whereas identities of partial N gene sequence were low. High nucleotide identity 98,7% and 98,5% were found when partial N gene sequence of strain SLO/99/05 was compared with those of vaccine strains H120 or D1466, respectively. Strain SLO/99/05 had very likely arisen by intergenic recombination between strain from QX genotype and attenuated vaccine strain H120 or D1466. Use of vaccines prepared from strain H120, is widespread in Asia and Europe (Cavanagh and Naqi, 2003), but it is known that vaccinated poultry is not protected against infection with strains of QX genotype (Liu and Kong, 2004). Infection of animals with strains from different genotypes may lead to the replacement of gene segments and to the emergence of new recombinant viruses (Brian and Spaan, 1997). Strains of Massachusetts serotype,


19

including the H120, have been often involved in recombination because they are frequently used as a vaccine strains (Wang et al., 1994; Jia et al., 1995; Kottier et al., 1995). Further studies of the N protein gene of other European QX strains would show whether they have been generated by recombination between the Asian strains and strains H120 or D1466. The evolution of the virus is continuing and living process (Holland et al., 1982; Kusters et al., 1989; Kusters et al., 1990; Jia et al., 1995), so it is possible, that the European strains, which are placed in QX genotype based on the S1 gene, would be classified in the other genotypes on the basis of other structural genes (Lee and Jackwood, 2000). Strain SLO/266/05 shared high partial S1 gene identity with the strain B1648 and strain 624/I, whereas sequence of partial N gene showed high identities with the strains H120, D1466 and 624/I. Phylogenetic analysis based on the partial S gene sequence grouped Slovenian strain SLO/266/05 together with the strain B1648 (Figure 2). Molecular analysis of partial S1 gene sequences suggested that strain SLO/266/05 has undergone an intragenic recombination event between strain from B1648 and 624/I genotypes. This proposal was based on the following findings: a) comparison of the protein S1 5’ region comprising HVR1 (region of protein S1 comprising 48 amino acids) showed that strain SLO/266/05 has less amino acid changes when compared to strain 624/I than to strain B1648. However, the other investigated part of the protein S1 (130 amino acids) comprising HVR2 has less amino acids changes when compared to strain B1648 (Figure 1); b) phylogenetic analysis performed on the both mentioned S1 gene regions independently, showed that classification of strain SLO/266/05 into genetic group was different, depending on the S1 region analyzed. Phylogenetic analysis performed on the S1 5’ region grouped strain SLO/266/05 together with 624/I genotype strains. Phylogenetic analysis based on the other investigated part of protein S1 revealed that SLO/266/05 was closely related with B1648 strain (results not shown). Recombination is rare in the region between HVR1 and HVR2 of S1 protein gene (Wang et al., 1997), but it could be proposed that S1 protein gene of strain SLO/266/05 was created by recombination between strain B1648 and 624/I and the accumulation of point mutations. Recombination event between strains from the 624/I and the B1648 genotypes could take place in the conserved region on the S1, which stretches between the amino acid at position 96 and 118 (Figure 1). Recombination and mutation are the mechanisms that cause the genetic diversity of IBV (Kusters et al., 1989; Kusters et al., 1990; Wang et al., 1993; Wang et al., 1994). Further investigation including sequencing of the whole S1 gene of SLO/266/05 strain as well as molecular analysis of the nucleotide and the deduced amino acid sequences have to be performed to confirm the preliminary findings. Phylogenetic analysis of the partial S1 gene sequences showed that nine Slovenian IBV strains isolated between 1991 and 1999 belong to the Italian 624/I genotype (Figure 2). Strains from 624/I genotype have been present in Slovenia since 1991 that was three


20

years before first detection in Italy (Capua et al., 1994). The last strains from the 624/I genotype were isolated in Slovenia as well as in Italy in 1999 (Bochkov et al., 2007). The first Slovenian strain of the genotype 624/I, the prototype strain SLO/682/91, was isolated from the broiler farm, located only thirty kilometers from the Italian border. Among the nine Slovenian strains of 624/I genotype this strain was the most closely related to the Italian strain 624/I, which was isolated in 1996 Figure 1, 2). Next four strains of this genotype were isolated in 1996. One strain was isolated from the broiler farm in the immediate vicinity of the site, where the first 624/I genotype strain was detected. The remaining three were isolated from geographically distant farms. Phylogenetic analysis of the partial S1 gene sequences of Slovenian 624/I strains revealed that Slovenian strains form two clusters within 624/I genotype independently of the time period and geographical origin of isolation (Figure 2). Comparison of the partial N gene nucleotide sequences and the deduced amino acid sequences showed high identities among the 624/I strains and strain H120, except the first strain (SLO/682/91) of this genotype isolated in Slovenia. The results showed that recombination events have probably occurred between attenuated vaccine strain H120 and the parental strains of 624/I genotype. The attenuated strain H120 was widely used for vaccination in Slovenia. These strains were suggested to provide large donor pool for IBV recombination (Wang et al., 1994; Jia et al., 1995; Kottier et al., 1995). Further molecular analysis of N gene of Italian 624/I strains that were isolated in 1990’s have to be performed to confirm this proposal. REFERENCES Beato MS, De Battisti C, Terregino C, Drago A, Capua I, Ortali G (2005). Evidence of

circulation of a Chinese strain of infectious bronchitis virus (QXIBV) in Italy. Vet Rec 28:720.

Bochkov YA, Batchenko GV, Shcherbakova LO, Borisov AV, Drygin VV (2006). Molecular epizootiology of avian infectious bronchitis in Russia. Avian Pathol 35: 379-93.

Bochkov YA, Tossi G, Massi P, Drygin VV (2007). Phylogenetic analysis of partial S1 and N gene sequences of infectious bronchitis virus isolates from Italy revealed genetic diversity and recombination. Virus Genes 35(1): 65-71.

Brian DA, Spaan WJM (1997). Recombination and coronavirus defective interfering RNAs. Semin Virol 8: 101-11.

Capua I, Gough RE, Mancini M, Casaccia C, Weiss C (1994). A ‘novel’ infectious bronchitis strain infecting broiler chickens in Italy. J Vet Med B 41: 83-9.

Cavanagh D. J (1983). Coronavirus IBV: structural characterization of the spike protein. Gen Virol. 64: 2577-83.

Cavanagh D, Davis PJ, Cook JK, Li D, Kant A, Koch G (1992). Location of the amino acid differences in the S1 spike glycoprotein subunit of closely related serotypes of infectious bronchitis virus. Avian Pathol 21(1): 33-43.


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Cavanagh D, Ellis MM, Cook JKA (1997). Relationship between sequence variation in the S1 spike protein of infectious bronchitis virus and the extent of cross-protection in vivo. Avian Pathol 26: 63-74.

Cavanagh D, Naqi SA (2003a). Infectious bronchitis. In: Saif YM, ed. Diseases of Poultry 11th ed. Ames: Iowa State University Press, 101-19.

Cavanagh D (2003b). Severe acute respiratory syndrome vaccine development: experiences of vaccination against avian infectious bronchitis coronavirus. Avian Pathol 32(6): 567-82.

Domanska-Blicharz K, Minta Z, Smietanka K, Porwan T (2006). New variant of IBV in Poland.Vet Rec 158: 808.

Enjuanes LD, Brian D, Cavanagh D, et al. (2000). Coronaviridae. In. Murphy FA, ed. Virus Taxonomy. New York: Academic Press, 835-49.

Farsang A, Ros C, Renström LH, Baule C, Soós T, Belák S(2002). Molecular epizootiology of infectious bronchitis virus in Sweden indicating the involvement of a vaccine strain. Avian Pathol. 31(3): 229-36.

Gelb J Jr, Keeler CL Jr, Nix WA, Rosenberger JK, Cloud SS (1997). Antigenic and S-1 genomic characterization of the Delaware variant serotype of infectious bronchitis virus. Avian Dis 41(3): 661-9.

Gelb J Jr. and MW Jackwood (1998). Infectious bronchitis. In: Swayne DE, Glisson JR, Jackwood MW, Pearson JE, Reed WM, eds. A laboratory manual for the isolation and identification of avain pathogens. 4th ed. American association of avian pathologists, 169-74.

Hopkins SR, Yoder HW Jr (1986). Reversion to virulence of chicken passaged infectious bronchitis vaccine virus. Avian Dis 30: 221-3.

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http://www.ncbi.nlm.nih.gov/entrez/viewer


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Kottier SD, Cavanagh D, Britton P (1995). Experimental evidence of recombination in coronavirus infection bronchitis virus. Virology 213: 569-80.

Kumar S, Tamura K, Nei M (2004). MEGA3: integrated software for molecular evolutionary genetics analysis and sequence alignment. Brief Bioinform 5(2):150-63.

Kusters JG, Niesters HG, Lenstra JA, Horzinek MC, van der Zeijst BA (1989). Phylogeny of antigenic variants of avian coronavirus IBV. Virology 169(1): 217-21.

Kusters JG, Jager EJ, Niesters HG, van der Zeijst BA (1990). Sequence evidence for RNA recombination in field isolates of avian coronavirus infectious bronchitis virus. Vaccine 8(6): 605-8.

Lee C-W, Jackwood MW (2000). Evidence of genetic diversity generated by recombination among avian corona virus IBV. Arch Virol 145: 2135-48.

Lai MM, Cavanagh D (1997). The molecular biology of coronaviruses. Adv Virus Res 48: 1-100.

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Meir R, Maharat O (2004). Sequence analysis of the S1 gene of infectious bronchitis viruses isolated in Israel in 2004. National center for biotechnology information, CoreNucleotide. http://www.ncbi.nlm.nih.gov/entrez/viewer. 15. 10. 2006

Nix WA, Troeber DS, Kingham BF, Keeler Jr CL, Gelb Jr J (2000). Emergence of subtype strains of the Arkansas serotype of infectious bronchitis virus in Delmavra broiler chickens. Avian Dis 44: 568-81.

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progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 22(22): 4673-80.

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Wang L, Xu Y, Collisson EW (1997). Experimental confirmation of recombination upstream of the S1 hypervariable region of infectious bronchitis virus. Virus Res 49: 139-45.



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Worthington KJ, Jones RC (2005). Molecular characterization of an infectious bronchitis virus from France, isolated in 2005, from tracheas of 10 day old commercia future breeder pullets with respiratory problems and whose parent flock had oviduct lesions. National center for biotechnology information, CoreNucleotide. http://www.ncbi.nlm.nih.gov/entrez/viewer. 15. 10. 2006

Zorman-Rojs O (1993). Kvantifikacija postvakcinalne imunosti infekcioznega bronhitisa perutnine. Ljubljana: Veterinarska fakulteta. Doktorska disertacija.

Table 1: Slovene IBV isolates used in the study.

Strain Year of isolation Animals Vaccination against IBV

SLO/54/90 1990 broilers no












SLO/267/99 1999 broiler breeders yes





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Figure 1: Amino acid sequence alignment of partial S1 protein sequences.


25

Figure 2: Phylogenetic relationship based upon partial nucleotide sequences of the S1 gene of the Slovene IBV strains (bold) and non-Slovene IBV strains generated by the neighbor-joining method with 1000 bootstrap replicates.


26

Figure 3: Phylogenetic relationship based upon partial nucleotide sequences of the N gene of the Slovene IBV strains (bold) and non-Slovene IBV strains generated by the neighbor-joining method with 1000 bootstrap replicates.


27

FIRST REPORT OF IBV QX-LIKE STRAINS IN SPAIN

DOLZ R 1, BERTRAN K 1and MAJÓ N1,2

1Centre de Recerca en Sanitat Animal (CReSA), 08193, Barcelona, Spain. 2Departament de Sanitat i Anatomia Animals, Universitat Autònoma de Barcelona,

08193, Barcelona, Spain. SUMMARY In Spain, since 2006, continuous surveillance studies are carried out in our laboratory based on the analysis of submitted samples from clinical cases suspected of having IB. IBV detection and identification is based on RT-PCR and S1 gene sequencing. Furthermore, IBV isolation is attempted from positive samples. In March 2008, QX-like strains were identified in two distinct farms of layer pullets and slow growing chickens. In both cases, birds were younger than 25 days of age and showed respiratory and renal disease. For several months after its first identification, this genotype was not further detected in any of the clinical cases submitted to our laboratory. Surprisingly, in October 2008 a QX-like strain was again detected in 40-day-old broilers with respiratory signs from a distinct geographic area of Spain. In 2009 IBVQX strains have been detected in two clinical cases. Based on these observations, QX-like strains are circulating in Spain, although up to now they are only sporadically involved in clinical cases. However, as it has been described in other countries, an increasing epidemiological importance of this new genotype in our country may be expected. INTRODUCTION The first data reported from QX infectious bronchitis virus (IBV) genotype, was the submission of the S1 gene sequence in Genbank database in 1999 (AF193423). In this submission, it was indicated that this strain had been isolated from Qingdao (China) and it was noted to be a proventriculus type. Subsequent epidemiological studies in China revealed this genotype as one of the most common genotypes in this country (Liu et al. 2006).


28

In 2005, presence of QX-like strains was reported for the first time in Europe due to an IBV surveillance programme carried out in Italy (Beato et al. 2005). In 2006, circulation of QX-like strains in several other European countries including France, Belgium, Germany, Holland and Poland was also reported (Domanska-Blicharz et al. 2006; Philipp and Voss 2006; Wit et al. 2006; Worthington and Jones 2006b). Based on these reports, first European QX-like strains were detected in the Netherlands in 2003 (Wit et al. 2006). However, the results obtained in our laboratory at that time did not indicate the circulation of this genotype in Spain, where Italy 02 was the predominant IBV genotype. Similarly, QX-like strains had not been identified in UK either at that time (Worthington and Jones 2006b). IBV situation in Spain was in-depth studied by a retrospective study carried out in our laboratory were IB viruses isolated between 1992 and 2005 were molecularly characterized. From this study it was revealed that during the nineties the prevalent genotype in field in Spain was the 4/91 genotype. However, in 1997 the new Italy 02 genotype appeared and after a period of co-circulation of both genotypes, Italy 02 displaced 4/91 genotype and became the predominant genotype. Since 2005, continuous surveillance of IBV genotypes is carried out by sequencing IBV isolates detected in clinical samples submitted by field veterinarians. Although during 2006 and 2007 all clinical outbreaks were related with Italy 02 isolates, the situation changed in 2008. In this paper, the current epidemiological situation of Spain (2008 and 2009) regarding IBV is revised with special attention to the first cases of QXIBV-like strains in our country. MATERIALS and METHODS IBV RT-PCR IBV detection was attempted from clinical samples (tracheas, kidneys or caecal tonsils) or clinical cases submitted to our center by veterinarians in which IBV was suspected based on clinical signs. Epidemiological information of clinical cases included in the study is summarized in Table 1. Viral RNA was extracted from tissues with the Nucleospin RNA Virus Kit (Macherey-Nagel) following manufacturers instructions. A reverse-transcriptase polymerase chain reaction (RT-PCR) amplifying a fragment of the 5’ non-coding region (UTR) was used to detect IBV within the samples as previously described by Adzhar and colleagues (Adzhar et al. 1996). S1 gene sequencing Partial S1 gene sequencing was carried out for all positive samples. Direct sequencing from clinical samples was first attempted by using S1UNI2+ and XCE2- or S1UNI2+ and S1PR2- primers (Adzhar et al. 1996; Dolz et al. 2006). When direct sequencing was not successful, clinical samples were inoculated in SPF chicken embryos for virus isolation. Allantoic fluid from each passage was tested for the presence of IBV, and also used for


29

the partial sequencing of the S1 gene. In addition, the S1 gene was entirely sequenced from the five clinical cases from which QXIBV-like strains had been detected by partial sequencing. An RT-PCR to amplify the complete S1 gene was carried out for the QXIBV-like strains as previously described (Adzhar et al. 1996). RT-PCR products were purified by QIAquick gel extraction Kit (Qiagen) according to the manufacturer’s instructions. Purified RT-PCR products were sequenced using ABI PRISM BigDye® Terminator v3.1 Cycle Sequencing Kit (PE Biosystems) as described by the manufacturer. Sequences were analyzed with an automated nucleic acid analyser (ABI PRISM 3100 Avant; PE Biosystems) Nucleotide and amino acid deduced sequence analyses Nucleotide and amino acid deduced sequences were aligned using ClustalX software included in BioEdit 5.0 package. Neighbour-Joining method with 1000 bootstraps replicates within the software MEGA version 3.0 (Kumar et al. 2004) was used to perform phylogenetic analyses. GenBank accession numbers S1 gene nucleotide and amino acid deduced sequences of the 5 Spanish QXIBV-like isolates were submitted to the GenBank nucleotide database. GenBank accession numbers for these isolates are presented in Table 1. GenBank accession numbers of the IBV isolates used in the phylogenetic comparisons included: Ark-99 (L10384); A2 (AY043312); Beaudette (X02342); B1648 (X87238); CK/CH/LHLJ/04V (DQ167139); CK/CH/LLN/98I (DQ167145); CK/CH/LSHH/03I (DQ167149); D1466 (M21971); D207 (M21969); D274 (X15832); D3896 (X52084); FR-CR88061-88 (AJ618986); FR-L1450L-05 (EF079117); FR-L1450T-05 (EF079118); FR-85131-85 (AJ618985); Gray (L14069); HBN (DQ070837); Holte (L18988); H120 (M21970); IBVQ (DQ480155); IS/1201 (DQ400359); Itay 02 (AJ457137); It-497-02 (DQ901377); JMK (L14070); K1019/03 (FJ807927); LX4 (AY189157); M41 (M21883); NL-L1449T-04 (EF079116); NL-L1449K-04 (EF079115); Spain/92/35 (DQ386091); Spain/96/334 (DQ064804); Spain/98/308 (DQ064807); Spain/98/328 (DQ386096); Spain/99/316 (DQ064809); Spain/00/336 (DQ386098); Spain/00/337 (DQ064813); Spain/04/221 (DQ386103); Spain/05/866 (DQ386102); UK-L633-04 (DQ901376); UK-1233-95 (AJ618984); UK/3/91 (Z83977); UK/7/91 (Z83975); 3654-VM (AY544776); 4/91 attenuated (AF093793); 4/91 pathogenic (AF093794). RESULTS IBV was detected in sixty-six clinical cases suspected of having IB from January 2008 until May 2009, from which forty-one isolates of IBV were genotyped by partial S1 gene sequencing. The predominant genotype in Spain during this period was still the Italy 02 genotype (29 isolates – 70.7%). Spanish Italy 02 strains isolated in 2008-2009 were a homogeneous group sharing 99.1% average nucleotide identities among them. All these isolates were grouped together with the more recent Spanish Italy 02 isolates (Figure 1) cluster.


30

In 5 clinical cases (12.2%) the sequences obtained showed maximum nucleotide (95.5%-99.1%) and amino acid sequence similarities (94% - 98.8%) and were clustered together in the phylogenetic tree with QXIBV genotype strains (Figure 1). When the complete S1 gene sequences of Spanish QXIBV strains were compared among them average nucleotide and amino acidic sequence identities were 98.9% and 98% respectively. Similar nucleotide and amino acidic homologies were obtained when they were compared with European and Korean QXIBV isolates (98.9% and 98.3% respectively). However, higher sequence distances were observed with Chinese QXIBV isolates (96.1% average nucleotide similarity and 95% average amino acid similarity) The two first QXIBV isolates were detected in March 2008 in pullet layers (IBV/La/SP/17/08) and slow growing chickens (IBV/Ck/SP/18/08) younger than 25 days of age. In both cases, birds showed similar lesions including dehydratation, fibrinous aerosaculitis, and enlarged and pale kidneys. Some birds also showed fibrinous poliserositis. Both cases were submitted the same day and both farms were closely located and shared the farmers, which probably was the main spreading source. Pullet layers were monitorized until the achievement of sexual maturity and no reproductive alterations were observed in that flock. For several months after the first detection of QXIBV, this genotype was not further detected in submitted samples. However, in October 2008 a QXIBV isolate was again identified in 41-day-old broilers experiencing continuous respiratory problems since 20 days of age that did not successfully respond to antibiotic treatment. The virus was isolated from tracheas and caecal tonsils. In 2009, QXIBV strains have been detected in two clinical cases of layer hens 33-week-old with reproductive alterations and in 10-day-old broiler chicks with severe respiratory problems. Therefore, IBVQX infections in Spain have been associated with renal and respiratory disease mostly in young birds. Moreover, in one clinical case, a sequence with maximum nucleotide and amino acid sequence similarities with D1466 isolates was identified (Figure 1). Samples were submitted from layer hens with severe respiratory disease. DISCUSSION Continuous molecular surveillance studies regarding IBV are carried out in our laboratory based on the analysis of samples from submitted clinical cases suspected of having IB which allows rapid identification of new isolates circulating in field. During the last years, results obtained in our laboratory and also in other laboratories suggested that Italy 02 was almost the only IBV genotype involved in clinical IB outbreaks in Spain. However, this scenario was contradictory with the one observed in most European countries, where Italy 02 had appeared but it had not been as successfully established as in Spain. For instance, presence of the new QXIBV genotype had been reported in most of these countries, with the exception of UK and Spain.


31

In UK, first report of QXIBV genotype was in 2007 in a Pekin bantam (Worthington and Jones 2006a). At that time in Spain, presence of QXIBV isolates were suspected because the observation of reproductive alterations associated with hydropic oviducts in two layer hens flocks. However, it was not possible to confirm IBV as the etiological agent of the lesions observed in those birds. In the present study, first confirmed cases of QXIBV genotype in Spain are reported. From our data, QXIBV strains are circulating and widely geographically distributed in our country. However, they are sporadically involved in clinical IB outbreaks, being Italy 02 still the predominant genotype in Spain. In Spain, QXIBV strains are mostly related with respiratory and renal problems in young birds. Although in one case has been detected in adult hens with drop in egg production, other infectious agents were also present in those birds, and therefore, it was not possible to determine the effect of the virus in that clinical case. Furthermore, during 2009 one isolate from D1466 genotype has also been detected from layer hens with respiratory problems. As far as authors know, this is the first report of QXIBV and D1466 strains in Spain. Thus, the epidemiological scenario regarding IBV in Spain is changing, and several IBV subpopulations are co-circulating in field. At the end of the nineties, after a period of IBV subtypes co-circulation in Spain, one genotype emerged as a predominant displacing previous genotypes in field. However, homologous vaccines against one of the genotypes existed at that time, which probably influenced in the final resulting situation. Nowadays, neither Italy 02 nor QXIBV homologous vaccines exist in Spain and heterologous vaccines are used in order to control IB outbreaks. Taking into account the experience in other European countries regarding IBVQX strains, an increasing epidemiological importance by this new genotype would be expected (Worthington and Jones 2006b). However, as it has been previously stated, epidemiological relevance of Italy 02 genotype in Spain is higher than in other countries, which may contribute to a low incidence of QXIBV clinical outbreaks. REFERENCES Adzhar A, Shaw K, Britton P, Cavanagh D (1996) Universal oligonucleotides for the

detection of infectious bronchitis virus by the polymerase chain reaction. Av Pathol 25:817-836.

Beato MS, De Battisti C, Terregino C, Drago A, Capua I, Ortali G (2005) Evidence of circulation of a Chinese strain of infectious bronchitis virus (QXIBV) in Italy. Vet Rec 156:720.

Dolz R, Pujols J, Ordonez G, Porta R, Majo N (2006) Antigenic and molecular characterization of isolates of the Italy 02 infectious bronchitis virus genotype. Avian Pathol 35:77-85.

Domanska-Blicharz K, Minta Z, Smietanka K, Porwan T (2006) New variant of IBV in Poland. Vet Rec 158:808.

Kumar S, Tamura K, Nei M (2004) Integrated software for Molecular Evolutionary Genetics Analysis and sequence alignment. Brief Bioinform 5:2.


32

Liu SW, Zhang QX, Chen JD, Han ZX, Liu X, Feng L, Shao YH, Rong JG, Kong XG, Tong GZ (2006) Genetic diversity of avian infectious bronchitis coronavirus strains isolated in China between 1995 and 2004. Arch Virol 151:1133-1148.

Philipp H-C, Voss M (2006) Infectious bronchitis virus infections in german chicken flocks - Recent field observations. In: Heffels-Redmann U, Kaleta EF (eds) V International Symposium on Avian Corona- and Pneumoviruses and Complicating Pathogens, pp 154-156. Rauischholzhausen, Germany: VVB Laufersweiler Verlag.

Wit JJd, Nieuwenhuizen J, Fabri THF (2006) Protection by maternally derived antibodies and vaccination at day of hatch against early challenge with IBV serotype D388. In: Heffels-Redmann U, Kaleta EF (eds) V International Symposium on Avian Corona- and Pneumoviruses and Complicating Pathogens, pp 314-318. Rauischholzhausen, Germany: VVB Laufersweiler Verlag.

Worthington KJ, Jones RC (2006a) New genotype of infectious bronchitis virus in chickens in Scotland. Vet Rec 159:291-292.

Worthington KJ, Jones RC (2006b) An update of the european RT-PCR IBV survey and recent findings on a novel IBV genotype. In: Heffels-Redmann U, Kaleta EF (eds) V International Symposium on Avian Corona- and Pneumoviruses and Complicating Pathogens, pp 176-188. Rauischholzhausen, Germany: VVB Laufersweiler Verlag.

Table 1. Epidemiological information of Spanish QX-like IBV isolates included in the study. Isolates are sorted by year of isolation.

ISOLATE ISOLATION YEAR

TYPE OF BIRD AGE

TISSUES FOR

ISOLATION

GENBANK ACCESSION

NUMBER IBV/La/SP/17/

08 2008 Layer pullets 20 d Kidney GQ253482

IBV/Ck/SP/18/08 2008 Slow growth

chickens 22 d Kidney GQ253483

IBV/Ck/SP/79/08 2008 Broiler

chickens 41 d Trachea, Tonsil GQ253484

IBV/La/SP/116/09 2009 Layer hens 33 w Tonsil GQ253485

IBV/Ck/SP/170/09 2009 Broiler

chickens 10 d Trachea GQ253486


33

Figure 1. Neighbor-Joining with 1000 bootstraps replicates phylogenetic tree constructed with 650 pb partial S1 gene sequences. Distances were estimated by the Kimura-2 parameter method. The length of the bar indicates number of substitutions per site. In colors, Spanish field viruses included in the study.

IBV/SP/31-2/08 IBV/SP/59-9/08 IBV/SP/59-4/08 IBV/SP/29-2/08 IBV/SP/45-2/08 IBV/SP/03/08 IBV/SP/138/09 IBV/SP/45-1/08 IBV/SP/163/09 IBV/SP/165/09 IBV/SP/146/09 IBV/SP/135/09 IBV/SP/139/09

IBV/SPl/119/09 IBV/SP/94-4/08 IBV/SP/155/09 IBV/SP/90-10/08 IBV/SP/126/09 IBV/SP/127/09

IBV/SP/122/09 IBV/SP/102/08 IBV/SP/156/09

Spain/04/221 Spain/00/337 Italy-02

Spain/99/316 UK-L633-04 It-497-02 4/91-pathogenic Spain/00/336

UK/3/91 FR-85131-85 IR-3654-VM Spain/92/35

D3896 D207 D274

GRAY JMK

HOLTE ARK99

CU-T2 B1648

BEAUDETTE H120 M41 Spain/96/334

IBVQ CK/CH/LSHH/03I

A2 LX4

CK/CH/LHLJ/04V HBN-DQ070837

CK/CH/LLN/98I UK/AV2150/07 IBV/La/SP/17/08 IBV/Ck/SP/18/08 IBV/Ck/SP/79/08 IBV/La/SP/116/09 IBV/Ck/SP/170/09 IS/1201

L1148 K1019/03 NL-L1449K-04 FR-L1450L-05

IBV/La/114/09 V1397

D1466

95 9

9

99 5

4 99

70

39

41

99

86

81 65 53

54 57

14 41

49

27 46

51

62

99

92 49

97

99

98

78 5

7 87

80

72

29

26

23 1

8 24 26

49

36

32

99

47 9

9

99

76

49

70

20

29

16

99

0.05

ITALY 02 GENOTYPE

QXIBV GENOTYPE

D1466 GENOTYPE


34

MOLECULAR SURVEY OF INFECTIOUS BRONCHITIS VIRUS IN EUROPE IN 2008

MONNE I, DRAGO A, FASOLATO M, CAPUA I and CATTOLI G

OIE/FAO and National Reference Laboratory for Newcastle Disease and Avian Influenza, Istituto Zooprofilattico Sperimentale delle Venezie, Viale dell’Università, 10,

35020 Legnaro, Padova, Italy

SUMMARY Molecular based survey of Infectious bronchitis viruses (IBVs) in Europe in 2008 was undertaken. More than 600 field samples collected from 10 distinct European countries were tested by a validated Real Time RT-PCR protocol. Two hundred and seventy-five IBV positive samples were further characterised by sequencing and phylogenetic analysis of the S1 gene. Co-circulation of several distinct genotypes was recognised in all the European countries involved in this monitoring activity. INTRODUCTION IBV is characterized by high frequency of appearance of new variants and the actual number of existent virus genotypes throughout the world is unknown. The continuing appearance of new IBV genotypes renders difficult to elaborate a definitive classification system and poses serious challenges for the control of the disease through vaccination. Previous works (Capua et al., 1999) have demonstrated that distinct serotypes can co-circulate in a region. Considering the high rate of evolution of this virus and its widespread, the identification of the circulating IBV field strains is extremely important for the selection of the more appropriate vaccination programme. The aims of this study were to identify the IBV strains circulating in European poultry flocks in 2008 and to recognise the possible emergence of new IBV genotypes. MATERIALS and METHODS 617 field samples (oropharyngeal/tracheal swabs and organs) have been collected in 10 European countries (United Kingdom, Belgium, France, Greece, The Netherlands, Italy,


35

Germany, Czech Republic, Ukraine) from poultry flocks showing clinical signs thought to be associated to IBV infection. Samples were screened by Real Time PCR targeting the conserved 5’-untranslated region of the viral genome (Callison et al., 2006). RNA of the samples tested positive by rRT-PCR was amplified by RT nested-PCR (Worthington et al., 2008) targeting the S1 spike gene and sequenced in order to distinguish the genotypes circulating in the monitored poultry farms. Since June 2008, types specific primers were used to detect the D1466 viruses (Cavanagh et al., 1999). RESULTS 271 samples were found to be positive for IBV. Different proportion of each known IBV genotype was recognised analysing the S1 sequences obtained in the framework of this survey. The 793B genotype followed by the Massachussets and D274 types were recognised as the most common European IBV genotypes. 84% of the viruses clustering within these genotypes possess 100% nucleotide identity with the sequences of vaccine strains. QX like IBV was identified in flocks of broilers and broiler breeders in six countries. Genotype 624/I, not detected in Western Europe during a previous survey in the period 2002-2006 (Worthington et al, 2008), was identified in the United Kingdom (UK) and in Romania. Italy02 genotype was identified only in Southern Europe (i.e. Greece and Italy). Identification of Arkansas genotype was mostly restricted to flocks using vaccine combination containing this IBV variant. The 2008 IBV monitoring did not detect any B1648 and D1466 variants. However, the type specific PCR for D1466 was applied starting from the second half of 2008. Two IBVs sharing nucleotide similarity ranging between 92 and 96 % with the Dutch V1397 IBV type were recognised in the UK and The Netherlands. The S1 sequences of these viruses were not closely related to each other, displaying a percentage of homology lower than 92%. DISCUSSION The results of this survey demonstrated that the 793B, Massachussets and D274 type IBVs were the most frequently detected variants. Based on the epidemiological data available and on the results of the sequence analysis, it may be suggested that a large percentage of these viruses were probably re-identifications of the vaccine viruses. However, the analyses applied for the present investigation do not allow to conclusively distinguish between vaccine and field viruses. Over the period examined, the QX, Italy02 and 624/I types were found to be the dominant field strains in several European countries covered by this survey. The European QX-type IBV was detected in Italy, Germany, Belgium, France, The Netherlands and Czech Republic from flocks of broilers and broiler breeders showing respiratory signs (56%), impaired growth (22%), enteritis (11%) and nephritis (11%). No QX-type was detected in the United Kingdom during this monitoring activity. Based on this data and on the results of a previous surveillance programme in the UK (Worthington et al, 2008), it may be suggested that the European QX variant did not spread in this country up to the end of 2008. In the present study, we


36

first detected the 624-I type in UK and Romania mostly from flocks of layers with drop in eggs production and from broilers showing increased mortality and poor growth. This IBV type was previously described in Italy (Capua et al 1994), in Poland (Capua et al., 1999) and in Russia (Bochkov et al., 2006). Further epidemiological and molecular investigations will be necessary to clarify the way of introduction of the 624/I type in these countries. The survey here described identified Italy02 type only in Italy and Greece. A survey carried out in Europe in the period 2002 -2006 (Worthington et al, 2008) showed that Italy02 was the third most common genotype in Western Europe. Since 2004, its prevalence was found to be declining in UK, France, Germany, The Netherlands and Belgium but not in Spain where an increasing number of Italy02 strains was identified. Our 2008 survey showed that the Italy02 remains a common finding in the Mediterranean countries covered by this survey and it is unclear the reason explaining the disappearance of this genotype from the remaining Western European countries. It has been demonstrated that protection against this specific field IBV type can be obtained using vaccine combination (Jones et al., 2005; Worthington, 2008). It may be that the wide use of this vaccine strategy has resulted in a progressive decrease of the Italy02 type circulation. Two viruses sharing the highest similarity with the V1397 variant (percentage of homology ranged between 92 and 96%) were identified. The V1397 variant was previously described in the Netherlands during the 80s. In the present study, V1397-like viruses were detected in samples collected in the Netherlands and in the UK respectively in September from layers with respiratory signs and October 2008 from broiler breeders with drop in eggs production. The low similarity existing between the sequences of the two 2008 V1397 variants suggests that they originated from a common progenitor but then they evolved separately in distinct environments. In conclusion, this study provides an update on the major IBV strains circulating in Europe and has shown the strategic role of the molecular survey to monitor the circulation of distinct IBV strains and to evaluate possible adjustments to existing vaccination programme. However, more extensive surveillance programmes should be undertaken in order to better elucidate the true distribution and epidemiology of the distinct IBV genotypes in Europe. REFERENCES Bochkov et al., 2006. Molecular epizootiology of avian infectious bronchitis in Russia.

Avian Pathol. 35(5):379-93. Callison et al., 2006. Development and evaluation of a real-time Taqman RT-PCR

assay for the detection of infectious bronchitis virus from infected chickens. J Virol Methods.;138(1-2):60-5.

Capua I et al.,1994. A 'novel' infectious bronchitis strain infecting broiler chickens in Italy. Zentralbl Veterinarmed B. 41(2):83-9.

Capua I, et al., 1999. Co-circulation of four types of infectious bronchitis virus (793/B, 624/I, B1648 and Massachusetts) Avian Pathology, 28, 587 – 592.


37

Cavanagh et al., 1999. Longitudinal field studies of infectious bronchitis virus and avian pneumovirus in broilers using type-specific polymerase chain reactions. Avian Pathology, 28, 593-605.

Jones et al., 2005. Efficacy of live infectious bronchitis vaccines against a novel European genotype, Italy 02. Veterinary Record, 156, 646-647.

Kusters et al., 1989. Phylogeny of antigenic variants of avian coronavirus IBV. Virology.;169(1):217-21.

Scott et al. (2006). Development and evaluation of a real-time Taqman RT-PCR assay for the detection of infectious bronchitis virus from infected chickens. JVM. 138, 60-65.

Worthington et al., (2008). A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. Avian Pathology 37, 247-257.

ACKNOWLEDGEMENTS The authors wish to thank Fort Dodge Animal Health (Huizerstraatweg 117, 1411 GM Naarden, The Netherlands) and Intervet Schering Plough Animal Health (Postbus 31 5830 AA Boxmeer, The Netherlands) for providing field samples which were essential for the realization of the present study.


38

FALSE LAYERS IB-CASE REPORT IN BROILER BREEDERS

BLOCK H

Gruppenpraxis Meyer-Block/ Thien, Am Rott 12, 49843 Uelsen, Germany SUMMARY This case report demonstrates the enormous financial consequences of very early IBV infections that cause false layers. INTRODUCTION Our vet practise is located in NorthWest Germany close to the Dutch border. In this area there is about 20% of the poultry production in Germany (mainly broiler and breeder) MATERIALS and METHODS Flock On the farm there were 80.000 ROSS 708 breeder birds of two different ages. The birds had been reared in Holland and had been moved to this farm at an age of 20 weeks. The IBV vaccination program followed the common recommendation out of the literature (H120/D274 at day of hatch, 4/91 at 10 days, IB Ma 5 at 10 weeks, IB multi at 14 weeks and IB vac 2 (H52) at 18 weeks). Figure 1 shows in red the standard production curve. The green and the black curves show the actual data from house 1 and 2. Houses 3-5 showed normal production parameters. Until week 26 there were no clinical signs for a disease.


39

RESULTS Post mortem At week 26 we collected the birds from these two houses. At the post mortem examination it showed that the oviducts had changed into big cysts. RT-PCR IBV and VNT The detection of IBV QX-virus through PCR (at 26 weeks of age) was negative. The virus neutralisation test showed a high level of antibodies against the D388 serotype. DISCUSSION Economically, there were two possibilities for the flocks of the two houses:

1. To cull all false layers (which we have done two times before in similar cases) 2. To slaughter the flock

After long discussion about the pro and cons of both possibilities we decided to slaughter 20.000 breeders at the age of 28 weeks. The costs have been round about 500.000 €. In our 10 cases that we had so far the origin of the hatching eggs was always the USA. In our experience, importing eggs with low or no MDA against D388 is a risk factor for getting false layers due to very early D388 infections.

laying curve

0

10

20

30

40

50

60

70

80

90

23 24 25 26 27 28 29 30 31 32 33

age in weeks

egg

prod

uctio

n in

%

standardhouse 1house 2


40

INFECTIOUS BRONCHITIS VIRUS in JORDAN: MOLECULAR SUBTYPE

ALROUSSAN DA, TOTANJI WS and KHAWALDEH GY

Provimi Jordan, P.O. Box 499, Amman 11118, Jordan

SUMMARY Infectious bronchitis virus (IBV) causes respiratory disease in chickens all over the world. Infectious bronchitis virus has many serotypes that do not confer cross protection against each other. The current study was designed to know which IBV types were circulating in Jordanian broiler chickens. Tracheal swabs from 175 broiler flocks at the acute phase of respiratory disease were collected. The swabs were subjected to RNA extraction and tested by reverse transcription PCR (RT-PCR). Specific-nested PCR were performed on RT-PCR products to detect and differentiate strains of Massachusetts, 4/91, and D274 types. The nucleic acid of IBV was detected in 105 out 174 (60%) broiler flocks by RT-PCR. Specific-nested PCR revealed that 35.2, 31.4, and 8.6% of these flocks had Massachusetts, 4/91, and D274, respectively, alone. In 24.8% of tested flocks, 2 types of IBV were detected. However, because the primers used in this study were designed specifically for 3 types of IBV, other types might have been present but not detected. Future work should include the isolation and molecular characterization of IBV in the region to adopt a suitable vaccination program using the common field serotypes as vaccines to protect against IBV caused disease. INTRODUCTION Infectious bronchitis (IB) is a major disease problem in the broiler industry. Many antigenic types of the causative agent, IB virus (IBV), exist, for example, the Massachusetts, D274, and 4/91 (also known as CR88 and 793/B) serotypes. The Massachusetts type was the first isolated in Europe in the 1940s (Cavanagh and Davis, 1992). The D274 type was the most common in several western European countries in the early and mid-1980s (Cook, 1984; Davelaar et al., 1984; Cavanagh et al., 1992). The 4/91 type was first identified in the United Kingdom in 1990/ 1991 (Gough et al., 1992; Parsons et al., 1992) but was retrospectively found to have been present in France since 1985 (Cavanagh et al., 1998). Cavanagh et al. (1999) detected the 4/91


41

type of IBV in swabs, collected in 1997 and 1998, from Saudi Arabia, Japan, Sweden, Denmark, Poland, Italy, France, and Argentina. Antibodies reactive with the 4/91 type have been detected in chickens in Thailand, Mexico, Greece, Britain, France, the Netherlands, Spain, and Germany (Cook et al., 1996).

In Jordan, chicken production is the most developed industry in the animal sector. There are serious respiratory diseases of unknown etiology that have caused catastrophic economic losses to farmers in the country. There is limited evidence in literature describing the prevalence of poultry respiratory diseases in Jordan. One report (Saad, 2006) describes the serotype of IBV (Massachusetts, Arkansas, Delaware variant 072, and JMK in poultry flocks in Jordan based on hemagglutination inhibition test and demonstrates the exposure of these flocks to Arkansas, Delaware variant 072, as well as Massachusetts-like serotypes of IBV. Therefore, this study was designed to know which IBV strains were circulating in Jordanian broiler chickens using type-specific reverse transcription PCR (RT-PCR) for Massachusetts, as well as other IBV strains (D274 and 4/91) not previously documented in Jordan. MATERIALS and METHODS Broiler Flocks During the period from September 2005 to November 2007, we examined 175 commercial broiler flocks from northern, southern, and central Jordan in which the chickens were suffering from respiratory disease. Five sterile swabs (Heinz Herenz Medizinalbedarf GmbH, Hamburg, Germany) were taken from 5 chickens from each flock at the acute phase of respiratory disease and sent to the Provimi Jordan laboratory where they were stored at 4°C until RNA was extracted. Vaccinated history included all flocks vaccinated by spray at 4 d of age against the M-41 strain of IBV (Intervet, Wim de Ko¨ rverstraat, AN Boxmeer, the Netherlands). In the majority of these flocks, signs of respiratory disease usually appeared at 33 to 35 d of age. Chickens suffered from severe gasping, coughing, conjunctivitis, nasal and ocular discharge, depression, weakness, and were reluctant to move. Gross lesions observed in these flocks included a moderate to severe congestion of trachea with or without mucopurelent exudates, airsaculitis, and pericarditis or perihepatitis. RNA Extraction Swabs from each flock were placed in 1,000 μl of PBS (pH 7.2) and were scraped on the side of the tube to facilitate removal of contents from the swab head. Extraction of RNA was performed on 60 μl of the pooled material for swab from each flock, using a Purescript RNA purification kit (Gentra Systems, Minneapolis, MN) according to the procedure of the manufacturer.


42

RT-PCR and Nested PCR The RT-PCR reaction was performed using one-step RT-PCR, an Access RT-PCR System kit (Promega Corp., Madison, WI), and primers XCE2− and XCE2+ (Alpha DNA, Montreal, Quebec, Canada; Table 1) according to the procedure of the manufacturer. Reverse transcription PCR was carried out in a DNA Engine thermal cycler (BioRad Laboratories Ltd., Mississauga, Ontario, Canada) for 1 reverse transcription cycle of 60 min at 45°C, followed by 94°C for 5 min, then 40 PCR cycles of 94°C for 45 s, 57°C for 45 s, and 72°C for 90 s, with a final extension cycle at 72°C for 5 min. Reverse transcription PCR produces a 466-bp fragment common to all IBV (Figure 1) that was used in 3 specific nested PCR with oligonucleotide XCE3−, which was designed to hybridize to RNA from all 3 strains, and oligonucleotides MCE1+, DCE1+, and BCE1+ (Table 1) that are specific for types Massachusetts, D274, and 4/91, respectively, and generates 295, 217, and 154-bp fragments, respectively (Figure 1). Nested PCR reaction contained 0.5 μl of RT-PCR product of positive reactions, 0.5 μl of Taq DNA polymerase (5 units/μl ; (Promega Corp.), 2 μl of deoxynucleoside triphosphate mix (10 mM; Promega Corp.), 5 μl of 10× PCR buffer (Promega Corp.), and 1 μl of each of the oligonucleotides MCE1+, DCE1+, and BCE1+ (50 Pmol/_l; Alpha DNA). A total reaction volume of 50 μl was obtained by adding nuclease-free water. The nested PCR were performed using the following conditions: 94°C for 1 min, 48°C for 2 min, and 72°C for 90 s, 35 cycles followed by a final extension cycle of 72°C for 10 min. The IBV (M41) antigen [Gezondheidsdienst voor Dieren B.V. (GD), Animal Health Service Deventer, the Netherlands] was used as a positive control for RNA extraction and RT-PCR. Negative control (nuclease-free water) was also used in each run. Agarose Gel Electrophoresis The RT-PCR and nested PCR products were electrophoresed on a 2% agarose gel in Tris-acetate-EDTA buffer (40 mM of Tris and 2 mM of EDTA, with a pH value of 8.0) containing ethidium bromide (Promega Corp.) for 45 min at 100 V and visualized under ultraviolet light (AlphaImager; Alphainnotech, San Leandro, CA). RESULTS All of the tested broiler flocks (175) that were received had a history of respiratory disease. The IBV were detected by RT-PCR in 105 out 175 (60%) of these flocks. Specific-nested PCR were performed on RT-PCR-positive products (105) to detect and differentiate strains of the Massachusetts, 4/91, and D274 types. The results of specific- nested PCR (Table 2) revealed that 35.2, 31.4, and 8.6% of these flocks had Massachusetts, 4/91, and D274, respectively, alone. On the other hand, 4.8, 7.6, and 12.4% of these flocks were infected with both Massachusetts + D274, D274 + 4/91, and Massachusetts + 4/91, respectively. The overall results of specific-nested PCR (Table 2) indicated the IBV Massachusetts and 4/91 types were found to be most prevalent (52.4 and 51.4%, respectively), whereas the IBV D274 type was found to be of low prevalence (21%). Figure 1 shows RT-PCR and nested PCR for IBV detection and subtyping (Massachusetts, 4/91, and D274).


43

DISCUSSION Current diagnosis of IB is commonly based on virus isolation in embryonating eggs, followed by immunological identification of the isolates. This procedure is time consuming and requires the use of specific polyclonal or monoclonal antibodies. Moreover, some isolates could be mixtures of different types of IBV that can confuse the interpretation of serotyping results. Reverse transcription PCR has been described previously using IBV, RNA extracted from allantoic fluid, and tracheal swabs. These techniques had been shown to be very efficient for the detection of IBV and for the identification of IBV types (Cavanagh et al., 1999; Handberg et al., 1999). In this study, we performed RT-PCR on the tracheal swabs from birds with a history of respiratory disease. Furthermore, the RT-PCR-positive reactions were subjected to specific-nested PCR to detect and differentiate strains of the Massachusetts, 4/91, and D274 types. The IBV had been detected in 60% of broiler flocks by RT-PCR. Nested PCR results indicated that the main IBV types circulating in the Jordan broiler population were the Massachusetts type and 4/91 type, which represent 52.4 and 51.4%, respectively, followed by D274, which represents 21% of tested flocks (Table 2). However, because the primers were selected specifically for these 3 types of IBV, other types might have been present but not detected. Massachusetts-type IBV was detected in 52.4% of the broilers tested. In a recent study, Cavanagh et al. (1999) reported that when Massachusetts-type IB vaccines were applied at 1 d old in the hatchery, vaccine virus could later be detected in all broiler flocks tested by RT-PCR on swabs, with maximal amounts during the first week of life. In our work, most of the Massachusetts type detected was from broilers showing respiratory signs beyond 4 wk of age. Our results suggest field exposure of these flocks to Massachusetts type alone or in combination with other type of IBV.

Twenty-six (24.8%) of tested flocks were infected with 2 types of IBV (Table 2). This is in agreement with previous observations showing that broiler flocks may be infected simultaneously with several types of IBV (Cavanagh et al., 1999). The results in a current study indicate a relatively high prevalence of 2 types of IBV (i.e., Massachusetts and 4/91 types), in addition to a low prevalence of the D274 type. This study and previous ones reported suggested that there is a possibility of the presence of several IBV stains in Jordan.

Like in many other parts of the world, Massachusettstype vaccines are the only officially authorized vaccines in Jordan. Despite the use of these IBV vaccines, it is common to find IB problems in vaccinated chickens. The results of this study may partially explain the failure of Massachusetts-type vaccines and necessitate revising the Jordanian vaccination program against IB. In this study, the sequence of PCR products was not determined, and therefore the origin of the isolates is not clear at present. In conclusion, by utilizing such diagnostic techniques, it is possible to conduct a detailed epidemiological study to determine the full economic effect of this disease. Future work should include the isolation, serotyping, and molecular characterization of IBV in the


44

region to adopt a suitable vaccination program using the common field serotypes as vaccines to protect against IBV-caused disease.

REFERENCES Adzhar, A., R. E. Gough, D. Haydon, K. Shaw, P. Britton, and D. Cavanagh. 1997.

Molecular analysis of the 793/B serotype of infectious bronchitis virus in Great Britain. Avian Pathol. 26: 625–640.

Cavanagh, D., and P. J. Davis. 1992. Sequence analysis of strains of avian infectious bronchitis coronavirus isolated during the 1960s in the UK. Arch. Virol. 130: 471– 476.

Cavanagh, D., K. Mawditt, P. Britton, and C. J. Naylor. 1999. Longitudinal field studies of infectious bronchitis virus and avian pneumovirus in broilers using type-specific polymerase chain reactions. Avian Pathol. 28: 593-605.

Cavanagh, D., P. J. Davis, J. K. A. Cook, D. Li, A. Kant, and G. Koch. 1992. Location of the amino-acid differences in the S1 spike glycoprotein subunit of closely related serotypes of infectiousbronchitis virus. Avian Pathol. 21: 33–43.

Cavanagh, D., K. Mawditt, R. Gough, J. P. Picault, and Britton. 1998. Sequence analysis of strains of the 793/B genotype (CR88, 4/91) of IBV isolated between 1985 and 1997. In E.F. Kaleta & U. Heffels-Redmann (Eds.) Proceedings of an International Symposium on Infectious Bronchitis and Pneumovirus Infections in Poultry (pp. 252–256). Giessen: Justus Liebig University.

Cook, J. K. A. 1984. The classification of new serotypes of infectious bronchitis virus isolated from poultry flocks in Great Britain between 1981 and 1983. Avian Pathol.13: 733–741.

Cook, J. K. A., S. J. Orbell, M. A. Woods, and M. B. Huggins. 1996. A survey of the presence of a new infectious bronchitis virus designated 4/91 (793B). Vet. Rec. 13: 178–180.

Davelaar, F. G., B. Kouwenhoven, and A. G. Burger. 1984. Occurrence and significance of infectious bronchitis virus variant strains in egg and broiler production in The Netherlands. Vet Q. 6: 114–120.

Handberg, K. J., O. L. Nielsen, M. W. Pedersen, and P. H. Jorgensen. 1999. Detection and strain differentiation of infectious bronchitis virus in tracheal tissues from experimentally infected chickens by reverse transcription polymerase chain reaction. Comparison with an immunohistochemical technique. Avian Pathol. 28: 327– 335.

Gough, R. E., C. J. Randall, M. Dagless, D. J. Alexander, W. J. Cox, and D. Pearson, 1992. A “new” strain of infectious bronchitis virus infecting domestic fowl in Great Britain. Vet.Rec. 130: 493-494.

Parsons, D., M. M. Ellis, D. Cavanagh, and J. K. A. Cook. 1992. Characterisation of an avian infectious bronchitis virus isolated from IB-vaccinated broiler breeder flocks. Vet. Rec. 131: 408–411.

Saad, M. G. 2006. Infectious bronchitis virus serotypes in poultry flocks in Jordan. Pre. Vet. Med. 78: 317-324.

VI.

INT.

SYM

PO

SIU

M O

N A

VIA

N C

OR

ON

A-

AN

D P

NE

UM

OV

IRU

SE

S, R

AU

ISC

HH

OLZ

HA

US

EN

, GE

RM

AN

Y, 2

009

45

Tabl

e 1:

Seq

uenc

es o

f olig

onuc

leot

ide

used

in re

vers

e-tra

nscr

ipta

se (R

T-P

CR

) and

nes

ted

PC

R

O

ligon

ucle

otid

ea Se

quen

ces

Spec

ifici

ty

Ban

d si

ze (b

p)

Ref

eren

ces

RT-

PCR

5’

-CA

CTG

GTA

ATTT

TTC

AGAT

GG

-3’

Adz

har e

t al.,

199

7 X

CE

2 +

XC

E2

– 5’

-CC

TCTA

TAA

ACA

CC

CTT

GC

A-3’

U

nive

rsal

46

6 A

dzha

r et a

l., 1

997

Nes

ted

PCR

X

CE

3 –

5’-C

AGAT

TGC

TTAC

AAC

CAC

C-3

’

A

dzha

r et a

l., 1

997

DC

E1 +

5’

-TTC

CAA

TTAT

ATC

AAA

CC

AGC

-3’

D27

4 21

7 A

dzha

r et a

l., 1

997

MC

E 1 +

5’

-AA

TAC

TAC

TTTT

AC

GTT

ACA

C-3

’ M

assa

chus

etts

29

5 A

dzha

r et a

l., 1

997

BC

E1

+ 5’

-AG

TAG

TTTT

GTG

TATA

AA

CC

A-3

’ 4/

91

154

Adz

har e

t al.,

199

7 a +

= G

e nom

e se

nse;

– =

ant

i-gen

ome

sens

e Ta

ble

2: In

fect

ious

bro

nchi

tis v

irus

(IBV

) typ

es a

mon

g 10

5 br

oile

r flo

cks

that

wer

e po

sitiv

e by

reve

rse-

trans

crip

tase

po

lym

eras

e ch

ain

reac

tion

(RT-

PC

R)1

IB

V ty

pe

No

of IB

V ty

pe (%

) M

assa

chus

etts

37

(35.

2)

Mas

sach

uset

ts +

4/9

1 13

(12.

4)

Mas

sach

uset

ts +

D27

4 5

(4.8

) To

t al M

assa

chus

etts

55

(52.

4)

4/91

+ D

274

8 (7

.6)

4/91

33

(31.

4)

Tota

l 4/9

1 54

(51.

4)

D27

4 9

( 8.6

) To

t al D

274

22 (2

1)

1 One

hun

dred

and

sev

enty

five

floc

ks h

ad re

spira

tory

dis

ease

and

wer

e te

sted

by

RT-

PC

R a

nd 1

05 o

f the

se fl

ocks

(60%

) w

ere

posi

tive.

__VI. INT. SYMPOSIUM ON AVIAN CORONA- AND PNEUMOVIRUSES, RAUISCHHOLZHAUSEN, GERMANY, 2009_


46

Figure 1. Reverse-transcriptase PCR and nested PCR for infectious bronchitis virus (IBV) detection and subtyping. Lanes M = 100-bp DNA ladder marker (Promega Corp., Madison, WI). Lane 1 = negative control for IBV. Lane 2 = XCE2+/ XCE2– primers set for general detection of IBV (positive; band at 466 bp). Lane 3 = Massachusetts type of IBV (positive; band at 295 bp). Lane 4 = D274 type of IBV (positive; band at 217 bp). Lane 5 = 4/91 type of IBV (positive; band at 154 bp).


47

RECOMBINATION, POINT MUTATIONS AND POSITIVE SELECTION ON THE BASIS OF THE MOLECULAR DIVERSITY OF BRAZILIAN STRAINS OF AVIAN

INFECTIOUS BRONCHITIS VIRUS

BRANDAO PE1, 4, SANDRI TL1, 4, SOUZA SP1, 4, KUANA SL3, RICHTZENHAIN LJ1,4 and VILLARREAL LYB2,4

1Department of Preventive Veterinary Medicine and Animal Health, College of Veterinary Medicine, University of São Paulo, Av. Prof. Dr. Orlando M. Paiva, 87,

CEP 05508-270, Sao Paulo, SP, Brazil 2Intervet Schering Plough Animal Health, Av. Sir Henry Wellcome, 335, CEP 06741-

050, Cotia, SP, Brazil 3Perdigão Agroindustrial, Brazil

4Coronavirus Research Group, Av. Prof. Dr. Orlando M. Paiva, 87, CEP 05508-270, Sao Paulo, SP, Brazil

SUMMARY This article describes the molecular diversity of Infectious bronchitis virus (IBV) amongst breeders, layers and broilers in Brazilian flocks and the molecular evolutionary events involved. Multiple organs samples from 20 flocks were submitted to the partial amplification of S gene (nucleotides 726-1071) of IBV. Fifteen out of the 20 strains sequenced segregated in an exclusive major cluster, apart from the archetypical sero/genotypes, subdivided in three other clusters (Brazil 01, 02 and 03). While three strains could be classified as Massachusetts genotype, the remaining two strains, originated from flocks with reproductive and respiratory disorders grouped within the 4/91 genotype cluster. Positive dN-dS values were found amongst strains in both Brazilian clusters 02 and 03, as well as in one of the 3 Brazilian strains in the Massachusetts cluster. Recombination was found in all Brazil 03 strains (nucleotides 764 to 812 of S). These results indicate that the IBV strains included in this study were under positive selection, possibly due to the fixation of random point mutations favored by the large number of birds per flock and high population density that turned out to be advantageous as a consequence of poor cross protection amongst field and the vaccine strains based on the Massachusetts serotype massively used for decades in Brazil in monotype vaccines. The relevance of the findings to poultry industry is discussed.


48

INTRODUCTION Infectious bronchitis (IB) is a highly complex infectious disease of poultry, with worldwide distribution and a high economic burden with the involvement of multiple serotypes of the causative agent Avian infectious bronchitis virus (IBV) (Cavanagh, 2007). A wide range of disorders in breeders, layers and broilers has been associated to IBV infection, including respiratory disease (Liu et al., 2008a), nephritis (Abdel-Moneim et al., 2006), reproductive failures in both male and female (Raj & Jones, 1997; Villarreal et al., 2007a), enteric disease (Villarreal et al, 2007b) and myopathy (Gough et al., 1992). A major issue for the control of IB is the low cross-protection between distantly related serotypes and the use of vaccines thus modulates the outcome of signs and disease spread as a factor of the genetic distance amongst vaccine and field strains based on S gene identity (Gelb et al., 2005).In Brazil, the main vaccine control strategies for IB are based solely on the massive use of Massachusetts serotypes attenuated vaccines, but, despite the wide use of vaccination, the disease still occurs at high frequency concurrently with the occurrence of antigenic groups divergent from the ones known worldwide (Di Fabio et al., 2000). The aims of this survey was to assess the molecular diversity of IBV amongst breeders, layers and broilers in Brazilian flocks and the molecular evolutionary events that drive such diversity for a more comprehensive knowledge of the IB situation at the field level. MATERIALS AND METHODS Source of viruses: A total of 20 flocks, including breeders, layers and broilers from the Southern, South-Eastern, Northeastern and Central-Western regions of Brazil were included in this study (Table 1). These regions represent the major Brazilian poultry regions, with high avian population density and flocks/farm clustering and frequencies of IBV-positive flocks ranging from 50 to 100%.Sampled flocks included birds presenting disorders of the enteric, respiratory, reproductive and renal tracts, already associated to IBV. Samples of lungs, tracheas, kidneys, reproductive organs and complete enteric contents were collected from each flock as organ-specific pools (3-5 birds/pool) between 2007 and 2008 and sent frozen to the laboratory. All birds had been vaccinated against IBV with vaccination schedules including attenuated Massachusetts vaccines (broilers) and attenuated plus killed Massachusetts vaccines (layer chickens and breeders). Also included in the study were two attenuated Massachusetts commercial vaccines from different manufacturers continuously used in the sampled flocks, named as Mass Vaccine 01 and Mass Vaccine 02. IBV screening and partial amplification of the spike gene: Pools were prepared as 50% (v/v) suspensions in DEPC-treated water and submitted to 3 freezing/thawing cycles in liquid nitrogen and 56C dry bath and clarified at 5,000g/15 minutes/4C. Total RNA was extracted from the supernatants with TRIzol™ reagent (Invitrogen, Carlsbad, CA, USA) following manufacture’s instructions. Each pool was surveyed for the presence of IBV by a reverse-transcription semi-nested PCR using primers targeted to a region of the 3’UTR, highly conserved amongst IBV genotypes, as described by Cavanagh et al. (2002). Partial reverse-transcription and amplification


49

of the spike gene of IBV was carried out as described by Worthington et al. (2008), resulting in amplicons of 390bp between nucleotides 705 and 1094 of the S1-coding region (regarding strain UK/7/93, Genbank Accession number Z83979). For this S1 RT-PCR, only one pool per flock was selected amongst those positive for the 3’UTR RT-PCR, preferentially the pool most directly related to the signs observed in a given flock (Table 1). Partial S gene analysis: The S1 390bp amplicons were purified from agarose gels using the GFX™ kit (GE Healthcare, Fairfield, CT, USA) and submitted to bi-directional DNA sequencing with Big Dye™ 3.1 (Applied Biosystems, Carlsbad, CA, USA) in a ABI-377 automatic sequencer (Applied Biosystems, Carlsbad, California USA). Sequences with PHRED scores higher than 20 (Ewing & Green, 1998) were assembled with Cap-Contig application and aligned with CLUSTALW included in Bioedit 7.0.9.0 software (Hall, 1999) with homologous sequences retrieved from the Genbank (accession numbers in Figure 1). A genealogic tree was built with the Neighbor-Joining distance algorithm and the Maximum Composite Likelihood substitution model with 1000 bootstrap replicates using MEGA 4 (Tamura et al., 2007). For rooting the tree, a bredavirus S gene sequence was used as an outgroup (Genbank accession number NC007447). Bootstrap values higher than 90 were considered as evidence of clustering. The difference between non-synonymous and synonymous substitutions (dN-dS) was calculated with the pairwise Nei-Gojobori/JC method using MEGA 4 (Tamura et al., 2007), being a positive dN-dS value considered as evidence for positive selection. The alignment was also used for the inference of recombination amongst the studied strains using the RDP, GENECONV, Bootscan, MaxChi, Chimaera, SiScan, Phylpro and LARD methods with window size=10 and highest p=0.05 with the software RDP3Beta34 (Martin et al., 2005). Accession numbers: Partial S1 sequences of the 20 field strains of IBV have been assigned Genbank accession numbers FJ791254 to FJ791273 (Table 1), while Mass Vaccine 01 and 02 received the accession numbers FJ791274 and FJ791275, respectively. RESULTS IBV screening and partial amplification of the spike gene: All the 20 flocks included in this study showed at least one organ pool positive for the RT-PCR to the 3’UTR, allowing the following amplification of the partial S1-coding region from the pools most related to each class of sign observed in these flocks, except for the breeders flock in which strainIBV/BRAZIL/2007/USP-32 was detected, which presented reproductive disorders by the time of sample collection (Table 1), but from which only the pool of enteric content was positive for IBV screening, which was then assumed as representative of the IBV population present in this flock for the purpose of S gene analysis. Genealogic analysis: Fifteen out of the 20 IBV strains detected segregated in a cluster unique to the Brazilian strains based on partial S1 sequences; from this major


50

cluster, other 3 subclusters can be depicted (Figure 1) with bootstrap values higher than 90, named Brazil 01 to 03. Amongst these 15 strains, no geographic, chronological or pathogenic pattern could be predicted from the tree topology. Unexpectedly, 2 strains (IBV/BRAZIL/2008/USP-31 and IBV/BRAZIL/2007/USP-32) from layers and breeders with respiratory and reproductive disorders were classified as belonging to the genotype 4/91, previously unknown amongst Brazilian poultry. The remaining 3 strains (IBV/BRAZIL/2008/USP-13, IBV/BRAZIL/2007/USP-14 and IBV/BRAZIL/2008/USP-15) detected in broilers and layers with respiratory, reproductive and enteric disorders, were classified as Massachusetts strains. Regarding the 3 Brazilian clusters, the most distant related were Brazil 01 and 03, with nucleotide and amino acid identities of 93.0 and 91.6 %, respectively (Table 2). Concerning the archetypical genotypes included in the analysis, the highest nucleotide/ amino acid identities for Brazil 01, 02 and 03 were related to the genotypes Massachusetts/ Connecticut, Arkansas/ D274 and 4/91/4/91, respectively (Table 2). For the three Brazilian strains typed as Massachusetts, IBV/BRAZIL/2008/USP-15 had 100% nucleotide identity with Mass Vaccine 01 and 97.9% with Mass Vaccine 02, while for the other 2 (IBV/BRAZIL/2008/USP-13 and IBV/BRAZIL/2008/USP-14) the nucleotide identity with Mass Vaccine 01 and 02 were 99.7/97.6 and 99.1/97%, respectively. Recombination analysis: Strain IBV/BRAZIL/2007/USP-18 in cluster Brazil 03 was identified as originated from recombination by the GENECONV method, with the major parent identified as strain IBV/BRAZIL/2008/USP-19 and nucleotides located at positions 764 to 812 of the spike gene (regarding strain UK/7/93, Genbank Accession number Z83979) as originated from a minor parent not found amongst the strains included in the analysis. Strains IBV/BRAZIL/2007/USP-17 and IBV/BRAZIL/2007/USP-16, also located in cluster Brazil 03 but in the most external nodes regarding the tree root, shared the same recombinant pattern as evidenced by the GENECONV method, except that in these the sequence of the minor parent was 1 nucleotide shorter (positions 764 to 811) and two point mutations when compared to strain IBV/BRAZIL/2007/USP-18 (A799T and C811T). BLAST/n analysis of the 48-49- nt recombinant sequences are homologous to strainsTP/64 dN-dS values: Positive pairwise dN-dS values of 0.004 were found between the following strains: IBV/BRAZIL/2008/USP-23 compared to IBV/BRAZIL/2008/USP-24, IBV/BRAZIL/2008/USP-21 and IBV/BRAZIL/2008/USP-22; IBV/BRAZIL/2008/USP-30 compared to IBV/BRAZIL/2008/USP-28 and IBV/BRAZIL/2008/USP-29; IBV/BRAZIL/2008/USP-15 compared to IBV/BRAZIL/2008/USP-13; and 0.008 for IBV/BRAZIL/2008/USP-24 compared to IBV/BRAZIL/2008/USP-21 and IBV/BRAZIL/2008/USP-22. DISCUSSION The existence of a major, characteristic Brazilian IBV genotype has already been described based on partial characterization of both the spike (Montassier et al., 2006; Villarreal et al., 2007b) and the N genes (Abreu et al., 2006). Nonetheless, the results obtained in the present investigation show that the genealogy of Brazilian strains of


51

IBV is much more complicated than previously known, as three consistent subclusters can be depicted in the topology of the tree in Figure 1. Despite the large geographic area the samples were collected from and the diverse range of pathologic features presented by the sampled flocks, the segregation of the 15 Brazilian genotype strains was independent of both these variables. This feature can be primarily attributed to the known role of the S1 subunit of the spike protein as a determinant of tropism rather than pathogenicity or virulence during IBV infection (Shen et al., 2004), resulting in IBV strains with high nucleotide/ amino acid identity to this region despite their pathological differences, as a result of high positive selection modulated not only by the immune response, but also by the cell membrane attachment factors to S1 known as heparan sulfate and syalic acid, to which the extended tropism of IBV has been associated (Madu et al., 2007). More accurate molecular markers for pathogenicity are the proteins 3a, implicated in IBV intracellular pathogeny (Liu et al., 2008b), or even the cytoplasmic tail of S, already associated to the efficiency of IBV replication (Youn et al., 2005), which could be useful targets to a more comprehensive understanding of the clinical features coupled to each of the strains described herein. It’s noteworthy that 2 strains typed as 4/91 have been collected from flocks presenting respiratory and reproductive disorders instead of chest muscle lesions classically associated to this type (Gough et al., 1992), in agreement to the wider tropism described to this genotype (Adzhar et al, 1997). Though already described worldwide (Cook et al., 1996; Seyfi Abad Shapouri et al., 2004; Xu et al., 2007; Roussan et al., 2008; Shimazaki et al., 2008; Worthington et al., 2008), this is a genotype previously not recognized in Brazilian poultry. Yet reasons for its emergence in this country can only be speculated at this moment, it could be due to the introduction of carriers during the importation of birds from countries where 4/91 is endemic or, as already suggested in cases of introduction of IBV types, to the role of migratory birds as sources of infection (Cavanagh, 2005; Liu et al., 2005). The most striking genetic feature found for the Brazilian genotype strains relates to the recombination that characterizes the cluster Brazil 03, shared by the three strains in this cluster starting at position 764 and ending at 812. The recombination event described in the present study has been determined as a derived from a major donor identified between a strain of the Brazilian genotype (IBV/BRAZIL/2008/USP-19) and a yet unidentified minor donor, without the involvement of any vaccine strain, allowing the speculation that other still undetected lineages might be found in the country. Natural recombination events in S1 with breakpoints at positions 82, 98, 100, 653, 1112 and 1460 of S, as well as intergenic S and N recombination have already been described as a result of the involvement of Arkansas and Massachusetts genotypes vaccines, resulting in increased genetic diversity of field strains of IBV (Wang et al., 1993; Bochkov et al., 2007). Nonetheless, similar patterns of natural recombination without the involvement of a vaccine strain have already been described (Chen et al., 2009) resulting in chimerical IBV genomes. It’s noteworthy that strains IBV/BRAZIL/2007/USP-16 and IBV/BRAZIL/2007/USP-17 share 100 identity in the recombinant region from the unknown origin (positions 764 to 811), emerging from a node derived from strain IBV/BRAZIL/2007/USP-18, the


52

major daughter in the recombination analysis, which, in it’s turn, differs from the first two in two nucleotide positions (799 and 811) in this area, which is also one-nucleotide longer. Taking into account the topology of the rooted tree in Figure 1, one can argue that strain IBV/BRAZIL/2007/USP-18 represents the most ancestral recombinant in the cluster Brazil 3 and that strains IBV/BRAZIL/2007/USP-17 and IBV/BRAZIL/2007/USP-18 harbors the chimerical S1 derivated from the first, which has now been fixated in Brazil 03. Regarding the comparison amongst the Brazilian genotype and the archetypical IBV genotypes, the fact that Massachusetts genotype was found as the closest to Brazil 01 only in terms of nucleotide sequences rather than amino acids, while this genotype was not the closest neither in terms of nucleotides nor amino acids for Brazil 02 and 03 (Table 02) is of high relevance as the region of S1 used for the analysis comprises sequences coding for conformational epitopes partially implicated in protection and virus neutralization (Ignjatovic & Sapats, 2005); taking into account that Massachusetts is the only genotype used for live vaccines in Brazil, a low protection can be predicted as a result of such vaccines in cases in which strains of the Brazilian genotype are involved. In fact, as already suggested (Lee et al. 2008), the massive use of monotype vaccines such as those based on H120 might allow the emergence of new IBV types as a result of strong immune pressure. A link between this suggestion and the epidemiological situation in the population under study is the discrete predominance of non-synonymous mutations in the S1-coding region under analysis for some strains of the Brazilian cluster, which might be interpreted as a trend for positive selection, possibly due to the fixation of random point mutations favored by the large number of birds per flock and high population density that turned out to be advantageous as a consequence of the above mentioned lower cross protection amongst field and the vaccine strains. Regarding the tree strains typed as Massachusetts, a vaccine origin could be assigned to strain IBV/BRAZIL/2008/USP-15, as the nucleotide identity between this strain and the one found in one of the commercial vaccines used in Brazil also included in this study (Mass Vaccine 01) was 100%. Nonetheless, for the two remaining strains (IBV/BRAZIL/2008/USP-13 and IBV/BRAZIL/2008/USP-14), this identity was bellow 100%, giving rise to the possibility that these are indeed wild field Massachusetts strains instead of vaccine strains. As for the relevance of the evolutionary patterns described herein at the field level, as all mutation events have been detected in a genetic region at least in part implicated in protection, it can be predicted that high endemic levels of IB will still occur in Brazil, with a continuous divergence of IBV lineages and increasing positively selected molecular and pathologic diversity. This knowledge is not only applicable for the suggestion of vaccine schedules but also to the proposition of evolutionary markers to the molecular epidemiology of IB. Continued studies on the in vitro and in vivo pathogenic and immunological features of the IBV strains detected during this survey, as well as the generation and analysis of molecular data from genes involved in IBV pathogeny will being carried out by the authors in order gather further insight on the impact of the IBV strains reported herein.


53

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Bochkov, Y.A., Tosi, G., Massi, P. & Drygin, V.V. (2007). Phylogenetic analysis of partial S1 and N gene sequences of infectious bronchitis virus isolates from Italy revealed genetic diversity and recombination. Virus Genes, 35, 65-71.

Cavanagh, D. (2005). Coronaviruses in poultry and other birds. Avian Pathology, 34, 439-48.

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Di Fabio, J., Rossini L.I., Orbell, S.J., Paul, G., Huggins, M.B., Malo, A., Silva, B.G. & Cook, J.K. (2000). Characterization of infectious bronchitis viruses isolated from outbreaks of disease in commercial flocks in Brazil. Avian Diseases, 44, 582-589.

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Hall, T. A. (1999) BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symposium Series, 41, 95-98.

Ignjatovic, J. & Sapats, S. (2005) Identification of previously unknown antigenic epitopes on the S and N proteins of avian infectious bronchitis virus. Archives of Virology, .150:1813-1831.

Lee, E.K., Jeon, W.J., Lee, Y.J., Jeong, O.M., Choi, J.G., Kwon, J.H. & Choi, K.S. (2008). Genetic diversity of avian infectious bronchitis virus isolates in Korea between 2003 and 2006. Avian Diseases, 52, 332-337.

Liu, S., Chen, J., Chen, J., Kong, X., Shao, Y., Han, Z., Feng, L., Cai, X., Gu, S. & Liu, M.(2005). Isolation of avian infectious bronchitis coronavirus from domestic peafowl (Pavo cristatus) and teal (Anas). Journal of General Virology, 86,719-725.

Liu, S., Wang, Y., Ma, Y., Han, Z., Zhang, Q., Shao, Y., Chen, J. & Kong, X.(2008a).Identification of a newly isolated avian infectious bronchitis coronavirus variant in China exhibiting affinity for the respiratory tract. Avian Diseases, 52, 306-314.

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Madu, I.G., Chu, V.C., Lee, H., Regan, A.D., Bauman, B.E. & Whittaker, G.R. (2007). Heparan sulfate is a selective attachment factor for the avian coronavirus infectious bronchitis virus Beaudette. Avian Diseases, 51, 45-51.

Martin, D.P., Williamson, C. & Posada, D. (2005). RDP2: recombination detection and analysis from sequence alignments. Bioinformatics, 21, 260-262..

Masters, P.S. (2006). The molecular biology of coronaviruses. Advances in Virus Research, 66:193-292.

Montassier, M.F.S., Brentano, L., Richtzenhain, L.J. & Montassier, H.J. (2006). Genetic diversity on S1 glycoprotein of avian infectious bronchitis virus strains isolated in Brazil between 1988–2000. In Proceedings of the V International Symposium on Avian Corona and Pneumoviruses (p119-131). Rauischholzhausen, Germany.

Moya, A., Elena, S.F., Bracho, A., Miralles, R. & Barrio, E. (2000). The evolution of RNA viruses: a population genetics view. Proceedings of the National Academy of Sciences,.97, 6967-6973.

Raj, G.D. & Jones, R.C. (1997) Infectious bronchitis virus: Immunopathogenesis of infection in the chicken. Avian Patholology, 26,677-706.

Roussan, D.A., Totanji, W.S. & Khawaldeh, G.Y. (2008) Molecular subtype of infectious bronchitis virus in broiler flocks in Jordan. Poultry Science, 87,661-664.

Seyfi Abad Shapouri, M.R., Mayahi, M., Assasi, K. & Charkhkar, S. (2004). A survey of the prevalence of infectious bronchitis virus type 4/91 in Iran. Acta Veterinaria Hungarica,52,163-166.

Shen, S., Law, Y.C. & Liu, D.X. (2004) A single amino acid mutation in the spike protein of coronavirus infectious bronchitis virus hampers its maturation and incorporation into virions at the nonpermissive temperature. Virology, 326,288-298.


55

Shimazaki, Y., Horiuchi, T., Harada, M., Tanimura, C., Seki, Y., Kuroda, Y., Yagyu, K., Nakamura, S. & Suzuki, S. (2008) Isolation of 4/91 type of infectious bronchitis virus as a new variant in Japan and efficacy of vaccination against 4/91 type field isolate. Avian Diseases, 52, 618-22.

Sungwhan, A., Maeda, A. & Makino, S. (1998) Coronavirus transcription early in infection. Journal of Virology, 72, 8517-8524.

Tamura, K., Dudley, J., Nei, M. & Kumar, S. (2007). MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Molecular Biology and Evolution, 24:1596-1599.

Villarreal, L.Y., Brandão, P.E., Chacón, J.L., Assayag, M.S., Maiorka, P.C., Raffi, P., Saidenberg, A.B., Jones, R.C., & Ferreira AJ. (2007a).Orchitis in roosters with reduced fertility associated with avian infectious bronchitis virus and avian metapneumovirus infections. Avian Diseases, 51, 900-904

Villarreal, L.Y., Brandão, P.E., Chacón, J.L., Saidenberg, A.B., Assayag, M.S., Jones, R.C., Ferreira, A.J. (2007b). Molecular characterization of infectious bronchitis virus strains isolated from the enteric contents of Brazilian laying hens and broilers. Avian Diseases, 51,974-978.

Wang, L., Junker, D. & Collisson, E.W. (1993) Evidence of natural recombination within the S1 gene of infectious bronchitis virus. Virology, 192, 710-716.

Worthington, K.J., Currie, R.J. & Jones, R.C.(2008). A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. Avian Pathology,37, 247-257.

Xu, C., Zhao, J., Hu, X. & Zhang, G. (2007).Isolation and identification of four infectious bronchitis virus strains in China and analyses of their S1 glycoprotein gene. Veterinary Microbiology, 122, 61-71.

Youn, S., Collisson, E.W. & Machamer, C.E.(2005).Contribution of trafficking signals in the cytoplasmic tail of the infectious bronchitis virus spike protein to virus infection. Journal of Virology, 79,13209-13217.


56

Table 1. Avian infectious bronchitis virus (IBV) field strains from Brazilian poultry included in the analysis of partial spike gene regarding bird type (broilers, layers and breeders), age (in weeks), signs present at the flock at the time of sample collection and sample from which the strains were recovered. Strain Genbank Type Age Signs Sample IBV/BRAZIL/2008/USP-13 FJ791254 broiler 6w Resb lungs IBV/BRAZIL/2007/USP-14 FJ791255 layers 28w Entc,

Repd female reproducvive tract

IBV/BRAZIL/2008/USP-15 FJ791256 NAa NA NA Pool of organs IBV/BRAZIL/2007/USP-16 FJ791257 broilers 5w Ent enteric content IBV/BRAZIL/2007/USP-17 FJ791258 layers 27w Ent,

Rep female reproducvive tract

IBV/BRAZIL/2007/USP-18 FJ791259 broilers Ni Rene kidney IBV/BRAZIL/2008/USP-19 FJ791260 Grand-

parents Ni Res,

Rep lung and trachea

IBV/BRAZIL/2008/USP-20 FJ791261 broilers 5w Ent enteric content IBV/BRAZIL/2008/USP-21 FJ791262 broilers 5w NA enteric content IBV/BRAZIL/2008/USP-22 FJ791263 broilers 3w NA enteric content IBV/BRAZIL/2008/USP-23 FJ791264 NA NA NA trachea IBV/BRAZIL/2008/USP-24 FJ791265 broilers 7w Ent enteric content IBV/BRAZIL/2008/USP-25 FJ791266 layers 23w Resp,

Rep female reproducvive tract

IBV/BRAZIL/2007/USP-26 FJ791267 broilers 6w Res trachea IBV/BRAZIL/2008/USP-27 FJ791268 broilers 5w Ent enteric content IBV/BRAZIL/2008/USP-28 FJ791269 broilers 5w Res lungs IBV/BRAZIL/2008/USP-29 FJ791270 broilers 6w Res trachea IBV/BRAZIL/2007/USP-30 FJ791271 breeders

(heavy) 45w Rep female

reproducvive tract IBV/BRAZIL/2008/USP-31 FJ791272 layers 29w Rep,

Res trachea

IBV/BRAZIL/2007/USP-32 FJ791273 breeders (heavy)

36w Rep enteric content

aNA = not available bRes = respiratory cEnt = enteric dRep = reproductive eRen = renal

VI.

INT.

SYM

PO

SIU

M O

N A

VIA

N C

OR

ON

A-

AN

D P

NE

UM

OV

IRU

SE

S, R

AU

ISC

HH

OLZ

HA

US

EN

, GE

RM

AN

Y, 2

009

57

Tab l

e 2.

Nuc

leot

ide/

amin

o ac

id id

entit

ies

(in %

) an

d re

spec

tive

stan

dard

dev

iatio

ns (

in b

rack

ets)

of p

artia

l spi

ke g

ene

sequ

ence

s (p

ositi

ons

726

to 1

071

of s

train

UK

/7/9

3, G

enba

nk A

cces

sion

num

ber

Z839

79) a

mon

gst t

hree

clu

ster

s of

Bra

zilia

n fie

ld s

train

s of

A

vian

infe

ctio

us b

ronc

hitis

viru

s (IB

V) a

nd a

rche

typi

cal g

enot

ypes

.

Clu

ster

Bra

z il 0

1 B

razi

l 02

Bra

zil 0

3 M

ass

D27

4 A

rk

Cal

D

E072

C

on

4/ 9

1

Bra

z il

01

99.6

(0.3

)/ 99

.1 (0

.6)

97.5

(0.3

)/ 95

.6 (0

.6)

93.0

(0.3

)/ 91

.6 (0

.9)

81.4

(0.5

)/ 7

9.3

(1.1

) 79

.5 (0

.3)/

78.9

(0.7

) 80

.4 (0

.3)/

77.3

(0.6

) 80

.6 (0

.5)/

79.1

(0.6

) 56

.8(0

.2)/

41.9

(0.4

) 80

.8 (0

.3)/

79.4

(0.5

) 78

.6 (0

.5)/

75.5

(0.9

)

Bra

z il

02

99

.7 (0

.2)/

99 (0

.7)

94.7

(0.4

)/ 94

.9 (0

.9)

81.7

(0.4

)/ 7

9.2

(1.0

) 80

.5 (0

.2)/

81.1

(0.5

) 81

.9 (0

.2)/

80.6

(0.6

) 81

.5 (0

.4)/

80.8

(0.6

) 58

.2 (0

.2)/

43.6

(0.4

) 81

.1 (0

.1)/

79.4

(0.5

) 80

.3 (0

.5)/

78.4

(1.0

)

Bra

zil

03

99.0

(0.5

)/ 98

.8 (0

.5)

80.0

(0.4

)/ 7

7.3

(1.0

) 79

.4 (0

.4)

79.2

(0.5

) 80

.1 (0

.2)/

79.7

(0.6

) 80

.4 (0

.3)/

79.8

(0.6

) 56

.3 (0

.3)/

43.7

(0.4

) 79

.7 (0

.2)/

78.3

(0.0

5)

80.5

(0.5

)/ 80

.5 (1

.1)



58

Figure 1. Rooted neighbor-joining distance tree with the MCL model for nucleotides 736 to1071 (regarding Genbank Accession number Z83979) of the spike glycoprotein gene of Avian infectious bronchitis virus (IBV) of archetypical genotypes and the 20 field strains from the present study (bold, underlined). Vaccine 01 and Vaccine 02 are two commercial vaccine strains also included in the study. Numbers at each node are bootstrap values (only those >50 are shown). The bar represents the number of nucleotide substitutions per site.

IBV/BRAZIL/2008/USP - 13 IBV/BRAZIL/2008/USP -14

MASS VACCINE 01 IBV/BRAZIL/2008/USP -15 AY851295.1M41

MASS VACCINE 02 DQ830980.1M41 AY561716.1Con L18990.1Con

X15832.1D274 AF169860.1Ark/213/96 AF169859.1Ark/15C/96 AF169858.1Ark/1535/95

DQ912832.1CAL99 AY942737.1Cal99 AY942738.1Cal99

IBV/BRAZIL/2008/USP -16 IBV/BRAZIL/2008/USP -17 IBV/BRAZIL/2008/USP -18

IBV/BRAZIL/2008/USP -19 IBV/BRAZIL/2008/USP -20

IBV/BRAZIL/2008/USP -21 IBV/BRAZIL/2008/USP -22 IBV/BRAZIL/2008/USP -23 IBV/BRAZIL/2008/USP -24

IBV/BRAZIL/2008/USP -25 IBV/BRAZIL/2008/USP -26 IBV/BRAZIL/2008/USP -27 IBV/BRAZIL/2008/USP -28 IBV/BRAZIL/2008/USP -29 IBV/BRAZIL/2008/USP -30

AJ457137.1Italy -02 IBV/BRAZIL/2008/USP -31

IBV/BRAZIL/2008/USP -32 AJ618987.1 -793B AJ618984.1 -793B

AJ618985.1 -793B AJ618986.1 -793B

AF093794.14 -91 AF093793.14 -91 EU359658.1DE072 EU359660.1DE072 EU359659.1DE072

NC007447Bredavirus 100

87

87

62

74

99

99

92 100

99

99

100

99

70

88

70

99

99

86 98

94

71 76 96

52

66

93 67

0.05

Massachusetts

Connecticut

Arkansas

California 99

Brazil 03 ( recombinant )

Brazil 02

Brazil 01

4/91

DE072


59

GENOTYPING AND SEROTYPING OF GUANGXI IBV ISOLATES DURING 1985~2008

WEI PING LI MENG, WEI ZHENG-JI, WANG XIU-YING, MO MEI-LAN and CHEN QIU-YING

Institute for Poultry Science and Health-Guangxi University-Nanning 530005, China SUMMARY Nucleocapsid protein-N protein-genes and S1 gene hypervariable region I (HVR I) of 26 IBV strains isolated in Guangxi between 1985 and 2008 were amplified by reverse transcriptase polymerase chain reaction (RT-PCR), and then were cloned, sequenced and compared with IBV reference strains. Deletion and insertion were not found in the N genes of the 26 isolates, but there were 4 and 3 amino-acid residues inserted at the sites of 33-4 and 34-35 within HVR I respectively in 18 of the isolates and isolate GX-NN6 had 4 amino-acid residues inserted at both sites. Phylogenetic tree based on amino acid sequences of HVR I showed that all the isolates were divided into 3 clusters and cluster I consisted 20 out of 26 isolates and was far from the vaccine strains in distance, phylogenetic tree based on amino acid sequences of N genes showed that there were 4 clusters and the majority of the isolates were grouped in cluster A, B and C, while only isolate GX-NN5 and the vaccine strains were in cluster D. Monovalent antisera against 7 field isolates and 3 commonly used vaccine strains of infectious bronchitis virus (IBV) were prepared in rabbits and were used in the virus neutralization test against the heterogeneous and homogeneous viruses on tracheal organ cultures (TOC). Relatedness values of each virus calculated from cross-neutralization antibody titer showed that 7 serotypes were distinguished among the 26 isolates tested. The relationship between genotyping and serotyping was also discussed. INTRODUCTION Infectious bronchitis (IB) is an acute, highly contagious viral disease causing by infectious bronchitis virus (IBV) a member of Coronavirus of Coronaviridae. IB is of major economic importance for it is a cause of poor weight gain and feed efficiency, death resulting from kidney damage, secondary and or mixed infections mainly in respiratory with other pathogens resulting death or increase of medication cost, declines in egg-production and egg-quality, and other un-well-known problems just like damages of muscle and gastrointestinal tract. Its highly transmissible nature and


60

the prevalence of multiple serotypes of IBV which may differ from time to time and from place to place, have been complicating and increasing the difficulty and cost of attempts to prevent the disease by vaccination (Cavanagh and Naqi, 1997). IBV strains vary greatly in their tissue tropism, and different strains can grow and result in damage at many epithelial surfaces including the respiratory tract, kidney, oviduct, muscle, and parts of gastrointestinal tract (Dhinakar Raj and Jones, 1997). IBV has four essential structural proteins, the three membrane proteins, the spike(S), integral membrane (M), and small envelope (E) proteins, and a phosphorylated, nucleocapsid (N) protein. The S protein interacts with cellular receptors and induces cell and viral membrane fusion Bosch et al (2003). The S glycoprotein is post-translationally cleaved into S1 and S2 subunits during viral maturation and the S1 protein determines serotype and is believed to play a major role in the induction of protective immunity. The mutation of the S1gene in the N-terminal amino acids directly leads to the emergence of new serotypes and the change of tissue tropism. N protein plays important role in virus replication, assembly, and cell immunity Liu et al (2009). IBV infections and its associated problems are still common in the field in Guangxi (Wei and Li, 1997; Mo et al., 2001; Wei et al., 2008) In the present study, the phylogenetic trees based on the nucleotide sequences of hypervariable region of S1 gene (HVR) of 26 field strains (Wei et al., 2008; Mo et al., 2009) which were isolated from Guangxi during the years of 1985 to 2008 were made and the serological relationship classified as serotypes of all the isolates are evaluated via virus neutralization on tracheal organ culture (TOC). The common used vaccine strains and some isolates of other places are also used as reference in the study. The relationship between genotyping and serotyping was also discussed. MATERIALS and METHODS Viruses and monovalent antisera Twenty-six IBV isolates isolated from dead or diseased chickens from different farms which had been inoculated with vaccines Ma5 or H120 in Guangxi provinces during the years of 1985-2008 and 4 reference strains H120, Ma5, M41 and 4/91 were also used in the study (Table 1). The viruses were propagated and passed in the allantoic cavities of 9-day-old specific pathogen free (SPF) chicken embryos and the allantoic fluids were collected after 72h post-inoculation at the fourth passage and used for the extraction of viral RNA, and the 10 of those of the viruses were inactivated and used as immunogens for the preparation of monovalent antisera in rabbits as described by Li et al (1991). RNA extraction and amplifications of genes S1 and N by RT-PCR Viral RNA was extracted from the infectious allantoic fluids in the method as described by Wei et al (2008). The primers used for the amplification of the N gene were NP1: 5’-CCATGGCAAGCGGTAAAGCAR -3, NP2 5’-CCACTCAAAGTTCATTCTCTCC -3’, and the anticipated amplification segment is about 1 230bp containing the entire N gene. The primers used for the amplification of the IBV S HVR I were B1 5’TGGTTGGCATTTACACGGGG -3’and B2 5’CAATGGGTAACAAACAC-3’and the anticipated amplification segment is about 228bp. The PCR conditions for the amplification S HVR I was performed as Mo et al


61

(2001). N gene was performed as Mo et al (2009). The products were analyzed on 0.8% agarose gel. Table 1 Sources and backgrounds of Guangxi isolates of IBV

Isolates Years Clinical problems GenBank accession No. S1 N

GX-C 1985 Abnormality of respiratory and urinary systems DQ859267 FJ767923

GX-G 1988 Abnormality of respiratory and urinary systems EF547363 FJ767924

GX-XD 1988 Abnormality of respiratory and urinary systems EF547364 FJ767925

GX-NN3 2004 Abnormality of respiratory and urinary systems DQ859271 FJ767914

GX-NN1 2005 Abnormality of respiratory systems DQ859268 GQ174473 GX-NN2 2005 Abnormality of respiratory systems DQ859272 FJ767913 GX-YL1 2005 Abnormality of urinary systems DQ859273 FJ767927 GX-YL3 2005 Abnormality of urinary systems DQ859274 FJ767929 GX-YL4 2005 Abnormality of urinary systems EF382353 FJ770461 GX-YL5 2005 Abnormality of urinary systems EF382354 FJ548847

GX-NN4 2005 Abnormality of respiratory and urinary systems EF382348 FJ767915



GX-NN7 2006 Abnormality of respiratory systems EF382349 NO GX-NN8 2006 Abnormality of urinary systems EF382350 FJ767918

GX-YL2 2006 Abnormality of respiratory and urinary systems DQ859275 FJ767928

GX-YL6 2006 Abnormality of respiratory systems EF382355 FJ767930 GX-YL7 2006 Abnormality of respiratory systems EF428326 FJ767931

GX-NN10 2006 Abnormality of respiratory systems EF382351 FJ767920


GX-LZ1 2006 Abnormality of respiratory and urinary systems EF428324 FJ767926


GX-YL8 2007 Abnormality of urinary systems FJ770462 FJ767932 GX-YL9 2007 Abnormality of urinary systems FJ770463 FJ767933

GX-NN12 2008 Abnormality of respiratory systems FJ770466 FJ767922

GX-GL1 2008 Abnormality of respiratory and urinary systems FJ770465 FJ767934

Gene sequence and analysis The products were purified and ligated into PMD18-T vector (TaKaRa, Japan). The resulting plasmids were designated T-vector-M and sequenced by Sangon bio-company (Shanghai, China).Sequences of T-vector-M and their deduced amino acids were compared with that of IBV reference strains derived from different regions or countries using the Lasergene software packageV5.0 (DNASTAR Inc, USA). The composition of the nucleotide sequence of N and HVR I genes was analyzed with DNAMAN software version4.0 (Lynnon BioSoft, Quebec, Canada). A phylogenetic tree was generated using an alignment of gene sequences in the Mregion from the above-mentioned IBV strains by the Lasergene software packageV5.0 (DNASTAR


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Inc, USA). Phylogenetic analysis was performed as previously described Wei et al (2008) Virus neutralization (VN) test VN tests were performed on TOC by a mode of fixed virus (100TOC-ID50) and serial diluted serum as previously described by Li et al (1991). Totally 30 IBV strains were used in the VN tests with the antisera of 10 different strains for the endpoint dilutions of each virus, and the viruses share the same endpoint dilutions with the antisera were considered as the same serotype. RESULTS The applications of genes S1 and N All of viruses got the RT-PCR amplified products of S1-HVR I in the sizes of 228nt or 249nt (Fig. 1) and of N gene in the size of 1230nt (Fig.2).

Fig. 1 Electrophoresis of PCR-amplified products of S1 gene of some isolates

Fig.2 Electrophoresis of PCR-amplified products of N gene of some isolates Alignment of amino acids and phylogenetic analysis of S1 gene of the IBV strains The amino acid sequences of the HVR I protein of the isolates and another eleven reference strains were aliged (Fig.3). There were 4 and 3 amino-acid residues inserted at the sites of 33-34 and 34-35 within HVR I respectively in 19 of the isolates, except that of isolate GX-NN6 was 4 amino-acid residues inserted at the both sites; A phylogenetic tree based on amino acid sequences of HVR I of all the viruses showed that they were classified into 4 distinct clusters. 20 out of IBV isolates

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23

250 bp 200 bp 242 bp

190 bp

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18

1230bp 1200bp 1000bp


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were grouped into clusterⅠ, isolates GX-YL1, GX-G, GX-XD and GX-NN2 had close relationship with Mass type vaccine strains, and they shared ClusterⅠ, isolates GX-YL6 and GX-NN7 of cluster Ⅰ had close relationship with the European strain 4/91

Fig. 3 Phylogenetic tree based on the deduced amino acid sequences of HVR I genes of Guangxi IBV isolates and reference IBV strains Alignment of amino acids and phylogenetic analysis of N gene of the IBV strains Deletion and insertion were not found in the N genes of 26 isolates, but it was found that point mutations inducing the amino acid substitutions were common in N genes of 26 isolates. A phylogenetic tree (Fig. 4) based on amino acid sequences of N of all the viruses showed that they were classified into 5 distinct clusters. 11 out of IBV isolates were grouped into cluster A. 10 out of IBV isolates were grouped into cluster B. Isolates GX-C, GX-G, X-XD and GX-NN6 were grouped into cluster C. GX-NN5 relationship with Mass type vaccine strains, and they shared Cluster D, had close GX-YL1 and GX-NN2, which shared 97.4% S1 gene amino acid identity with vaccine strain H120, did not clustered with the H120 strain based on N gene amino acid sequences. GX-NN7 and GX-YL6, which had 96.1% and 97.8% S1 gene amino acid identity with 4/91 respectively, did not clustered with the 4/91 strain based on N gene amino acid sequences. GX-C and GX-G, GX-XD, which belonged to two separate subgroups based on S1 gene amino acid sequences, clustered into the same subgroup based on N gene amino acid sequences. Recombinations were found in the genome of IBV isolate GX-NN5, isolated in 2005, which had 99.0% N gene amino acid identity with vaccine strain H120 and only had 52.6% S1 gene amino acid identity with H120. These results suggested that gene mutations and recombinations

Ⅰ

Ⅱ

Ⅲ

Ⅳ


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had occurred in the Guangxi IBV strains. The phylogenetic analysis based on the N gene sequences do not usually follows closely the phylogenetic clustering based on the S1 gene.

Fig. 4 Phylogenetic tree based on the deduced amino acid sequences of N genes of Guangxi IBV isolates and reference IBV strains Results of VN test Results of the VN test showed that 7 serotypes were classified from all the 30 strains: 13 isolates belong to serotype 1; 5 isolates belong to serotype 2; 5 strains including two isolates GX-NN10 and GX-YL2, two vaccine strains H120 and Ma5, and one reference strain M41 belong to serotype 3; isolate GX-YL1 belongs to serotype 4; isolates GX-GL1, GX-NN7, as well as one vaccine strain 4/91 belong to serotype 5; isolates GX-YL8 and GX-YL9 belong to serotype 6; since the neutralization titers of isolate GX-NN12 against all the antisera were relatively low, it may belong to another serotype (serotype 7) (Table 2). The comparison of the results of genotypings and Serotyping of the IBV strains When the results of genotypings of genes S1 and N were compared with results of Serotyping, only the results of 19 strains of S1 genotyping and of 14 strains of N genotyping out of total 30 strains were identical with the results of Serotyping (Table 2).

A

B

C

D


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DISCUSSION The results of gene sequencing of both S1 and N showed that the variations of genes are undergoing even the N gene which used to believe as conserved. There are 7 serotypes prevalence in Guangxi chickens and more and more serotypes are appeared in the recent years. Although the antigen variation is based on the variations of genes especially S1 gene, but the results of the study demonstrated that the results of genotyping can not fully represent the results of serotyping. The data of the study will provides a reliable scientific basis for catching up the emerged antigenic variation of the field viruses and developing more effective vaccine for the disease control in Guangxi.

Table 2 Relationship between the results of genotyping and serotyping of the viruses

Virus strains Serotypes Genotypes based on S1 gene Genotypes based on N gene

GX-YL5 1 B GX-NN4 1 B GX-NN1 1 A GX-NN3 1 B GX-NN5 1 D GX-NN6 1 C GX-NN9 1 B GX-NN11 1 A GX-YL3 1 B GX-YL4 1 A GX-YL6 1 B GX-YL7 1 A GX-LZ1 1 A GX-C 2 C GX-G 2 C GX-XD 2 C GX-NN2 2 B GX-NN8 2 A H120 3 D Ma5 3 D M41 3 D GX-NN10 3 A GX-YL2 3 A GX-YL1 4 A 4/91 5 D GX-NN7 5 A GX-GL1 5 A GX-YL8 6 B GX-YL9 6 B GX-NN12 7 B


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REFERENCES Bosch, B.J., Van Der Zee, R., DeHaan, C. A., Rottier, P. J., 2003. The corona virus

spike protein is a class I virus fusion protein: structural and functional characterization of the fusion core complex. J. Virol.77 (16), 8801-8811

Cavanagh, D., and S. A. Naqi. 1997 Infectious bronchitis. In: Diseases of Poultry. 10th ed. B.

W. Calneck et al. ed. Iowa State University Press, Ames, pp.511-526. Dhinakar Rai, G. and R. S. Jones. 1997. Infectious bronchitis virus: Immunogenesis of infection in the chicken. Avian Pathol. J. 26:677-706.

Li, K-R., P., Wei and M. F., Liang. 1991. Serotyping of avian infections bronchitis virus by neutralization test in tracheal organ cultures. Journal of Guangxi Agricultural College, J.10 (3):1-6.

Liu X-L. Su J-L. Zhao J-X. et al. Complete genome sequence analysis of a predominant infectious bronchitis virus (IBV) strain in China. J Virus Genes, 2009. 38:56–65

Mo, M-L, K-R Li and P. Wei. The applicaions of RT-PCR and RFLP analysis on the diagnosis and genotyping of S1 gene. Chinese J. of Preventive Veterinary Medicine. 2001(4): 277~281.

Mo M-L., Li M. Chen Q-Y., et al. Analysis of Biological Characteristics and Structural Genes of IBV Isolate from Chickens with Immunoprophylaxis Defeat. J. Genomics and Applied Biology, 2009, 28(2):275-280

Wei, P and K-R Li. 1997. The serotyping of Guangxi isolates of IBV in chickens by virus neutralization test on tracheal organ culture. 134th Annual Convention of American Veterinary Medical Association, July 19-24. Reno, Nevada, USA, PP32.

Wei, Z-J, P. Wei, M-L Mo et al. Genetic variation of S1 genes hypervariable region I of infectious bronchitis viruses isolated in different periods in Guangxi. Chinese J. of Virology, 2008, 24(2): 126~132.


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AVIAN INFECTIOUS BRONCHITIS VIRUSES FROM SLAUGHTERED CHICKENS EXHIBIT WIDE VARIETIES

CHEN HW, HUANG YP and WANG CH

School of Veterinary Medicine, National Taiwan University, No. 1, Sec. 4, Roosevelt Rd., Taipei 10617, Taiwan

SUMMARY Avian coronavirus infectious bronchitis (IBV) poses a major threat to the global poultry industry. New IBV geno- and serotypes are continually reported. However, information on IBV prevalence data is not frequently addressed. This study reports on a viral surveillance in Taiwan from 2005 to 2006 with sampling conducted in poultry slaughter houses. The genetic features of the obtained field isolates were investigated using sequence analysis and Simplot analysis. The one-directional neutralization test was performed to examine the antigenic variations. The selection pressures which may contribute to Taiwan IBVs evolution during the last decades were assessed. In the surveillance, 8 out of 47 flocks (17%) were IBV-infected, from which 13 IBV isolates were recovered. Based on the phylogenetic analysis of the S1 gene, 11 of 13 isolates (84.6%) were clustered into Taiwan group I. An IBV isolate experienced frequent recombination events with the China-like IBVs in the S gene. Another isolate demonstrated the China-like and H120-like genomes incorporation within the S2 gene and the E gene region, respectively. Some antigenic changes were found in the one-directional neutralization test. However, no positive selection pressures were related to those variations in the S1 genes among Taiwan IBVs. We suggest sampling chickens in poultry slaughter houses is effective and helpful in representing the viral prevalence covering sub-clinical infection circumstances. The IBV populations identified herein exhibited genetic and antigenic diversities. INTRODUCTION Infectious bronchitis virus (IBV) is the best-known avian corona virus (Cook, 2002). This disease was first reported in 1931 and was soon distributed worldwide (Cavanagh and Naqi, 2003). IBV is a highly infectious and contagious agent to chickens. The respiratory tract is the primary organ, while the urogenital system such as the kidney and oviduct are also affected. Young chickens suffering from IBV infection can acquire permanent damage to their oviducts. Older infected chickens may exhibit poor feed efficiency (Cavanagh, 2007; Cavanagh and Naqi, 2003). IBV


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has great economic importance owing to its effects on egg-laying performance and meat production (Cook, 2002). IBV is still the main infectious agent in chicken farms. The genome of avian coronaviruses is structured by a large sized ssRNA (-27.6 kb), from which four structural proteins are encoded. The spike glycoprotein (S), forming the surface projections of the virion, is described as a determinant of the host range and pathogenicity (Cavanagh, 2007). The pos-translationally cleaved subunits, S1 and S2 carry the epitopes for inducing virus neutralizing antibodies (Koch, et al., 1990). The S1 subunit is involved in virus entry and hemagglutination activity (Cavanagh and Davis, 1986; Koch and Kant, 1990). The envelope (E) and membrane (M) proteins are required with the virion assembly. The nucleocapsid (N) protein is closely associated with the RNA genome package (Lai and Holmes, 2001). IBV exists as dozens of sero- or genotypes. Like most RNA viruses, IBV undergoes mutations at a high frequency owing to the error prone RNA polymerase (Wege, et al., 1982). Moreover, IBV possesses a unique discontinuous transcription system and polymerase jumping phenomenon, which contribute to the high frequency of RNA recombination (Lai and Holmes, 2001). Over the last decade, the continuing emergence of variant strains has complicated disease control. A circulating serotype may experience antigenic shift via slight variations in the S1 gene (Cavanagh, et al., 1992). Poor cross-protection is conferred between serotypes (Cavanagh, 2007). Vaccination has been implemented to control IB for several decades. In developing live vaccines, virulent viruses are attenuated by passage in chicken embryos (Cavanagh, 2007). Massachusetts (Mass) type strains such as M41 and H120 have been widely used for immunization (Cavanagh and Naqi, 2003). However, animal exposure to live vaccines can cause damage by spreading viruses in large poultry populations (Farsang, et al., 2002). By acting as a heterologous RNA donor template, vaccine viruses may contribute to the genetic evolution of IBVs (Wang, et al., 1993), i.e. genomic recombination with vaccine strains can occur (Bochkov, et al., 2007; Jia, et al., 1995; Mondal and Cardona, 2007). IB field outbreaks still occur despite routine vaccine use (Cavanagh and Naqi, 2003). New IBV geno- and serotypes are continually reported. However, information on IB prevalence data is not frequently addressed. In Taiwan, apart from two local progeny strain types, Taiwan group I (TW-I) and Taiwan group II (TW-II), heterologous Mass serotype has been vaccinated for over a decade. A recent molecular investigation using the 6.8 kb structural gene regions of viruses revealed that Taiwan IBVs undergo recombination (Chen, et al., 2009). In such a vulnerable epizootiological situation, viruses may evolve rapidly under selection pressure. Therefore, monitoring disease status and analyzing the circulating viruses is critical. In the present study, an intensive IBV viral investigation program was performed in chickens. Samples were collected at a large scale from poultry slaughter houses. Disease prevalence was formulated using the virus isolation rate. Obtained field IBV isolates were molecularly characterized and the putative recombinants were further elucidated using the 6.8 kb fragment sequence. The selection pressures which may contribute to Taiwan IBVs evolution were assessed. In addition, a serological method was employed to study the antigenic varieties among the field isolates and reference strains.


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MATERIALS and METHODS Samples collection and virus isolation Four hundred and seventy chicken tracheal samples originating from 47 flocks nationwide were collected in two poultry slaughter houses during 2005-2006. Five to ten samples from each flock were pooled and homogenized in tryptose phosphate broth (Difco Lab., Detroit, MI). The homogenates were clarified using centrifugation at 3000g for 20 minutes and sterilized by passing through 0.45-μm syringe filters (Pall Corp., Ann Arbor, MI). Virus isolation was performed using the allantoic route in 9－11-day-old specific pathogen free (SPF) chicken embryos (Animal Health Research Institute, Tamsui, Taiwan). Each egg received 0.1 to 0.2 ml inoculation and was incubated at 37°C for 48 hours. After two blind passages, allantoic fluid was harvested for further study. Viral RNA extraction, RT-PCR and DNA sequencing Viral RNA was extracted from infected allantoic fluid using the Qiamp Viral RNA kit (Qiagen, Valencia, CA) following the manufacture’s instructions. Genes of interest were amplified with previously published primers (Huang and Wang, 2007) using one-step reverse-transcriptase polymerase chain reaction (RT-PCR). The obtained DNA products were submitted for sequencing in both directions with a commercial service (Tri-I Biotech, Taipei, Taiwan). Each nucleotide was confirmed from four identical results. Sequence analyses and recombination analyses Sequence analyses were conducted by Lasergene software package (DNAStar, Madison, WI). Phylogenetic trees were constructed using MEGA version 4 (Tamura, et al., 2007). SimPlot version 3.5.1 was employed to detect the inter-strain recombination events among the reference IBVs and field isolates. Selection pressure analysis on the S1 gene The selective pressures that contributed to the variations in the whole S1 gene of Taiwan IBVs, TW-I and TW-II, were assessed by calculating the average dS/dN ratio (synonymous substitutions per synonymous site to non-synonymous substitutions per non-synonymous site) with the web-based SNAP program (http://www.hiv.lanl.gov). A ratio lower than one indicates a specific gene region may be under positive pressures (Korber, 2000). Titration with viruses, antisera and one-directional neutralization test Viruses were ten-fold serially diluted and titrated in 9－11-day-old SPF chicken embryos. After seven days incubation, viral EID50 was determined according to the Reed and Muench method (1938). Antisera against two Taiwan IBVs, strains 2575/98 (TW-I) and 2296/95 (TW-II), respectively, were prepared in SPF chickens (Wang and Huang, 2000). Mass type strain antiserum was purchased from Charles River Laboratories (North Franklin, CT). Each antiserum was titrated against its homologous virus strain in SPF chicken embryos as previously described (Cowen and Hitchner, 1975). One-directional neutralization test was performed following the previous study (Wang and Huang, 2000). Briefly, an amount of 100 EID50 virus reacted with 20 units of antiserum in an equal volume at room temperature for one

http://www.hiv.lanl.gov


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hour. Ten 9-11-day-old SPF chicken embryos inoculated with the virus-serum mixtures were examined for IB lesions after seven days. A virus was considered to be of the same serotype with the antiserum if more than five embryos were protected from infection. Accession numbers The nucleotide sequence data reported in this paper have been submitted to GenBank Submission nucleotide sequence database and have been assigned the accession number from GQ229237 to GQ229258 (Table 1). The accession numbers and genes of interest in the IBV reference strains included in this study are listed as follows: 1171/92, DQ646406 (S1-N); 1211/92, AF250006 (S1); 2296/95, DQ646404 (S1); 2575/98, DQ646405 (S1); 2993/02, AY606316 (S1); 3025/02, AY606317 (S1); 3051/02, AY606318 (S1); 3071/03, AY606319 (S1); 3263/04, EU822338 (S1); 3468/07, EU822336 (S1); T03/01, AY606315 (S1); T07/02, AY606322 (S1); CK/CH/LDL/97I, EF030996 (S1) and EF602445 (S2-N); Gray, L18989 (S1) and M85245 (N); H120, EU822341 (S1-N); JMK, L14070 (S1). RESULTS IBV prevalence Among the 47 flocks sampled, eight were detected as positive for IBV infection after virus isolation. Thirteen IBVs were isolated and indicated in Table 1. The viral prevalence was shown to be 17% by flock, and 1.8% to 3.7% by chickens. Sequence analyses of the S1gene The S1 genes of the 13 isolates obtained in this study were directly sequenced. Based on the phylogenetic analysis of the full S1 gene (Figure 1), 11 from 13 isolates (84.6%) were clustered into TW-I. Those TW-I isolates shared 92.1－100% homology in the S1 gene with each other. In the other two isolates, the S1 gene of isolate 3374/05 was genetically related to the China CK/CH/LDL/97I-type strains (Chen, Huang and Wang, 2009). Isolate 3381/06 shared 97.9% and 97.2% with reference strains JMK and Gray, respectively. Sequence analyses of the partial N gene Partial N gene fragments (nt 178－755) of the 13 isolates were sequenced, from which 86.9-100% homology was shared among the 13 field isolates. Interestingly, the N gene from isolate 3382/06 was more similar (97.4% homology) to that of the Mass type H120, rather than that of the TW-I isolates (86.6%-89.1% homology). As illustrated in Figure 2, 3382/06 was clustered with the Mass group in terms of the N gene. The N gene sequence of the isolate 3381/06 showed 97.5% identity with reference strain Gray. Inter-strain recombination identification IBV 3374/05 was previously characterized as a natural recombinant between the Taiwan and China CK/CH/LDL/97I-type strains (Chen, Huang and Wang, 2009). A 3’ 6.8 kb gene length from isolate 3382/06 was further sequenced. The data shown in Table 2 reveals an abrupt shift in nucleotide identity when isolate 3382/06 was compared to the 1171/92 (TW-I), CH/CK/LDL/97I (China genotype VII) and H120


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(Mass), strongly suggesting that recombination events occurred. The Simplot analysis was used to illustrate the crossover events and to identify the putative recombination sites. Reference strains 1171/92, CH/CK/LDL/97I and H120 were served as the parental strains. As showed in Figure 3, the first crossover event between strains 1171/92 and CH/CK/LDL/97I was observed in the 3’ end of the S gene. The second putative recombination site was located in the middle of the E gene. The number in red indicates the genomic position in the 6.8 kb gene region. Selection pressure analyses The average dS/dN ratio was assessed from the S1 genes among the previous identified (from 1992 to 2007) local IBVs. The values obtained from TW-I (19 virus isolates, including the 11 recovered in this study) and TW-II (four virus isolates) were 4.27 and 3.34, respectively. Both values were higher than one. The results showed no positive selection pressures were related to those variations in the S1 genes from the two Taiwan IBV geno-groups. Neutralization test Based on the molecular characterization results, the two viral recombinants identified in this study, 3374/05 and 3382/06, were selected for use in performing the neutralization assays. Antisera from the three main serotypes circulating in Taiwan (TW-I, TW-II and Mass type) were employed. The results (Table 3) showed that the China-like recombinant 3374/05 possessed a distinct serotype with all of the tested antisera. The H120-like recombinant 3382/06 was of the same serotype as TW-I since most embryos can be protected by the TW-I antiserum. DISCUSSION This work indicated that IBV prevalence was 17% in Taiwan chicken flocks during 2005-2006, based on virus isolation. It was the first demonstration of IBV prevalent data in Taiwan. Compared with previous surveillance reports, the prevalence (17%) indicated here was not relatively high. According to Roussan et al., (2008), IBV was detected in up to 60% of the examined broiler flocks in Jordan. Also, 59% of the submitted clinical samples were positive for IBV in Western Europe (Worthington, et al., 2008). The RT-PCR diagnoses directly from tissues or samples employed in most surveillance situations were sensitive. However, it may not be possible to discriminate vaccine viruses. A higher IBV detection rate may result from investigations with samples or flocks that suffered from IB-suspected diseases. In this study, the tracheas of slaughtered chickens were collected in poultry slaughter houses, where sampled chickens were inspected as apparently normal. Biased sampling was therefore avoided. The data obtained from our study may cover the sub-clinical infection circumstances and represent a more factual disease status. The virus isolation based diagnosis performed in this work also made it possible to recover the live virulent viruses. IBV infection in Taiwan was recognized in 1968. A decade ago, local IBVs were molecularly characterized as two exclusive populations in the world (TW-I and TW-II) based on the RFLP of the S1 gene (Wang and Tsai, 1996). However, local IBVs keep evolving. Frequent recombination events with China CK/CH/LDL/97I-type strains were observed in several progeny strains (Chen, Huang and Wang, 2009).


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The 13 newly identified IBVs in this work were molecularly characterized. Among them, most isolates (11/13, 84.6%) were closely related to TW-I strains, suggesting that TW-I existed as a dominant geno-group in Taiwan. However, phylogenetic analyses revealed the isolate 3382/06 was clustered into TW-I based on the S1 gene, but not the N gene. In order to study the recombination site, a fragment covering the 3’ end of the structural protein gene region about 6.8 kb was further investigated. Surprisingly, in addition to the first crossover with the CK/CH/LDL/97I-type strains in the S gene, the abrupt shift in nucleotide identities among the M, 5a, 5b and N genes comparing the isolate 3382/06 to strain H120 strongly indicated another recombination event. Two putative H120-like recombinants were also reported in Italy. Different phylogeny profiles between the S1 and N gene regions toward H120 were described (Bochkov, Tosi, Massi and Drygin, 2007). In this study, we schematically present the crossover events in the plot and the genome positions of the putative recombination sites were readily observed. It was the first observation for vaccine virus genome incorporation of IBVs in Taiwan. For a decade, due to lack of proper homologous protection against local viruses, disease prevention in Taiwan relied on a cross-protective effect conferred by heterologous Mass type strains such as M41 and H120. In particular, live attenuated viruses are widely applied to many chicken types. Viruses likely shed and circulate in the field. Another heterologous isolate 3381/06 identified in this study was closely related with the American group strains, also a heterologous vaccine type approved in Taiwan. More challenging problems may arise if multiple genome incorporation occurs among field viruses. Thus the crucial observation from this study alerts the need for reconsideration on the policy of IB control. The S1 protein of IBV forms the virion surface and has several important biological functions. Mutations in the S1 gene may alter the antigenicity and pathogenecity of the virus (Lai and Holmes, 2001). To improve host adaptation ability, the virus has the greatest need to change the S1 protein. To address the relationship between the selection pressures and the long-term genetic variations in Taiwan IBVs, nearly all available S1 gene sequences from local viruses were analyzed with the dS/dN values. The results suggested no positive selection pressures might drive the evolution of the two Taiwan IBV geno-groups. Compared with the lower ratio 1.6 previously obtained in the molecular characterization on Italy 02 genotypes in Spain (Dolz, et al., 2006), our data, 4.27 (TW-I) and 3.34 (TW-II), were not suggestive of a high rate of non-synonymous changes in Taiwan IBVs. These observations may be explained by that current heterologous vaccine strains offer only partial protection and may not drive many non-synonymous substitutions on amino acid sequences in local viruses. In addition to the molecular characterization on the field isolates, examination of the antigenic aspect was also important, particularly when a new variant was reported. The two recombinant variants 3374/05 and 3382/06 were selected to further analyze their antigenic serotypes. The results showed no cross-neutralization effect was achieved between isolate 3374/05 and two groups of local viruses as well as Mass type, suggesting the isolate 3374/05 existed as a new serotype in Taiwan. Isolate 3382/06 possessed the same serotype with its S1 gene-homologous TW-I strains. Therefore, field chickens infected with those variants were probably not protected by the current preventive program. However, the disease history of the infected flocks from which variants 3374/05 and 3382/06 originated remained unavailable. The pathogenicity of those variants awaits further studies.


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Wang, L., Junker, D., Collisson, E.W., 1993, Evidence of natural recombination within the S1 gene of infectious bronchitis virus. Virology 192, 710-716.

Wege, H., Siddell, S., ter Meulen, V., 1982, The biology and pathogenesis of coronaviruses. Curr. Top. Microbiol. Immunol. 99, 165-200.

Worthington, K.J., Currie, R.J., Jones, R.C., 2008, A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. Avian Pathol. 37, 247-257.


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Table 1: IBVs isolated in this study.

Strain Year of isolation

Chicken type Location Genotypea Accession no.

of S1 gene Accession no.

of N gene

3339/05 2005 Taiwan country chicken

Yunlin TW-I GQ229237 GQ229248

3368/05 2005 Broiler Yilan TW-I GQ229238 GQ229249 3369/05 2005 Broiler Yilan TW-I GQ229239 GQ229250 3370/05 2005 Broiler Yilan TW-I GQ229240 GQ229251 3371/05 2005 Broiler Yilan TW-I GQ229241 GQ229252 3372/05 2005 Broiler Yunlin TW-I GQ229242 GQ229253 3373/05 2005 Broiler Yunlin TW-I GQ229243 GQ229254 3374/05 2005 Broiler Changhua China-like EU822337b EU822337b 3376/06 2006 Broiler Taoyuan TW-I GQ229244 GQ229255 3381/06 2006 Broiler Taoyuan JMK-like GQ229245 GQ229256 3382/06 2006 Broiler Taoyuan TW-I GQ229232b GQ229232b 3384/06 2006 Broiler Taoyuan TW-I GQ229246 GQ229257 3385/06 2006 Broiler Taoyuan TW-I GQ229247 GQ229258 a Genotype was determined based on the S1 gene. TW-I: Taiwan group I. b Sequence from S1 to N gene of isolate 3374/05 and 3382/06 were submitted. Table 2 Nucleotide identity between 3382/06 and respective putative parental strains.

Nucleotide identity (%) Parental strain S1 S2 3a 3b E M 5a 5b N 1171/92 91.2 91.5 82.8 87.2 91.6 90.7 86.4 96.0 88.1

CK/CH/LDL/97I 77.7 92.4 99.4 99.0 90.3 90.1 87.9 93.2 92.2 H120 82.0 84.2 85.6 86.2 87.9 93.0 100 100 97.6

Table 3 One-directional neutralization test of IBV isolates against antisera of TW-I, TW-II, and Mass type, respectively.

No. of protected embryos / No. of total embryos Isolate TW-I antiserum TW-II antiserum Mass antiserum PBS Control 3374/05 1/10 2/10 2/10 0/3 3382/06 9/10 1/10 0/10 0/5


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3368/05 3369/05 3384/06 3385/06

3339/05 3370/05 3371/05

3372/05 3373/05

2575/98 3071/03

3376/06 3382/06

TW-I

2296/95 3263/04 TW-II

Mass H120 3381/05

JMK Gray

American

3374/05 CK/CH/LDL/97I China VII

100

100

100

100

96

100

100

100

100

100

100

98

10096

0.02 Fig .1. Phylogenetic analysis on nucleotide sequence of whole S1 gene from the IBV isolates recovered in this study (●). The phylogenetic trees were constructed using the MEGA version 4 by the neighbor-joining method (bootstrapping for 1,000 replicates with its value > 70%).

3339/05 3370/05 3371/05

2296/95 3368/05 3369/05

3372/05 3373/05

3374/05 2575/98

3376/06 3384/06 3385/06 3071/03

3263/04

TW

3381/06 Gray American

3382/06 H120 Mass

China VII CK/CH/LDL/97I

100

100

96100

99

89

99

78

81

77

100

99

100

97

0.01 Fig. 2. Phylogenetic analysis on nucleotide sequence of partial N gene from the IBV isolates recovered in this study (●). The phylogenetic trees were constructed using the MEGA version 4 by the neighbor-joining method (bootstrapping for 1,000 replicates with its value > 70%).


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Fig. 3. Simplot analysis of the isolate 3382/06. Strains 1171/92 (pink), H120 (green), and the China strain CK/CH/LDL/97I (deep blue) were used as putative parental strains while isolate 3382/06 were queried. The similarity plot displays the consecutive nucleotide identity (%) from the S to N genes among the queried strain and parental strains. The breakpoint where the parental strains have equal identity to the query strain is the predicted recombination site. The 6.8 kb genome of 3382/06 was schematically assembled using Taiwan, China-like and H120-like sequence regions. The genomic positions of the crossover sites were indicated by numbers in red. The genomic scale was given at the top of the plot. IG: Intergenic region.


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A POLYMORPHISM STUDY ON INFECTIOUS BRONCHITIS VIRUS GENES S1 AND 3 USING RT-PCR AND RFLP

MAJDANI R 1; MARDANI K 2; MORSHEDI A 3, ALIABAD FN 1

and MARANDI MV 4

1 Institute of Bioscience and Biotechnology, Urmia University 2 Department of Food Hygiene and Quality Control, Faculty of Veterinary Medicine,

Urmia University 3 Department of Microbiology, Faculty of Veterinary Medicine, Urmia University

4 Department of clinical science, Faculty of Veterinary Medicine, The University of Teheran

SUMMARY Infectious bronchitis is a highly viral contagious disease of poultry worldwide. Prevention of the disease heavily depends on vaccination using attenuated viruses prepared according to circulating strains in the region. Rapid detection and differentiation of infectious bronchitis virus (IBV) involved in the disease outbreak is very important for controlling of the disease and developing new vaccines. In the present study, two fragments of 1.8 kb (S1 gene) and 1.2 kb (entire gene 3) of IBV vaccine strains and field isolates were amplified using reverse-transcription and polymerase chain reaction (RT-PCR). The amplified fragments were subjected to digestion with two restriction endonuclease enzymes AluI and RsaI. Digesting S1 gene using both enzymes generated four different RFLP patterns, grouping IBV strains into four similar groups, while for gene 3 both AluI and RsaI enzymes generated three RFLP patterns however with different groups. Strains 4/91 and 793/B of same serotype were differentiated from each other using two enzymes based on gene 3 and S1. Strains belonged to different serotypes showed different RFLP patterns for both S1 and 3 genes. S1 gene RFLP analysis using both of AluI and RsaI differentiate IB88, 793/B and 4/91 from each other. Analyzing S1 gene, both enzymes had the same discriminatory power and none of the enzymes differentiated 4/91 vaccine strain from field isolate. This is the first report on the molecular analysis of the gene 3 for IBV strain differentiation. All previous reports were focused on S1 gene analysis for strain differentiation. Our results revealed that RFLP analysis of S1 gene had higher discriminatory power than RFLP analysis of gene 3. This difference is may be due to the smaller size of the gene 3 that has been used in the study (1.8 for S1 and 1.2 for gene 3)


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INTRODUCTION Infectious bronchitis is an acute, highly contagious, viral disease of poultry with worldwide distribution (Cavanagh & Naqi, 2003; Cavanagh, 2005; Dolz, 2006). The primary tissue of IBV infection is the respiratory tract, though some isolates replicate in the kidney and oviduct, resulting in nephritis and reduced egg production (Liu et al., 2006). The genome of IBV, a member of the family Coronaviridae, contains a single-stranded positive sense RNA of 27.6 kb (Boursnell et al. 1987; Farsang et al., 2002). Four major structural proteins, the glycosylated spike (S) protein, the envelope or small membrane (E) protein, the membrane (M) protein, and the phosphorylated nucleocapsid (N) protein make up the IBV virions. Four non-structural proteins (3a, 3b, 5a, and 5b) are also encoded (Cavanagh & Naqi, 2003; Huang & Wang, 2007). Typically, the disease has been controlled with serotype-specific vaccines (Callison et al., 2001). Prevention of the disease heavily depends on vaccination using attenuated viruses prepared according to circulating strains in the region. Although commercial poultry flocks are routinely vaccinated for IBV, outbreaks of infectious bronchitis still happen due to naturally occurring variant viruses that continue to arise (Callison et al., 2005) and although many countries share some common antigenic types, IBV strains within a geographic region are unique and distinct (Bayry et al., 2005). Generally, different serotypes do not cross-protect. Therefore, the serotype of the virus causing the disease must first be determined so that the birds can be properly vaccinated. RT-PCR-RFLP is a rapid test and has led to the identification of a tremendous number of virus isolates, which was not possible with the traditional virus-neutralization test in embryonating eggs (Jack Wood, et al., 2005). The purpose of the present study was to genetically characterize vaccine strains and field isolates of IBV based on gene S1 and 3 RFLP analyses. Genes S1 and 3 of IBV viruses were amplified by RT-PCR. The amplified fragments were differentiated by RFLP analysis. MATERIALS AND METHODS Viruses Two field isolates and 5 vaccine strains were used in this study. Field isolate 793/B was obtained from faculty of veterinary medicine, Tehran University and field isolate 4/91 was provided by Razi Vaccine and Serum Research Institute, Karaj, Tehran. Vaccine strains were all obtained from vaccine manufactures. Viral RNA Extraction Viral RNA was extracted and purified using RNeasy Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. About 50 µl of each virus suspension was used for each extraction, and purified RNA was resuspended in 30 µl elution buffer and used immediately for cDNA synthesis or stored at -70˚C.


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Synthesis of cDNA cDNAs were synthesized in 25 µl reaction mixture, according to previous studies (Mardani et al., 2006) with minor modifications. For each cDNA synthesis reaction 5µl of extracted RNA was mixed with 1µl oligo(dt) (25 µM) (Fermentas, Cinnagen, Iran).The premix was incubated at 100˚C for 1min, subsequently the tubes were placed on ice for 5min and 19 µl premix containing 24U RNA guard(Fermentas, Cinnagen, Iran) 50 µM each of dATP, dTTP, dGTP and dCTP, 5µl of 5X reaction buffer(Fermentas) and 200U Moloney murine leukaemia virus reverse transcriptase (Fermentas, Cinnagen, Iran), was added. The reaction mixtures were incubated at 42˚C for 1hour followed by inactivation of the reverse transcriptase enzyme at 100˚C for 5min.The resultant cDNAs was immediately used in a PCR or stored at -70˚C for later use. Polymerase Chain Reaction Amplification of S1 gene The whole S1 gene (1.8kb) of the IBV strains was amplified using forward primer PolyF1 (GATTGTGCATGGTGGACAATG) reported in the previous study (Mardani et al., 2006), binding to the 3énd of polymerase gene (nucleotides 20,070 to 20,090, Beaudette strain) and reverse primer S1-R1 (5’ CCACCAGAAACTACAAACTG 3’) which designed in this study binding to the 3’ end of the S1 gene (nucleotides 21,857 to 21,876, Beaudette strain). The PCR reaction was carried out in 50µl mixture containing 50 µM each of dNTP, 0.5 µM each of primers, 5µl of 10X PCR buffer (CinnaGen, Iran), 2mM magnesium chloride, 5U taq DNA polymerase (CinnaGen, Iran) and 6µl cDNA as template. Amplification was performed using 35 cycles of incubation at 94˚C for 45s, 55˚C for 40s and 72˚C for 2min with a final extension at 72˚C for 5min.The obtained PCR products were separated on1.5% agarose gel and results were observed using ultraviolet transillumination. Amplification of gene3 For the amplification of whole gene 3, two primers S2-F1 (GGTGGAATGATACTAAGCATGG) and M-R1 (ACACCTACTGCAATGTTAAGGG) binding to the 3énd of the S2 gene and 5énd of the M gene respectively were designed according to IBV sequences data in the Genbank. The PCR reaction was carried out in 50 µl mixture containing 50 µM each of the dNTPs, 0.5 µM of each primer, 5 µl of 10X PCR buffer (CinnaGen, Iran), 2mM magnesium chloride, 5U Smartaq DNA polymerase (CinnaGen, Iran) and 6 µl of cDNA as template. Amplification was performed using 35 cycles of incubation at 94˚C for 45s, 55˚C for 40s and 72˚C for 90s, with a final extension at 72˚C for 5min. PCR product purification Before digestion, all PCR products were purified using DNA extraction Kit (Fermentas, Cinnagen, Iran), according to kit’s manufacture instructions. Restriction Fragment Length Polymorphism Purified PCR products for both genes S1 and 3 were digested using two restriction endonuclease enzymes AluI (Fermentas, Cinnagen, Iran) and RsaI (Fermentas, Cinnagen, Iran). For each digestion with AluI about 4µl of purified PCR product and


81

10U of AluI were mixed and incubated at 37˚C for 3 h. For RsaI digestion 5µl purified PCR product and 10U RsaI was mixed and then incubated at 37˚C for 3 h. Digested products were separated on 2% agarose gel and visualized using ultraviolet transillumination. RESULTS Amplification of genes S1 and 3 The first attempt to amplifying gene 3 (1.2Kb) of all IBV field and vaccine strains using was successful. All strains examined in this study gave only one band about 1.2Kb as it was expected. For amplifying S1 gene a few modifications in Mgcl2 concentration and annealing temperature was performed and finally PCR reaction was optimized for amplifying a fragment of 1.8 kb of S1 gene. Restriction endonuclease digestion of S1 gene The RFLP patterns generated from seven IBV strains by digestion with AluI and RsaI are shown in Figures 1 and 2 respectively. Each enzyme produced distinguishable fragments in the range of 100-1000 base pairs. Both enzymes generated four distinct RFLP patterns, grouping IBV strains used in this study in exactly similar groups. Strains H120, H52 and MA5 were generated identical RFLP patterns named pattern I, 4/91 field isolate and 4/91 vaccine strain had the same RFLP pattern named pattern II and strains IB88 and 793/B had two distinct RFLP patterns which named pattern III and IV respectively (Table 1). Restriction endonuclease digestion of gene 3 RFLP analysis of the gene 3 of the seven IBV strains using both AluI and RsaI enzymes generated three RFLP patterns (Figures 3 and 4). However the grouping of the IBVs according to generated RFLP patterns were different based on the enzyme used. Both enzymes grouped H120, H52 and MA5 together named pattern I, exactly same as the results obtained by analysing gene S1. 4/91 vaccine strain and its field isolate showed identical pattern (RFLP pattern II) using both enzymes, however AluI was not able to differentiate IB88 from 793/B (RFLP pattern III) while RsaI was able to discriminate these two strains from each other (Table 1) DISCUSSION One of the major problems with IBV is the frequent emergence of new variants. New variant strains of IBV have appeared all over the world in the last few years (Cook, 1984; Davelaar et al., 1984; King, 1988; Wang et al., 1994; Jia et al., 1995; Wang et al., 1996; Wang and Tsai, 1996; Hung, 2002). Because of low rate of cross-immunity between IBV strains, fast and easy identification and differentiation of new appeared strains in different geographical areas is very important for controlling the disease with appreciate attenuated vaccines against homologous strains. In the present study the RFLP patterns of two different regions of the IBV genome (genes S1 and 3) was compared. All previous reports were focused on S1 gene RFLP analysis for strain differentiation and to date the RFLP analysis of the gene 3 has not been reported elsewhere. In a report by Callison et al., (2001) in the United


82

States, 11 IBV strains from US foreign places analyzed based on their S1 gene RFLP patterns and they showed eight different RFLP patterns. In Korea 15 isolates was analyzed using RT-PCR RFLP analysis of the S1 gene and results revealed four different RFLP patterns (Lee, et al., 2003). Bouqdaoui et al., (2005) reported five different RFLP patterns based on S1 gene analysis of 30 IBV field isolates in Morocco. In this study, two field isolates and seven vaccine strains of IBV were analyzed based on RT-PCR RFLP analysis of genes S1 and 3. According to the S1 gene RFLP analysis using both restriction enzymes, strains H120, H52 and MA5 all from same serotype generated identical patterns while IB88 and 793/B again belong to the same serotype, generated different RFLP patterns which indicates the variation in the spike protein within the same serotype as reported by Cavanagh et al. (2005). According to the RT-PCR RFLP analysis of gene 3, IBV strains were classified in three different groups using both AluI and RsaI. Our results revealed that RFLP analysis of S1gene had higher discriminatory power than RFLP analysis of gene 3. This difference is may be due to the smaller size of the gene 3 that has been used in the study (1.8 for S1 and 1.2 for gene 3). Both genes had differentiated IBV strains and they can be used for IBV differentiation. Analysis of S1gene is better than the gene 3 for IBV discrimination but in the cases that S1 gene amplification was not successful because of the sequence variation gene 3 could be used for strain identification. For gene 3, using both enzymes will increase the discrimination power to the same level as S1 gene. ACKNOWLEDGMENTS We would like to sincerely thank Mr M. Azizi, Mr A. Kazemnia and Mr F. Farhangpagoh for their technical assistance. REFERENCES Bayry, J., Goudar, M.S., Nighot, P.K., Kshirsagar, S.G., Ladman, B.S., Gelb, J., Jr.,

Ghalsasi, G.R., Kolte, G.N., (2005). Emergence of a nephropathogenic avian infectious bronchitis virus with a novel genotype in India. Journal of clinical microbiology 43, 916-918.

Bochkov Y. A., Tosi, G., Massi, P. and Drygin, V. V. (2007) .Phylogenetic analysis of partial S1 and N gene sequences of infectious bronchitis virus isolates from Italy revealed genetic diversity and recombination, Virus Genes, 35:65–71.

Bouqdaoui, M.E., Mhand, R.A., Bouayoune, H. & Ennaji, M.M. (2005). Genetic Grouping of Nephropathogenic Avian Infectious Bronchitis. International Journal of Poultry Science 4, 721-727.

Callison, S.A., Jackwood, M.W., Hilt, D.A. (2001). Molecular characterization of infectious bronchitis virus isolates foreign to the United States and comparison with United States isolates. Avian Disease, 45, 492-499.


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Callison, S., Hilt, D., Jackwood, M., (2005). Using DNA shuffling to create novel infectious bronchitis virus s1 genes: implications for s1 gene recombination. Virus Genes 31, 5-11.

Cavanagh, D., (2005). Coronaviruses in poultry and other birds. Avian Pathology, 34, 439-448.

Cavanagh, D., Davis, P.J., (1986). Coronavirus IBV: removal of spike glycopolypeptide S1 by urea abolishes infectivity and haemagglutination but not attachment to cells. Journal of General Virology, 67, 1443-1448.

Cavanagh, D., Naqi, S.A., (2003), Infectious Bronchitis, In: Saif, Y.M., Barnes, H.J., Glisson, J.R., Fadly, A.M., Mcdougald, L.R., Swayne, D.E. (Eds.) Diseases of Poultry. Iowa State University Press, Ames, pp. 101-119.

Cavanagh, D., Picault, J.P., Gough, R., Hess, M., Mawditt, K., Britton, P., (2005). Variation in the spike protein of the 793/B type of infectious bronchitis virus, in the field and during alternate passage in chickens and embryonated eggs. Avian Pathology, 34, 20-25.

Chang, P.C., Lee, Y.L., H., S.J., K., S.H., (1997). Rapid differentiation of vaccine strains and field isolates of infectious laryngotracheitis virus by restriction fragment length polymorphism of PCR products. Journal of Virological Methods, 66, 179-186.

Dolz, R., Pujols, J., Ordonez, G., Porta, R., Majo, N., (2006). Antigenic and molecular characterization of isolates of the Italy 02 infectious bronchitis virus genotype. Avian Pathology, 35, 77-85.

Farsang, A., Ros, C., Renstrom, L.H., Baule, C., Soos, T., Belak, S., (2002). Molecular epizootiology of infectious bronchitis virus in Sweden indicating the involvement of a vaccine strain. Avian Pathology. 31, 229-236.

Gelb Jr, J., Weisman, Y., Ladman, B.S., Meir, R., (2005). S1 gene characteristics and efficacy of vaccination against infectious bronchitis virus field isolates from the United States and Israel (1996 to 2000). Avian Pathology. 34, 194-203.

Huang, Y.P., Wang, C.H., (2007). Sequence changes of infectious bronchitis virus isolates in the 3' 7.3 kb of the genome after attenuating passage in embryonated eggs. Avian Pathology, 36, 59-67.

Jackwood, M.W., Hilt, D.A., Lee, C.W., Kwon, H.M., Callison, S.A., Moore, K.M., Moscoso, H., Sellers, H., Thayer, S., (2005). Data from 11 years of molecular typing infectious bronchitis virus field isolates. Avian Diseases, 49, 614-618.

Kwon, H.M., Jackwood, M.W., Gelb, J., Jr., (1993). Differentiation of infectious bronchitis virus serotypes using polymerase chain reaction and restriction fragment length polymorphism analysis. Avian Diseases, 37, 194-202.

Lee, S.K., Sung, H.W., Kwon, H.M., (2004). S1 glycoprotein gene analysis of infectious bronchitis viruses isolated in Korea. Archives of Virology, 149, 481-494.

Liu, S., Han, Z., Chen, J., Liu, X., Shao, Y., Kong, X., Tong, G., Rong, J., (2007), S1 gene sequence heterogeneity of a pathogenic infectious bronchitis virus strain and its embryo-passaged, attenuated derivatives. Avian Pathology. 36, 231-234.

Liu, H.J., Lee, L.H., Shih, W.L., Lin, M.Y., Liao, M.H., (2003). Detection of infectious bronchitis virus by multiplex polymerase chain reaction and sequence analysis. Journal of Virolgical Methods, 109, 31-37.


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Liu, S.W., Zhang, Q.X., Chen, J.D., Han, Z.X., Liu, X., Feng, L., Shao, Y.H., Rong, J.G., Kong, X.G., Tong, G.Z., (2006). Genetic diversity of avian infectious bronchitis coronavirus strains isolated in China between 1995 and 2004. Archives of Virology, 151, 1133-1148.

Mardani, K., Browning, G.F., Ignjatovic, J., Noormohammadi, A.H., (2006). Rapid differentiation of current infectious bronchitis virus vaccine strains and field isolates in Australia. Australian Veterinary Journal, 84, 59-62.

Mardani, K., Noormohammadi, A.H., Ignatovic, J., Browning, G.F., (2006), Typing infectious bronchitis virus strains using reverse transcription-polymerase chain reaction and restriction fragment length polymorphism analysis to compare the 3' 7.5 kb of their genomes. Avian Pathology, 35, 63-69.

Mase, M., Tsukamoto, K., Imai, K., Yamaguchi, S., (2004). Phylogenetic analysis of avian infectious bronchitis virus strains isolated in Japan. Archives of Virology. 149, 2069-2078.

Meir, R., Rosenblut, E., Perl, S., Kass, N., Ayali, G., Perk, S., Hemsani, E., (2004), Identification of a novel nephropathogenic infectious bronchitis virus in Israel. Avian Diseases, 48, 635-641.

Wang, X. & Khan, M. (2000). Use of reverse transcriptase–polymerase chain reaction–restriction fragment length polymorphism to examine the interaction between infectious bronchitis virus strains Massachusetts 41 and JMK in ovo. Avian Pathology, 29, 441–448.

Table 1. RT-PCR RFLP patterns of IBV genes S1 and 3 generated using AluI and RsaI enzymes

Patterns Enzyme Genes

I. H120, H52, MA5 II. 4/91 (F), 4/91 (V) III. IB88 IV. 793/B

AluI

I. H120, H52, MA5 II. 4/91 (F), 4/91 (V) III. IB88 IV. 793/B

RsaI

S1

I. H120, H52, MA5 II. 4/91 (F), 4/91 (V) III. IB88, 793/B

AluI

I. H120, H52, MA5 II. 4/91 (F), 4/91 (V), 793/B III. IB88

RsaI

3


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Figure 1. RFLP patterns of the gene S1 of IBV vaccine strains and field isolates generated using AluI.

Figure 2. RFLP patterns of the gene S1 of IBV vaccine strains and field isolates generated using RsaI.


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Figure 3. RFLP patterns of the gene 3 of IBV vaccine strains and field isolates generated using AluI.

Figure 4. RFLP patterns of the gene 3 of IBV vaccine strains and field isolates generated using RsaI.


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INFECTIOUS BRONCHITIS VIRUS (IBV) INDUCES NF-KB SIGNALLING

McCRORY SA, MACDONALD A and HISCOX JA

Faculty of Biological Sciences, University of Leeds, UK. NF-KB signaling is part of the innate cellular response to virus infection. Little is known about this pathway in with regard to coronavirus infection with conflicting data from over-expression studies of severe acute respiratory coronavirus (SARS-CoV) proteins. The response to IBV was examined in model cell lines. NF-KB dependent promoter based assays coupled with immuno-fluorescence analysis revealed that NF-KB was both up regulated and trafficked from the cytoplasm to the nucleus in IBV infected cells. Both downstream and upstream components reveal how this pathway functions during infection.


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GENETIC AND PHENOTYPIC VARIATION OF INFECTIOUS BRONCHITIS VIRUS WITHIN THE HOST

GALLARDO RA, VAN SANTEN VL and TORO H

Department of Pathobiology, College of Veterinary Medicine, 264 Greene Hall, Auburn University, AL 36849, USA.

SUMMARY

The spike (S) protein, responsible for viral attachment, shows genetic and phenotypic variability among infectious bronchitis coronavirus (IBV) populations. We previously found different degrees of genetic heterogeneity among four commercial Ark-DPI-derived IBV vaccines before passage in chickens, reflected in the genes encoding the S1 subunit of the S protein. For three vaccines, a single subpopulation with an S gene sequence distinct from the vaccine predominant consensus was found in tears, trachea, and/or Harderian glands of chickens within 3 days after ocular vaccination. This finding suggests that a distinct virus subpopulation was positively selected by the chicken upper respiratory tract.

We hypothesized that the dominant genotype/phenotype further changes during host invasion as the environment of distinct tissues exert selective pressure on the replicating virus population. To address this hypothesis, we inoculated chickens with 105 50% egg infectious doses of an Ark-type IBV commercial vaccine via the ocular and nasal routes. The first 751 nucleotides of the S1 gene of IBV contained in lachrymal fluid, trachea, and oviduct/testis of individual chickens at different times post-inoculation were amplified by RT-PCR and sequences determined.

Based on the S1 consensus sequences obtained from the chicken tissues, six distinct predominant IBV populations with non-synonymous nucleotide differences were observed. The deduced amino acids at positions that define the predominant populations are shown in table 1. Consistent with our previous results (van Santen and Toro, 2008), the predominant IBV population contained in the vaccine (prior to inoculation), became a very minor or non-detectable population at all times in all tissues after replication in the vast majority (>90%) of the chickens. Interestingly, we observed significant differences in the incidence of IBV predominant populations in tears, trachea, and the reproductive tract in chickens (Fig.1). As seen in this figure, while the population named component 1 (C1) showed increased incidence in the tears and reproductive tract (oviduct/testis) of inoculated chickens, this population


89

was found in significantly fewer chicken tracheas (P<0.05). On the other hand, C5 was more frequent (P<0.05) in the reproductive tract of chickens than in either tears or trachea. C4 was highly selected in tears, trachea and reproductive tract without significant differences among these tissues (P>0.05).

These results corroborate previous observations that the predominant IBV population contained in the vaccine is rapidly negatively selected in the host. These results also indicate that intraspatial variation indeed occurs in the host and thus the dominant genotype/phenotype further changes during host invasion as the environment of distinct tissues exert selective pressure on the replicating virus population. REFERENCES Van Santen, V. L. and H. Toro (2008). Rapid selection in chickens of subpopulations

within ArkDPI-derived infectious bronchitis virus vaccines. Avian Pathology 37 (3):293-306.

Full length article is being submitted for publication elsewhere.


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Table 1. Primers used to amplify the first 751 nucleotides of the S1 gene of IBV.

Name Sequence Orientation Size

S17F 5’-TGAAAACTGAACAAAAGACCGACTTAG-3’ Forward 27

S18R 5’-GGATAGAAGCCATCTGAAAAATTGC-3’ Reverse 25 Table 2. Deduced amino acids at positions that define the predominant IBV populations detected in tissues of chickens inoculated with an Ark-type IBV vaccine strain.

Designation Position

nta 127 167 226 355 388 511 593 637 aab 43 56 76 119 130 171 198 213

Vaccinec Tyr Asn Leu Ser Ser Tyr Lys Ser

C1d His Asn Leu Ser Gly Tyr Lys Ala C2 His Asn Leu Ser Ser Tyr Lys Ala C3 His Asn Phe Pro Ser His Thr Ala C4 His Ser Leu Ser Ser Tyr Lys Ala C5 Tyr Asn Leu Ser Ser Tyr Lys Ala

a Nucleotide; b amino acid; c vaccine predominant IBV population prior to inoculation i.e. prior to host selection; d selected populations were designated C1 through C5.


91

Vaccin

e-L

Vaccin

e-T

Vaccin

e-R C1-L C1-T C1-R C2-L C2-T C2-R C3-L C3-T C3-R C4-L C4-T C4-R C5-L C5-T C5-R0

10

20

30

40

50

a

b

c

aa

b

Population

Sele

cted

(%)

Fig. 1. Frequency of distinct IBV subpopulations in tissues (T=trachea; R=oviduct/testis) or fluids (L=lachrymal fluid) detected in SPF chickens inoculated with an Ark-type IBV vaccine strain on day 14 of age by the ocular/nasal routes. Vaccine=Predominant sequence detected in the vaccine prior to inoculation. C1 through C5 = populations showing nucleotide changes resulting in non-synonymous changes (table 2). While IBV subpopulation named C1 shows an increased incidence in the tears and reproductive tract of most chickens, this population was found in the trachea of significantly fewer (P<0.05) birds. C5 was more frequent in the reproductive tract of chickens than in either tears or trachea (P<0.05). C4 was highly selected in tears, trachea and reproductive tract without significant differences (P>0.05) among these tissues.


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SIGNIFICANCE OF MINOR VIRAL SUBPOPULATIONS WITHIN ARK-TYPE INFECTIOUS BRONCHITIS VACCINES

VAN SANTEN VL, NDEGWA EN, JOINER KS, TORO H and VAN GINKEL FW

Auburn University, Department of Pathobiology, 264 Greene Hall, Auburn, AL, 36849-5519,

SUMMARY We previously found that commercial Ark-DPI derived vaccines contained different relative proportions of a minor subpopulation that was rapidly selected in the upper respiratory tract of chickens (van Santen & Toro, 2008). The minor subpopulation that was selected in chickens had an S1 gene sequence more similar to the virulent parental ArkDPI isolate than to the predominant vaccine population. Two Ark-DPI-derived vaccines (coded A and C) contained the selected subpopulation in amounts readily detectable in the RT-PCR product of the entire population. In vaccine C, the selected subpopulation was the predominant one. In another two vaccines (coded B and D), the selected subpopulation was rare, detectable only by RT-PCR with primers specific for the selected subpopulation. We hypothesized that the ArkDPI-derived vaccines containing higher proportions of the subpopulation(s) efficiently replicating in the upper respiratory tract of chickens (vaccines A and C) produce higher viral loads, more severe “vaccine reactions”, and a more rapid and/or vigorous immune response compared to the vaccines containing a very small proportion of the virus subpopulation able to efficiently replicate in chickens (vaccines B and D). We vaccinated four groups of 1-day-old SPF chickens via the ocular and nasal routes with 4 X 104 EID50 of each of the four vaccines and compared viral load [assessed by qRT-PCR (van Ginkel et. al, 2008)] in tears collected 3, 5, 8, 11, and 15 days post-vaccination (DPV); incidence of respiratory signs assessed daily 4-7 DPV, and 9, 11, and 14 DPV; and histopathological lesions in the trachea 3, 7, 10, 14, 17, and 28 DPV. Viral subpopulations present in each chicken were monitored by S1 gene sequence analysis. In addition, we monitored the mucosal immune response by assessing IgA and interferon gamma (IFN) mRNA expression in Harderian glands by qRT-PCR 3, 7, 10, 14, and 17 DPV. We found significantly higher (P<0.05) viral loads 5 and 8 DPV in tears of chickens vaccinated with vaccines A and C than in chickens vaccinated with vaccines B and D (Fig. 1A). Furthermore, when viral loads in tears of chickens of all vaccine groups were compared based on whether selected viral subpopulations predominated, significantly higher (P<0.05) viral loads were found in chickens with selected viral


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subpopulations at 3 and 5 DPV (Fig. 1B). Chickens vaccinated with vaccines A and C had consistently higher incidence of respiratory signs 4-7 DPV than chickens vaccinated with vaccine D (Fig 2). At 7 DPV, chickens vaccinated with vaccine C had more severe epithelial necrosis (Fig. 3A) and deciliation (Fig. 3B) in the trachea than chickens vaccinated with vaccine D, and more lymphocytic infiltration of the trachea (Fig. 3C) than chickens vaccinated with the other three vaccines. The relatively high incidence of respiratory signs in vaccine group B 5, 6 and 11 DPV and the relatively high levels of tracheal lesions in vaccine group B 6 DPV compared to vaccine group D were unexpected, because chickens vaccinated with vaccines B and D had similar viral loads in tears at all days tested. However, results for vaccine group B might be explained by the relative viral load in the tears not reflecting the viral load in the trachea. Chickens vaccinated with vaccines A and C had significantly higher IgA mRNA levels in Harderian glands 10 and 14 DPV than chickens vaccinated with vaccines B and D (Fig. 4A). In contrast, expected differences in IFN mRNA levels were not noted among groups (Fig. 4B). Thus, we demonstrated that IBV vaccines containing predominant viral subpopulations more efficiently replicating in the upper respiratory tract of chickens induce more severe respiratory signs and lesions, higher viral load, and increased mucosal IgA. From an applied perspective, these results indicate that the presence in Ark-DPI derived vaccines of varying levels of IBV able to efficiently replicate in chickens likely influences the outcome of vaccination in commercial operations. REFERENCES Van Ginkel, FW, van Santen, VL, Gulley, SL & Toro, H (2008) Infectious bronchitis

virus in the chicken Harderian gland and lachrymal fluid: viral load, infectivity, immune cell responses, and effects of viral immunodeficiency. Avian Diseases 52:608-617.

Van Santen, VL & Toro, H (2008) Rapid selection in chickens of subpopulations within ArkDPI-derived infectious bronchitis virus vaccines. Avian Pathology 37:293-306.


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Fig. 1: Viral load in tears. Tears were collected from 10 individual chickens in each group on the days post-vaccination indicated. The number of viral genomes in each sample was determined by qRT-PCR. The viral subpopulation(s) present in each sample were characterized by sequence analysis of the S1 gene. A. Samples are divided by vaccine group. Groups were compared for each day by ANOVA with Tukey’s post-test B. Samples are divided by whether the viral subpopulation was predominantly “selected” or not. Samples with selected virus were compared to those with unselected virus for each day by Students T –test. After 8 days post vaccination, insufficient samples without selected virus were available for statistical analysis. Error bars in both panels represent SEM.

Fig. 2: Incidence of respiratory signs. All chickens in each vaccine group were assessed individually on the days indicated. The presence or absence of respiratory signs (tracheal or nasal rales) was determined by bringing the head of each chicken close to the observer’s ear. At 4 DPV there were 30 chickens per group. Due to removal of 6 chickens/group for necropsy sampling every 3-4 days, the number of chickens per group was reduced to 12/group by day 14. Incidence of respiratory signs was compared between groups by Fisher’s exact test. For 3-7 DPV, group C had significantly higher (P=0.003) incidence of respiratory signs than group D. For 4-7 DPV, group A had consistently higher incidence of respiratory signs than group D (P=0.09).

A B


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Fig. 3: Histopathology of trachea. Six chickens in each vaccine group were necropsied on the days post-vaccination indicated and trachea samples routinely processed for histopathological examination. A, B. Lesion scores from 1 (normal) to 5 (severe) for necrosis (A) and deciliation (B) were assigned. Lesion scores for vaccine group C 6 DPV were significantly higher (P<0.05) than vaccine group D. C. The degree of lymphocytic infiltration was assessed by measuring the thickness of the lymphocytic infiltrate layer at five points for each trachea. The

lymphocytic infiltrate was significantly thicker for vaccine group C 6 DPV than for all other groups.

Fig. 4: Immune response gene expression in Harderian glands. Six chickens in each vaccine group were necropsied on the days post-vaccination indicated. cDNA to RNA extracted from Harderian glands of individual chickens was synthesized using random primers and the level of IgA heavy chain (A) and IFN (B) cDNA determined by qPCR. Resuls were normalized to ß-actin cDNA levels. Most samples had undetectable levels of IgA cDNA 7 DPV and IFN cDNA 3 DPV.

A B

C

A B


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ARE THE STRUCTURAL AND ACCESSORY GENES OF INFECTIOUS BRONCHITIS VIRUS RESPONSIBLE FOR PATHOGENESIS?

ARMESTO M, BRITTON P and CAVANAGH D

Coronavirus Group, Institute for Animal Health, Compton, Newbury,

Berkshire RG20 7NN, UK. SUMMARY Utilizing our reverse genetics system for IBV we have shown that replacement of the spike gene of the non-pathogenic strain Beaudette (Beau-R) with that of either M41 or UK/4/91 did not alter pathogenicity in chickens. In order to determine whether any of the other IBV structural protein genes, envelope protein (E), membrane protein (M) and nucleoprotein (N) or the two accessory gene clusters, gene 3 encoding the 3a and 3b proteins or gene 5 encoding the 5a and 5b proteins were involved in pathogenicity or in the loss of virulence associated with attenuation in Beaudette, we replaced all the genes downstream from the replicase gene of Beau-R with those of the pathogenic strain M41 to produce a chimaeric IBV: Beau-Rep-M41-Struct. We assessed the pathogenicity of Beau-Rep-M41-Struct in vivo but found no differences in pathogenicity between the chimaeric Beau-Rep-M41-Struct and its apathogenic parent Beau-R (based on snicking, nasal discharge, wheezing, watery eyes, rales and ciliostasis in trachea). Interestingly, Beau-Rep-M41-Struct did not cause ciliostasis in tracheal organ cultures although the virus was able to replicate. Multiple passages (n= 25) of the virus in chicken kidney cells (CKC) resulted in slightly better growth but still did not cause ciliostasis. Sequence analysis showed that there were no changes either in the 3’ or the 5’-end of the passed chimaeric virus. Our studies suggest that one or more of the 15 non-structural proteins encoded by the IBV replicase gene are involved in pathogenicity. INTRODUCTION We have previously shown using our reverse genetics system (Armesto et al., 2008; Britton et al., 2005; Casais et al., 2001) that replacement of the Beaudette S gene with the corresponding S gene from the virulent M41 strain of M41 resulted in a recombinant IBV (rIBV) that had the cell tropism associated with M41 but did not alter the pathogenicity. Similar experiments but using the S gene from the pathogenic UK/4/91 strain produced similar results indicating that although the S gene may play a role in virulence it was unable to restore virulence in a non-pathogenic strain


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suggesting that the attenuation associated with the Beaudette strain resides elsewhere within the virus genome. Loss of virulence for other coronaviruses has been shown to result from modification or deletion of the other structural or accessory genes. To determine whether the other Beaudette structural or the Beaudette accessory genes may be responsible for attenuation we decided to replace all the Beaudette structural and accessory genes with the corresponding sequences from M41 using our IBV reverse genetics system. MATERIALS and METHODS Viruses The Beaudette-CK (Beau-CK; (Cavanagh et al., 1986)) strain of IBV is apathogenic in chickens resulting from multiple passages in embryonated eggs, the virus has been virus adapted for growth in chick kidney (CK) cell cultures and can be grown in Vero cells, an African green monkey cell line. Beau-R, a recombinant IBV produced from an infectious RNA transcribed from a full-length cDNA of Beau-CK (Casais et al., 2001) and the same properties as Beau-CK. The M41-CK strain of IBV is an isolate derived from M41 (Darbyshire et al., 1979) following adaption to growth on CK cells. The virus is pathogenic in chickens resulting symptoms associated with the disease infectious bronchitis. Although M41-CK can grow in CK cells it does not produce infectious virus in Vero cells. Recombination and rescue of rIBVs The rIBV Beau-Rep-M41-Struct was produced using our IBV reverse genetics system in which the structural and accessory genes of the Beau-R cDNA within the vaccinia virus genome were deleted and replaced with the corresponding sequences from M41-CK by homologous recombination using the transient dominant selection (TDS; (Armesto et al., 2008; Britton et al., 2005)). The modified IBV cDNA in the vaccinia virus genome was then used for the rescue of rIBVs in CK cells using rFPV/T7 (Britton et al., 1996) for the generation of infectious IBV RNA (Armesto et al., 2008; Britton et al., 2005; Casais et al., 2001). Resultant chimaeric rIBVs were passed three times in CK cells before being used in subsequent experiments. Characterization of recombinant IBVs The criteria used to measure pathogenicity were snicking, tracheal rales, wheezing, nasal discharge, watery eyes and ciliary activity of the trachea following eye drop and nasal inoculation of 1-day-old chicks (Hodgson et al., 2004). Chicks were observed daily for clinical signs; snicks, the presence of tracheal rales, nasal discharge, watery eyes and wheezing. Tracheas were removed from randomly selected chickens from each group at 4 and 7 days post-inoculation for assessment of ciliary activity.


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RESULTS and DISCUSSION The results of our experiments will be described in full elsewhere. Following the replacement of the Beau-R sequence encoding the structural and accessory protein genes with the corresponding sequence from M41-CK in the cDNA within the vaccinia virus genome, we used the vaccinia virus-derived DNA to rescue several rIBVs consisting of the replicase gene from Beaudette and the structural and accessory genes from M41-CK. One of the rIBVs, Beau-Rep-M41-Struct, was passaged thee times in CK cells and sequenced to confirm that the virus contained the chimaeric genome and used for experiments to determine the pathogenicity of the rIBV. Analysis of chicks either mock-infected or infected with Beau-R, M41-CK or the rIBV Beau-Rep-M41-Struct showed that only chicks infected with M41-CK showed clinical signs associated with an IBV infection. The tracheas isolated from either mock-infected chicks or chicks infected with Beau-R or Beau-Rep-M41-Struct had >95% ciliary activity. In contrast the tracheas taken from chicks infected with M41-CK showed 0% ciliary activity (100% ciliostasis). Analysis of tracheal epithelial cells scraped from the tracheas either for the presence of virus or IBV-derived RNA by RT-PCR confirmed that only the tracheas taken from chicks infected with M41-CK contained virus by titration on tracheal organ cultures or detectable RNA by RT-PCR. In conclusion, from the parameters used to assess pathogenicity, our results have demonstrated that the chimaeric rIBV, Beau-Rep-M41-Struct consisting of the replicase derived from Beau-R and the structural and accessory genes from M41-CK, was not pathogenic. Our studies showed that replacement of the Beaudette structural and accessory genes did not restore virulence indicating that one or more of the 15 non-structural proteins encoded by the IBV Beaudette replicase gene are involved in loss of virulence associated with this virus. ACKNOWLEDGEMENTS This work was supported by the Department of Environment, Food and Rural Affairs (DEFRA) project codes OD0714 & OD0717 and the Biotechnology and Biological Sciences Research Council (BBSRC).


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REFERENCES Armesto, M., Casais, R., Cavanagh, D. & Britton, P. (2008). Transient dominant

selection for the modification and generation of recombinant infectious bronchitis coronaviruses. In SARS- and Other Coronaviruses: Laboratory Protocols, pp. 255-273. Edited by D. Cavanagh: Humana Press.

Britton, P., Evans, S., Dove, B., Davies, M., Casais, R. & Cavanagh, D. (2005). Generation of a recombinant avian coronavirus infectious bronchitis virus using transient dominant selection. Journal of Virological Methods 123, 203-211.

Britton, P., Green, P., Kottier, S., Mawditt, K. L., Pénzes, Z., Cavanagh, D. & Skinner, M. A. (1996). Expression of bacteriophage T7 RNA polymerase in avian and mammalian cells by a recombinant fowlpox virus. Journal of General Virology 77, 963-967.

Casais, R., Thiel, V., Siddell, S. G., Cavanagh, D. & Britton, P. (2001). Reverse genetics system for the avian coronavirus infectious bronchitis virus. Journal of Virology 75, 12359-12369.

Cavanagh, D., Davis, P. J., Pappin, D. J. C., Binns, M. M., Boursnell, M. E. G. & Brown, T. D. K. (1986). Coronavirus IBV: partial amino terminal sequencing of spike polypeptide S2 identifies the sequence Arg-Arg-Phe-Arg-Arg at the cleavage site of the spike precursor propolypeptide of IBV strains Beaudette and M41. Virus Research 4, 133-143.

Darbyshire, J. H., Rowell, J. G., Cook, J. K. A. & Peters, R. W. (1979). Taxonomic studies on strains of avian infectious bronchitis virus using neutralisation tests in tracheal organ cultures. Archives of Virology 61, 227-238.

Hodgson, T., Casais, R., Dove, B., Britton, P. & Cavanagh, D. (2004). Recombinant infectious bronchitis coronavirus Beaudette with the spike protein gene of the pathogenic M41 strain remains attenuated but induces protective immunity. Journal of Virology 78, 13804-13811.


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IMPORTANCE OF SIALIC ACID FOR THE INFECTION OF THE TRACHEAL EPITHELIUM BY DIFFERENT STRAINS OF INFECTIOUS BRONCHITIS VIRUS

ABD EL RAHMAN S1, 2, NEUMANN U3, HERRLER G1 and WINTER C1,3

1Institute of Virology, 3Clinic of Poultry, University of Veterinary Medicine Hannover, Bünteweg 17, 30559 Hannover, Germany.

2Department of Virology, Faculty of Veterinary Medicine, Mansoura University, Mansoura, Egypt.

SUMMARY Avian infectious bronchitis virus (IBV) is a major pathogen in commercial poultry flocks. We recently demonstrated that sialic acid serves as a receptor determinant for IBV on the tracheal epithelium. Here we compared the IBV strains Beaudette, 4/91, Italy02, and QX for their sialic acid-binding properties. We demonstrate that sialic acid binding is important for the infection of primary chicken kidney cells and the tracheal epithelium by all four strains. There were only slight differences between the four strains, indicating the universal usage of sialic acids as receptor determinants by IBV. In addition, we analysed the primary target cells in the respiratory epithelium of the four different strains and found that all of them infected ciliated cells and goblet cells. INTRODUCTION Avian infectious bronchitis virus (IBV) causes an economically important disease mainly in the chicken and is clinically associated with respiratory symptoms. IBV uses the respiratory epithelium of the upper respiratory tract as primary target cells. In the course of infection, the virus can spread to several other organs including kidney and oviduct. Belonging to the family Coronaviridae, IB virions comprise four structural proteins: the nucleoprotein N, the membrane proteins M, E, and S. The S protein is composed of two subunits, the S1 ‘‘head’’ and the S2 ‘‘stalk’’, and is responsible for binding to and fusion with host cells membranes. The important viral binding domain appears to be located on the S1 subunit of the S protein. We recently demonstrated that sialic acid serves as receptor determinant for infection of cells (Winter et al., 2006, 2008). One major characteristic of IBV virions is the high variability of the S1 subunit and the occurrence of new strains. Most serotypes differ by 20% to 25% from each other but some serotypes show a difference of 50% in the S1 sequence (reviewed in Cavanagh, 2007).


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MATERIALS and METHODS Viruses IBV strains Beaudette, 4/91, Italy 02, and QX were propagated in specific pathogen-free 10-day-old embryonated chicken eggs. The allantoic fluid was harvested, clarified by low-speed centrifugation, frozen in liquid nitrogen and stored at -80oC. The virus titre was determined by titration in primary chicken embryo kidney cells. All strains were kindly provided by Hans Philipp (Lohmann Tierzucht, Cuxhaven, Germany) except for the Beaudette strain, which was kindly given by Dave Cavanagh (Institute for Animal Health, Compton, UK). Cells Primary chick embryo kidney cells were prepared from 20-day-old specific pathogen-free chicken embryos as described previously (Winter et al., 2006). Preparation of tracheal organ cell culture The tracheal organ cell cultures (TOCs) were prepared as described previously (Winter et al., 2008). The rings were incubated at 37oC on a rotator, and on the next day the rings were screened for selection of TOCs with 100% ciliary activity. Preparation of cryosections Tracheas were prepared from 4- to 6-week-old specific pathogen-free chickens, cut into small pieces of 1 cm length, washed with phosphate-buffered saline (PBS) and infected by one of the following IBV inocula: strains Beaudette and QX were applied at a titre of 105 plaque-forming units/ring, and strains Italy 02 and 4/91 applied at 5 x105 plaque-forming units/ring. The infected rings were incubated at 37oC on a rotator. After 24 h they were mounted on small filter papers with tissue-freezing medium (Jung, Heidelberg, Germany), frozen in liquid nitrogen and preserved at –80 oC prior to cutting. Sections of 10 mm thickness were obtained with a cryostat machine (Reichert-Jung, Nußloch, Germany). The sections dried overnight at room temperature and kept at –20 oC until they were subjected to the staining procedure. Plaque reduction assay Primary chick embryo kidney cells grown on cover slips were treated with 50 mU neuraminidase of Clostridium perfringens type 5 (5 or V?) ( Sigma-Aldrich, St Louis, Missouri, USA) diluted in medium 199, or mock-treated, for 1 h at 37oC. After washing, the cells were infected by the indicated IBV strains at a multiplicity of infection of 0.001. Methylcellulose was added after an adsorption period of 1 h at 37oC. After 24 h, the cells were fixed with 3% paraformaldehyde and permeabilized with 0.2% Triton X-100. Plaques were stained using an anti-IBV polyclonal serum raised in rabbits and a FITC-labelled secondary antibody. After counting the plaques, the outcome of the mock-treated cells (without neuraminidase) was set as 100%. Data shown are the mean values of three experiments. Neuraminidase treatment and infection of TOCs Two groups of four TOCs were either treated with 50 mU neuraminidase of C. perfringens type 5 (5 or V?) per ring or incubated with medium alone. After incubation at 37oC for 1 h, the rings were washed with PBS and infection with the indicated virus


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strain was performed using 1 x 104 (pfu?) per ring. Following three washes with PBS, the TOCs were incubated with medium at 37oC on a rotator. All experiments were performed three times. Ciliary activity was analysed. Briefly, each ring was observed at daily intervals under a microscope to monitor the ciliary activity. The ring was divided virtually into 10 portions and each portion was analysed to see whether ciliary activity was detectable. The number of portions with visible ciliary activity was multiplied by 10 to give the percentage ciliary activity of the respective TOCs. Immunofluorescence analysis of cryosections The sections were fixed with 3% paraformaldehyde for 15 min and were permeabilized with 0.2% Triton X-100 for 5 min followed by three washing steps with PBS. All antibodies were diluted in 1% bovine serum albumin and incubated with the sections for 1 h at room temperature in an incubation chamber. For detection of infected cells, monoclonal antibody Ch/IBV 48.4 directed against the N protein of IBV (ID Lelystad, The Netherlands) or a polyclonal IBV antiserum raised in rabbits was used. Mucus producing goblet cells were stained with anti-MUC-5AC antibody (Acris, Hiddenhausen, Germany). Cilia were detected by CY3 labelled anti-ß-tubulin antibody (Sigma-Aldrich). Bound antibodies were visualized by FITC-labelled and Cy3-labelled anti-rabbit antibodies (Sigma-Aldrich), or anti-mouse antibodies (Acris). For lectin staining, biotinylated Maackia amurensis Lectin II (MAAII) was used after preincubation of sections with the Avidin/Biotin Blocking kit (both from Vector Laboratories, USA). Detection of lectin was carried out with streptavidin Cy3 (Sigma-Aldrich). Fluorescence microscopy was performed with a Leica inverted-2 confocal microscope. RESULTS Plaque reduction after neuraminidase treatment To analyse the importance of sialic acids for the infection of cultured cells for the strains Italy02, 4/91 and QX, we chose primary embryo chick kidney cells because they are sensitive to infection with all strains. The Beaudette strain which has a broader tropism on cultured cells was used as a reference strain. For this strain the importance of sialic acid for infection of embryo chick kidney cells has been described recently (Winter et al., 2006). As shown in Figure 1, desialylation of the cells with 50 mU neuraminidase prior to infection by either of the different strains decreased the number of infected cells by about 50% for Beaudette, 4/91 and QX. After infection with Italy 02, the decrease in the plaque number was about 75%. Importance of sialic acids for the infection of TOCs We have recently shown that desialylation of TOCs results in a delay of ciliostasis after infection by IBV. Here we confirmed this effect with the isolates Italy02, 4/91 and QX (Figure 2). All strains induced ciliostasis after infection of TOCs with 104 plaque-forming units. In the case of QX, the epithelium had completely lost ciliary activity by 2 days post-infection, whereas infection with the other strains resulted in complete ciliostasis at around 5 days post-infection. Prior desialylation of the TOCs prevented complete ciliostasis. At 5 days post infection, more than 50% of the ciliary activity was retained for all strains. This shows the universal importance of sialic acids for the infection of the tracheal epithelium by all IBV strains analysed.


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Infection of the epithelial cells in TOCs We were interested to analyse whether there are strain-dependent differences in the infection of primary target cells by IBV. For this purpose cryosections were prepared from infected TOCs and were stained for virus antigen. In parallel, the sections were immunostained for tubulin (Figure 3), mucin (Figure 4), and for reactivity with the lectin MAA (Figure 5) to visualize ciliated cells, mucus-producing cells, or cells expressing 2,3-linked sialic acids, respectively. Co-staining of cells with antibodies against IBV N protein and ß-tubulin indicated that all strains were able to infect ciliated cells (Figure 3). Co-staining of cryosections for virus antigen and MUC5Ac demonstrated that all strains were able to infect mucus-producing cells (Figure 4). However, the strains differed in their efficiency to infect the tracheal epithelium. Infection by Beaudette and QX resulted in a larger number of infected cells compared with Italy 02 and 4/91 when TOCs were infected with the same amount of virus, which was determined by titration on primary chick kidney cells. To achieve comparable pictures, TOCs infected with Italy 02 and 4/91 were infected with an infectious dose that was five times higher compared with the other strains Sialic acid expression of infected cells When TOCs were analysed for reactivity with lectins, we found an intensive staining of the tracheal epithelium with the lectin MAAII that recognizes 2,3-linked sialic acids (Figure 5). We observed staining of the basal cell layer as well as mucus in goblet cells and a clear staining of the apical membrane. This result showed that in the tracheal epithelium there were many cells expressing 2,3-linked sialic acids. These are potential receptor determinants for the different IBV strains. Interestingly, a co-staining of infected TOCs for virus antigen and sialic acids revealed a weaker MAAII binding in infected tissues, especially on the apical membrane, compared to uninfected samples. DISCUSSION Infection of poultry with IBV is still a matter of economic concern, mainly due to the occurrence of new serotypes with low cross-protection by conventional vaccines. We analysed three major serotypes, which show a high incidence in Europe, for their ability to use sialic acids as a receptor determinant. One characteristic of IBV is that most strains grow only in primary cells derived from their natural host, the chicken. A major factor determining this tropism might be the binding to structures on the cell surface that allow internalization of the virions. In previous work (Winter et al., 2006), we demonstrated that primary chicken kidney cells became resistant to infection by the two IBV strains Beaudette and M41 after sialic acids had been removed from the cell surface by neuraminidase treatment. Here we show for the strains 4/91 and QX that desialylation of chick embryo kidney cells decreases the number of infected cells to the same extent as for the control strain Beaudette. For the strain Italy 02 we observed an even stronger effect. This may be explained by a different efficiency of this strain in the recognition of sialic acid molecules on the cell surface. Interestingly, the protective effect of the neuraminidase treatment observed in Italy 02-infected TOCs was not stronger than that of TOCs infected by the other strains. Strain Italy 02 appears to replicate in TOCs less efficiently than the three other strains. A weaker binding of strain Italy 02 to the sialoglycoconjugates on the tracheal epithelium may


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explain our results. The virus with the strongest ciliostatic effect in TOCs was the QX strain followed by the Beaudette strain. This finding is in accordance with the fact that, with these two viruses, a five times lower amount of virus was required for infection of TOCs to get a similar number of infected cells, compared with Italy 02 and 4/91. This difference makes it all the more astonishing that, for both strains QX and Beaudette, at least 50% of the ciliary activity was retained at 5 days post infection of desialylated cells, similar to the values determined for Italy 02 and 4/91. The finding that neuraminidase treatment could not prevent ciliostasis completely can be explained by incomplete removal of sialic acids from the epithelial cells, so that some cells still are sensitive to infection. We did not observe any differences in the choice of the primary target cells by the analysed strains. Antigen of all strains was detected in ciliated cells as well as in goblet cells. It appears that there is no preference for a certain cell type that could explain the differences in the increased infectivity of QX and Beaudette. The analysis of the tracheal epithelium with the lectin MAAII showed that 2,3-linked sialic acids are abundantly expressed in the epithelial cells. The interesting finding that the MAAII binding to the apical membrane was always reduced after infection by the different IBV strains might be a consequence of an altered glycocalix by the infection. Spike proteins expressed on the cell surface may interact with 2,3-linked sialic acids and thus prevent the lectin from binding to this sugar molecule. The interaction of S protein with 2,3-linked sialic acids may also result in down regulation of the respective sialoglycoconjugates from the cell surface. Such a phenomenon might also explain the observed effect of virus interference after vaccination with different attenuated strains at the same time point (Winterfield & Fadly, 1975). Sialic acids are used by a broad range of pathogenic organisms as a ligand for attachment to cells. IBV appears to follow a similar strategy as, for example, avian influenza viruses to bind to epithelial cells. The fact that all analysed IBV strains use sialic acids to bind to epithelial cells leads to the conclusion that, despite the high variation among the different S proteins, the sialic acid binding activity is a conserved feature of IBV. This does not necessarily mean that all strains bind with the same strength to specific glycans. It could well be that some strains prefer different sialoglycoconjugates than others, and hence have a slightly different organ tropism within the host. The analysis of the recognized sialylated glycans and the identification of the sialic acid binding site in the S protein will be interesting in future work. REFERENCES Cavanagh, D. (2007). Coronavirus avian infectious bronchitis virus. Veterinary

Research, 38, 281_297. Cavanagh, D. & Davis, P.J. (1986). Coronavirus IBV: removal of spike

glycopolypeptide S1 by urea abolishes infectivity and haemagglutination but not attachment to cells. Journal of General Virology, 67, 1443_1448.

Winter, C., Schwegmann-Wessels, C., Cavanagh, D., Neumann, U. & Herrler, G. (2006). Sialic acid is a receptor determinant for infection of cells by avian Infectious bronchitis virus. Journal of General Virology, 87, 1209_1216.

Winter, C., Herrler, G. & Neumann, U. (2008). Infection of the tracheal epithelium by infectious bronchitis virus is sialic acid dependent. Microbes and Infection, 10, 367_373.


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Winterfield, R.W. & Fadly, A.M. (1975). Potential for polyvalent infectious bronchitis vaccines. American Journal of Veterinary Research, 36, 524_526.

Worthington, K.J., Currie, R.J. & Jones, R.C. (2008). A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. Avian Pathology, 37, 247_257.

Fig. 1: Effect of pretreatment of cells with neuraminidase on the infection by different strains of IBV. Chick embryo kidney cells were incubated in the presence (open boxes) or absence (black boxes) of neuraminidase from Clostridium perfringens and then infected by either of the four IBV strains Bd, 4/91, Italy 02, and QX at a multiplicity suitable for a plaque assay. The reduction in the plaque number was used to determine the effect of the enzyme treatment on the infectivity of IBV.


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Fig. 2: Effect of neuraminidase treatment on the infection of TOCs by different strains of IBV. TOCs were incubated in the presence or absence of neuraminidase from Clostridium perfringens and then infected by either of the four IBV strains Bd (A), 4/91 (B), QX (C), and Italy 02 (D) or mock-infected (A-D). Up to five days p.i., TOCs were analyzed for ciliary activity at daily intervals.

Fig. 3: Immunofluorescence analysis of cryosections prepared from IBV infected tracheal organ cultures. At 24 h.p.i., sections were stained with an anti ß-tubulin antibody to detect cilia (red) and with a monoclonal anti-N antibody to visualize virus antigen (green).

4/91


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Fig. 4: Immunofluorescence analysis of cryosections prepared from tracheal organ cultures infected by either of four IBV strains (Bd, 4/91, Italy02, QX). At 24 h.p.i, sections were stained for goblet cells using an anti Muc5AC antibody (red) and for virus antigen using a polyclonal IBV serum (green). Areas of costaining are indicated by white arrows (white arrow at QX in correct position?).

Fig. 5: Immunofluorescence analysis of cryosections prepared from IBV-infected tracheal organ cultures infected by either of four IBV strains (Bd, 4/91, Italy02, QX). At 24 h.p.i, sections were incubated with the lectin MAAII to detect α2,3-linked sialic acid (red). The apical side of the ciliated epithelium is indicated by an arrow in the top left panel. Virus antigen was detected with a monoclonal anti-N anitbody (green). Interestingly, the apical staining with MAAII is always reduced in infected tissues.


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HIGH THROUGHPUT PROTEOME SCREENING OF INFECTIOUS BRONCHITIS VIRUS INFECTED CELLS REVEALS NOVEL HOST-CELL INTERACTIONS

EMMOTT E and HISCOX JA

Faculty of Biological Sciences, University of Leeds, UK.

High throughput proteome screening using stable isotope labeling in cell culture (SILAC) coupled to quantitative MS/MS analysis was used to identify cellular changes in response to infection with the avian coronavirus infectious bronchitis virus (IBV). To reduce sample complexity the cell was separated into nuclear, nucleolar and cytoplasmic fractions. Several thousand proteins were identified with specific functional groups showing ablation and enrichment in different cellular compartments. These included members of cellular signaling cascades, RNA processing enzymes and protein degradation pathways. Additionally this analysis revealed the presence of two viral replicase components within the nuclear compartment.


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THE USE OF FTA® CARDS TO TRANSPORT SAMPLES FOR DIAGNOSIS OF INFECTIOUS BRONCHITIS VIRUS

AND AVIAN METAPNEUMOVIRUS BY RT-PCR

SAVAGE CE, COWLEY K and JONES RJ

Department of Veterinary Pathology, University of Liverpool, Leahurst Campus, Neston CH64 7TE UK

SUMMARY The transporting of samples for the diagnosis of avian respiratory viruses across international borders is complicated because of the possible accidental importation of virulent forms of Newcastle Disease virus (NDV) or avian influenza virus (AIV). FTA® cards (Flinders Technology Associates) by inactivating viruses but allowing their presence to be demonstrated by RT- PCR, provide a convenient way to obviate this problem. Preliminary studies have shown that over time, the detectability of viral RNA declines. We have previously reported that treating infected swabs in a microwave oven for 20 seconds also inactivates IBV and aMPV (in addition to NDV and AIV) but allows detection by RT-PCR. By inoculating untreated filter paper and then microwaving we compared the survival time at three temperatures of the viral RNA on FTA cards and on filter paper after microwaving. Both viruses could be detected after both treatments for at least six weeks. INTRODUCTION Until recently, it has been problematical to send clinical material which may contain live notifiable viruses over international borders. All countries have strict regulations concerning the importation of infectious materials from human, animal or plant sources. The USA originally required that material had to be treated with phenol or formalin (Snyder, 2002) while in the UK an Import Licence has to be granted. In an attempt to obviate this problem we have previously shown that treating swabs infected with poultry respiratory viruses by microwave for 20 seconds inactivates virus but allows detection by RT-PCR (Elhafi et al., 2004). However, FTA cards are now the generally accepted commercial answer to the safe transportation of infectious material. Our laboratory conducted a European RT-PCR Survey on IBV genotypes (Worthington et al., Av. Path. 2008) and receiving the samples did not present any problems as all came from within the European Union. More recently we have been


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receiving samples from further afield, including countries in Latin America and all have all been sent on FTA cards. The number of detections of both IBV and aMPV has been fewer than were expected, and we suspected that there had been deterioration of the RNA during transportation. Thus it was decided to investigate the length of persistence of the viral RNA at three different temperatures on these cards and to compare this with virus inactivated by microwaving. MATERIAL and METHODS Viruses IBV strain M41, and aMPV, subtype B were both grown in tracheal organ culture (TOC) and diluted to 3.4 log10 median ciliostatic doses (CD50)/ml for inoculation. FTA®cards and filter papers Flinders Technology Associates cards were developed by the Whatman Corporation and are designed to be used to transport and store a variety of biological materials including viral and bacteriological samples, blood and imprints of tissue. Infectious material applied to the cards is inactivated on contact as the cells are lysed and nucleic acid is denatured. The samples are preserved at room temperature for PCR analysis. ‘Classic’ FTA cards were used. Each one has four circles for application of samples. Filter paper of the same type and size as in FTA cards were used for microwaving and circles of the same diameter as on FTA cards were drawn on them. Experimental design Both FTA cards and filter papers were inoculated with 60µl of the above dilution of each virus. After drying at room temperature, the filter papers were placed in a domestic microwave for 20 seconds on full power. 5mm circles were punched from both the FTA cards and the filter papers. These were divided up and stored at three temperatures: RT (230C), fridge (40C) or -200C. At each collection time, weekly for 6 weeks, two small discs of each type were taken into elution buffer for 10 minutes at room temperature and RNA was extracted immediately. The RT-PCR based on the S1 spike gene was performed (Worthington et al., 2008) In a separate study to test the sensitivity of FTA cards at room temperature three dilutions of aMPV were used to inoculate the cards and two samples were collected on days 1, 6, and 20 post inoculation. The RNA was extracted by using either Qiagen RNAesy mini Kit or by the phenol/chloroform method. RESULTS IBV could be detected on both the FTA cards and on filter papers after microwave treatment for at least six weeks, the full length of the trial (Table 1) at all three temperatures. aMPV could also be detected for at least 6 weeks but the results were inconsistent in some of the samplings, unlike IBV (Table 2).


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When the sensitivity of FTA cards for aMPV was tested with different dilutions of virus, ability to detect virus was found to be dose-related (Table 3). Thus a titre of 3.6 log10 CD50/ml could be detected using both extraction methods for at least 20 days. The 2.6 log titre was detected at 6 days by both extraction methods. Of the lower titres only 1.6log could be detected at 1 day and only by the Qiagen kit extraction method; 0.6log titre of virus could not be detected at any of the sampling times. Apart from at the 1.6 log dilution, the two detection methods were identical in sensitivity. DISCUSSION The ability of both FTA cards (Moscoso, et al., 2005; Perozo et al., 2006; Purvis et al., 2006) and microwave treatment (Elhafi et al., 2004) to inactivate these and other avian viruses but allowing nucleic acid to remain detectable by RT-PCR, has been shown previously. The results shown here demonstrate that RNA of both viruses is capable of persisting for several weeks for subsequent RT-PCR detection, by both FTA card application and microwave treatment. However, the sensitivity test on aMPV indicated that the ability to detect is dose-specific. Both viruses were tested in tracheal organ culture fluids for simplicity and for ease of titration. However in practice, clinical material such as tracheal or cloacal swabs or tissue smears would be applied to the FTA cards. A study with Newcastle disease virus (NDV) (Perozo et al., 2006) showed that under experimental conditions, although virus could be detected on FTA card samples taken from the trachea, lung, caecal tonsils and cloacal faeces, the length of time for which detection was possible varied according to the organ. For example, while samples taken from the caecal tonsils were detectable up to 7 days after inoculation, those from the cloacal faeces were positive for only two days. Further work is clearly required to determine if the presence of clinical materials such as tissues or faeces whether normal or diseased would compromise the detection of IBV or aMPV RNA by inactivation or by reducing access in some way. Elhafi et al. (2004) showed that tracheal swabs from infected chicks were suitable for RT-PCR detection after microwave treatment. In the main trial, aMPV results were less consistent than those for IBV. This may have been due to the ‘stickiness’ of aMPV (C. J. Naylor, personal communication), perhaps preventing even distribution of virus on the filter papers. The FTA card system is validated and accepted by authorities responsible for importation of clinical material across national boundaries. However the materials are not inexpensive. In practical terms, microwave treatment appears to be as efficient as FTA card use and expense is minimal, since 20 seconds in a domestic microwave is all that is needed for virus inactivation. Unfortunately, it is not possible to conclusively prove that the treatment has been done, although we have found that re-microwaving on receipt would still leave RNA available for testing (Elhafi et al., 2004).


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REFERENCES Elhafi, G., Naylor, C.J., Savage, C.E. and Jones, R.C. (2004) Microwave or autoclave

treatments destroy the infectivity of infectious bronchitis virus and avian pneumovirus but allow detection by reverse transcriptase-polymerase chain reaction. Avian Pathology, 33, 303-306.

Moscoso, H., Raybon, E.O., Thayer, S.G. and Hofacre, C.L. (2004) Molecular detection and serotyping of infectious bronchitis virus from FTA filter paper. Avian Diseases, 49, 24-29.

Perozo, F., Villegas, P., Esteves, C., Alvarado, I. and Purvis, L.B. (2006) Use of FTA filter paper for the molecular detection of Newcastle disease virus. Avian Pathology, 36, 93-98.

Purvis, L.B., Villegas, P. and Perozo, F. (2006) Evaluation of FTA paper and phenol for storage, extraction and molecular characterization of infectious bursal disease virus. Journal of Virological Methods, 138, 66-69.

Snyder, J. W. (2002) Packaging and shipping of infectious substances. Clinical Microbiology Newsletter, 12, 89-92.

Worthington, K. J. Currie, R. J. W and Jones, R.C. (2008) A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. Avian Pathology 37, 247-257,

Table 1. Detection of infectious bronchitis virus on FTA cards or after microwave treatment after storage at different temperatures

Weeks after inoculation Sample 1 2 3 4 5 6

FTA cards

RT + + + + - + Fridge + + + + + - -200C + + + + + +

Microwave

RT + + + + - + Fridge + + + + - + -200C - + + + + +


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Table 2. Detection of avian metapneumovirus virus on FTA cards or after microwave treatment after storage at different temperatures.

Weeks after inoculation

1 2 3 4 5 6

FTA cards

RT + + - - + - Fridge + + - + + + -200C + - + - + -

Microwave

RT - - - - - - Fridge + - - - + - -200C + + - - + -

Table 3. Sensitivity of detection of different dilutions of avian metapneumovirus on FTA cards.

Titre 3.6a 3.6 2.6 2.6 1.6 1.6 0.6 0.6

Extraction Method

Kit b Phenol Kit Phenol Kit Phenol Kit Phenol

Days

1 + + + + + - - - 6 + + + + - - - - 20 + + - - - - - -

a Log10 TCID50/ml; bQiagen RNeasy minikit


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MOLECULAR DETECTION AND TYPING OF AVIAN INFECTIOUS BRONCHITIS VIRUS

LÜSCHOW D, DE QUADROS VL and HAFEZ HM

Institute of Poultry Diseases, Faculty of Veterinary Medicine Freie Universität Berlin, Königsweg 63, 14163 Berlin, Germany

SUMMARY In the present study a system for molecular detection and typing of infectious bronchitis virus (IBV) was established. For the general diagnosis of IBV a universal RT-PCR in form of a real time assay with an internal heterologous control on the basis of a GFP in vitro transcript was developed. For further typing of IBV positive samples two conventional subtype specific RT-PCR assays as well as a RT-PCR system in combination with restriction enzyme analysis (REA) or sequence analysis of PCR products were carried out. INTRODUCTION Infectious bronchitis (IB) is an acute, rapidly spreading disease of chickens affecting the respiratory, reproductive and renal system. The diseases is characterized by respiratory signs, drop in egg production and poor egg quality or nephritis. The causative agent, infectious bronchitis virus (IBV) is a member of the genus Coronavirus, within the family Coronaviridae. The genome of IBV consists of single-stranded positive sense RNA coding for four structural proteins: the nucleocapsid protein – N, the membrane protein – M, the small membrane protein – E, and the spike protein – S which consists of the two subunits S1 and S2. The high genetic variation within the S1 protein is responsible for multiple serotypes and variant strains which have been identified throughout the world. The occurrence of IBV serotypes and strains vary from country to country as well as from region to region. IB is difficult to differentiate from many other respiratory diseases and conditions related to reduced egg production. Clinical signs and gross lesions are only indicative and a definitive and accurate diagnosis based on direct antigen detection or isolation of IBV or on indirect detection of antibodies. Using conventional methods many limitations can influence the efficacy of the diagnosis. For example, in cases of direct detection using electron microscopy or immunofluorescence a sufficient amount of antigen in the samples is necessary, whereas the virus isolation in chicken embryos


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is very time consuming. In addition, with the widespread use of vaccines, serology has limited value in the diagnosis of the infection. Recently, molecular biological methods like polymerase chain reaction (PCR) systems have become increasingly important for a fast and sensitive diagnosis of infectious bronchitis. The use of IBV universal primers allowed the general diagnosis of IBV infection, the various subtypes and variant strains can be differentiated by IBV type specific primers or rather by restriction enzyme analysis of PCR products or sequence analysis of selected genome regions. The objective of the present study was the development of a molecular system for detection and typing of IBV field strains circulating in German poultry flocks. MATERIALS and METHODS Detection of IBV For the general diagnosis of IBV a universal real-time reverse transcriptase polymerase chain reaction (RT-qPCR) with an internal heterologous control (IC) was developed. The selected IBV specific primers and a FAM labelled probe were located within conserved regions of the M gene of IBV. As IC a green fluorescent protein (GFP) in vitro transcript based on a GFP primer probe Cy5 system was generated in accordance to Hoffmann et al. (2005; 2006). The IC spiking of samples was done before RNA extraction or within the PCR setup. The sensitivity of the assay was validated by an IBV in vitro RNA within a uniplex (IBV detection) and the duplex (IBV and IC detection) PCR assay. The specificity was evaluated by different IBV subtypes as well as by a panel of non related avian pathogens. Typing of IBV For typing of IBV two conventional subtype specific RT-PCR assays were performed. The detection of subtype 4/91 IBV was carried out with primers described by Handberg et al. (1999) For the detection of QX-like IBV two primers located within the S1 gene were designed. Further molecular typing was performed by a RT-PCR system combined with restriction enzyme analysis (REA) or sequence analysis of PCR products. The initially selected primers correspond to relative conserved parts of the S1 gene, enclosing the S1 gene hypervariable region (Kwon et al., 1993; Yu et al., 2001). Subsequently a further primer pair and a set of nested primers were used for amplification of almost all field samples (Bochkov et al., 2007; Dolz et al., 2006). Obtained PCR products were digested with at least two different restriction enzymes or sequenced directly in a commercial DNA sequencing service. RESULTS and DISCUSSION In the present study we reported about a molecular system for detection and typing of IBV field strains circulating in German poultry flocks. This system was composed of two different steps. For the general diagnosis of IBV infection a universal RT-qPCR assay was developed. To avoid false negative results due to PCR inhibitors or failed RNA extraction an internal heterologous control (IC) on the basis of a GFP in vitro transcript was co-amplified within a duplex RT-qPCR assay (Hoffmann et al., 2006). The sensitivity of the assay was determined with a 10-fold dilutions series of an IBV in vitro RNA. Investigation within the IBV uniplex assay revealed a detection


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limit of 102 copies / well with a PCR efficiency of about 94%. The same detection limit was obtained by investigation of an IBV in vitro RNA, which was spiked with the IC within the duplex assay. The IC was amplified in nearly all IBV dilutions steps, an inhibition was found only for the highest amount of IBV RNA. The specificity of the assay was confirmed by testing different avian pathogens which revealed negative results. In contrast, in all investigated IBV subtypes IBV RNA was detected success-fully. By investigation of field samples with this universal RT-qPCR a tolerable, competitive inhibition of the IC amplification due to high IBV RNA amounts was observed for some IBV positive samples. However, in all IBV negative samples the IC was amplified, which allowed an exclusion of false negative results due to presence of PCR inhibitors. For typing of IBV universal RT-PCR positive samples a subtype specific 4/91 RT-PCR was applied. The used primers were located within the S1 gene region and allowed an exclusive hybridization with subtype 4/91 IBV RNA (Handberg et al., 1999). In addition, recent investigations reported about the incidence of the new subtype QX-like IBV in Europe (Beato et al., 2005; Worthington et al., 2008). In this view, a subtype specific RT-PCR for fast and sensitive detection of QX-like IBV was developed. The designed primer pair amplified a fragment of about 220 bp within the S1 gene region of IBV. The specificity of both assays was confirmed by investigation of different IBV strains, which revealed positive results only for the corresponding subtype. For further typing of almost all subtypes and variant strains of IBV a RT-PCR system combined with REA or sequence analysis was established. For this purpose a primer pair was chosen, which was localised on the one hand within relative conserved parts of the S1 gene and enclosed on the other hand the S1 gene hypervariable region. Initial investigations revealed that selected primers were able to amplify a fragment of the expected size from all available subtypes and variant strains of IBV like H120/H52, Beaudette, 4/91, D274, and D8880. These strains were differentiable by the use of two different restriction enzymes. However, for some IBV universal PCR positive field samples the used primer pair delivered only very weak or negative results. So, in consequence of the high sequence variation of some IBV strains the system was expanded by a further primer pair and a set of nested primers and subsequent sequence analysis of the obtained PCR products. So far, different viruses of the Massachusetts type as well as 4/91, D1466, V1397, and QX-like IBV, which was the predominant subtype in 2008, were detected from German IBV field samples. Altogether, the combination of a universal IBV RT-PCR and a following differentiation system represents an effective procedure for the molecular diagnosis of IBV. An advance for the generic IBV diagnosis represents the development of a specific and sensitive IBV RT-qPCR system. By application of an internal control false negative results due to PCR inhibitors or failed RNA extraction can be excluded.


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REFERENCES Beato, M. S., De Battisti, C., Terregino, C., Drago, A., Capua, I. & Ortali, G. (2005).

Evidence of circulation of a Chinese strain of infectious bronchitis virus (QXIBV) in Italy. Veterinary Record, 156, 720.

Bochkov, Y. A., Tosi, G., Massi, P. & Drygin, V. V. (2007). Phylogenetic analysis of partial S1 and N gene sequences of infectious bronchitis virus isolates from Italy revealed genetic diversity and recombination. Virus Genes, 35, 65-71.

Dolz, R., Pujols, J., Ordonez, G., Porta, R. & Majo, N. (2006). Antigenic and molecular characterization of isolates of the Italy 02 infectious bronchitis virus genotype. Avian Pathology, 35, 77-85.

Handberg, K. J., Nielsen, O. L., Pedersen, M. W. & Jorgensen, P. H. (1999). Detection and strain differentiation of infectious bronchitis virus in tracheal tissues from experimentally infected chickens by reverse transcription-polymerase chain reaction. Comparison with an immunohistochemical technique. Avian Pathology, 28, 327– 335.

Hoffmann, B., Beer, M., Schelp, C., Schirrmeier, H. & Depner, K. (2005). Validation of a real-time RT-PCR assay for sensitive and specific detection of classical swine fever. Journal of Virological Methods, 130, 36-44.

Hoffmann, B., Depner, K., Schirrmeier, H. & Beer, M. (2006). A universal heterologous internal control system for duplex real-time RT-PCR assays used in a detection system for pestiviruses. Journal of Virological Methods, 136, 200-209.

Kwon, H. M., Jackwood, M. W. & Gelb, J., Jr. (1993). Differentiation of infectious bronchitis virus serotypes using polymerase chain reaction and restriction fragment length polymorphism analysis. Avian Diseases, 37, 194-202.

Worthington, K. J., Currie, R. J. & Jones, R. C. (2008). A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. Avian Pathology, 37, 247-257.

Yu, L., Wang, Z., Jiang, Y., Low, S. & Kwang, J. (2001). Molecular epidemiology of infectious bronchitis virus isolates from China and Southeast Asia. Avian Diseases, 45, 201-209.


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TEST FOR EXTRANEOUS AGENTS IN AVIAN INACTIVATED VACCINES USING PCR: DETECTION OF INFECTIOUS BRONCHITIS VIRUS

MOTITSCHKE A and JUNGBÄCK C

Paul-Ehrlich-Institut, Paul-Ehrlich-Str. 51-59, 63225 Langen SUMMARY In this study the sensitivity of a PCR method for the detection of Infectious bronchitis virus (IBV) was compared to the sensitivity of the serological test described in the European Pharmacopoeia (Ph. Eur.). For this purpose, different dilutions of an inactivated IBV vaccine were prepared and groups of SPF chickens were vaccinated with a double dose of the prepared vaccine dilutions. After a period of 21 days, the animals were revaccinated with a single dose. Two weeks later, serum samples from each animal were tested for IBV antibodies using an Idexx ELISA. In parallel, samples of the diluted vaccine were tested for IBV by PCR. It was found that the sensitivity of the PCR method used is comparable to or even slightly better than the serological test. Thus this PCR method fulfils the requirements on sensitivity of the Ph. Eur. and could be used as an alternative test for the detection of extraneous agents in final batches of inactivated vaccines. INTRODUCTION Final product batches of avian inactivated vaccines must be tested for the presence of extraneous agents as required by the Ph. Eur. (6th Ed. 2009). The currently most common method for the testing for extraneous agents in final product batches of inactivated avian vaccines is the vaccination of chickens with multiple doses of the vaccine to be tested and the subsequent testing of the sera for antibodies against the antigens listed in Ph. Eur. The vaccine has passed the test when no antibodies to extraneous agents are detected. As an alternative to the serological detection, a PCR method can be used. The Ph. Eur. allows the use of PCR if the method is shown to be comparable to the conventional method with regard to specificity and sensitivity. While the specificity of the PCR was already tested by the Institute of Virology and Immunoprophylaxis (IVI, Mittelhäusern, Switzerland) who has established the PCR,


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data to compare the sensitivity of PCR and serological assay in chicken by ELISA to detect IBV are missing. MATERIALS and METHODS Vaccines An inactivated oil-emulsion vaccine containing the IBV strains M41 and D274 was diluted 10-1 to 10-4 in another oil-emulsion vaccine from the same manufacturer containing Egg drop syndrome virus (EDSV) antigen. The latter was chosen as diluent because the two vaccines were of the same composition except for the different antigen. A sufficient volume of each dilution was prepared for the vaccination of the chickens and for PCR testing. In order to simulate a contamination of an inactivated oil-emulsion vaccine with live IBV a live virus vaccine (IBV strain H52) was diluted in the inactivated EDS vaccine in a 10-fold serial dilution up to 10-7. Immunisation of chickens Immunisation with dilutions of IBV inactivated vaccine Groups of 10 SPF chickens each were vaccinated at the age of 4 weeks with a double dose (2 x 0.5 ml) of a 10-fold dilution series (neat to 10-4) of the prepared IBV vaccine in the breast muscle. After a period of 21 days, the animals were revaccinated with a single dose (0.5 ml) by the same route. Two weeks after the last injection, serum samples were taken and tested for IBV antibodies by ELISA. Immunisation with dilutions of live IBV vaccine SPF chickens of the same hatch and age were used. As only 31 chickens of the hatch were left for this study, it was decided to vaccinate only four groups of chickens (7-8 animals per group). Dilutions of 10-4 to 10-7 of the live IBV vaccine preparations were selected for injection. IBV-antibody-ELISA A commercially available Infectious Bronchitis Disease antibody test kit (FlockCheck IBV, Idexx) was used to detect antibodies in the chicken sera. The ELISA was done as prescribed by the manufacturer. Briefly: Chicken sera from treated chicken were diluted 1:500 in sample diluent buffer. 100 µl of the prediluted samples were dispensed into the wells of the IBV antigen coated ELISA plates in duplicate. Plates were then incubated for 30 min at room temperature. After incubation the plates were washed 3 times with 300 µl of Aqua bidest. 100 µl goat anti-chicken horseradish peroxidise conjugate were dispensed into each well and incubated for 30 min at room temperature. Following a washing step as described above 100 µl of TMB substrate were added to the plates. The enzymatic reaction was stopped after 15 min with 100 µl of stop solution. OD values were measured at 650 nm against air as blank. The S/P ratios and endpoint titres were calculated as described by the manufacturer. S/P ratios greater than 0.2 or titres greater than 396 are considered to be positive. Calculation of titres was performed according to the following equation: Log10titre = 1.09 (Log10S/P) + 3.36.


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IBV-PCR The PCR performed was established by the IVI using primers already described by Falcone et al. (1997). The sequence of primer IBV3 is 5’-GCC-CCA-GCT-CCA-GTC-AT-3’ and of IBV4 is 5’-CCA-AGC-ATC-TGG-GAC-TGG-T-3’. The concentration of the primers in the final reaction mix (50 µl) was 0.625 pmol/µl. The specificity of the PCR was already validated by Bruckner (2007). Testing at the Paul-Ehrlich-Institut (PEI) also confirmed that the primers are able to detect all tested IBV vaccine strains in live and inactivated avian viral vaccines (CR 88121, Massachusetts H120, H52, 1263, IBV 4-91, D274, Ma5, M41). Extraction of viral RNA was performed using the QIAamp Viral RNA Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. A volume of 140 µl of each dilution step of the vaccine was extracted. For the RT-reaction 2 µl of the extracted RNA were mixed with 48 µl of the PCR reagents of a RT-PCR OneStep Kit (Qiagen, Hilden, Germany). PCR was performed as a touch down PCR: The programming of the thermocycler was in a way that the annealing temperature decreases in steps of 2°C from +60°C down to +48°C. With each annealing temperature two cycles were performed. The following cycles were 35x (95°C, 30 sec., 48°C, 30 sec., 72 °C, 30 sec). Finally a prolonged extension step of 72°C for 10 min was programmed before the samples were left at 4°C. For analysis, 2 µl of loading buffer were added to 10 µl of PCR product. 10 µl of this mixture were analysed on a 2% agarose gel containing SYBR-Safe in TBE buffer. In order to control the repeatability of the PCR method samples of diluted vaccines were tested in parallel at both laboratories. Both laboratories used the same standard operating procedure and pair of primers. RESULTS Immunisation of chickens with dilutions of IBV inactivated vaccine Results were summarised in Table 1. IBV antibodies could be detected by ELISA in all sera of the chickens immunised with the undiluted inactivated vaccine. In the group of chickens immunised with a 1:10 dilution of the vaccine 5 out of 10 sera reacted positively. All further dilutions of the vaccine did not induce measurable IBV antibodies. Control chickens and the group treated with the vaccine used a diluent only were negative for IBV antibodies. Immunisation of chickens with dilutions of live IBV vaccine Results were summarised in Table 2. None of the live vaccine dilutions 10-4 to 10-7 were able to induce antibodies against IBV.


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PCR Results were summarised in Tables 1, 2 and Fig. 1. At the PEI laboratory the undiluted and 10-1 diluted inactivated IBV vaccine produced a single specific DNA band of 240 bp. At IVI also the 10-2 dilution of the vaccine gave a positive result. Dilutions up to 10-4 of the live IBV vaccine could be detected by PCR at PEI laboratory but none of the dilutions 10-5 to 10-7. At IVI, who only received the dilutions 10-4 to 10-7 for testing, none of the samples reacted positively. DISCUSSION Batch testing of inactivated viral poultry vaccines includes a test for extraneous agents. This test is usually performed by detecting antibodies against a list of various antigens after immunisation of chickens (serological assay). Alternatively PCR can be used provided that the method is at least as sensitive as the serological assay and of appropriate specificity. As the specificity was already proven by the IVI who established the PCR this study focuses on the sensitivity of the PCR method to detect IBV as a possible extraneous agent and compare the results to the serological assay. For this purpose a series of dilutions of an inactivated vaccine containing IBV strains M41 and D274 was used to immunise groups of 10 chickens per dilution. Samples of the same dilutions were tested by PCR for the presence of IBV. The results showed that all chickens immunised with the undiluted inactivated vaccine reacted positively in the IBV ELISA. Only 5 out of 10 chickens produced antibodies when immunised with a 10-1 dilution of the vaccine. No antibodies could be detected in chickens that received a 10-2 dilution of the vaccine. The PCR method was able to detect inactivated IBV up to 10-2 at the PEI laboratory and 10-3 at IVI which tested the samples in parallel. In a preliminary run at the PEI laboratory the dilution of 10-3 was also tested positive. The results show that the sensitivity of the PCR method to detect IBV in inactivated vaccines is at least as sensitive as the serological assay using chickens. Taking into account the low sample volume of 140 µl for PCR compared to the total volume of 1.5 ml for the immunisation of the chickens, the sensitivity of the PCR is even higher than that of the serological assay. In order to simulate a contamination of inactivated vaccine with live IBV a vaccine containing live IBV (strain IB H52) was diluted up to 10-7 in an inactivated oil-emulsion vaccine. SPF chickens were immunised with dilutions 10-4 up to 10-7. None of the chickens reacted positively in the ELISA. By means of PCR it was possible to detect IBV in dilutions up to 10-4 when tested at the PEI laboratory. At IVI none of the four samples (10-4, 10-5, 10-6, 10-7) reacted positively. The latter demonstrated that the detection limit of the PCR was reached using these dilutions. With increasing dilutions the probability to receive a positive PCR reaction decreases. Thus it is likely


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that the same sample one time reacts positively and another time negatively after repeated measurements. While the PCR can react positively at a dilution 10-4 but none of the 8 chickens reacted positively in the ELISA it can be assumed that the PCR is at least as sensitive as the serological assay using chickens. The results of this study show that the sensitivity of the IBV PCR fulfils the requirement of the Ph. Eur. and therefore can be used as an alternative method for the detection of IBV as an extraneous agent in inactivated avian viral vaccines. Using a PCR method for extraneous agents testing has undoubtedly advantages but some critical aspects must be considered as well. A major advantage is that animal testing can be replaced by PCR. Another major advantage is that PCR results can be obtained within 1 or 2 days while the serological assay in chickens takes at least 5 weeks. This time aspect is important both for manufacturers and authorities to reduce the time needed for quality testing and batch release of vaccines. A point of discussion when using PCR is its usually higher sensitivity compared to the serological assay. It may happen that a batch of avian viral vaccine is tested positive by PCR but negative in the serological assay. In that case the question arises how to deal with this kind of results. A possible solution is to use PCR as a screening test. The relevance of a positive result should be critically assessed by means of a serological assay. A more general aspect when comparing the sensitivity of extraneous agents test methods is that the results are influenced by the test systems used and other factors. In the serological assay, for example, the results depend on the sensitivity of the ELISA as well as the immunogenicity of the viral strain used for immunisation. Various parameters of the PCR method affect the sensitivity of a PCR test system (primers, temperature, composition of the reaction mix, quality of polymerase, sensitivity of detection system …). Therefore each PCR test system used for the detection of extraneous agents in batch testing of viral vaccines has to be validated at least for sensitivity, specificity and repeatability. There is a strong need for PCR reference materials for the list of avian viruses which have to be tested for extraneous agents according to the current Ph. Eur. Established reference materials would allow to define the minimum amount of virus to be detected irrespective of the PCR system used. This is important because the Ph. Eur. does not provide any specifications regarding the primers or PCR systems to be used. The availability of PCR reference materials would help authorities and vaccine manufacturers to evaluate and compare the suitability of a newly developed PCR method with regard to its sensitivity


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REFERENCES Bruckner L, Ottiger H.P., Pre-validation Study for Testing of Avian Viral Vaccines for

Extraneous Agents by PCR. Pharmeuropa Bio 2007-1, 15-17 Falcone E, D’Amore E et al. Rapid diagnosis of avian infectious bronchitis virus by

the polymerase chain reaction. J. Virol. Methods 1997; 64:125-130 Falcone E, D’Amore E, Di Trani L, Puzzeli S & Tollis M. Detection of avian infectious

bronchitis virus in poultry vaccines by the polymerase chain reaction. In Animal Alternatives, Welfare and Ethics (ed. L.F.M. van Zuphten & M. Balls), Amsterdam, The Netherlands: Elsevier, 1997, 1007-1012

Methods of analysis, Ph. Eur. 6th edition. Strasbourg, France: Council of Europe; 2009

ABBREVATIONS Ab Antibodies EDS(V) Egg Drop Syndrome (Virus) IB(V) Infectious Bronchitis (Virus) IVI Institute of Virology and Immunoprophylaxis ND(V) Newcastle Disease (Virus) PEI Paul-Ehrlich-Institut Ph. Eur. European Pharmacopoeia Table 1 Serological and PCR results obtained with dilutions of an inactivated IBV vaccine Dilutions of inactivated IBV vaccine undiluted

vaccine 10-1 10-2 10-3 10-4 diluent only unvaccinated chickens

IBV Ab ELISA

+ (10/10)

+ (5/10)

- (0/10)

- (0/10)

- (0/10)

- (0/10)

- (0/10)

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Table 2 Serological and PCR results obtained with dilutions of a live IBV vaccine Dilutions of live IBV vaccine (in an inactivated vaccine) undiluted

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Figure 1: Reverse transcriptase PCR amplification of different dilutions of avian viral IBV vaccines. The sensitivity of the PCR was determined by testing 10-fold serial dilutions of an inactivated oil emulsion IB vaccine (inac vaccine) and of a live IB vaccine (live vaccine). Both vaccines were diluted in an inactivated oil emulsion vaccine (vaccine diluent). M: 100 bp molecular weight marker.


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THE AVIAN CORONAVIRUS INFECTIOUS BRONCHITIS VIRUS AS A MODEL FOR AN IN-HOUSE VIRUS DETECTION MICROARRAY

ABU-MEDIAN AA and BRITTON P

Coronavirus Group, Institute for Animal Health, Compton, Newbury,

Berkshire RG20 7NN, UK SUMMARY The early detection and identification of the aetiological agent(s) is a crucial factor in the control of a disease. Existing parallel assay detection/screening techniques aimed at identifying single or a few known pathogens, often rely on knowledge gained from disease symptoms. These techniques are time-consuming and suffer from severe limitations. Detection microarrays have the advantage of analysing a single field or clinical sample for the presence of multiple viruses in a single operation without prior knowledge of the identity of the pathogens. As part of a UK government-funded project, three generations of in-house oligonucleotide-based microarrays have been developed. The third generation microarray comprised 2884 oligonucleotide probes (70-mer) derived from 308 virus species from 36 families. Oligonucleotide probes covering the three coronavirus groups, with probes spanning the avian coronavirus infectious bronchitis virus (IBV) M41 genome, all other known avian viruses and exotic mammalian viruses were included. The array has been validated with different strains of IBV - predominantly M41 - as a model for the array development and evaluation. Following scanning of microarrays, raw data were normalised, statistically analysed and visualised using ‘DetectiV’ custom software (http://www.biochip-deteciv.co.uk). Analyses revealed different hybridisation profiles of the different IBV strains (cell culture-grown or from experimental infection) with IBV-derived oligonucleotides and some other avian coronaviruses on the array. Representatives of the three coronavirus groups were also tested successfully. This study shows that a detection microarray offers a rapid diagnostic and surveillance tool. INTRODUCTION With the current threat imposed by emerging and zoonotic viruses, it is essential to have a rapid and specific assay for virus detection and characterisation. Many detection methods are aimed at identifying single or a few known targets often relying on knowledge gained from disease symptoms. Existing techniques to screen for a

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wider spectrum of viruses, or for the detection of novel or emerging viruses, suffer from severe limitations. Pan-viral detection microarrays were developed predominantly targeting human pathogens (Wang et al. 2002). Since then, several arrays were developed for specific human and plant pathogens. A great advantage of a diagnostic microarray is that a single field or clinical sample can be analysed for the presence of multiple viruses in a single operation without prior knowledge of the identity of the pathogens. A single detection/characterisation platform specific for veterinary viruses was not developed. Here, we report the development and validation of the first pan-viral microarray targeting viruses of veterinary importance. Three generations of oligonucleotide-based microarrays were developed in-house. The third generation microarray (an expansion of the second generation) comprised 2884 oligonucleotides derived from 308 virus species from 36 families, and being validated to detect veterinary viruses. In this study, the avian coronavirus, infectious bronchitis virus (IBV) M41 has been used as a model for validating the array. The array has also been interrogated with other strains of IBV, representatives of the three groups of coronaviruses, representatives of nidoviruses and some avian and mammalian viruses. MATERIALS and METHODS 70-mer oligonucleotides (probes) were designed from fully sequenced viral genomes, partial sequences, and from sequences generated at the Institute for Animal Health (IAH), using publicly available software. Of these, 371 probes were derived from coronaviruses belonging to the 3 groups comprising the family Coronaviridae, with 96 probes derived from group 3 coronaviruses, of which 67 probes were derived from IBV with 22 probes spanning the genome of IBV M41. Oligonucleotides were synthesised commercially and randomly spotted in duplicate onto glass slides. Total RNA was extracted from virus-infected cell cultures or from tissues of experimentally-infected animals. RNA was reverse-transcribed and the resulting cDNA randomly amplified and labelled with Cy3-dCTP, and hybridised for 2 to 4 hours with probes printed onto slides. Slides were scanned and fluorescence was quantified using scanner software. Raw data were normalised, statistically analysed and visualised using ‘DetectiV’ custom software (http://www.biochip-detectiv.co.uk) which has been developed at IAH (Watson et al. 2007). RESULTS and DISCUSSION The results of our experiments will be described in full elsewhere. The array was developed by three laboratories; the IAH, Veterinary Laboratories Agency (VLA) Weybridge and Centre for Environment, Fisheries and Aquaculture Science (CEFAS) Weymouth. Probes derived from coronaviruses comprised around 12% of the array (Fig. 1). Analyses of raw data following the scanning of microarrays hybridised independently with labelled cDNA randomly generated from different IBV strains revealed specific detection of each strain with different hybridisation profiles between the strains (Fig. 2). The labelled IBV cDNA also detected oligonucleotides derived from the 3’ end of the newly identified avian coronaviruses isolated from ducks, geese and pigeons, indicating a close relationship to IBV (Fig. 2). In addition,

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probes spanning the IBV M41 genome were detected following hybridisation with the labelled cDNA derived from IBV M41. This confirmed that the random amplification method used to generate the virus-derived cDNA is a suitable approach for diagnostic microarrays in particular when the source of the sample is unknown. Hybridisation was also specific with the mammalian coronaviruses HCoV NL63, TGEV and MHV; the fish torovirus WBV and the arterivirus EAV (both belong to order Nidovirales); and avian influenza H7N7. ACKNOWLEDGEMENTS This work was supported by the Department of Environment, Food and Rural Affairs (DEFRA) project codes SE4102 & SD0443 and the Biotechnology and Biological Sciences Research Council (BBSRC). REFERENCES Wang, D., Coscoy, L., Zylberberg, M., Avila, P. C., Boushey, H. A., Ganem, D. and

DeRisi, J. L. (2002) Microarray-based detection and genotyping of viral pathogens. PNAS 99, 15687-15692.

Watson, M., Dukes, J. P., Abu-Median, A., King, D. P. and Britton, P. (2007) DetectiV: visualisation, normalisation and significance testing for pathogen-detection microarray data. Genome Biology 8:R190 (doi:10.1186/gb-2007-8-9-r190).


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Figure 1. Percentage of virus families respresented in the IAH-VLA-CEFAS veterinary microarray.

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Adenoviridae Arenaviridae Arteriviridae Asfarviridae Astroviridae Baculoviridae Birnaviridae Bornaviridae Bunyaviridae Caliciviridae Caulimoviridae Circoviridae Coronaviridae Dicistroviridae Filoviridae Flaviviridae Geminiviridae Hepadnaviridae Hepeviridae Herpesviridae Iridoviridae Nimaviridae Nodaviridae Orthomyxoviridae Papillomaviridae Paramyxoviridae Parvoviridae Picornaviridae Polyomaviridae Poxviridae Reoviridae Retroviridae Rhabdoviridae Roniviridae Togaviridae Tombusviridae Control Host Genes

Coronaviridae 12.86%

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Figure 2. Hybridisation profiles, corresponding to the group 3 probes, following analysis of the randombly generated cDNA from three different strains of IBV (M41-CK, Italy O2 and QX by the virus microarray.


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RAPID DETECTION AND OBJECTIVE CHARACTERISATION OF IBV ISOLATES USING HIGH RESOLUTION MELT CURVE ANALYSIS

AND A MATHEMATICAL MODEL

HEWSON KA1, NOORMOHAMMADI AH 2, DEVLIN JM 3, MARDANI K4 and IGNJATOVIC J5

1* Corresponding author, Department of Veterinary Science, The University of Melbourne, 250 Princes Highway Werribee, Victoria, Australia

2 Department of Veterinary Science, The University of Melbourne, 250 Princes Highway Werribee, Victoria, Australia

3 Department of Veterinary Science, The University of Melbourne, Parkville, Victoria, Australia

4 Faculty of Veterinary Medicine, The University of Urmia, Nazloo, Urmia, West Azarbaijan, Iran

5 Department of Veterinary Science, The University of Melbourne, 250 Princes Highway Werribee, Victoria, Australia

SUMMARY Classification of infectious bronchitis virus (IBV) in field samples is important for implementation of vaccination strategies to control the disease in commercial poultry. The lengthy process of sequencing variable sections of IBV structural genes is often used for IBV strain characterisation. A protocol for real-time PCR combined with an emerging technology, high resolution melt (HRM) curve analysis, was developed for rapid detection and classification of IBV. Initially, HRM curves were generated from 230 - 435 bp PCR products of the 3’UTR of 20 Australian IBV reference field strains and subjected to further analysis using a mathematical model. It was shown that a combination of HRM curve analysis and the mathematical model successfully characterised 189 out of 190 comparisons of pairs of IBV strains, corresponding to their structural gene identities. This new technique has detected and differentiated novel and vaccine-related Australian strains. Real-time PCR/HRM curve analysis is a rapid, reproducible, accurate and non-subjective system for detection and differentiation of IBVs in field samples. INTRODUCTION Infectious bronchitis virus (IBV) is a group 3 Coronavirus, which causes predominately respiratory signs, but can also be associated with or reproductive


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signs in poultry (Cavanagh and Naqi 2003). The IBV strains identified in Australia to date have been classified as either subgroup 1, 2 or 3, or ‘other’ field strains (Ignjatovic et al. 2006; Hewson et al. 2009). At present, the most common method for IBV detection and characterisation involves nucleotide sequencing of the S1 gene, which is time-consuming and costly. Also, the S1 gene is one of the most variable regions of the IBV genome and is subject to recombination events (Jia et al. 1995; Kottier et al. 1995; Lee and Jackwood 2000), making interpretation of the nucleotide sequencing difficult and often unreliable. High resolution melt (HRM) curve analysis has been used successfully for detection of mutations in cancer (Krypuy et al. 2006) and for classification of viral, bacterial and parasitic pathogens (Odell et al. 2005; Cheng et al. 2006; Price et al. 2007; Vandersteen et al. 2007; Lin et al. 2008). HRM curve analysis utilises a saturating fluorescent dye which fluoresces when bound to dsDNA. This dye is added to the real-time PCR mixture before standard real-time PCR followed by HRM, which is performed on a real-time thermocycler (with HRM capacity). The HRM step involves gradually melting double-stranded DNA, over a set temperature range and increment (ramp), to single-stranded (non-fluorescing) DNA. The pattern of the decrease in fluorescence during the melt process is recorded and plotted. The plots vary depending on the length and sequence of the DNA amplicon, with accuracy reported at a single base change in a 400bp amplicon (Reed and Wittwer 2004). The aim of this study was to evaluate the reliability of HRM curve analysis of the 3'UTR of the IBV genome for the detection, differentiation and characterisation of IBV strains, and evaluate its application for routine diagnosis of field cases.

MATERIALS and METHODS IBV strains and primers used in this study Australian reference field strains Q1/99, Q4/99, N1/62, Q1/73, Q1/76, H104, V1/71, V5/90 (Mardani et al. 2006), Q3/88, N1/03, V6/92, V18/91, N1/88 (Sapats et al. 1996; Ignjatovic et al. 2006; Mardani et al. 2008), V1/02 (Mardani et al. 2006) and V1/07 (Hewson et al. 2009) and all Australian IBV vaccine strains (A, I, VicS (Fort Dodge Australia Pty Ltd), S (Intervet Australia Pty Ltd) and B (Vaxsafe IB®, Bioproperties Australia Pty Ltd.)) were analysed. Forty field samples from suspected IBV cases in Australia were submitted for testing with the 17 returning positive results analysed in this study. Primers All1-F, sense primer binding to nucleotides 26930-26948 of the IBV Beaudette strain (GenBank accession number NC001451), and Del1-R (Mardani et al. 2006), anti-sense primer binding to nucleotides 27362-27344 were used in real-time PCR. Real-Time PCR of the IBV 3’UTR RNA from all reference strains and field submissions was extracted using spin columns bedded with QiaexII suspension matrix beads, subsequently used to generate cDNA and subjected to real-time PCR as previously described (Hewson et al. 2009). Real-time PCR and HRM curve analysis were performed as previously described (Hewson et al. 2009), using a Rotorgene 6000 (Corbett Life Science, NSW, Australia).


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Nucleotide sequencing and analysis Nucleotide sequences for all strains were obtained for the IBV 3’UTR corresponding to nucleotides 26977-27351 of the Beaudette strain. For the reference strains, the S1 gene sequence corresponding to nucleotides 20488-22108 of the Beaudette strain was obtained, while the maximum possible S1 gene sequence was obtained from the field samples. Nucleotide sequence identities were then determined using ClustalW2 (http://www.ebi.ac.uk). HRM curve analysis The optimal ramp and temperature range for differentiation of the 3’UTR PCR products was determined. A HRM genotype confidence percentage (GCP) is a comparison of a test strain to a reference strain and is expressed as a percentage, with 100% representing an exact match. GCPs were generated for each of the Australian reference strains and a cut-off value determined using these values for known related strains (based on nucleotide sequencing). GCP values for the 17 field samples were obtained and the cut-off value determined above was used to classify each sample as subgroup 1, 2, 3 or ‘other’. RESULTS HRM analysis of the 3’UTR Amplicons of 230-435bp were generated from the 3’UTR of all strains using real-time PCR with the All1-F and Del1-R primers. A ramp of 0.3°C over a temperature range of 75°C – 85°C, resulted in the most distinct, while consistent, conventional melt curves for Australian IBV reference strains. The conventional melt curves produced by the reference strains from each subgroup are presented in Figure 1. Based on nucleotide sequence identities, no unrelated strains produced the same shape curve. VicS vaccine, from subgroup 1, produced a slightly different conventional melt curve shape. Ten of the 17 field samples produced conventional melt curves matching subgroup 1, 2 matched subgroup 3, while the remaining 5 samples produced unknown (‘other’) conventional melt curves. A mathematical system to relate or differentiate IBV strains GCP values are generated from the normalised HRM curves shown in Figure 2. These curves provide an alternative way to display the same data used to produce the conventional melt curves. Using GCPs for pairs of reference strains that had an S1 gene identity ≥ 95% and 3’UTR identity of ≥ 97%, a cut-off value was generated as a mathematical model to assess the relationship of the IBV strains without visual interpretation by the operator (non-subjective). The average of the GCPs was 93.09 with a SD of 7.88; thus the GCP range for related strains was determined to be 80.13 - 100. Using this range, 168 out of a possible 190 GCP values for the reference strains used were typed in accordance with the identities of their S1 gene and 3'UTR sequences, with 94% of these 168 values differentiating unrelated strains (i.e. a GCP value <80.13). Of the 22 GCPs that were not typed or differentiated in accordance to their 3’UTR and/or S1 gene identities, visual examination of the conventional melt curves for 21 of these comparisons resulted in accurate typing, due to the distinctive nature of the subgroup conventional melt curves.

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Classification of the 17 field submissions using GCP/HRM curve analysis was verified by their 3’UTR and S1 gene nucleotide sequence identities. Three of the 5 ‘other’ strains were classified as subgroup 3 variants, 1 was a subgroup 1 variant, while the remaining strain was determined to be a recombinant strain. DISCUSSION

The system described in this study (Hewson et al. 2009) represents a considerable advancement in IBV detection and classification, using HRM curve analysis. This system has proved to be highly successful in identifying field strains of IBV in Australia, including the detection of novel variants and recombinants. Using the GCPs in conjunction with the shape of the conventional melt curves, all strains under investigation in this study were correctly typed, with one exception. The results of this investigation are consistent with previous studies exploring the relationships between Australian IBV strains (Sapats et al. 1996; Ignjatovic and Sapats 1997; Mardani et al. 2006; Mardani et al. 2006), with no unrelated strains being typed as similar. Monitoring the shape of the conventional melt/HRM curves, in addition to analysis of their GCPs, enabled more reliable typing as each subgroup produces considerably different melt curves. It was observed that the GCPs for the reference IBV strains fell either within the GCP range of 80.13 - 100, just outside this range (GCP of 60-80), or far below this range (GCP value <30). More than three quarters of the reference strain comparisons in this study fell into this latter category, meaning that in most cases extensive analysis of data may not be necessary. Classification of the 17 field submissions using their 3’UTR and S1 gene sequence identities correlated with the initial classification using the GCP/HRM analysis. There is a large disparity in processing time between these 2 techniques to identify IBV isolates, as the HRM analysis technique (from receipt of infected tissue to HRM analysis) can be completed in less than five hours. The described technique utilises a small PCR product that was successfully generated directly from infected tissues/swabs to determine that 17 of the original 40 field samples were positive for IBV. The VicS vaccine used in this study produced a GCP and a conventional melt curve that was inconsistent with other subgroup 1 strains, despite a high 3’UTR nucleotide sequence identity (99-100%). Evidence in our laboratory suggests the presence of 2 populations of virus in the VicS vaccine preparation. This ultimately demonstrates a further capacity of the HRM curve analysis for the possible detection of more than one strain in a single preparation. This new system is faster and more cost-effective than the currently established methods for detecting and differentiating IBV isolates and has many potential benefits for rapid control of IBV outbreaks. ACKNOWLEDGEMENTS Funding to support this research was provided by the Australian Egg Corporation Limited (AECL). The authors thank Naomi Kirkpatrick and Denise O’Rourke for technical assistance throughout this study. The authors would also like to thank Garry Anderson for help with statistical analyses.


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a) Subgroup 1 strains

b) Subgroup 2 strains

c) Subgroup 3 strains

d) ‘Other’ Australian strains

Figure 1 Conventional melt curves for Australian IBV subgroup strains. a) Subgroup 1 strains, which include 4 of the 5 Australian vaccine strains, and various field reisolates, b) Subgroup 2 strains, previously referred to as ‘novel’ Australian strains, c) Subgroup 3 strains, recently emerged field strains, d) various ‘other’ Australian IBV strains that were not classified in a subgroup. Graphs generated using TeeChart Office.


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Figure 2 Normalised HRM curve for various Australian IBV strains. The genotype confidence percentage (GCP) values are derived from this graph.


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REFERENCES

Cavanagh, D. and Naqi, S.A. (2003). Infectious Bronchitis. Diseases of poultry. Y. M. Saif, H. J. Barnes, J. R. Glissonet al, Iowa State Press: p.101-119.

Cheng, J.-C., Huang, C.-L., Lin, C.-C., Chen, C.-C., Chang, Y.-C., Chang, S.-S. and Tseng, C.-P. (2006). "Rapid Detection and Identification of Clinically Important Bacteria by High-Resolution Melting Analysis after Broad-Range Ribosomal RNA Real-Time PCR." Clin. Chem. 52(11): 1997-2004.

Hewson, K., Noormohammadi, A., Devlin, J., Mardani, K. and Ignjatovic, J. (2009). "Rapid detection and non-subjective characterisation of infectious bronchitis virus isolates using high-resolution melt curve analysis and a mathematical model." Arch. Virol. 154(4): 649-660.

Ignjatovic, J., Gould, G. and Sapats, S. (2006). "Isolation of a variant infectious bronchitis virus in Australia that further illustrates diversity among emerging strains." Arch. Virol. 151(8): 1567-1585.

Ignjatovic, J. and Sapats, S.I. (1997). "A long-term study of Australian infectious bronchitis viruses indicates a major antigenic change." Avian Pathol. 26(3): 535-553.

Jia, W., Karaca, K., Parrish, C.R. and Naqi, S.A. (1995). "A novel variant of avian infectious bronchitis virus resulting from recombination among three different strains." Arch. Virol. 140(2): 259-271.

Kottier, S.A., Cavanagh, D. and Britton, P. (1995). "Experimental Evidence of Recombination in Coronavirus Infectious Bronchitis Virus." Virology 213(2): 569-580.

Krypuy, M., Newnham, G.M., Thomas, D.M., Conron, M. and Dobrovic, A. (2006). "High resolution melting analysis for the rapid and sensitive detection of mutations in clinical samples: KRAS codon 12 and 13 mutations in non-small cell lung cancer." BMC Cancer 6: 295-307.

Lee, C.W. and Jackwood, M.W. (2000). "Evidence of genetic diversity generated by recombination among avian coronavirus IBV." Arch. Virol. 145(10): 2135-2148.

Lin, J.-H., Tseng, C.-P., Chen, Y.-J., Lin, C.-Y., Chang, S.-S., Wu, H.-S. and Cheng, J.-C. (2008). "Rapid Differentiation of Influenza A Virus Subtypes and Genetic Screening for Virus Variants by High-Resolution Melting Analysis." J. Clin. Microbiol. 46(3): 1090-1097.

Mardani, K., Browning, G.F., Ignjatovic, J. and Noormohammadi, A.H. (2006). "Rapid differentiation of current infectious bronchitis virus vaccine strains and field isolates in Australia." Aust. Vet. J. 84(1-2): 59-62.

Mardani, K., Noormohammadi, A.H., Hooper, P., Ignjatovic, J. and Browning, G.F. (2008). "Infectious bronchitis viruses with a novel genomic organization." J. Virol. 82(4): 2013-2024.

Mardani, K., Noormohammadi, A.H., Ignatovic, J. and Browning, G.F. (2006). "Typing infectious bronchitis virus strains using reverse transcription-polymerase chain reaction and restriction fragment length polymorphism analysis to compare the 3' 7.5 kb of their genomes." Avian Pathol. 35(1): 63-69.

Odell, I.D., Cloud, J.L., Seipp, M. and Wittwer, C.T. (2005). "Rapid Species Identification Within the Mycobacterium chelonae-abscessus Group by High-Resolution Melting Analysis of hsp65 PCR Products." Am. J. Clin. Pathol. 123(1): 96-101.


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Price, E.P., Smith, H., Huygens, F. and Giffard, P.M. (2007). "High-Resolution DNA Melt Curve Analysis of the Clustered, Regularly Interspaced Short-Palindromic-Repeat Locus of Campylobacter jejuni." Appl. Environ. Microbiol. 73(10): 3431-3436.

Reed, G.H. and Wittwer, C.T. (2004). "Sensitivity and specificity of single-nucleotide polymorphism scanning by high-resolution melting analysis." Clin. Chem. 50(10): 1748-1754.

Sapats, S.I., Ashton, F., Wright, P.J. and Ignjatovic, J. (1996). "Sequence analysis of the S1 glycoprotein of infectious bronchitis viruses: identification of a novel genotypic group in Australia." J. Gen. Virol. 77(3): 413-418.

Vandersteen, J.G., Bayrak-Toydemir, P., Palais, R.A. and Wittwer, C.T. (2007). "Identifying Common Genetic Variants by High-Resolution Melting." Clin. Chem. 53(7): 1191-1198.


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PATHOGENIC PATTERNS IN CHICKEN CHALLENGED WITH VARIANT STRAINS OF INFECTIOUS BRONCHITIS VIRUS ISOLATED FROM CHICKEN

FLOCKS WITH DIFFERENT CLINICAL MANIFESTATIONS

CHACON JL, ASSAYAG MS; REVOLLEDO L; IVO M; VEJARANO MP; PEDROSO AC and FERREIRA A

Department of Pathology, College of Veterinary Medicine, University of São Paulo, São Paulo, Brazil.

Avenida Professor Doutor Orlando Marques de Paiva 87, Cidade Universitária, CEP 05508-900, São Paulo, SP, Brazil.

SUMMARY Infectious bronchitis virus (IBV) is the causative agent of avian infectious bronchitis, which is characterized by respiratory, reproductive and renal signs. Two IBV samples, which were isolated from commercial chickens with respiratory disease (USP-78), and severe enteric disturbs and mild respiratory disease (USP-22) were inoculated in 26-day-old chickens and characterized by partial sequencing of the S1 gene. The sample USP-22 was inoculated by the oral and oculonasal routes (group 1 and 2, respectively) and the sample USP-78 was inoculated by the oculonasal route (group 3). The three groups showed similar clinical signs and microscopic lesions. The clinical signs included depression, coughing, sneezing, tracheal rales, open mouth breathing and mild watery faeces. The macroscopic and microscopic lesions were more evident in trachea. The nucleotide sequences revealed 98.3% identity between the both field isolates and between 52.5 to 76% identity when they were compared with vaccine strains available worldwide. INTRODUCTION Infectious bronchitis (IB) is an acute, highly contagious viral respiratory disease of chickens. The primary tissue of IB virus (IBV) infection is the respiratory tract, though some isolates replicate in the kidney, oviduct, testis and intestine, resulting in nephritis, fertility disturbs and reduced egg production (Cavanagh & Naqi, 2003). Frequently, the IB has been controlled with serotype-specific vaccines, but IB outbreaks still occur, because vaccines offer little or any cross-protection between serologically distinct viruses (Hofstad, 1981). The S1 subunit of spike glycoprotein of IBV is responsible for inducing neutralizing and serotype-specific antibodies in chickens and mutations in the antigenically important spike glycoprotein S1 subunit


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leads to the emergence and proliferation of variant serotypes associated with disease outbreaks (Moore et al., 1998). In Brazil, Massachusetts serotype is the unique vaccine strain used and variant IBVs were isolated from commercial chicken flocks with respiratory disease, reduced egg production and fertility disturbs. Last years variant IBV was detected in broilers which showed mild respiratory signs, and severe enteric disturbs with decreased growth rates. The purpose of this study was to know the pathogenic patterns produced by two variant viruses isolated from broiler flocks with different symptoms in SPF and commercial chickens. In addition, this study aimed to determine genetic relationships between the two Brazilian isolates and reference strains by sequence data analysis. MATERIALS AND METHODS Viruses The USP-78 and USP-22 samples isolated from broiler flocks with respiratory disease and enteric disturbs respectively, were included in this study. These viruses were previously characterized as variant isolates. Experimental design This study was repeated twice and the experiments were carried out in isolators. The first experiment was carried out in commercial chickens free of antibodies against IBV and in the second experiment SPF chickens were used. Eighty 26 day-old chickens were divided in four groups. The birds from the group 1 and 2 were infected with the sample USP-22 by oral and oculonasal route respectively. The chickens from the group 3 were inoculated with the USP-78 sample by oculonasal route. The group four was the negative control. The clinical signs were recorded during two weeks. Samples of conjunctive, trachea, lungs, kidneys, testis and enteric content were collected from two birds at 5, 6, 7, 8 and 9 days post-inoculation for RNA detection and histopathology. At 3 weeks post-infection sera were collected from all groups. Molecular characterization RNA of the two isolates used in this study was submitted to a RT-PCR assay for molecular characterization. S1 gene was amplified using the primer S1OLIGO 5’ and CK2 (Know and Jackwood (1995), Keeler et al., (1998)). Phylogenetic analysis was performed by the neighbour-joining method with 1000 bootstrap replicates with MEGA 3.0. RESULTS Clinical signs In all infected groups of the two experiments clinical signs were observed since the second day post-inoculation. No differences were observed between the groups infected with the two isolates. In the three infected groups, respiratory-type signs were more evident. It was possible to see depression, lacrimation, swollen face, nasal discharge, sneezing and tracheal rales. Mild watery faeces were observed in all groups after 7 days post infection and during 6 days. The signs were more severe in commercial chickens.


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Microscopic findings Similar to clinical signs observed during the experiments, the more important microscopic lesions were observed in trachea. The lesions included congestion, loss of cilia, mucous glandular depletion and hyperplasia, epithelial hyperplasia and epithelium degeneration. Mild lesions could be observed in lungs and kidneys. Mild epithelium desquamation and hypercellularity were observed. No lesions were observed in testis in any group. No differences were observed among groups infected with virus isolated from birds with respiratory or enteric disturbs. No differences were observed between groups infected by oculonasal or oral route with the enteric virus. RNA detection IBV RNA was detected in all three inoculated group in all birds during 5 to 9 days-post-inoculation in conjunctive, trachea, lungs and kidney. The virus was detected in testis and enteric content too, but not in all sampled birds from the three infected group. No virus was detected from the negative control group. DNA sequencing and phylogenetic analysis The IBV Brazilian isolates used in this study showed high genetic similarity between each other (nucleotide sequences revealed 98.3% identity) and with other Brazilian isolates published previously. The IBV Brazilian isolates show low genetic relationship with references serotypes used as vaccine in other countries (Table 1). The IBV Brazilian variant used in this study formed one unique branch together other Brazilian isolates published previously. Serology Antibodies against IBV were detected in all infected birds. No significant differences were observed among the infected groups. DISCUSSION IBV replicates in the upper respiratory tract, mainly in the trachea disseminating via the blood stream to different tissues like intestine, ovary, testis and kidneys remaining in these tissues for long periods of time (Cavanagh & Naqi, 2003). In this study, IBV was detected in all analyzed tissues in the groups infected with IBV originated from broilers with different clinical manifestations. However, in all the infected groups, the more severe lesions were observed in trachea. The two isolates produced similar respiratory symptoms. This indicates that the two isolates have respiratory tropism and the differences observed in the field might be produced by concomitant infections. No differences were observed in the clinical signs nor microscopic lesions when the USP-22 was inoculated by oculonasal and oral route. This indicates that pathological pattern is not influenced by infection route. The two isolates showed high genetic similarity. In addition, nucleotide and amino acid sequence analysis showed high molecular similarity between the isolates used in this study and other Brazilian variants published previously. On other hand, Brazilian IBV strains showed low genetic similarity when they were compared with all reference serotypes used as vaccine worldwide.


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REFERENCES Cavanagh D. & Naqi, S. 2003. Infectious bronchitis. In Saif, Y.M.; Barnes, J.R.;

Glisson, J.R.; Fadly, A.M.; McDougald L.R. & Swayne D. (Eds.), Diseases of poultry, 11th ed. (pp.101-119). Ames: Iowa State Press.

Hofsad, M.S. 1981. Cross-immunity in chickens using seven isolates of avian infectious bronchitis virus. Avian Dis. 25: 650-654.

Keeler, C.L.; Reed, K.L.; Nix, W.A. & Gelb, J. 1998. Serotype identification of avian infectious bronchitis virus by RT-PCR of the peplomer (S1) gene. Avian Dis, 42, 275-284.

Know, H.M.; Jackwood, M.W. 1995. Cloning and sequence comparison of the S1 glycoprotein of the Gray and JMK strains of avian infectious bronchitis virus. Virus Genes, 9, 219-229.

Moore K.M.; Bennett, J.D.; Seal, B.S.; Keeler, C.L.; Jackwood, M.W. 1998. Sequence comparison of avian infectious bronchitis virus S1 glycoproteins of the Florida serotype and five variant isolates from Georgia and California. Virus Genes, 17, 63-83.

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INFECTIOUS BRONCHITIS VIRUS INDUCES ACUTE INTERFERON-GAMMA PRODUCTION THROUGH POLYCLONAL STIMULATION OF CHICKEN

LEUKOCYTES

JANSEN1 C, ARIAANS1 M, VAN HAARLEM1 D, VAN DE HAAR1 P, DE WIT2 JJ and VERVELDE1 L

1 Dept. Infectious Diseases and Immunology, Faculty Veterinary Medicine, University Utrecht, Yalelaan 1, 3584 CL Utrecht, The Netherlands; 2 GD-Animal Health Service, Deventer, The Netherlands

SUMMARY Infectious Bronchitis Virus, a member of the Coronaviridae, is a major cause of economic losses in the poultry industry and can be involved in respiratory disease, nephritis, and poor egg production and quality. We found that in vitro stimulation with IBV resulted in chicken interferon gamma (chIFN-γ) production in splenocytes and peripheral blood leukocytes of both infected birds and uninfected birds. The non-specific stimulation did not occur when other avian viruses or other coronaviruses were used or when mammalian cells were stimulated with IBV. The aim of the preliminary study described here, was to investigate whether the induction of chIFN-γ also occurred in vivo after intransal and intra-ocular inoculation of M41, and whether Natural Killer (NK) cells might be activated and responsible for the rapid production of chIFN-γ. The number of cells isolated from lungs of infected birds that secreted chIFN-γ was increased at 24 hr p.i., suggesting that the rapid induction not only occurred in vitro but also in vivo. Activation of NK cells was found at both 24 hr and 48 hr p.i. based in the increased percentage CD107+ CD3- cells in lungs of infected birds compared to control birds. INTRODUCTION Infectious Bronchitis Virus is a coronavirus that is a major cause of economic losses in the poultry industry and can be involved in respiratory disease, nephritis, and poor egg production and quality. As well as being an economically relevant pathogen in poultry, IBV also bears close resemblance to the human pathogen severe acute respiratory syndrome coronavirus (SARS-CoV; Cavanagh, 2003). The most efficient and rapid host response against viruses consists of the production of type I IFNs (IFN-α and IFN-β), an essential part of the antiviral innate immune system (reviewed by Kawai and Akira, 2006). Many if not most nucleated cells are


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capable of producing type I IFNs, though differences in the production of IFN-α or IFN-β have been described (reviewed by Thiel & Weber, 2008). Secreted IFNs stimulate adjacent cells to express potent antiviral proteins, stimulate macrophages and NK cells to elicit antiviral responses. IFN-γ, a type II IFN, is predominantly produced by NK, NKT cells, dendritic cells and Th1 CD4+ and CD8+ effector cytotoxic T lymphocytes once antigen specific immunity develops. IFN-γ increases antigen presentation on macrophages and expression of MHC class I molecules on normal cells, promotes NK cell activity and leukocyte adhesion and binding required for migration. Thus the interferon production triggered by the first contact with the viral intruder slows down the virus multiplication, buys the organism time, and helps to establish an adaptive immune response. In a previous study, we showed that chIFN-γ production was not only significantly increased in splenocytes of IBV-infected chickens after in vitro restimulation with IBV, but also in splenocytes of IBV-uninfected chickens (Ariaans et al., 2009). Based on these findings, we investigated the possibility of polyclonal stimulation by IBV in more details. The aim of this pilot study was to investigate whether the rapid chIFN-γ production also occurred in vivo after infection with IBV-M41, and whether chicken NK cells are activated suggesting that this cell population might be the possible producers of the chIFN-γ. Avian NK cells have been described as a population of cells in the chicken embryonic spleen at a developmental stage where T cells have not yet migrated to the periphery. These cells express surface CD8 homodimers, but no T (CD3) or B-cell specific antigens and are able to kill the NK-susceptible cell-line LSCC-RP9 (Göbel et al., 1994). The frequency of avian NK cells in peripheral blood lymphocytes (PBL) and spleen was very low, ranging from 0.5 to 1.0% (Göbel et al., 2001). This is in sharp contrast to NK cell frequencies in mammals, which have approximately around 10% of NK cells. To measure the secretion of chIFN-γ ex vivo upon IBV infection we used an ELISPOT assay (Ariaans et al., 2008) and a CD107 assay to study NK cell activation (Jansen et al. 2009). MATERIALS and METHODS Chickens and virus One-day-old SPF layer chickens were housed in isolators. The chickens were given commercially available food according to the manufacturer’s instructions. Drinking water was supplied ad libitum. At an age of 31 days, the chickens of group A were challenged with 10^4.0 EID50 of IBV M41 by oculo-nasal route (one droplet of 0.05 ml on the eye, one droplet on the nostril). The chickens of group B were challenged with sterile water (one droplet of 0.05 ml on the eye, one droplet on the nostril). At 24 and 48 hours post inoculation (hr p.i.), 3 birds per group were euthanized using CO2/O2 and subsequent bleeding. Peripheral blood, trachea, lung and spleen were collected and stored in RPMI medium supplemented with 100 U/ml penicillin/streptomycin (P/S), 2 mM glutamax and 5% FCS for FACS analysis and elispot assays. PBMC were purified using, slow spin centrifugation (61xg, 20 min). Spleen tissue was squeezed through 70 μm mesh in RPMI medium to prepare a single cell suspension. Lung tissue was cut into small pieces and digested in RPMI containing collagenase and DNAse for 30 min at 37°C, and subsequently squeezed through a 70 μm mesh to prepare a single cell suspension. Splenocytes and lung leukocytes were isolated by density gradient


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centrifugation using FICOLL-Hypaque. The trachea was cut longitudinally and the mucosa was scraped off using a microscope slide. This tissue was squeezed through a 70 μm mesh to prepare a single cell suspension. Cells were counted using trypan blue exclusion. Elispot assay The ELISPOT assay has been described by Ariaans et al. (2008a), minor modifications are described here. Briefly, MultiScreentm-IP 96-well plates were treated with 70% ethanol for 1 min, washed and coated with mouse-anti-chIFN-γ, blocked and splenocytes were seeded at 2 x 105 cells/well in triplicate in culture medium. Cells were incubated with either culture medium to measure spontaneous release of chIFN-γ or medium supplemented with phorbol 12-myristate 13-acetate (PMA; 50 ng/ml) and ionomycin (500 ng/ml) as a positive control for the assay. The plates were incubated for 24 or 48 hr at 41 °C, 5% CO2. ChIFN-γ was detected by incubation with biotinylated mouse-anti-ChIFN- and poly-HRP streptavidin (Sanquin). The assay was developed using TMB substrate, and analyzed using the A·EL·VIS machine and the Eli.Analyse software (Version 4.0) that allows for automated counting of the number of spots based on size and intensity. Real-time quantitative RT-PCR (qRT-PCR) Total RNA was isolated using the RNeasy Mini Kit and DNase treated using the RNase-free DNase Set following manufacturer’s instructions (Qiagen Benelux B.V.). cDNA was generated with reverse transcription using iScript cDNA Synthesis Kit (Biorad). Real-time qRT-PCR was performed using the TaqMan Universal PCR Master Mix (Applied Biosystems, AB). Detection of IFN-α, IFN-β, and 28S was described by Ariaans et al. (2008b). Primers were used at 600 nM and probes at 100 nM concentration. Corrections for variation in RNA preparation and sampling were performed according to Eldaghayes et al. (2006). CD107 assay The CD107 assay that has been described to study NK cell activation in humans (Betts et al., 2005; Penack et al., 2005) was adapted for the chicken. Lung cells were resuspended in IMDM medium supplemented with 2% heat inactivated FCS; 8% heat inactivated chicken serum, P/S and 2 mM glutamax at a concentration of 1 x 106

cells/ml. Cells were incubated for 4 hr at 37C, 5% CO2 in the presence of 1µl/ ml Golgistop (BD) and anti-chCD107 mAb. After incubation, cells were washed in PBS supplemented with 0.5% BSA, stained with anti-CD3 mAb (SB) and flowcytometry was performed. RESULTS Birds were inoculated by the oculo-nasal route with IBV M41 and 24 hr and 48 hr p.i. chIFN-γ secretion measured using elispot assay. In the lungs of infected birds 24 hr p.i. spontaneous release of chIFN-γ was much higher than in uninfected birds (fig. 1). However, in spleen and PBMC, no increase in chIFN-γ was found. At 48 hr p.i. no significant difference in the number of chIFN-γ spots was found compared to uninfected birds in lung, spleen and PBMC (data not shown). For all birds at both


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time points the positive control, cells stimulated with PMA/ionomycin, resulted in high numbers of chIFN-γ spots, lowest in PBMC and highest in lung. The number of cells that could be isolated from trachea was too low to perform the elispot assay in triplicate, though it was evident that the number of cells isolated from infected birds was consistently higher than from uninfected birds. ChIFN-γ was produced in the trachea, but based on a single well per bird we cannot draw any conclusions except that the assay can also be applied for the trachea. To investigate whether IBV M41 also affected production of type I IFNs in the lung, chIFN-α and chIFN-β, mRNA expression was measured, because we do not yet have an elispot available for these cytokines. No increase in chIFN-α and chIFN-β was found at 24 hr and 48 hr p.i. (fig. 2). At 24 hr p.i. even a decrease was found in infected birds compared to control birds. At early stages of the immune response, IFN-γ can be produced by NK cells. Activation of NK cells was measured by CD107 expression (fig. 3) combined with staining for CD3 to discriminate between NK cells (CD3-) and cytotoxic T cells (CD3+). At 24 hr p.i. CD107 expression was increased on CD3- cells isolated from lungs of infected birds, but hardly increased on CD3+ T cells. No differences were found in the percentage cells that expressed CD107 at 24 hr or at 48 hr p.i. with M41, nor in the subpopulation of cells. DISCUSSION Birds were inoculated with IBV-M41 by the oculo-nasal route and at 24 and 48 hr p.i. chIFN-γ production was measured using elispot assay to ensure that the active protein was measured that can have a biological effect within the bird. In lungs of infected birds at 24 hr p.i. spontaneous release of chIFN-γ was much higher than in uninfected birds. However, in spleen and PBMC, no increase in chIFN-γ was found suggesting the response occurred at the site of infection. At 48 hr p.i. no significant difference was found in infected compared to uninfected birds in lung, blood and spleen. Although the number of birds was low, these preliminary data suggest that the chIFN-γ production did not remain stable. A kinetic study will be performed to determine whether waves of secretion might occur. These finding are in contrast to SARS-CoV that induces IFN-γ slightly later, 48 hr p.i., and remains rather stable for 5 days, although individual differences between human subjects were found (Castilletti et al. 2005). The effect of M41 on type I IFNs was measured by qPCR. mRNA expression of both chIFN-α and chIFN-β at 24 hr p.i. was slightly decreased in infected birds, but returned to control levels at 48 hr p.i.. This in accordance with the findings that M41 failed to yield interferon in primary monolayer cultures of chick kidney cells and in organ cultures of chick embryo trachea using Semliki Forest virus for the tests (Holmes & Darbyshire, 1978). However, in vitro studies describing induction and sensitivity of IBV for IFNs have been inconsistent (Otsuki et al. 1979, 1988), and data from infected birds are lacking. The interferon production triggered by the first contact with the viral intruder slows down the virus multiplication, however, many viruses including coronaviruses are capable to cope with the IFN system. It has been described for mouse hepatitis virus (MHV) and SARS-CoV, type 2a and 2b coronoviruses respectively, that they can passively escape from getting sensed by cytoplasmic pattern recognition receptors (PRRs) by formation of double membrane


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vesicles at perinuclar sites within the cytoplasm where RNA synthesis takes place and actively escape by ORF3b, ORF6, and N protein that are able to inhibit interferon regulatory factor 3 (IRF3; reviewed by Thiel & Weber, 2008). Whether evolutionary conserved IFN antagonists are encoded by all coronaviruses, including IBV, remains to be confirmed. After infection with M41, the production of type II IFN seemed to preceed that of type I IFN, which is rather surprising. However, an apparent discrepancy in type I IFN production upon Toll like receptor (TLR) triggering in chickens has been described. It was shown that upon triggering of TLR7 on a macrophage cell line HD11, up regulation of chIL-1β, chIL-6, chIL-8, but not chIFN-α or chIFN-β mRNA was found (Philbin et al., 2005). The lack of type I IFN response was unexpected, since mammalian blood lymphocytes respond to TLR7/8 agonists by up regulation of these molecules (Lee et al., 2003). Moreover, exposure of chickens or chicken splenocytes to the imidazoquinolinamine, S-28828, led to an increase in type I IFN-like activity (Karaca et al., 1996). The apparent discrepancy in type I IFN production may be resolved by f.e. the differential activity of different TLR7 agonists. Nonetheless, the implications of reduced type I IFN responses after TLR7 activation are potentially very important in protection against viral pathogens. The rapid production of chIFN-γ might outweigh the lack or reduced type I IFNs and in its turn be responsible for the induction of a rapid antiviral state, slowing down the viral replication and spread. A subpopulation of cells that can be rapidly activated and recruited upon viral invasion are NK cells. In most farm animals, the definition of NK cells is difficult due to the lack of specific markers (Evans & Jaso-Friedmann, 1993). Chicken NK cells express surface CD8 homodimers, but no T (CD3) or B-cell specific antigens (Göbel et al., 1994). To measure whether the NK cells were present and activated, a CD107 assay was performed using lung cells of infected and uninfected control birds. Cytotoxic cells contain cytolytic vesicles with proteins like granzyme and perforin. Lining the membrane of these vesicles is CD107 and upon activation the vesicles degranulate and merge with the surface membrane resulting in relocation of CD107 to the surface membrane (Alter et al., 2004). Upon infection with IBV-M41, CD107 expression was up regulated on CD3- cells in the lungs, suggesting that NK cells were activated. Expression of CD107 on CD3+ cytotoxic T cells (CTL) did not increase substantially indicating that, as expected, within 48 hr p.i. antigen-specific CTLs were not yet present in the lungs. At later time points CTLs play an important role in controlling IBV infection (Collison et al., 2000). In future experiments we will investigate whether these NK cells are responsible for the rapid secretion of chIFN-γ or whether other cells such as dendritic cells also play a role. Based on the number of chIFN-γ spots found in lungs of control birds at 24 hr p.i., it is possible that another cell population is present and capable of producing chIFN-γ, but this population is not present in the spleen and peripheral blood based on the low background number of spots. In conclusion, our preliminary data suggest that IBV-M41 induces rapid production of chIFN-γ in the lung within 24 hr p.i. by the oculo-nasal route, as previously found in in vitro cultures of splenocytes stimulated with IBV-M41. A cell population that might be responsible for the rapid production of chIFN-γ in the lung could be NK cells, as expression of CD107 on CD3- cells is increased compared to uninfected birds within 24 hr p.i. and remains stable for at least 48 hr. In a follow we will perform a kinetic


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study with more birds per time points, in which we will determine the phenotype of the cells that produce chIFN-γ in the lung and trachea in more detail. REFERENCES Alter, G., Malenfant, J.M., Altfeld, M. 2004. CD107a as a functional marker for the

identification of natural killer cell activity. J. Immunol. Meth. 294, 15-22. Ariaans, M.P., Van de Haar, P.M., Lowenthal, J.W., Van Eden, W., Hensen, E.J., and

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Castilletti, C., Bordi, L., Lalle, E., Rozera, G., Poccia, F., Agrati, C., Abbate, I., and Capobianchi, M. R., 2005. Coordinate induction of IFN-alpha and -gamma by SARS-CoV also in the absence of virus replication. Virology 341, 163-169.

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Evans, D.L. and Jaso-Friedmann, L. 1993. Natural killer (NK) cells in domestic animals: phenotype, target cell specificity and cytokine regulation. Vet. Res. Commun. 17, 429-447.

Göbel, T.W., Chen C.L., Shrimpf J. et al. 1994. Characterization of avian natural killer cells and their intracellular CD3 protein complex. Eur. J. Immunol. 24, 1685-1691.

Göbel, T.W., Kaspers B., and Stangassinger M. 2001. NK and T cells constitute two major, functionally distinct intestinal epithelial lymphocyte subsets in the chicken. Int. Immunol. 13, 757-762.

Holmes, H.C. and Darbyshire, J.H. 1978. Induction of chicken interferon by avian infectious bronchitis virus. Res. Vet. Sci. 25, 178-81.


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Jansen, C., Van de Haar, P. Van Haarlem, D., Viertlböck, B., Göbel, T., and Vervelde, L. 2009. Are NK cells involved in the defense against avian influenza virus? 7th International Symposium on Avian Influenza, Athens, Georgia, USA.

Karaca, K., Sharma, J.M., Tomai, M.A. and Miller, R.L. 1996. In vivo and In vitro interferon induction in chickens by S-28828, an imidazoquinolinamine immunoenhancer. J. Interferon Cyto. Res. 16, 327–332.

Kawai, T. and Akira, S. 2006. Innate immune recognition of viral infection. Nat. Immunol. 7, 131-137.

Lee, J., Chuang, T.H., Redecke, V., She, L., Pitha, P.M., Carson, D.A., Raz, E. and Cottam, H.B. 2003. Molecular basis for the immunostimulatory activity of guanine nucleoside analogs: activation of Toll like receptor 7. Proc. Natl. Acad. Sci. USA 100, 6646–6651.

Otsuki, K., Maeda, J., Yamamoto, H., and Tsubokura, M. 1979. Studies on avian infectious bronchitis virus (IBV) III. Interferon induction by and sensitivity to interferon of IBV. Arch. Virol. 60, 249–255.

Otsuki, K., Nakamura, T., Kawaoka, Y., and Tsubokura, M. 1988. Interferon induction by several strains of avian infectious bronchitis virus, a coronavirus, in chickens. Acta Virol. 32, 55–59.

Penack O, Gentilini C, Fischer L et al. 2005. CD56dimCD16neg cells are responsible for natural cytotoxicity against tumor targets. Leukemia 19, 835-840.

Philbin, V.J., Iqbal, M., Boyd, Y., Goodchild, M.J., Beal, R.K., Bumstead, N., Young, J. and Smith, A.L. 2005. Identification and characterization of a functional, alternatively spliced Toll-like receptor 7 (TLR7) and genomic disruption of TLR8 in chickens. Immunology 114, 507–521.

Thiel, V. and Weber, F. 2008. Interferon and cytokine responses to SARS-coronavirus infection. Cyto. & Growth Factor Rev. 19, 121-132.


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Figure 1. Enhanced production of chIFN-γ in the lung, 24 hr after infection with IBV-M41. The number of IFN-γ producing cells (mean of triplicate wells per bird +SEM) from uninfected (white, n=3) and IBV infected (black, n=3) was measured using an elispot assay. Cells were isolated from blood, spleen and lung 24 hr after IBV infection and were incubated with medium only to measure spontaneous release of chIFN-γ.


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Figure 2. No increase in type I IFN mRNA in lung, 24 and 48 hr after infection with IBV-M41. Real-time qRT-PCR of mRNA levels normalised to 28S of chIFN-α and chIFN-β expressed as 40-Ct (mean of triplicate wells per bird +SEM). No significant differences between uninfected (white, n=3 per time point) and IBV infected (black, n=3 per time point) birds were found.

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Figure 3. Enhanced expression of chCD107 (LAMP-1) shows activation of chicken NK cells upto 48 hours after IBV-M41 inoculation. CD107 expression in lung cells from uninfected (white, n=6) and IBV infected (black n=5) chickens was analysed by flowcytometry. Lung cells from IBV infected chickens showed significant (p<0.05) increased CD107 expression in CD3- cells. Median CD107 expression is shown together with the interquartile range.


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THE EXPERIMENTAL PRODUCTION OF BIVALENT INFECTIOUS BRONCHITIS (IB) NEWCASTLE DISEASE (ND) VACCINE (793/B IBV SEROTYPE / CLONE 30

NDV)

KHALESI B, MASOUDI S and MOGHADDAMPOUR M

Razi Vaccine & Serum Research Institute Razi Vaccine & Serum Research Institute, Hasarak, Karaj, Iran.

SUMMARY Infectious bronchitis is an acute, rapidly spreading, viral disease of chickens characterized by respiratory signs, decreased egg production, and poor egg quality. Some strains of the causative virus, infectious bronchitis virus (IBV), are nephropathogenic. The latter strains produce interstitial nephritis resulting in significant mortality. IBV, a coronavirus, is worldwide in distribution and has numerous serotypes. Two or more serotypes may be seen simultaneously in one geographic region. IBV is shed by infected chickens in respiratory discharges and feces. Naturally infected chickens and those vaccinated with live IBV may intermittently shed virus for many weeks or even months.

Newcastle disease virus (NDV) - A type strain for avian paramyxoviruses. Members of this family have a single stranded, linear, RNA, with an elliptical symmetry. The total genome is roughly 16,000 nucleotides. Replication of the virus takes place in the cytoplasm of the host cell. NDV is a contagious and fatal viral disease affecting most species of birds. Clinical signs are extremely variable depending on the strain of virus, species and age of bird, concurrent disease, and preexisting immunity. Four broad clinical syndromes are recognized by scientists. They are Viscerotropic velogenic, Neurotropic velogenic, Mesogenic, and Lentogenic. NDV is so virulent that many birds die without showing any clinical signs. A death rate of almost 100 percent can occur in unvaccinated poultry flocks. IB and ND are of major economic importance to commercial chicken producers worldwide.

A bivalent poultry vaccine is provided for having two live biological agents or microbial components. Each live biological agent or microbial component is effective in preventing or treating an avian disease, and the bivalent vaccine is safe and effective for immunizing poultry flocks.

In this study, 250 SPF chickens have divided into 5 groups each containing fifty chickens. The chickens have vaccinated singly by infectious bronchitis (793/B IBV) Newcastle disease (Clone 30 NDV) (group one and two) and in combination (bivalent 793/B IBV serotype / Clone 30 NDV group 3); also, two other groups have considered as control (group 4) and challenge group (group 5).


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Reactivity and immune response was determined using hemagglutination inhibition (HI) and serum neutralization (SN) test. Five weeks post vaccination all groups were submitted to challenge with the M41- challenge strain 106 EID50 /bird and NDV challenge strain 107 EID50 /bird also the ciliostasis test has performed.

The respective serum antibody titers obtained in the hemagglutination inhibition (HI) and serum neutralization test (SN) after vaccination with bivalent IB/ND vaccine. Protection, as measured by assessing ciliary activity of the tracheal epithelium following challenge, was excellent with the vaccine schedule used in this trial.

Each dose of potential vaccine contains at least 106 EID50/bird of live attenuated Newcastle virus vaccine clone 30 strain and 10 3.5 EID50/bird of live attenuated Infectious Bronchitis virus vaccine, 793/B IBV strain. INTRODUCTION Infectious bronchitis (IB), as a very contagious respiratory disease of chickens, was firstly described in North Dakota, USA, in 1930 (Cavanagh and Naqi, 2003). Nowadays, it has been spread all over the world (Cook, 2001; Cavanagh and Naqi, 2003). Prevention of the disease by immunization is worthwhile due to contagious nature of the disease and occurrence of numerous serotypes of IB virus (IBV) (Cavanagh and Naqi, 2003). Nevertheless, prevention of the disease by vaccination has been associated with partial success. Histopathologic studies of IBV have well addressed (Kotani et al., 2000). Newcastle disease (ND), for the first time was described in Java, Indonesia. In 1927, isolation of ND virus (NDV) was reported in England. There was another report from Betava, Indonesia. Very limited studies have addressed the effect of NDV on IB vaccination. In a comparative study, Thornton and Muskett, (1973), examined immunization of ND vaccine alone or in combination with IB vaccine. The results of this study indicated that in those chickens received B1 vaccine alone, the mean haemagglutination inhibition (HI) titer against ND was lower than those received mixed vaccine (B1 and H120). In the other study the difference, however, was not significant. EID50 of the challenge virus for the group received B1 vaccine alone was significantly higher than the group received mixed B1 and H120 (10 1 0. 13 vs 10 9 . 01 ) (Thornton and Muskett, 1973). Results of this study indicated that protection of chickens against challenge of virulent NDV is higher in those chickens received B1alone, as compared to those received B1 and H120. The tolerable EID50 for the chicks received ND vaccine and the chicks received ND in combination with IB vaccine were 109 .6 5 and 107. 9 8, respectively (P<0.05) (Thornton and Muskett, 1973). The aim of the present study is to investigate effect of bivalent infectious bronchitis (IB) Newcastle disease (ND) vaccine (793/B IBV serotype / Clone 30 NDV) on SPF chickens.


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MATERIALS and METHODS Vaccines lentogenic Newcastle disease virus (NDV) strain Clone 30 and IB vaccine serotype 4/91. Both vaccins are commercial available in Intervet America Inc. Each dose of potential vaccine contains at least 106 EID50/bird of live attenuated Newcastle virus vaccine clone 30 strain and 10 3.5 EID50/bird of live attenuated Infectious Bronchitis virus vaccine, 793/B IBV strain. Experimental design 250 one-day-old SPF chickens were divided into 5 groups and placed in negative pressure isolators. SPF eggs have provided from Leghorn Company of Germany.The expermental groups including as followes: Group I was vaccinated by eye-drop with 104 EID50 /bird of the793/B IBV serotype. Group II was vaccinated by eye-drop with minimal 105.8 EID50 /bird of the Clone 30 NDV. Group III were vaccinated by eye-drop with a combination of the bivalent infectious bronchitis (IB) Newcastle disease (ND) vaccine (793/B IBV serotype / Clone 30 NDV) (with the same dose as in the single vaccination). Groups IV and V were considered as control and challenge group. Clinical symptoms and ciliostasis test Four and eight days post-vaccination six birds per group were removed clinical symptoms were recorded and their tracheas were examined in the ciliostasis test. Challenge programs Five weeks post-vaccination the birds vaccinated by eye-drop were submitted to a challenge with the M41-challenge strain (106 EID50 per bird, eye-drop). Four days after the M41-challenge, clinical symptoms have recorded and tracheas had taken for the ciliostasis test.

The birds vaccinated by eye-drop were submitted to a NDV-challenge five weeks post-vaccination (NDV-Herts-challenge strain, 107 EID50 per bird; intramuscular) and clinical symptoms and mortality were recorded daily during 12 days.

From the birds which received a challenge with the USDA-IBV-M41-challenge strain clinical symptoms were recorded four days post-challenge; the birds were then killed and tracheas were collected for virus recovery attempts and for observation of cilial movement. The birds which received a challenge with the USDA-NDV-challenge strain were observed for mortality and clinical signs during 10 days post-challenge.


155

RESULTS The experimental result is presented in Table 1,2,3,4. It is clear, that the combination with the spontaneously hemagglutinating IB strain (4/91) provides better protection against challenge with both a M41-challenge strain and a NDV-challenge strain than the combination with the IB strain which is not spontaneously hemagglutinating chicken erythrocytes (H120). The HI and SN results showed that the best time for vaccination agnst IB and ND in boriler flock is 7 days old. DISCUSSION Administration of mixed vaccine has demonstrated to induce deciliation, hyperplasia, hyperemia and some lesions in the tracheal epithelial mucosa (Nakamura et al., 1992). There is a considerable variation in response of birds to mixed vaccines. Smith (2003) reported that this variation was associated to the IBV strain. Cholakova (1985) studied the immune response induced by live mixed ND (Lasota) and IB (H120 or H52) vaccines. The results showed that simultaneous use of these three vaccines had not any positive or negative effects on the immune response as compared to the administration of single or combined vaccines. Cavanagh and Naqi (2003) have declared that excess IBV particles in the vaccine may interfere with NDV immune response. By virtue of this, it is suggested that combined vaccines are preferred to mixed single vaccines (Cook, 2001). As a general rule, it is indicated that two distinct live vaccines should not be mixed for use and it ought to be administered separately with 14 (at least 7) days interval to avoid any possible interference (Cook, 2001). Monitoring the level of serum antibodies is commonly used as an index to assess the amount of protection induced by vaccination against IBV (Cook et al., 1991). In IBV infections, the level of serum antibodies plays a very important role in prevailing the infection. However, a direct relationship between the serum antibody titer and the level of protection against the infection was still not shown. In fairness to this, the chickens with very low level of serum antibodies were protected against the pathogenic virus (Chabra and Peters, 1985; Cook et al., 1991). It has been recently shown that mucosal immunity in the respiratory system acts as the first line of defense against IBV challenge and resistance against IBV infections may be due to either tracheal mucosal immunity (Chabra and Peters, 1985) or cell mediated immunity (Pakpinyo and Sasipreeyajan, 1993). There are some reports indicating a significant relationship between mucosal (local) IgA and resistance against infection with IBV. Therefore, measuring local IgA could be a suitable alternative for monitoring the level of protection after IBV vaccination or infection (Beard, 1968; Sharma and Adlakha, 1994). Despite the fact that there is not any report regarding the effect of mixed ND + IB vaccine on IB antibody titers, the result of this study indicated that mixed ND + IB vaccine induced higher systemic and local antibody responses as compared to the administration of IB vaccine alone. However, because of histopathologic lesions induced, this method may not be recommended. According to mentioned results the produced (IB+ND ;793/B IBV serotype / Clone 30 NDV) vaccine is useful for protection boliler flocks poultry aginst ND and IB disease which is recommed to use at 7 days old by eye drop method.


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However making a good conclusion about the protection is impossible and further investigation and a field experiment is required.

REFERENCES Beard, CW (1968). An interference type of serological test for infectious bronchitis

virus using Newcastle disease virus. Avian Dis., 12: 658-665. Cavanagh, D and Naqi, SA. (2003). Infectious bronchitis. In: Saif, YM (Ed.),

Diseases of poultry. (11th. Edn.), Ames, Iowa, Iowa State . Chabra, JH and Peters, RW. (1985). Humoral antibody response and assessment of

protection following primary vaccination of chickens with maternally derived antibody against avian infectious bronchitis virus. Res.Vet. Sci., 38: 14-21.

Cholakova, R (1985). Associated vaccination of poultry agains infectious bronchitis, Newcastle disease and infections bursitis.Vet. Med., 22: 32-38.

Cook, JKA; Davison, TF; Huggins, MB and McClaughlan, P. (1991). Effect of in ovo bursectomy on the course of an infectious bronchitis virus infection in line C white leghorn chickens. Arch. Virol., 118: 225-234.

Cook, J (2001). Coronaviridae. In: Jordan, F; Pattison, M; Alexander, D and Faragher, T (Eds.), Poultry diseases. (5th. Ed.), London, W. B. Saunders. PP: 298-306.

Cavanagh, D and Naqi, SA (2003). Infectious bronchitis. In: Saif, YM (Ed.), Diseases of poultry. (11th. Ed.), Ames, Iowa, Iowa State University Press. PP: 101-121.

Kotani, T; Wada, S; Tsukamoto, Y; Kuwamura, M; Yamate, J and Sakuma, S (2000). Kinetics of lymphocytic subsets in chicken tracheal lesions infected with infectious bronchitis virus. J. Vet. Med. Sci., 62: 397-401.

Nakamura, K; Narita, M; Imai, K; Matsumura T; Maeda, M and Tanimura, T (1992). The effect of mixed live vaccines of Newcastle disease and infectious bronchitis on the chicken respiratory tract. J. Comp. Pathol., N. 106: 341-350.

Pakpinyo, S and Sasipreeyajan, J (1993). Efficacy of live lasota Newcastle disease vaccine simultaneously vaccinated with various types of inactivated vaccine. Thaio J. Vet. Med., 23: 1, 35-47.

Sharma, SN and Adlakha, SC (1994). Text book of veterinary virology. 1st. Ed., Vikas Publishing Co., PP: 286-296.

Smith, JA (2003). Impact of Newcastle vaccines on IBV control. Poultry Digest Online. 3: 1-14.

Thornton, DH and Muskett, JC (1973 ). Comparison of immunity to Newcastle disease after vaccination with Newcastle disease vaccine given alone or together with infections bronchitis vaccine. Vet. Rec., 92: 373-374.


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Table 1: Clinical symptoms 1 and 7 days post vaccination

Percentage of birds with clinical symptoms

Group Vaccination

schedule (eye drop) 4 day post

vaccination P.V.

8 day post vaccination

P.V. 1 day 1 0.5

I:IB 793/B IBV serotype 7 day 0.5 0

1 day 1 0.5 II: Clone 30 NDV)

7 day 0.5 0

1 day 2 1.5 III:IB +ND 7 day 1 0 1 day 0 0

Control: 7 day 0 0

Table 2: Percentage of birds with cilia stoping 1 and 7 days postvaccination

Persentage of birds with cilia stoping 1 and 7

Group Vaccination

schedule (eye drop)

4 days post vaccination

8 days post vaccination

1 day 2.5% 1% I:IB 793/B IBV serotype 7 day 2.5% 0.5% 1 day 0 0 II: Clone 30 NDV 7 day 0 0 1 day 16% 18% III:IB +ND 7 day 32% 35% 1 day 0 0

Control: 7 day 0 0


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Table 3: antibody serum post vaccinaton (HI) (SN) 1 and 7 days after post vaccination

Group Vaccination

schedule (eye drop)

Serum antibody using HI

Serum antibody using SN

1 day 0 2 I:IB 793/B IBV serotype 7 day 0 1.8

1 day 3.2 - II: Clone 30 NDV) 7 day 5 - 1 day 3.8 2 III:IB +ND 7 day 5.2 3.6 1 day 0 0

Control: 7 day 0 0

Table 4: Protection combined vaccine against M41 challange strain five weeks post-vaccination

Group Vaccination schedule (eye drop)



Average score cilia

stoping after challenge

1 day 3.8 2 2.5 III:IB +ND 7 day 5.2 3.6 3 1 day 0 0 -

Control: 7 day 0 0 -

Table 5: Protection combined vaccine against ND challange strain five weeks post-vaccination

Group Vaccination schedule (eye drop)



1 day 3.8 2 III:IB +ND 7 day 5.2 3.6 1 day 0 0

Control: 7 day 0 0


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ENHANCED EFFICACY OF THE USE OF A MONOVALENT INFECTIOUS BRONCHITIS VIRUS INACTIVATED VACCINE IN LAYERS PRIMED WITH H120 AND 793B LIVE IBV VACCINES TO INCREASE THE PROTECTION AGAINST

CHALLENGE WITH 3 EUROPEAN SEROTYPES OF IBV

DE WIT1 JJ, VAN DE SANDE1 H and PRANDINI2 F

1 GD (Animal Health Service), Deventer, the Netherlands 2 Merial Italia S.p.A., Milanofiori - Strada 6 - Palazzo E/5 20090 Assago (MI) Italy

SUMMARY This study investigated the efficacy of boosting pullets vaccinated with the live Infectious Bronchitis Virus (IBV) vaccines H120 (Massachusetts type strain) and GALLIVAC® IB88 (CR88 strain belonging to the 793B variant group) with the monovalent IBV M41 component of the multivalent inactivated vaccine GALLIMUNE® 407 ND+IB+EDS+ART (Merial) against M41, D388, and 793B challenges. Respiratory signs, serological response, ciliostasis and egg production were monitored. The protection afforded by the live vaccinations alone, against the M41 challenge was already close to 100%, so a potential enhanced efficacy of the booster dose could not be detected. The laying hens that had been boosted with the inactivated vaccine displayed a low level of ciliostasis after the 793B and D388 challenges, 5% and 11% respectively, whereas the non-boosted laying hens showed 19% and 23% ciliostasis respectively. The laying hens that were boosted with the inactivated vaccine showed (very) limited drops in egg production after the 793B and D388 challenges, 15% and 5% respectively, whereas the non-boosted laying hens showed drops in egg production of 30% and 23% respectively. It was concluded that the use of this monovalant IBV vaccine increased the level of protection against the heterologous challenges with 793B and D388 viruses. INTRODUCTION Infections with Infectious Bronchitis Virus (IBV) are a known cause of respiratory problems, drops in egg production, poor egg shell quality, drops in hatchability and sometimes nephritis and false layers in chickens (Cavanagh & Gelb, 2008). Many papers report the findings of experimental infections in young birds using a large variety of IBV challenge strains. The number of papers that report the results of experimental infections of hens in lay are relatively scarce whereas vaccination


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challenge studies in laying birds are even scarcer. Most of these papers are several decades old and report the results of vaccination and challenge experiments with the IBV strains that were most prevalent at that time. At the present time, these strains represent only a minority of the IBV strains that are currently being detected in the field. The number of types (usually serotypes or genotypes) of strains that have been detected and reported since then has increased significantly (Davelaar et al., 1984; Cook et al., 1999; Di Fabio et al., 2000; Cavanagh, 2003; Jackwood et al., 2005; Ignjatovic et al., 2006; Liu et al., 2006; Villarreal et al., 2007; Worthington et al., 2008; Chen et al., 2009). To achieve a broad protection against damage by challenge with IBV variants at several weeks post vaccination, the use of certain combinations of live IBV vaccines has been shown to be able to induce a high level of protection against challenges with several heterologous strains (Cook et al., 1999; Gelb Jr et al., 2005), although the same combinations can be unsuccessful for other heterologous strains (Ladman et al., 2002). In order to achieve an increased level of protection during the laying period of commercial layers and parent stocks, the use of inactivated IBV vaccines after a priming with live IBV vaccines has been shown to be effective against homologous Massachusetts challenges (Gough et al., 1977; Box et al., 1980; Gough et al., 1981; Timms & Bracewell, 1983; Box & Ellis, 1985; Muneer et al., 1987; Box et al., 1988). The efficacy of increasing the level of protection against heterologous challenges in the laying period has rarely been reported. Birds that had been vaccinated twice with a live Massachusetts type vaccine and had been boosted with a killed oil-emulsion vaccine containing a Massachusetts strain showed no protection against a challenge with a strain of the Arkansas type (Muneer et al., 1987). In Europe, many variant IBV strains have been detected in recent years (Worthington et al., 2008). One of these strains, D388 (serotype) also known as the QX-like strain (genotype) has been associated with respiratory problems, nephritis, drops in egg production and a ‘false layers’ syndrome. In Europe, the D388 strain has first been detected in a Dutch broiler flock, at 18 days of age, suffering from nephritis in November 2003. This was only a few months after the large scale import of pullets from other parts of Western and Eastern Europe to compensate for the hens that had been culled in the spring of 2003 due to the highly pathogenic H7N7 avian influenza epidemic in the Netherlands. Since then this strain has been detected in many other European countries. In 2007 and 2008, it was by far the most detected IBV field strain in the Netherlands (data AHS Deventer, unpublished). At the moment, no homologous D388/QX vaccine is available. It has been reported by several groups that a vaccination schedule using a live Massachusetts vaccine at day 1 of age followed by a 793B-like vaccine at 14 days, is able to induce a considerable cross protection against a QX field strain in young SPF birds (Worthington & Jones, 2006; Terregino et al., 2008). There are no reports on how best to protect hens against a D388/QX challenge during the laying period. This paper reports the results of a vaccination-challenge study in laying hens. The efficacy against homologous M41 and 793B and a heterologous D388 IBV challenges was investigated in pullets vaccinated with live H120 (1 vaccination) and GALLIVAC IB88 (serotype 793B live vaccine) (2


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vaccinations) and boosted with the monovalent IBV M41 component of a commercially available multivalent inactivated vaccine GALLIMUNE® 407 ND+IB+EDS+ART. The parameters monitored were egg production, respiratory signs, ciliostasis and serological response (using the virus neutralization test (VNT)). For two of the challenges, the immunoperoxidase test (IPT) for the detection of IBV in kidneys and uterus were carried out as well. MATERIALS and METHODS Hens, experimental design and housing 18-day-embryonated eggs from a well performing commercial Lohmann Brown layer breeder flock were transported to GD. After disinfection, the eggs were hatched at the animal facility of GD. After hatching, pullets were housed in a floor pen (HEPA filtered air, strict hygiene barrier) in the animal facility of GD. All pullets were vaccinated at day of hatch with a live vaccine of the Massachusetts serotype (H120), and at 28 and 70 days with a vaccine of the 793B serotype (CR88). The pullets were not beak-trimmed. At week 15, the birds were individually marked using tags and divided into 2 different groups (separated by a fence), named Primed and OEV (see table 1). The pullets in the group OEV were vaccinated with an inactivated oil emulsion vaccine (OEV) containing the M41 strain. The other birds (Primed) were not revaccinated. At 24, 26, and 32 weeks of life, 20 laying hens from the group OEV were placed in an isolator with a laying nest and were challenged with an IBV strain. At the same ages, 2 groups of 20 laying hens from group ‘Primed’ were placed in 2 separate isolators with a laying nest. One group (Primed-Chal) was challenged with IBV, the other group (Primed-Neg) was inoculated with allantoic fluid containing no IB virus as controls. The hens were challenged 2 days after transfer into the isolators and monitored for 8 days (except for the M41 challenge that was monitored for 5 days). Feeding and water Water and feed was supplied ad libitum Lighting According to advice of breeding company Vaccines and application The following vaccines (Merial) were used: BIORAL® H1201: freeze dried live vaccine, batch 83983 (2000 doses per vial). The content of one vial was dissolved in 200 ml of cold (4°C) sterile, demineralised water. One dose of vaccine was applied to each of the one-day-old chicks by an eye-drop of 0.05 ml and one droplet of 0.05 ml into the nostril. GALLIVAC IB882: freeze dried live vaccine of the 793B serotype, batch L196716 (2000 doses per vial). The content of one vial was dissolved in 200 ml of cold (4°C) sterile, demineralised water. One dose of vaccine was applied by one droplet (0.05

1 ® BIORAL is a registered trademark of Merial in the United States of America and elsewhere. 2 ® GALLIVAC and GALLIMUNE are registered trademarks of Merial in the United States of America and elsewhere.


162

ml) into the eye and one droplet (0.05 ml) into the nostril of each of the pullets at the age of 28 and 70 days. GALLIMUNE 407 ND+IB+EDS+ART (lot L206232) is an inactivated oil emulsion vaccine that contains the M41 strain of IBV: hens of group OEV were injected with 0.3 ml of the vaccine into the breast muscle, using a calibrated injection syringe. Challenge viruses and application The laying hens of Primed-Chal and OEV groups were challenged using low passage pathogenic strains from 3 different serotypes of IBV. Each challenge strain had been used in earlier experiments causing 100% ciliostasis in unvaccinated chickens. The IBV strains used, challenge dose per bird, application route and age at challenge with the different viruses were: IBV M41 (GD, HSL 514-02) titre of 107.2 EID50, eye drop, 24 weeks. IBV D388 (GD, HSL 514-01) titre of 107.0 EID50 eye drop, 26 weeks. IBV CR88121 (Merial, serotype 793B) titre of 106.9 EID50 intratracheal, 32 weeks. The negative control group (Primed-Neg) was inoculated with IBV-negative allantoic fluid. Serology and blood sampling At several time points during the study, blood samples were collected from the birds. All sera was stored at -20°C until testing. All sera was tested in the same test run. The sampling timings and bird numbers were as follows:

1. Day of hatch, 30 birds. The sera was pooled into 5 pools. 2. 28 days of life, 30 birds. The sera was pooled into 5 pools. 3. 15 weeks of life, 20 birds were blood sampled and the sera was tested

individually. 4. 22 weeks of life, 30 hens from groups OEV and Primed were blood sampled

and the sera was tested individually. The samples (pooled or individual) were tested for the presence and amount of virus neutralising antibodies against IBV using M41, 793B, and D388 antigens. The test was performed using primary chicken kidney cells in presence of the appropriate controls. The starting dilution of the VNT was the 1:128 dilution of the serum, which is used routinely by GD for older birds that have been vaccinated with different serotypes of IBV. Ciliostasis At 5 days post challenge, tracheal protection was measured in 10 birds from group OEV and in 5 birds from groups Primed-Chal and Primed-Neg. The level of tracheal protection was determined 5 days post challenge (d.p.c.) using the ciliostasis test on 5 tracheal rings per bird. The tracheas were placed in Hanks medium immediately after euthanasia of the chickens. Subsequently, 5 rings (equally divided over the total length of the trachea) were cut and placed in the medium at 37ºC. The level of ciliostasis in each sample, was determined independently by two technicians between 1 and 4 hours after euthanasia. The degree of beating of the cilia per ring was expressed as 0 (100% beating of cilia), 1 (75 - 99% still beating), 2 (50 – 75 % still beating), 3 (25 - 50% beats) or 4 (0 - 25% of the cilia beats). One bird can score between 0 and 20 (5 rings x score 4).


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Egg production Initially the number of eggs that were collected during the 5 days post challenge was used to measure the egg production of each group. However, several groups started egg pecking which undermined the reliability of the number of collected eggs as parameter for the egg production. Therefore, an alternative method was used for the D388 and 793B challenge groups and the observation period was extended from 5 to 8 days post challenge. At post mortem at day 5 and 8, the activity of the ovary and oviduct was used as an approximate guide to egg production. A hen was considered to be in production on that day if she had an active ovary and oviduct. If the hen had a degenerated ovary, the hen was considered to be out of production. If the hen had a degenerated ovary, but also an egg in the oviduct, the hen was considered to be in production at 50%. Statistical analyses The serological results were statistically analysed by using the programme Statistix, using the two-tailed Student T-test. The groups were tested for equal or unequal variances. The differences in mean titres were considered to be significant when P < 0.05. RESULTS Serology The average levels of virus neutralising antibodies against the 3 challenge IB viruses at the different ages are summarized in table 2. The VNT titres from the 1-day-old chickens showed that they had a high level of IBV maternally derived antibodies as can be expected from Dutch layer breeders. The 15-week-old Primed hens had detectable VNT titres against the 3 serotypes of IBV. The VNT titres from group Primed at 22 weeks of age were on average 0.9 log2 lower than the titres at 15 weeks of age. Two titres had dropped significantly (M41 (P = 0.04), D388 (P = 0.01)). At 22 weeks of age, 7 weeks after the administration of the inactivated vaccine in group OEV, only the M41 titre had increased significantly with 2.3 log2 (P = 0.0000) compared to the titres at 15 weeks of age. Ciliostasis The hens from the group Primed-Neg that had been challenged with IBV-negative allantoic fluid did not show any sign of ciliostasis at 5 d.p.c. (table 3). Hens from the group Primed-Chal showed an average protection against ciliostasis at 5 d.p.c. of 77% (D388), 81% (793B) and 96% (M41) with an average of 85%. Birds of group OEV showed an average protection against ciliostasis at 5 d.p.c. of 89% (D388), 95% (793B) and 99% (M41) with an average of 94%. Egg production The egg production at 5 and 8 days post challenges is reported in table 4. All birds of group Primed-Neg were producing an egg at 5 and 8 days post inoculation of the IBV-negative allantoic fluid. The M41 challenge did not result in a drop in egg production in any of the groups. The D388 challenge resulted in a drop in egg production at 5 d.p.c. of 40% in the Primed-Chal group and a drop of 10% in the OEV


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group. At 8 d.p.c. the drop in egg production of the Primed-Chal group was 7% and 0% in the OEV group. The 793B challenge resulted in a drop in egg production at 5 d.p.c. of 60% in the Primed-Chal group and a drop of 10% in the OEV group. At 8 d.p.c. the drop in egg production of the Primed-Chal group was 0% and 20% in the OEV group. IPT IBV The Immuno-peroxidase test for IBV was carried out at 5 and 8 d.p.c. on the kidneys and uterus of the birds that had been challenged with D388 and 793B. The results are listed in tables 5 and 6. IB viruses D388 or 793B were not detected in any of the uteruses of the challenged birds. 793B was also not detected in any of the tested kidneys. D388 was detected at 5 d.p.c. in 0% and 80% of the kidneys of the Primed-Neg and Primed-Chal respectively. At 8 d.p.c. it was detected by IPT in 0, 30, and 20% of the kidneys of groups Primed-Neg, Primed-Chal, and OEV respectively. DISCUSSION This study investigated the efficacy of boosting pullets vaccinated with the live IB vaccines H120 and CR88 (793B like) with a monovalent IB M41 component of the multivalent inactivated vaccine GALLIMUNE 407 ND+IB+EDS+ART (Merial) against a M41, D388, and 793B type challenges. Regarding the serological response against the M41 strain, a significant rise in VNT M41 titres was observed after use of the inactivated vaccine. However, since the protection against ciliostasis and drop in egg production afforded by the live vaccinations alone was already 100%, it was impossible to demonstrate a further increase in the protection level against that challenge. Concerning the serological response against the D388 strain, no effect was observed on the level of VNT D388 titres after use of the inactivated vaccine. However, the average degree of ciliostasis after the D388-QX challenge of the OEV group was 11% versus 23% in the Primed–PC group. The average drop in egg production at 5 and 8 days post challenge was also lower in the OEV group (5%) compared with the drop in egg production of 23% in the Primed-Chal group. The percentage of birds that was positive in the Imunoperoxidase test on kidneys was also much lower in the OEV group than in the Primed-Chal group. Considering the serological response against the 793B strain, no effect was observed after use of the inactivated vaccine on the level of VNT 793B titres. However, in the OEV group the average degree of ciliostasis after the 793B challenge was 5% versus 19% in the Primed–PC group. The average drop in egg production at 5 and 8 days post challenge was also lower in the OEV group (15%) compared with the drop in egg production of 30% in the Primed-Chal group. Based upon these results it can be concluded that besides an increase of homologous protection as shown by the 793B groups, cross protection against the heterologous D388(QX-like) serotype can be achieved with a vaccination programme including immunisation with live vaccines against Mass and 793B serotypes boosted by an OEV which contains only the IBV Mass component.


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REFERENCES Box, P. G., Beresford, A. V. & Roberts, B. (1980) Protection of laying hens against

infectious bronchitis with inactivated emulsion vaccines. Vet Rec, 106, 264-268.

Box, P. G. & Ellis, K. R. (1985) Infectious bronchitis in laying hens: interference with response to emulsion vaccine by attenuated live vaccine. Avian Pathol, 14, 9-22.

Box, P. G., Holmes, H. C., Finney, P. M. & Froymann, R. (1988) Infectious bronchitis in laying hens: the relationship between haemagglutination inhibition antibody levels and resistance to experimental challenge. Avian Pathol, 17, 349-361.

Cavanagh, D. (2003) Severe acute respiratory syndrome vaccine development: experiences of vaccination against avian infectious bronchitis coronavirus. Avian Pathol, 32, 567-582.

Cavanagh, D. & Gelb, J. J. (2008). Infectious Bronchitis. In Y. M. Saif, Fadly, A.M., Glisson, J.R., McDougald, L.R., Nolan, L.K. and Swayne, D.E. (Ed), Diseases of Poultry (12th ed., pp. 117-135).

Chen, H. W., Huang, Y. P. & Wang, C. H. (2009) Identification of Taiwan and China-like recombinant avian infectious bronchitis viruses in Taiwan. Virus Res, 140, 121-129.

Cook, J. K. A., Orbell, S. J., Woods, M. A. & Huggins, M. B. (1999) Breadth of protection of the respiratory tract provided by different live-attenuated infectious bronchitis vaccines against challenge with infectious bronchitis viruses of heterologous serotypes. Avian Pathology, 28, 477-485.

Davelaar, F. G., Kouwenhoven, B. & Burger, A. G. (1984) Occurrence and significance of infectious bronchitis virus variant strains in egg and broiler production in the Netherlands. Vet Q, 6, 114-120.

Di Fabio, J., Rossini, L. I., Orbell, S. J., Paul, G., Huggins, M. B., Malo, A., Silva, B. G. & Cook, J. K. (2000) Characterization of infectious bronchitis viruses isolated from outbreaks of disease in commercial flocks in Brazil. Avian Dis, 44, 582-589.

Gelb Jr, J., Weisman, Y., Ladman, B. S. & Meir, R. (2005) S1 gene characteristics and efficacy of vaccination against infectious bronchitis virus field isolates from the United States and Israel (1996 to 2000). Avian Pathol, 34, 194-203.

Gough, R. E., Allan, W. H. & Nedelciu, D. (1977) Immune response to monovalent and bivalent Newcastle disease and infectious bronchitis inactivated vaccines. Avian Pathol, 6, 131-142.

Gough, R. E., Wyeth, P. J. & Bracewell, C. D. (1981) Immune responses of breeding chickens to trivalent oil emulsion vaccines: responses to infectious bronchitis. Vet Rec, 108, 99-101.

Ignjatovic, J., Gould, G. & Sapats, S. (2006) Isolation of a variant infectious bronchitis virus in Australia that further illustrates diversity among emerging strains. Arch Virol, 151, 1567-1585.

Jackwood, M. W., Hilt, D. A., Lee, C. W., Kwon, H. M., Callison, S. A., Moore, K. M., Moscoso, H., Sellers, H. & Thayer, S. (2005) Data from 11 years of molecular typing infectious bronchitis virus field isolates. Avian Dis, 49, 614-618.

Ladman, B. S., Pope, C. R., Ziegler, A. F., Swieczkowski, T., Callahan, C. J., Davison, S. & Gelb, J., Jr. (2002) Protection of chickens after live and


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inactivated virus vaccination against challenge with nephropathogenic infectious bronchitis virus PA/Wolgemuth/98. Avian Dis, 46, 938-944.

Liu, S. W., Zhang, Q. X., Chen, J. D., Han, Z. X., Liu, X., Feng, L., Shao, Y. H., Rong, J. G., Kong, X. G. & Tong, G. Z. (2006) Genetic diversity of avian infectious bronchitis coronavirus strains isolated in China between 1995 and 2004. Arch Virol, 151, 1133-1148.

Muneer, M. A., Newman, J. A., Halvorson, D. A., Sivanandan, V. & Coon, C. N. (1987) Effects of avian infectious bronchitis virus (Arkansas strain) on vaccinated laying chickens. Avian Dis, 31, 820-828.

Terregino, C., Toffan, A., Beato, M. S., De Nardi, R., Vascellari, M., Meini, A., Ortali, G., Mancin, M. & Capua, I. (2008) Pathogenicity of a QX strain of infectious bronchitis virus in specific pathogen free and commercial broiler chickens, and evaluation of protection induced by a vaccination programme based on the Ma5 and 4/91 serotypes. Avian Pathol, 37, 487-493.

Timms, L. M. & Bracewell, C. D. (1983) Cell mediated and humoral immune response of chickens to inactivated oil-emulsion infectious bronchitis vaccine. Res Vet Sci, 34, 224-230.

Villarreal, L. Y., Brandao, P. E., Chacon, J. L., Saidenberg, A. B., Assayag, M. S., Jones, R. C. & Ferreira, A. J. (2007) Molecular characterization of infectious bronchitis virus strains isolated from the enteric contents of Brazilian laying hens and broilers. Avian Dis, 51, 974-978.

Worthington, K. J., Currie, R. J. & Jones, R. C. (2008) A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. Avian Pathol, 37, 247-257.

Worthington, K. J. & Jones, R. C. (2006) New genotype of infectious bronchitis virus in chickens in Scotland. Vet Rec, 159, 291-292.

Table 1. Study design

Vaccination (age in days) challenge Group 1 28 70 105

Primed NC H120 793B 793B - - Primed Primed C H120 793B 793B - M41, 793B or D388 OEV H120 793B 793B OEV M41 M41, 793B or D388 Table 2. VNT titres of the different groups at different ages

Mean VNT (log2) titres against the serotypes (challenges)

Group Age (weeks)

M41 D388 793B - 0 11.4 10.2 12.6

15 7.4 9.2 9.3 22 6.5 8.1 8.5

Primed

∆ 22-15 -0.8 -1.1 -0.7 22 9.7 8.6 9.6 OEV

∆ 22-15 +2.3 -0.6 +0.3


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Table 3. Results of the ciliostasis test at 5 days post challenge

Group challenge Average protection (%) against ciliostasis at 5 days post challenge with

M41 D388 793B mean Primed-Neg No 99 100 100 100 Primed-Chal Yes 96 77 81 85

OEV Yes 99 89 95 94 Table 4. Egg production at 5 and 8 days post challenge

Group Days post challenge

Average egg production at 5 and 8 days post challenge with IBV strain

M41 D388 793B mean 5 100 100 100 100 Primed-Neg* 8 - 100 100 100 5 100 60 40 67 Primed-Chal 8 Nd** 93 100 97 5 100 90 90 93 OEV 8 Nd** 100 80 90

* Challenged with IBV-negative allantoic fluid ** Not done Table 5. IPT IBV on kidney and uterus at 5 and 8 d.p.c. with D388

Percentage of birds that was IPT IBV negative Kidney uterus

Group challenge D388

5 d.p.c. 8 d.p.c. 5 d.p.c. 8 d.p.c. Primed-Neg no 100 100 100 100 Primed-Chal yes 20 70 100 100

OEV yes ND 80 ND 100 Table 6. IPT IBV on kidney and uterus at 5 and 8 d.p.c. with 793B strain 88121

Percentage of birds that was IPT IBV negative Kidney uterus

Group challenge 793B

5 d.p.c. 8 d.p.c. 5 d.p.c. 8 d.p.c. Primed-Neg no 100 100 100 100 Primed-Chal yes 100 100 100 100

OEV yes 100 100 100 100


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DEVELOPEMENT OF IN OVO VACCINE AGAINST INFECTIOUS BRONCHITIS IN POULTRY FLOCKS

KHALESI B, MASOUDEI S and MOGHADAMPOUR M

Razi Vaccine & Serum Research Institute Razi Vaccine & Serum Research Institute, Hasarak, Karaj, Iran.

SUMMARY The Infectious Bronchitis virus (IBV), a Coronavirus, targets not only the respiratory tract but also the urogenital tract. The virus can spread to different organs throughout the chicken. Infection initially causes respiratory disease in the infected birds and also drops in egg production in layers and breeders. Kidney damage can also occur. Although morbidity is high, the mortality rate in an infected flock can vary, depending on standards of management and on possible involvement of secondary bacterial infections. Control is best achieved by improved biosecurity and vaccination. Commercially-available vaccines for IB are not administered in ovo. Rather, they are administered post-hatch in a variety of formats. Briefly, such vaccines are typically administered by the labor-intensive methods of spraying (e.g., handspray, knapsack spray, or automated spray. In ovo vaccination is an emerging trend in the poultry industry because of its advantages like negligible manpower involvement, induction of neonatal resistance and better protection. This study describes that following in ovo administration of live, attenuated H120 IB virus in a freeze-dried environment. The potential vaccine virus replicated in embryos and newborn chicks, did not hamper hatchability, successfully induced antibody response and conferred protection against the disease in broiler birds. Experimental results establish the safety and efficacy of the in ovo vaccine (IOV) relative to in ovo administration to chickens. Vaccine virus has the potential for application as in ovo vaccine against infectious bronchitis in commercial chicken. INTRODUCTION Infectious bronchitis virus (IBV) is one of the primary causes of respiratory disease in domestic fowl. IBVs primarily and initially infect the trachea though they can also infect the kidney and oviduct and other epithelial surfaces. Infection with IBV reduces the performance of broilers and in laying birds drops in egg production and egg


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quality can occur. Live-attenuated vaccines against infectious bronchitis are generally given to chicks at 1 day of age in the hatchery by spray, or later in the field by either drinking water or spray. However, as with any live-attenuated vaccine administered via the respiratory tract, which must replicate in order to be effective, vaccination at an early age can damage the epithelial lining of the trachea and depending on the extent of this damage, secondary bacterial infections can occur. For this reason it is important to assess post vaccinal reactions when preparing vaccine strains. Vaccination in ovo against Marek’s disease and infectious bursal disease is commonplace in the USA [1]. In ovo delivery gives the advantage that all vaccinations can be done prior to hatching as long as the vaccines are shown to be compat ible and safe. Whilst considerable work has been done on in ovo delivery of Marek and infectious bursal disease vaccines, little work has been done on delivering IBV vaccines in ovo. This is due to the fact that most strains of IBV are embryo lethal, both in the 9–12-day-old embryos that are used for producing IB vaccines, and in 18-day-old embryos.One group has shown that in ovo vaccination with IBV can be successfully accomplished, although the strain used had residual pathogenicity even at low titres. As uniform mass application of IBV vaccines post hatch can be problematic with regard to obtaining even distribution of spray or ensuring all birds receive the correct dosage from drinking water, delivering IBV in ovo, via which a precise dosage can be achieved, could bring significant advantages to the poultry industry. Commercially available vaccines for IB are not administered in ovo. Rather, they are administered post-hatch in a variety of formats. Briefly, such vaccines are typically administered by the labor-intensive methods of spraying (e.g., hand spray, knapsack spray, or automated spray equipment) or in drops (eye or nose). A method of vaccinating a poultry animal against infectious bronchitis virus (IBV), said method comprising administering an IBV vaccine in ovo to a developing chick in a fertilized egg that has not yet hatched; wherein said IBV vaccine comprises a solution comprising a live attenuated strain of IBV of 10−1.0 EID50 to 100.0 EID50 per dose per egg; wherein at least 80% of eggs receiving said vaccine hatch; and wherein at least 89% of chicks hatched from said eggs are protected against a challenge of virulent IBV administered to said chicks, and wherein the percent of chicks protected from said challenge is determined by a cilia stopping test (CST). As more fully explained below, the in ovo vaccines of the present paper provide distinctive advantages over the inconvenient and time-consuming post-hatch routes of administration presently available.


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MATERIALS and METHODS Assessment of clinical signs post hatch Examination of general health was done daily during the course of the experiments. Nasal discharge was assessed by gently squeezing the nares of the chicks and determining if any fluid was visible. Experiment 1: Safety study for in ovo chicken vaccination Embryonated eggs Chick starter crumbs and water were provided Specified Pathogen Free (SPF) White Leghorn chicken embryonated eggs (Lohmann, Germany) were used. All chicks were hatched in a contained environment and transferred to negative pressure isolators for the remainder of the chick starter crumbs and water were provided ad libitum during the course of the experiments. Specific-pathogen-free (SPF) chicken eggs were commercially obtained from Lohman company of Germany. In brief, SPF eggs were incubated within appropriate facilities. At 18 days of incubation, 4 groups of 25 eggs were administered in ovo vaccinations. Vaccination procedures The IOV of the present study were prepared as follows. A commercially-available H120 IB vaccine from Razi vaccine and serum institute was obtained. This vaccine contains live, attenuated IB virus in a freeze-dried environment. Prior to its use herein, this vaccine contained a titer of 10 6.4 EID50 IB virus per vial. Next, this vaccine was reconstituted in saline and then further admixed with saline until the following concentrations were obtained: solutions containing, respectively, titers of 102.0, 101.0, 100.0, and 10−1.0 of IB virus/vaccine per dose (size: 0.05 ml) were prepared. At 18 days of incubation, Groups 1-4 (each consisting of 25 eggs) were injected in ovo with a dose of 0.05 ml per egg of the vaccines of the present invention containing, respectively, the following titers of IB virus: titers of 102.0, 101.0, 100.0, and 10−1.0 of IB virus/vaccine per dose. As a control, Group 5 (also consisting of 25 eggs) did not receive any in ovo injections at day 18 of incubation. To administer the injections, one ml glasses syringe have been used. Until hatching, both the inoculated eggs (i.e., 100 eggs total in Groups 1-4@25 eggs/group) and the control eggs (i.e., 25 eggs in Group 5) were incubated within the same incubator. The number of hatched eggs per group was experimentally recorded at days 20, 21, and 22 of incubation. Experiment 2: Efficacy study for in ovo vaccination of commercial chicken eggs. Commercial chicken eggs for broilers were obtained from Karaj province of IRAN. These eggs were incubated within appropriate facilities. After 18 days incubation, all eggs were candled, and 4 groups of 28-30 eggs were inoculated with graded doses of the in ovo vaccine. The in ovo vaccines administered, hatching and/or husbandry conditions, the manner of egg injections, the preparation of the challenge virus, serological analysis, and the determination of protection using the CST methodology.

mailto:1-4@25


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In brief, 6 days post-challenge (hereinafter, “PC”), the chicks were killed and their tracheas removed. One part per trachea was collected for microscopic examination and cillary activity was assessed. This assessment used the following scale:+=full movement;±=impaired movement; and−=no movement. In cases of impairment, additional trachea parts were taken to confirm this finding. The percentage of protection against challenge with this virulent strain of IB was calculated using the following formula: protection % = (A+½B)(100)/C wherein A=# chicks assessed+; B=·chicks assessed±; and C=total# chicks. RESULTS The results were all conducted in experiments 1 and 2. Tables 1- 2 below present the results from this study of in ovo vaccination with IB virus of commercial chicken eggs. Results of experiment 1 Hatchability in the inoculated eggs ranged from 70% to 82% in comparison to 95% for the negative control eggs of Group 5. All of these observed hatchability percentages were within customary limits. No systemic effects between the inoculated groups (i.e., Groups 1-4) were observed. Based upon the results of Experiment 1 as set forth above, it was concluded that in ovo vaccination at incubation day 18 using dosages ranging from a low of 10 −1.0 EID 50 IB vaccine to a high of 10 2.0 EID 50 IB vaccine was safe relative to hatchability. Results of experiment 2 As set forth above in Tables 5 and 6 hatchability in the inoculated groups (i.e.,Groups 1-4) was good,with customary limits, and ranged from 84% to 91%. Hatchability in the control group (i.e., Group 5) was 84%. Table 5 above sets forth the conditions and mortality of the chicks from Groups 1-5 prior to challenge with a virulent strain of IB virus. In Table 5, “BC” is used as an abbreviation for bad condition. To summarize the results: 2 chicks in Group 1 died (1 from yolk sac inflammation); and 2 control chicks from Group 5 were in bad condition (1 died on day 9 post-hatch and the other 1 chick was killed on day 9 post-hatch because it could not stand upright). Table 5 does not present clinical respiratory signs. Several chicks in groups given dosages of 100.0 or greater (i.e., Groups 1-3) showed mild respiratory signs probably due to IB virus replication from 6 days of age onward until challenge at 3 weeks of age (Groups 1 and 2) or until 12 days of age (Group 3). The observed respiratory signs were mild and no damage was seen at 6 days post-challenge as determined by the CST methodology (presented below). Tables 6 and 7 presents the results of the challenge study. In Tables 8 and 9, 1 chick in Group 2 was in bad condition from the day after challenge until its death at 5 days post-challenge. This death was not attributed to either the in ovo vaccination or the subsequent challenge. As determined by the CST methodology, the protection afforded commercial eggs for broilers by in ovo vaccination at incubation day 18 with IB virus/vaccine derived from Poulvac® IB MM


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against exposure to a virulent challenge IB virus at 3 weeks of age was excellent. The protection percentages ranged from a low of 89% to a high of 100%. The control chicks (Group 5) had no protection against challenge with a virulent IB virus at 3 weeks of age. Serological analysis yielded the results set forth below in tables 8 and 9. As set forth above in Tables 8 and 9, serological analysis of serum samples revealed mean antibody titers in all inoculated groups were only slightly higher than those of the control chicks (i.e., Group 5). However, within the inoculated groups, the highest mean antibody titer was measured in Group 1 chicks; i.e., the chicks which had received the strongest dose of the in ovo vaccines. Additionally, in the vaccinated chicks, a considerable number showed antibody titers at or above the detection level. Based upon the entirety of this experiment 2 as described above, it was concluded that in ovo vaccination at day 18 of incubation of commercial chicken eggs for broilers with IB virus/vaccine at dosages of ranging from a low of 10 −1.0 EID 50 per egg to a high of 10 2.0 EID 50 per egg was efficacious in protecting chicks against exposure to a challenge at 3 weeks of age with virulent IB M41 virus. DISCUSSION As the poultry industry is moving towards greater use of in ovo vaccination, the purpose of this study was to determine if IBV vaccines based on the Beaudette strain could be delivered via this route. An objective specific to the poultry industry is the production of vaccines that can be administered in ovo i.e. to embryos of 18 days of age, three days prior to hatch. The technology exists to apply vaccines very precisely in this way, more efficiently than vaccinating hatched birds by spray or drinking water. To date this objective has been achieved only in respect of Marek’s disease virus. Vaccine strains of other viruses, including IBV, adversely affect hatchability. Our molecularly cloned IBV is remarkably benign when given to 18-day-old embryos; hatchability is unaffected. Furthermore, preliminary experiments have shown that our spike-swapped chimaeras induce protective immunity following in ovo vaccination. The quantity of pathogenic agent to be included in the vaccine to be administered to the host egg can vary, depending upon the particular pathogenic agent, and also the size of the animal (larger animals may require larger quantities of agent). A desired quantity is within the range of about 10 −1.0 EID 50 to about 10 2.0 EID 50 of pathogenic agent, e.g. virus (in particular IB), per vaccine dose. A quantity within the range of about 10 0.0 EID50 to about 10 1.0 EID 50 pathogenic agent per vaccine dose is also useful herein. Throughout this application, “EID 50 ” refers to a 50% egg infectious dose. In addition to the foregoing, the vaccines may be formulated with known additives, including adjuvants. Examples of desirable adjuvants include polymers and copolymers of acrylic acid, as well as those derived from other alkyl esters. Other constituents include media such as water, saline solution, or water-in-oil emulsions in quantities sufficient to top off the dose. The pathogenic agent may be dissolved or suspended in the media just described. In a preferred embodiment of the invention, the vaccine is formulated with substantially no virus neutralizing factor.


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A vaccine dose is typically within the range of about 0.001 mL to about 1.0 mL, and more preferably within the range of about 0.01 to 0.1 mL, with about 0.05 mL being even more preferred. The dosing regimen for the vaccine most desirably includes administration in ovo to a developing chick in a fertilized egg that has not yet hatched. One administration of the dose is typically preferred, but more than one is within the scope of the invention. The dosing schedule chosen should ensure both the safety of the developing animal, as well as efficacy of immunization. A dose of vaccine may be administered in ovo during a time period which is within the range of about day 1 up to and including about a few minutes before hatching. More preferably, a dose is delivered in ovo within the time period of about day 5 to about day 25. Even more desirably, the dose is administered during the period of about day 10 to about day 20. A dosing at about day 18 may be particularly desirable. Administration of the vaccine may be done by hand, but is more typically and economically administered using commercially available egg injection equipment, such as that available from Embrex, Inc. of North Carolina. An advantage of in ovo vaccine according to the invention is that the vaccine is applied to each individual bird (egg). This translates into better accuracy as compared with more traditional vaccination programs (non-in ovo). This is reflected in the high percentages of protection against challenge in the vaccinated birds. In addition, because the in ovo birds are vaccinated at a considerably earlier age than are those who receive the inoculation post-in ovo, there is more time for the birds to develop their immunity before exposure to outside ambient conditions in which virulent strains of the virus may be present. This result has been unexpected. Normally, introduction of live viruses into embryos would have been expected to generate fairly lethal results. A developing embryo is a highly fragile organism, and the presence of a live virus such as IB would have normally killed the developing animal. At best, much of the art has mandated the use of a virus neutralizing factor to prevent such an occurrence when chicks are inoculated in ovo. Conversely, introduction of what would have been considered exceedingly minute quantities of live virus, while not killing the embryo, would not have been expected to impart satisfactory immunogenic properties to the organism. To confirm the potential use of these candidate vaccines in the field, future work will concentrate on the effect of maternally derived antibody on the efficacy of these viruses as vaccine candidates following in ovo vaccination. REFERENCES Ricks CA, Avakian A, Bryan T, Gildersleeve R, Haddad E, Ilich R, et al. 19990 .In ovo

vaccination technology. Adv Vet Med;41:495–515. Wakenell PS, Sharma JM. Chicken embryonal vaccination with avian infectious

bronchitis virus. Am J Vet Res 1986;47:933–8. Geilhausen HE, Ligon FB, Lukert PD. The pathogenesisof virulent and avirulent

avian infectious bronchitis virus. Arch Gesamte Virusforsch 1973;40:285–90. Chew et al., Avian Disease 41:598-603, 1997, Pathogenicity of Attenuated Infectious

Bronchitis Virus for Oviducts of Chickens Exposed in Ovo.


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Wakenell, et al., Am J Vet Res, vol. 47, No. 4, Apr. 1986, Chicken embryonal vaccination with avian infectious bronchitis virus.

Wakenell, et al., Avian Diseases 39:752-765, 1995, Embryo Vaccination of Chickens with Infectious Bronchitis Virus: Histologic and Ultrastructural Lesion Response and Immunologic Response to Vaccination.

Tarpey I., Orbell S.J., Casais R., Hodgson T., Lin F., Hogan E., Cavanagh D., Safetyand efficacy of an infectious bronchitis virus used for chicken embryo vaccination, Vaccine (2006) 24:6830–6838.

Table 1: Calculated Hatchability after in ovo vaccination at incubation day 18 with a dose of 0.05 ml/egg of IB vaccine to Groups 1- 5 (25 eggs/group)

Group EID 50 # hatched % hatched 1 10 2.0 17 72 2 10 1.0 21 82 3 10 0.0 17 70 4 10 −1.0 20 80 5 None 24 95

Table 2: Raw Hatchability results after in ova vaccination at incubation day 18 with a dose of 0.05 ml/egg of IB vaccine to Groups 1-5 (25 eggs/group)

Group EID 50 # hatched (day 20)

# hatched (day 21)

# hatched(day 22)

total # hatched

1 10 2.0 — 12 5 17 2 10 1.0 — 10 11 21 3 10 0.0 — 12 5 17 4 10 −1.0 — 17 3 20 5 none — 24 — 24

Table 3: Calculated Hatchability results after in ovo vaccination at incubation day 18 with a dose of 0.05 ml/egg of IB vaccine to Groups 1-5 (28-30 eggs/group).

Group EID 50 # hatched (total # eggs) % hatched 1 10 2.0 25 (29) 86 2 10 1.0 26 (28) 93 3 10 0.0 25 (28) 89 4 10 −1.0 27 (30) 90 5 none 24 (28) 86


175

Table 4: Raw Hatchability results after in ovo vaccination at incubation day 18 with a dose of 0.05 ml/egg of IB vaccine to Groups 1-5 (28-30 eggs/group).

Group EID 50 # hatched(day 20) # hatched(day 21) # hatched total # 1 10 2.0 - --- (day 22) hatched 2 10 1.0 — 22 3 25 3 10 0.0 2 24 — 26 4 10 −1.0 18 7 — 25

Table 5: Clinical Signs In Non-Challenged Chicks post-hatch (PH) mortality prior to challenge

# days PH Post hatch

Group 1 (n = 25)

Group 2 (n = 26)

Group 3 (n = 25)

Group 4 (n = 27)

Group 5 (n = 24)

1 1 died 1 BC (bad codition)

5 1 died (yolk sac)

7 2 BC 9 1 died 1 killed # live chicks 23 26 25 27 22

(day 21 PH)

Table 6: Calculated Results Of Post-challenge Protection Protection against challenge at 3 weeks of age with virulent IB virus as determined by CST

Group (EID 50 ) # chicks protection % 1(10 2.0 ) 23 100 2(10 1.0 ) 25 100 3(10 0.0 ) 25 96 4(10 −1.0 ) 27 89 5(none) 22 0

Table 7: Raw Results Of Post-Challenge Protection, post-challenge assessments of protection with CST techniques Group(EID 50 ) # chicks +(full CST motion) ±(impaired motion)CST −(no CST motion 1(10 2.0 ) 23 23 0 0 2(10 1.0 ) 26 25 0 0 3(10 0.0 ) 25 23 2 0 4(10 −1.0 ) 27 23 2 2 5(none) 22 0 0 22


176

Table 8:Calculated serological results ,GMT against M41 antigen at 3 weeks of age using the HI test of serum samples.

( EID 50 ) # chicks 4.5 23 4.0 22 4.1 24 4.0 24 3.8 22

Table 9 : Raw Serological Results # chicks with indicated 2 log HI titers to IB M41 antigen at 3 weeks of age as determined from serum Groups Age of chicks (week)

Numbers (EID 50) 3 4 5 6 7 Group 1 (10 2.0 ) 8 2 9 2 2 Group) 2 (10 1.0 ) 11 3 5 3 4 Group 3 (10 0.0 ) 8 7 7 2 3 Group 4 (10 −1.0 ) 9 7 7 1 2 Group 5 (none) 11 5 5 1 1


177

EFFICACY OF COMBINED VACCINES AT DAY OF HATCH AGAINST A D388 CHALLENGE IN SPF AND COMMERCIAL CHICKENS

DE WIT JJ and VAN DE SANDE H

GD (Animal Health Service), P.O. Box 9, 7400 AA Deventer, the Netherlands SUMMARY Strains of infectious bronchitis virus (IBV) serotype D388 (genotype QX) were first detected in the Netherlands in December 2003 and have shown to be a major cause of respiratory problems and nephritis in broilers and other young chickens and of poor egg production in breeders and layers in the Netherlands and surrounding countries. In this study, the efficacy was determined of combining vaccines at day of hatch against an early challenge with IBV D388 of SPF and commercial birds. The efficacy was high in the SPF birds, in contrast to the results in the commercial birds. INTRODUCTION Since December 2003, a new serotype of IBV (D388), genotype QX has been detected in the Netherlands and surrounding countries. It has been involved in respiratory disease, nephritis and drops in egg production and egg quality. Part of flocks that were infected at very young age showed false layers at later age. Experiments by GD (Animal Health Service, Deventer, The Netherlands) showed that a). D388 infections in young birds caused serious damage in respiratory tract, kidney and oviduct, b). Mass vaccines or 793B-like vaccines at day of hatch provided only moderate protection against D388 (genotype QX), and c). Mass vaccines (incl. combi) at day of hatch and 793B-like vaccine at 14 days of age (D14) provided a reasonable to high protection at 4 weeks of age. This broad protection of the vaccination schedule using a Mass strain at day of hatch and a 793B-like vaccine at D14 has been reported for many strains (Cook et al., 1999) and also specifically for an Italian QX strain (Terregino et al., 2008) Two experiments were performed in SPF birds and commercial broilers to study the efficacy of combinations of several Mass vaccines and a 793B-like vaccine on the day of hatch against a challenge with D388 at 14 and 28 days of age.


178

MATERIALS and METHODS Vaccinations and serology Seven groups of 1-day-old SPF layer pullets and 10 groups of 1-day-old commercial broilers were housed in negative pressure isolators and vaccinated according to tables 1 and 2. The hatching eggs were obtained from a healthy broiler breeder flock with “normal” levels of neutralizing antibody levels against the strains that were included in the vaccines. All vaccines were applied by eye/nose drop (1 dose per bird). Challenge All birds were challenged by eye/nose-drop (104 EID50 D388 in 0,1 mL per bird) at 14 or 28 days of life. Determination of protection The level of protection in the trachea was determined using the ciliostasis test on 5 trachea rings per bird. The tracheas are placed in HMEM medium immediately after electrocuting of the chickens. Subsequently, 5 rings (equally divided over the total length of the trachea) are cut and placed in medium and placed at 37ºC. The level of ciliostasis is determined independently by two technicians between 1 and 4 hours after electrocution. The level of beating of the cilia per ring was expressed as 0 (0% beating of cilia), 1 (> 0 - 25% still beats), 2 (> 25 - 50% still beats), 3 (> 50 - 75% beats) or 4 (> 75 - 100% of the cilia beats). One bird can score between 0 and 20 (5 rings x score 4). The level of protection in the kidney was determined using the immuneperoxidase test. RESULTS SPF birds Seven (out of 200) birds showed severe respiratory distress after the vaccination resulting in a very low body weight at 14 days of life (table 3). Six of these birds had been vaccinated with the combination of vaccines. Four of these birds (group D en E, table 4) also showed complete ciliostasis at 14 days of life. Almost all other birds selected for the ciliostasis test at day of challenge showed no or very low level of ciliostasis. Groups E and G (combined vaccines at day of hatch) showed a high level of protection against ciliostasis and infection of the kidney after challenge at 14 and 28 days, respectively. The results of these groups were comparable to that of the group F (Mass/D274 at day of hatch) and 793B (D14). Application of Mass/D274 alone (group D) resulted in a protection level of 49% in the trachea. Fifty percent of the birds of group D had detectable replication of D388 in the kidney at 8 d.p.c. Commercial broilers The hatching eggs were obtained from a healthy broiler breeder flock with “normal” levels of neutralizing antibody levels against the strains that were included in the vaccines. The mean VNT antibody titres (log2) to serotypes M41, D274, 793B and D388 were 11.5, 9.5, 10, and 9.5 respectively. Non of the broilers showed severe respiratory distress after the vaccination. The vaccines that were combined with a


179

793B-like vaccine at day of hatch included a Mass/D274 combination, a Mass vaccine and the H120 vaccine. At 14 days of live, several of the birds of the 3 groups with the combination of vaccines at day of hatch, still had a certain level of ciliostasis. All 4 vaccinated groups (D, E, G and I) that were challenged at 14 days of life, showed an average tracheal protection level between 34 and 40%. The 3 groups (E, G and I) that had been vaccinated with the combined vaccines did not show more protection in the trachea than group D. All 3 vaccinated groups (F, H and J) that were challenged at 28 days of life, showed a tracheal protection level between 21 and 71%, which was lower than has been achieved by the separate application of the same vaccines (day of hatch and D14) in earlier experiments. The groups that were vaccinated at day of hatch with the combination of vaccines showed a level of protection against detectable replication in the kidney that was between 60% and 90% (different days post challenge) at 14 and 28 days of life DISCUSSION Combining 2 IBV vaccines resulted in severe respiratory distress in 6 (out of 100) SPF birds. The efficacy of this combination at DAY OF HATCH at tracheal and kidney level was high at 14 and 28 d.p.i. and comparable to the protection induced by the separate application of the vaccines at DAY OF HATCH and D14. However, the induced levels of protection of the same combination and 2 other combinations applied at DAY OF HATCH in commercial broilers with MDA against the vaccine strains were considerately lower than in the SPF birds. The results were also considerately lower than seen in earlier experiments when the vaccines were applied separately. The same kind of results have been reported before (Cook et al., 1999). These experiments showed that a separate application of the Mass vaccine and 793B-like vaccine is to be preferred to induce a relevant level of protection against challenge with D388/QX strain. For an early protection against the challenge which can result in nephritis and false layers, a high level of maternally derived virus neutralizing antibodies remains very important (De Wit et al., 2006). ACKNOWLEDGEMENTS This research was supported by a grant from the National Board for Poultry and Eggs (PPE) of the Netherlands.


180

REFERENCES Cook, J. K. A., Orbell, S. J., Woods, M. A. & Huggins, M. B. (1999) Breadth of

protection of the respiratory tract provided by different live-attenuated infectious bronchitis vaccines against challenge with infectious bronchitis viruses of heterologous serotypes. Avian Pathology, 28, 477-485.

De Wit, J. J., Nieuwenhuizen, J. & Fabri, T. H. F. (2006) Protection by maternally derived antibodies and vaccination at day of hatch against early challenge with IBV serotype D388. In: H.-R. a. Kaleta (Ed). Proceedings of the V. International symposium on Corona- and pneumovirus infections (314-318). Rauischholzhausen, Germany.

Terregino, C., Toffan, A., Beato, M. S., De Nardi, R., Vascellari, M., Meini, A., Ortali, G., Mancin, M. & Capua, I. (2008) Pathogenicity of a QX strain of infectious bronchitis virus in specific pathogen free and commercial broiler chickens, and evaluation of protection induced by a vaccination programme based on the Ma5 and 4/91 serotypes. Avian Pathol, 37, 487-493.

VI.

INT.

SYM

PO

SIU

M O

N A

VIA

N C

OR

ON

A-

AN

D P

NE

UM

OV

IRU

SE

S, R

AU

ISC

HH

OLZ

HA

US

EN, G

ER

MA

NY,

200

9

181

Tabl

e 1.

Stu

dy d

esig

n va

ccin

atio

n-ch

alle

nge

trial

of S

PF

bird

s ha

ndlin

g Va

ccin

atio

n (o

c-i.n

., 1

dose

) D

388

chal

leng

e 10

0 - %

cili

osta

sis

num

ber o

f bird

s Pe

rcen

tage

neg

ativ

e in

de

IPT

kidn

ey,

num

ber o

f bird

s G

rou p

(S

PF)

Day

0

Day

14

day

0 d.

p.c.

5

d.p.

c.

0 d.

p.c.

5

d.p.

c.

8 d.

p.c.

11

d.p

.c.

A

- -

14

n=4

n=10

n=

4 n=

10

n=10

n=

10

B

- -

28

n=6

n=10

n=

6 n=

10

n=10

n=

10

C

- -

14 n

eg a

ll / 2

8 ne

g al

l n=

5 / n

=5

n=5

/ n=5

n=

5 / n

=5

n=5

/ n=5

n=

5 / n

=5

n=5

/ n=5

D

M

ass/

D27

4*

14

n=

5 n=

10

n=5

n=10

n=

10

n=10

E

M

ass/

D27

4 +

793B

14

n=5

n=10

n=

5 n=

10

n=10

n=

10

F M

ass/

D27

4 79

3B

28

n=5

n=10

n=

5 n=

10

n=10

n=

10

G

Mas

s/D

274

+ 79

3B

28

n=

3 n=

10

n=3

n=10

n=

10

n=10

* 1

dos

e of

an

com

mer

cial

ly a

vaila

ble

Inac

tivat

ed IB

vac

cine

with

M41

and

D27

4 an

tigen

Ta

ble

2. S

tudy

des

ign

vacc

inat

ion-

chal

leng

e tri

al o

f com

mer

cial

bro

ilers

ha

ndlin

g Va

ccin

atio

n (o

c-i.n

., 1

dose

) D

388

chal

leng

e 10

0 - %

cili

osta

sis

num

ber o

f bird

s Pe

rcen

tage

neg

ativ

e in

de

IPT

kidn

ey

Gro

u p

(bro

ilers

) D

ay 0

da

y 0

d.p.

c.

5 d.

p.c.

0

d.p.

c.

5 d.

p.c.

8

d.p.

c.

11 d

.p.c

. A

-

14

n=5

n=10

n=

5 n=

10

n=10

n=

10

B

- 28

n=

3 n=

10

n=3

n=10

n=

10

n=10

C

-

14 n

eg a

ll /2

8 ne

g al

l n=

5 / n

=5

n=5

/ n=5

n=

5 / n

=5

n=5

/ n=5

n=

5 / n

=5

n=5

/ n=5

D

M

ass/

D27

4*

14

n=5

n=10

n=

5 n=

10

n=10

n=

10

E

Mas

s/D

274

+ 79

3B

14

n=4

n=10

n=

4 n=

10

n=10

n=

10

F M

ass/

D27

4 +

793B

28

n=

4 n=

10

n=4

n=10

n=

10

n=10

G

M

ass

+ 79

3B

14

n=5

n=10

n=

5 n=

10

n=10

n=

10

H

Mas

s +

793B

28

n=

5 n=

10

n=5

n=10

n=

10

n=10

I

H12

0 +

793B

14

n=

5 n=

10

n=5

n=10

n=

10

n=10

J

H12

0 +

793B

28

n=

5 n=

10

n=5

n=10

n=

10

n=10

* 1

dos

e of

an

com

mer

cial

ly a

vaila

ble

Inac

tivat

ed IB

vac

cine

with

M41

and

D27

4 an

tigen


VI.

INT.

SYM

PO

SIU

M O

N A

VIA

N C

OR

ON

A-

AN

D P

NE

UM

OV

IRU

SE

S, R

AU

ISC

HH

OLZ

HA

US

EN, G

ER

MA

NY,

200

9

182

Tabl

e 3.

Num

ber o

f sev

ere

resp

irato

ry re

actio

ns a

fter t

he v

acci

natio

n of

the

SPF

bird

s ha

ndlin

g Va

ccin

atio

n (o

c-i.n

., 1

dose

) D

388

chal

leng

e G

rou p

(S

PF)

Day

0

Day

14

day

Num

ber o

f bird

s w

ith v

ery

low

BW

at 1

4 da

ys b

y cl

inic

al re

actio

n on

vac

cina

tion

A

- -

14

0 B

-

- 28

0

C

- -

14 n

eg a

ll / 2

8 ne

g al

l 0

D

Mas

s/D

274

14

1

E

Mas

s/D

274

+ 79

3B

14

3

(3x

kidn

ey IB

pos

) F

Mas

s/D

274

793B

28

0

G

Mas

s/D

274

+ 79

3B

28

3

Tabl

e 4.

ha

ndlin

g Va

ccin

atio

n

(oc-

i.n.,

1 do

se)

D38

8 ch

alle

nge

100

- % c

ilios

tasi

s Pe

rcen

tage

neg

ativ

e in

de

IPT

kidn

ey,

num

ber o

f bird

s G

rou p

(S

PF)

Day

0

Day

14

day

0 d.

p.c.

5

d.p.

c.

0 d.

p.c.

5

d.p.

c.

8 d.

p.c.

11

d.p

.c.

A

- -

14

100

0 10

0 60

20

/ 55

* 85

B

- -

28

100

0 10

0 44

50

/ 62

90

C

- -

14 n

eg a

ll / 2

8 ne

g al

l 99

/ 99

10

0 / 9

3 10

0 / 1

00

100

/ 100

10

0 / 1

00

100

/ 100

D

Mas

s/D

274

14

4x

100

/ 1x

0

49

100

90

50 /

73

80

E

Mas

s/D

274

793B

14

2x 1

00 /

3x 0

95

40

10

0 90

/ 93

90

F M

ass/

D27

4 79

3B

28

89

94

100

100

80 /

93

100

G

Mas

s/D

274

793B

28

100

94

100

100

100

/ 100

10

0

* Ave

rage

per

cent

age

of 5

, 8 a

nd 1

1 da

ys p

ost c

halle

nge


VI.

INT.

SYM

PO

SIU

M O

N A

VIA

N C

OR

ON

A-

AN

D P

NE

UM

OV

IRU

SE

S, R

AU

ISC

HH

OLZ

HA

US

EN, G

ER

MA

NY,

200

9

182

Tabl

e 5.

ha

ndlin

g Va

ccin

ati o

n (o

c-i.n

., 1

dose

) D

388

chal

leng

e 10

0 - %

cili

osta

sis

nu

mbe

r of b

irds

Perc

enta

ge n

egat

ive

in d

e IP

T ki

dney

G

roup

(b

roile

rs)

Day

0

day

0 d.

p.c.

5

d.p.

c.

0 d.

p.c.

5

d.p.

c.

8 d.

p.c.

11

d.p

.c.

A

- 14

10

0 0

100

70

10 /

30

10

B

- 28

10

0 0

100

55

10 /

33

35

C

- 14

neg

all

/ 28

neg

all

100

100

/ 100

10

0 10

0 10

0 / 1

00

100

D

Mas

s /D

274

14

100

34

100

90

35 /5

7 45

E

M

ass /

D27

4 +

793B

14

86

40

10

0 80

80

/ 73

60

F

Mas

s /D

274

+ 79

3B

28

81

71

100

100

100

/ 93

90

G

Mas

s +

793B

14

35

35

10

0 90

10

0 / 9

3 10

0

H

Mas

s +

793B

28

99

21

80

10

0 70

/ 88

95

I

H12

0 +

793B

14

84

39

10

0 90

80

/ 83

90

J

H12

0 +

793B

28

98

47

10

0 10

0 80

/ 83

80



183

IMMUNE RESPONSES IN CHICKS AFTER SINGLE OR DUAL VACCINATION WITH LIVE INFECTIOUS BRONCHITIS MASSACHUSETTS AND VARIANT

VACCINES: SOME PRELIMINARY FINDINGS

GANAPATHY1 K, ROTHWELL

2 L, LEMIERE

3 S, KAISER

2 P and JONES

1 RC

1Department of Veterinary Pathology, University of Liverpool,

Leahurst Campus, Neston, South Wirral, CH64 7TE, UNITED KINGDOM 2Institute for Animal Health, Compton, Berkshire RG20 7NN, United Kingdom

3Merial SAS, 13b Avenue Albert Einstein, 69100 Villeurbanne, France

SUMMARY Infectious bronchitis (IB) is an economically important viral disease in chickens, mainly affecting the respiratory, urinary and reproductive systems. For the prevention of the disease, live and inactivated vaccines have been used for many decades but due to the ability of the virus to mutate or to form recombinants, occasionally novel variant viruses can penetrate through the protection conferred by existing vaccines. To overcome this, it has been shown that when chicks are vaccinated with two different serotypes, there is a broader protection against a range of unrelated IBV challenge viruses compare to homologous vaccines. Despite the widespread use of such vaccination regimes, the underlying immune mechanisms involved are not known. This paper reports on some preliminary findings on humoral and cell-mediated immune responses of chicks vaccinated first with live Massachusetts (H120) vaccine then followed by a live 793B (CR88) vaccine. In addition, protection conferred against virulent IBV M41, 793B, QX, Italy-02 and D1466 were assessed using an in vitro tracheal organ culture challenge model. INTRODUCTION Infectious bronchitis (IB) caused by a coronavirus, is arguably the most important respiratory viral disease in chickens in regions where avian influenza and Newcastle disease are absent. It mainly affects the respiratory, urinary and reproductive systems causing substantial economic losses (Cavanagh and Gelb, 2008; Dhinakar Raj and Jones, 1997). For the prevention of the disease, live and inactivated vaccines have been used for many decades but the disease persists through out the world. The challenge posed


184

by IB is mainly related to the ability of the virus to mutate or to form recombinants (Cavanagh et al., 1992; Kottier et al., 1995; Wang et al., 1997), which occasionally breaks-through the protection conferred by existing vaccines. With decreased host ability to clear an infection, virulent viruses tend to persist in the flocks, and occasionally variant strains may emerge. In recent years, variant genotypes such as 793B, Italy-02 and QX have been reported in Europe (Worthington et al., 2008). For control of IB, it has been shown that when chicks are vaccinated with two different serotypes/genotypes, there is a broader protection against various unrelated IBV challenge viruses compare to use of homologous serotype of live vaccines (Cook et al., 1999; Terregino et al., 2008; Worthington et al., 2008). Despite the broad protection, the underlying immune mechanisms are not known. Thus, the objective of this study is to examine the immune responses of chicks vaccinated with Massachusetts (H120) at day old followed by a 793B (CR88) at 13 days old. MATERIAL AND METHODS Vaccine preparation IBV H120 and CR88 live vaccines were provided by Merial Animal Health Limited (Europe). One vial of each vaccine was thoroughly mixed with 100 ml of SW (Sterile water). Chicks Day-old Rhode Island Red specific pathogen free chicks from Institute of Animal Health (Compton) were used. Experimental design Day-old chicks were randomly divided into 6 groups as shown in the Table 1. They were inoculated with 0.1 ml of SW or prepared vaccines according to schedule outlined in Table 1. Each chick was inoculated by ocular (0.05 ml) and nasal (0.05 ml) routes. Clinical signs Chicks were monitored daily for clinical signs. Sampling A number of samples were collected including for virus detection (RT-PCR and isolation), serology, cytokine detections, immunohistochemistry and in vitro tracheal protection. Some of the related samplings are outlined below. Sera Blood samples were collected at 1, 6, 13, 19 and 26 days old. Sera were used for routine ELISA and HI antibody assays. For ELISA, a commercial ELISA kit (BioChek, Gouda, Holland) was used and data were analysed as recommended by the manufacturer. For HI, serum antibody levels against M41, 793B, D1466, QX and Italy-02 were determined using 4HA units of IB antigens. The IBV HI antigens were kindly provided by Ruth Manvell (Veterinary Laboratory Agency, Weybridge, UK).


185

Tissues: At 6, 13, 19 and 26 days old, 5 chicks from vaccinated and 3 chicks from unvaccinated groups were sacrificed. Samples of trachea, lung, kidneys, caecal tonsil and spleen were removed and immediately placed in RNA later solution. These were stored at – 20 °C until further use. Tissues were processed and various cytokines were determined using quantitative real-time RT-PCR (Powell et al., 2009). This was carried out at the Institute of Animal Health, Compton. Tracheal Organ Culture (TOC) and in-vitro challenge At 26 days old, tracheas were harvested from five vaccinated and 3 unvaccinated chicks. After cleaning off excess fat, TOC were prepared as described before (Cook et al., 1976). After overnight incubation at 37 oC in a rotating rack, rings with 100% ciliary movement were used for in vitro protection studies. After separation into groups, the rings were inoculated with 0.1 ml of virulent M41, 793B, D1466, It-02 or QX IB viruses. The rings were read daily and scored for ciliostasis. Number of days taken for 50% of the rings to become non-viable (ndTOCNV50%) was obtained after plotting graphs. RESULTS Following the first vaccination at day-old, mild respiratory signs were observed only in the group given H120 and CR88 simultaneously. However, the signs disappeared before 13 days old. No clinical signs were observed in any other groups. Humoral antibody titres varied depending on the assay used. With IBV-specific ELISA, highest antibody titres were detected in the group that received heterologous vaccines apart, H120 at day old and CR88 at 13 days old. Interestingly, second highest levels of antibodies were detected in the group that received H120 vaccine at day-old and again at 13 days old. The group of chicks that had been given CR88 vaccine at 13 days old showed moderate levels of antibodies. In contrast, HI antibodies against M41 and 793B of all the vaccinated groups showed high and ‘clustered’ levels. However, the levels were markedly higher against M41 compare to the 793B antigen. The levels of HI against It-02, QX and D1466 were variable, ranging from high against It-02 but only trace levels against QX and D1466. For cell-mediated immune responses, the levels of mRNA in various tissues were detected in samples of trachea, kidney and spleen. The results of tracheal analysis showed a storm of IFN-γ, IL-1 and IL-6 after primary but not after the second vaccination. For IL-13, cytokine storm was only found in the group given H120 and CR88 simultaneously. No IL-10 was detected. Results of other tissues are currently being analysed. In vitro TOC challenge showed that the ndTOCNV50% was the highest among the homologous challenges, particularly against M41 and 793B. For heterologous challenges, the ndTOCNV50% differed depending on the vaccination group and challenge virus used. Broadly, ndTOCNV50% for the group given H120 at day-old followed by CR88 at day 13 was the highest.


186

DISCUSSION In this study, for the first time, it was attempted to evaluate the underlying immunological mechanisms for broader protection conferred by chicks vaccinated with heterologous compare to homologous IBV vaccines. The heterologous vaccines (H120 and CR88) were administered apart (H120 at day old and CR88 at 13 days old) or simultaneously at day old, and other groups were included as vaccinated and unvaccinated controls for comparison. Based on clinical signs observed, it appears that it is unwise to administer the H120 and CR88 vaccines simultaneously at day old, as post-vaccination reactions were seen until 13 days old. If such clinical signs can develop in SPF chicks kept under experimental condition, it is likely that much more severe respiratory disease may develop under the field condition. Humoral antibodies were assayed for ELISA and HI antibodies. In both assays, the level of antibodies in the chicks that received H120 at day old and CR88 at 13 days old were the highest. This may have contributed to enhanced protection seen in this group. Humoral antibodies, even though not directly correlated with respiratory protection, play an important role in limiting the spread of the virus to visceral tissues (Yachida et al., 1985) . Cellular immune responses were monitored by assaying for various cytokines in the spleen, trachea, lungs and caecal tonsil. To date, IFN, IL-1B, IL-6, IL-10 and IL-13 were determined in the trachea. It appears that following the first vaccination, there were cytokine storms at day 6 and 13 days old. For IL-13, cytokine storms were not seen. For IL-10, only trace amount were detected in few of the chicks. Further work is in progress to establish the significance of these findings. This study highlights kinetics of underlying humoral and cell-mediated immune responses following the heterologous live IBV vaccination. Broadly, it further strengthens the wider use of H120 followed by 793B vaccine. Table 1: Experimental groups

Vaccination (days) Group Number of chicks 1 13

1 35 H120 H120 2 35 NONE CR88 3 35 H120 CR88 4 35 H120 NONE 5 35 H120&CR88 NONE 6 21 NONE NONE


187

REFERENCES Cavanagh, D., Davis, P.J., Cook, J.K.A., Li, D., Kant, A. and Koch, G., 1992.

Infectious bronchitis virus: evidence for recombination within the Massachusetts serotype. Avian Pathol 21, 33-43.

Cavanagh, D. and Gelb, J., Jr., 2008. Infectious Bronchitis. In Disease of Poultry. Saif et al. (Eds). Pp 117 - 135. Blackwell Publishing Professional, Ames, IA.

Cook, J.K., Darbyshire, J.H. and Peters, R.W., 1976. The use of chicken tracheal organ cultures for the isolation and assay of avian infectious bronchitis virus. Arch Virol 50, 109-18.

Cook, J.K.A., Orbell, S.J., Woods, M.A. and Huggins, M.B., 1999. Breadth of protection of the respiratory tract provided by different live-attenuated infectious bronchitis vaccines against challenge with infectious bronchitis viruses of heterologous serotypes. Avian Pathol 28, 477-485.

Dhinakar Raj, G. and Jones, R.C., 1996. Protectotypic differentiation of avian infectious bronchitis viruses using an in vitro challenge model. Vet Microbiol 53, 239-52.

Dhinakar Raj, G. and Jones, R.C., 1997. Infectious bronchitis virus: immunopathogenesis of infectious in chicken. Avian Pathology 26, 667-706.

Gomez, L. and Raggi, L.G., 1974. Local immunity to avian infectious bronchitis in tracheal organ culture. Avian Dis 18, 346-68.

Kottier, S.A., Cavanagh, D. and Britton, P., 1995. Experimental evidence of recombination in coronavirus infectious bronchitis virus. Virology 213, 569-80.

Powell, F.L., Rothwell, L., Clarkson, M.J. and Kaiser, P., 2009. The turkey, compared to the chicken, fails to mount an effective early immune response to Histomonas meleagridis in the gut. Parasite Immunol 31, 312-27.

Terregino, C., Toffan, A., Beato, M.S., De Nardi, R., Vascellari, M., Meini, A., Ortali, G., Mancin, M. and Capua, I., 2008. Pathogenicity of a QX strain of infectious bronchitis virus in specific pathogen free and commercial broiler chickens, and evaluation of protection induced by a vaccination programme based on the Ma5 and 4/91 serotypes. Avian Pathol 37, 487-93.

Wang, L., Xu, Y. and Collisson, E.W., 1997. Experimental confirmation of recombination upstream of the S1 hypervariable region of infectious bronchitis virus. Virus Res 49, 139-45.

Worthington, K.J., Currie, R.J. and Jones, R.C., 2008. A reverse transcriptase-polymerase chain reaction survey of infectious bronchitis virus genotypes in Western Europe from 2002 to 2006. Avian Pathol 37, 247-57.

Worthington, K.J. and Jones, R.C., 2006. New genotype of infectious bronchitis virus in chickens in Scotland. Vet Rec 159, 291-2.

Yachida, S., Sawaguchi, K., Aoyama, S., Iritani, Y. and Hayashi, Y., 1985. Appearance of haemagglutinability of infectious bronchitis virus following in vitro and in vivo tracheal passages. Zentralbl Veterinarmed B 32, 736-43.


188

PREDICTIVE VALUE OF THE RESULTS OF AN α-IBV IgM ELISA FOR THE EFFICACY OF IBV VACCINATIONS IN THE FIELD

DE WIT JJ, VAN DER MEULEN A and FABRI THF

Animal Health Service (GD), P.O. Box 9, 7400 AA Deventer, the Netherlands SUMMARY Infectious bronchitis virus (IBV) is, in spite of vaccination(s), a major cause of respiratory problems in broilers and of poor egg production in breeders and layers in the Netherlands. A possible cause of failure of the protection induced by vaccination(s) is an inadequate application of the vaccine. The availability of a simple and cheap method to check or estimate the efficacy of an IBV vaccination would be an asset to the poultry industry. In this field study, the efficacy of IBV vaccinations was compared with the results of the α-IBV IgM ELISA on sera that were collected at several intervals post vaccination of the same broilers. The predictive value of the IgM ELISA was determined for the efficacy of the vaccination. The results that groups which showed at least 50% positive sera at 10 days post vaccination in the IgM ELISA had a protection of at least 89% against challenge. Groups of broilers with a low level of IgM positives showed an average protection of 43% with a range from 0% to 85%. These results show that the IgM ELISA can be used as an indicator of the efficacy of the IBV vaccination. INTRODUCTION Infectious bronchitis virus (IBV) is, in spite of vaccination(s), a major cause of respiratory problems in broilers and of poor egg production in breeders and layers in the Netherlands. Possible reasons of failure of the protection induced by vaccination(s) are e.g. a.) heterologous challenges, b.) immunesuppression, c.) very short or long interval between vaccination and challenge and d.) inadequate vaccine application. The mass application of IBV vaccines in the field is known for its many variations in a.) application technique (eyedrop, spray, water, atomist), b.) water (quantity, quality, temperature), c.) dosage, and d.) combination of different vaccines (e.g. IBV with Newcastle Disease vaccines). Many of these factors can have a negative effect on the efficacy of the vaccination. Therefore, a simple and cheap method to check the vaccination would be an asset to the poultry industry.


189

To determine the protection induced by the vaccination in the field by challenge experiment is not practical. If a vaccination induces a humeral response, serology can be used for checking the vaccination by detection of a seroconversion using paired serum samples. This strategy is hampered for vaccinations in presence of maternally derived antibodies (MDA). An alternative strategy for serodiagnosing an IBV vaccination would be to look for the presence of an IBV-specific IgM response. In contrast to the two other immuno-globulin classes, IgG and IgA, that are produced in response to IBV vaccination and infection, IBV-specific IgM response is short-lived (Gillette, 1974; Mockett & Cook, 1986; Martins et al., 1990, 1991; Toro et al, 1994; De Wit et al, 1998) Detection of IgM is therefore indicative of a recent infection or vaccination. Results of a IgM-capture ELISA, specific for IgM directed against IBV, were reported in Avian Pathology (De Wit et al, 1998). The specificity of the -IBV-IgM ELISA was 99.0%, the sensitivity based on an experimental vaccination (H120) of 9-week-old SPF chickens was 83 to 100%, depending on the days post vaccination. The IgM responses were rapid (first IgM after 3 to 5 days) and transient (about 2 weeks) and therefore indicative for an acute IBV infection or vaccination. A field study (De Wit, 1998) showed that about half of 80 broiler flocks did not show a detectable IgM response after IBV vaccination, which might indicate that part of the vaccinations might not have been successful. In this study, the correlation between the efficacy of IBV vaccinations and the IgM response after vaccination was determined in 12 groups of broilers originating from 6 broiler flocks. The IgM response after vaccination of 129 flocks was also determined. MATERIALS and METHODS IgM ELISA The ELISA was performed as described by De Wit et al. (1998). Briefly, a monoclonal antibody specific to chicken IgM was used as catching antibody and coated onto microtitre plates. Subsequent steps included the addition of test serum, IBV-antigen or control-antigen, enzyme labelled anti-IBV monoclonal antibody, and enzyme substrate. Vaccinations From 6 broiler flocks that were going to be (for the first time) vaccinated with a Mass (containing) vaccine at about 14 days of age, 10 birds per flock were transported to GD and housed in negative pressure isolators. These birds were vaccinated with one dose of the same vaccine as the other birds of that flock at 14 days of age by eye-drop. From each of the flocks, 10 birds that had been vaccinated at the farm (spray or water application), were transported to GD and housed in isolators. Blood samples were taken at day of vaccination, 7, 10 and 14 d.p.v. and stored in -20°C until testing. Challenge All birds were challenged between 14 and 20 days after vaccination by eye-drop (104

EID50 M41 in 0,1 mL per bird).


190

Determination of protection The level of protection was determined using the ciliostasis test on 5 trachea rings per bird. The tracheas are placed in HMEM medium immediately after electrocuting of the chickens. Subsequently, 5 rings (equally divided over the total length of the trachea) are cut and placed in medium and placed at 37ºC. The level of ciliostasis is determined independently by two technicians between 1 and 4 hours after electrocution. The level of beating of the cilia per ring was expressed as 0 (0% beating of cilia), 1 (> 0 - 25% still beats), 2 (> 25 - 50% still beats), 3 (> 50 - 75% beats) or 4 (> 75 - 100% of the cilia beats). One bird can score between 0 and 20 (5 rings time score 4). Field sera From 129 flocks (120 broiler, 6 pullets, 3 rearing broiler breeder flocks) at least 10 sera were collected between 7 and 14 days post IBV vaccination. The broiler flocks were all vaccinated between 14 and 19 days of age. One rearing flock was vaccinated at 11 days of age, the other flocks were vaccinated between 11 and 14 weeks of age. The variables included age (11 days to 14 weeks), vaccine (7 different IBV vaccines were used), dosage (0.5 to 2.1 dose per bird), use of stabiliser, way of application (eye-drop, spray, atomist or water), and combinations of vaccines (non, 2 IBV vaccines, one IBV and a ND vaccine, or 2 IBV vaccines and a ND vaccine). RESULTS The 6 groups of broilers that had been vaccinated in the isolator by eye-drop showed at least 50% positive sera at 10 d.p.v in the IgM ELISA and all groups showed a protection between 89% and 100% against challenge (Figure 1, ■). The 6 groups of broilers that had been vaccinated at the farm (Figure 1, ♦ ) showed on average 7% positives in the IgM ELISa and an average protection of 43% with a range from 0% to 85%. The results of the IgM ELISA in sera of the 129 flocks sampled between 7 and 14 days post vaccination are listed in figure 2. It showed that 44% of these flocks had less than 10% positives after vaccination. 26% of the flocks had at a IgM response between 50 and 100%. DISCUSSION The comparison of the IgM response post IBV vaccination and the efficacy against challenge in the 12 groups of broilers showed that groups which showed at least 50% positive sera at 10 days post vaccination in the IgM ELISA had a protection of at least 89% against challenge. Groups of broilers with a low level of IgM positives showed an average protection of 43% with a range from 0% to 85%. Although the absence of detectable amounts of IgM post vaccination does not prove the vaccination didn’t induce (local) protection, it is not considered to be a good sign. It could mean that the vaccination was not or less effective. The results of the 129 vaccinated flocks showed that 44% of these flocks had less than 10% positives after vaccination. The results of these 2 field studies suggest that the application of IBV vaccines in the field is less simple than many people think.


191

More important, the average efficacy of the IBV vaccinations could be improved considerably. The main question is however, how to achieve this. With the use of the IgM ELISA, it is relatively easy and cheap to compare and optimize different IBV mass application methods under field conditions. The IgM response of large groups of farms with different application methods (application method, application machines, dosage, amount of water, additives, etc) can be compared to find the critical factors that influence the efficacy of IBV vaccinations. This might result in a significantly higher average efficacy of IBV vaccinations under field conditions, what normally would result in less respiratory disease outbreaks, less secondary infections and condemnations, less need of antibiotics, more first quality eggs and progeny. REFERENCES Davelaar, F.G., Noordzij, A., & Donk, J.A. van der (1982). A study on the synthesis

and secretion of immunoglobulins by the Hardian gland of the fowl after eyedrop vaccination against infectious bronchitis at 1 day-old. Avian Pathology, 11: 63-79.

De Wit, J.J. (1998). Detection of IgM by ELISA after IBV vaccination. Proceedings of the International symposium on infectious bronchitis and pneumovirus infections in Poultry. Rauischholzhausen, Germany, pp 361-365.

De Wit, J.J., Mekkes, D.R., Koch, G. & Westenbrink, F. (1998). Detection of specific IgM antibodies to infectious bronchitis virus by an antibody-capture ELISA. Avian Pathology, 27, 155-160.

Gillette, K.G. (1974). Avian infectious bronchitis: demonstration of serum IgG and IgM neutralising antibody by sucrose density-gradient centrifugation and mercapto-ethanol reduction. Avian Diseases, 18: 515-525.

Martins, N.R. da Silva, Mockett, A.P.A., & Cook, J.K.A. (1990). A method for the rapid purification of serum IgM for the diagnosis of recent viral infections of chickens. Journal of Virological Methods, 29: 117-126.

Martins, N.R. da Silva, Mockett, A.P.A., Barrett, A.D.T., & Cook, J.K.A. (1991). IgM responses in chicken serum to live and inactivated infectious bronchitis virus vaccines. Avian Diseases, 35: 470-475.

Mockett, A.P.A., & Cook, J.K.A. (1986). The detection of specific IgM to infectious bronchitis virus in chicken serum using an ELISA. Avian Pathology, 15: 437-446.

Toro, H., Hidalgo, H., & Collingwood-Selby, A.N. (1994). Lachrymal antibody response of specific pathogen free chickens and chickens with maternal immunity after infectious bronchitis virus vaccination. Preventive Veterinary Medicine, 18: 267-274.


192

Figure 1.

Correlation between IgM respons at 10 days post IBV vaccination and protection against challenge (M41)

0

20

40

60

80

100

0 20 40 60 80 100Percentage of IgM IBV ELISA positive sera

Leve

l of p

rote

ctio

n

Figure 2.

Results IgM ELISA at 7-14 days post IBV vaccination (129 young flocks)

0

10

20

30

40

50

60

0-10 11-20 21-30 31-40 41-50 51-60 61-70 71-80 81-90 91-100

Percentage of IgM IBV positive samples per flock

num

ber o

f flo

cks


193

IMMUNE RESPONSE IN CHICKENS TO RECOMBINANT S1 AND N PROTEINS OF INFECTIOUS BRONCHITIS VIRUS

MEIR R 1, MAHARAT O 1, KRISPEL S 2 and KATZ E 3

1Division of Avian and Aquatic Animal Diseases, Kimron Veterinary Institute, P. O. Box 12, Bet Dagan, 50250 Israel

2 Migal, 10200 Kiryat Shmona, Israel 3 Institute of Medical Research, Faculty of Medicine, The Hebrew University of

Jerusalem, Israel

SUMMARY

Prevention of infectious bronchitis (IB) by live attenuated vaccines is of limited efficacy due to the frequent emergence of novel IB strains and variants. The viral proteins S1 and N, expressed in a prokaryotic system, (S1r and Nr) were tested for their ability to induce humoral and cell-mediated immune response in SPF chicks vaccinated by the intraocular and intranasal route. The proteins were tested alone and with the addition of the modified E. coli enterotoxin nLT, known to have mucosal adjuvant properties. In addition, the vaccinated chickens were tested for their resistance to challenge by a live virus. Vaccination by S1r or Nr alone induced cell-mediated responses but no detectable humoral response on day of challenge. The addition of LT improved IgA-specific antibody levels in tracheal lavage, but had no impact on systemic humoral or cell-mediated responses. S1r provided partial protection against the homologous strain, and Nr provided some protection only when chemically bound to nLT. Further investigation is required to find a better vector/adjuvant and optimize proportions of S1r and Nr to achieve satisfactory protection. INTRODUCTION The continuous changes in infectious bronchitis viruses (IBV) pose a great challenge to the vaccine industry. The efficacy of live attenuated vaccines is usually limited to diseases caused by homologous viruses and in most cases in vivo challenge trials are needed to determine the field strain protectotype (Gelb et al. 1991; Gelb et al. 2005). A possible solution to prevention of infectious bronchitis (IB) caused by novel IBV variants is the production of non-infective, viral protein-based vaccines, keeping in mind the production feasibility and ease of administration to the hatched chick.


194

The S1 and N proteins have been studied for their immunogenic properties; S1 is known to inducing humoral immune responses, particularely the production of neutralizing antibodies (Ignjatovic and Galli 1993; Moore et al. 1997), while the N protein is known to induce cytotoxic T cell (CTL) responses, critical for virus clearance from the infected bird (Collisson et al. 2000; Seo et al. 1997b). In this study we tested the ability to induce immune responses by the recombinant proteins S1r and Nr, of the M41 strain, expressed in a prokaryotic system. In addition, the modified E. coli enterotoxin LT (nLT) was tested for its ability to facilitate presentation of exogenous proteins via the cross-protection pathway (Basta and Alatery 2007), and as a mucosal adjuvant. Systemic and local IBV antibody levels, the cell mediated immune response, and the protection efficacy of S1r and Nr were examined. MATERIALS and METHODS Protein expression The genes encoding for the S1 and N proteins of the M41 strain, and the modified non-toxic enterotoxine nLT (Vasserman and Pitcovski 2006), were cloned in the expression vector pQE-30 and expressed in E. coli JM 109. The expressed proteins were identified in a Western blot assay, with anti poly-Histidine antibodies. Protection study SPF chicks were divided into seven vaccine groups, 16 chicks each, and placed in positive pressure Horsfal-Bauer isolators. The chicks were vaccinated by the intraocular and intranasal route at 1, 14, and 28 days of age, with the following proteins; S1r, a mix of S1r+nLT, Nr, a mix of Nr+nLT and chemically bound Nr+nLT. The positive control group was vaccinated twice by the intraocular route, at 1 and 14 days of age, with the live attenuated vaccine H120. The negative control was unvaccinated. At 35 days of age, all chicks were bled, six chicks from each group were sacrificed, the spleens were removed, and a tracheal lavage was sampled. The remaining chicks were challenged by the live M41 virus, by the intraocular route. Five days post challenge, tracheal swabs were samples from all birds, and tested directly for the presence of IBV using a real-time RT-PCR assay (unpublished). Humoral immune response The systemic humoral response was tested at day of challenge and 10 days PC using a commercial IBV ELISA kit (IDEXX), the HI assay, and an ELISA assay for the detection of M41 specific IgG and IgA antibodies. The local immune response was tested in tracheal lavages for M41 specific IgG and IgA antibodies. Cell-mediated immune response Splenocytes sampled at day of challenge were tested for their ability to secret IFN γ in response to in vitro stimulation by the inactivated M41 strain, using the ELISPOT assay (unpublished). All samples were tested in triplicates of 2X106 cells/well. Con A was used as positive control of cell viability, and cell-culture medium containing FCS, as negative control. Results were calculated as index of IFN-secreting cells, based on the net ratio of sample/positive-control counts.


195

RESULTS Protection study One week after the third protein administration, S1r provided 40% protection, compared to 75% and 10% in the positive and negative control groups, respectively. The addition of nLT lowered the protection efficacy significantly. Both Nr, and Nr+nLT mix vaccines did not protect, and the chemically bound Nr+nLT provided 30% protection. Humoral immune response Chicks vaccinated with S1r or a mix of S1r+nLT did not produce detectable local or systemic antibodies (Table 1). The addition of nLT to S1r increased IgA levels in the tracheal lavages, but did not affect systemic antibody levels. The Nr protein alone, and with the addition of nLT increased systemic IgA levels at day of challenge. Chicks vaccinated with Nr or the chemically bound Nr+nLT, had a secondary humoral response after challenge, similar to that of the positive control(results not shown). In tracheal lavages, Nr alone had no effect on IgG and IgA levels but the addition of nLT, both as a mix and in a chemical bond, increases IgA levels (Fig 1). Cell-mediated immune response The cell-mediated response was tested in two separated experiments. Both S1r and Nr induced a cell-mediated response in vitro as was demonstrated by the ELISPOT assay (Fig.2). When stimulated by the inactivated M41, splenocytes from chicks vaccinated with S1r or Nr had a secretion index of 89, and 200, respectively, compared to 56 and 86 in the positive controls. The addition of nLT reduced the cellular response significantly on both accounts. DISCUSSION The possibility of using recombinant viral proteins as an alternative to live attenuated vaccines has been previously reviewed (Jackwood 1999). While viral vectors such as fowlpox virus or baculovirus containing IBV structural proteins have been tested with different degrees of success (Song et al. 1998; Wang et al. 2002), the safety of commercial use of a live recombinant virus is still unknown. In this study we examined the option of vaccinating chicks with viral proteins synthesized by a high efficiency prokaryotic expression vector, with and without the addition of the non-infective, non-toxic mucosal adjuvant nLT. Vaccination with S1r provided partial protection against challenge with live M41, the homologous virus. The low level of protection could be attributed to the lack of systemic and local humoral responses to this protein, compared to that of the live vaccine. On the other hand, S1r was more efficacious than the live vaccine in inducing a cell-mediated response. Based on these findings it was deduced that S1r lacks B cell epitopes but has linear T cell epitopes involved in the immune response. Three such epitopes were identified previously by Ignjatovic. and Sapats as capable of inducing a delayed type hypersensitivity response (Ignjatovic and Sapats 2005). Vaccination with Nr did not provide any protection against challenge even though it induced a strong cell-mediated response in vitro, and a secondary systemic humoral response, indicating the existence of both B and T-cell linear epitopes on this protein.


196

The enterotoxin LT was tested for its potential to increase the immune response to the recombinant viral proteins, by adding the modified enterotoxin nLT as a mix or chemically bound to Nr. The known properties of nLT as a mucosal adjuvant were manifested by an increase of IgA levels in tracheal lavages of S1- and Nr-vaccinated chicks. Its impact on the cell-mediated response varied, depending on the preparation technique; when added in a mix with either one recombinant protein, nLT reduced significantly the number of IFN γ secreting splenocytes, but had a positive impact when chemically bound to Nr, as was demonstrated by the cell-mediated response and the partial protection. These findings indicate that other binding techniques should be tested, before a final conclusion can be drawn. In addition, other non-infective adjuvants should be examined. Although both S1r and Nr induced a specific immune response, the protection efficacy achieved by either protein was lower than that of the homologous live attenuated vaccine. To improve the degree of protection, the proteins should be tested in a dose response manner, separately and in association with each other. REFERENCES Basta, S. and Alatery, A., 2007. The cross-priming pathway: a portrait of an intricate

immune system. Scand J Immunol. 65, 311-319. Collisson, E.W., Pei, J., Dzielawa, J. and Seo, S.H., 2000. Cytotoxic T lymphocytes

are critical in the control of infectious bronchitis virus in poultry. Dev Comp Immunol. 24, 187-200.

Gelb, J., Jr., Wolff, J.B. and Moran, C.A., 1991. Variant serotypes of infectious bronchitis virus isolated from commercial layer and broiler chickens. Avian Dis. 35, 82-87.

Gelb, J.J., Weisman, Y., Ladman, B.S. and Meir, R., 2005. S1 gene characteristics and efficacy of vaccination against infectious bronchitis virus field isolates from the United States and Israel (1996-2000). Avian Pathol. 34, 194-203.

Ignjatovic, J. and Galli, L., 1993. Structural proteins of avian infectious bronchitis virus: role in immunity and protection. Adv Exp Med Biol. 342, 449-453.

Ignjatovic, J. and Sapats, S., 2005. Identification of previously unknown antigenic epitopes on the S and N proteins of avian infectious bronchitis virus. Arch Virol. 2, 2.

Jackwood, M.W., 1999. Current and future recombinant viral vaccines for poultry. Adv Vet Med. 41, 517-522.

Moore, K.M., Jackwood, M.W. and Hilt, D.A., 1997. Identification of amino acids involved in a serotype and neutralization specific epitope within the S1 subunit of avian infectious bronchitis virus. Arch Virol. 142, 2249-2256.

Seo, S.H., Wang, L., Smith, R. and Collisson, E.W., 1997b. The carboxyl-terminal 120-residue polypeptide of infectious bronchitis virus nucleocapsid induces cytotoxic T lymphocytes and protects chickens from acute infection. J Virol. 71, 7889-7894.

Song, C.S., Lee, Y.J., Lee, C.W., Sung, H.W., Kim, J.H., Mo, I.P., Izumiya, Y., Jang, H.K. and Mikami, T., 1998. Induction of protective immunity in chickens vaccinated with infectious bronchitis virus S1 glycoprotein expressed by a recombinant baculovirus. J Gen Virol. 79, 719-723.


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Vasserman, Y. and Pitcovski, J., 2006. Genetic detoxification and adjuvant-activity retention of Escherichia coli enterotoxin LT. Avian Pathol. 35, 134-140.

Wang, X., Schnitzlein, W.M., Tripathy, D.N., Girshick, T. and Khan, M.I., 2002. Construction and immunogenicity studies of recombinant fowl poxvirus containing the S1 gene of Massachusetts 41 strain of infectious bronchitis virus. Avian Dis. 46, 831-838.

Table 1. Systemic humoral response at 35 days of age (day of challenge). Vaccine ELISA IBV

(IDEXX) HI Abs IgG (άM41) IgA (άM41)

S1r - - - - S1r+nLT (mix) - - - - Nr - - - + Nr+nLT (mix) - - - + Nr+nLT (chemical bond) - - - + H120 - +/- + - neg. control - - - -

0,00

0,10

0,20

0,30

0,40

0,50

0,60

H120

S1r

S1r+nL

T Nr

Nr+nLT

*Nr+nL

T

unva

ccina

ted

Vaccine

OD

620

nm IgA

IgG

Figure 1. M41 antibody levels in tracheal lavages at day of challenge.

Figure 2. IFN γ secretion of in vitro stimulated splenocytes by inactivated M41. Nr+nLT* - chemically bound.

0

50

100

150

200

250

Vaccine

Inde

x IF

N se

cret

ing

cells

H120 S1r S1r+nLT Nr+nLT* unvaccinated

0

50

100

150

200

250

Vaccine

Ind

ex IF

N s

ecre

tin

g c

ells

H120 N N+LT unvaccinated


198

MANIPULATION OF THE INFECTIOUS BRONCHITIS CORONAVIRUS GENOME FOR VACCINE DEVELOPMENT

BRITTON P, ARMESTO M, CASAIS R and CAVANAGH D

Coronavirus Group, Institute for Animal Health, Compton, Newbury, Berkshire RG20 7NN, UK

SUMMARY Reverse genetic (infectious clone) systems have been developed for several coronaviruses and offer a rational way for manipulating the genome to produce defined live vaccine strains. We have developed a reverse genetics system based on infectious bronchitis virus (IBV), the single most economically costly infectious disease of domestic fowl in the UK. The IBV reverse genetics system has been used to swap the surface spike protein (S) gene, to determine whether the recombinant IBVs, expressing the heterologous S proteins can be used as vaccinal strains suitable for protection against challenge. In order to determine whether IBVs expressing heterologous S proteins can be used as vaccines we have replaced the S gene of our cloned non-pathogenic Beaudette strain with the S genes from either the pathogenic Massachusetts M41 or UK/4/91 strain. In chickens the spike-swapped recombinants had the same non-pathogenic phenotype as Beaudette but induced immunity to challenge with either M41 or UK/4/91 virus. Interestingly there was no cross protection against chickens challenged with M41 that had been previously vaccinated with the IBV BeauR-4/91(S), the rIBV expressing the 4/91 S protein. These results are promising for vaccine development by the gene-swapping approach and that genetic manipulation of the IBV genome has real potential for the production of a new generation of genetically stable vaccines. INTRODUCTION The coronavirus S protein is a type I glycoprotein, consisting of four domains: a signal sequence that is cleaved during synthesis; the ectodomain, which is present on the outside of the virion particle; the transmembrane region responsible for anchoring the S protein into the lipid bilayer of the virion particle; and the cytoplasmic tail, which oligomerises in the endoplasmic reticulum (Vennema et al., 1990) to form trimers (Delmas & Laude, 1990) which constitute the coronavirus virion spikes observable by electron microscopy. The S protein is responsible for binding to the


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target cell receptor and fusion of the viral and cellular membranes, fulfilling a major role in the infection of susceptible cells. In addition, neutralising antibodies raised against epitopes within the S protein are responsible for protecting infected animals against infection. The IBV S glycoprotein (1162 amino acids) is cleaved into two subunits, S1 (535 amino acids, 90-kDa) and S2 (627 amino acids, 84-kDa). The S1 subunit contains the receptor-binding activity of the S protein (Koch et al., 1990; Schultze et al., 1992). In addition to the transmembrane and C-terminal cytoplasmic tail domains, the S2 subunit ectodomain region contains a fusion peptide-like region (Luo & Weiss, 1998) and two heptad repeat regions involved in oligomerisation of the S protein (De Groot et al., 1987). The shorter heptad repeat is adjacent to the transmembrane region and consists of a leucine zipper motif (Britton, 1991). We have shown, using our IBV reverse genetics system (Armesto et al., 2008; Casais et al., 2001; Cavanagh et al., 2007), that replacement of the S protein of the avirulent Beaudette strain of IBV with the S protein from the virulent M41 strain (Britton et al., 2005; Casais et al., 2003) resulted in a rIBV that had the tropism of M41 in vitro but was still non-pathogenic in chickens (Hodgson et al., 2004). Vaccination of chicks with the rIBV induced protective immunity against challenge with M41. In order to further test the system we replaced the Beau-R S gene with the corresponding S gene from IBV UK/4/91 (793/B serotype) a pathogenic field isolate that does not belong to the same serotype, Massachusetts, as Beau-R and M41. Our results demonstrated that the resultant rIBV, BeauR-UK/4/91(S) was still non-pathogenic and was able to induce protective immunity against challenge with IBV UK/4/91 but not against M41, confirming that the S protein alone is able to offer a protective immune response as the two rIBVs were isogenic apart from the S protein. In addition, our results demonstrate that spike-swapping offers a way of generating a new generation of IBV vaccines. MATERIALS and METHODS Viruses The Beaudette-CK (Beau-CK; (Cavanagh et al., 1986)) strain of IBV is apathogenic in chickens resulting from multiple passages in embryonated eggs, the virus has been virus adapted for growth in chick kidney (CK) cell cultures and can be grown in Vero cells, an African green monkey cell line. Beau-R, a recombinant IBV produced from an infectious RNA transcribed from a full-length cDNA of Beau-CK (Casais et al., 2001) and the same properties as Beau-CK. The M41-CK strain of IBV is an isolate derived from M41 ((Darbyshire et al., 1979)) following adaption to growth on CK cells. The virus is pathogenic in chickens resulting symptoms associated with the disease infectious bronchitis. Although M41-CK can grow in CK cells it does not produce infectious virus in Vero cells. IBV UK/4/91 (also known as 793/B) was isolated in the UK in 1991 following an out break of IBV, and grows very poorly in CK cells, presumably from lack of adaption to these cells (Adzhar et al., 1997; Cavanagh et al., 2005). Recombination and rescue of rIBVs The rIBV BeauR-UK/4/91(S) was produced using our IBV reverse genetics system as previously described for the generation of BeauR-M41(S), the rIBV expressing the


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M41 S protein (Armesto et al., 2008; Britton et al., 2005; Casais et al., 2003). The overall strategy for swapping the Beaudette S gene for another S gene from a different strain is highlighted in Figure 1. Inoculation of chickens Groups of 1-week-old specific pathogen free Rhode Island Red chickens were housed in positive-pressure, HEPA-filtered isolation rooms with each group in a separate room. The chicks were inoculated with 3.0 log10 CD50 of BeauR-UK/4/91(S) virus via the conjunctival (eye-drop) and intranasal routes (Hodgson et al., 2004). Three groups of chicks were given a challenge inoculation in the same manner 21 days after the primary inoculation, either consisting of UK/4/91 or M41 virus. Mock-infected controls were inoculated with 0.1 ml of 0.2% BSA in PBS. Assessment of pathogenicity Chicks were monitored daily for clinical signs, in which the criteria used to measure pathogenicity were snicking, tracheal rales, wheezing, nasal discharge, watery eyes and ciliary activity of the trachea following eye drop and nasal inoculation of the chicks (Hodgson et al., 2004). Tracheas were removed from randomly selected chickens from each group at 4 to 7 days post-inoculation for assessment of ciliary activity. RESULTS Assessment of pathogenicity Chicks infected with the rIBVs, BeauR-M41(S) and BeauR-UK/4/91(S), or the parental viruses Beau-R, M41 and UK/4/91 were assessed along with mock-infected chicks for clinical signs associated with an IBV infection. The tracheas were removed from randomly selected chicks 4 to 7 days post infection and assessed for ciliary activity. As can be seen from Figure 2, none of the chicks infected with either of the two recombinant viruses exhibited ciliostasis when compared to the virulent M41 and UK/4/91 strains of IBV. The tracheas taken from mock infected chicks and those infected with Beau-R did not show any ciliostasis. This observation demonstrated that replacement of the Beaudette S protein with that from two virulent strains of IBV did not result in Beaudette becoming more virulent. No clinical signs associated with IBV were observed in chicks infected with the two rIBVs or Beau-R in contrast to those infected with M41 or UK/4/91 which did show clinical signs. Assessment of protection Following inoculation of groups of chicks with the rIBVs at 21 days post vaccination they were challenged with either M41 or UK/4/91 wild type viruses. The chicks were monitored for clinical signs and tracheas removed from randomly selected chicks for assessment of ciliary activity. Chicks, previously infected with either M41 or UK/4/91 that had recovered from infection, were used as controls for assessment of protection by the rIBVs. Analysis of the tracheas for ciliary activity, Figures 3 (BeauR-M41(S)) and 4 (BeauR-UK/4/91(S)), showed that the rIBVs were able to protect against challenge with homologous virus (donor of the S gene), to a similar level as observed from chicks that had recovered from infection with either of the two wild type viruses, M41 and UK/4/91. Interestingly, rIBV BeauR-UK/4/91 did not protect the chicks from


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challenge with M41; this lack of cross-protection is similar to previous results where chicks were previously infected with UK/4/91 and subsequently challenged with M41 following recovery from infection with UK/4/91 (Cook et al., 1999). In addition no clinical signs were observed in the chicks that had been vaccinated with either of the rIBVs and challenged with homologous wild-type virus; Figure 5 shows, as an example, the results for the analysis of snicking in chicks previously vaccinated with BeauR-UK/4/91 and subsequently challenged with UK/4/91. DISCUSSION We had previously used our IBV reverse genetics system (Casais et al., 2001) to determine whether replacement of the Beaudette S gene in our molecular clone, Beau-R, with a corresponding S gene from a virulent stain of IBV (M41) would (1) alter the pathogenicity of Beaudette and (2) whether such a rIBV could have the potential to be a basic vaccinial strain. We generated such a rIBV, BeauR-M41(S), that had the same attenuated phenotype as Beau-R (Beaudette) but did acquire the tissue tropism associated with M41 (Casais et al., 2003). Furthermore, we were able to show that BeauR-M41(S) was able to protect chicks against challenge with homologous wild type M41, the donor strain for the S gene (Hodgson et al., 2004). We developed a quicker process for replacing the Beau-R S gene based on homologous recombination for modifying the Beau-R cDNA cloned in vaccinia virus (Britton et al., 2005). We have subsequently replaced the Beau-R S gene with that from a virulent field isolate of IBV, UK/4/91, generating the rIBV BeauR-UK/4/91(S) in order to test whether such spike swaps have the potential for generating new vaccines. To establish the principle that spike-swapped rIBVs have the potential for vaccine development, we inoculated chickens, by eye-drop and intranasally, and challenged them three weeks later using either the M41 or UK/4/91 strains. Using prevention of clinical signs, such as snicking, and retention of tracheal ciliary activity as measures of protection, Beau-R (Beaudette) did not induce protection against M41 or UK/4/91 whereas the appropriate spike-swapped rIBV or the appropriate wild type virus did do so. Our results demonstrated that spike-swapped rIBVs, belonging to two different serotypes, have the capability of inducing an effective protective immune response to challenge against homologous virus. The rIBV, BeauR-UK/4/91(S) was unable to generate an immune response that protected against challenge with the non-homologous M41 strain; this was an expected result as M41 and UK/4/91 belong to different serotypes and previous results have demonstrated that there is no cross-protection between these two serotypes. However, in our experiment the rIBVs were isogenic, apart from the different S genes. Therefore, protection in our hands could only have been through the S gene as all the other genes were identical; thus confirming that a protective immune response can be induced from the S gene alone. We had previously tested rIBVs Beau-R and BeauR-M41(S) for their potential as vaccines for 18-day-old embryos (Tarpey et al., 2006). The rIBVs were inoculated in ovo and the pathogenicity was assessed by observing the effect on hatch, plus clinical signs and effect on the tracheal ciliary activity post hatch. Neither of the two rIBVs strains reduced hatchability or caused nasal discharge, and caused minimal damage to the ciliated epithelium of the trachea. In contrast, a commercial vaccine drastically reduced the hatch (35%) and caused substantial clinical signs. In addition


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the rIBVs induced a serological response and BeauR-M41(S) gave a high level of protection of the embryos against virulent M41 challenge. These results are promising for the development of embryo-safe efficacious IBV vaccines for in ovo application. In summary, this work demonstrates the utility of the reverse genetics approach for preparing recombinant IBVs and has demonstrated that potential vaccines based on rIBVs of the Beaudette strain, are safe and effective for protecting chicks and for the in ovo vaccination of SPF embryos. Our results have demonstrated, the extensive antigenic and genomic variations of IBV can be addressed by spike-swapping. ACKNOWLEDGEMENTS This work was supported by the Department of Environment, Food and Rural Affairs (DEFRA) project codes OD0714 & OD0717, the Biotechnology and Biological Sciences Research Council (BBSRC) and Intervet. REFERENCES Adzhar, A., Gough, R. E., Haydon, D., Shaw, K., Britton, P. & Cavanagh, D. (1997).

Molecular analysis of the 793/B serotype of infectious bronchitis virus in Great Britain. Avian Pathology 26, 625-640.

Armesto, M., Casais, R., Cavanagh, D. & Britton, P. (2008). Transient dominant selection for the modification and generation of recombinant infectious bronchitis coronaviruses. In SARS- and Other Coronaviruses: Laboratory Protocols, pp. 255-273. Edited by D. Cavanagh: Humana Press.

Britton, P. (1991). Coronavirus Motif. Nature 353, 394. Britton, P., Evans, S., Dove, B., Davies, M., Casais, R. & Cavanagh, D. (2005).

Generation of a recombinant avian coronavirus infectious bronchitis virus using transient dominant selection. Journal of Virological Methods 123, 203-211.

Casais, R., Dove, B., Cavanagh, D. & Britton, P. (2003). Recombinant avian infectious bronchitis virus expressing a heterologous spike gene demonstrates that the spike protein is a determinant of cell tropism. Journal of Virology 77, 9084-9089.

Casais, R., Thiel, V., Siddell, S. G., Cavanagh, D. & Britton, P. (2001). Reverse genetics system for the avian coronavirus infectious bronchitis virus. Journal of Virology 75, 12359-12369.

Cavanagh, D., Casais, R., Armesto, M., Hodgson, T., Izadkhasti, S., Davies, M., Lin, F., Tarpey, I. & Britton, P. (2007). Manipulation of the infectious bronchitis coronavirus genome for vaccine development and analysis of the accessory proteins. Vaccine 25, 5558-5562.

Cavanagh, D., Davis, P. J., Pappin, D. J. C., Binns, M. M., Boursnell, M. E. G. & Brown, T. D. K. (1986). Coronavirus IBV: partial amino terminal sequencing of spike polypeptide S2 identifies the sequence Arg-Arg-Phe-Arg-Arg at the cleavage site of the spike precursor propolypeptide of IBV strains Beaudette and M41. Virus Research 4, 133-143.

Cavanagh, D., Picault, J. P., Gough, R., Hess, M., Mawditt, K. & Britton, P. (2005). Variation in the spike protein of the 793/B type of infectious bronchitis virus, in


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the field and during alternate passage in chickens and embryonated eggs. Avian Pathology 34, 20-25.

Cook, J. K. A., Orbell, S. J., Woods, M. A. & Huggins, M. B. (1999). Breadth of protection of the respiratory tract provided by different live-attenuated infectious bronchitis vaccines against challenge with infectious bronchitis viruses of heterologous types. Avian Pathology 28, 471-479.

Darbyshire, J. H., Rowell, J. G., Cook, J. K. A. & Peters, R. W. (1979). Taxonomic studies on strains of avian infectious bronchitis virus using neutralisation tests in tracheal organ cultures. Archives of Virology 61, 227-238.

De Groot, R. J., Luytjes, W., Horzinek, M. C., Van der Zeijst, B. A. M., Spaan, W. J. M. & Lenstra, J. A. (1987). Evidence for a coiled-coil structure in the spike proteins of coronaviruses. Journal of Molecular Biology 196, 963-966.

Delmas, B. & Laude, H. (1990). Assembly of coronavirus spike protein into trimers and its role in epitope expression. Journal of Virology 64, 5367-5375.

Hodgson, T., Casais, R., Dove, B., Britton, P. & Cavanagh, D. (2004). Recombinant infectious bronchitis coronavirus Beaudette with the spike protein gene of the pathogenic M41 strain remains attenuated but induces protective immunity. Journal of Virology 78, 13804-13811.

Koch, G., Hartog, L., Kant, A. & van Roozelaar, D. J. (1990). Antigenic domains of the peplomer protein of avian infectious bronchitis virus: correlation with biological function. Journal of General Virology 71, 1929-1935.

Luo, Z. L. & Weiss, S. R. (1998). Roles in cell-to-cell fusion of two conserved hydrophobic regions in the murine coronavirus spike protein. Virology 244, 483-494.

Schultze, B., Cavanagh, D. & Herrler, G. (1992). Neuraminidase treatment of avian infectious bronchitis coronavirus reveals a hemagglutinating activity that is dependent on sialic acid-containing receptors on erythrocytes. Virology 189, 792-794.

Tarpey, I., Orbell, S. J., Britton, P., Casais, R., Hodgson, T., Lin, F., Hogan, E. & Cavanagh, D. (2006). Safety and efficacy of an infectious bronchitis virus used for chicken embryo vaccination. Vaccine 24, 6830-6838.

Vennema, H., Rottier, P. J., Heijnen, L., Godeke, G. J., Horzinek, M. C. & Spaan, W. J. (1990). Biosynthesis and function of the coronavirus spike protein. Advances in Experimental Medicine and Biology 276, 9-19.


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Receiver IBV cDNA in Vaccinia virus

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Figure 1. Schematic diagram outlining the principle of spike swapping. The Beau-R S gene within the vaccinia virus genome was deleted apart from the transmembrane and cytoplasmic tail. The ectodomain of the donor S gene, the complete S gene apart from the transmembrane and cytoplasmic domains, is inserted into the transfer/recombination vector (pGPTNEB193; (Armesto et al., 2008; Britton et al., 2005)) and inserted into the Beau-R cDNA replacing the S gene by homologous recombination using transient dominant selection (Britton et al., 2005). The Beau-R cDNA within the vaccinia virus genome with the new S gene sequence is used to generate rIBVs (Armesto et al., 2008; Britton et al., 2005; Casais et al., 2003; Casais et al., 2001).


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Figure 3. Comparison of the induction of protective immunity by IBV strains M41 (donor of the S protein gene), Beau-R (receiver of the S protein gene), and BeauR-M41(S) (Beau-R with the S protein gene of M41). Chicks were mock infected or vaccinated with the outlines at the top of the figure and challenged 21 days post vaccination with M41. Tracheas were removed six days post challenge from randomly selected chicks with in each group and assessed for ciliary activity. A high ciliary activity was indicative of good protection, whereas a low ciliary activity (a measure of ciliostasis indicative of virulent virus destroying the epithelial cells of the trachea) indicated lack of protection. The rIBV, BeauR-M41(S) induced protection against challenge with M41 to a similar level observed for chicks that had recovered from infection with M41. The Beau-R S protein was unable to protect against M41 even though the proteins only differ by 5%.


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Figure 4. Comparison of the induction of protective immunity by IBV strains UK/4/91 (donor of the S protein gene), Beau-R (receiver of the S protein gene), and BeauR-UK/4/91(S) (Beau-R with the S protein gene of UK/4/92). Chicks were mock infected or vaccinated with BeauR-UK/4/91challenged 21 days post vaccination with either UK/4/91 or M41. Tracheas were removed, at 4 to 6 days post challenge, from randomly selected chicks with in each group and assessed for ciliary activity. A high ciliary activity was indicative of good protection, whereas a low ciliary activity (a measure of ciliostasis indicative of virulent virus destroying the epithelial cells of the trachea) indicated lack of protection. The rIBV, BeauR-UK/4/91(S) induced protection against challenge with homologous UK/4/91 virus (donor of the S gene) but was unable to protect against challenge with M41.


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Figure 5. Comparison of the induction of protective immunity by IBV strains UK/4/91 (donor of the S protein gene), Beau-R (receiver of the S protein gene), and BeauR-UK/4/91(S) (Beau-R with the S protein gene of UK/4/92). Chicks were mock infected or vaccinated with BeauR-UK/4/91challenged 21 days post vaccination with either UK/4/91 or M41. Chicks were monitored 3 to 7 days post challenge for clinical signs associated with an IBV infection; in this example snicking. Only the non-vaccinated chicks challenged with UK/4/91 displayed higher rates of snicking and therefore clinical signs associated with IBV. The rIBV, BeauR-UK/4/91(S) induced protection against challenge with homologous UK/4/91 virus (donor of the S gene) and there appeared to be some attenuation of clinical signs associated following challenge with M41. Indicating that although there was no protection against ciliostasis with BeauR-UK/4/91(S) against challenge with M41 there was some attenuation of pathogenicity associated with clinical signs.


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MOLECULAR IDENTIFICATION AND CHARACTERIZATION OF A TURKEY CORONAVIRUS IN FRANCE

MAUREL S1, TOQUIN D1, QUEGUINER M1, LE MEN M1, ALLEE C1, LAMANDE J1, BERTIN J2, RAVILLION L3, RETAUX C2, TURBLIN V4, MORVAN H5,

PICAULT JP1and ETERRADOSSI N1

1 French Agency for Food Safety (AFSSA), Avian and Rabbit Virology Immunology and Parasitology Unit (VIPAC), BP 53, 22440 Ploufragan, France

2 Coopérative Le Gouessant, BP 228, 22400 Lamballe, France 3 Laboratoire BOCAVET, 79140 Cerizay, France

4 MC Vet Conseil, 72300 Sablé-sur-Sarthe, France 5 Laboratoire de Développement et d’analyse des Côtes d’Armor, BP54, 22440

Ploufragan, France

SUMMARY Poult enteritis complex (PEC) and Poult enteritis and mortality syndrome (PEMS) were first recognized as a major cause of losses among turkey poults in the USA in the early nineties. PEMS is an infectious, transmissible and multifactorial enteric disease affecting turkey poults up to 28 days of age. It is characterized by diarrhoea, anorexia, dehydration, marked growth depression and weight loss, immune dysfunction and acute mortality. A coronavirus was shown to be involved with turkey enteritis in the USA a couple of decades ago. Some strains of turkey coronavirus (TCoV) have been proposed, possibly with other bacterial or viral agents, to contribute to the etiology of PEMS. In addition to the USA, TCoV was reported as being prevalent in Brazil, Canada, Italy and the UK. In France since 2003, an increasing number of turkey flocks affected by enteric disorders clinically evocative of PEC or PEMS have been observed. To investigate the possible presence of TCoV in such flocks, we developed and validated a TaqMan-based real-time RT-PCR assay for the rapid detection and quantification of the avian coronavirus viral RNA in clinical samples. To avoid false negative results due to genetic variation, the RT-PCR primers were designed into conserved regions within the N gene of avian coronaviruses. Using this assay, a coronavirus was detected in the organs and/or intestinal contents from PEMS-suspected turkey flocks. To identify more precisely the detected coronavirus, we amplified the full length S gene (including its variable S1 part) from several isolates detected in different flocks, by using classical RT-PCR and either direct sequencing or cloning of the PCR products prior to sequencing. Sequence analyses allowed to


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identify a TCoV-related coronavirus. On the basis of the sequence of the S gene, the phylogenetic relationships of the French TCoV isolates with other avian coronaviruses were determined. To the best of the authors’ knowledge, these data represent the first full length sequence for the S gene of European TCoV isolates. INTRODUCTION Poult enteritis complex (PEC) includes several infectious intestinal disorders of young turkeys up to 7-10 weeks of age. Clinical signs include diarrhea, stunting, dehydration, anorexia, weight loss and immune dysfunction (Barnes et al. (2000)). When associated with mortality, this disease is designated as poult enteritis and mortality syndrome (PEMS). PEMS is an infectious, transmissible and multifactorial disease and a coronavirus was shown to be one of the causative agents involved in its etiology in the USA in 1991 (Guy (2003)). Turkey coronavirus (TCoV) belongs to the genus coronavirus in the family Coronaviridae, a group of enveloped and positive-stranded RNA viruses that infect a wide range of mammalian and avian species. Coronaviruses are divided into three groups (I, II and III) based on the genome structure and organization. Three of the virus structural proteins have been identified as the transmembrane glycoprotein (M), phosphorylated nucleocapsid (N) and spike glycoprotein (S). The spike protein contributes to the peplomers on the viral surface and contains neutralizing and group-specific epitopes that are the basis for antigenic variation. As a consequence, the spike protein is highly variable among different coronaviruses while the M and N proteins are more conserved between the coronavirus antigenic groups. Numerous techniques for the detection of TCoV based on immunohistochemistry (Breslin et al. (2000); Cardoso et al. (2008); Guy et al. (1997); Teixeira et al. (2007)), enzyme-linked immunosorbent assay (Gomaa et al. (2008a); Gomaa et al. (2009); Guy et al. (2002); Ismail et al. (2001); Loa et al. (2000)) and RT-PCR (Breslin et al. (2000); Circella et al. (2007); Ismail et al. (2001); Lin et al. (2002a); Lin et al. (2004); Loa et al. (2006); Sellers et al. (2004); Spackman et al. (2005); Teixeira et al. (2007); Velayudhan et al. (2003)) have been developed since a decade. It has been shown that close antigenic relationships exist between TCoV and infectious bronchitis virus (IBV) but they are different enough to separate them into the group 3 of coronaviruses (Akin et al. (2001); Breslin et al. (1999); Cavanagh et al. (2001); Gomaa et al. (2008b); Guy (2000); Guy et al. (1997); Lin et al. (2002a); Lin et al. (2004); Lin et al. (2002b); Loa et al. (2000)). So far, only North-American strains of TCoV have been extensively sequenced and characterized at the molecular level (Cao et al. (2008); Gomaa et al. (2008b)) whereas only short nucleotide sequences corresponding to the 3’ end of the S gene, the gene 3 and the 5’ end of gene 4 region and the gene 5 and the 5’ end of gene 6 region have been published for European TCoV strain from UK (Cavanagh et al. (2001)). Since 2003, an increasing number of turkey flocks exhibiting clinical signs compatible with PEC have been observed in France. To investigate the presence of TCoV in these flocks, we developed molecular methods. To avoid false negative results due to possible molecular variation between North-American and European strains, we


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first designed and validated an avian coronavirus-specific TaqMan-based real-time RT-PCR (AvCoV Q RT-PCR) assay for the rapid detection of viral RNA in clinical samples, based on conserved sequences in the N gene. Then we further characterized the coronavirus positive samples by cloning and sequencing the full length S gene of the detected viruses. Finally we identified more precisely the detected virus by comparing these sequences with GenBank sequences. MATERIALS and METHODS Viruses, clinical samples and RNA extraction The virus isolates used in the validation of the Av CoV Q RT-PCR are listed in Table 1 for avian coronaviruses and in Guionie et al. (2007) for the 21 viruses not belonging to the coronavirus genus. Clinical samples (duodenum, jejunum, cæcum, kidney, spleen, liver, bursa of Fabricius or/and thymus) from 59 turkey flocks suspected of PEC were received for virological investigation. The samples were ground and suspended w/v in PBS. The suspensions were centrifuged at 3500 rpm for 15 min and the supernatants were collected for RNA extraction using the QIAamp® Viral RNA Mini Kit (Qiagen), according to the manufacturer’s instructions. The purified RNA was eluted in 60 µl of AVE buffer. Development of the AvCoV Q RT-PCR The primers and Taqman probe were designed from a conserved region, in the middle of nucleocapsid gene of Turkey coronavirus and screened by NCBI nucleotide BLAST to exclude any cross reaction with cellular sequences or other pathogen targets (e.g. virus or bacteria). Primers and probe were synthesized by Applied Biosystems (Cheshire, UK). The one-step Taqman AvCoV Q RT-PCR assay was run using the QuantitectTM probe RT-PCR Kit (Qiagen). Briefly, each mix contained 1X (12.5 µl) of QuantitectTM RT-PCR master mix, 0.25 µl of Quantitect RT mix, 300 nM of each primer, 100 nM of probe, 5 µl of template RNA and RNase-free water to a final volume of 25 µl. cDNA was amplified using a Taqman 7000 thermocycler (Applied Biosystems) under the following cycling condition : 48°C for 30 min (reverse transcription), 95°C for 10 min (RT inactivation and activation of the HotStartTaq DNA polymerase); 40 cycles combining 95°C, 15 s (denaturation) and 60°C 1 min (annealing, extension step, and fluorescence data collection). Data were analysed with the SDS software version 1.3.1 (Applied Biosystems). To assess the analytical sensitivity of detection, the AvCoV Q RT-PCR assay was tested using 10-fold dilution series of RNA runoff transcripts corresponding to a fragment of the N gene. The transcripts were generated from a plasmid containing this fragment of a TCoV cloned downstream to the T7 promoter into the pcDNA 3.1Directional TOPO expression vector (Invitrogen). The pcDNA-TCoV constructs were linearized at the Eco RV (New England Biolabs) restriction site located downstream of the inserts. Runoff transcripts were synthesized using the T7 RNA polymerase (Promega) according to the standard protocol. Following the transcription reaction, the DNA templates were removed by digestion with the RQ1 RNase-free DNase (Promega). The lack of residual contaminating DNA was assessed by performing Q RT-PCR assay without the reverse transcriptase enzyme. The in vitro


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transcripts were extracted with phenol/chloroform, resuspended in nuclease-free water, aliquoted and stored at –70°C. They were quantitated by measuring the A260 with a spectrophotometer. The exact number of RNA molecules was calculated using the formula:

)/(10023.6159)5.320((

)/( 23 µlmoleculesYsnucleotideinlengthtranscript

RNAµlgX

S gene sequencing from samples positive in the AvCoV Q RT-PCR Three samples that proved positive in AvCoV Q RT-PCR and originated from different geographical locations were used in RT-PCR with primers defined in conserved regions flanking the S gene, upstream (in the 3’ end of 1b gene) and downstream (in the 5’ end of 3a gene), to amplify the CoV spike glycoprotein gene for sequencing. The PCR products were purified and cloned into pGEM®-T easy vector (Promega) and transformed into JM109 competent cells according to the manufacturer’s instructions. The recombinant plasmids were extracted with the QIAprep Spin miniprep kit (Qiagen) and screened by SacI digestion (Biolabs). At least 3 positive clones for each sample were sequenced in both directions using the Big Dye® terminator cycle kit and the Genetic Analyzer 3130 sytem according to the manufacturer’s recommendations (Applied biosystems). Due to the hypervariability of the S gene, we used the ‘gene walking’ method to sequence the S gene in its entirety. Sequence analysis The nucleotide-nucleotide BLASTn and protein-protein BLASTp search analysis were performed on-line at the National Center of Biotechnology Information (http://blast.ncbi.nlm.nih.gov/Blast.cgi). To examine the similarity of the different TCoV isolates with other coronaviruses, phylogenetic analysis were conducted based on the deduced amino acid sequences using ClustalW (MEGA software). Bootstrap was performed with 1000 replicates and trees were generated using the maximum likelihood (PhyML software), the maximum parsimony and neighbor-joining analysis (MEGA software). RESULTS Specificity of the AvCoV Q RT-PCR All the coronavirus-positive samples (Table 1) were detected in the AvCoV Q RT-PCR with a Ct lower than 30, whereas all the non-coronavirus virus controls (Guionie et al. (2007)) produced Ct higher than 40. The specifity of the AvCoV Q RT-PCR was thus validated with 15 strains of IBV, TCoV strain Indiana and 21 non-coronaviruses. Sensitivity of the AvCoV Q RT-PCR The detection and quantification limits were determined using Ct values obtained for each reaction containing from 5x109 to 5x102 copies of the standard RNA. The values were plotted against the log of the number of templates copies and a linear equation (Y= -3.47X + 47.1) with a R² value = 0.9986 was generated (Fig. 1). The assay maintained linearity for at least eight orders of magnitude from 1x109 copies per µl to

http://blast.ncbi.nlm.nih.gov/Blast.cgi


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1x102 copies per µl. Using the slope from the linear equation, the overall efficiency of the assay was estimated to be 94.13%. To exclude any false positive results, we decided to take in account only value less than 35 cycles threshold, corresponding to 2300 molecules of template RNA per µl of tested sample. Application to clinical samples of the AvCoV Q RT-PCR Out of 52 flocks presenting with signs compatible with PEC, 18 flocks were positive for avian coronavirus (34.6 %) whereas the samples from 7 flocks without enteritis were negative for avian coronavirus. The virus was found predominantly in cæcum (37.0 %) and jejunum (34.8 %) contents and bursa of Fabricius (32.6 %). These results are consistent with previous studies showing that TCoV is mainly found in the bursa of Fabricius, intestine and droppings (Breslin et al. (2000); Guy et al. (1997)). We selected three samples from geographically distant flocks with more than 5x107 viral RNA copies per µl for further identification of the detected coronavirus. Genetic analysis of the coronavirus isolates Based on the entire S sequence, the three isolates share 97.4 % of nucleotidic identity and 96.7 % of amino acid identity. The three isolates thus appeared very closely related but most nucleic acid mutations resulted in changes in the amino acid sequence and there were few synonymous mutations. BLASTn searches using the full nucleotide sequence of the S gene of the studied TCoV isolates resulted in alignment with TCoV isolates from North America references TCoV-ATCC, -Gh, -GI, -540 and -MG10 (Accession Numbers EU022526, AY342356, AY342357, EU022525 and EU095850, respectively). The similarity was limited to nucleotides 1588 to 3597 of the full S gene sequence, corresponding to the S2 region, and reached at most 75 % identity. Nucleotides at positions 1785-2010 of the S2 nucleotidic sequence shared 47 % identity with the more similar IBV strain (isolate AL/5364/00 n°EU359656). Amino acid sequence analysis using the full S protein with other group 3 coronaviruses showed a similarity close to 60 % with North American TCoV and 38 % with IBV (Table 2). We found significant alignments of the S1 French isolate amino acid sequence with Quail CoV (QCoV) from Italy (Circella et al. (2007)), but with a limited (42 %) similarity (Table 2). The S1 subunit shared 39 to 41 % similarity with TCoV from North America and only 19 % with IBV (Table 2). The S2 subunit shared 74 – 77 % identity with North American TCoV and approximately 52 % with the IBV strains tested (Table 2). The phylogenetic tree based on the full S glycoprotein is presented in Figure 2. CONCLUSION We have developed and validated a quantitative one-step RT-PCR specific for avian coronaviruses. We applied it on clinical samples from PEC-positive turkey flocks and found that 35 % of these flocks were positive for avian coronaviruses whereas seven turkey flocks without enteritis tested were negative for avian coronaviruses. As expected, the virus was found predominantly in the intestinal content and the bursa of Fabricius.


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The full length S gene of 3 isolates was cloned and sequenced. The region encoding the S2 subunit shared 74.1 - 77.3 % similarity with the North-American TCoV published sequences thus demonstrating the detected viruses share close genetic relationships with TCoV. The genetic region encoding the hypervariable S1 subunit proved very different from all sequences previously published and shared only 39.4 - 42.4 % amino acid similarity with the S1 sequences related to TCoV previously established from turkeys in North America (Cao et al. (2008); Gomaa et al. (2008b); Jackwood et al. (2004)) or from quail in Italy (Circella et al. (2007)). ACKNOWLEDGEMENTS The authors gratefully acknowledge the financial support of Conseil Régional de Bretagne, Conseil Régional des Pays de Loire, France Agrimer, and Comité Interprofessionnel de la Dinde Française (CIDEF) and the help of Pôle Agronomique de l’Ouest for project coordination. REFERENCES Akin A, Lin TL, Wu CC, Bryan TA, Hooper T, Schrader D. (2001) Nucleocapsid

protein gene sequence analysis reveals close genomic relationship between turkey coronavirus and avian infectious bronchitis virus. Acta Virol. 45(1): 31-8

Barnes HJ, Guy JS, Vaillancourt JP. (2000) Poult enteritis complex. Rev Sci Tech. 19(2): 565-88

Breslin JJ, Smith LG, Fuller FJ, Guy JS. (1999) Sequence analysis of the turkey coronavirus nucleocapsid protein gene and 3' untranslated region identifies the virus as a close relative of infectious bronchitis virus. Virus Res. 65(2): 187-93

Breslin JJ, Smith LG, Barnes HJ, Guy JS. (2000) Comparison of virus isolation, immunohistochemistry, and reverse transcriptase-polymerase chain reaction procedures for detection of turkey coronavirus. Avian Dis. 44(3): 624-31

Cao J, Wu CC, Lin TL. (2008) Complete nucleotide sequence of polyprotein gene 1 and genome organization of turkey coronavirus. Virus Res. 136(1-2): 43-9

Cardoso TC, Castanheira TL, Teixeira MC, Rosa AC, Hirata KY, Astolphi RD, Luvizotto MC. (2008) Validation of an immunohistochemistry assay to detect turkey coronavirus: a rapid and simple screening tool for limited resource settings. Poult Sci. 87(7): 1347-52

Cavanagh D, Mawditt K, Sharma M, Drury SE, Ainsworth HL, Britton P, Gough RE. (2001) Detection of a coronavirus from turkey poults in Europe genetically related to infectious bronchitis virus of chickens. Avian Pathol. 30(4): 355-68

Circella E, Camarda A, Martella V, Bruni G, Lavazza A, Buonavoglia C. (2007) Coronavirus associated with an enteric syndrome on a quail farm. Avian Pathol. 36(3): 251-8

Davelaar FG, Kouwenhoven B, Burger AG. (1984) Occurrence and significance of infectious bronchitis virus variant strains in egg and broiler production in the Netherlands. Vet Q. 6(3): 114-20


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Gomaa MH, Yoo D, Ojkic D, Barta JR. (2008a) Seroprevalence of turkey coronavirus in North American turkeys determined by a newly developed enzyme-linked immunosorbent assay based on recombinant antigen. Clin Vaccine Immunol. 15(12): 1839-44

Gomaa MH, Barta JR, Ojkic D, Yoo D. (2008b) Complete genomic sequence of turkey coronavirus. Virus Res. 135(2): 237-46

Gomaa MH, Yoo D, Ojkic D, Barta JR. (2009) Virus shedding and serum antibody responses during experimental turkey coronavirus infections in young turkey poults. Avian Pathol. 38(2): 181-6

Guionie O, Toquin D, Sellal E, Bouley S, Zwingelstein F, Allee C, Bougeard S, Lemiere S, Eterradossi N. (2007) Laboratory evaluation of a quantitative real-time reverse transcription PCR assay for the detection and identification of the four subgroups of avian metapneumovirus. J Virol Methods. 139(2): 150-8

Guy JS. (2000) Turkey coronavirus is more closely related to avian infectious bronchitis virus than to mammalian coronaviruses: a review. Avian Pathol. 29(3): 207-12

Guy JS. (2003) Turkey coronavirus enteritis. In: 11th Ed. Diseases of Poultry. (Iowa State Press) Ames, IA. Saif YM, Barnes HJ, Glisson JR, Fadly AM, McDougald LR & Swayne DE. pp. 300-307.

Guy JS, Barnes HJ, Smith LG, Breslin J. (1997) Antigenic characterization of a turkey coronavirus identified in poult enteritis- and mortality syndrome-affected turkeys. Avian Dis. 41(3): 583-90

Guy JS, Smith LG, Breslin JJ, Pakpinyo S. (2002) Development of a competitive enzyme-linked immunosorbent assay for detection of turkey coronavirus antibodies. Avian Dis. 46(2): 334-41

Ismail MM, Cho KO, Hasoksuz M, Saif LJ, Saif YM. (2001) Antigenic and genomic relatedness of turkey-origin coronaviruses, bovine coronaviruses, and infectious bronchitis virus of chickens. Avian Dis. 45(4): 978-84

Jackwood MW, Hilt DA, Boynton T, Callison SA. (2004) Molecular analysis of TCoV, SARS-CoV, and IBV: how are they related. 4th International Symposium on Avian Corona- and Pneumovirus Infections. Rauischholzhausen, Germany. pp. 158-165.

Jungherr EL, Chomiak TW, Luginbuhl RE. (1956) Immunological differences in strains of infectious bronchitis virus. 60th United States Livestock Sanitary Association. pp. 203-209.

Lin TL, Loa CC, Wu CC. (2002a) Existence of gene 5 indicates close genomic relationship of Turkey coronavirus to infectious bronchitis virus. Acta Virol. 46(2): 107-16

Lin TL, Loa CC, Wu CC. (2004) Complete sequences of 3' end coding region for structural protein genes of turkey coronavirus. Virus Res. 106(1): 61-70

Lin TL, Loa CC, Wu CC, Bryan T, Hooper T, Schrader D. (2002b) Antigenic relationship of turkey coronavirus isolates from different geographic locations in the United States. Avian Dis. 46(2): 466-72

Loa CC, Lin TL, Wu CC, Bryan TA, Hooper TA, Schrader DL. (2006) Differential detection of turkey coronavirus, infectious bronchitis virus, and bovine coronavirus by a multiplex polymerase chain reaction. J Virol Methods. 131(1): 86-91


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Loa CC, Lin TL, Wu CC, Bryan TA, Thacker HL, Hooper T, Schrader D. (2000) Detection of antibody to turkey coronavirus by antibody-capture enzyme-linked immunosorbent assay utilizing infectious bronchitis virus antigen. Avian Dis. 44(3): 498-506

Parsons D, Ellis MM, Cavanagh D, Cook JK. (1992) Characterisation of an infectious bronchitis virus isolated from vaccinated broiler breeder flocks. Vet Rec. 131(18): 408-11

Picault JP. (1987) Thesis: Contribution à l'étude des coronaviroses dans l'espèce gallus gallus en France. Université Claude Bernard / Ecole Nationale Vétérinaire de Lyon. Lyon, France.

Picault JP, Guittet M, Bennejean G, Drouin P, Lamandé J, Allée C. (1988) A pragmatic approach for the diagnosis of Infectious Bronchitis (IB) variant virus infections. 1st International Symposium On Infectious Bronchitis. Rauishholzhausen, Germany. pp. 229-241.

Picault JP, Drouin P, Guittet M, Bennejean G, Protais J, L'Hospitalier R, Gillet JP, Lamande J, Bachelier AL. (1986) Isolation, characterisation and preliminary cross-protection studies with a new pathogenic avian infectious bronchitis virus (strain PL-84084). Avian Pathol. 15(3): 367-83

Sellers HS, Koci MD, Linnemann E, Kelley LA, Schultz-Cherry S. (2004) Development of a multiplex reverse transcription-polymerase chain reaction diagnostic test specific for turkey astrovirus and coronavirus. Avian Dis. 48(3): 531-9

Spackman E, Kapczynski D, Sellers H. (2005) Multiplex real-time reverse transcription-polymerase chain reaction for the detection of three viruses associated with poult enteritis complex: turkey astrovirus, turkey coronavirus, and turkey reovirus. Avian Dis. 49(1): 86-91

Teixeira MC, Luvizotto MC, Ferrari HF, Mendes AR, da Silva SE, Cardoso TC. (2007) Detection of turkey coronavirus in commercial turkey poults in Brazil. Avian Pathol. 36(1): 29-33

Velayudhan BT, Shin HJ, Lopes VC, Hooper T, Halvorson DA, Nagaraja KV. (2003) A reverse transcriptase-polymerase chain reaction assay for the diagnosis of turkey coronavirus infection. J Vet Diagn Invest. 15(6): 592-6


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Table 1. List of the avian coronavirus strains used for the specificity test of the AvCoV Q RT-PCR. Pathogen virus AFSSA reference Reference or sourceInfectious Bronchitis Virus PL 84084/5.5 Picault et al. (1986)Infectious Bronchitis Virus strain Connecticut CR Conn/5.1 Jungherr et al. (1956)Infectious Bronchitis Virus CR 84221/6.1 Picault JP thesis (1987)Infectious Bronchitis Virus CR 84222/6.1 Picault JP thesis (1987)Infectious Bronchitis Virus CR 85131/13.1 Picault JP thesis (1987)Infectious Bronchitis Virus CR 88121/11.4 Picault et al. ( 1988)Infectious Bronchitis Virus CR 88061/8.3 Picault et al. (1988)Infectious Bronchitis Virus CR 4-91/3.1 Parsons et al. (1992)Infectious Bronchitis Virus (vaccine) CR B48/1.1 Cevac® Mass L (CEVA Santé animale, France)Infectious Bronchitis Virus (vaccine) CR D274b/6.3 Davelaar et al. (1984)Infectious Bronchitis Virus CR D212/56 Davelaar et al. (1984)Infectious Bronchitis Virus CR D3128/64 Davelaar et al. (1984)Infectious Bronchitis Virus CR D3896/52 Davelaar et al. (1984)Infectious Bronchitis Virus strain Beaudette VBBPa/3.8 Picault JP thesis (1987)Infectious Bronchitis Virus strain Beaudette VBBSa/2.2 Picault JP thesis (1987)Turkey Coronavirus strain Indiana TCoV INp4/1.1 Kind gift from Prof CC Wu & YM Saïf Table 2. Amino acid sequence identity of the entire S protein or subunits S1 / S2 of French TCoV isolate with other group 3 coronaviruses. Sequences were aligned using the ClustalW method.

n/a: not available


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Figure 1. Sensitivity of the AvCoV Q RT-PCR. In the left panel, the amplification plot of 10-fold serial dilutions of in vitro transcribed RNA is represented. It shows the evolution of the fluorescence (DeltaRn) of the 10-fold serial diluted RNA (A: 5x109

copies to H: 5x102 copies) in function of the cycle number. In the right panel, the corresponding standard curve is represented with the evolution of the cycle threshold (Ct) as a function of the log10 of the number of RNA copies. The slope and the correlation coefficient (R²) are indicated.

Figure 2. Representative phylogenetic tree of the full S glycoprotein for French isolate and TCoV strains Gh, Gl, 540, MG10 and ATCC, an IBV strain Beaudette and the SARS-CoV isolate civet014. Trees were generated from amino acid sequences using Neighbor-joining method with 1000 bootstraps replicates with the SARS-CoV as an outgroup. Accession numbers are indicated.


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SUBGENOMIC RNA TRANSCRIPTION OF TURKEY CORONAVIRUS

CAO JZ, WU CC and LIN TL

The present study was carried out to determine subgenomic mRNA (sgRNA) transcription of turkey coronavirus (TCoV) genome. Northern blotting and RT-PCR were used to examine the production of positive (+) and negative (-) sgRNA from total RNA that was isolated from TCoV-infected turkey small intestines or baby hamster kidney (BHK) cells. Northern blotting revealed five sgRNA bands hybridized with [α-32P]UTP-labeled RNA probe synthesized from TCoV nucleocapsid gene. The abundance of (+) and (-) sgRNA was found to be similar by both Northern blotting and RT-PCR when total RNA was isolated from the TCoV-infected intestines. However, the abundance of (+) sgRNA was much more than that of (-) sgRNA when total RNA was isolated from TCoV-infected BHK cells. Five sgRNAs were identified for spike gene of TCoV genome. These five sgRNAs differed at the transcription-regulating sequence. In addition, a novel sgRNA (sg7) was revealed in the predicted 3’ UTR region by RT-PCR. The sg7 encodes an open reading frame of 74 nucleotide residues and was identified from TCoV isolates ATCC, 540, and 310. Thus, a total of six sgRNAs was produced by TCoV. The sg7 was also detected from another avian coronavirus, infectious bronchitis virus (IBV) strains Ark99 and Connecticut, but not from M41, suggesting that the 3’UTR of IBV may be shorter than previously reported.


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COMPARATIVE GENOMICS ON AVIAN CORONAVIRUSES; ORIGIN AND DIVERGENCE ASSOCIATED WITH HOST AND PATHOGENIC SHIFTS

JACKWOOD1 MW, PATERSON2 AH, KISSINGER3 JC, D. HILT1 A, MCCALL1 A W, MCKINLEY1 ET and BOYNTON1 TO

1Department of Population Health, College of Veterinary Medicine, University of Georgia, 953 College Station Road, Athens, GA, 30602, USA,

2Department of Crop and Soil Science, College of Agricultural and Environmental Sciences, University of Georgia, 111 Riverbend Road, Athens GA 30602

3Department of Genetics, Center for Tropical and Emerging Global Diseases, University of Georgia, 500 D. W. Brooks Drive, Athens GA 30602

SUMMARY About 12 years ago, turkey coronavirus (TCoV) emerged as a new virus causing enteric disease in turkeys in North Carolina, USA. Nucleic acid sequence analysis indicated that different isolates of TCoV were genetically related and similar to infectious bronchitis virus. In this study, we report the complete genomic sequences for 4 strains of Turkey coronavirus (TCoV) that segregate into different phylogenetic groups. Phylogenetic analysis of the genomic sequences was used to reconstruct the changes associated with the emergence of TCoV as well as the divergence among the viruses. The genetic changes reported herein for this group III coronavirus correspond to a host as well as pathogenic shift and are directly related to a double recombination event surrounding the spike glycoprotein gene. The source of this genetic material remains unknown. INTRODUCTION Coronaviruses are worldwide in distribution, highly infectious, and extremely difficult to control because they have extensive genetic diversity, a short generation time and a high mutation rate. They can cause respiratory, enteric, and in some cases hepatic and neurological diseases in a wide variety of animals and humans. An enormous, previously unrecognized gene pool for coronaviruses exists among animals, which is actually not unlike the gene pool existing for influenza viruses in animals. Since coronaviruses have been shown, both experimentally and in nature, to undergo genetic recombination by a genomic template switching mechanism at a rate similar to that of influenza viruses, it is not surprising that host switching leading to new diseases have occurred among coronaviruses. As an example, the abrupt


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emergence of the severe acute respiratory syndrome coronavirus (SARS-CoV) in southern China in 2002 quickly spread to 27 countries (including the United States), and resulted in 8096 cases and 744 deaths (http://www.who.int/csr/sars/country/table2004_04_21/en/index.html). Phylogenetic analysis of recent sequence data indicates that a SARS-like bat coronavirus could be the progenitor of SARS-CoV (Li et al., 2005). Animals, like palm civet cats and raccoon dogs in which SARS-like coronaviruses have also been detected are commonly found in the market place in China and are likely intermediate hosts. Approximately 12 years ago, Guy et al. (1997) reported that a coronavirus isolate from North Carolina turkeys (NC95) was similar to infectious bronchitis virus (IBV). Their conclusions were based on cross reactivity of polyclonal antibodies in the fluorescent antibody test and immnoperoxidase procedures. In addition, they found that a monoclonal antibody directed against the membrane protein of IBV reacted with the NC95 isolate of TCoV. Since that report, several other studies conducted on TCoV isolates, showed that the order of the genes at the 3’ end of the genome was similar to IBV (Breslin et al., 1999; Cavanagh et al., 2001; Guy, 2000; Li et al., 2005; Stephensen et al., 1999; Velayudhan et al., 2003). However, our analysis of sequence data for the spike gene indicated that it was not related to IBV (GenBank Acc. # AY342356 and AY342357). In this study, we present the full genome sequences for 4 TCoVs and use them to reconstruct the changes associated with tissue tropism and host shifts, which lead to an emerged coronaviral disease in turkeys. MATERIALS AND METHODS Viruses Four isolates of TCoV designated TCoV/VA-74/03, TCoV/TX-GL/01, TCoV/IN-517/94, and TCoV/TX-1038/98 (Lin et al., 2002) were obtained from Mr. Tom Hoper (Purdue University, West Lafayette, IN, USA). The viruses were propagated in 1-day old specific pathogen free (SPF) turkeys and in 18 to 22 day old embryonating turkey eggs. Coronavirus genome amplification Viral RNA was purified with the High Pure RNA extraction kit (Roche Diagnostics Corporation, Indianapolis, IN) and used directly in the amplification reaction. The TaKaRa RNA LA PCR kit (Takara Bio Inc., Otsu, Shiga, Japan) was used in a strand displacement amplification reaction to randomly amplify the viral genomic RNA. Whole genome nucleotide sequencing and analysis Plasmid DNA from the libraries of cloned cDNA fragments for each virus were isolated using an alkaline lysis method modified for a 96-well format, and incorporating both Hydra and Tomtek robots (http://www.intl-pag.org/11/abstracts/P2c_P116_XI.html). Cycle sequencing reactions were performed using the BigDye Terminator® Cycle Sequencing Kit Version 3.1 (Applied Biosystems, Foster City, CA) and MJ Research (Watertown, MA) thermocyclers. Each viral genome was shotgun sequenced to approximately 10X coverage.

http://www.who.int/csr/sars/country/table2004_04_21/en/index.html

http://www.intl-


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Chromatogram files and trace data were read and assembled using SeqMan Pro, and genome annotation was conducted with SeqBuilder (Lasergene v. 8.0.2 DNASTAR, Inc., Madison, WI). Sequences generated in this study for TCoV and reference sequences TCoV/MG10 (EU095850) TCoV/540 (EU022525), TCoV/ATCC (EU022526), Peafowl/GD/KQ6/03 (AY641576), IBV/Beaudette (NC 001451), IBV/H120 (FJ888351), IBV/CA99 (AY514485), IBV/Arkp11 (EU418976), IBV/Arkp101 (EU418975), IBV/CK/CH/LSD/051 (EU637854), IBV/SAIBK (DQ288927), IBV/BJ (AY319651), Partridge/GD/S14/03 (AY646283), IBV/A2 (EU526388), beluga whale/CoV/SW1 (NC010646), MuniaCoV/HKU13 (FJ376622), ThrushCoV/HKU12 (FJ376621), BulbulsCoV/HKU11 (FJ376619) and IBV/Mass41 (AY851295) were aligned with ClustalV (DNAStar) and ClustalW (MEGA 4.0.2, Tamura et al., 2007). Phylogenetic trees (bootstrap test of phylogeny= 1000 replicates) were constructed with the Neighbor-Joining method, Minimum Evolution method, Maximum Parsimony method, and Unweighted Pair-Group Method with Arithmetic Mean (UPGMA) using MEGA 4.0.2. SimPlot version 1.3 was used to examine the genomes for recombination (Lole et al., 1999). RESULTS AND DISCUSSION Genome organization. The sizes of the genomes of TCoV/VA-74/03, TCoV/TX-GL/01, TCoV/TX-1038/98, and TCoV/IN-517/94 are 27,771, 27,619, 27,782, and 27,665 nucleotides respectively. The order of the genes was the same for all of the viruses examined and was; 5’ UTR-1a (1ab)-spike-3a-3b-envelope-membrane-ORF X-5a-5b-Nucleocapsid-3’ UTR. The genome organization, location and size of the open reading frames for each of the viruses were similar to previously published data for TCoVs (Cao et al., 2008; Gomaa et al., 2008; Lin et al., 2004). Phylogenetic analyses. Alignments were generated with both ClustalV and ClustalW and all of the phylogenetic trees generated with the methods described had similar topography. Although limited in number, when all of the available group 3 coronavirus full-length genomes were examined, three major clades were observed supporting recently published data indicating that the group 3 coronaviruses can be divided into three subgroups (Woo et al., 2009). Subgroup 3a containing TCoV and IBV isolates, subgroup 3b containing viruses isolated from a bulbul, a thrush, and a munia, and subgroup 3c containing a virus isolated from a beluga whale (SW1). Based on our analysis of spike sequence, we further divide the TCoV isolates into different genetic groups. Studies on the hypervariable regions of IBV spike have shown that different genetic groups can represent different serotypes of the virus (Lee et al., 2003). Phylogenetic analysis of the coding sequences for spike, membrane, RNA dependent RNA polymerase (RdRp), nucleocapsid, 3CLpro, and helicase for the TCoVs sequenced in this study as well as TCoV/MG10 (EU095850), TCoV/540 (EU022525), TCoV/ATCC (EU022526), and other representative group 3 coronaviruses including IBV/Arkp11 (EU418976), beluga whale/CoV/SW1 (NC010646), ThrushCoV/HKU12 (FJ376621), and IBV/Mass41 (AY851295), showed that for all of the genes, the beluga whale and thrush CoVs do not cluster with the other viruses or with each other. In addition, the TCoVs cluster with IBV/Mass41 and


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IBV/Arkp11 for all of the genes except spike. For spike, the IBV isolates clearly fall outside the cluster of TCoV viruses. Recombination events and molecular evolution. Recombination detection analysis (SimPlot http://sray.med.som.jhmi.edu /SCRoftware/ simplot/) on TCoV isolates, showed that recombination occurred in the 3’ end of gene 1ab, and in the 3’ end of spike, essentially inserting a different spike gene into TCoV. Similarities between spike glycoprotein sequences and the locations of the cross-over sites among TCoVs isolated over time, suggests that this was a one time event which lead to the emergence of TCoV from its closest IBV ancestor. Subsequently BLASTn and BLASTp (http://blast.ncbi.nlm.nih.gov/Blast.cgi) analyses using all and parts of the recombined region were conducted and no sequences in the GenBank or Swissprot databases matched the sequences. A codon-based test of positive selection (Z-test, MEGA4) was used to analyze the numbers of synonymous and nonsynonymous substitutions per site (dS/dN ratio) in the spike gene and it showed that positive selection was occurring between the TCoV/IN-517/94 and the TCoV/TX-1038/98 isolates, but not between more recent TCoV isolates. Similar to what was observed for the emergence and evolution of the SARS-CoV, a high rate of molecular evolution occurs when viruses switch hosts, which slows once adaptation to the new host occurs (Song et al., 2005). In conclusion, TCoVs emerged through a recombination event in which the spike gene of an unknown coronavirus or other virus replaced the spike gene of IBV. Replacing the spike gene of IBV resulted in both a pathogenicity change from an upper-respiratory disease to an enteric disease as well as a host shift from chickens to turkeys. Adaptation of TCoV to the new host (turkeys) resulted in additional mutations in the spike gene leading to viruses that segregate into different genetic groups. REFERENCES Breslin, J. J., Smith, L. G., Fuller, F. J. & Guy, J. S. (1999). Sequence analysis of the

turkey coronavirus nucleocapsid protein gene and 3' untranslated region identifies the virus as a close relative of infectious bronchitis virus. Virus Res 65, 187-193.

Cao, J., Wu, C. C. & Lin, T. L. (2008). Complete nucleotide sequence of polyprotein gene 1 and genome organization of turkey coronavirus. Virus Res 136, 43-49.

Cavanagh, D., Mauditt, K., Sharma, M., Drury, S. E., Ainsworth, H. L., Britton, P. & Gough, R. E. (2001). Detection of coronavirus from turkey poults in Europe genetically related to infectious bronchitis virus of chickens. Avian Pathoglogy 30, 355-368.

Gomaa, M. H., Barta, J. R., Ojkic, D. & Yoo, D. (2008). Complete genomic sequence of turkey coronavirus. Virus Res 135, 237-246.

Guy, J. S. (2000). Turkey coronavirus is more closely related to avian infectious bronchitis virus than to mammalian coronaviruses: a review. Avian Pathoglogy 29, 207-212.

Lee, C. W., Hilt, D. A. & Jackwood, M. W. (2003). Typing of field isolates of infectious bronchitis virus based on the sequence of the hypervariable region in the S1 gene. J Vet Diagn Invest 15, 344-348.

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Li, W., Shi, Z., Yu, M., Ren, W., Smith, C., Epstein, J. H., Wang, H., Crameri, G., Hu, Z., Zhang, H., Zhang, J., McEachern, J., Field, H., Daszak, P., Eaton, B. T., Zhang, S. & Wang, L. F. (2005). Bats are natural reservoirs of SARS-like coronaviruses. Science 310, 676-679.

Lin, T. L., Loa, C. C. & Wu, C. C. (2004). Complete sequences of 3' end coding region for structural protein genes of turkey coronavirus. Virus Res 106, 61-70.

Lin, T. L., Loa, C. C., Wu, C. C., Bryan, T., Hooper, T. & Schrader, D. (2002). Antigenic relationship of turkey coronavirus isolates from different geographic locations in the United States. Avian Dis 46, 466-472.

Lole, K. S., Bollinger, R. C., Paranjape, R. S., Gadkari, D., Kulkarni, S. S., Novak, N. G., Ingersoll, R., Sheppard, H. W. & Ray, S. C. (1999). Full-length human immunodeficiency virus type 1 genomes from subtype C-infected seroconverters in India, with evidence of intersubtype recombination. J Virol 73, 152-160.

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TURKEY CORONAVIRUS MOLECULAR BIOLOGY AND PATHOGENICITY OF A GROWTH-ROBBING

PATHOGEN

GOMAA1 MH, YOO2 D, OJKIC3 D and BARTA4 JR

1Department of Virology, Faculty of Veterinary Medicine, Kafrelsheikh University, Egypt; 2Department of Pathobiology University of Illinois at Urbana-Champaign,

Urbana, IL, USA; 3Animal Health Laboratory, 4Department of Pathobiology, University of Guelph, ON, Canada.

SUMMARY Turkey Coronavirus (TCoV) is an important enteric pathogen affecting commercial turkeys. Using an Ontario isolate, the complete genome sequence (27,657 bases) was determined and assigned as the reference sequence for TCoV (GenBank NC_010800). The genome organization of this virus was determined to be 5’-UTR-Pol-S-3a-3b-E-M-ORF-X-5a-5b-N-UTR-3’. Based on these findings, TCoV was assigned to group III coronaviruses. Two structural proteins (nucleocapsid and truncated spike glycoprotein) were expressed in E. coli and, after purification, used to develop two novel diagnostic enzyme linked immunosorbent assays (ELISAs) for serodiagnosis of TCoV. Serum samples from Ontario turkey farms tested using both recombinant protein ELISAs revealed high seroprevalence of TCoV. To study TCoV infection, an experimental trial was conducted in which day-old turkey poults Hummel et al. (1992) were inoculated orally with the same Ontario TCoV isolate. Birds developed clinical signs of enteritis such as depression, anorexia and obvious diarrhea as well as significantly decreased body weight gain in comparison to the uninfected controls. Virus shedding was monitored using TCoV-specific RT-PCR. TCoV shedding in the feces of infected birds started at 24 hours post infection, continued for a period of up to 17 days post-infection but ceased completely by 21 dpi. Following the antibody titers against TCoV using the recombinant TCoV antigen-based ELISAs revealed that seroconversion was detectable reliably at 10 14 days post-infection and that high titers of TCoV-specific antibody were present in the serum of all infected poults by 3 weeks post infection. Poults recovered from an infection acquired at day-of-age were refractory to re-infection by oral/intramuscular challenge at 21 days of age indicating that protective acquired immunity is elicited relatively quickly. TCoV is a widespread and serious pathogen affecting young turkey poults.


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INTRODUCTION Turkey coronavirus (TCoV) is a group III coronavirus closely related to infectious bronchitis virus (IBV) of chickens Gomaa et al. (2008a). TCoV is one of the most important causative agents of diarrhea in turkey poults. Infection of immunologically naïve poults with TCoV produces diarrhea and as a consequence negatively impacts their growth rates causing significant economic losses. Although TCoV was identified as the causative agent of ‘Bluecomb disease’ of turkey poults over 50 years ago Tumulin et al (1957), vaccines are not available to control the disease. Coronaviruses possess 4 major structural proteins: spike (S) glycoprotein, which consists of two subunits (S1 and S2), membrane (M) protein, small envelope (E) protein, and nucleocapsid (N) protein. The N protein is highly conserved among group III coronaviruses, and different strains of IBV and TCoV share 97% or greater identity at the amino acid sequence level Gomaa et al (2008a). During IBV infections in chickens, the IBV N protein is expressed at high levels and produces antibodies that react with a variety of IBV serotypes, Seah et al. (2000). The B cell epitopes have been mapped in the carboxyl terminal region of the N protein Boots et al., (1991), Seo et al., (1997). The N protein is also involved in the cell mediated immunity (CMI) and protection of chickens from IBV infection. As with IBV Lugovskaya e al., (2006), ELISA tests have been developed using recombinant N protein for various viruses such as measles virus Hummel et al., (1992), vesicular stomatitis virus Ahmed et al., (1993), Newcastle disease virus Erington et al., (1995) and severe acute respiratory syndrome coronavirus (SARS-CoV) Qiu et al. (2005). Recently, the full-length genomic sequence of TCoV was completed Gomaa et al. (2008a). and provided the genetic information necessary to produce recombinant antigens for development of diagnostic tests for TCoV in turkeys. The objective of this study was to develop and validate ELISA for TCoV based on N and S recombinant antigens. These ELISAs were subsequently applied to determine the seroprevalence of TCoV in breeder and meat turkey flocks in Ontario, Canada and Arkansas, USA. MATERIALS and METHODS Virus TCoV was isolated from an Ontario turkey suffering from acute enteritis and diarrhea. Intestines from affected birds were homogenated in PBS and then clarified by centrifugation at 4,000×g for 15 minutes. The supernatant was filtered through a 0.22μm-membrane filter (Millipore, Bedford, Mass.). This isolate was named TCoV MG10 Griener et al (1995) and used for this study. Complete genomic sequencing of TCoV-MG10 The complete genome sequence and the established virus organization had been done (see Gomaa et al (2008a). Cloning of the TCoV-N and TCoV-S54-395 protein genes Clonining and expression of TCoV-N and TCoV-S54-395 protein genes had been done according to (Gomaa et al (2008b, 2009b).


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ELISA The already developed two novel ELISAs described earlier have been used to conduct a seroprevelance of TCoV among North American turkey farms as discussed by (Gomaa et al (2008b, 2009b). Statistical analyses Differences in TCoV seroprevalence between breeder and meat turkey flocks were tested with one-tailed T-tests assuming unequal variances. In all cases, differences were considered significant at p<0.05. Spearman Rank Order Correlation calculated using the Free Statistics Software (38) was used to assess the correlation of OD405 values, and the seropositive or seronegative status determined by cut-off OD405 values established for the TCoV-N- and TCoV-S54-395–based ELISAs. The OD405 values from 360 paired field serum samples from Ontario commercial turkeys were included in this correlation analysis. Experimental turkeys Thirty (30) one day old turkey poults used in this study were wing tagged and divided into 2 separately housed groups, 10 infected and 20 control birds. Day-old birds were orally inoculated with 100 µL of clarified virus-containing supernatant. At 21 days of age, remaining poults in the infected group received a secondary challenge of TCoV-MG10 via oral gavage (100 µL of clarified TCoV MG10). At 28 days of age, eight birds from the control group were moved to the room housing the infected poults and orally inoculated with TCoV MG10 as described above for the day of age poults. RNA extraction, RT-PCR and PCR RNA extraction was carried out from the filtered cloacal swab suspensions using Qiagen RNA mini-easy Kits (Qiagen, Valencia, California, USA) according to the manufacturers’ instructions. Complementary DNA (cDNA) was generated with random hexamers and PCR with nucleocapsid gene-specific primers (NF1, 5?-AGGGAAATTTTGGTGATGAC-3?; NR1, 5?-ATGGGCGTCCTTGTGCTGTA-3?) was carried out as previously described (Gomaa et al., 2008a). PCR amplification products were separated by electrophoresis through 1% agarose, stained with ethidium bromide and visualized by ultraviolet light. Reactions were considered positive for TCoV if an amplification product of 330 bp was visible in the agarose gel. To confirm the identity of the amplicon as TCoV, randomly selected positive samples were purified using Qiagen PCR purification kits (Qiagen) and then sequenced using the NF1 and NR1 primers. The cDNA from each cloacal swab sample was also tested for turkey astrovirus using Astro-For (5?-AATAAGGTCTGCACAGGT-3?) and Astro-Rev (5?-TGGCGGCGAACTCCTCAACA-3?) primers (Sellers et al., 2004) as well as for avian reovirus using ARV S4 P4 (5?-GTGCGTGTTGGAGTTTC-3?) and ARV S4 P5 (5?-ACAAAGCCAGCCATRAT-3?) primers (Bruhn et al., 2005). Positive avian astrovirus (TAstV-2) and reovirus (S1133) controls were generously provided by Dr Stacey Schultz-Cherry (Southeast Poultry Research Laboratory, USDA-ARS, Athens, Georgia, USA). For all PCR reactions, both known positive and negative control samples were included for quality control.


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RESULTS Complete genomic sequence of turkey coronavirus The full-length genome of TCoV-MG10 was 27,592 nucleotides, excluding the 3’ poly(A) tail and approximately 42 nucleotides at the 5’ terminus. Two open reading frames, ORFs 1a and 1b, resided in the first two thirds of the genome, and 9 additional downstream ORFs were identified. A gene for hemagglutinin-esterase was not present in TCoV. The region between the membrane (M) and nucleocapsid (N) protein genes contained 3 potential small ORFs: ORF-X, a previously uncharacterized ORF with an associated putative TRS within the M gene (both apparently shared among all group III coronaviruses), and previously described ORFs 5a and 5b. The TCoV genome is organized as follows: 5’UTR-replicase (ORF1a and ORF1b)-spike (S) protein-ORF3 (ORFs 3a and 3b)- small envelop (E or 3c) protein- membrane (M) protein- ORF-X, ORF5 (ORFs 5a, and 5b) -nucleocapsid (N) protein-3’UTR-poly(A). TCoV genome structure and sequence was most similar, but distinct from, avian infectious bronchitis virus (IBV). This is the first complete genome sequence for a TCoV and confirms that TCoV belongs to group III coronaviruses see Gomaa et al., (2008a). Seroprevalence of Turkey Coronavirus in Turkeys in North America Established using Two Newly Developed Recombinant Antigen TCoV-Specific Antibody ELISAs Although TCoV had been identified more than 50 years ago but little is known about its prevalence in the field. To address this, nucleocapsid and truncated spike glycoproteins of TCoV carrying relevant B cell epitopes were cloned and expressed in E. coli. Expressed nucleocapsid protein produced two distinct proteins (52 and 43 KD); their specificity was confirmed by Western blot using two different monoclonal antibodies.Truncated S protein representing the S1 subunit region (amino acid positions 54-395) was likewise expressed and confirmed as a 30 KD protein using hyperimmune serum. Both proteins were purified and used as antigens to develop two ELISAs for TCoV serosurveillance that were then validated using experimentally derived turkey serum. Both recombinant ELISAs showed high sensitivity (97% and 95% for N- and S1-based, respectively) and specificity (93% and 92% for N and S1-based, respectively) for TCoV, which was significantly higher than an infectious bronchitis coronavirus (IBV)-based commercial test for TCoV. To assess the utility of these ELISA tests, 360 serum samples from turkey farms in Ontario and 81 serum samples from Arkansas were tested for TCoV-specific antibodies using the two recombinant TCoV ELISAs. High seroprevalence of TCoV was found on the Ontario farms with 73.89% or 71.11% of breeders and 60.00% or 56.67% of meat turkeys testing seropositive using the N- and S-based ELISAs, respectively. Similarly high field prevalence was found in Arkansas where 64.20% or 79.01% of the serum samples tested seropositive using the N- and S-based ELISAs, respectively. Virus Shedding and Serum Antibody Responses during experimental Turkey Coronavirus infections in young turkey poults The course of turkey coronavirus (TCoV) infection in young turkey poults was examined using a field isolate (TCoV-MG10) from a diarrheal disease outbreak on a commercial turkey farm in Ontario, Canada. Two day old and 28-day-old poults were


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inoculated orally with TCoV-MG10 to examine the effect of age on viral shedding and serum antibody responses to the virus. The presence of coronavirus particles measuring 105.8±21.8 nm in the cloacal contents was confirmed using transmission electron microscopy. The pattern of cloacal TCoV shedding was examined by RT PCR amplification of the nucleocapsid (N) gene fragment. TCoV serum antibody responses were assessed with two recently developed TCoV ELISAs that used TCoV N and S1 polypeptides as coating antigens. Poults were found equally susceptible to TCoV infection at 2 days of age and at 4 weeks of age and turkeys of either age shed virus in their feces starting as early as 1 day post-inoculation up to 17 days post-inoculation. Poults infected at 2 days of age were immunologically protected against subsequent challenge at 20 days post inoculation. The protection was associated with measurable serum antibody responses to both the nucleocapsid and S1 structural proteins of TCoV that were detectable as early as one week post-infection see Gomaa et al., (2009a). Infections with a pathogenic turkey coronavirus isolate negatively affect growth performance and intestinal morphology of young turkey poults Turkey coronavirus (TCoV) is an important viral pathogen causing diarrhea of young turkey poults that is associated with sizeable economic losses for the turkey industry. Using a field isolate that tested negative for turkey astrovirus and avian reovirus we were able to reproduce the clinical disease associated with TCoV. Clinical signs and weight gain of poults during experimental infections were compared with age matched uninfected controls. Poults infected at 2 days of age had 100% morbidity and 10% mortality and birds infected at 28 days of age showed 75% morbidity and no mortalities. Diarrhea was consistently seen in infected poults at 2-3 days post infection (dpi) with duration of about 3-5 days. Mean body weights of birds infected at 2 or 28 days of age were significantly reduced compared to uninfected birds by 7 days post-infection and remained significantly lower for the duration of the study. At 44 days of age, poults infected at 2 or 28 days of age weighed only 68.1% or 77.7%, respectively, compared to uninfected turkeys of the same age on the same diet, a mean difference in body weights of 683 or 477g, respectively. Infected birds had profound villus atrophy with some compensatory crypt hyperplasia at 5 to 7 dpi. Villus heights in the duodenum were significantly reduced at 7 dpi. We were able to reproduce enteric disease using only a pathogenic field isolate (MG10) of TCoV that negatively affected growth performance and intestinal morphology of young turkey poults Gomaa et al., (2009b). DISCUSSION TCoV is the causative agent of Bluecomb disease of turkey poults. Since the original description of ‘mud fever’ in the USA caused by TCoV, it has since been found in Canada, Dea et al., (1991); Gomaa et al., (2008a), Great Britain : Cavanagh et al. (2005), and Brazil Teixeira et al., (2007). Despite the initial uncertainty regarding its relationship to other coronaviruses, TCoV has been shown to be a group III coronavirus closely related to IBV of chickens Gomaa et al., (2008a). A study has separated group III coronaviruses into three subgroups, and TCoV is classified as a group IIIa coronavirus along with IBV, Woo et al., (2008). Using a field isolate of a pathogenic turkey coronavirus, the full genome sequence was obtained and, based


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on these sequence data, diagnostic methods (ELISAs) based on two recombinant antigens were developed. Using these tests, seroprevalence of TCoV in turkey farms and pathogenesis of TCoV infection were studied in experimentally infected turkey poults. The complete genome sequence of a pathogenic TCoV determined in the current study was used to clarify the genetic relatedness of TCoV with other coronaviruses. A pathogenic isolate of turkey coronavirus, designated TCoV-MG10, was recovered from an outbreak of acute enteritis in young turkeys in Ontario Canada and the full-length genomic sequence was determined. The full-length genome was 27,632 nucleotides plus the 3’ poly(A) tail. Two open reading frames, ORFs 1a and 1b, resided in the first two thirds of the genome, and 9 additional downstream ORFs were identified. A gene for hemagglutinin-esterase was not identified in TCoV. The region between M and N protein genes contained 3 potential ORFs. A novel ORF was located immediately downstream the M gene and upstream of gene 5 and was designated ORF-X. Further studies are required to identify the nature of this particular ORF. ORF-X has an associated putative TRS within the M gene; ORF-X and its putative TRS are shared among all group III coronaviruses examined thus far. ORFs 5a and 5b were located immediately upstream from the N gene and immediately follow ORF-X. The TCoV genome is organized as follows: 5’UTR-replicase (ORF1a and ORF1b)-spike (S) protein-ORF3 (ORFs 3a and 3b)- small envelop (E or 3c) protein- membrane (M) protein- ORF-X, ORF5 (ORFs 5a, and 5b) -nucleocapsid (N) protein-3’UTR-poly(A). The TCoV genome structure and sequence were most similar to IBV. Based on the data obtained from the current study and that of other studies, group III coronavirus can be divided into three subgroups. Group IIIa includes the IBV and TCoV; group IIIb includes Whale CoV (SW1), while group IIIc includes the most recently identified avian coronaviruses bulbuls CoV (BuCoV-HKU11), Thrush CoV (ThCoV-HKU12) and Munia CoV (MuCoVHKU13) Woo et al., (2008). TCoV is one of the most important etiological agents of diarrhoea in young turkey poults Cavanagh, (2005) and is believed to be involved in PEMS (Poult Enteritis and Mortality Syndrome). First efforts to determine seroprevalence of TCoV were based on virus neutralization Pomeroy et al., (1975) and fluorescent antibody tests Patel et al., (1976). Subsequently, ELISA based detection methods were attempted; however, determining the seroprevalence of TCoV in commercial turkey populations remained difficult because TCoV cannot be propagated in cell culture Aly and Reynolds, (1998), Guy et al., (1997); Breslin et al., (2001); unpublished observations), although some isolates can be propagated in turkey embryos Pomeroy et al., (1975). Thus, a variety of antigens have been used to determine the seroprevalence of TCoV in turkey flocks. These include IBV whole virus antigen for antibody detecting ELISA, Patel et al., (1976), recombinant TCoV-N protein derived from a prokaryotic expression system, Loa et al., (2000) and the baculovirus expression system for ELISA or competitive ELISA tests Guy et al., (2002). Two recombinant antigen ELISAs were developed in the present study. The efficacies of the two ELISAs were compared to those of a commercially available IBV ELISA, which is currently used for detecting TCoV infections. The recombinant N protein was expressed, purified and used as an antigen to develop an ELISA for serological detection of TCoV. The N-based ELISA showed high sensitivity (97%) and specificity (93%) for TCoV, and


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these were significantly higher than those of the IBV based commercial test for TCoV. In addition to N, a portion of the S1 subunit of the spike glycoprotein of TCoV was cloned to carry B cell epitopes (amino acid positions 54-395) Gomaa et al., (2009a) and expressed in E. coli. This protein fragment was purified and used to develop an indirect ELISA for detection of antibodies against TCoV. This ELISA also showed high sensitivity (95%) and specificity (92%) for TCoV. To further evaluate the potential of the ELISA, 360 and 81 serum samples were collected from commercial turkey farms in Ontario and from Arkansas, USA, respectively, and tested for TCoV-specific antibodies. High seroprevalence of TCoV was found in 71.11% of breeders and 56.67% of meat turkeys. A significantly positive correlation was found with a TCoV N protein-based ELISA, and there was little to no correlation with the whole IBV antigen-based ELISA. High seroprevalence of TCoV was also found on the Ontario farms with 73.9% of breeders and 60.0% of meat turkeys testing seropositive using the N based ELISA. Similarly, high field prevalence was found in Arkansas where 64.2% of the serum samples tested seropositive. Only a few epidemiological surveys have been conducted to determine TCoV seroprevalence. Using IFA or the commercially available IBV ELISA (IDEXX), it was determined that in Indiana 175/325 (53.84%) or 163/325 (50.15%) of field samples were positive for TCoV, respectively, Loa et al., (2004). In a second study using a competitive ELISA and a recombinant TCoV N protein, 28% of sampled turkey poults were found to possess antibodies to TCoV. Our recombinant TCoV N based ELISA detected 73.89% and 60.00% positive sera from breeder turkey farms and meat turkey farms in Ontario, Canada, respectively, and the TCoV S54 395 based ELISA found 71.11% and 56.67% positive with the same samples. Field serum samples obtained in Arkansas showed similar seroprevalence. The higher seroprevalence detected using our recombinant ELISAs compared to results using the commercially prepared IBV whole virus ELISA plate (Loa et al, 2000) and our results using the IDEXX IBV antigen plates suggests that TCoV specific antibodies cross-react with IBV antigens but to a lesser extent than ELISAs based on the homologous antigen. The prevalence of TCoV antibodies was high in both breeder turkey and meat turkey flocks as demonstrated by our data, but the prevalence was significantly higher in the turkey breeder flocks. This is perhaps because these birds in the latter group were older and thus more likely to have been exposed to the virus to mount a repeated antibody response. The course of TCoV infection in young turkey poults was examined using a field isolate (TCoV-MG10) from a diarrheal disease outbreak on a commercial turkey farm in Ontario, Canada. Two-day-old and 28-day-old poults were inoculated orally with TCoV-MG10 to examine the effect of age on viral shedding and serum antibody responses to the virus. The presence of coronavirus like particles measuring 105.8±21.8 nm in the cloacal contents was confirmed using transmission electron microscopy. The pattern of cloacal TCoV shedding was examined by RT PCR amplification of the nucleocapsid (N) gene fragment. TCoV serum antibody responses were assessed with two recently developed TCoV ELISAs that used TCoV N and S1 polypeptides as coating antigens. Poults were found equally susceptible to TCoV infection at 2 days of age and at 28 days of age and turkeys of either age shed virus in their feces starting as early as 1 day post-inoculation and for up to 17 days post-inoculation. Poults infected at two days of age were


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immunologically protected against subsequent challenge at 20 days post inoculation. The protection was associated with measurable serum antibody responses to both the nucleocapsid and S1 structural proteins of TCoV that were detectable as early as one week post-infection. We were able to reproduce enteric disease using only a pathogenic field isolate (MG10) of TCoV that negatively affected growth performance and intestinal morphology of young turkey poults. Clinical signs and weight gain of poults during experimental infections were compared with age matched-uninfected controls. Poults infected at 2 days of age had 100% morbidity and 10% mortality and birds infected at 28 days of age showed 75% morbidities and no mortalities. Diarrhea was seen in infected poults at 2-3 (dpi) with duration of about 3-5 days. Mean body weights of birds infected at 2 or 28 days of age were significantly reduced compared to uninfected birds by 7 days post-infection and remained significantly lower for the duration of the study. At 44 days of age, poults infected at 2 or 28 days of age weighed only 68.1% or 77.7%, respectively, compared to uninfected turkeys of the same age on the same diet, a mean difference in body weights of 683 or 477g, respectively. Infected birds had profound villus atrophy with some compensatory crypt hyperplasia at 5 to 7 dpi. Villus heights in the duodenum were significantly reduced at 7 dpi. Infected birds showed signs of enteritis in the form of ballooning of the intestine, which was filled with frothy material. The intestine had a thin wall and was flaccid. These lesions are consistent with what is expected during the enteric virus infections as previously reported, Ismail et al., (2003). In the same manner, the microscopic lesions were in the form of shortening of the villi with increasing in the crypt depth together with infiltration of heterophils mainly eosinophils as an indicator of the host response to the viral infection. In an attempt to study the age susceptibility in the TCoV infection, 8 birds from the control group were randomly selected at 28 days of age and were inoculated orally with TCoV. These poults developed diarrhoea 24 hours post infection, which lasted for 3-5 dpi then stopped completely. There was a slight decrease in the body weights of infected birds in comparison to uninfected birds. The immunological profile of these birds was the same but they developed higher antibody titter in a shorter duration compared to the birds that were infected at early stage of life. This result was found to be consistent with some earlier studies which reported that TCoV infection is usually associated with varying mortalities in young turkey poults while it is usually associated with drop in the productivity of the affected turkey flocks either for meat or for egg production Guy, (2000); Guy, (2003). Our results revealed the seroconversion of the birds to the viral infection as we were able to detect antibodies against TCoV as early as one week post infection reaching a peak 3 weeks after inoculation. In order to study the effect of a recurrent infection of the same flock of turkey with the same virus as may happen under field conditions, we gave the birds another dose of virus in the same manner at the 21 days of age. Challenged birds were not observed to develop diarrhoea after infection and the curve of the antibodies reached to a very high peak after 3 weeks after this challenge infection. These data indicated that the antibodies developed in response to the first infection were able to prevent the clinical diarrhoea and other clinical signs upon challenge with virulent TCoV. The second exposure to the virus acted as a booster dose that increased the titer of TCoV-specific antibodies to a high level.


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Lugovskaya , N. N., A. V. Scherbakov, A. S. Yakovleva, M. A. Tsyvanyuk, N. S. Mudrak, V. V. Drygin, and A. V. Borisov. 2006. Detection of antibodies to avian infectious bronchitis virus by a recombinant nucleocapsid protein-based enzyme-linked immunosorbent assay. J. Virol. Methods 135:292–296.

Patel, B. L., Deshmukh, D. R. and Pomeroy, B. S. 1975. Fluorescent antibody test for rapid diagnosis of coronaviral enteritis of turkeys (bluecomb). Am. J. Vet. Res. 36: 1265-1267.

Pomeroy, B. S., Larsen, C. T. Deshmukh, D .R. and Patel, B. L. 1975. Immunity to transmissible (coronaviral) enteritis of turkeys (blue comb). Am. J. Vet. Res. 36: 553-555.

Qiu, M., Wang, J., Wang, H., Chen, Z., Dai, E., Guo, Z., Wang, X., Pang, X., Fan, B., Wen, J., Wang., J., and Yang, R. 2005. Use of the COOH portion of the nucleocapsid protein in an antigen-capturing enzyme linked immunosorbent assay for specific and sensitive detection of severe acute respiratory coronavirus. Clin. Diagn. Lab. Immunol. 12: 474 476.

Seah, J. N., L. Yu, and J. Kwang. 2000. Localization of linear B-cell epitopes on infectious bronchitis virus nucleocapsid protein. Vet. Microbiol. 75:11-16.

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Teixeira, M. C. B, Luvizo, M. C. R., Ferari, H. F., and Mendes, A. R. da Silva, S. E. L., and Cardoso, T. C. 2007. Detection of turkey coronavirus in commercial turkey poults in Brazil. Avian. Pathol. 36: 29-33.

Tumlin, J.T., B. S. Pomeroy, and R. K. Lindorfer. 1957. Bluecomb 540 disease of turkey. IV. Demonstration of a filterable agent. J. Am. Vet. Med. Assoc.130:360-365.

Woo, P. C., Lau, S. K., Lam, C. S., Lai, K. K., Huang, Y., Lee, P., Luk, G. S., Dyrting, K. C., Chan, K. H. and Yuen, K. Y 2008. Comparative analysis of complete genome sequences of three novel avian coronaviruses reveals a novel group 3c coronavirus. J. Virol. In Press. [doi:10.1128/JVI.01977-08].


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PATHOLOGY AND VIRUS TISSUE DISTRIBUTION OF TURKEY CORONAVIRUS (TCOV) IN EXPERIMENTALLY INFECTED CHICKS AND TURKEY POULTS

GOMES1 DE, HIRATA KY 1, TEIXEIRA MCB 1, FERRARI HF2,. VICENTE RM1, LUVIZOTTO MCR2, GAMEIRO R 1 and CARDOSO1 TC

São Paulo State University (UNESP), 1 Laboratory of Animal Virology, DAPSA, and 2 Laboratory of Veterinary Pathology, DCCRA, Veterinary School, Clóvis Pestana

Street 793, 16.050-680, Araçatuba, São Paulo, Brazil

SUMMARY Fifteen specific pathogen free chicks and commercial poults aged 1 day were inoculated with Brazilian strain of turkey coronavirus (TCoV). The purpose was to elucidate the pathogenicity and viral distribution in TCoV-infected chicks and poults over a period of 7 days post-inoculation (dpi), by histopathology analysis, in situ reverse transcriptase chain reaction (in situ RT-PCR) and immunohistochemistry (IHC). At 2-7 dpi, the TCoV-antigens were detected in paranasal sinus, lachrymal accessory gland (Harderian gland) of infected chicks; ileum, ileum-cecal junction and ceca portions of infected poults. In all positive tissues, an intense inflammatory infiltrate, considered as multifocal distribution of lymphocytes, were described. Positive cells typically exhibit a brown reaction product (in situ RT-PCR and IHC) or red deposit (in situ RT-PCR) the cytoplasm of infected cells. TCoV was re-isolated from a pool of tissue, corresponding to nasal concha, paranasal sinus, larynx, cranial portion of trachea (chicks), jejunum, ileum-cecal junction and ceca (poults) after 3 consecutive passages on 28-old embryonated turkey eggs. The results suggested that TCoV, isolated from field case in Brazil, was able to replicate in respiratory epithelial and glandular cells of experimental infected chicks. INTRODUCTION The Brazilian turkey industry ranks second producer in the world, and last year more than 187 millions of carcases were produced. In 2006, an outbreak of Poult Enteritis Mortality Syndrome was detected for the first time occasioned by a group III Coronavirus (Teixeira et al., 2007). Since then, the respective virus was isolated (BR/TCoV/2006) and partial sequence from the spike gene has been sequenced (Cardoso et al., 2008b). According to Cavanagh (2005) the primary disease presentation of PEMS is diarrhoea, restlessness, and a general poor condition of the poult, associate to high mortality and morbility. The disease has been described


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worldwide (Adams and Hofstad, 1971; Dea et al., 1991; Brown et al., 1997; Cavanagh et al., 2001; Teixeira et al., 2007; Pantin-Jackwood et al., 2008) and recently, reported in Great Britain resulting in a first report of this disease (Culver et al., 2006). Previous works has demonstrated that turkey coronavirus (TCoV) is closely related to infectious bronchitis virus (IBV) of chickens however, not so many studied the pathogenesis of virus infection in different hosts (Breslin et al., 1999a, b; Cavanagh et al., 2001). Besides, the TCoV also was successfully propagated in turkeys and chicken embryos as well when inoculated into the amniotic cavity (Gough et al., 1988; Guy, 2008). Coronaviruses had often been described to be fastidious. This claim arose from the difficulty to find type of cells in which TCoV can propagate in vitro, explained by the use of turkey embryos as source of infective virus in the laboratory (Guy, 2008). Based on the fact that coronavirus is not necessary limited to replicating in, or causing disease in a single host, this study aimed to describe if TCoV can replicate in chickens. The pathological changes and virus distribution from 1-day specific pathogen free chicks and commercial poults, experimentally infected by TCoV isolated were described.

MATERIAL and METHODS Virus The TCoV strain (BR/TCoV/2006 accession number FJ188401), isolated from field cases of PEMS in 2007 (Teixeira et al., 2007), purified and characterized in 2008 (Cardoso et al., 2008b), was used for experimental inoculation of specific pathogen free (spf) chicks and commercial poults. The virus was propagated in our laboratory with 24-day-old embryonated commercial poults inoculated via the amniotic sac rout (Guy, 2008). The intestines of inoculated embryos were collected 48h after inoculation, homogenized, and clarified by centrifugation at 3000 x g for 30 min. The supernatants were filtered through 0.45- and 0.25-µm syringe filters (Corning), respectively. Aliquots of 1ml were frozen at -86°C, and then thawed out and used for titration before experimental inoculation. The spf chicks were obtained from a free of detectable chicken pathogen flock, raised for vaccine production in Brazil (Biovet). The birds were kept in wire cages inside high-security isolation rooms provided with feed and water ad libitum. In order to certify that the virus recovered was TCoV, sequencing of 3ÙTR and spike gene position 1178 reverse primer S2 +, and position 2073 forward primer S2 – .was performed. Experimental inoculation of turkey and chickens Fifteen 1-day-old spf chicks and commercial poults were inoculated with102.3 embryo infective dose [EID/50] of TCoV per bird by oral route. The same number of birds were kept into separated room and served as unexposed controls. For both inoculations the birds were observed twice a day for clinical signs. Equal number of chicks and poults (n=5.0) from each inoculation were euthanized by CO2 overdose at 2, 5 and 7 days post-inoculation (dpi). Tissues from both infected species corresponded to nasal concha, paranasal sinus, larynx, cranial portion of trachea, ileum, ileum-cecal junction and ceca were collected.


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Virus isolation Tissues suspension were prepared from a pool of nasal concha, paranasal sinus, larynx and cranial portion of trachea and pool of ileum, ileum-cecal junction and ceca fragments from all infected chicks and poults at different dpi. All samples were homogenized with twofold volume of Minimal Essential Medium (MEM), clarified by centrifugation at 2,500 x g for 20 min, and filtered twice through 0.45- and 0.22-µm syringe filters. Twenty-eight-day-old embryonated turkey eggs were inoculated with 300 µL of each tissue suspension by amniotic sac route and harvested after 48h. Harvested tissues were processed and propagated serially as described above for three consecutive passages. Tissue suspensions of the third passage were processed, homogenized, clarified, filtrated and a suspension of 200 ml was kept at -86 C for reverse polymerase chain (RT-PCR) reaction search (Teixeira et al., 2007; Culver et al., 2008; Pantin-Jackwood et al., 2008). Histopathology and Immunohistochemistry Paraffin-embedded tissues were sectioned, mounted, stained with hematoxylin and eosin (HE), and examined. Examiner was blind to the bird species and scored the lesions as follow: -, no lesion; +/-, minimal; +, mild; ++, moderate; +++, severe for severity of inflammatory reactions and distribution of lesions as focal, multifocal and diffuse. Unstained sections (4m) were used for direct immunohistochemical examination after deparafinisation, rehydratation and washes in buffered saline added by 0.1% Tween 80. Just before staining, slides were treated three times with 3% of hydrogen peroxide diluted into methanol for 30 min to inactivate endogenous peroxidase. The slides were then washed for 5 x 10 min in buffered saline to remove the residues between each step of the reaction, and the nonspecific binding was blocked using dried 15% nonfat milk for 90 min. The viral antigen was demonstrated by the avidin-biotin complex (ABC) immunoperoxidase method as described before, with some modifications (Cardoso et al., 2008b). The optimum primary antibody dilution determined by previous titration on indirect ELISA was 1:200 in PBS plus 10% of nonfat dried milk. Slides were covered by 200l of diluted antibody overnight at 4C in a humidified chamber. After 5 washes, 100l/slide of streptavidin-peroxidase complex (Sigma-Aldrich, St Louis, MA; cat # S-5512) was added and incubated for 1h at 37C. In addition, substrate made fresh in the dark, by mixing equal volumes of 0.02% hydrogen peroxide and 0.6mg DAB (3,3´- diaminobenzidine tetrahydrochloride, Gibco BRL cat # 15972-011), was added to the slides for 30 min at room temperature. After staining, sections were counterstaining with hematoxylin, air dried and coverslipped. Positive controls consisted of tissue sections from TCoV-infected poults in which the infection has been confirmed by RT-PCR amplification. Negative controls consisted of tissues collected from no-inoculated poults, for which infection status was confirmed to be negative. The intensity of staining in each section was scored as follows: -, no antigen staining; +/-, rare; +, infrequent; ++, common; +++, widespread staining. In-situ Reverse Transcription Polymerase Chain Reaction (in situ-RT-PCR) In this study it was used the modified primer combination UTR11- /UTR41+ which are incriminated to produce a very sensitive RT-PCR and was used to detect TCoV in gut contents on oropharyngeal swabs (Cavanagh et al., 2001). Fixed tissues were


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dehydrated through an auto-dehydrator, embedded in paraffin, cut into 5-µm sections and placed on glass slides pre-treated with poly-lysine (Sigma-Aldrich). Sections were dried overnight at 37°C and stored at 4ºC. All reverse primers were synthesized as being labeled to biotin (Invitrogen, Brazil). The sections were first de-paraffinized in xylene and re-hydrated through graded concentrations of ethanol followed by proteinase K treatment (5µg/ml) for 5 min at 37ºC. After, the slides were rinsed in PBS and hydrated in ethanol 70%, 50% and ultra-pure water. Before start the in situ RT-PCR reaction, all the slides were submitted to heat at 95°C during 5 min in a Thermal Cycler (Mastercycler Eppendorf, Germany), and left on ice until start the reaction. A 50µl volume of RT-PCR reaction mixture (containing 500µM MgCl2, 250 dNTP mix, 2µl each primer, 0.5U Superscript III RT enzyme, 2.5U/µl of TAQ polymerase platinum high fidelity and ultra-pure water) was added around each section, limited by in situ frame. After covering with a cover slip, the sections were placed into the humidified chamber and the in situ RT-PCR conditions followed. These conditions were: 42°C for 60min, 94°C for 1 min, 56°C for 1 min and 72°C for 2 min, during 34 cycles. The amplified products were extended by incubation at 72ºC for 10min. The amplified products were revealed by addition of ExtrAvidin-alkaline phosphatase, diluted 1:100 and incubated 30 min at 37°C under humidified chamber. In addition, substrate made fresh in the dark, SIGMAFAST Fast Red TR/Naphtol As-Mix (cat # B5655) was added to the slides for 30 min at room temperature. The reaction was stopped by washing with distilled water and the specific red and blue color, respectively was revealed after counterstained with aqueous hematoxylin. A distinguish intensive red, respectively was considered positive and the negative controls consisted of sections treated with biotin IBV labeled primers and omission of RT step (Cardoso et al., 2008a) Micrographs Analysis Photomicrographs of positive infected cells were observed under transmitted light Axio Imager A.1 microscope connected to AxioCam MRc3 (Carl Zeiss Oberkochen, Germany). The micrographs were processed with Axiovision 4.7 software (Carl Zeiss) and documented. RESULTS Virus isolation The TCoV could be recovered from tissue suspensions after three consecutive passages in embryonated turkey embryos, from a pool of nasal concha, paranasal sinus, larynx and cranial portion of trachea of inoculated chicks and also from ileum, ileum-ceca and ceca of infected poults. The confirmation was performed by amplification of 3ÚTR region and Spike gene of TCoV region using RT-PCR (Fig. 1A and B). Histopathology findings As suspected, no clinical signs neither microscopic lesion could be observed from any uninfected birds. However, infected poults showed slight watery diarrhoea at 5 and 7 dpi. In comparison, the infected chicks showed mild to moderate focal inflammatory cells, close only to paranasal sinus (Fig. 2A) and lachrymal accessory gland (Fig. 1B), also denominated as Harderian gland. At 2 dpi, for both chick and


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poult infected with TCoV, no lesions were observed. Regarding to intestine microscopic achieves in infected poults, multifocal enteritis, predominantly in ileum-ceca junction, consisting of infiltration of mononuclear cells, mostly lymphocytes was visible (Fig. 3A). At 5 and 7 dpi, disruption of enterocytes villous and congestion could be also observed (Fig. 3B). In a panoramic view of intestinal lumen, exfoliated enterocytes and differences in villous height/crypt depth could be observed at 5 and 7 dpi (Fig. 3B). No lesions were detected in any other tissue or in any organ of uninfected controls. Viral distribution in infected chicks and poults as demonstrated by IHC and in situ-RT-PCR Epithelial and glandular cells from paranasal sinus and Harderian gland, from TCoV-infected chicks at 5 and 7 dpi presented both viral antigen and respective genome characterized by dark-brownish granules confirmed by IHC and in situ RT-PCR, respectively. No specific peroxidise-positive staining was seen in the order tissues or of the uninfected control. Positive staining was observed in the cytoplasm of enterocytes at the tip and middle section of affected villous, distributed in all portions from TCoV-infected poults at 5 and 7 dpi by IHC and in situ RT-PCR. Table 1 summarized the number of positive results (TCoV nucleic acid and antigen) in 5 specific tissues from the subgroups of five infected poults and chicks killed at 3 intervals days after inoculation. The sensitivity and specificity of in situ RT-PCR and IHC were calculated from the data in Table 1. The sensitivity of each of the two methods, determined by dividing the number of TCoV nucleic-acid and antigen-positive ileum, ileum-cecal junction and ceca related to a total of 15 samples, was 73% and 60%, at 5 and 7 dpi respectively. In fact, the two methods did not present significant difference in sensitivity relates to ileum and ileum-cecal junction collected at 5 dpi (Table 1). The sensitivity of each of two methods to detect TCoV nucleic-acid and antigen-positive paranasal sinus and Harderian gland was 66% and 26% for 5-7 dpi (Table 1). TCoV nucleic acid and antigen were not detected in nasal concha, larynx and cranial portion of trachea of any inoculated chick or poult. The specificity of the two methods, determined by dividing the number of control samples negative for TCoV nucleic acid or antigen by the number of control birds, was 100% in each dpi. Positive cells typically exhibited a brown reaction product (in situ RT-PCR and IHC) in the cytoplasm. However when substrate of alkaline phosphatase enzyme was used positive red deposit was evidenced. TCoV nucleic acid and antigen were not detected in poult paranasal sinus and Harderian gland. The same was observed for ileum, ileum-ceca junction and cecum of all inoculated chicks. Intense and specific positive signals were most often seen within paranasal sinus and Harderian gland of chicks and ileum-cecal junction of poults. DISCUSSION The BR/TCoV/2006 accession number FJ188401 and FJ9557899 strain of turkey coronavirus, which caused the first episode of PEMS in Brazil, appeared to be able to replicate in cells different from intestinal epithelium. This conclusion was consistent with the positive results from virus re-isolation and in situ RT-PCR and IHC approaches. TCoV nucleic acid and antigen were detected in respiratory tissue, most

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consistent labelling in paranasal sinus and Harderian gland of infected chicks. Simultaneous detection of viral protein and nucleic acid provided molecular evidence of viral replication in these cells. The large amounts of viral protein expressed in villous enterocytes, together with the presence of high copy numbers of virus-specific RNA, strongly confirmed, at least in poults, that villous enterocytes support TCoV replication and represent a main target of virus, as reported before (Dea et al., 1991; Heggen et al., 1998; Ismail et al., 2003; Teixeira et al., 2007). The virus re-isolation confirmed these achieves, however it was difficult to precise, by the reason of tissue amount, which from nasal concha, paranasal sinus, larynx and cranial portion of trachea chicks tissues were individually infected. So, by pooling the samples, it was possible to recover viable virus after 3 consecutive passages, which could be explained by low replication rate of TCoV in these cells. According to previous studies, the samples prepared from ileum, ileum-cecal junction and ceca revealed infective virus in all individual portions (Teixeira et al., 2007; Cardoso et al., 2008b; Guy, 2008). It has been reported that PEMS caused alterations in cells of immune system, including down regulation of macrophage function, reduction expression of lymphocytes subpopulations in the thymus, and spleen (Heggen et al., 1998). However, no remarkable histological changes in bursa and thymus could be observed in the present study. Moreover, according to retrospective review, the propagation of TCoV on turkey and chicken embryos via amniotic route has been documented and suggested that TCoV is not pathogenic to chickens, but replication of the virus in the guts raises a question about the potential role of chickens as carriers of the virus (Ismail et al., 2003). Interesting, the bovine Coronavirus has been also described able to replicate in poults, but not in chickens (Ismail et al., 2001). The number of avian species in which coronaviruses have been detected has doubled in the past couple of years and the host range of Group III coronavirus does extend beyond the chickens (Cavanagh, 2005). Clearly, by retrospective view, there is a potential for the emergence of new coronavirus diseases in domestic birds, from both avian and mammalian sources. In situ RT-PCR and IHC detect TCoV nucleic acid and proteins and enable comparisons to be made of transcriptional and translational events in cells infected with TCoV. The slightly greater sensitivity of in situ RT-PCR has been attributed to higher quantities of TCoV nucleic acid than of proteins at 5-7 dpi. The current understanding of the replication cycle of coronavirus is that large quantities of mRNA begin forming within 12-24 h of cellular infection; this is then followed by proteins synthesis (Cavanagh, 2005). In summary, the data presented in this study, further corroborate previous findings (Teixeira et al., 2007) in demonstrating that poults are susceptible to BR/TCoV/2006 infection, by their pathological changes and virus distribution. In addition, our data also represents the first description of TCoV replication in respiratory cells of paranasal sinus and Harderian gland of chicks. ACKNOWLEDGEMENTS This research was carried out with co-operation of Biovet Laboratories (São Paulo, Brasil), supported by FAPESP (grants 05/52994-3; 07/56041-6; 07/53090-6; 08/50380-6; 08/09945-0; 09/50800-8) and CNPq (grant 472226 / 2007-0).

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Brown, T. P., Garcia, A. P. and Kelly, L. (1997). Spiking mortality of turkey poults: I. Experimental reproduction in isolation facilities. Avian Diseases, 41, 604-609.

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Cardoso, T. C., Castanheira, T. L., Teixeira, M. C., Rosa, A. C., Hirata, K. Y., Astolphi, R. D. and Luvizotto, M. C. (2008b). Validation of an immunohistochemistry assay to detect turkey coronavirus: a rapid and simple screening tool for limited resource settings. Poultry Science, 87, 1347-1352.

Cavangh, D. (2005). Coronaviruses in poultry and other birds. Avian Pathology, 34, 439-448.

Cavanagh, D., Mawditt, K., Sharma, M., Drury, S. E., Ainsworth, H. L., Britton, P. and Gough, R.E. (2001). Detection of a coronavirus from turkey poults in Europe genetically related to infectious bronchitis virus of chickens. Avian Pathology, 30, 365-378.

Culver, F., Britton, P. and Cavanagh, D. (2008). RT-PCR detection of avian coronaviruses of galliforms birds (chicken, turkey, pheasants) and in parrots. Methods Molecular Biology, 454, 35-42.

Culver, F., Dziva, F., Cavanagh, D. and Stevens, M. P. (2006). Poult enteritis and mortality syndrome in turkeys in Great Britain. Veterinary Record, 159, 209-210.

Dea, S., Verbeek, A. and Tijssen, P. (1991). Transmissible enteritis of Turkeys: experimental inoculation studies with tissue-cultured-adapted turkey and bovine coronaviruses. Avian Diseases, 35, 767-777.

Gough, R. E., Alexander, D. J., Lister, M. S. and Cox, W. J. (1988). Routine virus isolation or detection in the diagnosis of diseases of birds. Avian Pathology, 17, 893-907.

Guy, J. S. (2003). Turkey Coronavirus Enteritis. In: Disease of Poultry, 11th Edit., Y. M. Saif, H. J. Barnes, J. R. Glisson, A. M. Fadrly, L. R. McDougald and D. E. Swayne, Eds, Iowa State University Press, Ames, pp. 300-307.

Guy, J. S. (2008). Isolation and propagation of coronavirus in embryonated eggs. Methods Molecular Biology, 454, 109-117.

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Heggen, C. L., Qureshi, M. A., Edens, F. W., Barnes, H. J. and Havenstein, G. B. (1998). Alterations in lymphocytic and mononuclear phagocytic systems of turkey poults associated with exposure to pout enteritis and mortality syndrome. Avian Diseases, 42, 711-720.

Ismail, M. M., Cho, K. O., Ward, L. A. and Saif, Y. M. (2001). Experimental bovine coronavirus in turkey poults and young chickens. Avian Diseases, 45, 157-163.

Ismail, M. M., Tang, Y. and Saif, Y. M. (2003). Pathogenicity of turkey coronavirus in turkeys and chickens. Avian Diseases, 47, 515-522.

Pantin-Jackwood, M. J., Day, J. M., Jackwood, M. W. and Spackman, E. (2008). Enteric viruses detected by molecular methods in commercial chicken and turkey flocks in the United States between 2005 and 2006. Avian Diseases, 52, 235-244.

Teixeira, M. C, Luvizotto, M. C. R., Ferrari, H. F., Mendes, A. R., da Silva S. E. and Cardoso, T. C. (2007). Detection of turkey coronavirus in commercial turkey poults in Brazil. Avian Pathology, 36, 29-33.

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Figure 1- Positive HIS results in chickens experimentally infected with TCoV. A)

Brownish deposits in the respiratory tract; B) Sequencing of spike gene of TCoV recovered from respiratory tract.

Figure 2- Histopathological changes in infected chicks. (A) Paranasal sinus and (B)

lachrymal accessory gland (Harderian gland) sections showing multifocal inflammatory infiltrate at 5-7 dpi. HE.

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Figure 3- Histopathological changes in infected poults. (A) Ileum-cecal junction sections showing vacuolization of enterocytes and congestion. HE. (B) Ileum-ceca villous showing differences in height and crypt depth, detachment of superficial enterocytes at 5-7 dpi. HE.

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MATRIX METALLOPROTEINASES EXPRESSION IN EXPERIMENTALLY INFECTED POULTS WITH TURKEY CORONAVIRUS

CARDOSO1 TC, GOMES DE, ASTOLPHI1 RD, NOVAIS1 JB, GUEDES1 ACR, FERRARI2 HF, SILVA-FRADE1 C and LUVIZOTTO2 MCR

São Paulo State University (UNESP), 1 Laboratory of Animal Virology, DAPSA, and 2 Laboratory of Veterinary Pathology, DCCRA, Veterinary School, Clóvis Pestana

Street 793, 16.050-680, Araçatuba, São Paulo, Brazil

SUMMARY

Poult enteritis and mortality syndrome (PEMS) is an acute transmissible infectious intestinal disease accompanied by high mortality and morbidity. The etiology of this multifactorial disease remains to be elucidate; however, Turkey Coronavirus (TCoV) was initially assumed to be one the primary agent involved. Severity of this enteric disorder is most of the time associated to bacterial secondary infections. Many reports have demonstrated the association between virus infection and extracellular matrix (EM) breakdown. The EM is composed by fibrillar and no fibrillar collagens, fibronectin, laminin, and basement membrane glycoprotein. In general, inducers such as EGF (Epithelium Growth Factor), IL-1 or TNF (Tumor Necrosis Factor) enhance MMP production. Here we search for the expression of MMP-1, 2 and -9 in BM from intestine sections prepared from experimentally infected turkey embryos with TCoV. In all analyzed intestine sections, MMP9 expression was intensively visualized along the BM suggesting an important role in mediating TCoV infection and pathogenesis however the exact pathway remains unknown. INTRODUCTION

Matrix metalloproteinases (MMPs) are peptides that play an important role in maintaining integrity of extracellular matrix (ECM) of several tissues (Aijaz et al., 2006). MMPs belong to a large family of zinc- and calcium-dependent enzymes initially characterized by their capacity to degrade components of the ECM. Moreover, gelatinases MMP2 and MMP9 and collagenases (MMP1, MMP8 and MMP13) are the most studied. Transcriptional regulation by pro-inflammatory cytokines increases lymphocyte derived MMP9 levels in the airway lumen of asthmatics (Vermeer et al., 2009). The secretion of metalloproteinase tissue inhibitors (TIMPs) in a specific molar ratio of 1:1 inactivates MMPs function (Rossel et al., 2009). The mechanism by which MMP9 activity leads to asthma pathogenesis


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and remodeling remains unclear. Using a model of well-differentiated human airway epithelia, it has been found that apical MMP9 significantly increases transepithelial conductance. Moreover, apical MMP9 (also denominated gelatinase B) treatment decreased immunostaining of tight junction proteins suggesting disruption of barrier

function (Vermeer et al., 2009). Consistent with this, a hypothesis of how viruses gained access to the epithelial basolateral surface is supported by the action of MMP9, which increased infection efficiency. The inflammatory cells as neutrophils and macrophages, and also epithelium cells can synthesize the MMP9 (Vermeer et al., 2009). Fortunately, these effects can be blocked by TIMPs produced by other components still no defined. In addition, loss of epithelial integrity correlates with increased epithelial cell death. Thus we hypothesized that MMP9 exerts its effects on the epithelium by cleaving one or more components of cell-cell junctions. To asses the MMPs 1, 2 and 9 and TIMPs (1 and 2) expression, turkey embryos aged 23-25-days of incubation were infected with Turkey Coronavirus (TCoV). MATERIAL and METHODS Virus and infection The TCoV strain (TCoV/Brazil/2006 accession number FJ188401), isolated from field cases of PEMS in 2007 (Teixeira et al., 2007), purified and characterized in 2008 (Cardoso et al., 2008), was used for experimental inoculation of commercial poults. The virus was propagated and experimentally used in our laboratory to infect 24-day-old embryonated commercial poults via the amniotic sac rout. After 48h, the embryos were sacrificed and the sections from ileum, ileum-cecal junction and ceca were fixed in buffered formaldehyde and embedded in paraffin. Immunohistochemistry Paraffin-embedded tissues were sectioned, mounted, stained with hematoxylin and eosin (HE), and examined by light microscopy. Examiner was blind to the bird specie. Unstained sections (4m) were used for direct immunohistochemical examination after deparafinisation, rehydratation and washes in buffered saline added by 0.1% Tween 80. Just before staining, slides were treated three times with 3% of hydrogen peroxide diluted into methanol for 30 min to inactivate endogenous peroxidase. The slides were then washed for 5 x 10 min in buffered saline to remove the residues between each step of the reaction, and the nonspecific binding was blocked using dried 15% nonfat milk for 90 min. The MMPs and TIMPs were demonstrated by the avidin-biotin complex (ABC) immunoperoxidase method as described before, with some modifications (Teixeira et al., 2007). The optimum primary antibody dilution was MMP-1 1:100; MMP-2 1:200, MMP-9 1:1000, TIMP-1 1:200 and TIMP-2 1:300, respectively. In order to detect the TIMP1 and 2 the immunofluorescence method described by Cardoso et al. (2008) was performed. Slides were covered by 200l of diluted antibody overnight at 4C in a humidified chamber. After 5 washes, 100l/slide of streptavidin-peroxidase complex (Sigma, S-5512) was added and incubated for 1h at 37C. In addition, substrate made fresh in the dark, by mixing equal volumes of 0.02% hydrogen peroxide and 0.6mg DAB (3,3´- diaminobenzidine tetrahydrochloride, Gibco BRL cat # 15972-011), was added to the slides for 30 min at room temperature. After IHC staining, sections were counterstaining with


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hematoxylin, air dried, coverslipped, and examined. Positive controls consisted of tissue sections from TCoV-infected poults in which the infection has been confirmed by RT-PCR amplification. Negative controls consisted of tissues collected from no-inoculated poults, for which infection status was confirmed to be negative. The intensity of staining in each section was scored as follows: -, no antigen staining; +/-, rare; +, infrequent; ++, common; +++, widespread staining. Zymography Gelatine-substrate gel electroforesis was used to detect gelatinases (MMP-2 and MMP-9). Suspension of embryos intestine were separated by electrophoresis on a 10% sodium dodecyl sulphate polyacrilamide gel containing 0.3 mg/ml gelatine (SDS-PAGE). The gel was then incubated for 30min on a shaker with 2.5% Triton X-100 and washed three times in distilled water for 30min. Subsequently, the gel was incubated for 24h in a MMP-substrate buffer (50mM Tris-HCl, 5mM CaCl2, pH8) to allow proteinaeses to degrade gelatine. The gel was stained by Comassie blue and destaining solution revealed the expression of gelatinases as clear bands against a blue background. Statistical analysis Statistical analysis was performed using Statistical Analysis System (SAS) for Windows Software version 9.1 (SAS Institute). Analysis of normalized data was performed using the Wilcoxon rank test with the significance level P<0.05. RESULTS and DISCUSSION Coronaviruses had often been described as being fastidious. This claim arose from the difficulty that virologists had experienced in finding types of cells in which to grow coronaviruses in vitro (Cavanagh, 2005). TCoV cannot be grown in cell culture, so the turkey embryos are the only "in vivo" model to propagate the virus. MMP and TIMP expression were well visualized by IHC technique. However, only MMP-9 and TIMP-2 were intensively and widespread staining in ileum, ileum-cecal junction and ceca (Fig. 1B and D). In contrast, the activity of proMMP-2 protein (72 kDA) decreased in infected embryos, differently from proMMP9 (90 kDA), which was present in infected and uninfected poults in zymography assay. Negative results were obtained for MMP-1 and TIMP-1 search in all analysis. In response to cytokines, MMPs together with a number of potent chemokines, play an important role in orchestrating leuckocyte extravasation into the inflammatory focus. Indeed, of the major MMPs controlling the basement membrane turnover and tight-junction efficiency, MMP-2 and -9 have been shown to enhance leukocyte infiltration into the central nervous system in viral encephalitis (Wang et al., 2008). In this study we have partially demonstrated, for the first time, that experimental infection with TCoV display the MMP 2 and -9 expressions associated to TIMP-2 production in the turkey embryo model. In spite of these results, further studies are undergoing to establish a cell line (intestinal enterocytes) to elucidate the exact role of MMPs in the TCoV pathogenesis.

H


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ACKNOWLEDGEMENTS This research was carried out with co-operation of Biovet Laboratories (São Paulo, Brasil), supported by FAPESP (grants 05/52994-3; 07/56041-6; 07/53090-6; 08/50380-6; 08/09945-0; 09/50800-8) and CNPq (grant 472226 / 2007-0). REFERENCES Cardoso, T.C., Castanheira, T.L., Teixeira, M.C., Rosa, A.C., Hirata, K.Y., Astolphi,

R.D., Luvizotto, M.C. (2008). Validation of an immunohistochemistry assay to detect turkey coronavirus: a rapid and simple screening tool for limited resource settings. Poultry Science, 87, 1347-1352.

Cavanagh, D. (2005). Coronaviruses in poultry and other birds. Avian Pathology, 34, 439-448.

Puff, C., Krudewig, C., Imbschweiler, I., Baumgärtner, W., Alldinger, S. (2008). Influence of persistent canine distemper virus infection on expression of RECK, matrix-metalloproteinases and their inhibitors in a canine macrophage/monocytic tumor cell line (DH82). The Veterinary Journal, doi:10.1016/j.tvjl.2008.03.026.

Sulik, A., Chyczewski, L. (2008). Immunohistochemical analysis of MMP-9, MMP-2 and TIMP-1, TIMP-2 expression in the central nervous system following infection with viral and bacterial meningitis. Folia Histochemica et Cytobiologica, 46, 437-442.

Vermeer, P.D., Denker, J., Estin, M., Moninger, T.O., Keshavjee, S., Karp, P., Kline, J.N., Zabner, J. (2009). MMP-9 modulates tight junction integrity and cell viability in human airway epithelia. American Journal Physiology Lung Cellular Molecular Physiology. 296, 751-762.

Rossel, A., Alvarez-Sabín, J., Arenillas, J.F., Rovira, A., Delgado, P., Fernández-Cadenas, I., Penalba, A., Molina, C.A., Montaner, J. (2009). A matrix metalloproteinase protein array reveals a strong relation between MMP-9 and MMP-13 with diffusion-weighted image lesion increase in human stroke. Stroke, doi: 10.1161/01.STR.0000170641.01047.

Teixeira MC, Luvizotto MC, Ferrari HF, Mendes AR, da Silva SE, Cardoso TC. (2007). Detection of turkey coronavirus in commercial turkey poults in Brazil. Avian Pathology, 36, 29-33.


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Figure 1. Intestinal slides showing: A) TIMP- 1 negative results; B) TIMP-2 positive

immunofluorescence signals; C) MMP-2 positive signals inside enterocytes cytoplasm; D) Strong MMP-9 labelled enterocytes prepared from infected poults.

A B

TIMP1

TIMP2

C D


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DNA-MEDIATED VACCINATION AGAINST CHALLENGE INFECTION BY TURKEY CORONAVIRUS

LIN TL, ABABNEH M, HSIEH MK, CHEN YN and WU CC

Department of Veterinary Pathobiology and Animal Disease Diagnostic Laboratory, 406 South University Street, Purdue University, West Lafayette, IN, 47907-2065,

USA

Turkey coronavirus (TCoV) causes acute highly contagious enteric disease of turkeys, characterized by acute atrophic enteritis, diarrhea, death, or decreased body weight gain. Currently there is no effective vaccination to prevent the infection. DNA vaccination has emerged as a new approach for protection against infectious diseases in that DNA vaccine encodes a targeted gene segment from the infectious agent to enable the host to generate specific humoral and/or cellular immune response against the infection. The objective of the present study was to determine the immune response and protective efficacy of DNA vaccines based on TCoV nucleocapsid (N) and spike (S) genes in turkeys. Both TCoV N and S1 genes were cloned into pTriEX-3 expression vector, respectively. Groups of One-day-old turkeys were intramuscularly inoculated with PBS or 750 µg of pTriEX-3 vector three times at weekly interval. The other groups of one-day-old turkeys were inoculated with one dose of 750 µg of TCoV N gene-based DNA first, followed by inoculating with 750 µg of TCoV S1 gene-based DNA two times at weekly interval. The group that received one dose of N gene-based DNA and two doses of S1 gene-based DNA had significantly increased stimulation index for TCoV N protein-specific lymphocyte proliferation at 21 days for trial 1 and at 14 and 21 days for trial 2 after the first inoculation (P<0.05) or for Concanavalin A (Con A) induced lymphocyte proliferation at 14 and 21 days after the first inoculation (P<0.05). The same group had significantly increased nitrite concentration in interferon- bioassay at 14 days for trial 1 and at 21 days for trial 2 after the first inoculation (P<0.05). ELISA titers to TCoV N protein or TCoV S1 fragment were low for the group receiving one dose of N gene-based DNA and two doses of S1 gene-based DNA, but they were significantly higher than those of the control group (P<0.05). Virus neutralization was detected in the serum samples from the group that received one dose of N gene-based DNA and two doses of S1 gene-based DNA. The group receiving one dose of N gene-based DNA and two doses of S1 gene-based DNA showed significant reduction or absence of the bright green fluorescence for TCoV antigens in the enterocytes of intestinal epithelium by immunofluorescent antibody assay (P<0.05). These results indicated DNA-mediated vaccination with TCoV N protein expressing DNA and TCoV S1 protein expressing DNA can elicit specific immune response to TCoV and reduce TCoV infectivity in turkeys.


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SUBTYPES OF AVIAN METAPNEUMOVIRUS CIRCULATING IN BRAZILIAN COMMERCIAL FLOCKS

CHACON JL1; PEDROSO AC1; TOQUIN2 D; ETERRADOSSI N2; PATNAYAK D3;

GOYAL S3 and FERREIRA AJP1

1Department of Pathology, College of Veterinary Medicine, University of São Paulo, São Paulo, Brazil.

2French Agency for Food safety (AFSSA), Avian and Rabbit Virology Immunology and parasitology Unit (VIPAC), Ploufragan, France.

3Deparment of Veterinary Population Medicine, College of Veterinary Medicine, University of Minnesota, USA.

Avenida Professor Doutor Orlando Marques de Paiva 87, Cidade Universitária, CEP 05508-900, São Paulo, SP, Brazil.

SUMMARY The avian Metapneumovirus (aMPV) causes a widespread disease of turkeys and chickens. Molecular analysis of the genome and viral antigenic characteristics had enabled classification of the isolates into four subtypes: A, B, C and D. Molecular characterization of aMPV detected between 2004 to 2008 from broiler, breeder, layer and turkey flocks with or no history of vaccination against aMPV revealed dissemination of subtypes A and B in Brazil. On the other hand, subtypes C and D were not detected. DNA sequencing of seven samples showed that detected aMPV-B were not from vaccine origin. These Brazilian subtype B strains formed one monophyletic group, apart from the subtype B vaccine strain used in the country and other subtype B isolates from other different countries. The high homogeneity among Brazilian isolates indicates that all they have a common origin. This work shows the circulation of subtypes A and B in Brazil. INTRODUCTION Avian metapneumovirus (aMPV) causes acute rhinotracheitis in turkeys and plays an important role in swollen head syndrome (SHS) in chickens. In laying birds, it causes a transient drop in egg production, along with mild respiratory tract illness. Uncomplicated infections have low mortality, but cases complicated by secondary infections can result in up to 25% mortality (Jones, 1996). Based on antigenic differences and nucleotide sequence analysis of the attachment glycoprotein (G) gene, aMPV has been classified in four subtypes (A to D). Subtype C is prevalent in


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the US whereas subtypes A, B and D are found mainly in Europe (Seal et al., 2000; Cook & Cavanagh, 2002). Serological evidences indicate that aMPV is widespread in commercial flocks from different regions of Brazil. Subtypes A and B were isolated from turkeys and chickens which presented respiratory disease. The purpose of this work was to detect and differentiate the aMPV subtypes circulating in Brazilian commercial flocks. In order to able to detect the four subtypes known worldwide, a RT-PCR assay was developed and validated. MATERIALS and METHODS Samples Two hundred twenty eight samples collected from broiler, breeder, layer, turkey flocks from the main producer regions of Brazil were analyzed in this study. During 2004 to 2008, field samples were collected from non-vaccinated and vaccinated birds against aMPV which showed respiratory disease, reduced egg production and fertility disturbs. aMPV screening The primers published by Bäyon et al., (1999) amplifying a fragment of the nucleocapside (N) gene were modified in order to detect the four subtypes known world-wide. Reference strains of the four subtypes were used to validate the assay. PCR products of each subtype were sequenced to confirm the identity. After the validation, all field samples were submitted to the standardized RT-PCR assay. aMPV subtyping The aMPV-positive samples in the screening assay were submitted to other RT-PCR assay that amplifies a fragment of G gene and differentiate subtypes A and B (Cavanagh et al., 1999). In order to know the origin of the detected aMPV, nine samples were sequenced and phylogenetic analysis was carried out. RESULTS RT-PCR assay standardization and validation Using the primers designed in this study, we amplified DNA of all the four aMPV subtypes. The PCR products were sequenced and a phylogenetic tree was constructed. Each sequence could be grouped into their corresponding subtype. No DNA of other avian respiratory pathogens was amplified with these primers. aMPV screening From the 228 analyzed samples, 38 were aMPV-positive. aMPV was detected in broiler, breeder, layer and turkey flocks from six Brazilian states. aMPV subtyping Only subtypes A and B were detected. Thirteen samples belonged to subtype A and 25 were characterized as subtype B. Both subtypes A and B were detected in one layer flock. In five cases, the detected subtype was different from the subtype used in the farm for vaccination.


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Phylogenetic analysis Nine aMPVs isolates were sequenced and seven could be differentiated from vaccine strains. The Brazilian isolates were grouped into two subtypes A and B. In each subtype, the Brazilian isolates showed high genetic similarity among them. DISCUSSION The RT-PCR assay developed for aMPV screening could detect the four world-wide recognized subtypes. The phylogenetic analysis showed that is possible subtyping aMPVs using the N gene because the sequenced samples could be grouped into their corresponding subtype. This study shows that aMPV is highly spread in Brazilian commercial flocks and affects broilers, breeders, laying hens and turkeys. Only subtypes A and B could be detected in vaccinated and non-vaccinated flocks. In two cases, aMPV vaccine strains were detected from non-vaccinated flocks. This demonstrates vaccine strain capacity of horizontal transmission. The Brazilian subtype B isolates formed one monophyletic group which is different from the subtype B vaccine strain used in the country and is different from subtype B isolates of other countries. The high homogeneity of Brazilian subtype B isolates might indicate a common origin. Subtype A was the first detected subtype in Brazil (1995). After that, massive vaccination using this subtype was established in commercial flocks. In 2005, subtype B aMPV was isolated from chicken flocks showing respiratory signs (Chacón et al., 2007). In the last two years of the study, the majority of the detected aMPVs belonged to subtype B. It might suggest a change in the aMPV population in Brazil in the recent years. REFERENCES Bäyon-Auboyer, M.H., Jestin, V., Toquin, D., Cherbonnel, M. & Eterradossi, N.

(1999). Comparison of F-, G- and N-based RT-PCR protocols with conventional virological procedures for the detection and typing of turkey rhinotracheitis virus. Archives of Virology, 144, 1091-1109.

Cavanagh, D., Mawditt, K., Britton, P. & Naylor C.J. (1999). Longitudinal field studies of infectious bronchitis virus and avian pneumovirus in broilers using type-specific polymerase chain reactions. Avian Pathology, 28, 593-605.

Chacón, J.L.V., Brandão, P., Buim, M., Villarr, L.Y., Ferreira, A.J.P. (2007). Detection by RT-PCR and molecular characterization of subtype B avian metapneumovirus isolated in Brazil. Avian Pathology, 36, 383-387.

Cook, J.K.A. & Cavanagh D. (2002). Detection and differentiation of avian pneumoviruses (metapneumoviruses). Avian Pathology, 31, 117-132.

Jones, R. (1996). Avian pneumovirus infection: questions still unanswered. Avian Pathology, 25, 639-648.

Seal, B.S., Sellers, H.S. & Meinersmann, R.J. (2000). Fusion protein predicted amino acid sequence of the first US avian pneumovirus isolate and lack of heterogeneity among other US isolates. Virus Research, 66, 139-147.


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Figure. Phylogenetic relationships among Brazilian aMPV isolates and reference strains. Following alignment of sequences from the G gene. Bootstrap confidence levels are presented.

S H S B R 6 6 8 / 0 3

T R T B R 1 6 9

C V L 1 4 / 1

1 5 5 6

L A H A

S H S B R 6 6 2 / 0 3

c h ic k e n / A / B R / 1 2 1 / 9 5

S H S B R 6 6 9 / 0 3

c h ic k e n / A / B R / 1 1 9 / 9 5

8 5 4 4

a M P V / A / c h ic k e n / C H N / 6 3 1 8 / 2 0 0 4

a M P V / c h ic k e n / N ig e r ia / N I3 8 9 / 2 0 0 6

a M P V / A / B ra z i l -0 5 / U S P -0 1

a M P V / A / B ra z i l -0 7 / U S P -0 3

S u b t y p e A va c c in e u s e d in B ra z i l

F r / 8 5 / 2

F r / 8 5 / 1

a M P V / c h ic k e n / N ig e r ia / N IR 8 9 / 2 0 0 6

2 1 1 9

Is r / 1 7 0 8 / 0 2

8 7 2 S

6 5 7 4

S u b t y p e B va c c in e u s e d in B ra z i l

a M P V / B / B ra z i l -0 7 / U S P -0 6

a M P V / B / B ra z i l -0 7 / U S P -0 5

a M P V / B / B ra z i l -0 7 / U S P -0 8

a M P V / B / B ra z i l -0 6 / U S P -0 7

a M P V / B / B ra z i l -0 5 / U S P -0 4

a M P V / B / B ra z i l -0 5 / U S P -0 3

a M P V / B / B ra z i l -0 5 / U S P -0 2

P L -2 K o re a P h e a s a n t

P L -1 K o re a P h e a s a n t

1 5 a C a n a d a G o o s e

M in n e s o t a 7

M in n e s o t a 2 A

C o lo ra d o

M n -1 a

M n -2 a

9 9 1 7 8 M u s c o vy d u c k F ra n c e

9 9 3 5 0 M u s c o vy d u c k F ra n c e

0 0 0 9 4 W h o t e P e k in d u c k F ra n c e

4 3

2 6

9 9

2 96 6

3 56 0

6 5

9 9

4 6

3 5

3 2

3 9

7 9

5 8

5 1

3 9

5 4

9 8

4 8

9 9

3 6

5 7

8 4

0 .5

Subtype

Subtype

Subtype

Subtype

Brazilian field isolates

Brazilian field isolates


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FIELD OBSERVATIONS AFTER NATURAL INFECTION OF BRAZILIAN LAYER CHICKENS WITH A PHYLOGENETICALY DIVERGENT LINEAGE

OF SUBTYPE B aMPV

VILLARREAL LYB1,3, SANDRI TL2,3, ASSAYAG MS4, RICHTZENHAIN LJ2,3, MALO A1 and BRANDÃO PE2,3

1Intervet Schering Plough Animal Health, Av. Sir Henry Wellcome, 335, CEP 06741-050, Cotia, SP, Brazil

2Department of Preventive Veterinary Medicine and Animal Health, College of Veterinary Medicine, University of São Paulo, Av. Prof. Dr. Orlando M. Paiva, 87,

CEP 05508-270, Sao Paulo, SP, Brazil 3Coronavirus Research Group, Av. Prof. Dr. Orlando M. Paiva, 87, CEP 05508-270,

Sao Paulo, SP, Brazil 4Sadia

SUMMARY This survey describes the clinical manifestations in a Brazilian commercial layer chicken farm with the involvement of a previously unknown lineage of aMPV of the B subtype. Three flocks of the farm received two doses of a live subtype A aMPV vaccine and, simultaneously, two other similar flocks at the same farm were not vaccinated against aMPV (control flocks). Control flocks showed lesions on frontal sinuses and at the posterior part of the head with a caseous content with Staphylococcus aureus, together with respiratory signs, alterations of the eggs, atrophy/ degeneration of the ovaries and low production rates. Vaccinated flocks showed no clinical signs during the experiment. The phylogenetic tree showed that the subtype B aMPV lineages involved in the outbreak were grouped in an exclusive cluster, separate from other subtype B strains already detected in Brazil, suggesting the existence of at least two subpopulations of this pathogen in a possible geographic pattern. INTRODUCTION Avian metapneumovirus (aMPV) is subdivided in four different subtypes (A, B, C, and D). In commercial layers, aMPV infection may affect the quality of eggs and cause a range of reproductive tract abnormalities, including egg peritonitis, folded shell membranes in the oviduct, misshapen eggs, and ovary and oviduct regression. aMPV infections are also associated with swollen head syndrome in chickens


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(Bäyon-Auboyer et al., 2000, Gough, 2003, Hess,et al., 2004) . In Brazil, the occurrence of these signs, as well as subtypes A and B of aMPV, have already been described (D'Arce et al., 2005; Chacón et al., 2007; Villarreal et al., 2007), though more complete studies including molecular typing and its association of this with vaccine trials, serology and field observations are scarce. This survey describes the clinical manifestations observed in a Brazilian commercial layer chicken farm with the involvement of a previously unknown lineage of aMPV of the B subtype. MATERIALS and METHODS In 2008, in an aMPV-endemic region of Brazil, a multi-age commercial layer poultry farm with 800,000 multi-age birds housed in side-by-side houses was chosen for a longitudinal study following an aMPV vaccination program. Three multi-age flocks at the farm received two doses of a live subtype A aMPV vaccine at 3 and 9 weeks of age via the ocular route, followed by one dose of multivalent inactivated vaccine containing subtype A aMPV, Mass+D274 IBV, NDV and EDSV at 14 weeks of age. Simultaneously, two other similar flocks at the same farm were not vaccinated against aMPV (control flocks). At different times after the time of administration of the killed vaccine (20/22, 25/27 and 39 weeks), organ-specific pools (trachea, kidneys, lungs and reproductive tract, 6 birds per pool) and individual serum samples were collected from both vaccinated and control flocks for the detection of aMPV in the organs was done by RT-PCR to gene G (Cavanagh et a., 1999) (organs) and the serum samples were tested for antibodies against aMPV and IBV by ELISA and against Mycoplasma gallisepticum by slide agglutination. G gene amplicons were sequenced and the sequences were then used to build a amino acids tree. RESULTS Control flocks showed a characteristic lesion located above the frontal sinuses and at the posterior part of the head with a caseous content, together with respiratory signs and alterations of the eggs shell and albumen quality, low production rates and atrophy and degeneration of the ovaries. Vaccinated flocks showed no clinical signs during the experiment. Head lesions from unvaccinated birds were tested for bacteriological isolation and pure cultures of Staphylococcus aureus were found. Both control and vaccinated groups had very high aMPV antibodies titres, ranging from 9,403 to 45,014, with a decreasing trend in the vaccinated flocks, while titres for IBV antibodies were low. All samples were also negative for Mg antibodies (Table 1). By RT-PCR, subtype B aMPV was found in all two control flocks. The amplicons were purified from agarose gels and submitted to bi-directional DNA sequencing for the generation of a distance NJ tree with the MCL model and 1000 bootstrap replicates. The phylogenetic tree showed that the subtype B aMPV lineages involved in the outbreak were grouped in an exclusive cluster, separate from other subtype B strains already detected in Brazil (Figure 1), suggesting the existence of at least two subpopulations of this pathogen in a possible geographic pattern.


257

DISCUSSION Comparing the vaccinated flocks with the control flocks, despite being in contact with flocks not vaccinated against aMPV, the first showed no signs of aMPV disease while controls showed low production rates, egg shell alterations and respiratory signs including facial edema, what, in association with PCR-negative results, allow one to conclude that the vaccination schedule used was effective in preventing the manifestation of disease despite the high challenges as inferred by the high ELISA titers. Also, PCR-negative results in the vaccinated flocks suggest that the live vaccine prevented aMPV colonization in the trachea as a result of local immunity, while the killed vaccine might have prevented aMPV systemic dissemination that can lead to reproductive disease. Regarding the decreasing ELISA titers of the vaccinated flocks, it can be speculated that, as the immune status of the birds increases, a low number of susceptible individuals is available for an effective aMPV replication and transmission. Therefore, the vaccinated flocks would not show an increase in seroconversion but instead have shown a decreasing trend. Finally, PCR and phylogenetic analysis showed that a subtype B lineage of aMPV was involved in the disease. This lineage was found divergent from other lineages already described, suggesting that subpopulations of aMPV are present in Brazil and that the vaccination schedule was effective in preventing the disease despite the divergent aMPV involved. Nonetheless, as already described by Cecchinato et al. (2008), escape mutants of subtype B of aMPV can emerge for which vaccine can confer insufficient protection, a situation that might be seen in Brazil in the future as a consequence of the amino acids mutations that are accumulating in Brazilian strains of this virus as shown in the present survey. REFERENCES Bäyon-Auboyer, M.H., Arnauld, C., Toquin, D. & Eterradossi, N (2000). Nucleotide

sequences of the F, L and G protein genes of two non-A/non-B avian pneumoviruses (APV) reveal a novel APV subgroup. Journal of General Virology, 81,2723-2733.

Cavanagh, D., Mawditt, K., Welchman, D.B., Britton, P. & Gough, R.E. (2002). Coronaviruses from pheasants (Phasianus colchicus) are genetically closely related to Coronaviruses of domestic fowl (Infectious Bronquitis Virus) and turkeys. Avian Pathology, 31, 81-93.

Cavanagh, D., Mawditt, K., Britton, P & Naylor, C. J. (1999). Longitudinal field studies of infectious bronchitis virus and avian pneumovírus in broilers using type-specific polymerase chain reactions. Avian Pathology, 28, 593-605.

Cecchinato, M., Catelli, E., Lupini, C., Rixxhizzi, E., Brown, P. & Naylor, C.J. (2008)Field avian metapneumovirus evolution avoiding vaccine induced immunity. In Proceedings of the 7th International Symposium on turkey diseases, p. 176-179. Berlin, Germany.

Chacón, J.L., Brandão, P.E., Buim, M., Villarreal, L. & Ferreira, A.J. (2007). Detection by reverse transcriptase-polymerase chain reaction and molecular


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characterization of subtype B avian metapneumovirus isolated in Brazil. Avian Pathology, 36, 383-387.

D'Arce, R.C., Coswig, L.T., Almeida, R.S., Trevisol, I.M., Monteiro, M.C., Rossini, L.I., Di Fabio, J., Hafez, H.M. & Arns CW. (2005).Subtyping of new Brazilian avian metapneumovirus isolates from chickens and turkeys by reverse transcriptase-nested-polymerase chain reaction. Avian Pathology, 34, 133-136.

Gough, R. E. (2003). Avian pneumoviruses. In Saif. Y. M., Barnes, H.J., Glisson, J. R., Fadly, A. M., McDoulgald, L. R. & Swaine, D. E. Diseases of Poultry, 11th ed., Iwoa State University Press, Ames, IA, p. 308-317.

Hess, M., Huggins, M. M., Mudzamiri, R. & Heinez, V. (2004). Avian pneumovírus excretion in vaccinated and non-vaccinated specific pathogen free laying chickens. Avian Pathology, 33, 35-40.

Villarreal, L.Y., Brandão, P.E., Chacón, J.L., Assayag, M.S., Maiorka, P.C., Raffi, P., Saidenberg, A.B., Jones, R.C. & Ferreira, A.J. (2007). Orchitis in roosters with reduced fertility associated with avian infectious bronchitis virus and avian metapneumovirus infections. Avian Diseases, 51, 900-904.

Table 1. aMPV ELISA, PCR and clinical signs in the control and vaccinated flocks with the respective ages (in weeks)

GROUP FLOCK AGE SIGNS aMPV PCR aMPV ELISA (GMT)

22w + (subtype B) 45,014 1 Control

27w

Egg drop (6.5%), low egg quality, respiratory + (subtype B) 42,613

2 control 39w Egg drop (7.9%), low egg quality, respiratory, swollen head

+ (subtype B) 37,700

20w No - 31,000 3 Vaccinated

25w No - 10,500


25w No - 11,371


25w No - 9,403

+= positive


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Figure 1. Amino acids tree for the partial attachment (G) glycoprotein (aa 59 to 124) of avian metapneumovirus (aMPV) with the four subtypes. The four strains detected in the present study with the respective Genbank accesion numbers are shown in the dotted box. Numbers at each node are 1,000 rplicates bootstrap values. The bar represents the number of substitutions per site. BRSV is the bovine respiratory sincytial virus used as an outgroup.


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TURKEY RHINOTRACHEITIS OUTBREAK IN 7 WEEK OLD TURKEYS CAUSED BY A VACCINE DERIVED AVIAN METAPNEUMOVIRUS

RICCHIZZI E1, CATELLI E1, CECCHINATO M2, LUPINI C1, BROWN P3 and NAYLOR CJ3

1 Università di Bologna, Dipartimento di Sanità Pubblica Veterinaria e Patologia Animale, Via Tolara di Sopra 50, 40064 Ozzano Emilia (BO), Italy;

2 Università di Padova, Dipartimento di Sanità Pubblica, Patologia Comparata ed Igiene Veterinaria, Agripolis - Viale dell'Università 16, 35020 Legnaro (PD), Italy;

3 University of Liverpool, Department of Veterinary Pathology, Jordan Building, Leahurst, CH64 7TE Neston, United Kingdom;

SUMMARY A virus was isolated in association with respiratory disease typical of turkey rhinotracheitis, from turkeys which had been vaccinated with a B subtype Avian Metapneumovirus (AMPV) vaccine. Sequencing of the virus showed that, as in a previous similar report, the virus had originated from a live A subtype vaccine. In this instance the disease was much later, at 50 days of age, and there had been no recent history of use of the vaccine. This may indicate that AMPV vaccines are able to circulate in the environment for longer than was previously envisaged. INTRODUCTION Avian Metapneumovirus (AMPV) is a negative sense RNA virus belonging to the family Paramyxoviridae genus Metapneumovirus (Pringle, 1998). Nucleotide sequencing has shown there to be 4 virus subtypes (A,B,C and D) (Juhasz & Easton, 1994; Bayon-Auboyer et al., 2000; Seal et al., 2000) which can cause moderate disease due to upper respiratory tract infection that can become severe when complicated by secondary pathogens (Cook et al., 1991). Italian AMPV field studies from 1987 had detected predominantly subtype B viruses, and in general, subtype B vaccines were used for its control (Catelli et al., 2004). Subtype A vaccine was also used to a more limited degree, though no subtype A field viruses had been detected up to 2003. Live vaccines offered good protection but their instability can lead to a return to virulence after only a small number of back passages. In experimental conditions, disease has been seen after 4-10 back passages of vaccine in naive turkeys (Naylor & Jones, 1994). In a subsequent study, subtype A vaccine was applied to turkeys in the hatchery at one day old, prior to being placed on farms.


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Disease was observed at 3 weeks of age and a subtype A virus was detected. Sequencing of the full genome of this virus and the identification of nine vaccine specific mutations showed unequivocally that this was vaccine-derived (Catelli et al., 2006). We now report details of a subtype A AMPV isolated during an outbreak in Italy in 2003 where disease typical of AMPV infection was seen on a farm of 20,000 male turkeys. Birds had been vaccinated with subtype B vaccine at 7 days of age and the disease occurred some 6 weeks later during which period, virus was isolated and later characterized. MATERIALS and METHODS Farm Samples were collected in a turkey farm during an outbreak of a respiratory disease. All the birds were vaccinated with a live AMPV B type vaccine at 7 days old by oculo-nasal administration. The symptoms observed started at 45 day of age and were characterized by sneezing, ocular discharge and nasal exudates. All the flocks gradually showed respiratory disease and sometimes this included mortalities (10-11%) probably due to secondary bacterial infection. Collection of material 2 out of 4 flocks were sampled. In each group 10 birds were swabbed from birds showing very early respiratory symptoms. From each animal oro-pharingeal swabs were simultaneously taken for RT nested PCR and virus isolation. Swabs for RT nested PCR were air dried for 30 minutes then stored at ambient temperature prior to testing. Swabs for virus isolation were immediately immersed in a medium transport with antibiotics to suppress bacterial and Mycoplasma growth then kept refrigerated during transportation then stored at -80°C until processing. All the swabs were processed as pools. RT nested PCR A subtype specific RT nested PCR, based on G gene sequence and able to differentiate A and B subtypes, was used to detect and subtype AMPV from dry swabs and to confirm the virus isolates as AMPV. The methods for RNA extraction and RT nested PCR were performed according to Cavanagh et al. (1999). Virus isolation Virus isolation was performed in chicken embryo tracheal organ cultures (TOC) (Cook et al., 1976). Swabs were pooled and the supernatants were used to inoculate TOCs after filtration through 0.2 μm membrane filters. Ciliostasis was taken as indicator of AMPV detection. AMPV identification and subtyping was determined by RT nested PCR previously described, the RNA was extracted from the inoculated medium. Differentiation of vaccinal and field type AMPV The genome of isolated virus has been sequenced as previously described by Catelli et al. (2006) to cover specific mutations unique to the vaccine.


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Virulence assessment of AMPV isolate 259-01/03 Virulence was determined by the assessement of clinical signs. Thirdy 1-day-old commercial turkeys were housed in three positive pressure isolators with 10 poults in each. Poults in isolator 1 were inoculated with AMPV at a dose of 3,5 Log10 ID50 per bird. Poults in group 2 were inoculated with the commercial vaccine sequenced by Catelli et al. (2006). The doses represented a ten time overdose compared to the manufacturer’s recommendations. The third group were inoculated with sterile water and kept as negative control. Clinical signs were daily scored as previously described by Naylor & Jones (1994) and outlined below

0. no clinical signs 1. clear nasal exudates 2. turbid nasal exudates 3. swollen infra-orbital sinuses and/or frothy eyes.

RESULTS Isolation on TOCs of a cicliostatic agent was confirmed to be AMPV subtype A via RT nested PCR and it was named 259-01/03. It possessed 8 out of 9 of the sequence identification markers and was hence concluded to have derived from the vaccine (Table 1). At position 3553 the sequence matched that of the vaccine progenitor and other subtype A viruses. The strain was able to cause disease when inoculated in poults in experimental conditions (Figure 1). DISCUSSION The study has made one very important finding about the characteristics and use of the AMPV vaccine and additionally confirms 2 other findings. Not for the first time, a virulent vaccine derived AMPV virus has been detected which is associated with disease typical of AMPV infection. However in this instance the virus was isolated on a farm where no the A type vaccine had been applied. This means that at very least, the virus persisted in the immediate environment from a previous A type vaccination, but it is likely that the virus may have been present in the broader environment. The presence of 8 out of 9 vaccine markers is an interesting finding. The probabilities overwhelmingly imply that the virus was indeed vaccine derived. The virus has been shown to be virulent, then the back mutation at position 3553 may be involved in the reversion to virulence, and deserves further investigation. A further finding of the study is its confirmation of the limitations of cross protection between subtypes. Van de Zande et al. (2000) showed that the duration of heterologous protection declined while homologous protection was maintained. In this instance, the A subtype virus was found some 6 weeks after vaccination at a time when heterologous protection might have critically deteriorated.


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REFERENCES Bayon-Auboyer M-H., Arnauld C., Toquin D., Eterradossi N. (2000) Nucleotide

sequence of the F, L and G protein genes of two non-A/non-B avian pneumoviruses (APV) reveal a novel APV subgroup. J Gen Virol, 81, 2723-2733.

Catelli E., Cecchinato M., Ortali G., De Matteo P., Savage C.E., Jones R.C., Naylor C.J., 2004. Avian Pneumovirus in Italy. Proceedings of the 4th International Symposium on Avian Coronavirus and Pneumovirus Infections; 2004 June 20-24, Rauischholzhausen, Germany. WB Laufersweiler Verlag, Wetten berg, Germany; 2004. p. 275-281.

Catelli E., Cecchinato M., Savage C.E., Jones R.C., Naylor C.J. (2006). Demonstration of loss of attenuation and extended field persistence of a live Avian Metapneumovirus vaccine. Vaccine, 24, 6476-6482.

Cavanagh D., Mawditt K., Britton P., Naylor C.J. (1999). Longitudinal field studies of infectious bronchitis virus and avian pneumovirus in broiler using type-specific polymerase chain reactions. Avian Pathology, 28, 593–605.

Cook J.K.A., Darbyshire J.H., Peters R.W. (1976). The use of chicken tracheal organ cultures for the isolation and assay of avian infectious bronchitis virus. Archives of Virology, 50, 109–118.

Cook J.K.A., Ellis M.M., Huggins M.B. (1991). The pathogenesis of turkey rhinotracheitis virus in turkey poults inoculated with a virus alone or together with two strains of bacteria. Avian Pathology, 20, 155-166.

Juhasz K., Easton A.J. (1994). Extensive sequence variation in the attachment (G) protein gene of Avian Metapneumovirus: evidence for two distinct subgroups. Journal of General Virology, 75, 2873-80.

Naylor C.J., Jones R.C. (1994). Demonstration of a virulent subpopulation in a prototype live attenuated turkey rhinotracheitis vaccine. Vaccine, 12(13), 1225–30.

Pringle C.R.(1998). Virus Taxonomy-San Diego 1998. Archives of Virology, 143(7), 1449-59

Seal B. (1998). Matrix Protein gene nucleotide and predicted amino acid sequence demonstrate that the first US avian pneumovirus isolate is distinct from European strains. Virus Res, 58, 45-52.

Van de Zande S., Nauwynck H., Naylor C.J., Pensaert M. (2000). Duration of cross-protection between subtypes A and B avian pneumovirus in turkeys. The Veterinary Records, 147, 132–4


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Table 1. Vaccine markers. Nucleotides and genome positions

Nucleotides Genome position a

Vaccine* progenitor Vaccine* 259-01/03

2941 U A A 3553 U C U 3825 G A A 5055 A G G

5140 U C C

5929 A G G

6358 U C C

10022 U G G

11624

U C C a antigenome * da Catelli et al. (2006)

Fig. 1 Mean daily clinical scores after inoculation of isolate 259/-01/03 compared to a ten times overdose of vaccine and uninoculated controls.

0

0,5

1

1,5

2

2,5

1 2 3 4 5 6 7 8 9 10 11 12 13 14

259-01/03 Negative controls Subtype A

Mea

n da

ily c

linic

al s

core

s

Days post infection

3


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HUMAN METAPNEUMOVIRUS INFECTION IN TURKEYS

NAGARAJA KV1, VELAYUDHAN BT1, HALVORSON1 DA and GRAY GC 2

1University of Minnesota, Saint Paul, Minnesota, USA 2University of Iowa, Iowa City, Iowa, USA

SUMMARY Turkey poults free of antibodies to avian metapneumovirus (aMPV-c) were exposed to the human metapneumovirus virus (hMPV). Birds exposed to hMPV virus and those non exposed controls were monitored. Nasal turbinate, trachea and lungs were examined for viral RNA, lesions, and presence of viral antigen by RT-PCR. Inaddition, H&E and immunohistochemical staining was done on tissues collected from exposed and nonexposed birds. We developed specific RT-PCR for human metapneumovirus. We raised human metapneumovirus antiserum in rabbits for immunohistochemistry for this study. Birds were kept in the BL2 facilities of University of Minnesota at the Saint Paul. Campus. Sham-inoculated and aMPV-C inoculated birds were included in the study as controls. Birds were monitored daily for clinical signs. Birds infected with human metapneumovirus showed clinical signs similar to that of avian pneumovirus. Specific human metapneumovirus RNA was detected in the nasal turbinate of infected birds. Nasal turbinate and trachea showed inflammatory changes in birds infected with human virus. Immunohistochemistry demonstrated viral antigen in nasal turbinate of birds infected with human virus. In summary, human metapneumovirus caused a transient infection in birds exposed to the virus INTRODUCTION Human metapneumovirus (hMPV) is a single-stranded, negative-sense, nonsegmented RNA virus of the genus Metapneumovirus of subfamily Pneumovirinae of the family Paramyxoviridae, hMPV is noted for causing respiratory tract infections in children (1). The virus is distributed worldwide, and has been identified in the United States, Canada, United Kingdom, Italy, Germany, France, Israel, Australia, Asian countries, and Peru (3,4). The disease by hMPV vary from mild upper respiratory tract infection to severe bronchiolitis or bronchitis It affects all age groups, but it is more severe in young, elderly, and immunocompromised persons (5). Serologic surveys indicate that the virus is ubiquitous in nature and that new infections can occur throughout life because of incomplete protection and


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genetic heterogeneity of the virus (2,6,7). Recent studies have shown that avian metapneumovirus (aMPV) subtype C isolates from domestic turkeys and wild birds in the United States show high sequence homology to hMPV (8,9). Both viruses belong to genus Metapneumovirus and share a projected amino acid identity of 56%–88% (9). This study was conducted to examine whether human metapneumovirus (hMPV) will infect turkeys. MATERIALS and METHODS Four genotypes of hMPV obtained from the University of Iowa and aMPV C isolated from the nasal turbinates of turkeys with acute upper respiratory tract infection were used in this study. Turkey poults of 2-weeks old were divided into 6 groups of 20 each. Birds in each group were inoculated oculonasally with 1 of the 4 genotypes of hMPV or aMPV C (positive controls). Poults were monitored daily for clinical signs. Two poults from each group were killed for necropsy and sample collection at days 3, 5, and 7 postexposure. Nasal turbinates, tracheas, and lungs were tested for viral RNA by reverse-transcription (RT) PCR with specific primers (consensus primers for all of the 4 genotypes) for hMPV and aMPV. Tissue sections were stained with hematoxylin and eosin and examined for histopathologic lesions and for viral antigen by mmunohistochemical methods (10). Sera collected from poults in each group at days 14 and 21 postexposure were tested with aMPV-ELISA (11). No convincing evidence exists for any cross-reactivity between hMPV and aMPV. Briefly, poults in each group were given a score of 0 when they showed no clinical signs; a score of 1 for unilateral nasal discharge; 2 for bilateral nasal discharge; or 3 for thick, copious, bilateral nasal discharge. Unilateral sinus swelling was recorded as score 1, bilateral sinus swelling as 2, unilateral conjunctivitis as 1, and bilateral conjunctivitis as 2. The total score for each bird was expressed as the sum of individual scores mentioned above. Nasal turbinates, tracheas, and lungs were used for RT-PCR. Viral RNA was extracted from the tissue homogenate supernatant. One-step RT-PCR was performed by using a commercially available 1-step RT-PCR kit (Qiagen). Two primers designed on the basis of the hMPV fusion protein gene, 5′-GAGCAAATCCCAGACA-3′and 5′-GAAAACTGCCGCACAACATTTAG-3′, were used as forward and reverse primers, respectively (13). Attempts to isolate virus from homogenate of Nasal turbinate from poults exposed to the 4 genotypes of hMPV was made in LLC-MK2 cells. Tissue sections from nasal turbinates, tracheae, and lungs were stained with hematoxylin and eosin and examined for histopathologic changes. An immunoperoxidase procedure (10) originally developed to detect aMPV antigen was modified to detect hMPV antigen in formalin-fixed nasal turbinate, trachea, and lung tissues by using hMPV B2 polyclonal sera from rabbits. The tissue sections were also tested by using aMPV polyclonal sera from rabbits (10). RESULTS Poults inoculated with any of the 4 hMPV genotypes had unilateral or bilateral nasal discharge which varied from watery to thick mucus. Clinical signs started on day 4 postexposure and stopped. The severity varied between the 4 isolates of hMPV


267

used. Poults inoculated with aMPV-c showed more severe clinical signs and sinus swelling. At day 3 postexposure, RT-PCR detected hMPV viral RNA in the nasal turbinates of poults in each group exposed to hMPV. Control nonexposed birds were negative for hMPV. The aMPV RNA was not detected in unexposed or hMPV-exposed poults. Those infected with aMPV showed viral RNA in nasal turbinate. At days 3 and 5 postexposure, each of the 4 genotypes of hMPV had caused mild to moderate histopathologic lesions in the nasal turbinates of poults. Changes were more severe in those birds inoculated with one of the hMPV strain than with any of the other 3 genotypes. Immunohistochemical observation showed hMPV antigen in the epithelial surface of nasal turbinates of poults in each group inoculated with hMPV on day 3 postexposure. DISCUSSION All 4 genotypes of hMPV tested in this experiment did infect turkeys, as shown by clinical signs, RT-PCR results, immunohistochemical observations and histopathology of nasal turbinates and tracheas of exposed turkeys. The severity of clinical signs and lesions varied with different genotypes, possibly because of differences in titers inoculated, which can potentially influence virus dissemination and pathogenicity. The clinical signs were similar to those of birds experimentally infected with aMPV C (12). The main clinical sign observed in birds infected with aMPV C is watery to thick mucous discharge (12), and our poults exposed to hMPV showed watery to thick mucous nasal discharge. REFERENCES Van den Hoogen BG, de Jong JC, Groen J, Kuiken T, de Groot R, Fouchier RA, et al.

A newly discovered human pneumovirus isolated from young children with respiratory tract disease. Nat Med 2001;7:719–24. [PubMed: 11385510]

Hamelin ME, Boivin G. Human metapneumovirus: a ubiquitous and long-standing respiratory pathogen. Pediatr Infect Dis J 2005;24:S203–7. [PubMed: 16378047]

Hamelin ME, Abed Y, Boivin G. Human metapneumovirus: a new player among respiratory viruses. Clin Infect Dis 2004;38:983–90. [PubMed: 15034830]

Gray GC, Capuano AW, Setterquist SF, Sanchez JL, Neville JS, Olson J, et al. Human metapneumovirus, Peru. Emerg Infect Dis 2006;12:347–50. [PubMed: 16494771]

Boivin G, Abed Y, Pelletier G, Ruel L, Moisan D, Cote S, et al. Virological features and clinical manifestations associated with human metapneumovirus: a new paramyxovirus responsible for acute respiratory-tract infections in all age groups. J Infect Dis 2002;186:1330–4. [PubMed: 12402203]

Agapov E, Sumino KC, Gaudreault-Keener M, Storch GA, Holtzman MJ. Genetic variability of human metapneumovirus infection: evidence of a shift in viral genotype without a change in illness. J Infect Dis 2006;193:396–403. [PubMed: 16388487]

Boivin G, Mackay I, Sloots TP, Madhi S, Freymuth F, Wolf D, et al. Global genetic diversity of human metapneumovirus fusion gene. Emerg Infect Dis 2004;10:1154–7. [PubMed: 15207075]


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Njenga MK, Lwamba HM, Seal BS. Metapneumoviruses in birds and humans. Virus Res 2003;91:163–9. [PubMed: 12573494]

Van den Hoogen BG, Bestebroer TM, Osterhaus AD, Fouchier RA. Analysis of the genomic sequence of a human metapneumovirus. Virology 2002;295:119–32. [PubMed: 12033771]

Jirjis FF, Noll SL, Halvorson DA, Nagaraja KV, Shaw DP. Pathogenesis of avian pneumovirus infection in turkeys. Vet Pathol 2002;39:300–10. [PubMed: 12014494]

Chiang S, Dar AM, Goyal SM, Sheikh MA, Pedersen JC, Panigrahy B, et al. A modified enzymelinked immunosorbent assay for the detection of avian pneumovirus antibodies. J Vet Diagn Invest 2000;12:381–4. [PubMed: 10907873]

Velayudhan BT, McComb BC, Bennett RS, Lopes VC, Shaw DP, Halvorson DA, et al. Emergence of a virulent type C avian metapneumovirus in turkeys in Minnesota. Avian Dis 2005;49:520–6. [PubMed: 16404993]

Falsey AR, Erdman D, Anderson LJ, Walsh EE. Human metapneumovirus infections in young and elderly adults. J Infect Dis 2003;187:785–90. [PubMed: 12599052]


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IDENTIFICATION OF TWO REGIONS WITHIN THE SUBTYPE-A AVIAN METAPNEUMOVIRUS FUSION PROTEIN (AMINO ACIDS 211 - 310 AND 336 –

479) RECOGNIZED BY NEUTRALIZING ANTIBODIES

BROWN PA1, BONCI M2, RICCHIZZI E2, JONES RC1 and NAYLOR CJ1

1Department of Veterinary Pathology, University of Liverpool, Leahurst, Neston, Cheshire, CH64 7TE, UK,

2 Department of Veterinary Public Health and Animal Pathology, University of Bologna, Italy

SUMMARY Six subtype A avian metapneumovirus (AMPV) fusion (F) protein fragments were expressed in E.coli.Virus neutralization tests following adsorption of sera with each F fragment, identified two regions within the F protein that were recognized by neutralizing antibodies. These fragments reduced neutralizing endpoint dilutions of homologous subtype A antiserum by 2.0 and 3.0 log2 dilutions respectively and by 1.0log2 dilution with subtype B antiserum. The regions identified were similar to those previously reported for the F protein of respiratory syncytial virus (RSV). Results suggests that the equivalent regions of the F proteins in subtype A, B and C AMPV may share similar functions and that this similarity may extend to more distantly related viruses including RSV.

INTRODUCTION

Currently, the main approach for control of AMPV infection is through the use of live-attenuated vaccines, supported by subsequent use of killed vaccines in birds requiring longer periods of protection (Buys et al., 1989; Cook et al., 1989; Cook et al., 1989; Williams et al., 1991; Cook et al., 1995; Cook et al., 1996). Although field outcomes have been generally good, the minimal coding sequence changes needed for their reversion to virulence (Naylor and Jones, 1994; Catelli et al., 2006) has been one factor in the quest for improved live vaccines. In order to generate an optimal live AMPV vaccine, it is probable that a range of viral modifications will be needed and some of these are likely to involve changes to key antigenic regions. Unfortunately to date little is known so far about the subtype A AMPV regions important for induction of protective immunity. One study showed the importance of attachment protein gene expression levels (Naylor et al., 2007) and


270

another demonstrated that AMPV fusion (F) protein expressed from a fowlpox vector, was able to induce significant protection against virulent challenge (Qingzhong et al., 1994). A further study also showed that an AMPV which grew in turkeys but did not induce protection against subsequent challenge, became protective once its F protein was exchanged for an F protein from a known protective strain (Naylor, 2006). More generally, the F protein of viruses within the Paramyxoviridae family is recognized as one of the major immunogenic proteins and has been the target of many studies on aspects of immunity and protection. The current study further investigates the immunogenic importance of the F protein by determining regions of that protein likely to be important in inducing immunity. The F protein was expressed in six sections and their abilities to bind to antibodies, known to neutralize AMPV in cell culture, were determined, both for homologous neutralizing antibodies as well as neutralizing antibodies of other AMPV subtypes. This was achieved by observing each fragment’s ability to bind to neutralizing antibodies present in AMPV sera raised during experimental infections by a natural route. The virus used in the tests did not have either SH or G proteins (Naylor et al., 2004) so any neutralization involving viral surface proteins would be due to interactions with F. MATERIALS and METHODS Expression vector development The full length F gene of AMPV, subtype A was amplified in six individual sections (see Figure 1) using PCR. These sections were ligated into the EcoRV restriction enzyme site of plasmid vector pET 30 (Novagen) (See figure 2) and then used to transform E.coli. Positive clones were identified using specific PCR primers. Expression and purification E.coli cells (DE3 PlysS invitrogen) specialized for expressing recombinant proteins were transformed with the F section vectors. Expression cultures were generated and expression induced for three hours. Cell pellets from each culture were collected then lysed using detergents and sonication. Recombinant proteins were captured using metal affinity chromatography that was specific for an engineered region of six Histidine (His) amino acids placed at the amino terminal of each F section (see Figure 3). Virus neutralization (VN) In brief, A VN test was designed to identify potential neutralizing epitopes within the expressed regions of the fusion protein. Neutralization tests were performed in vero cells using an SH-G deletion recombinant (ΔSH-G) derived from subtype-A AMPV (Naylor et al., 2004) and subtype A, B and C antisera raised during infection by a natural route. On two occasions similar neutralization tests were performed following adsorption of the same subtype A, B and C antisera with each of the F fragments and a negative control (p-ve). This control sample was taken from an E.coli clone


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containing unmodified p-ET30 plasmid which had been subjected to the same procedures as the f section vectors. RESULTS At day 4 pi, typical ΔSH-G CPE (Naylor et al., 2004) could be seen in the positive control and at log2 13 and 14 dilutions for subtype A and B antisera and at log2 7 to 14 dilutions for subtype C antisera (Table 1). End points were taken as log2 13 for subtype A and B antisera and log2 7 for subtype C antisera. These end point dilutions provided the basis for the range of dilutions used in tests involving F fragments. Adsorption of the subtype A antiserum with each of the F fragments (Fp1, 2, 3, 4, 5 and 6) resulted in a reduction of neutralizing endpoint from log2 13 to log2 10 with Fp5 and to log2 11 with Fp4 (Table 2). No change in neutralizing endpoints was observed with Fp1, 2, 3 or 6 or p-ve. A reduction in neutralizing endpoints was observed with subtype B anti-serum to log2 12 with Fp5 and Fp4 but again no change was observed with Fp1, 2, 3 or 6 or p-ve (Table 3). No reductions in neutralizing endpoints were observed with subtype C anti-serum with any F fragment or p-ve (Table 4). In all cases the duplicate tests (tests 1 and 2) produced near identical results and can be seen in tables 2 to 4 DISCUSSION This study showed that F sections 4 and 5, which form part of the AMPV F1 ectodomain, were able to neutralize AMPV neutralizing antibodies present in serum raised during experimental infections by a natural route with subtype A and B viruses. This implies that those neutralizing antibodies bound specifically to these F regions and were very likely to have been induced by those regions during the infections in which they were generated. No reductions were observed using F sections 1, 2, 3 or 6. Interestingly, (Lounsbach et al., 1993) and (Werle et al., 1998) using F protein fragments of RSV expressed in E.coli and baculovirus respectively demonstrated neutralizing epitopes between aa residues 190 – 289, a region that overlaps with APV F section 4 in the current study. Moreover, this region in HMPV has been found to behave similarly (Ulbrandt et al., 2008). Other F regions of the AMPV ectodomain were not recognized by neutralizing antibodies but caution must be taken in interpreting those findings. The sections were generated in E.coli and hence were not subject to post translational modification, such as would normally occur when virus grows in its eukaryotic host. Moreover components of neutralizing epitopes might be present which only have effect when combined with other parts of the protein when folded into its native conformation. This is the first report identifying regions within the F protein of subtype A AMPV associated with a protective immune response and it is reasonable to infer that these same regions might stimulate or enhance such a neutralizing response in birds. Immunity to these sections could be facilitated in a number of ways ranging through


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injection of the peptides with suitable adjuvants, perhaps using a prime-boost vaccination strategy as described by (Liman et al., 2007), expression of sections in a vector organism such as fowlpox and DNA vaccination. However, a more likely use of the information about the importance of these sections of F will be in modification of live virus for the purpose of generating candidate live vaccines via reverse genetics. The F protein has already been shown to be important in inducing immunity (Qingzhong et al., 1994) and the comparison of the results of the current study with those from the previous F gene exchange study (Naylor, 2006) indicate that all four F mutations associated with this earlier study occurred between aa residues 295 and 458, and hence all are were within sections 4 and 5 of the current study. This correspondence adds weight to the notion that sections 4 and 5 are indeed influential in inducing effective immunity against virulent AMPV challenge. It further suggests that focus on these regions is likely to be influential in developing live vaccines which induce higher levels of protection than are available in current alternatives. ACKNOWLEDGEMENTS The authors wish to acknowledge the excellent technical help of Ms Jayne Clubbe and Mrs. Carol Savage and Dr Z. Woldehiwet and Dr R. Birtles for technical advice. REFERENCES Buys, S. B., du Preez, J. H. and Els, H. J. (1989). "The isolation and attenuation of a

virus causing rhinotracheitis in turkeys in South Africa." Onderstepoort J Vet Res 56(2): 87-98.

Catelli, E., Cecchinato, M., Savage, C. E., Jones, R. C. and Naylor, C. J. (2006). "Demonstration of loss of attenuation and extended field persistence of a live avian metapneumovirus vaccine." Vaccine.

Cook, J. K., Huggins, M. B., Woods, M. A., Orbell, S. J. and Mockett, A. P. (1995). "Protection provided by a commercially available vaccine against different strains of turkey rhinotracheitis virus." Vet Rec 136(15): 392-3.

Cook, J. K. A., Ellis, M. M., Dolby, C. A., Holmes, H. C., Finney, P. M. and Huggins, M. B. (1989). "A live attenuated turkey rhinotracheitis virus vaccine. I. Stability of the attenuated strain." Avian Pathol 18: 511-522.

Cook, J. K. A., Holmes, H. C., Finney, P. M., Dolby, C. A., Ellis, M. M. and Huggins, M. B. (1989). "A Live Attenuated Turkey Rhinotracheitis Virus-Vaccine .2. The Use of the Attenuated Strain as an Experimental Vaccine." Avian Pathology 18(3): 523-534.

Cook, J. K. A., Orthel, F., Orbell, S., Woods, M. A. and Huggins, M. B. (1996). "An experimental turkey rhinotracheitis (TRT) infection in breeding turkeys and the prevention of its clinical effects using live-attenuated and inactivated TRT vaccines." Avian Pathology 25(2): 231-243.

Liman, M., Peiser, L., Zimmer, G., Propsting, M., Naim, H. Y. and Rautenschlein, S. (2007). "A genetically engineered prime-boost vaccination strategy for oculonasal delivery with poly(D,L-lactic-co-glycolic acid) microparticles against infection of turkeys with avian Metapneumovirus." Vaccine 25(46): 7914-26.


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Lounsbach, G. R., Bourgeois, C., West, W. H., Robinson, J. W., Carter, M. J. and Toms, G. L. (1993). "Binding of neutralizing monoclonal antibodies to regions of the fusion protein of respiratory syncytial virus expressed in Escherichia coli." J Gen Virol 74 ( Pt 12): 2559-65.

Naylor, C. J. (2006). A method of exchanging RNA sequences between protective and unprotective candidate APV vaccines reveals some regions important in stimulating protection. V. International Symposium on Avian Corona and Pneumoviruses and Complicating Pathogens, University of Giessen Rauischholzhausen, Germany, VVB Laufersweiler Verlag.

Naylor, C. J., Brown, P. A., Edworthy, N., Ling, R., Jones, R. C., Savage, C. E. and Easton, A. J. (2004). "Development of a reverse-genetics system for Avian pneumovirus demonstrates that the small hydrophobic (SH) and attachment (G) genes are not essential for virus viability." J Gen Virol 85(Pt 11): 3219-27.

Naylor, C. J. and Jones, R. C. (1994). "Demonstration of a virulent subpopulation in a prototype live attenuated turkey rhinotracheitis vaccine." Vaccine 12(13): 1225-30.

Naylor, C. J., Ling, R., Edworthy, N., Savage, C. E. and Easton, A. J. (2007). "Avian metapneumovirus SH gene end and G protein mutations influence the level of protection of live-vaccine candidates." J Gen Virol 88(Pt 6): 1767-75.

Qingzhong, Y., Barrett, T., Brown, T. D., Cook, J. K., Green, P., Skinner, M. A. and Cavanagh, D. (1994). "Protection against turkey rhinotracheitis pneumovirus (TRTV) induced by a fowlpox virus recombinant expressing the TRTV fusion glycoprotein (F)." Vaccine 12(6): 569-73.

Ulbrandt, N. D., Ji, H., Patel, N. K., Barnes, A. S., Wilson, S., Kiener, P. A., Suzich, J. and McCarthy, M. P. (2008). "Identification of antibody neutralization epitopes on the fusion protein of human metapneumovirus." J Gen Virol 89(Pt 12): 3113-8.

Werle, B., Bourgeois, C., Alexandre, A., Massonneau, V. and Pothier, P. (1998). "Immune response to baculovirus expressed protein fragment amino acids 190-289 of respiratory syncytial virus (RSV) fusion protein." Vaccine 16(11-12): 1127-30.

Williams, R. A., Savage, C. E. and Jones, R. C. (1991). "Development of a Live Attenuated Vaccine against Turkey Rhinotracheitis." Avian Pathology 20(1): 45-55.


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Metal affinity capture of His tagged recombinant protein Table 1. Standard virus neutralization test using the ΔSH-G deletion recombinant subtype A AMPV (Naylor et al., 2004) with subtype A, B and C positive antisera.

log2 dilutions

Antisera subtype

5 6 7 8 9 10 11 12 13 14 Positive control

Cells only

A 0 0 0 0 0 0 0 0 1a 2 2 0 B 0 0 0 0 0 0 0 0 2 2 2 0

0 0 2 2 2 2 C

2 2

2 2

2 0

a: number of wells from 2 showing CPE


276

Table 2. Neutralization of ΔSH-G deletion recombinant subtype A AMPV with subtype A antisera, following adsorption with subtype A F fragments.

log2 dilutions

Test 1 7 8 9 10 11 12 13 14

Fp 1 0 0 0 0 0 1a 4 4

Fp 2 0 0 0 0 0 0 4 4 Fp 3 0 0 0 0 0 0 4 4 Fp 4 0 0 0 0 4 4 4 4 Fp 5 0 0 0 4 4 4 4 4 Fp 6 0 0 0 0 0 0 2 4 p-ve 0 0 0 0 0 0 4 4 Test 2 Fp 1 0 0 0 0 0 0 4a 4 Fp 2 0 0 0 0 0 1 3 4 Fp 3 0 0 0 0 0 0 3 4 Fp 4 0 0 0 0 3 4 4 4 Fp 5 0 0 0 3 4 4 4 4 Fp 6 0 0 0 0 0 0 4 4 p-ve 0 0 0 0 0 0 4 4

a: number of wells from 4 showing CPE Table 3. Neutralization of ΔSH-G deletion recombinant subtype A AMPV with subtype B antisera, following adsorption with subtype A F fragments.

log2 dilutions

Test 1 7 8 9 10 11 12 13 14

Fp 1 0 0 0 0 0 0 3a 4

Fp 2 0 0 0 0 0 0 3 4 Fp 3 0 0 0 0 0 0 4 4 Fp 4 0 0 0 0 0 3 4 4 Fp 5 0 0 0 0 0 4 4 4 Fp 6 0 0 0 0 0 0 4 4 p-ve 0 0 0 0 0 0 4 4 Test 2 Fp 1 0 0 0 0 0 0 4a

4 Fp 2 0 0 0 0 0 0 4 4 Fp 3 0 0 0 0 0 0 3 4 Fp 4 0 0 0 0 0 4 4 4 Fp 5 0 0 0 0 0 4 4 4 Fp 6 0 0 0 0 0 0 4 4 p-ve 0 0 0 0 0 0 4 4



277

Table 4. Neutralization of ΔSH-G deletion recombinant subtype A AMPV with subtype C antisera, following adsorption with subtype A F fragments.

log2 dilutions

Test 1 5 6 7 8 9 10 11 12

Fp 1 0 0 3a 4 4 4 3 4 Fp 2 0 0 3 3 4 4 3 4 Fp 3 0 0 4 4 4 4 4 4 Fp 4 0 0 3 3 4 4 4 4 Fp 5 0 0 3 4 4 4 4 4 Fp 6 0 0 3 4 4 4 4 4 p-ve 0 0 3 4 4 4 4 4

Test 2

Fp 1 0 0 4a 4 4 4 4 4 Fp 2 0 0 3 3 4 4 4 4 Fp 3 0 0 4 4 4 3 4 4 Fp 4 0 0 3 4 4 4 4 4 Fp 5 0 0 2 4 3 4 4 4 Fp 6 0 0 3 4 4 4 4 4 p-ve 0 0 4 4 4 4 4 4



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AVIAN METAPNEUMOVIRUSES IN ITALY: EVIDENCE OF ATTACHMENT PROTEIN EVOLUTION COINCIDENT WITH MASS LIVE VACCINE

INTRODUCTION

CECCHINATO M1, CATELLI E2, LUPINI C2, RICCHIZZI E2, CLUBBE J3 and NAYLOR CJ3

1 Dipartimento di Sanità Pubblica, Patologia Comparata e Igiene Veterinaria Faculty of Veterinary Medicine, University of Padua, Viale dell’Università, 16, 35020

Legnaro (PD), Italy; 2 Dipartimento di Sanità Pubblica Veterinaria e Patologia Animale, Faculty of

Veterinary Medicine, University of Bologna, Via Tolara di Sopra, 50, 40064 Ozzano Emilia (BO), Italy;

3 Department of Veterinary Pathology, University of Liverpool, Jordan Building, Leahurst, CH64 7TE Neston, United Kingdom;

SUMMARY Avian metapneumoviruses (AMPV) of subtype B dominate over other subtypes on Italian poultry farms in northern Italy. AMPVs from the Veneto region of Italy between 1987 and 2007 were sequenced in their attachment (G) and fusion (F) protein genes, together with other subtype B AMPVs from other parts of Western Europe, collected prior to 1994. All viruses in the survey had very similar predicted F sequences whereas the predicted G protein sequences found in Italy from 2001 were distinctly different from those found in pre 1994 viruses. Nonetheless pre 1994 Italian AMPVs were more similar to post 2000 AMPVs than other pre 1994 viruses from other parts of Europe, thereby showing that the later viruses had probably evolved from early Italian viruses. The occurrence of the later variants followed introduction of the mass administration with a single subtype B vaccine and its G protein sequence placed it clearly in the pre 1994 cluster. The possibility of the vaccine having driven field virus evolution in order to evade immune pressure cannot be excluded. INTRODUCTION The vast majority of Avian Metapneumoviruses (AMPVs) detected in Italy have proved to be of subtype B (Catelli et al., 2004a; Catelli et al., 2004b) but the genetic relatedness between detected viruses has not been previously investigated. An early sequence study in 1994 (Juhasz & Easton, 1994) showed that G genes from Italy, Hungary and Spain were very similar and a further study of a UK virus in 1994 again


279

showed that its G protein was very similar (Cavanagh et al., 1999). The current study investigates the genetic relatedness of F and G genes of subtype B AMPVs and especially focuses on those in circulation in the Veneto region of northern Italy from 1987 to 2007. The area has an unusually high density of poultry farms and the majority of these have been operated by a single commercial company. Disease typical of AMPV infection started to become a problem in turkeys in the 1990s and this led to the use of a live subtype B vaccine in an experimental manner on perceived problem farms. The vaccine selected was the only subtype B live vaccine available at that time and this remained the case for the duration of the investigation. For several years in the 1990s a range of vaccine doses were applied to turkeys by either eyedrop or spray so as to determine best practise for the particular environment. In late 2001, one-day-old live vaccination became the standard for all turkeys in the region. The study determines and compares the sequences of the G and F proteins present in viruses over the period and in discussing the findings, considers possible effects of live vaccination. MATERIAL and METHODS Viruses. Reference subtype B viruses were assembled from a range of European countries for the period between 1986 and 1994. These had generally undergone a limited number of passages in chicken tracheal organ cultures or other culture systems. These included UK 8-94, UK 11/94, Italy 16/91, Italy 2119, France 147, Spain 149, Netherlands 6726/90 and France 38/86. In Italy between 1987 and 2007 seven AMPVs (Table 1) were characterized as subtype B by the RT-PCR method of Cavanagh et al. (1999). Four of the seven viruses were recovered from turkeys with TRT while the remainder derived from broiler chicken flocks displaying mild respiratory disease. Six AMPVs were isolated and grown in chicken embryo tracheal organ cultures (TOCs) (Cook et al., 1976). In one case (IT/Ck/1348-01/07) virus was characterized using RNA extracted from dry swabs as reported below. A single commercial subtype B vaccine was widely used in the area after 2000 and this was included in the study.

RNA extraction and RT-PCR RNA was extracted using a commercial kit (QIAamp viral mini kit, Qiagen). In order to sequence the entire F and G protein genes, one reverse transcription and 3 overlapping independent PCRs were performed for each gene. Oligonucleotide primers specific for subtype B AMPVs within F and G genes were based on previously published sequences while primers in flanking regions were based on either previously determined partial sequences or comparisons between subtype A AMPVs and respiratory syncytial virus sequences. The primer sequences and approximate genome positions are reported in Table 2. Extracted RNA was used in a RT reaction (Improm-II™ Reverse Trascriptase, Promega, under recommended conditions) and the cDNA was amplified by PCR (GoTaq® DNA Polymerase, Promega, under recommended conditions).


280

DNA sequencing and phylogenetic analysis. Amplified DNA products were purified using the Wizard® SV Gel and PCR Clean-Up System (Promega). Sequencing was performed at BMR Genomics (Padua, Italy) with the 3730xl DNA Analyzer (Applied Biosystems, USA). Nucleotide sequences were edited and assembled using Bioedit software then aligned against previously published complete sequences of the B subtype using Clustal X. Phylogenetic analysis was carried out under distance criterion, with neighbour-joining as algorithm, using MEGA4 software. Bootstrap values were obtained with 1,000 replicates. Branches with bootstrapping values >70 were considered significant, corresponding to a confidence interval >95%. RESULTS Amplicons of the expected sizes were obtained from all samples with the exception of IT/Ck/1348-01/07 where only the G gene was amplified. When comparing sequences, predicted amino acids were chosen in preference to nucleotides because these would be expected to more closely relate to the antigenicity of each respective protein. The F protein sequences were highly conserved. The average percentage amino acid identity differences when comparing all pre 1994 viruses to all post 2000 viruses was 0.77%, while comparing between pre 1994 viruses or post 2000 viruses gave 0.59% and 0.32% respectively. However for the G sequences identity differences were higher. Average percentage amino acid identity differences when comparing all pre 1994 viruses to all post 2000 viruses was 4.75% while comparing between pre 1994 viruses or post 2000 viruses gave 2.26% and 1.00% respectively. Phylogenetic dendrograms (Figure 1) showed that pre 1994 viruses clustered closely together some distance from the other main cluster formed by the post 2000 Italian viruses. While the pre 1994 Italian viruses clustered with the other pre 1994 viruses, these were genetically closer to the post 2000 Italian viruses than the remainder of the pre 1994 cluster. At the detailed amino acids sequence level (Figure 2) this is well illustrated by the exclusive presence of a serine residue at position 387 in all Italian viruses. A further mutation exclusive to the Italian viruses was also found beyond the G gene in the polymerase (L) gene start but the L gene was not included in the phylogenetic analysis. Numerous coding mutations were seen and all of these either affected charged amino acids (K, R, D, E and H) or those amino acids able to accept O-linked glycan during post translational modification (S and T). For the latest viruses detected, this affected 6 charged (aa 149, 173, 265, 273, 274 and 383) and 15 glycan associated (aa 18, 92, 145, 151, 179, 193, 228, 237, 273, 276, 326, 329, 362, 365 and 387) residues. DISCUSSION The study shows that over a 20 year period attachment proteins in AMPVs have undergone major amino acid substitutions while F protein sequences remained largely unchanged. The phyolgenic analysis indicates that while the early pre 1994 Italian viruses were highly similar to the other pre 1994 European viruses investigated, they are also highly likely to have been the progenitor(s) for the viruses seen in Italy after 2000. These later viruses displaced progenitor viruses over a period of 20 years on farms in the Veneto region, and many detections were made


281

on farms separated by distances of up to 40km. The precise detail of mutations accumulated over the period suggests a sequence of progressive changes to a relatively homogeneous pre 1994 progenitor virus pool rather than a series of totally unrelated changes at different locations to the common progenitor. A precise mechanism for these changes cannot be fully described from the data available. At one extremity, changes might have been due to unforced mutation of viruses in poultry free of AMPV and with no prior immunity to the virus. Alternatively changes might have occurred due to immune pressure induced either by exposure to circulating field viruses or that induced by the subtype B vaccine in common use in the Veneto region since the 1990s. Unfortunately because no sequence data are available for the critical period between 1994 and 2000, it is not possible to say whether mutation rates accelerated after vaccine introduction. REFERENCES Catelli E., Cecchinato M., Delogu M., De Matteo P., Ortali G., Franciosi C., De Marco

M.A., Naylor C.J., 2004a. Avian Pneumovirus infection in turkey and boiler farms in Italy: a virological, molecular and serological field survey. Italian Journal of Animal Science, 3 (3): 286-292.

Catelli E., Cecchinato M., Ortali G., De Matteo P., Savage C.E., Jones R.C., Naylor C.J., 2004b. Avian Pneumovirus in Italy. Proceedings of the 4th International Symposium on Avian Coronavirus and Pneumovirus Infections; 2004 June 20-24, Rauischholzhausen, Germany. WB Laufersweiler Verlag, Wettenberg, Germany; 2004. p. 275-281.

Cavanagh D., Mawditt K., Britton P., Naylor C.J., 1999. Longitudinal field studies of infectius bronchitis virus and avian pneumovirus in broiler using type-specific Polymerase chain reactions. Avian Pathology, 28: 593-605.

Cook J.K.A., Darbyshire J.H., Peter R.W., 1976. The use of chicken tracheal organ cultures for the isolation and assay of avian infectious bronchitis virus. Archives of Virology, 50: 109-118.

Juhasz K. & Easton A.J., 1994. Extensive sequence variation in the attachment (G) protein gene of avian pneumovirus: evidence for two distinct subgroups. Journal of General Virology, 75: 2873-2880.


282

Table 1. Details of year of collection and species of host of Italian AMPVs used in the study

Strain Year of detection Species IT/Ty/Vr240/87 1987 Turkey IT/Ty/2a/01 2001 Turkey IT/Ck/33a/02 2002 Chicken IT/Ck/34a/02 2002 Chicken IT/Ty/129-18/04 2004 Turkey IT/Ty/205-16/04 2004 Turkey IT/Ck/1348-01/07 2007 Chicken

Table 2. Sequences of oligonucleotide primers used for reverse transcription copying and PCR amplification of AMPV F and G genes.

Primer name and sequences (5’-3’) PCR

Gene Positiona Product size (bp)

MB1+ b GAGGACTAGGTATGTCCTGAAG M 2852 F1- CCTGCACTATCAGAGAATTG F 3578

726

F5+ CCTCGAAATAGGGAATGTTGAGAAC F 3090 F3- CCTATGGAGCAACTTACAC F 4121

1031

F7+ GACACCCTGTCAGTATGGATC F 4043 M2:3- CATGATCAGGCCAGGACCAATAATTAT M2 4686

643

SH1+ b GCTTTGATCTTCCTTGTTGC SH 5507 G6- CTGACAAATTGGTCCTGATT G 6460

953

Gstart+ CAAGTATCCAGATGGGGTC G 5914 GB1- GTACAGCACCACTCAATCAG G 7159

1245

G15+ GACTGCACACTATCTGATCCAG G 7000 Lb6- CAGTTGCTAACCCAGACACAC L 7926

926

a anti-genome sense b also used for reverse transcription


283

88

78

93

75

100

88 74

Figure 1. Radial phylogram of 19 subtype B AMPVs. The phylogram is based on predicted attachment protein sequences using MEGA4 software. Branches lengths are proportional to the estimated genetic differences. The bar represents the number of nucleotide substitutions per site. Only bootstrapping values > 70 are reported.


284

Figure 2. Deduced amino acid sequences of the complete G proteins of European AMPVs collected between 1986 and 2007. The oldest virus, France-38/86, is shown in full and other residues are only specified where different to this sequence.

10 20 30 40 50 60 70 80 90 100 110 120 ....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....| France 38/86 GTSIQMGSELYIIEGVSSSEIVLKQVLRRSKKILLGLVLSALGLTLTSTIVISICISVEQVKLRQCVDTYWAENGSLHPGQSTENTSTRDKTTTKDPRRLQATGAGKFESCGYVQVVDGD NL/6726/90 ........................R..................................................................................L............ UK 8-94 ..............................Q......................................................................................... UK 11/94 ...........T............................................................................................................ Spain 872S ..............................Q....R.....................................................G.............................. B vaccine ...................................R.....................................................G.............................. France 147 .........................................................................................G.............................. Spain 149 ...................................R.....................................................G.............................. Hungary 657/4/89 ..............................Q..........................................................G.............................. Israel 1708/02 ----X...................................I............................................................................... Italy 2119 ..............................Q......................................................................................... IT/Ty/Vr240/87 ........................................................................................................................ Italy 16/91 ........................................................................................................................ IT/Ty/2a/01 ...........................................................................................I............................ IT/Ck/33a/02 ...........................................................................................I............................ IT/Ck/34a/02 ...........................................................................................I............................ IT/Ty/129-18/04 ...........................................................................................I............................ IT/Ty/205-16/04 ...................................E.......................................................I............................ IT/Ck/1348-01/07 .................P.........................................................................I............................ 130 140 150 160 170 180 190 200 210 220 230 240 ....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....| France 38/86 MHDRSYAVLGGVDCLGLLALCESGPICQGDTWSEDGNFCRCTFSSHGVSCCKKPNSKATTAQRNSKPANSKSTPPVHSDRASKEHNPSQGEQPRRGPTSSKTTIASTPSTEDTAKPTISK NL/6726/90 ....................................I................................................................................... UK 8-94 ..................................G...................K..................................................G.............. UK 11/94 ...H..............................G...................K........................................E.........G.............. Spain 872S ..........................................L...........K................................................................. B vaccine ..........................................L...........K................................................................. France 147 ......................................................K................................................................. Spain 149 ......................................................K................................................................. Hungary 657/4/89 ......................................................K................................................................. Israel 1708/02 ...........................R.....K....................K................................................................. Italy 2119 ......................................................K................................................................. IT/Ty/Vr240/87 ......................................................K................................................................. Italy 16/91 ......................................................K................................................................. IT/Ty/2a/01 ........................S...R.A.......................K.................I......................R....................M... IT/Ck/33a/02 ........................S...R.A.......................K.................I...........................................M... IT/Ck/34a/02 ........................S...R.A.......................K.................I...........................................M... IT/Ty/129-18/04 ........................S...R.A.....................E.K...I.............I...........................................M... IT/Ty/205-16/04 ........................S...R.A.....................E.K...I.............I...........................................M... IT/Ck/1348-01/07 ........................S...R.A.....................E.K.................I..................................S........M... 250 260 270 280 290 300 310 320 330 340 350 360 ....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....|....| France 38/86 PKLTIRPSQRGPSGSTKAASSTPSHKTNTRGTSKTTDQRPRTGPTPGRPRQTHSTATPPPTTPIHKGRAPTPKPTTDLKVNPREGSTSPTAIQKNPTTQSNLVDCTLSDPDEPQRICYQV NL/6726/90 ..............N........................................................................................N................ UK 8-94 ........K....S................................E......................................G...............P.................. UK 11/94 ........K....S.........N......................E......................................G...............P...........P...... Spain 872S ..............................................E......................................................................... B vaccine ..............................................E......................................................................... France 147 ..............................................E......................................................................... Spain 149 ..............................................E......................................................................... Hungary 657/4/89 ..............................................E......................................................................... Israel 1708/02 ........K.............S....D..........K.......V.................Q............P.......G.......T.......................... Italy 2119 ........................................................................................................................ IT/Ty/Vr240/87 .........................................I.............................................................................. Italy 16/91 .................................................................................................I...................... IT/Ty/2a/01 ........................Y.......RE.I.................................................N.................................. IT/Ck/33a/02 ........................Y.......RE.I.............................................L...N.................................. IT/Ck/34a/02 ........................Y.......RE.I.................................................N.................................. IT/Ty/129-18/04 ........................Y.......RE.I.................................................N..S............................... IT/Ty/205-16/04 ........................Y.......RE.I.................................................N..S............................... IT/Ck/1348-01/07 ........................Y.......RE.I.................................................N.................................. 370 380 390 400 410 420 ....|....|....|....|....|....|....|....|....|....|....|....| France 38/86 GTYNPSQSGTCNIEVPKCSTYGHACMATLYDTPFNCWRRTRRCICDSGGELIEWCCTSQ* NL/6726/90 ...........................................................* UK 8-94 ...........................................................* UK 11/94 ......................Y....................................* Spain 872S ...........................................................* B vaccine ...........................................................* France 147 ...........................................................* Spain 149 ...........................................................* Hungary 657/4/89 ...........................................................* Israel 1708/02 .......L..............Y....................................* Italy 2119 ..........................S................................* IT/Ty/Vr240/87 ..........................S................................* Italy 16/91 ..........................S................................* IT/Ty/2a/01 ......................Y...S................................* IT/Ck/33a/02 ......................Y...S................................* IT/Ck/34a/02 ......................Y...S................................* IT/Ty/129-18/04 ......................Y...S................................* IT/Ty/205-16/04 ......................Y...S................................* IT/Ck/1348-01/07 .A..S.................Y...S................................*


285

AN INVESTIGATION INTO MOLECULAR DIFFERENCES BETWEEN AVIAN METAPNEUMOVIRUSES (AMPVS) OF CHICKEN AND TURKEY ORIGIN

CLUBBE J, JONES RC1, LE GALLUDEC H2 and NAYLOR CJ1

1University of Liverpool, Department of Veterinary Pathology, Leahurst, Neston, South Wirral, CH64 9PL, UK and 2Fort Dodge Animal Health.

SUMMARY Since the emergence of avian metapnuemoviruses (AMPVs) a range of live attenuated vaccines derived from chicken and turkey isolates have been developed in a bid to control the disease. It has been hypothesised that AMPV vaccines will be more efficacious in the species from which the progenitor virus was derived, however protection studies have proven difficult. A more fundamental approach was taken in this study, comparing chicken and turkey derived AMPVs at their underlying genetic level to identify if species-specific regions exist. A full length RT-PCR and sequencing system was developed for subtype A and B viruses. Complete nucleotide sequences from a range of subtype A and B European field strains from both species, including commercially available vaccines, were aligned and compared. Furthermore, in an attempt to encourage viruses to evolve towards possible species specificity, turkey derived viruses of known sequence were multiply passaged in chicken tracheal organ cultures (TOC), then re-sequenced. While numerous sequence differences between viruses were identified, none was specific for the host species. INTRODUCTION Avian metapneumovirus (AMPV) is an important respiratory pathogen affecting turkeys and chickens worldwide. Since its emergence in the 1980s, a range of live attenuated subtype A and B vaccines derived from both species have been developed in a bid to control the disease. Studies have shown that subtype A and B vaccines offer good cross protection against challenge with either subtype (Cook et al., 1995; Cook et al., 2000; Eterradossi et al., 1995; Toquin et al., 1996) and that these vaccines also confer protection against subtype C (Cook et al., 1999). Commercially, species-specific vaccines have been marketed in the belief that they are more efficacious when used in the species from which the progenitor field virus was derived. In support of this, some experimental studies have indicated


286

contrasting outcomes following infection with chicken or turkey-derived AMPVs in terms of pathogenesis, susceptibility and protection (Buys et al., 1989a; b; Cook et al., 1993). The lack of a fully reproducible chicken challenge model means such clinical differences can be difficult to reproduce and other virus related variables not related to species of origin such as virulence, tropism or the culture system are likely to play a more significant role. To clarify these issues, this study compared chicken and turkey-derived subtype A and B AMPVs at their underlying genetic level. RNA and protein sequences of complete viruses were compared to see whether species specific regions exist. An RT-PCR and sequencing system was used to obtain full length nucleotide sequence from a range of isolates, and the amino acid sequences of proteins were deduced and compared. Comparisons were also made of non-coding regions as mutations here have been shown to alter viral properties (Naylor et al., 2007). Furthermore, in an attempt to encourage mutation towards possible species specificity, turkey-derived AMPVs of known sequence were each passaged ten times in chicken TOC, re-sequenced and compared to the original. MATERIALS and METHODS Virus strains The following European field strains were isolated between 1986 and 2004. All isolates had undergone a small number of passages in TOC. Numbers in superscript refer to the numbers in tables 1-8 for identification of samples: Chicken derived subtype A: CP1 (UK) Chicken derived subtype B: 38/91/OC4 (France)1, 27/91/OC4 (Holland)2, 149/90 (Spain)4

Turkey derived subtype A: 3B (UK), LTZ (Germany), CVL (UK) Turkey derived subtype B: VR 240 (Italy)2, 205-16(Italy)3, 16/91/OC4 (Italy)4, 147/86/OC5 (France)5

Vaccines Live-attenuated freeze-dried commercial vaccines were reconstituted with 2ml of sterile de-ionised water. Turkey subtype A: Poulvac TRT (Fort Dodge), Nobilis TRT, (Intervet), Turkadin (Pitman-Moore) Turkey subtype B: Aviffa -RTI (Merial)1

Chicken subtype B: Nobilis Rhino CV (Intervet)3, Nemovac (Merial)5

RNA extraction Viral RNA was extracted from TOC fluid using QIAamp Viral RNA kit (Qiagen) according to manufacturer’s instructions.


287

RT-PCR Extracted RNA was reverse transcribed into 3 and 4 overlapping sections for subtypes A and B respectively. Subsequent cDNA amplification using subtype specific primers generated overlapping regions of approx 2kb throughout the whole genome. Sequencing and analysis DNA PCR products were prepared for sequencing as recommended by Cogenics (UK). Sequences were analysed using Chromas (Version 1.45) and Bioedit Sequence Alignment Editor (Version 7.0.9.0). TOC Embryonated SPF chicken eggs (Lohmann Animal Health, Cuxhaven, Germany) were used to prepare TOCs from 19 day old embryos as previously reported (Cook et al., 1976). Viable rings were selected after 24 hours, inoculated with turkey derived AMPV and incubated at 370C. At cilliostasis, or after seven days, rings were freeze thawed and centrifuged. Supernatant was used to inoculate new TOCs. RESULTS Comparisons of chicken and turkey subtype B AMPVs identified random individual sequence differences between tested isolates, with the highest number occurring in the G protein. However, the genetics of the two species were highly similar and in the five chickens and five turkey-derived protein sequences compared. No consistent regions associated with either host were identified. This also applied to sequence of the non-coding regions. The amino acid sequences for each protein of the chicken and turkey isolates were identical apart from the differences shown in tables 1-8. Due to insufficient numbers of chicken derived AMPVs, a full comparison study for subtype A could not be performed. Just one available chicken isolate (CP1) was compared to six turkey isolates. However, CP1 showed only 11 amino acid differences in this comparison. Passage of turkeys AMPVs in chicken TOC did not produce any species specific sequence changes. However it was noted that mutations occurred in all three passaged vaccines but only in one of the four field strains (Table 9). DISCUSSION In this study, full length sequences of chicken and turkey derived AMPVs were aligned and compared to highlight possible regions of species specificity. Comparisons were made of nucleotide and coded protein sequence plus non coding regions. While numerous individual sequence differences between viruses were observed, no consistent changes related to either host were found with subtype B. Due to the close relationship of subtype A and B viruses it is reasonable to infer that a full comparison of subtype A AMPVs would reveal a similar outcome but attempts to acquire chicken subtype A isolates were unsuccessful.


288

Lack of subtype A detections compared to B may be because it is less prominent in chickens or because isolation is more difficult. A study comparing pathogenesis of both subtypes in broilers suggested subtype B may be more invasive and observed clinical signs in more birds, a wider tissue distribution pattern and longer persistence than subtype A (Aung et al., 2008). However it is difficult to know what part other differences such as virulence may have played. Passage of turkey AMPVs in chicken TOC was performed to investigate the potential of turkey AMPVs to acquire chicken AMPV characteristics, but no species specific sequences were seen. In contrast, it is interesting to note that all three cell culture attenuated vaccine strains mutated over the course of ten TOC passages compared with only one of the four field strains which were derived from respiratory cells. This suggests that virus is more influenced by the tissue of the culture system than whether the host is chicken or turkey. In conclusion, the rational for vaccine species matching appears unfounded because virus differences between chickens and turkeys appear not to exist. There remains the highly unlikely but theoretical possibility that certain combinations of apparently unrelated mutations might cause some species specificity but nonetheless factors such as vaccine subtype and level of attenuation are likely to be of greater importance. REFERENCES Aung, Y. H., Liman, M., Neumann, U. & Rautenschlein, S. (2008). Reproducibility of

swollen sinuses in broilers by experimental infection with avian metapneumovirus subtypes A and B of turkey origin and their comparative pathogenesis. Avian Pathol 37, 65-74.

Buys, S. B., du Preez, J. H. & Els, H. J. (1989a). The isolation and attenuation of a virus causing rhinotracheitis in turkeys in South Africa. Onderstepoort J Vet Res 56, 87-98.

Buys, S. B., du Preez, J. H. & Els, H. J. (1989b). Swollen head syndrome in chickens: a preliminary report on the isolation of a possible aetiological agent. J S Afr Vet Assoc 60, 221-222.

Cook, J. K., Huggins, M. B., Orbell, S. J. & Senne, D. A. (1999). Preliminary antigenic characterization of an avian pneumovirus isolated from commercial turkeys in Colorado, USA. Avian Pathology 28, 607-617.

Cook, J. K., Darbyshire, J. H. & Peters, R. W. (1976). The use of chicken tracheal organ cultures for the isolation and assay of avian infectious bronchitis virus. Arch Virol 50, 109-118.

Cook, J. K., Huggins, M. B., Woods, M. A., Orbell, S. J. & Mockett, A. P. (1995). Protection provided by a commercially available vaccine against different strains of turkey rhinotracheitis virus. Vet Rec 136, 392-393.

Cook, J. K., Kinloch, S. & Ellis, M. M. (1993). In vitro and in vivo studies in chickens and turkeys on strains of turkey rhinotracheitis virus isolated from the two species Avian Pathology 22, 157-170.


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Cook, J. K. A., Chester, J., Orthel, F., Woods, M. A., Orbell, S. J., Baxendale, W. & Huggins, M. B. (2000). Avian pneumovirus infection of laying hens: experimental studies. Avian Pathol 29, 545-556.

Eterradossi, N., Toquin, D., Guittet, M. & Bennejean, G. (1995). Evaluation of different turkey rhinotracheitis viruses used as antigens for serological testing following live vaccination and challenge. Veterinary Medicine 42, 175-186.

Naylor, C. J., Ling, R., Edworthy, N., Savage, C. E. & Easton, A. J. (2007). Avian metapneumovirus SH gene end and G protein mutations influence the level of protection of live-vaccine candidates. J Gen Virol 88, 1767-1775.

Toquin, D., Eterradossi, N. & Guittet, M. (1996). Use of a related ELISA antigen for efficient TRT serological testing following live vaccination. Vet Rec 139, 71-72.

ACKNOWLEDGMENTS This work was supported by Fort Dodge Animal Health.


290

N P M

aa position 107 141 145 284 389 227 162 179 217 250 Chicken 1 R A K E K A A A G V Chicken 2 R A K E K A A A G V Chicken 3 R A K E K A S A G V Chicken 4 R A – E E A A S G V Chicken 5 Q A K K K A A A S V Turkey 1 R A K E E A A S G V Turkey 2 R A K E K A A A G I Turkey 3 R A K E K T A A G V Turkey 4 R V K E K A A A G V Turkey 5 R A K E K A A A G V Table 1. Amino acid differences between chicken and turkey AMPV strains in the nucleocapsid protein (N), phosphoprotein (P) and matrix protein (M)

F

aa position 89 149 178 188 253 296 323 499 506

522 Chicken 1 I S N R R R K A I S Chicken 2 I S N R R R E V I S Chicken 3 I S N R K R E V I P Chicken 4 F S N R R R E V V S Chicken 5 I S K R R R K A I S Turkey 1 F S N R R R E V V S Turkey 2 I S N R R R E V I S Turkey 3 I S N K R K E V I S Turkey 4 I S N R R R E V I S Turkey 5 I T N R R R E V V S Table 2. Amino acid differences between chicken and turkey AMPV strains in the fusion protein (F)


291

M2:1 M2:2

aa position 84 182 184

10 12 15 19 64 65

66 Chicken 1 E G G V A R L E F V Chicken 2 E G S V A R L E F V Chicken 3 E D S M A K S E F V Chicken 4 E G S V A R L E F V Chicken 5 E G G V A R L L N L Turkey 1 E G S V A R L E F V Turkey 2 E G S V A R L E F V Turkey 3 D G S V V R L E F V Turkey 4 E G S V A R L E F V Turkey 5 E G S V A R L E F V Table 3. Amino acid differences between chicken and turkey AMPV strains in the M2:1 and M2:2 proteins

SH

aa position 27 84 85 87 99 105 112 113 144 157 175

Chicken 1 R N Q H T N Q R I N N Chicken 2 R N Q H T N R R I N N Chicken 3 R N H Y T D Q R I N N Chicken 4 R N Q Y T D Q R I N N Chicken 5 R N Q H T N Q R I N N Turkey 1 R N Q Y T D Q R I N N Turkey 2 R N Q Y T N Q R I N N Turkey 3 Q S Q Y A N Q K I S S Turkey 4 R N Q Y T N Q R I N N Turkey 5 R N Q Y T D Q R V N N Table 4. Amino acid differences between chicken and turkey AMPV strains in the small hydrophobic protein (SH)


292

G

aa position 7 20 31 85 87 103 106 119 146 150 152 168 170 174 188 211 221 232

Chicken 1 I Q G D T F C R T D N K N T T G S T Chicken 2 I R G D T L C R T D I K N T T G S T Chicken 3 T Q G D T F C H T G N K K T T E G T Chicken 4 I Q R G T – – R T D N K K T T G S T Chicken 5 I Q G D T F C R T D N K N T T G S T Turkey 1 I Q R G T F C R T D N K K T T G S T Turkey 2 I Q G D T F C R T D N K K T T G S T Turkey 3 I Q E D I F C R A D N E K I I G S M Turkey 4 I Q G D T F C R T D N K K T T G S T Turkey 5 I Q G G T F C R T D N K K T T G S T Table 5. Amino acid differences between chicken and turkey AMPV strains in the glycoprotein (G)

G

aa position 244 249 250 259 260 268 269 271 277 282 321 324 329 333 337 339 349 378

Chicken 1 Q G S S H S K T T G S P K T L D Q H Chicken 2 Q G N S H S K T T G S P K T L N Q H Chicken 3 K S S N H S K T T E G P K T P D P Y Chicken 4 Q G S S H S K T T E S P K T L D Q H Chicken 5 Q G S S H S K T T G S P N T L D Q H Turkey 1 Q G S S H S K T T E S P K T L D Q H Turkey 2 Q G S S H S K T I G S P K T L D Q H Turkey 3 Q G S S Y R E I T G N S K T L D Q Y Turkey 4 Q G S S H S K T T G S P K I L D Q H Turkey 5 Q G S S H S K T T E S P K T L D Q H Table 6. Amino acid differences between chicken and turkey AMPV strains in the glycoprotein (G)


293

L

aa position 62 72 157 312 426 495 738 877 895 1174 1335 1355 1365 1468 1472

Chicken 1 G N V Y G P L R K K M V A S G Chicken 2 G S V Y G P L R K K M V A S G Chicken 3 S N I H S P L R K K V V A G G Chicken 4 G N V Y G P Q R Q K V – A S G Chicken 5 G N V Y G P L R K K M V A S G Turkey 1 G N V Y G P Q R Q K V V A S G Turkey 2 G N V Y G P Q R K K V V A S G Turkey 3 G N V Y G L Q K K R M V A S R Turkey 4 G N V Y G P Q R K K V V T S G Turkey 5 G N V Y G P Q R Q K V V A S G Table 7. Amino acid differences between chicken and turkey AMPV strains in the polymerase protein (L)

L aa position 1475 1476 1493 1530 1562 1606 1637 1644 1650 1676 1741 1874 1888 1927

Chicken 1 D I R E T T T G Y R I A I G Chicken 2 D I C G T A T G Y R I A I G Chicken 3 N I R G T T T G Y R I A I S Chicken 4 D I R G T T T G H R I T I S Chicken 5 D I R E T T T G Y R I A I G Turkey 1 D I R G T T T G H R I T I S Turkey 2 D I R G T T T R Y R I A I G Turkey 3 D V R G I T A G Y K V A M G Turkey 4 D I R G T T T G Y R I A I G Turkey 5 D I R G T T T G H R I A I S Table 8. Amino acid differences between chicken and turkey AMPV strains in the polymerase protein (L)

Subtype Field strain (F) or Vaccine strain (V) Amino acid changes

A V 7 A V 4 B V 5 B F 4 B F 0 B F 0 B F 0

Table 9. Number of amino acid changes occurring in turkey AMPVs after 10 passages in chicken TOC.


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USE OF REVERSE GENETICS TO DEVELOP A POSITIVE CONTROL VIRUS FOR RT-NESTED PCR DETECTION OF SUBTYPE A AND B AVIAN

METAPNEUMOVIRUS (AMPV)

FALCHIERI M1, BROWN PA 1, CATELLI E 2 and NAYLOR CJ1

1Department of Veterinary Pathology, University of Liverpool, Leahurst Campus, Neston, Cheshire, CH647TE, UK.

2 Department of Veterinary Public Health and Animal Pathology,University of Bologna, via Tolara di Sopra 50, Ozzano dell’Emilia (BO), 40064, Italy.

SUMMARY A standardized RT-nested PCR of high sensitivity has been in use in our laboratory for some time to detect avian metapneumovirus subtype A and B. Until now we have avoided the use of positive control viruses (CV) because of the risk of contamination leading to sample false positives. The paper describes the production and testing of a modified virus which in our standard PCR produces RT-nested PCR amplicons of increased sizes compared to those generated from unmodified viruses, thus enabling cross contamination to the sample tested to be instantly detectable. A DNA copy of an AMPV subtype A genome was modified by site directed mutagenesis to introduce the subtype B G gene primer sequence (G9+B) at the equivalent position in the subtype A G gene. To increase amplicon sizes, an insertion was introduced between binding sites for the opposing primer pairs (outer pair G1+ and G6-, inner pairs G8+A and G5- or G9+B and G5-). PCR from the modified DNA produced nested amplicons of 463 and 556 bp compared to 268 for unmodified subtype A and 361 for unmodified subtype B viruses. Using reverse genetics the modified DNA generated a virus which, after RNA extraction, gave the same 463 and 556 bp amplicons in RT-nested PCRs. For further convenience virus was absorbed onto filter paper, dried and inactivated by microwave treatment, then stored in flip top tubes. RNA extracted from papers were henceforth used as reliable, efficient and distinguishable RT-nested PCR positive controls. INTRODUCTION A standardized RT-nested PCR of high sensitivity has been in use in our laboratory for some time to detect and distinguish AMPV subtypes A and B. This protocol is based on sequences of the G gene and it consists of an RT step and two subsequent PCRs (Cavanagh et al., 1999).


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Because PCR is a very sensitive technique, false positive results can sometimes occur due to cross contamination from the positive control template. The paper describes the production and testing of a modified virus which in our standard PCR produces final amplicons of increased sizes compared to those generated from unmodified viruses, thus allowing rapid recognition in the event of such a cross contamination. The control virus was made by modifying a full length AMPV DNA copy which was subsequently rescued in an AMPV reverse-genetics system (Naylor et al., 2004). This was adsorbed onto filter paper and inactivated to allow potential safe use and transport of the material as had been previous demonstrated by Elhafi et al. (2004). MATERIALS and METHODS Construction of the positive control (Fig. 1) Virus was modified so that a standard RT-nested PCR (Cavanagh et al., 1999) would produce bands of increased size. A cloned full-length DNA copy of an AMPV subtype A genome was modified by site direct mutagenesis (SDM) to introduce the subtype B G gene primer sequence (G9+B) at the equivalent position in the subtype A G gene. To increase amplicon sizes, an insertion was introduced between opposing primer binding sites. (outer pair G1+ and G6-, inner pairs G8+A and G5- or G9+B and G5-). The insert, of about 200 bp, was amplified from the AMPV M2 gene by PCR using primers which introduced Xho1 sites at both ends. A Sal1 site was introduced between G8+A and G5- in the plasmid by site direct mutagenesis. After digestion of the plasmid and of the insert with Sal1 and Xho1 respectively, they were ligated together using T4 DNA ligase, in presence of both enzymes. The modified plasmid was amplified in E.coli and purified with “QIAprep Spin Miniprep Kit” (Qiagen). Virus Rescue Rescue of virus from the full length DNA copy was performed as described by Naylor et al. (2004). Presence of virus was shown by low power microscopy (Fig. 2). Viral inactivation and storage The modified virus was adsorbed onto filter paper, dried and inactivated by microwave treatment. Small pieces (0.5 x 3 cm) were cut and stored in 1.5 ml flip top tubes at 4°C. Testing the control virus RT-nested PCR was carried out on the inactivated modified virus together with unmodified subtype A and B viruses.


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RESULTS The modified virus produced bands of 463 and 556 bp. Unmodified subtype A and B viruses produced 268 bp and 361bp bands respectively (Fig. 3). DISCUSSIONS A positive control is desirable in PCR because negative results due to absence of suitable template can be distinguished from those produced due to the diagnostic system malfunctioning. The disadvantage is that contamination from that control material can lead to false positives samples. (Kwok & Higuchi, 1989). We have created a positive control virus, using reverse genetics, that produces amplicons about 200 bp larger than those produced by the unmodified viruses (463 bp for A subtype and 556 bp for B subtype, against 268 and 361 bp). Bands are clearly and immediately recognizable on agarose gels as different to test positives. The virus grew readily on Vero cells. The inactivation of the virus allows for its easy storage for relatively long periods. Equally, because the virus is a genetically modified organism, inactivation avoids restrictions in movement and use which would apply if it remained viable. Our positive control has proved to be a reliable and efficient way to eliminate the risk of false positive samples due to contamination from positive controls. REFERENCES Cavanagh D., Mawditt K., Britton P., Naylor C.J.. Longitudinal field studies of

infectious bronchitis virus and avian pneumovirus in broiler using type-specific polymerase chain reactions. Avian Patholology, 1999; 28:593-605.

Elhafi G., Naylor C.J., Savage C.E., Jones R.C.. Microwave or autoclave treatments destroy the infectivity of infectious bronchitis virus and avian pneumovirus but allow detection by reverse transcriptase-polymerase chain reaction. Avian Pathology, 2004; 33 (3): 303-306.

Kwok S., Higuchi R.. Avoiding false positives with PCR. Nature, 1989; 18 May 339 (6221): 237-238.

Naylor C.J., Brown P.A., Edworthy N., Ling R., Jones R.C., Savage C.E., Easton A.J.. Development of a reverse-genetic system for Avian pneumovirus demonstrates that the small hydrophobic (SH) and attachment (G) genes are not essential for virus viability. Journal of General Virology, 2004; 85: 3219-3227.


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Fig. 1: Modification of the full length DNA copy of AMPV

Fig. 2: AMPV Cytopathic effect on Vero cells


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Fig. 3: Subtype A (268 bp) Subtype B (361 bp) Positive control virus (463 and 556 bp) Negative control (NC)


299

THE EFFECT OF SH GENE MODIFICATIONS ON CYTOPATHIC EFFECTS SEEN IN VERO CELLS

BROWN PA and NAYLOR CJ Department of Veterinary Pathology, University of Liverpool, UK

SUMMARY Previously subtype A and B avian metapneumoviruses have been shown to produce either focal or syncytial cytopathic effects in Vero cells. Development of subtype A AMPV reverse genetics led to viruses lacking the entire SH gene which caused unique giant syncytial formations in the same cells. The current study further examines this phenomenon by making and testing viruses selectively expressing shortened SH proteins. The 173 amino acid membrane associated SH protein has a hydrophobobicity profile suggesting a type 1 membrane protein with a membrane domain closest to the N terminus. Using reverse genetics, 9 viruses were made with SH proteins shortened from either end. Shortening of the C terminus end was made by introduction of premature stop codons into the SH open reading frame. Progressive shortenings to the N terminus were made by deletions of the relevant section combined with introduction of substitute transcription and translation start signals closer to the the C terminus. All constructs were shown to produce the desired SH message and some were quantified using PCR. Infection of Vero monolayers with the viruses led to giant syncytial formations in all cases, with the exception of the virus where the SH gene had a stop codon following base 238 and therefore could express the only first 80 amino acids of the protein. However the presumed untranslated section of the gene was still present, so there remained a small probability that the remainder of the gene was being expressed by unknown mechanisms. To eliminate this possibility bases 238 to the 3’ gene end were deleted, and this also gave the wild type CPE in vero cells. It is not currently possible to draw clear conclusions about which section(s) of the SH gene is (are) responsible for the maintenance of the wildtype CPE. MATERIALS and METHODS Modification of virus A full length DNA copy cloned of a European subtype A AMPV cloned in E Coli was modified so as to remove or reduce the size of the SH gene open reading frame using standard molecular biology techniques. One modified copy had the SH gene removed in its entirety. In the other eight variants reduced SH protein sections coding amino acids 1 to 38, 1 to 78, 1 to111, 1 to 144, 132 to 173, 86 to 173, 59 to 173 and 29 to 173 were present. For reduction of SH from the C terminal, protein truncation


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was achieved by introduction of premature stop codon. For N terminal reductions, sections between the SH transcription start and newly introduced late start codon were removed. One C terminal truncation was made by removal of the relevant section of the genome (coding for aa 79 to 173). Constructs sizes are summarised in fig 1 Fig 1

All clones were used to rescue virus in 12 well plates of confluent vero cells as previously described (Naylor et al, 2004) and outlined in Figure 2. Fig 2

132-173

86-173

59-173

29-173

Unmodified 1-173

1-38

1-78

1-111 1-144

NEW VIRUS

M2 protein

Phosphoprotein

Nucleocapsid

N

M

F

M2

P

L

SH G

N

M

F

M2

P

L

SH G

DNA copy of genome

Polymerase


301

After 1 week, contents of rescue wells were freeze thawed then used to infect T25 flasks of freshly confluent Vero cells. Effect of rescued virus on vero cells Cell sheets were observed by light microscopy and photographed. Transcription An RT- PCR on extracted viral RNA using oligo dT and SH specific oligo nucleotide was used to determine whether SH message had been made in the modified viruses. RESULTS Effect of rescued virus on vero cells All viruses with shortened or absent SH open reading frames produced giant syncytia (fig 3) where large sections of the cell sheet were fused together. This was with the exception of the two viruses having the SH gene section coding for amino acids 1-78 (premature stop codon or 79-173 deletion) which produced CPE typical of the same virus with an intact SH gene (Figure 4). Uninfected Vero cells are shown in figure 5. Fig 3 typical giant syncytia seen with SH deletion viruses

Fig 4 Syncytial type seen with mutants 1-78 and unmodified virus

Fig 5 uninfected vero cells


302

Transcription All viruses produced the expected mRNA transcripts as illustrated in figure 6. Fig 6 Detection of SH mRNA in viruses

DISCUSSION Characteristic giant syncytia have been previously described for AMPVs lacking an SH protein (Ling et al, 2008, Naylor et al 2004) and the current study has extended that work to show that viruses with partial SH deletions show a similar CPE, with the exception of viruses expressing amino acids 1-78 of the SH open reading frame. The giant syncytia frequently contain readily observable multiple nuclei. The syncytia appear to be large single cells made from the individual cells of the cells monolayer and the individual multiple nuclei presumably came from those individual cells. The syncytia appear to have arisen due to increased fusion activity by the SH modified viruses. If this is the case, it implies that the SH protein has some role in regulation of fusion and this presumably offers some advantage to the virus. One advantage already seen is that higher virus titres result from cells infected with unmodified virus but the mechanism remains unknown. It is not clear why the viruses expressing amino acids 1-78 have a normal CPE. It seems possible that aa 1-78 may represent the minimum active section of the SH protein but it is then difficult to explain why viruses with longer but still incomplete SH

AAAAAAAAA

Selective mRNA RT-PCR

mRNA transcript

Virus 1-38 stop

Virus 1-78 stop

Virus 1-111 stop

Virus 1-144 stop

Virus 29-173

Virus 1-78 deletion

Virus 59-173

Virus 132-173

Virus 86-173

Virus 1-173

TTTTTTTT

SH gene


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proteins do not behave similarly. More work will be needed before this phenomenon can be explained. REFERENCES Ling, R., Sinkovic, S., Toquin, D., Guionie, O., Eterradossi, N. & Easton, A. J. (2008).

Deletion of the SH gene from avian metapneumovirus has a greater impact on virus production and immunogenicity in turkeys than deletion of the G gene or M2-2 open reading frame. J Gen Virol 89, 525-533.

Naylor, C. J., Brown, P. A., Edworthy, N., Ling, R., Jones, R. C., Savage, C. E. & Easton, A. J. (2004). Development of a reverse-genetics system for Avian pneumovirus demonstrates that the small hydrophobic (SH) and attachment (G) genes are not essential for virus viability. J Gen Virol 85, 3219-3227.


304

CONSTRUCTION OF GFP AVIAN METAPNEUMOVIRUS (AMPV) RECOMBINANT LACKING THE SMALL HYDROPHOBIC PROTEIN GENE

LUPINI C1, CATELLI E1, CECCHINATO M2 and NAYLOR CJ3

1 Università di Bologna, Dipartimento di Sanità Pubblica Veterinaria e Patologia Animale, Via Tolara di Sopra 50, 40064 Ozzano Emilia (BO), Italy;

2 Università di Padova, Dipartimento di Sanità Pubblica, Patologia Comparata ed Igiene Veterinaria, Agripolis - Viale dell'Università 16, 35020 Legnaro (PD), Italy;

3 University of Liverpool, Department of Veterinary Pathology, Jordan Building, Leahurst, CH64 7TE Neston, United Kingdom;

SUMMARY An AMPV reverse-genetics system was used to create a virus lacking the entire SH open reading frame which produced unusual giant syncytial formations in cell culture. The replacement of the gene with GFP resulted in virus which also produced the same syncytial formations. The intrinsic fluorescent properties of the virus may prove useful in pathogenesis studies. The similarity of size of the 2 genes suggests that the altered cytopathic effect did not result from the shortening of the genome and possible consequential effects on the transcription or translation of the following attachment and polymerase protein genes. INTRODUCTION Avian Metapneumovirus (AMPV) is a member of the Paramyxoviridae family and is responsible for Turkey Rhinotracheitis in turkeys which is an upper respiratory tract infection. This can cause severe disease and losses where secondary pathogens exacerbate the situation. An AMPV reverse-genetics system have been developed which enables specific mutations to be introduced into the virus genome and the subsequent phenotypic consequences determined (Naylor et al., 2004). This system was used to show that infection with a recombinant virus, unable to express the SH gene, resulted in the production of unusually large syncytia in Vero cells. It is possible that this was due to a change in the pattern of genome transcription resulting from the loss of a transcription unit. In this work we replace the SH gene with GFP which is a gene of similar size, and in which expression can be confirmed by fluorescence of its gene product, in suitable UV irradiation.


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MATERIALS and METHODS Construction of ΔSH-AMPV-GFP The ΔSH-AMPV plasmid was modified using Site-direct mutagenesis (Quikchange; Stratagene) to insert a SalI restriction site immediately following the gene start transcription signal. XhoI restriction sites were added to each end of the GFP gene by PCR copying of the gene from plasmid pEcoli-6xHN-GFPuv (Clonetech) using primers containing the XhoI recognition site. Ligation of SalI-cut ΔSH-AMPV and XhoI-cut GFP in the presence of SalI and XhoI, was performed and resulted in the final construct ΔSH-AMPV-GFP. This was selected and amplified by transformation of Stb12 competent cells followed by PCR screening and sequencing, further culture, DNA purification and final RE screening. Virus rescue The transfection was performed in Vero cells inoculated with a Fowlpox recombinant virus expressing T7 polymerase. The cells were transfected with the full length plamid ΔSH-AMPV-GFP and other plamids expressing Nucleocapsid (N), Phosphoprotein (P), Matrix 2nd (M2) and Polimerase (L) genes using Lipofectamine 2000. The transfection mixture was removed after over night of incubation at 37°C, and the cells were washed and maintained with medium containing 5% fetal calf serum The transfected cell sheet was viewed, freeze-thawed and the clarified material was used to infect new cells. Resultant cell sheets were examined daily for citopathic effect (CPE) and monitored by fluorescent microscopy for the expression of GFP. RESULTS Green fluorescent cells were observed two days after transfection of ΔSH-AMPV-GFP but not ΔSH-AMPV. In both instances, CPE appeared as scattered groups of cells which later developed into giant syncytia. The syncytia induced by the two viruses were similar and were indistinguishable. Further fluorescence microscopy of virus which had received 3 serial passages showed that the GFP gene continued to be expressed. These results indicated that the recovered recombinant viruses exhibited growth properties similar to those of the wild-type virus in tissue culture and that the insertion of a foreign gene does not noticeably affect the in vitro replication characteristics of ΔSH-AMPV. DISCUSSION The similarity of size of the SH and GFP genes suggests that the altered cytopathic effect did not simply result from removal of an expressing gene and the possible consequential effects on the transcription or translation of the following attachment and polymerase protein genes. This shows that the SH protein is responsible for maintaining the wild type CPE generally seen with AMPVs. In particular, for AMPV, the absence rather than the presence of the SH protein enhances fusion. It is not known whether the large syncytia in the SH-gene-deleted virus are due to the SH protein having an inhibitory effect on fusion or whether the


306

SH protein is required for alteration of a default giant syncytial CPE to form the focal type. The study also shows that AMPV is able to accept, express and maintain a foreign gene over at least 3 passages. Similar recombinant viruses expressing other genes are likely to find numerous applications. In particular, the expression of genes from other respiratory pathogens are likely to enable the construction of promising recombinant live respiratory vaccines in both turkeys and chickens. GFP recombinant may prove useful in pathogenesis studies. REFERENCES Naylor C.J., Brown P.A., Edworthy N., Ling R., Jones R. C., Savage C. E., Easton A.

J.(2004) Development of a reverse-genetics system for Avian pneumovirus demonstrates that the small hydrophobic (SH) and attachment (G) genes are not essential for virus viability. Journal of General Virology. 85: 3219-3227.


307

GROWTH IN VITRO AND IN VIVO OF A RECOMBINANT AMPV EXPRESSING eGFP

EDWORTHY1NE, TOQUIN2 D, ZWINGELSTEIN2 F, ETERRADOSSI2 N and EASTON1 AJ

1 Department of Biological Sciences, University of Warwick, Coventry, UK. 2 Agence Française de Securite Sanitaire des Aliments (AFSSA) Avian and Rabbit

Virology Immunology and Parasitology Unit (VIPAC), BP53, 22440 Ploufragan, France

SUMMARY A cDNA copy of the group A strain CVL14/1 AMPV genome was modified by introduction of novel restriction sites and the insertion of a gene encoding the enhanced green fluorescent protein. The eGFP gene was flanked by AMPV transcriptional initiation and termination signals and was inserted immediately upstream of the virus SH gene. The cDNA was used to recover infectious virus, designated mutant AMPVeGFP which was used to infect tissue culture cells and turkey poults. The addition of eGFP as an extra gene in genome AMPVeGFP did not decrease the level of mRNA produced from the downstream G gene compared to the control CF2 genome which did not contain the eGFP gene. Confocal microscopy time-lapse experiments were used to monitor the spread of virus in cell culture, demonstrating the spread during the formation of syncytia. Following intranasal inoculation of SPF turkey poults quantitative real-time PCR for the AMPV N gene detected the highest signals in the sinuses for birds infected with AMPVeGFP at both three and five days p.i. (109.9 and 1010.2 copies / g of tissues, respectively) with detectable RNA levels also seen in all of the respiratory tissues on both days (106.9 to 108.2 copies / g of tissues). RNA was also seen in spleen, liver, thymus, bursa of Fabricus, and M. pectoralis at 3 days p.i. (106.7 to 108.0 copies / g of tissues), and in all tissues except for the liver and bursa of Fabricius at 5 days p.i. No RNA was detected in any of the mock infected controls. Overall, the genome of AMPVeGFP appears to reach all deeper organs and muscle tissues but the expression of eGFP was detected only in sinus tissue from infected birds. The recombinant virus expressing eGFP offers a method to follow the spread of virus in infected animals.


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INTRODUCTION As with all pneumoviruses, avian metapneumovirus (AMPV) is thought to be primarily a respiratory pathogen though the exact tropism of the virus has been shown to be strain and host (chickens or turkeys) dependent (Cook et al., 1993). AMPV subgroup A virus was found in the turbinates, sinuses, upper and lower trachea, lungs and air sacs of experimentally infected turkeys periodically up to 10 days post infection (d.p.i.) whereas AMPV subgroup B was found mainly in the upper respiratory tract (Buys et al., 1989, Cook et al., 1991, Jones et al., 1988, McDougall & Cook, 1986, Van de Zande et al., 1999). Indirect immunofluorescence experiments confirmed the presence of AMPV antigen in the turbinates and trachea but not lungs or air sacs if infected turkeys (Jones et al., 1988). Analysis of turkey poults and chickens experimentally infected with AMPV showed that the virus associated with the epithelial ciliated cells of the turbinates and trachea (Majo et al., 1995). In chickens, in vitro experiments have shown ciliostasis in tracheal organ cultures and syncytia formation in chicken embryo cell cultures following of AMPV infection (Gough et al., 1988). In experimentally infected chickens, virus was isolated from the respiratory tract (nasal tissue, sinuses, trachea, and lung), though it was found most abundantly in the upper respiratory tract tissues. In these experiments, no virus was isolated from the kidney, liver, duodenum, bursa of Fabricius or caecal tonsils. Using immunoperoxidase staining, virus was found most consistently in nasal turbinates up to 5 d.p.i. and was found occasionally in sinuses and trachea on days 4 and 5 d.p.i. Associated histological analysis of the upper respiratory tract tissues showed changes most strikingly in the nasal turbinates and less severely in the sinuses and trachea (Catelli et al., 1998). The association of AMPV infection with egg drop syndrome suggested that the virus was capable of infecting the reproductive tract. However, direct experimental evidence is mixed, with Giraud et al. (1986) finding no evidence of transmission of the virus to eggs in turkeys, but several other groups finding AMPV in the reproductive tract of broiler breeders and laying turkeys (Anon., 1985, Lister & Alexander, 1986, O'Brien, 1985, Wyeth et al., 1987). In chickens, Jones et al. (1988) found AMPV in the reproductive tract of laying hens, as well as in the upper respiratory tract, but not in blood or ovaries and AMPV antigen has been detected by immunoperoxidase staining of antigen in the oviduct in laying hens (Cook et al., 2000). Shedding of small amounts of AMPV from turkeys up to 14 d.p.i. has been reported (Cook et al., 1991), again suggesting that AMPV is able to infect tissues outside of the respiratory tract. Reverse genetics systems have been described for AMPV which offer the possibility of introducing specific mutations or other genetic alterations into the AMPV genome (Govindarajan et al., 2006; Naylor et al., 2004; Naylor et al., 2007). This approach has been utilised to determine the function of naturally occurring mutations isolated from field strains (Naylor et al., 2007). The insertion of a reporter gene, such as green florescent protein (GFP), into the viral genome can serve as a marker of recombinant virus production and expression of GFP by a recombinant virus also allows visualization of virus in vitro and in vivo, thereby increasing the speed of analysis and production of slow growing viruses (Govindarajan et al., 2006; Techaarpornkul et al., 2001). AMPV has three glycoproteins expressed on its surface; the F (fusion) protein, the G (glycoprotein) protein and the SH (small hydrophobic) protein. While the roles of the


309

F and G proteins have been well defined the function of SH protein is unknown. Recombinant viruses have been created which lack one or more of the viral glycoproteins. A recombinant AMPV, lacking the SH gene produced a syncytial plaque phenotype, suggesting a role in regulation of the cell-cell fusion process. Recombinant AMPV lacking an SH or G gene were also shown to replicate poorly in turkeys compared to wildtype AMPV (Ling et al., 2008). We have used as reverse genetics approach to insert an eGFP ORF either in place of the SH gene or after the SH gene in the CF2 AMPV cloned genome (Naylor et al., 2007). These recombinant viruses were then used to follow virus replication in vitro and the AMPVeGFP virus was used to determine sites of replication in vivo. This was complemented by using quantitative real time PCR to determine the levels of AMPV-specific RNA in the tissues of experimentally infected turkey poults. MATERIALS and METHODS Generation of recombinant AMPV genomes The SH gene of the CF2 AMPV genome is flanked by the sequences recognised by the restriction enzymes AgeI and EagI. A fragment containing the complete coding sequence of the enhanced green florescent protein (eGFP) derived from the jellyfish Aequorea Victoria was generated by PCR from the plasmid pEGFP-C1 (BD Biosciences Clontech, GenBank Accession #: U55763) using one primer containing the site for AgeI and another containing the site for EagI together with the appropriate SH gene transcription start and end sequences.. The PCR product was inserted into a plasmid containing a complete copy of the CF2 AMPV genome cDNA which had been previously digested with AgeI and EagI. The resulting plasmid was used to generate an infectious virus in which the SH gene had been replaced with that of eGFP. The virus was named AMPVeGFPΔSH. A virus containing all of the AMPV genes but also containing a copy of the eGFP gene was generated by inserting a copy of the AMPV SH gene into the eGFP-G intergenic region of the AMPVeGFPΔSH genome The cDNA containing the AMPV-ΔSH genome was digested with restriction enzymes EagI and SmaI which are located immediately downstream of the eGFP gene. A PCR amplified fragment containing the complete AMPV SH gene flanked by EagI and SmaI recognition sites was generated and inserted into the AMPVeGFPΔSH genome. The resulting recombinant virus contained all of the AMPV genes together with an eGFP gene located between the SH and G genes. This virus was named AMPVeGFP. All recombinant viruses were recovered from cDNA as described previously (Ling et al., 2008). A diagrammatic representation of the recombinant virus genomes is shown in Figure 1. Infection of tissue culture cells and visualisation of the recombinant AMPV. BSC-1 cells were infected with AMPVeGFP at a multiplicity of infection of 2 plaque forming units per cell for 1hr at 33ºC. The medium was removed, and the cells overlaid with Leibovitz’s L-15 medium (Invitrogen). The cells were visualised using a Leica SP5 confocal microscope. Photographs were taken between 5hr and 33hr 20min p.i. every 10min in a heated chamber (33°C) with a 489nm Argon laser. Infection was monitored over 2300min, with the majority of eGFP expression occurring between 300 and 2000min (5hr and 33hr 20min).


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Infection of turkeys Specific pathogen free (SPF) turkeys were hatched and kept in isolators, blood samples taken, and one group of nine birds with one control group of six birds were established. The experimental group was then infected via intranasal inoculation with 0.1ml AMPVeGFP per bird, and the SPF birds with 0.1ml of MEMH medium as the mock-inoculated controls. At three and five days post inoculation two birds from each group were culled and organs taken for histology and quantitative real-time PCR experiments. These tissues were stored in 10% paraformladehyde. The organs taken were: sinuses, upper trachea, lower trachea (with bronchi), lung, oviduct (in females), spleen, liver, thymus, bursa of Fabricus, and M. pectoralis. The remaining birds were culled at 21 days p.i. Detection of eGFP expression in vivo The respiratory tissue samples (sinuses, lung, upper trachea, and lower trachea) from infected and control animals were examined for eGFP expression. A sample of each organ was removed from paraformladehyde and soaked for at least 24hours in PBS before being snap frozen in Tissue-Tek O.C.T.™ compound (Sakura Finetek). The embedded tissues were then sliced using a freezing microtome (OTF 5000, Bright) and 5µm sections were mounted with Gel Mount™ Aqueous Mounting Medium (Sigma-Aldrich) on slides. The slide cover slips were immediately sealed and examined for eGFP fluorescence using a Leica SP5 confocal microscope. Four or more slices were examined for each bird for all respiratory organs. Detection of virus RNA by real time PCR. RNA was prepared and AMPV N gene RNA was specifically detected using the procedure described by Guionie et al. (2006). RESULTS Detection of eGFP expression in vitro and in vivo Tissue culture cells infected with the recombinant AMPVs expressing eGFP were analysed using confocal microscopy. In both cases expression of eGFP was clearly observable throughout the cytoplasm of the infected cells, with no significant differences between the viruses either containing or lacking the SH gene (Figure 2). Following infection with AMPVeGFP, a time-lapse film created from a compilation of images prepared every 10 minutes over a time course from 5hr to 33hr 20min p.i. was prepared (not shown). This demonstrated that the virus infection spread outwards from the initial point of infection to neighbouring cells, probably by cell-cell fusion. Analysis of sinus tissues from two birds infected with virus AMPVeGFP showed clear eGFP expressing cells from both birds on day 3 and from one of the birds on day 5 (Figure 3). Sinus tissues from birds mock infected showed no GFP expression. No other organ studied showed eGFP expression. Detection of AMPV RNA in vivo Tissue samples taken from SPF turkeys infected with AMPVeGFP were taken and RNA extracted. The RNA was used in a real time PCR designed to detect the virus N gene RNA. The results are shown in Table 1.


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DISCUSSION Two recombinant AMPV genomes were generated, each containing a copy of the eGFP gene flanked by appropriate virus transcription initiation and termination sequences. One virus contained a complete copy of the genome with the eGFP gene inserted prior to the virus SH gene while the second had the SH gene replaced with the eGFP gene (Figure 1). These viruses were used to infect tissue culture cells and eGFP gene expression was monitored by confocal microscopy. Both viruses were able to express the eGFP gene (Figure 2). The time-lapse experiments showed the spread of virus in cell culture and the development of syncytia formation (not shown). Introduction of the eGFP gene into the AMPV genome also showed that the genome is capable of accepting large foreign gene insertions, suggesting that recombinant AMPV genomes may be used in the creation of multivalent vaccine. The eGFP expression of AMPVeGFP also allowed visual identification of virus-infected cells within turkey tissue samples. Visible levels of eGFP were found in the sinus tissues of two infected birds at 3 days p.i. and one bird at 5 days p.i., demonstrating virus replication in the respiratory tract (Figure 3). In contrast, while quantitative real-time PCR confirmed the presence of the virus at the highest levels in the respiratory tract, this technique also showed low levels of virus RNA in all tissues sampled in at least one bird on day 3 or day 5 p.i. (Table 1). These data demonstrate that recombinant viruses can be used to follow the spread of virus infection both in vitro and in vivo. The expression of eGFP was sufficiently sensitive and specific to identify tissues which support virus gene expression. These data suggest that the virus gene expression may be restricted to the sinus region of the respiratory tract, though it was not possible to identify the specific cell type involved. In apparent contradiction, the real time PCR data indicate that the virus RNA can be isolated from a very wide range of tissues. However, these data do not indicate whether the widely distributed virus RNA represents true spread of infectious virus or simply passive transfer of non-replicating virus to other sites. This question remains the subject of further study. REFERENCES Anon. (1985). Turkey rhinotracheitis of unknown aetiology in England and Wales

(Preliminary report from the British Veterinary Poultry Association). Vet. Record 117, 653-654.

Buys, S. B., Du Preez, J. H. & Els, H. J. (1989). The isolation and attenuation of a virus causing rhinotracheitis in turkeys in South Africa. Onderstepoort J.Vet. Res. 56, 87-98.

Catelli, E., Cook, J. K., Chesher, J., Orbell, S. J., Woods, M. A., Baxendale, W. & Huggins, M. B. (1998). The use of virus isolation, histopathology and immunoperoxidase techniques to study the dissemination of a chicken isolate of avian pneumovirus in chickens. Avian Pathol. 27, 632-640.

Cook, J. K., Ellis, M. M., and Huggins, M. B. (1991). The pathogenesis of turkey rhinotracheitis virus in turkey poults inoculated with the virus alone or together with two strains of bacteria. Avian Pathol. 20, 155-166.


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Cook, J. K. A., Kinloch, S. & Ellis, M. M. (1993). In vitro and in vivo studies in chickens and turkeys on strains of turkey rhinotracheitis virus isolated from the two species. Avian Pathol. 22, 157-170.

Giraud, P., Bennejean, G., Guittet, M. & Toquin, D. (1986). Turkey rhinotracheitis in France: Preliminary investigations on a ciliostatic virus. Vet. Record 119, 606-607.

Gough, R. E., Collins, M. S., Cox, W. J. & Chettle, N. J. (1988). Experimental infection of turkeys, chickens, ducks, geese, guinea fowl, pheasants and pigeons with turkey rhinotracheitis virus. Vet. Record 123, 58-59.

Govindarajan, D,, Buchholz, U. J. & Samal, S. K. (2006). Recovery of avian metapneumovirus subgroup C from cDNA: cross-recognition of avian and human metapneumovirus support proteins. J. Virol. 80, 5790-5797.

Guionie, O., Toquin, D., Sellal, E., Bouley, S., Zwingelstein, F., Allée, C., Bougeard, S., Lemière, S. & Eterradossi, N. (2007). Laboratory evaluation of a quantitative real-time reverse transcription PCR assay for the detection and identification of the four subgroups of avian metapneumovirus. J. Virol. Methods. 139, 150-158.

Jones, R. C., Williams, R. A., Baxter-Jones, C., Savage, C. E. & Wilding, G. P. (1988). Experimental infection of laying turkeys with Rhinotracheitis virus: Distribution of virus in the tissues and serological response Avian Pathol. 17, 841-850.

Ling, R., Sinkovic, S., Toquin, D., Guionie, O., Eterradossi, N. & Easton, A. J. (2008). Deletion of the SH gene from avian metapneumovirus has a greater impact on virus production and immunogenicity in turkeys than deletion of the G gene or M2-2 open reading frame. J. Gen. Virol. 89, 525-33.

Lister, S. A. & Alexander, D. J. (1986). Turkey Rhinotracheitis: A Review. Veterinary Bull. 56, 637-663.Majo, N., Allan, G. M., O'Loan, C. J., Pages, A. & Ramis, A. J. (1995). A sequential histopathologic and immunocytochemical study of chickens, turkey poults, and broiler breeders experimentally infected with turkey rhinotracheitis virus. Avian Dis. 39, 887-896.

McDougall, J. S. & Cook, J. K. (1986). Turkey rhinotracheitis: preliminary investigations. Vet. Record 118, 206-207.

Naylor, C. J., Brown, P. A., Edworthy, N., Ling, R., Jones, R. C., Savage, C. E. & Easton, A. J. (2004). Development of a reverse-genetics system for Avian pneumovirus demonstrates that the small hydrophobic (SH) and attachment (G) genes are not essential for virus viability. J. Gen. Virol. 85, 3219-3227.

Naylor, C. J., Ling, R., Edworthy, N., Savage, C. E. & Easton, A. J. (2007). Avian metapneumovirus SH gene end and G protein mutations influence the level of protection of live-vaccine candidates. J. Gen. Virol. 88, 1767-7175.

O'Brien, J. D. (1985). Swollen head syndrome in broiler breeders. Vet. Record 117, 619-620.

Techaarpornkul, S., Barretto, N. & Peeples, M. E. (2001). Functional analysis of recombinant respiratory syncytial virus deletion mutants lacking the small hydrophobic and/or attachment glycoprotein gene. .J Virol. 75, 6825-6834.

Van de Zande, S., Nauwynck, H. & Pensaert, M. (2001). The clinical, pathological and microbiological outcome of an Escherichia coli O2:K1 infection in avian pneumovirus infected turkeys. Vet. Microbiol. 81, 353-365.


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Wyeth, P. J., Chettle, N. J., Gough, R. E. & Collins, M. S. (1987). Antibodies to TRT in chickens with swollen head syndrome. Vet. Record 120, 286-287.

Figure 1.: Diagrammatic representation of recombinant AMPV genomes containing n eGFP gene flanked by transcription initiation and termination signals. A. Recombinant AMPVeGFPΔSH in which the eGFP gene has replaced the AMPV SH gene. B. recombinant AMPVeGFP in which the eGFP gene is located between the SH and G genes.

Figure 2.: Tissue culture cells infected with mutant AMPV expressing eGFP. Panels A and C show phase contrast. Panels B and D show eGFP fluorescence visualised by confocal microscopy. Panels A and B are cells infected with recombinant AMPVeGFPΔSH and panels C and D are cells infected with recombinant AMPVeGFP.


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Figure 3.: Detection of eGFP expression in sinus tissue taken from turkeys infected with recombinant AMPVeGFP. A. Sinus tissue prepared 3 days post infection. B. Sinus tissue prepared 5 days post infection.

Table 1.: Real-time quantitative PCR determination of number of copies of the AMPV N gene in tissues taken from turkeys infected with either AMPVeGFP or AMPVeGFPΔSH. The numbers of RNA molecules are shown as log 10 copies per 10mg of tissue. Samples were prepared from turkeys at either 3 or 5 days post infection (d.p.i.). Values in parenthesis were outside of the linear range of the real-time PCR control and undetectable reactions (Ct>40) are marked as zero. Duplicate values are presented where available.

3 d.p.i

Sinus Upper Trachea

Lower Trachea

Lungs Spleen Liver Thymus Bursa of Fabricius

Pectoralis muscle

AMPVeGFP 7.48 8.59

6.00 6.63

6.81 7.31

4.64 5.00

5.14 5.98 5.07 5.62

5.61 5.32

5.32AMPV-eGFPΔSH

7.92 7.64

5.49 5.49

4.90 (4.35)

4.83 4.97

4.67 4.78 (4.26) 5.02 5.31

5 d.p.i

AMPVeGFP 8.51 8.82

5.25 5.68

6.33 6.74

(3.70) 5.33

6.27 4.62 5.37 5.64

5.37 5.39

(4.47)

AMPVeGFPΔSH

7.79 7.82

6.31 6.30

6.30 6.69

4.66 6.51

(4.31) (4.01)

0 4.88 0 5.53


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GENERATION AND BIOLOGICAL ASSESSMENT OF RECOMBINANT AVIAN METAPNEUMOVIRUS SUBGROUP C (AMPV-C) VIRUSES CONTAINING

DIFFERENT LENGTH OF THE G GENE

YU Q, ESTEVEZ C, KAPCZYNSKI D and ZSAK L

USDA-ARS, Southeast Poultry Research Laboratory, 934 College Station Road, Athens, GA 30605, U.S.A

SUMMARY Genetic variation in length of the G gene among different aMPV-C isolates has been reported. However, its biological significance in virus replication, pathogenicity and immunity is unknown. In this study, we developed a reverse genetics system for aMPV-C and generated two Colorado (CO) strain-based recombinant viruses containing either the full-length G gene derived from a Canadian goose isolate or a C-terminally truncated G gene of the CO strain. The truncated short G (sG) gene encodes 252 amino acids (aa), which is 333 aa smaller than the full-length G (585 aa). The biological properties of these two recombinant G variants were assessed in Vero cells and in specific-pathogen-free (SPF) turkeys. The results demonstrated that the large portion (333 aa) of the extracellular domain of the viral attachment protein is not essential for virus viability both in vitro and in vivo, but may play a role in enhancing virus attachment specificity and immunity in a natural host. INTRODUCTION The glycoprotein (G) of avian metapneumovirus (aMPV) has been much of scientific curiosity owning its role in viral attachment, and possible role in viral virulence and protective immunity. The G protein of aMPV isolates exhibits extensive genetic and antigenic variations that are the primary criteria for aMPV subgroup classification and for consideration of vaccine development (Cook, 2000; Juhasz and Easton, 1994; Patnayak et al., 2002). Among the aMPV subgroups, aMPV-C isolates possess the most extensive sequence divergence from other subgroups (Toquin et al., 2006). Furthermore, the G gene length reported by different researchers for aMPV-C strains isolated from the United States was strikingly different, ranging from 783 nts (252 aa) to 1798 nts (585 aa) (Alvarez et al., 2003; Bennett et al., 2005; Govindarajan et al., 2004; Lwamba et al., 2005). Recent studies have shown that the G gene length variation for those U.S. isolates was resulted from a truncation of the G gene during serial passages of the virus in Vero cells or circulation in a host in the field (Kong et


316

al., 2008; Velayudhan et al., 2008). However, the biological significance of the G gene length variation in virus replication, pathogenicity and immunogenicity remains to be defined. In the present study, we developed a reverse genetic system for aMPV-C and generated two Colorado strain-based recombinant viruses containing either the full-length G gene derived from a Canadian goose isolate or a C-terminally truncated G gene of the CO strain. The biological properties of these two G recombinant variants were characterized in Vero cells and SPF turkeys. MATERIALS and METHODS Virus and RNA preparation The aMPV-C Colorado strain (aMPV/CO/96/C) was kindly provided by Dr. Bruce Seal (USDA, ARS, RRC, Athens, GA, USA). The virus was propagated for not more than five times in Vero cells at 370C in 5 % CO2 in D-MEM medium supplemented with 10 % fetal bovine serum (FBS) and antibiotics (Invitrogen, Baltimore, MD) and used as stock. A virulent challenge virus stock was prepared by three serial passages of the aMPV-CO stock in one-week-old SPF turkeys as described previously (Velayudhan et al., 2007). At the third passage, the tracheas of infected turkeys were harvested and homogenized in BHI medium containing antibiotics (Sigma). The supernatant of the tracheal tissues was used as a virulent challenge virus, designated as aMPV-C Tr. Total cellular RNA from infected Vero cells or tracheal tissues was extracted using either Trizol-LS reagent (Invitrogen, Carlsbad, CA) or a MagMAX™ AI/ND Viral RNA Isolation kit (ABI, Austin, TX) according to the manufacturer’s instruction. Construction of full-length cDNA clones Five overlapped cDNA fragments spanning the entire genome of aMPV-CO were generated by RT-PCR amplification of genomic RNA with five pairs of specific primers using a SuperscriptTM One-Step RT-PCR kit with Platinum Taq Hi-Fi Polymerase (Invitrogen, Carlsbad, CA) (Figure 1). These cDNA fragments were then cloned into a modified pBluescript plasmid (Stratagene, La Jolla, CA). One of the assembled clones containing a truncated short G gene (783 nts, coding for 252 aa) was selected by sequencing analysis. This clone was designated as paMPV-C sG. The open reading frame (ORF) of the short G gene in paMPV-C sG was substituted with that of a full-length G gene (1798 nts, coding for 585 aa) from a Canadian goose isolate (Bennett et al., 2005) using an In-FusionTM PCR Cloning kit (Clontech, Mountain View, CA). The resulting clone was named as paMPV-C gG. Construction of plasmids expressing the N, P, M2-1, and L proteins cDNAs coding for the N, P, M2-1, or L protein of the aMPV-CO strain were generated by RT-PCR from viral RNA, and then cloned into a cloning site of pcDNA 3.1/Zeo expression vector (Invitrogen, Carlsbad, CA) (Figure 2). The viral gene expression was under the control of the T7 RNA polymerase promoter. These viral gene plasmids were designated as pN, pP, pM2-1, or pL, respectively.


317

Nucleotide sequencing All RT-PCR products and purified plasmids were sequenced directly with M13 universal primers or gene-specific primers using the Applied Biosystems-PRISM fluorescent big dye sequencing kit and the ABI 3730 DNA sequencer (ABI, Foster City, CA). Nucleotide sequence editing, assembling and comparison were accomplished using the DNASTAR program (Madison, WI). Virus rescue and propagation A mixture of 1 ug of paMPV-C sG or paMPV-C gG, 1 ug of pN, 0.5 ug of pP, 0.1 ug of pL and pM2-1 plasmids was transfected into HEp-2 cells that were infected with MVA/T7 recombinant virus (Wyatt et al., 1995) using LipofectamineTM 2000 reagent according to the manufacturer’s protocol (Invitrogen). A negative control monolayer was transfected as described above, but omitting the pL plasmid, thus not supplying the essential viral polymerase gene. At 6 h post transfection, the cells were washed 1x with PBS and maintained in D-MEM medium supplemented with 2% FBS and antibiotics (Invitrogen). At 72 h post transfection, the transfected HEp-2 cells were scraped and co-cultured onto an existing 50% monolayer of Vero cells. The co-cultured cells were incubated at 370C in 5 % CO2 for 5 days. The rescued virus in the co-cultured cells was harvested by freeze and thawing three times and propagated in Vero cells for 3-5 passages until aMPV-induced cytopathic effects (CPE) were observed. Animal experimental design Forty one-week-old SPF turkey poults were randomly divided into four groups of ten birds. Birds in group 1 received 100 ul of PBS via intranasal (IN) and intraocular (IO) routes per bird as a control. Birds in groups 2, and 3 were inoculated with 100 ul of raMPV-C sG (3.16x106 TCID50/mL) or raMPV-C gG (3.16x106 TCID50/mL) per bird, respectively, via IN and IO routes. Each bird in group 4 was infected with 100 ul of aMPV-C Tr, the challenge virus stock, via IN and IO routes as a positive virus control. After inoculation, birds were observed daily for recording clinical signs of aMPV disease. Intra-tracheal swabs were collected at 3, 5, and 7 days post-inoculation (DPI) from each bird for detection of virus replication or viral RNA shedding in the respiratory tract. Each of the swabs was stored in 1 ml of BHI medium containing antibiotics (Sigma) at -700C. At 14 DPI, serum was taken from each of the birds for detection of antibody responses by ELISA. Immediately after serum collection, birds in all groups were challenged with 100 ul (per bird) of aMPV-C Tr via IN/IO routes. The challenged birds were monitored daily for clinical signs. Tracheal swabs were collected from each of the challenged birds at 3 and 5 days post-challenge (DPC) for detection of virus shedding. Virus titration and detection of viral RNA The titers of various aMPV-C virus stocks and time course samples were determined by infecting Vero cell monolayers with 100 µl of serial 10-fold dilutions of the viruses as previously described (Kapczynski and Sellers, 2003). Titers were calculated by the Reed and Muench method (Reed and Muench, 1938) and expressed as 50% tissue culture infective dose (TCID50). Virus replication or viral RNA shedding in the turkey tracheal tissues was detected by RT-PCR with a pair of aMPV-C N gene-specific primers.


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Detection of antibody response by ELISA Anti-aMPV-C antibody response of turkeys after infection was determined by an ELISA test (Kapczynski, 2004) with some modifications. Sucrose gradient purified aMPV-CO virus was used as antigen. Turkey sera were diluted (1:100) and individually tested in triplicates. Coating, washing, blocking and diluting reagents were purchased from Immunochemistry Technologies (Bloomington, MN). Goat anti-turkey IgG horseradish peroxidase-conjugate and LumiGLO, a luminol-based chemiluminescent substrate, were supplied by KPL, Inc. (Gaithersburg, MD). The chemiluminescence relative light unit (RLU) was measured on a Biotek Synergy HT Microtiter Plate Luminometer (Biotek Instruments, Winooski, VT). A standard negative serum pool (collected from turkeys inoculated with PBS) was included in each plate of the ELISA test as a negative control. The positive anti-aMPV-C serum pool (collected from the virulent challenge virus infected turkeys) was two-fold diluted from 1:100 to 1:3200 and used to establish a standard curve for determining antibody titers of the tested sera. Sera were deemed positive when the mean RLU value was greater than the mean RLU plus 2x standard deviation of the three dilutions (1:100-1:400) of the negative turkey serum pool. RESULTS Generation of recombinant aMPV-C sG and gG gene variants Two FLCs containing a different length of the G gene (Figure 1), and four supporting plasmids expressing the N, P, M2-1, and L proteins of the aMPV-CO strain (Figure 2) were constructed. After co-transfection of the full-length clones and the supporting plasmids in HEp-2 cells and amplification in Vero cells, two aMPV-C infectious viruses were rescued. One of the rescued G gene variants, raMPV-C gG, contained a full-length G gene derived from a Canadian goose isolate (Bennett et al., 2005). The goose G (gG) consists of 1798 nts, coding for 585 aa, which is the same length as that for the aMPV-C Colorado and Minnesota strains reported by Govindarajan and Samal (Govindarajan and Samal, 2004). Whereas the other recombinant G variant, raMPV-C sG, contained a truncated G gene (783 nts) coding for 252 aa, such that the C-terminal 333 aa of the G protein had been deleted. This short G (sG) gene possesses the same length and sequences as that of the Colorado strain reported by Lwamba et al. (Lwamba et al., 2005). The fidelity of the rescued viruses was confirmed by sequencing analysis (data not shown). Biological characterization of the recombinant G variants in cell cultures To determine if the G gene length variation affects virus replication and growth characteristics in vitro, the recombinant G variants and the parental CO strain were examined in Vero cells for growth dynamics and cytopathic effects (CPE). Both G variants induced typical cytopathic effects that were indistinguishable from those seen with the parental aMPV-CO infection (data not shown). Virus titers of samples collected at different time points post-infection showed that the sG variant displayed a similar growth dynamics as the parental aMPV-CO strain. However, the gG appeared to grow slower (about one log10 titer lower) than the sG variant and the parental aMPV-CO strain during the first 72 h post-infection (Figure 3).


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Pathogenic assessments of the recombinant G variants in turkeys Experimental infection of one-week-old SPF turkeys was carried out to evaluate the pathogenic effects of the G gene length variation in vivo. From 3 DPI, the turkey poults infected with the virulent aMPV-C Tr displayed typical clinical signs of the disease, showing nasal exudates when squeezed, nasal discharge, or frothy eyes. These clinical signs gradually disappeared after 8 DPI. However, the birds infected with either the short G or goose G variant did not show any clinical signs (Table 1). RT-PCR data to detect viral RNA in tracheal tissues (Table 1) indicated that lower percentage of birds infected with the short G mutant had detectable virus replication at 3 and 5 DPI (22%, 22%) than the gG variant (55%, 75%) and the challenge virus (100%, 100%). However, at 7 DPI the percentage of birds with detectable virus replication in the sG variant-infected group increased to a similar level (63%) compared to the challenge virus infected group (60%). Antibody response and protection of the recombinant G variant-vaccinated turkeys against challenge As shown in Table 2, at 14 DPI, all birds vaccinated with the gG variant or infected with the challenge virus had positive serum conversion with a mean titer of 6.6 and 7.9 log2, respectively. Six of eight (75%) turkeys vaccinated with the sG mutant had positive antibody response with a mean titer of 6.4 log2. All sera collected from unvaccinated birds (inoculated with PBS) were negative. To examine if the immune response of turkeys induced by the G variants was protective against aMPV-C disease, the vaccinated birds were challenged with aMPV-C Tr. As shown in Table 3, seven of eight (87%) birds vaccinated with the sG variant, and eight of nine (89%) birds vaccinated with the gG variant were protected against the challenge. Whereas all unvaccinated birds (PBS control) showed typical clinical signs of the disease. The birds inoculated with aMPV-C Tr were fully protected against the same virus challenge. Detection of challenge virus replication in tracheal tissue (Table 3) showed that 63% of the sG vaccinated birds and 89% of the gG vaccinated birds had a detectable level of virus RNA shedding at 3 DPC. After 5 DPC the challenge virus RNA shedding was decreased to 22% for the gG vaccinated birds or below a detectable level for the sG vaccinated birds. In contrast, all unvaccinated birds had challenge virus shedding at 3 and 5 DPC, whereas the aMPV-C Tr virus inoculated birds did not have any detectable level of virus shedding following challenge. DISCUSSION In the present study, we developed a reverse genetics system for aMPV-C, and generated two recombinant G gene variants, representing the full-length G (585 aa) and a C-terminally truncated short G (252 aa) viruses. The in vitro biological characterization showed that both G variants were viable in Vero cells and induced typical cytopathic effects as seen with the parental virus although the gG growth was slightly delayed in the first 72 hours of infection. This demonstrated that the C-terminal 333 aa, about 60% of the full size G protein, is not essential for virus viability in Vero cells. This is not surprise since a complete G gene deletion from a rescued aMPV subgroup A recombinant virus did not decrease virus viability in Vero cells


320

(Naylor et al., 2004). Interestingly, to date all pneumovirus G proteins studied by using the reverse genetic technology seem to be dispensable for virus viability in cell cultures (Biacchesi et al., 2004; Naylor et al., 2004; Techaarpornkul et al., 2001). This suggests that an alternative surface protein, most likely the F protein, of members of the pneumovirus subfamily, may play a role in mediating virus attachment to the cultured cells for infection. The clinical signs observed in the animal experiment have shown that both G variants were attenuated in turkeys and caused no clinical disease. The exact reasons for this attenuation are currently not known. It is possible that unintentional amino acid changes were introduced into the viral genome during construction of rescued viruses. An example for this is that the sG variant has two aa changes found in the M and L proteins. Although they are not located in any known functional domains (Poch et al., 1990), we can not rule out the possibility of their contribution to the attenuation. It has also been reported that cloned virus population for negative strand RNA viruses seems to be less virulent than its unpurified quasi-species population (Duarte et al., 1994a; Duarte et al., 1994b). Since our G variant stocks were prepared by propagation in Vero cells at a low level of passages (5-8 times), the virus population may still be relatively homogenous. The detection of viral RNA shedding in the tracheal tissue revealed that both G variants were infectious in turkeys, but attenuated as evidenced by the lower percentage (22% to 75%) of birds with detectable viral RNA than that (100%) infected with the virulent challenge virus at 3 and 5 DPI. The short G variant appeared to grow slower at early days of infection than the full-length G variant. These results demonstrated that the 333 aa extracellular domain of the G protein is not essential for virus viability in turkeys, but it may be required for efficient virus replication. This is consistent with an earlier observation that a C-terminal truncation of the G gene in a rescued respiratory syncytial virus (RSV) mutant was viable but attenuated in mice (Elliott et al., 2004). Our data clearly showed that both G variants were immunogenic and induced anti-aMPV-C antibody response. However, two of eight birds inoculated with the sG variant did not have a positive serum conversion. This can be explained with a lower level and delayed sG virus replication in these birds as it was detected by RT-PCR from the tracheal tissue. Together, the lower percentages of virus shedding and positive serum conversion for the sG variant compared to those observed with the gG variant indicated that the deleted region from the G gene may play a role in enhancing virus replication and antibody response, perhaps through increasing the attachment specificity. Importantly, both the sG and gG variants provided a significant protection against the virulent aMPV-C Colorado strain. About 90% of the challenged birds did not show any clinical signs and challenge virus shedding was reduced at 3 and 5 DPC. Despite the 100% positive serum conversion induced by the gG variant, it did not provide a full protection against the virulent Colorado strain challenge. About 10% of the gG vaccinated birds showed clinical signs and 90% of the birds shed detectable challenge virus RNA at 3 DPC. It is possible that a small antigenic difference in the G protein between the goose isolate and the Colorado


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strain may account for the incomplete protection (Alvarez et al., 2004; Bennett et al., 2005). In conclusion, we successfully developed a reverse genetics system for examining the effects of the G gene length variation among aMPV-C isolates on virus viability, virulence and immunity. Our results show that the C-terminal 333 aa (more than 60% of the extracellular domain) of the viral attachment G protein is not essential for virus viability in vitro and in vivo, but may play a role in enhancing virus attachment specificity and immunity in a natural host. Both the short and the full length G containing variants were attenuated in turkeys, yet immunogenic and induced immune response that provided a significant protection against virulent aMPV-CO challenge. ACKNOWLEDGEMENTS The authors wish to thank Xiuqin Xia and Fenglan Li for excellent technical assistance, Bernard Moss for the gifts of MVA/T7 recombinant virus, and Melissa Scott and Joyce Bennett for performing the automated nucleotide sequencing. This research was supported by USDA, ARS CRIS project 6612-32000-056-00D. REFERENCES Alvarez, R., Jones, L.P., Seal, B.S., Kapczynski, D.R. and Tripp, R.A. (2004)

Serological cross-reactivity of members of the Metapneumovirus genus. Virus Res. 105(1), 67-73.

Alvarez, R., Lwamba, H.M., Kapczynski, D.R., Njenga, M.K. and Seal, B.S. (2003) Nucleotide and predicted amino acid sequence-based analysis of the avian metapneumovirus type C cell attachment glycoprotein gene: phylogenetic analysis and molecular epidemiology of U.S. pneumoviruses. J. Clin. Microbiol. 41(4), 1730-5.

Bennett, R.S., LaRue, R., Shaw, D., Yu, Q., Nagaraja, K.V., Halvorson, D.A. and Njenga, M.K. (2005) A wild goose metapneumovirus containing a large attachment glycoprotein is avirulent but immunoprotective in domestic turkeys. J. Virol. 79(23), 14834-14842.

Biacchesi, S., Skiadopoulos, M.H., Yang, L., Lamirande, E.W., Tran, K.C., Murphy, B.R., Collins, P.L. and Buchholz, U.J. (2004) Recombinant human Metapneumovirus lacking the small hydrophobic SH and/or attachment G glycoprotein: deletion of G yields a promising vaccine candidate. J Virol 78(23), 12877-87.

Cook, J.K. (2000) Avian rhinotracheitis. Rev. Sci. Tech. 19(2), 602-13. Duarte, E.A., Novella, I.S., Ledesma, S., Clarke, D.K., Moya, A., Elena, S.F.,

Domingo, E. and Holland, J.J. (1994a) Subclonal components of consensus fitness in an RNA virus clone. J Virol 68(7), 4295-301.

Duarte, E.A., Novella, I.S., Weaver, S.C., Domingo, E., Wain-Hobson, S., Clarke, D.K., Moya, A., Elena, S.F., de la Torre, J.C. and Holland, J.J. (1994b) RNA virus quasispecies: significance for viral disease and epidemiology. Infect Agents Dis 3(4), 201-14.


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Elliott, M.B., Pryharski, K.S., Yu, Q., Parks, C.L., Laughlin, T.S., Gupta, C.K., Lerch, R.A., Randolph, V.B., LaPierre, N.A., Dack, K.M. and Hancock, G.E. (2004) Recombinant respiratory syncytial viruses lacking the C-terminal third of the attachment (G) protein are immunogenic and attenuated in vivo and in vitro. J Virol 78(11), 5773-83.

Govindarajan, D. and Samal, S.K. (2004) Sequence analysis of the large polymerase (L) protein of the US strain of avian metapneumovirus indicates a close resemblance to that of the human metapneumovirus. Virus Res. 105(1), 59-66.

Govindarajan, D., Yunus, A.S. and Samal, S.K. (2004) Complete sequence of the G glycoprotein gene of avian metapneumovirus subgroup C and identification of a divergent domain in the predicted protein. J. Gen. Virol. 85(Pt 12), 3671-3675.

Juhasz, K. and Easton, A.J. (1994) Extensive sequence variation in the attachment (G) protein gene of avian pneumovirus: evidence for two distinct subgroups. J. Gen. Virol. 75, 2873-80.

Kapczynski, D.R. (2004) Development of a virosome vaccine for protection in turkeys against avian metapneumovirus subtype C. Avian Dis. 48(3), 332-343.

Kapczynski, D.R. and Sellers, H.S. (2003) Immunization of turkeys with a DNA vaccine expressing either the F or N gene of avian metapneumovirus. Avian Dis 47(4), 1376-83.

Kong, B.W., Foster, L.K. and Foster, D.N. (2008) Species-specific deletion of the viral attachment glycoprotein of avian metapneumovirus. Virus Res 132(1-2), 114-21.

Lwamba, H.C., Alvarez, R., Wise, M.G., Yu, Q., Halvorson, D., Njenga, M.K. and Seal, B.S. (2005) Comparison of the full-length genome sequence of avian metapneumovirus subtype C with other paramyxoviruses. Virus Res. 107(1), 83-92.

Naylor, C.J., Brown, P.A., Edworthy, N., Ling, R., Jones, R.C., Savage, C.E. and Easton, A.J. (2004) Development of a reverse-genetics system for Avian pneumovirus demonstrates that the small hydrophobic (SH) and attachment (G) genes are not essential for virus viability. J. Gen. Virol. 85(Pt 11), 3219-27.

Patnayak, D.P., Sheikh, A.M., Gulati, B.R. and Goyal, S.M. (2002) Experimental and field evaluation of a live vaccine against avian pneumovirus. Avian Pathol. 31(4), 377-382.

Poch, O., Blumberg, B.M., Bougueleret, L. and Tordo, N. (1990) Sequence comparison of five polymerases (L proteins) of unsegmented negative-strand RNA viruses: theoretical assignment of functional domains. J Gen Virol 71 ( Pt 5), 1153-62.

Reed, L.J. and Muench, H. (1938) A simple method for estimating fifty percent endpoints. American Journal of Hygiene 27, 493-497.

Techaarpornkul, S., Barretto, N. and Peeples, M.E. (2001) Functional analysis of recombinant respiratory syncytial virus deletion mutants lacking the small hydrophobic and/or attachment glycoprotein gene. J Virol 75(15), 6825-34.

Toquin, D., Guionie, O., Jestin, V., Zwingelstein, F., Allee, C. and Eterradossi, N. (2006) European and American subgroup C isolates of avian metapneumovirus belong to different genetic lineages. Virus Genes 32(1), 97-103.


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Velayudhan, B.T., Noll, S.L., Thachil, A.J., Shaw, D.P., Goyal, S.M., Halvorson, D.A. and Nagaraja, K.V. (2007) Development of a vaccine-challenge model for avian metapneumovirus subtype C in turkeys. Vaccine 25(10), 1841-7.

Velayudhan, B.T., Yu, Q., Estevez, C.N., Nagaraja, K.V. and Halvorson, D.A. (2008) Glycoprotein gene truncation in avian metapneumovirus subtype C isolates from the United States. Virus Genes 37(2), 266-72.

Wyatt, L.S., Moss, B. and Rozenblatt, S. (1995) Replication-deficient vaccinia virus encoding bacteriophage T7 RNA polymerase for transient gene expression in mammalian cells. Virology 210(1), 202-5.

Table 1. Observation of clinical signs and detection of virus shedding at different days post inoculation Inoculums No of birds showing clinical signs No and % of birds shedding viral RNA 3 5 7 10 3 5 7 PBS 0/10 0/10 0/10 0/10 0/10 (0%) 0/10 (0%) 0/10 (0%) aMPV-CO Tr 7/10 9/10 8/10 0/10 10/10 (100%) 10/10 (100%) 6/10 (60%) raMPV-C sG 0/9 0/9 0/9 0/9 2/9 (22%) 2/9 (22%) 5/8 (63%) raMPV-V gG 0/10 0/10 0/10 0/10 5/9 (55%) 6/8 (75%) 1/8 (13%)

Table 2. Serum antibody response in turkeys at day 14 post inoculation Inoculums No of Ab positive sera Mean ± s.d. and % in group (log2) PBS 0/10 (0%) aMPV-CO Tr 9/9 (100%) 7.9 ± 0.7 raMPV-C sG 6/8 (75%) 6.4 ± 0.4 raMPV-C gG 8/8 (100%) 6.6 ± 0

Table 3. Observation of clinical signs and detection of virus shedding at different days post challenge

Inoculums No of birds showing clinical signs No and % of birds shedding viral RNA 3 5 7 9 3 5 PBS 9/10 9/10 2/10 0/10 10/10 (100%) 10/10 (100%) aMPV-CO Tr 0/9 0/9 0/9 0/9 0/9 (0%) 0/9 (0%) raMPV-C sG 1/8 1/8 0/8 0/8 5/8 (63%) 0/8 (0%) raMPV-V gG 0/9 1/9 1/9 0/9 8/9 (89%) 2/9 (22%)


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Figure 1. Construction of aMPV-C full-length cDNA clones. Five cDNA fragments, marked with F1, F2, F3, F4, and F5, respectively, were generated by RT-PCR from genomic RNA of the aMPV-CO strain. These cDNA fragments were sequentially cloned and assembled into a modified pBluescript vector by using naturally occurred or engineered restriction enzyme sites that are marked by black arrows. The resulting full-length cDNA clone (FLC) is designated as paMPV-C sG. The truncated G gene in paMPV-C sG, highlighted with the pink filling, was substituted with the full-length goose G gene, highlighted with the green filling, by RT-PCR. The resulting FLC is named as paMPV-C gG. T7 promoter direction is indicated by a red bold arrow. Hepatitis delta virus ribozyme (HDV Rz) cleavage site is marked by a black arrow. T7Φ represents the T7 terminator sequence.

N L G M F P M2 SH

TTrr

LLee aMPV-CO genome

N L G M F P M2 SH TTrr LLee

HDV Rz T7 T7

Cleavage site AatII

AatII

N L gG M F P M2 SH TTrr LLee

HDV Rz T7 T7

Cleavage site

3’ 5’

paMPV-C sG

paMPV-C gG

Sub-cloning and assembly

RT-PCR

N M P

NNoottII//TT77

F

NNhheeII

G M2 SH

KKppnnII

L

PPaaccII AAvvrrIIII NNaarrII

Swapping truncated G with full-length goose G

TTrr

F1 F2 F3 F4 F5

LLee


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aMPV-CO genome

Figure 2. Construction of viral gene expression plasmids. cDNAs coding for the N, P, M2-1, or L protein were generated by RT-PCR from viral RNA of the aMPV-CO strain, and cloned into a cloning site in pcDNA3.1 expression vector. The viral gene expression was under the control of T7 RNA polymerase promoter (T7), and followed by the bovine growth hormone poly A site (BGH poly A). T7 promoter direction is indicated by a red bold arrow. The resulting viral gene expression plasmids are designated as pN, pP, pM2-1, or pL, respectively.

Growth dynamics of raMPV in vero cells

0

1

2

3

4

5

6

7

8

0 12 24 36 48 60 72 96 120 144

Hours post infection

Tite

r (lo

g1oT

CID

50/m

l)

aMPV-CO

raMPV-C sG

raMPV-C gG

Figure 3. Growth dynamics of the recombinant G variants and the aMPV-CO strain in cells. Vero cell monolayers were infected with aMPV-CO, raMPV-C sG, or raMPV-C gG at 0.1 m.o.i.. At every 12 hours post infection, the monolayers were harvested. Virus titers at each time point, in duplicate, were determined by TCID50 titration in Vero cells. The mean titer of the two tests at each time point is expressed in log10 TCID50/mL.

pN

pP

pL

pM2-1

poly A site

poly A site poly A site

N gene

M2-1 gene P gene

NN LL GG MM FF PP MM22 SSHH TTrr LLee 33’’ 55’’

poly A site L gene T7 T7

T7 T7

pN

pP

pL

pM2-1

BGH poly A

BGH poly A BGH poly A

N gene

M2-1 gene P gene

NN LL GG MM FF PP MM22 SSHH TTrr LLee 33’’ 55’’

BGH poly A L gene T7 T7

T7


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AVIAN METAPNEUMOVIRUS – NEARLY 30 YEARS OF VACCINATION

COOK JKA

Huntingdon, Cambs. UK. PE29 1XQ

SUMMARY Avian metapneumovirus (aMPV) was first reported in South Africa in the late 1970s, in Western Europe some 5 years later and then spread to many other parts of the world, eventually occurring in the USA, but not until 1996. aMPV causes a clearly defined disease in turkeys and is also involved as a pathogen in chickens, but its role in complex disease situations in that species is less clearly defined. Vaccines were quickly developed to control and prevent infections caused by this virus. This paper will review the different types of vaccine developed, their use in both turkey and chickens and consider how successful they have been. INTRODUCTION In the late 1970s an apparently new and severe respiratory infection was reported for the first time in turkeys in South Africa (Buys & Du Preez, 1980). The disease, characterised by sneezing, tracheal râles, swollen infraorbital sinuses and nasal and often frothy ocular discharge, was named turkey rhinotracheitis (TRT). This disease had a devastating effect on the turkey industry of South Africa, from which it never recovered. Several years later, a disease with similar clinical signs was reported in France (Giraud et al., 1986) and then in England where the causal agent was isolated (McDougall & Cook, 1986; Wilding et al., 1986; Wyeth et al., 1986) and characterised as a pneumovirus (Cavanagh & Barrett, 1988). It was the first, and is still, the only avian pneumovirus (APV) to have been described and is the type strain of a new genus, Metapneumovirus. Having initially been called TRT, the virus then became known as APV, but is now known by the more correct name, avian metapneumovirus (aMPV). Shortly after its appearance in England and France, aMPV was also reported from other parts of Europe, the Middle and Far East and Latin America and it was soon recognised as a major disease threat in both turkeys and chickens in many parts of the world. However, it was not until 1996 that it was reported in USA, in the state of


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Colorado (Cook et al., 1999), and subsequently in Minnesota (Goyal et al., 2000). Currently, Australia and Canada are probably the only two major areas of the world where the disease has not been reported. Only a single serotype is recognised, but based on molecular sequencing of the G glycoprotein (Juhasz & Easton, 1994) and neutralisation tests using monoclonal antibodies (Mabs) that recognised the G glycoprotein (Cook et al., 1993a) two subtypes, designated A and B were identified. Initially it was believed that subtype A strains were found in England and South Africa and subtype B ones in mainland Europe (Cook et al., 1993a). However, it is now clear that subtype A strains have been present in Europe for a long time (van de Zande et al., 1998) and that subtype B strains are present in England (Naylor et al., 1997a). The isolate from USA, belongs to a distinct subtype, designated subtype C (Seal, 1998). A fourth subtype (D) has been reported in Muscovy ducks in France (Toquin et al., 1999). CONTROL OF aMPV INFECTIONS It quickly became apparent that control of aMPV infections by means of strict hygiene procedures combined with the use of good biosecurity was not adequate and the devastating effect of the disease on the South African turkey industry indicated that urgent intervention was necessary. Maternally derived immunity does not protect against infection (Naylor et al., 1997b). Eradication of the disease was not generally possible, probably due to the size and complexity of the poultry industry in most areas. However, eradication of aMPV in the initial outbreak in Colorado, USA did prove effective. The reasons for this probably included the relatively small size and isolated location of the outbreak there, combined with availability of expertise concerning the epidemiology of the disease in England. However, whilst eradicated from the turkey population in Colorado, aMPV infections continued to be a serious problem within the much larger turkey industry in Minnesota and some neighbouring states. Because of the urgent need to control the devastating effects of aMPV infections in the turkey population, as an interim measure, controlled exposure and the use of autogenous vaccines was commonly employed both in Europe and USA, but development of effective vaccines became a major requirement. VACCINE DEVELOPMENT Very soon after the disease was reported in South Africa, the first prototype vaccine was developed there (Buys et al., 1989a). The virus was attenuated by serial passage of sinus material from infected turkeys via the chorioallantoic (CAM) route, CAMs being collected 7 days later for subsequent passage. Material collected after 17 CAM passages was used as a prototype vaccine with promising results. However, no further development work was undertaken because the disease had already led to the serious decline of the turkey industry in South Africa. Once aMPV infection was diagnosed in Western Europe, vaccine development became the urgent goal of several research groups. The common approach was to attempt attenuation of the virus by serial passage on a number of different substrates including chick embryo fibroblasts and VERO cultures (Williams et al., 1991; Giraud et al., 1987) or alternate passage in embryonated eggs and tracheal organ cultures


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[TOC] (Cook et al., 1989; Cook & Ellis, 1990). Interestingly, continual passage in TOC alone, even as many as 100 times, resulted in no attenuation of the virus. The major Pharmaceutical Companies continued development work and a number of commercial vaccines were quickly in production and very widely used. When administered carefully and correctly they provided and continue to provide, excellent protection for turkeys of all ages, being used in meat turkeys and to prime future layers and breeders for injection prior to onset of lay of the inactivated vaccines that were soon developed. FACTORS AFFECTING SUCCESSFUL VACCINATION These vaccines provided immediate benefit in reducing the welfare and economic effects of aMPV infections in turkey flocks. However, it soon became clear that, to those not used to vaccinating turkeys against respiratory pathogens at a very young age, this species presented new challenges. Vaccination of turkeys in the hatchery, or at the farm, whilst still in boxes, gave the best results since the importance of very careful administration to ensure that all birds receive an adequate dose of vaccine at the same time quickly became apparent. Vaccination is usually by spray or ideally by eye drop, but for the future, the in ovo route might prove beneficial (Worthington et al., 2003; Hess et al., 2004). A single vaccination may be sufficient to protect meat turkeys throughout their life. However, reinfection can occur late in life and in some situations meat turkeys that are reared beyond 10 to 12 weeks are revaccinated. Apparent vaccination failures were attributed to a number of factors – over attenuation of the virus strains (inability to induce a good immune response), under attenuation (causing too severe a reaction), poor vaccine administration, leading to rolling infections and the suggestion of reversion to virulence (Catelli et al., 2006). Despite these possible problems, vaccination continues to be seen as highly beneficial, provided careful attention is paid to the method of administration. Whilst there is strong evidence of excellent cross protection between the A and B subtypes (Cook et al., 1995; Eterradossi et al., 1995), the suggestion arose that apparent vaccine failures may have been due to infections with the subtype not included in the vaccine. By this time, live attenuated vaccines of both subtypes had been developed and so it became possible to use the two types of vaccine on the same farm. However, this raises the possibility of persistence of both subtypes in a particular area. With the emergence of the subtype C virus in USA, it was soon shown that there is good cross protection between the A and B subtype vaccines and the Colorado isolate (Cook et al., 1999). However, of necessity, vaccine development work continued in USA independently of that being undertaken in Europe. Attempts to develop live attenuated vaccines involved serial passage of the virus using a number of different methods (Jirjis et al., 2001; Patnayak et al., 2002; 2003). The 63rd passage of VERO-adapted virus was developed into the live-attenuated commercial vaccine that is currently available for use in the endemic areas in Minnesota and neighbouring states (J. Sharma, personal communication). Interestingly, whilst in Europe it was considered essential to aim for 100% vaccination of a flock, in USA successful field trial results were reported in which only two birds per 1000, in turkey


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flocks each with 20,000-50,000 birds, were given the vaccine (Gulati et al., 2001). A further difference between USA and elsewhere was that whilst extensive aMPV vaccination of turkeys quickly became routine in England and Europe, only affected areas or farms were vaccinated in Minnesota and vaccine administration was reduced as outbreaks came under control (K Nagaraja, personal communication). Developments in the field of molecular biology have led to different vaccination strategies being considered. The F protein gene of aMPV, expressed in a fowl poxvirus vector (Qingzong et al., 1994) or a DNA vaccine expressing either the F or N gene (Kapczynski & Sellers, 2003) have been shown experimentally to provide protection but have apparently not been pursued further. The development of a reverse genetics system for aMPV (Naylor et al., 2004) offers the opportunity for novel approaches to vaccine development to be applied in the future. THE DISEASE IN CHICKENS Shortly after the initial reports of infection in turkeys in South Africa a similar condition was reported in chickens there (Buys et al., 1989b). Whilst in South Africa and some other countries, for example parts of Latin America, aMPV causes a distinct disease condition in chickens, its role elsewhere as a pathogen in that species has remained unclear. In breeders there is evidence that it behaves as a primary pathogen, but in broilers it is probable that it plays a role in complex disease syndromes, for example Swollen Head Syndrome, rather than being a primary pathogen. However, the benefits seen from using aMPV vaccines in chickens (see below) support the evidence that it can be an important pathogen in chickens of all ages. VACCINATION OF CHICKENS TO CONTROL aMPV INFETIONS The difficulty in confirming the involvement of aMPV in clinical conditions reported in chickens, particularly in broilers, has led to often erroneous reports that aMPV vaccines are not efficacious in controlling infections in chickens. However, where aMPV is known to be involved in a disease syndrome, the use of aMPV vaccines has proved highly efficacious, particularly in layers and breeders. Nevertheless, good diagnosis is essential since aMPV vaccines will not improve the performance of flock where aMPV is not involved in the syndrome reported. Initially there was some concern that aMPV strains isolated from turkeys and from chickens were antigenically different. However, whilst there may be differences in the susceptibility of chickens to isolates for the two species, in terms of antigenicity differences do not exist (Cook et al., 1993b) and under experimental conditions, the commercially available vaccines protect both species equally well. However, because of the possibility that isolates from chickens may replicate more efficiently in that species (Cook et al., 1993b) the perceived advantage of using chicken-origin strains led to development of vaccines incorporating strains of aMPV isolated from chickens for use in that species. A further complicating factor in chickens is the need to incorporate Infectious Bronchitis (IB) and Newcastle Disease (ND) vaccines in the control programme.


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There is a risk of interference between these viruses and aMPV since they all target the same cells in the respiratory tract, although published data do not agree on whether or not this is a serious problem in the field (Cook et al., 2001; Ganapathy et al., 2006;Tarpey et al., 2007). However, it is advisable, in commercial flocks, to consider leaving an interval of approximately one week between day-old vaccination against IB and ND and administration of aMPV vaccine. PROTECTION OF LAYING AND BREEDING BIRDS In addition to the live attenuated vaccines, inactivated aMPV vaccines are widely used for the protection of breeding and laying turkeys and chickens. These may be monovalent vaccines or incorporate antigens to other pathogens in multivalent vaccines. Highly effective inactivated aMPV vaccines are now widely used in vaccination programmes following priming with live-attenuated vaccines. When performed carefully and correctly such programmes provide excellent lifelong protection against drops in egg production involving aMPV in both turkeys and chickens (Giraud et al., 1987; Cook et al., 1996; 2000). It has been shown both experimentally (Cook et al., 2000) and under field conditions that the use of inactivated vaccines without live priming can provide good protection against the effect of aMPV challenge on egg production and egg quality, although some clinical signs may be seen for a short time after challenge. Therefore, in some countries where it is difficult to licence live-attenuated aMPV vaccine, control programmes including inactivated vaccine as the only aMPV component are used successfully. CONCLUSIONS In the 40 years since aMPV was first described in turkeys and then in chickens in South Africa, rapid strides have been made in minimising the damaging effects of the virus. In chickens the situation has never been clear-cut, although aMPV vaccines have certainly been beneficial in situations where aMPV is confirmed as being involved in the disease syndrome reported. Currently, vaccines are not used routinely in broilers in Europe, although they continue to be considered beneficial in breeding and laying chickens. However, in turkeys the continual use of the vaccines available has led to the disease being well controlled, provided that the vaccines are applied carefully and correctly. This may be particularly true in USA, where aMPV infections are now no longer considered to be a major concern. It is interesting to speculate on this difference between USA and elsewhere and to wonder if the different situation in USA is a result of the different aMPV type found there, the different vaccination policy used, or to the absence of the virus from the chicken population in that country.


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REFERENCES Buys, S.B. & Du Preez, J.H. (1980). A preliminary report on the isolation of a virus

causing sinusitis in turkeys in South Africa and attempts to attenuate the virus. Turkeys, 28, 36.

Buys, S.B., Du Preez, J.H. & Els, H. J. (1989a). The isolation and attenuation of a virus causing rhinotracheitis in turkeys in South Africa. Onderstepoort Journal of Veterinary Research, 56, 87-98.

Buys, S. B., Du Preez, J.H. & Els, H. J. (1989b). Swollen head syndrome in chickens: a preliminary report on the isolation of a possible aetiological agent. Journal of the South African Veterinary Association, 60, 221-222.

Catelli, E., Cecchinato, M., Savage, C.E., Jones, R.C. & Naylor, C.J. (2006). Demonstration of loss of attenuation and extended field persistence of a live avian metapneumovirus vaccine. Vaccine, 24, 6476-6482.

Cavanagh, D. & Barrett, T. (1988). Pneumovirus-like characteristics of the mRNA and proteins of turkey rhinotracheitis virus. Virus Research, 11, 241-256.

Cook, J.K.A. & Ellis, M.M. (1990). Attenuation of turkey rhinotracheitis virus by alternative passage in embryonated chicken eggs and tracheal organ cultures. Avian Pathology, 19, 181-185.

Cook, J.K.A., Ellis, M.M., Dolby, C.A., Holmes, H.C., Finney, P.M. & Huggins, M.B. (1989). A live attenuated turkey rhinotracheitis virus vaccine. 1. Stability of the attenuated strain. Avian Pathology, 18, 511-522.

Cook, J.K.A., Jones, B.V., Ellis, M.M., Jing Li. & Cavanagh, D. (1993a). Antigenic differentiation of strains of turkey rhinotracheitis virus using monoclonal antibodies. Avian Pathology, 22, 257-273.

Cook, J.K.A., Kinloch, S. & Ellis, M.M. (1993b). In vitro and in vivo studies in chickens and turkeys on strains of turkey rhinotracheitis virus isolated from the two species. Avian Pathology, 22, 157-170.

Cook, J.K.A., Huggins, M. B., Woods, M.A., Orbell, S.J. & Mockett, A.P.A. (1995). Protection provided by a commercially available vaccine against different strains of turkey rhinotracheitis virus. Veterinary Record, 136, 392-393.

Cook, J.K.A., Orthel, F., Orbell, S., Woods, M.A. & Huggins, M.B. (1996). An experimental turkey rhinotracheitis (TRT) infection in breeding turkeys and the prevention of its clinical effects using live-attenuated and inactivated TRT vaccines. Avian Pathology, 25, 231-243.

Cook, J. K. A., Huggins, M. B., Orbell, S.J. & Senne, D. A. (1999). Preliminary antigenic characterization of an avian pneumovirus isolated from commercial turkeys in Colorado, USA. Avian Pathology, 28, 607-617.

Cook, J. K. A., Chesher, J., Orthel, F., Woods, M. A., Orbell, S. J., Baxendale, W. & Huggins, M. B. (2000). Avian pneumovirus infection of laying hens: experimental studies. Avian Pathology, 29, 545-556.

Cook, J. K. A., Huggins, M. B., Orbell, S. J., Mawditt, K. & Cavanagh, D. (2001). Infectious bronchitis virus vaccine interferes with the replication of avian pneumovirus vaccine in domestic fowl. Avian Pathology, 30, 233-242.

Eterradossi, N., Toquin, D., Guittet, M. & Bennejean, G. (1995). Evaluation of different turkey rhinotracheitis viruses used as antigens for serological testing following live vaccination and challenge. Journal of Veterinary Medicine, B, 42, 175-186.


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Ganapathy, K., Todd, V., Cargill, P., Montiel, E. & Jones, R.C. (2006). Interaction between a live avian pneumovirus vaccine and two different Newcastle disease virus vaccines in broiler chickens with maternal antibodies to Newcastle disease virus. Avian Pathology, 35, 429-434

Giraud, P., Bennejean, G., Guittet, M. & Touquin, D. (1986). Turkey rhinotracheitis in France: Preliminary investigations on a ciliostatic virus. Veterinary Record, 119, 606-607.

Giraud, P., Le Gros F.X., Guittet, M. & Bennejean, G. (1987). Turkey rhinotracheitis: isolation of a viral agent and first trials with experimental inactivated or attenuated vaccines. In: Proceedings of the 36th Western Poultry Disease Conference, Davis, California, p 94.

Goyal, S.M., Chiang, S-J., Dar, A. M., Nagaraja, K. V., Shaw, D. P., Halvorson, D. A. & Kapur, V. (2000). Isolation of avian pneumovirus from an outbreak of respiratory illness in Minnesota turkeys. Journal of Veterinary Diagnostic Investigation, 12, 166-168.

Gulati, B.R., Patnayak, D.P., Sheikh, A.M., Poss, P.E. & Goyal, S.M. (2001). Protective efficacy of high-passage avian pneumovirus (APV/MN/turkey/1-a/97) in turkeys. Avian Diseases, 45, 593-597.

Hess, M., Huggins, M. B. & Heincz, U. (2004). Hatchability, serology and virus excretion following in ovo vaccination of chickens with an avian metapneumovirus vaccine. Avian Pathology, 33, 576-580.

Jirjis, F. F., Noll, S.L., Martin, F., Halvorson, D. A., Nagaraja, K. V. & Shaw, D. P. (2001). Vaccination of turkeys with an avian pneumovirus isolate from the United States. Avian Diseases, 45, 1006-1013.

Juhasz, K. & Easton, A.J. (1994). Extensive sequence variation in the attachment (G) protein gene of avian pneumovirus: evidence for two distinct subgroups. Journal of General Virology, 75, 2873-2880.

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McDougall, J.S. & Cook, J.K.A. (1986). Turkey rhinotracheitis: Preliminary investigations. Veterinary Record, 118, 206-207.

Naylor, C., Shaw, K., Britton, P. & Cavanagh, D. (1997a). Appearance of type B avian pneumovirus in Great Britain. Avian Pathology, 26, 327-338.

Naylor, C.J., Worthington, K.J. & Jones, R.C. (1997b). Failure of maternal antibodies to protect young turkey poults against challenge with turkey rhinotracheitis virus. Avian Diseases, 41, 968-971.

Naylor, C.J., Brown, P.A., Edworthy, N., Ling, R., Jones, R.C., Savage, C.E. & Easton, A.J. (2004). Development of a reverse-genetics system for avian pneumovirus demonstrates that the small hydrophobic (SH) and attachment (G) genes are not essential for virus viability. Journal of General Virology, 85, 3219-3227.

Patnayak, D.P., Sheikh, A.M., Gulati, B.R, & Goyal, S.M. (2002). Experimental and field evaluation of a live vaccine against avian pneumovirus. Avian Pathology, 31, 377-382.

Patnayak, D.P., Gulati, B.R., Sheikh, A.M. & Goyal, S.M. (2003). Cold adapted avian pneumovirus for use as live attenuated vaccine in turkeys. Vaccine, 28, 1371-1374.


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Qingzhong, Y., Barrett, T., Brown, T. D. K., Cook, J. K. A., Green, P., Skinner, M. A. & Cavanagh, D. (1994). Protection against turkey rhinotracheitis pneumovirus (TRTV) induced by a fowlpox virus recombinant expressing the TRTV fusion glycoprotein (F). Vaccine, 12, 569-573.

Seal, B. (1998). Matrix protein gene nucleotide and predicted amino acid sequence demonstrate that the first US avian pneumovirus isolate is distinct from European strains. Virus Research, 58, 45-52.

Tarpey, I., Huggins, M.B. & Orbell, S.J. (2007). The efficacy of an avian metapneumovirus vaccine applied simultaneously with infectious bronchitis and Newcastle disease virus vaccines to specific-pathogen-free chickens. Avian Diseases, 51, 594-596.

Toquin, D., Bäyon-Auboyer, M. H., Eterradossi, N. & Jestin, V. (1999). Isolation of a pneumovirus from a Muscovy duck. Veterinary Record, 145, 680.

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Wilding, G. P., Baxter-Jones, C. & Grant, M. (1986). Ciliostatic agent found in rhinotracheitis. The Veterinary Record, 118, 735.

Williams, R.A., Savage, C.E. & Jones, R.C. (1991). Development of a live attenuated vaccine against turkey rhinotracheitis. Avian Pathology, 20, 45-55.

Worthington, K.J., Sargent, B.A., Davelaar, F.G., & Jones, R.C. (2003). Immunity to avian pneumovirus infection in turkeys following in ovo vaccination with an attenuated vaccine. Vaccine, 21,1355-1362.Wyeth, P.J., Gough, R., Chettle, N. & Eddy, R. (1986). Preliminary observations on a virus associated with turkey rhinotracheitis. Veterinary Record, 119, 139.


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FIELD AVIAN METAPNEUMOVIRUS EVOLUTION AVOIDING VACCINE INDUCED IMMUNITY

CATELLI E1, CECCHINATO M2, LUPINI C1, RICCHIZZI E1and NAYLOR CJ3

1 Dipartimento di Sanità Pubblica Veterinaria e Patologia Animale, University of Bologna, Via Tolara di Sopra, 50, 40064 Ozzano Emilia (BO), Italy;

2 Dipartimento di Sanità Pubblica, Patologia Comparata e Igiene Veterinaria University of Padua, Viale dell’Università, 16, 35020 Legnaro (PD), Italy;

3 Department of Veterinary Pathology, Jordan Building, Leahurst, Neston, Cheshire, CH64 7TE, United Kingdom

SUMMARY Infection of turkeys and chickens with Avian Metapneumovirus (AMPV) leads to disease and serious economic losses in unprotected birds. In the late 1980s, live AMPV vaccines became available in Europe and these largely brought the disease under control. However disease has still been encountered and much of this remains unexplained. There is evidence that in some instances disease might be due to the involvement of another subtype or in some cases the disease may be caused by the vaccine itself. However many disease outbreaks remain unexplained especially in situations where they are observed after apparently successful vaccination with virus of the same subtype. Longitudinal studies were undertaken in Italian turkey farms and demonstrated that subtype B AMPV was frequently detected some period after subtype B vaccination. Sequencing showed that these later viruses were not derived from the previously applied vaccine, and have genome differences in key antigenic regions. An experimental infection of birds was carried out to discover whether the consistent mutations observed in the more recent Italian isolates might allow the viruses to avoid an immune response induced by the common subtype B vaccine, which has virus sequence close to AMPV strains isolated in Italy in the late 80s. The results of the study demonstrated that the more recent field viruses were able to overcome vaccine induced immunity and are consistent with the hypothesis that the field virus has changed in key antigenic regions in order to thrive within a groups of well vaccinated birds.


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INTRODUCTION Infection of turkeys and chickens with Avian Metapneumovirus (AMPV) leads to disease and serious economic losses in unprotected birds. In the late 1980s, live AMPV vaccines became available in Europe and these largely brought the disease under control. However disease has still been encountered and much of this remains unexplained. There is evidence that in some instances disease might be due to the involvement of another subtype (Naylor et al., 1997; Van de Zande et al., 2000) or in some cases the disease may be caused by the vaccine itself (Catelli et al., 2006). However many disease outbreaks remain unexplained especially in situations where they are observed after apparently successful vaccination with virus of the same subtype. A long term survey was carried out in Northern Italy in turkey farms. After the introduction of mass vaccination, AMPV detections decreased but there was an increase in the number of later age detections following vaccination with the same subtype (Catelli, 2006). It had previously been hypothesized that later outbreaks might be due to poor initial vaccination or losses of protection with time. At that time we had not discovered the extent of changes in the field viruses but we have now shown that Italian subtype B viruses fall into 2 distinct clusters (“old” and “new”) based on attachment protein sequences, with the subtype B vaccines forming a part of the “old” group (Cecchinato et al., 2009). Furthermore two longitudinal studies showed that the later subtype B viruses could be detected following subtype B vaccination (Catelli, 2006). In this paper we examine the possibility that these changes in AMPV field strains might be sufficient to avoid the protective response induced by subtype B vaccine. After vaccination with a commercial B subtype vaccine we challenged poults with either “old” or “new” viruses and measured protection based on clinical disease and virus shed. MATERIALS and METHODS Vaccine virus The AMPV B-subtype commercial vaccine was given at dose recommended by the manufacturer by eye drop route. Challenge viruses and dose inoculated Isolates IT/Ty/Vr240/87 and IT/Ty/205-16/04 were isolated in Italian turkey farms respectively in 1987 and in 2004. These viruses were titrated and end points were calculated by the method of Reed and Muench (1938). For inoculation both viruses were used at doses of 3.6 log10 CD50 per poult. Poults Unvaccinated 1-day-old poults were obtained from a commercial turkey hatchery with enforced a high level of biosecurity. Experimental design Fifty 1-day-old commercial turkeys were housed in three positive pressure isolators with 20 poults in two of these (isolator 1 and 2) and the remaining used as negative control in another one (isolator 3). Each birds were identified using leg labels.


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At 1 day old, poults in isolator 1 were vaccinated and poults in the other two isolators were inoculated with sterile water. At 20 days of age half of the birds of isolator 1 were exchanged with half birds of the other isolator (isolator 2). At 21 days of age birds in isolator 1 were inoculated with IT/Ty/205-16/04, in isolator 2 with IT/Ty/Vr240/87 and in isolator 3 with sterile water. Clinical signs were observed daily up to 12 days post challenge (d.p.c.). Oro-pharyngeal swabs were daily collected for virus detection from 3 to 9 and 11 d.p.c. At 13 d.p.c. all birds were humanely killed. Monitoring clinical signs Clinical signs were scored as previously described (Naylor & Jones, 1994) and reported below: 0. no signs; 1. clear nasal exudate; 2. turbid nasal exudate; 3. swollen infra-orbital sinuses and/or frothy eyes. A score 2 or above was considered to show that the bird displayed severe disease. Virus shedding Swabs were individually analysed. RNA extraction and RT-nested PCR were performed as reported by Cavanagh et al. (1999). RESULTS No disease was observed in birds in isolator 3 (negative control). As expected severe disease was observed in unvaccinated birds challenged with both IT/Ty/205-16/04 and IT/Ty/Vr240/87. For the vaccinated birds, those challenged with the “old” B subtype (IT/Ty/Vr240/87) showed no severe clinical signs while sixty percent of the group challenged with “new” B subtype (IT/Ty/205-16/04) showed severe clinical signs for at least one day during the trial. Results of RT-nested PCR, showed that vaccinated birds challenged with the “new” B subtype shed more virus (30 positive samples) compared to the birds challenged with the “old” one (21 positive samples). DISCUSSION The sequences of the G genes of subtype B field Avian Metapneumoviruses detected in Italy showed that later viruses contained a consistent set of mutations and furthermore, these mutations all affected charged amino acids, or amino acids to which glycan might be added during post translational modification (Cecchinato et al. 2009). An experimental infection of birds was carried out to discover whether the consistent mutations observed in the more recent Italian isolates might allow the viruses to avoid an immune response induced by the common subtype B vaccine, which has the earlier virus sequence. The results of the study demonstrated that the “new” field viruses were able to overcome vaccine induced immunity and are consistent with the hypothesis that the field virus has changed in key antigenic regions in order to thrive within a groups of well vaccinated birds. The sequencing of complete genomes of “old” and “new” viruses is underway and will show whether there are extensive sequence differences to be discovered outside to the F and G genes.


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REFERENCES Catelli, E., (2006) Dati epidemiologici sulle infezioni da Pneumovirus Aviare in Italia.

In: Atti della giornata di studio “Malattie respiratorie e problemi di produzione” Giornata di Studio INTERVET (19-23), Bologna, Italy. Italian.

Catelli, E., Cecchinato, M., Savage, C.E., Jones, R.C. & Naylor, C.J. (2006). Demonstration of loss of attenuation and extended field persistence of a live avian metapneumovirus vaccine. Vaccine, 24, 6476-6482.

Cavanagh, D., Madwitt, K., Britton, P. & Naylor, C.J. (1999). Longitudinal field studies of infectious bronchitis virus and avian pneumovirus in broilers using type-specific polymerase chain reactions. Avian Pathology, 28, 593-605.

Cecchinato M., Catelli E., Lupini C., Ricchizzi E., Clubbe J., Naylor C.J.. (2009) Avian metapneumoviruses in Italy: evidence of attachment protein evolution coincident with mass live vaccine introduction. Proceedings of the 6th International symposium on Avian corona- and pneumoviruses And complicating pathogens; 2009 June 14-17; Rauischholzhausen, Germany. In press.

Naylor, C.J., & Jones, R.C., 1994. Demonstration of a virulent subpopulation in a prototype live attenuated turkey rhinotracheitis vaccine. Vaccine, 12, 1225-1230.

Naylor, C., Shaw, K., Britton, P. & Cavanagh, D. (1997). Appearance of type B avian pneumovirus in Great Britain. Avian Pathology, 26, 327-338.

Reed, L.J,. & Muench, H. (1938). A simple method of estimating fifty percent end points. American Journal of Hygiene, 27, 493-497.

Van de Zande, S., Nauwynck, H., Naylor, C. & Pensaert, M. (2000). Duration of cross-protection between subtypes A and B avian pneumovirus in turkeys. Veterinary Record, 147, 132-134.


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LOW AVIAN METAPNEUMOVIRUS (AMPV) VACCINE PERFORMANCE DUE TO TURKEY ASTROVIRUS PERSISTENCE INFECTION: FILED STUDY, BRAZIL

CARDOSO1 TC, FERREIRA3 HL, DA SILVA1 SEL, FERRARI1 HF, TEIXEIRA2 MCB and LUVIZOTTO1 MCR

1 São Paulo State University, DAPSA, Faculdade de Odontologia, Curso de Medicina Veterinária, Rua Clóvis Pestana, 793, Universidade Estadual Paulista, Araçatuba,

SP, CEP 16.050-680, Brasil, 2. São Paulo University, Department of Preventive Veterinary Medicine and Animal

Health, School of Veterinary Medicine, University of São Paulo, CEP 05508-270, São Paulo, SP, Brazil

3 CODA-CERVA-VAR, Avian Virology and Immunology, Groeselenberg 99, B-1180 Uccle, Brussels, Belgium

SUMMARY A field study was conducted to detect avian Metapneumovirus (AMPV) vaccine interference due to turkey Astrovirus (TAstV-2) type 2 persistent infection in the field. Intestinal contents were collected from turkeys in one commercial operation and molecular detection methods were used to screening astrovirus, rotavirus, reovirus and turkey coronavirus by reverse transcriptase and polymerase chain reaction (RT-PCR). All birds were positive for TAstV-2 in intestinal contents confirmed by RT-PCR and sequencing. After collected the respective sera low rates of AMPV antibodies were found ( 10GMT). Comparison of infected birds and AMPV vaccine efficacy, suggests that 75% of the birds presented low AMPV antibodies and decreased to 5% at 10 weeks after vaccination. At necropsy, birds presenting low AMPV antibodies and positive RT-PCR for TAstV-2 were euthanized and bursa of Fabricius (BF) showed mild lymphoid depletion and edema, with no lesions in the intestine. By our results we can speculate that TAstV-2 infection may affect vaccine response in turkeys. The full impact of our findings needs to be further determined by experimental infection with all environmental variables controlled. INTRODUCTION Turkey rhinotracheitis, now commonly termed avian Metapneumovirus (AMPV) infection, is associated with serious welfare and economic problems in susceptible populations of turkeys and probably also of chickens (Catelli et al., 2006; Njenga et al., 2003). The infection principally affects the upper respiratory tract, although egg-


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laying performance may also be affected in breeding turkeys. Secondary infections exacerbate the effects of the primary virus infection (Cook et al., 2001; Patnayak et al., 2002; Patnayak and Goyal, 2004). The virus persists for only a short time both in the host and in the environment and is not known to be transmitted via the egg (Njenga et al., 2003). Highly effective vaccines are available to control APV infections, and hence good biosecurity and careful use of these vaccines should enable infection to be controlled and spread restricted (Cook et al., 2001; Hess et al., 2004; Tarpey and Huggins, 2007). Diagnosis and surveillance are normally performed serologically using enzyme-linked immunosorbent assays (ELISAs) (Hess et al., 2004). Turkey astrovirus (TAstV), a positive sense single-stranded RNA (ssRNA) virus, is an important causative agent of enteritis of poults (Brown et al., 1997; Culver et al., 2006; Pantin-Jackwood et al., 2008). The detection and diagnosis of astroviruses have been mainly dependent on electron microscopy (EM). Reverse transcription-polymerase chain reaction (RT-PCR) has high specificity and sensitivity (Cavanagh et al., 1997). Recently, it has been demonstrated that astroviruses and/or rotaviruses may be persist within a turkey flock through their entire life (Pantin-Jackwood et al., 2008). In addition, experimental infections have demonstrated mild lesions in bursa of Fabricius, which can compromise the immune system (Behling-Kelly et al., 2002). Some studies revealed the interference between AMPV and other avian viruses (Cook et al., 2001; Ganapathy et al., 2005). Nevertheless, the interference of astrovirus infection on AMPV vaccine performance, or other pathogens, was never mentioned before. MATERIAL and METHODS History The flock (n=10.000 birds) belonged to a farm that was managed by high biosecurity instructions was investigated. Birds at 1 week of age, vaccinated against AMPV at first day presented no symptoms of enteric disorder, however showed low growth performance and deep depression (Fig. 1A). All samples, consisted by (n=50) cloacal swabs, (n=500) sera and eventually, the entire bird (n=20), were preserved at -86°C until shipment on wet ice (by express mail) to São Paulo State University, Veterinary School, Virology section. All birds were searched for turkey reovirus, coronavirus, and rotavirus, by conventional RT-PCR. Necropsy and pathology analysis was conducted. Serology was also performed for AMPV antibodies response. Total RNA extraction The total RNA, for all RT-PCR assays, was extracted by Trizol standard protocol guanidinium thiocyanate and acid-phenol with some modifications (Sellers et al., 2004). Two hundred microliters of clinical suspension were mixed with 500ml of Trizol reagent and incubated 10 min at room temperature. After addition of 200l of chloroform, it was mixed vigorously for 10 s and centrifuged at 13, 000 x g for 10 min. The upper aqueous phase was mixed with equal volume of cold isopropanol and incubated on ice 10 min. The total RNA precipitate was centrifuged at 13, 000 x g for 10 min and washed with ethanol. The total RNA was dissolved in 30l of diethyl-pyrocarbonate (DEPC) treated sterile double-distilled water and stored at –20C until use.


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RT-PCR assay for turkey coronavirus (TCoV) and astrovirus (TAstV-1 and 2) The RT-PCR, applied to detect TCoV, TAstV-1 and/or TAstV-2, was performed following the instructions of commercial kit described by Teixeira et al., (2007) and Silva et al., (2008), respectively. A total of 10l of PCR products were electrophoresed at 100V for 1 h in 1.5% agarose gel in 1 x Tris-borate EDTA (TBE) buffer and visualized by ethidium bromide staining and ultraviolet (UV) transluminator. Gel images were captured using Kodak DC290 digital camera and ADOBE 6.0 software. The specificity of the both RT-PCR was tested by addition of other common avian RNA virus (NDV La Sota strain). RT-PCR for turkey rotavirus and reovirus A set of primers was synthesized for turkey rotavirus (GenBank accession number ES204132 and EF204143) reported by Pantin-Jackwood et al. (2007) able to amplify a 630 bp product. Primers used for amplify S3 gene segment of turkey-origin reovirus reported by Spackman et al. (2005) were used. The RT-PCR was done following the same protocol used for amplifying astrovirus and coronavirus. Serology The AMPV antibodies were measured by enzyme-linked immunosorbent assay, ELISA (KPL, Gaithersburg, MD, USA), following all the manufacture's instructions. The 500 sera samples from each week were assayed and the result were expressed by the media of Geometric Mean Titre (GMT), where serum 10 was considered positive expressed as (log2) (+/- standard deviation) (Cook et al., 2001). Sequencing All TAstV-2 related RT-PCRs products were submitted to bi-directional DNA sequencing with DYEnamic Dye Terminator Kit (GE Healthcare) in a MegaBace 1000 automatic DNA sequencer (GE Healthcare). The sequences obtained were aligned with homologous sequences from GenBank with CLUSTAL/W method with Bioedit v.5.0.9. FINAL REMARKS In Brazil, the AMPV infection is currently controlled by vaccination at one day of age, and has been genetically well characterized that subtype A and B are the viruses circulating among flocks (DÀrce et al., 2005). In spite of many reports on AMPV in our country, the only report concerned to turkey Astrovirus, in Latin America, was recently published in 2008 (Silva et al., 2008). Face to the great importance of Brazilian turkey industry, this study was conducted to elucidate the low performance of a turkey flock with no history of enteric disorder. Members of the astrovirus family have been implicated in enteric disease in some mammalian species, including human (Njenga et al., 2003; Velaydhan et al., 2006). In turkeys, clinical depression and mild to moderate diarrhea, most of the times, characterized as frothy yellow feces and mild atrophy of primary lymphoid organs, have been described (Behling-Kelly et al., 2002). In addition, previous studies have demonstrated that astrovirus may be present within a turkey flock life (Pantin-Jackwood et al., 2007). However the direct effect of this persistent infection remains unclear. Our findings revealed low AMPV vaccine performance associated to TAstV-


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2 RNA detection in a particular flock. According to previous reports, the duration of immunity produced by AMPV vaccination present the antibodies peak at 7 weeks post-vaccination and in some cases an extended filed persistence of live AMPV vaccine was observed (Catelli et al., 2006; Hess et al., 2004; Patnayak and Goyal, 2004). After that point, specific antibodies decline. Healthy poults, analyzed in the present study, revealed normal GMT titres, range from 65 to 67. In contrast, poults positive for TAstV-2, and no another virus associated, demonstrated low levels of vaccinal antibodies, 10. Necropsy findings, revealed mild atrophy in bursa of Fabricius with mononuclear infiltration (Fig. 1B, C and D). Other changes were minimal and were noticeably normal. Similar features were described in the first description of TAstV-2 infection in Brazil (Silva et al., 2008). In addition, associations between live AMPV vaccines with Newcastle and Infectious bronchitis virus have been reported previously (Ganapathy et al., 2005; Cook et al., 2001). Finally, further investigation is extremely necessary to determine the real effect of Astrovirus infections on humoral response produced by AMPV vaccination. REFERENCES Behling-Kelly, E., Schultz-Cherry, S., Koci, M., Kelley, L., Larsen, D., Brown, C.

(2002). Localization of Astrovirus in experimentally infected turkeys as determined by in situ hybridization. Veterinary Pathology, 39, 595-598.

Brown, T.P., Garcia, A.P., Kelly, L. (1997). Spiking mortality of turkey poults: I. Experimental reproduction in isolation facilities. Avian Diseases, 41, 604-609.

Catelli, E., Cecchinato, M., Savage, C.E., Jones, R.C., Naylor, C.J. (2006). Demonstration of loss of attenuation and extended field persistence of a live avian metapneumovirus vaccine. Vaccine, 24, 6476-6482.

Cavanagh, D., Mawditt, K., Shaw, K., Britton, P., Naylor, C. (1997). Towards the routine application of nucleic acid technology for avian disease diagnosis. Acta Veterinaria Hungarica, 45, 281-298.

Cook, J.K.A., Huggins, M.B., Orbell, S.J., Mawditt, K., Cavanagh, D. (2001). Infectious bronchitis virus vaccine interferes with the replication of avian penumovirus vaccine in domestic fowl. Avian Pathology, 30, 233-242.

Culver, F., Dziva, F., Cavanagh, D., Stevens, M.P. (2006). Poult enteritis and mortality syndrome in turkeys in Great Britain. Veterinary Record, 159, 209-210.

DÁrce, R.C.F., Coswig, L.T., Almeida, R.S., Trevisol, I.M., Monteiro, M.C.B., Rossini, L. I., Di Fabio, J., Hafez, H.M., Arns, C.W. (2005). Subtyping of new Brazilian avian metapneumovirus isolates from chickens and turkeys by reverse transcriptase-nested polymerase chain reaction. Avian Pathology, 34, 133-136.

Ganapathy, K.,Cargill, P., Montiel, E., Jones, R.C. (2005). Interaction between live avian pneumovirus and Newcastle disease virus vaccines in specific pathogen free chickens. Avian Pathology, 34, 297-302.

Hess, M., Huggins, M.B., Heincz, U. (2004). Hacthability, serology and virus excretion following in ovo vaccination of chickens with an avian metapneumovirus vaccine. Avian Pathology, 33, 576-580.

Njenga, M.K., Lwamba, M.H., Seal, B.S. (2003). Metapneumovirus in birds and humans. Virus Research, 163-169.


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Pantin- Jackwood, M.J., Spackman, E., Day, M.J., Rives, D. (2008). Periodic monitoring of commercial turkeys for enteric viruses indicates continuous presence of astrovirus and rotavirus on the farms. Avian Diseases, 51, 674-680.

Patnayak, D.P., Goyal, S.M. (2004). Duration of immunity produced by a live attenuated vaccine against avian pneumovirus type C. Avian Pathology, 33, 465-469.

Patnayak, D.P., Sheikh, A.M., Gulati, B.R., Goyal, S.M. (2002). Experimental and field evaluation of a live vaccine against avian penumovirus. Avian Pathology, 31, 377-382.

Sellers, H., Koci, M.D., Linnemann, E., Kelley, L.A., Shultz-Cherry, S. (2004). Development of a multiplex reverse transcription-polymerase chain reaction diagnostic test specific for turkey Astrovirus and Coronavirus. Avian Diseases, 48, 531-539.

Tarpey, I., Huggins, M.B. (2007). Onset of immunity following in ovo delivery of avian metapneumovirus vaccines. Veterinary Microbiology, 124, 134-139.

Velayudhan, B.T., Nagaraja, K.V., Thachil, A.J., Shaw, D.P., Gray, G.C., Halvorson, D.A. (2006). Human Metapneumovirus in Turkeys Poults. Emergent Infectious Disease, 12, 1853-1859.


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Figure 1- A) Aspect of depressed poults; B) Macroscopical findings, principally edema, of bursa of Fabricius; C-D) Microscopic lesions characterized by lymphocytic depletion and mononuclear infiltration. ACKNOWLEDGEMENTS This research was supported by FAPESP (grants 05/52994-3; 07/56041-6; 07/53090-6; 08/50380-6; 08/09945-0; 09/50800-8) and CNPq (grant 472226 / 2007-0).

C


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THE ROLE OF HUMORAL AND CELL-MEDIATED IMMUNITY IN THE CONTROL OF AVIAN METAPNEUMOVIRUS INFECTION IN TURKEYS

RUBBENSTROTH D and RAUTENSCHLEIN S

Clinic for Poultry, University of Veterinary Medicine Hannover, Bünteweg 17, 30559 Hannover, Germany; [email protected];

SUMMARY Field observations and experimental studies indicated that circulating antibodies may not be sufficient to prevent infection of turkeys with avian Metapneumovirus (aMPV). It is speculated that the cell-mediated immunity may play an important role in the pathogenesis and control of aMPV. We investigated the protective role of passively transferred aMPV antibodies against challenge with a virulent aMPV subtype A strain in aMPV-maternal antibody free turkeys. Although, aMPV antibodies were detected on the mucosal surfaces of the respiratory tract and in the circulation after antibody transfer, turkeys were not protected against aMPV challenge. Clinical signs and the virus clearance rate were comparable between aMPV antibody-positive birds and antibody-negative controls. Furthermore, it was shown that the depletion of functional T cells affected the susceptibility of aMPV-antibody-free turkeys for aMPV-infection and delayed clinical recovery and virus clearance from the respiratory tract. These observations provided circumstantial evidence that functional T cells play an important role in the control of aMPV-infections in turkeys, and they should be specifically targeted to improve current vaccination strategies. INTRODUCTION Avian Metapneumovirus (aMPV) is known to cause an acute respiratory disease referred to as Turkey Rhinotracheitis (TRT) in turkeys or Avian Rhinotracheitis (ART) in other bird species. The disease is characterized by respiratory symptoms such as sneezing, ocular discharge and swelling of the infraorbital sinus (Gough, 2003). Virus replication in the respiratory epithelium leads to influx of lymphoid cells and mucosal damage such as epithelial desquamation and loss of ciliary activity (Majo et al., 1995; Liman & Rautenschlein, 2007). A systemic immunosuppression has been proposed as an additional consequence of aMPV-infection (Timms et al., 1986; Chary et al., 2002a; Chary et al., 2002b). Due to damage of the respiratory epithelium and possibly systemic immunosuppressive effects aMPV-infection may be followed by



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secondary infections leading to great economic losses to the poultry industry (Van de Zande et al., 2001; Marien et al., 2005; Rubbenstroth et al., 2008). Current vaccination regimes against aMPV infection are mainly based on attenuated live or inactivated vaccines and have proven to be useful tools for the control of the disease (Jones, 1996). Nevertheless, they often do not provide sufficient protection. In order to be able to improve current vaccination strategies it is necessary to understand more about the means of aMPV-protection. Not much is known about the role of humoral and cell-mediated immune parameters in the control of aMPV-infection. Previous experimental and field observations indicated that humoral immunity may not be sufficient for aMPV protection. Experimental studies have demonstrated that maternally derived antibodies did not prevent virus replication and clinical disease (Naylor et al., 1997). On the other hand studies in turkeys had shown that aMPV-infection induces a transient stimulation of local humoral immunity and the accumulation of T cells in the Harderian gland (Chary et al., 2002a; Cha et al., 2007; Liman & Rautenschlein, 2007). The goal of this study was to understand more about the role of humoral and cell-mediated immunity in the control of aMPV-infection. Our objectives were to investigate the development of clinical disease, virus clearance and the induction aMPV-antibodies in aMPV-maternal antibody negative commercial turkey poults a) after passive transfer of aMPV-neutralizing antibodies; and b) after chemical suppression of the T cell responsiveness with Cyclosporin A (CsA). MATERIAL and METHODS Experimental design Experiment 1: Eighty-three 14-day-old commercial female Big-6 turkey poults, which were confirmed to be free of maternal antibodies against aMPV, were randomly assigned to four groups of 18 to 24 turkeys. Birds of two groups (AC, 24 birds and AV, 18 birds) were intravenously inoculated with aMPV-A-specific antibodies by two consecutive injections of 1.4 ml of concentrated aMPV-specific antibodies (aMPV-Ab+) at days 14 and 15 of life. The two remaining groups (CC, 23 birds and CV, 18 birds) received a similar treatment with the aMPV-Ab negative preparation (aMPV-Ab-). Turkeys of groups CV and AV were oculonasally inoculated with 103 CD50 of the virulent aMPV-strain BUT 8544 (Liman & Rautenschlein, 2007) per bird fifteen minutes after the second antibody injection. Groups CC and AC received virus-free TOC-supernatant. At the same time, five and six turkeys of groups CC and AC, respectively, were sacrificed for necropsy. Six turkeys of each group were sacrificed at days 5, 9 and 14 post inoculation (pi). At necropsy choanal swabs were taken for aMPV-genome detection by RT-PCR (Cavanagh et al., 1999), and serum and lacrimal fluid was collected for antibody detection in the virus-neutralizing test and ELISA. Choanal swabs and serum samples were collected immediately before the first antibody injection, fifteen minutes after the second antibody injection and at days 1, 3, 5, 7, 9, 11 and 14 pi (n = 6-10). Lacrimal fluid was collected fifteen minutes after the second antibody injection and at days 5, 9 and 14 pi (n = 6-10). Experiment 2 and 3: Twenty-eight (Experiment 2) and sixty (Experiment 3) one-day-old commercial female Big-6 turkey poults, which were confirmed to be free of maternal antibodies against aMPV, were randomly assigned to two groups of equal


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numbers of turkeys. Beginning at day 6 (Exp. 2) or 4 (Exp. 3) post hatch, one group (T) was repeatedly treated every three to four days by intramuscular route with 100 mg/kg body weight of Cyclosporin A (Sandimmun, Novartis, Switzerland). At 28 days post hatch, five to eight (Exp. 2) or twelve to sixteen (Exp. 3) turkeys of each group were oculonasally inoculated with 103 CD50 of the virulent aMPV-strain BUT 8544 (Liman & Rautenschlein, 2007). Clinical signs were monitored daily. In Experiment 2, at the day of virus-inoculation, and at 4, 7, 10-11, and 14 days post virus inoculation (pi) choanal swabs were collected and tested for aMPV by RT-PCR. At the day of virus-inoculation, 7, 10-11 and 14 days pi serum samples were collected to detect aMPV antibodies in the ELISA and virus neutralization test. At 14 days pi, the experiment was terminated and all birds humanely sacrificed. Peripheral blood leukocytes were collected randomly from birds of the CsA-treated groups (n= 6-8) at one day before virus-inoculation, 7 and 13-14 days pi, and the relative (Exp. 2) and absolute (Exp. 3) numbers of CD4+ and CD8a+ T cells were determined by flow cytometric analysis. Antibody preparations Turkey serum free of detectable aMPV-specific antibodies was collected from turkey poults reared under isolated conditions. For the production of anti-aMPV hyperimmune serum three male turkeys were first inoculated intranasally with virulent BUT 8544 at 8 weeks post hatch and intramuscularly with an inactivated aMPV-subtype A vaccine at the age of 8, 11, 13, 15 and 17 weeks. One week after the last booster, serum was harvested and stored at -70°C until further use. The turkey sera were heat inactivated at 56°C for 30 minutes and total immunoglobulin (Ig) concentration was increased by ammonium sulphate precipitation (Lebacq-Verheyden et al., 1974). The resulting concentrated antibody preparations obtained from the hyperimmune sera (aMPV-Ab+) had VN log-2 titres of 10.3. Parallel preparations from aMPV-antibody free turkey sera (aMPV-Ab-) also treated with ammonium sulphate were confirmed to be free of aMPV-specific antibodies by VNT and ELISA. Serology aMPV-specific IgG antibodies were detected by Avian Rhinotracheitis Antibody Test Kit (BioChek, Gouda / Netherlands) following the manufacturers’ instructions. The different samples were diluted in the provided dilution buffer as follows: serum 500-fold, and lacrimal fluid 18-fold. ELISA-results are presented as sample to positive control (S/P) ratios. Detection of virus neutralizing (VN) antibodies was performed on primary chicken embryo fibroblast cells following previously published methods (Baxter-Jones et al., 1989). Results are presented as VN log-2 titres. Detection of aMPV by RT-PCR RNA was isolated from choanal swabs using Trifast GOLD (Peqlab, Erlangen / Germany) according to the manufacturers’ instructions. The RT reaction was performed using the ImProm-II© RT system (Promega, Madison / USA) according to the manufacturer’s directions with random primers (Invitrogen, Karlsruhe / Germany). The nested PCR was performed following previously published procedures (Cavanagh et al., 1999).


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Clinical score Clinical signs were recorded as individual scores per animal. A score of 0 (no signs) to 3 (severe signs) was assigned to each of the following respiratory symptoms: nasal exudate, ocular discharge and infraorbital swelling. The sum of these scores resulted in a total score of 0 to 9 for each individual turkey. Flow cytometric analysis of peripheral blood leukocytes For the detection of relative T-Lymphocyte numbers (Exp. 2), peripheral blood leukocytes (PBL) were isolated by sucrose gradient centrifugation with Biocoll and stained with fluorescence-labelled monoclonal antibodies directed against the T-lymphocyte antigens CD4 and CD8a (Southern Biotech., Birmingham, Alabama, USA). Labelled cells were analysed by flowcytometry and results are presented as percent CD4 or CD8a positive cells of the gated lymphocyte population. For the detection of total T-Lymphocyte numbers (Exp. 3), EDTA-supplemented and diluted blood was fixed with Transfix (Cytomark, Buckingham, UK) and stained with CD4 and CD8a specific antibodies. Before flowcytometric analysis, a defined number of fluorescent beads (FlowCount, Beckmann Coulter, Galway, Ireland) was added to each sample. Results were calculated as number of CD4 or CD8a positive cells per µl blood. Statistical analysis Statistical analysis of ELISA and VNT results was performed with Statistix 7.0 software, using One-way analysis of variants (ANOVA) and comparison of means by Tukey Test. P-values of P < 0.05 were considered to be significant. RESULTS and DISCUSSION Effect of passive aMPV-antibody transfer on the development of aMPV-infection Serum antibody levels after passive transfer Following passive transfer of aMPV-specific antibodies, VN (data not shown) and ELISA antibodies were detected in turkey sera (Fig. 1). aMPV-specific ELISA IgG antibodies were also detected on mucosal surfaces of conjunctivae (Fig. 1). This is in congruence with previous data, showing that IgG was transferred through the epithelial barrier in turkeys and chickens (Toro et al., 1993; Suresh & Arp, 1995). aMPV-challenged birds without passively transferred aMPV-specific antibodies (CV) developed significantly increased ELISA antibody levels in sera and lacrimal fluid, starting at day 9 pi (Fig. 1; P<0.05). In turkeys with passively transferred aMPV-specific antibodies (AV) ELISA-IgG levels did not significantly increase after challenge infection. Antibody levels achieved by passive antibody transfer at the time of aMPV challenge were comparable to or even higher than peak antibody responses induced by virulent aMPV infection. If aMPV-antibodies would play a role in protection these antibody levels achieved by passive transfer in this study were sufficiently high enough to test this hypothesis. Clinical signs Clinical signs were recorded in Experiment 1 daily throughout the experiment using a scoring system from 0 to 9. Virus-inoculated turkeys developed respiratory


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symptoms, such as nasal and ocular discharge and swelling of the infraorbital sinus. Symptoms were first observed at day 3 pi and were most severe at days 6 and 7 pi. Clinical signs had completely vanished by day 12 pi. Both infected groups (CV and AV) showed comparable development of clinical symptoms (Table 1). These results reveal that even in the presence of high levels of local and circulating antibodies, turkeys were not protected against clinical disease after oculonasal challenge with a homologues aMPV-A strain. Detection of aMPV by RT-PCR aMPV was detected from choanal swabs by a subtype specific RT-PCR (Cavanagh et al., 1999). All samples collected from the aMPV-inoculated groups (CV and AV) in Experiment 1 were positive for aMPV subtype A between day 3 and 7 pi. Detection rates declined starting at day 9 pi. in both challenged groups. At day 14 pi only one sample from group CV was aMPV-positive by RT-PCR (Table 1). These results demonstrate that aMPV-antibody positive turkeys did not show protection against aMPV infection nor improved virus clearance compared to antibody negative animals. Effect of CsA-treatment on the development of aMPV-infection Depletion of T-lymphocyte subpopulations In Exp. 3 at all investigated time points total numbers of CD4 and CD8a positive T-lymphocytes in peripheral blood were significantly reduced (P < 0.05) after repeated CsA-treatment compared to untreated controls (Fig. 2). These results confirm the T-lymphocyte suppressing effect of CsA in turkeys, as already described by Suresh & Sharma (1995). aMPV-infection did not affect T-lymphocytes numbers in peripheral blood compared to virus-free groups (P > 0.05). In Exp. 2 similar CsA-mediated T cell suppressive effects were detected by flow cytometric analysis, when relative numbers of T-lymphocytes were determined in isolated PBL or spleen leukocytes as compared to Exp. 3 where total numbers were detected (data not shown). Clinical signs aMPV-inoculated turkeys showed TRT-like symptoms, beginning at day 3 pi in Exp. 2 and 3. Untreated and CsA-treated groups had comparable mean clinical scores during the first six to seven days pi. Thereafter, recovery of CsA-treated aMPV-infected turkeys was delayed by about two days in both experiments as compared to T cell intact infected birds (Table 2). Turkeys of the uninfected group did not express signs of TRT. These observations indicate that CsA-treatment affected the recovery of aMPV-infected birds suggesting a of role functional T cells in aMPV pathogenesis. It has been shown in other pneumovirus infection models that the protective role of T cells may depend on the infectious dose (Frey et al., 2008). For pneumovirus infection in mice (PVM) it was shown that T cells were important for the control of virus infection after a low dose but that disease and mortality were independent of T cells after lethal high-dose PVM infection. We speculate that both T-cell-dependent and -independent pathways may contribute to aMPV in turkeys and that the influence of the viral dose has to be tested in the future.


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Detection of aMPV by RT-PCR In Exp. 2 ninety to hundred percent of choanal swabs collected from both aMPV-inoculated groups at days 4 and 7 pi were aMPV-positive. At day 10 pi CsA-treated, aMPV-inoculated birds had a higher rate of aMPV-positive swabs compared to the untreated group. At day 14 pi 38 % of the choanal swabs from the CsA-treated group were still aMPV-positive, whereas no aMPV was detected in the untreated group. No aMPV was detected in either uninfected group. These observations may explain the delayed recovery from clinical respiratory signs observed in both experiments. It can be speculated that T cells control aMPV clearance. Specifically CD8+ T cells are known to play a significant role in the control of pneumovirus infections of other animals (Fu et al., 2009). It is thought that effective respiratory syncytial virus clearance requires the induction of balanced Th1-type immunity, involving the activation of IFN-gamma-secreting cytotoxic T cells (Bueno et al., 2008). Development of serum antibodies Following aMPV-inoculation CsA-treated and untreated turkeys developed significantly increased levels of aMPV-specific antibodies in sera by day 7 pi as compared to virus-free birds. At day 10 and 14 pi aMPV-infected CsA-treated turkeys had significantly higher ELISA and VN serum antibody levels compared to the infected T cell-intact control group (P < 0.05; Fig. 3). It can be speculated that the longer presence of viral antigen in T cell deficient birds leads to a stronger stimulation of the humoral immune response resulting in higher aMPV-antibody levels in CsA-treated turkeys as compared to the T cell-intact controls. CONCLUSIONS The presented study demonstrates that intravenously administered antibodies did not protect turkeys from aMPV-infection and respiratory disease, although they were readily transferred to the mucosal surfaces. Our results support previous observations that serum antibodies acquired after vaccination were not indicatory for actual protection (Cook et al., 1989; Williams et al., 1991a; Williams et al., 1991b; Sharma et al., 2004; Kapczynski et al., 2008). Antibody induction is therefore considered to be an insufficient parameter for estimating the degree of protection provided by an aMPV vaccine. The results of Exp. 2 and 3 demonstrate that CsA-treatment of turkeys led to reduced numbers of circulating CD4 and CD8a positive T-lymphocytes. Recovery from clinical signs and clearance of aMPV was delayed in chemically T-lymphocyte-suppressed turkeys. This indicates that T-cells play an important role in the control of aMPV-infection in turkeys. They should therefore be considered as important targets for the development of improved aMPV vaccines. ACKNOWLEDGEMENTS The authors like to thank Dorothee Schmalstieg and Christine Haase for their valuable technical assistance and Sonja Bernhardt and Martina Koschorrek for the help with the animal experiments. The project is funded by the German Research Society (DFG, RA 767/3-1).


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methods for detecting antibodies to turkey rhinotracheitis virus. Avian Pathology, 18, 91-98.

Bueno, S. M., Gonzalez, P. A., Cautivo, K. M., Mora, J. E., Leiva, E. D., Tobar, H. E., Fennelly, G. J., Eugenin, E. A., Jacobs, W. R., Riedel, C. A. & Kalergis, A. M. (2008). Protective T cell immunity against respiratory syncytial virus is efficiently induced by recombinant BCG. Proceedings of the National Academy of Sciences of the United States of America, 105, 20822-20827.

Cavanagh, D., Mawditt, K., Britton, P. & Naylor, C. J. (1999). Longitudinal field studies of infectious bronchitis virus and avian pneumovirus in broilers using type-specific polymerase chain reactions. Avian Pathology, 28, 593-605.

Cha, R. M., Khatri, M. & Sharma, J. M. (2007). B-Cell Infiltration in the Respiratory Mucosa of Turkeys Exposed to Subtype C Avian Metapneumovirus. Avian Diseases, 51, 764-770.

Chary, P., Rautenschlein, S., Njenga, M. K. & Sharma, J. M. (2002a). Pathogenic and immunosuppressive effects of avian pneumovirus in turkeys. Avian Diseases, 46, 153-161.

Chary, P., Rautenschlein, S. & Sharma, J. M. (2002b). Reduced efficacy of hemorrhagic enteritis virus vaccine in turkeys exposed to avian pneumovirus. Avian Diseases, 46, 353-359.

Cook, J. K. A., Holmes, H. C., Finney, P. M., Dolby, C. A., Ellis, M. M. & Huggins, M. B. (1989). A live attenuated turkey rhinotracheitis virus-vaccine. 2. The use of the attenuated strain as an experimental vaccine. Avian Pathology, 18, 523-534.

Frey, S., Krempl, C. D., Schmitt-Graff, A. & Ehl, S. (2008). Role of T Cells in Virus Control and Disease after Infection with Pneumonia Virus of Mice. Journal of Virology, 82, 11619-11627.

Fu, Y. H., He, J. S., Zheng, X. X., Wu, Q., Zhang, M., Wang, X. B., Wang, Y., Xie, C., Tang, Q., Wei, W., Wang, M., Song, J. D., Qu, J. G., Zhang, Y., Wang, X. & Hong, T. (2009). Intranasal immunization with a replication-deficient adenoviral vector expressing the fusion glycoprotein of respiratory syncytial virus elicits protective immunity in BALB/c mice. Biochemical and Biophysical Research Communications, 381, 528-532.

Gough, R. E. (2003) Avian pneumoviruses. In Y. M. Saif. Diseases of Poultry 11th edn (pp. 92-99): Iowa State Press.

Jones, R. C. (1996). Avian pneumovirus infection: Questions still unanswered. Avian Pathology, 25, 639-648.

Kapczynski, D. R., Perkins, L. L. & Sellers, H. S. (2008). Mucosal vaccination with formalin-inactivated avian metapneumovirus subtype C does not protect turkeys following intranasal challenge. Avian Diseases, 52, 28-33.

Lebacq-Verheyden, A. M., Vaerman, J. P. & Heremans, J. F. (1974). Quantification and distribution of chicken immunoglobulins IgA, IgM and IgG in serum and secretions. Immunology, 27, 683-692.


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Liman, M. & Rautenschlein, S. (2007). Induction of local and systemic immune reactions following infection of turkeys with avian Metapneumovirus (aMPV) subtypes A and B. Veterinary Immunology and Immunopathology, 115, 273-285.

Majo, N., Allan, G. M., O'Loan, C. J., Pages, A. & Ramis, A. J. (1995). A sequential histopathologic and immunocytochemical study of chickens, turkey poults, and broiler breeders experimentally infected with turkey rhinotracheitis virus. Avian Diseases, 39, 887-896.

Marien, M., Decostere, A., Martel, A., Chiers, K., Froyman, R. & Nauwynck, H. (2005). Synergy between avian pneumovirus and Ornithobacterium rhinotracheale in turkeys. Avian Pathology, 34, 204-211.

Naylor, C. J., Worthington, K. J. & Jones, R. C. (1997). Failure of maternal antibodies to protect young turkey poults against challenge with turkey rhinotracheitis virus. Avian Diseases, 41, 968-971.

Rubbenstroth, D., Ryll, M., Behr, K.-P. & Rautenschlein, S. (2008). Avian Metapneumovirus supports experimental Riemerella anatipestifer infection in turkeys. In H. M. Hafez (Ed.). Proceedings of the 7th International Symposium on Turkey Diseases (pp. 197-201). Berlin, Germany

Sharma, J. M., Chary, P., Khatri, M., Cha, R. & Palmquist, J. M. (2004). Pathogenesis and control of avian pneumovirus. In U. Heffels-Redmann & E. F. Kaleta (Ed.). Proceedings of the IV. Symposium on Avian Corona- & Pneumovirus Infections (pp. 318-321). Rauischholzhausen, Germany

Suresh, M. & Sharma, J. M. (1995). Hemorrhagic Enteritis Virus-Induced Changes in the Lymphocyte Subpopulations in Turkeys and the Effect of Experimental Immunodeficiency on Viral Pathogenesis. Veterinary Immunology and Immunopathology, 45, 139-150.

Suresh, P. & Arp, L. H. (1995). A time-course study of the transfer of immunoglobulin-G from blood to tracheal and lacrimal secretions in young turkeys. Avian Diseases, 39, 349-354.

Timms, L. M., Jahans, K. L. & Marshall, R. N. (1986). Evidence of immunosuppression in turkey poults affected by rhinotracheitis. Veterinary Record, 119, 91-92.

Toro, H., Lavaud, P., Vallejos, P. & Ferreira, A. (1993). Transfer of IgG from serum to lacrimal fluid in chickens. Avian Diseases, 37, 60-66.

Van de Zande, S., Nauwynck, H. & Pensaert, M. (2001). The clinical, pathological and microbiological outcome of an Escherichia coli O2 : K1 infection in avian pneumovirus infected turkeys. Veterinary Microbiology, 81, 353-365.

Williams, R. A., Savage, C. E. & Jones, R. C. (1991a). Development of a live attenuated vaccine against turkey rhinotracheitis. Avian Pathology, 20, 45-55.

Williams, R. A., Savage, C. E., Worthington, K. J. & Jones, R. C. (1991b). Further studies on the development of a live attenuated vaccine against turkey rhinotracheitis. Avian Pathology, 20, 585-596.


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ELISA antibodies in serum

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(A) ELISA antibodies in lacrimal fluid

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Figure 1: Detection of aMPV-ELISA antibodies after passive transfer of in serum (A) and lacrimal fluid (B). Turkeys were treated with an ammonium sulphate-treated aMPV-hyperimmune serum or aMPV-antibody free serum and subsequently challenged with a virulent aMPV-subtype A strain. Values marked with different superscript letters at the same experimental day are significantly different from each other (One Way ANOVA and Tukey test, P < 0.05); n = 6-10. Table 1: Effect of passive transfer of aMPV-specific antibodies on the development of clinical disease and virus clearance after inoculation of turkeys with a virulent aMPV subtype A

Groups

aMPV-specific antibody transfer

Development of clinical disease presented as mean clinical score/group and aMPV detection by RT-PCR (% positive

birds/group) at days pi

1 3 5 7 9 11 14 Virus-free - 0a

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0.7 (70)

0 (50)

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aMPV A + 0 (100)

0.7 (100)

2.5 (100)

3.1 (100)

0.7 (90)

0.2 (33)

0 (0)

a Mean clinical score per group and day; n = 6-18; clinical signs were nasal and ocular discharge and swelling of the infraorbital sinus.

b percent of aMPV-positive swabs per group and day; n = 6-8.


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CD4 positive T-lymphocytes

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Figure 2: Effect of Cyclosporin A on total numbers of T-lymphocytes in peripheral blood (Exp. 3). (A) CD4 positive T-cells. (B) CD8a positive T-cells. Values marked with different superscript letters at the same experimental day are significantly different from each other (One Way ANOVA and Tukey test, P < 0.05); n = 6-8; ND: not done. Table 2: Effect of Cyclosporin-A induced suppression of T cells on the development of clinical disease and virus clearance after inoculation of turkeys with a virulent aMPV subtype A (Experiment 2 as a representative experiment)

Groups CsA-treatment

Development of clinical disease presented as mean clinical score/group and aMPV detection by RT-PCR (% positive

birds/group) at days pi 3 4 5 6 7 8 9 10 14

Virus-free - 0a

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0.1 (57)

0 (38)

a Mean clinical score per group and day; n = 5-8; clinical signs were nasal and ocular discharge and swelling of the infraorbital sinus.

b ND = not done c percent of aMPV-positive swabs per group and day; n = 5-8.


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Serum ELISA antibodies

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Figure 3: Effect of CsA-treatment on the aMPV-antibody response (Exp. 2); (A) ELISA antibodies; (B) virus neutralising (VN) antibodies. Values marked with different superscript letters at the same experimental day are significantly different from each other (One Way ANOVA and Tukey test, P < 0.05); n = 5-8;


355

ACTIVITY AND EFFICACY OF AN EXPERIMENTAL INACTIVATED OIL-ADJUVANTED SUBGROUP C AVIAN METAPNEUMOVIRUS (AMPV-C) ANTIGEN

PREPARATION IN WHITE PEKING AND SPF MUSCOVY DUCKLINGS

TOQUIN1 D, ALLEE1 C, LE BRAS1 MO, AMELOT2 M and ETERRADOSSI1 N

French Agency for Food Safety (AFSSA), BP 53, 22440 Ploufragan, France, 1 Avian and Rabbit Virology Immunology and Parasitology Unit (VIPAC),

2 Experimental Service for Avian Pathology (SEEPA) SUMMARY As a contribution to the control of AMPV-C infections in ducks, an experimental inactivated oil-adjuvanted immunizing antigen was prepared from a French AMPV-C isolate propagated in Vero cells. A similar mock immunizing antigen was prepared as a control from non-infected cells. Two groups of thirty ducklings, either 3-day-old White Peking ducks without maternal antibodies to AMPV-C, or day-old specific pathogen free Muscovy ducks (Afssa-Ploufragan), were housed in an A2 containment cell. At day 14, half the birds in each group were transferred to another containment cell and received 2 doses per bird of either the AMPV-C immunizing antigen (10 birds) or the control mock antigen (5 birds). The White Peking and SPF Muscovy ducks remaining in the first containment cell were kept as non-immunized controls. Three weeks later, all ducks in both containment cells were submitted to serological testing and the immunized ducklings received a booster injection with one dose per bird of the same antigen as received previously. Two weeks later, all the birds in both containment cells were blood sampled for serological testing, then they were challenged by the intranasal route with 103,5TCID50 per bird of an heterologous French AMPV-C isolate. At days 3, 5, 7 and 11 post challenge, tracheal swabs were sampled in all birds for qRT-PCR assessment of virus excretion. Activity of the immunizing antigens was assessed serologically using an in-house ELISA for the detection of anti AMPV-C antibodies. The injection of one or two doses of the AMPV-C immunogen elicited a strong anti-AMPV-C antibody response in White Peking ducks (mean ELISA ratio > 0.800) and a moderate one in Muscovy ducks (mean ELISA ratio > 0.300). Background responses elicited by the mock immunizing antigen were moderate in both duck species (< 0.450 and 0,100 in White Peking and Muscovy ducks, respectively). No antibodies against APMV-C were detected in non-immunized control birds prior to challenge. Due to the challenge being performed at 7 weeks of age and the resulting limited clinical signs, the


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challenge was validated by ELISA testing in the unvaccinated challenged groups, which in both duck species exhibited a sharp rise in anti-AMPV-C antibodies two weeks post challenge. Virus replication as assessed by qRT-PCR on tracheal swabs was detected as soon as 3 days post challenge in all Muscovy ducks, irrespective of their immunization schedule, whereas in White Peking ducks, it was only detected in the non-immunized and challenged control group at 11 days post challenge, and not in the groups that had previously been immunized with the AMPV-C or mock antigens. In summary, the AMPV-C antigen preparation was able to elicit specific anti-AMPV C antibodies in both Muscovy and White Peking ducks, however these antibodies did not control the tracheal replication of an heterologous challenge AMPV-C virus in Muscovy ducks. In White Peking ducks, the late detection of the challenge virus in non-immunized birds only impedes any conclusions regarding the efficacy of the immunization scheme in this species. INTRODUCTION Subgroup C avian metapneumoviruses (AMPV-C) were first detected in turkeys with respiratory signs in the USA (Senne et al., 1997; Cook et al., 1999). They were then isolated in France in farmed breeder Muscovy or White Peking ducks experiencing a drop in egg production after respiratory signs (Toquin et al., 1999) and have also been serologically detected in younger birds with an history of “infectious bronchitis-like” respiratory disease which can be experimentally reproduced in SPF Muscovy ducks (Jestin et al., 2000). AMPV-C have also been isolated from wild ducks and geese in the USA (Bennett et al., 2002), thus showing that AMPV-C viruses are most likely commonly found in duck species worldwide, although the French and US isolates of AMPV-C belong to different genetic lineages (Toquin et al., 2006b). The repeated isolation of AMPV-C from ducks is in sharp contrast with results obtained with other AMPV subgroups (AMPV-A, -B or –D), originally isolated in turkeys, which do not seem to infect Muscovy duck as experimentally demonstrated by the lack of clinical signs, the lack of virus re-isolation from the trachea, and the lack of serovonversion after the inoculation of these viruses by a natural route (Toquin et al., 2006a). As already implemented in turkey and chicken breeders (Cook et al., 1996, 2000), vaccination could be considered in ducks as an approach to control the respiratory and reproductive signs associated with AMPV infections. However, no live attenuated vaccine derived from a European AMPV-C isolate is currently available. As no AMPV-C have yet been detected in turkeys in Europe, the importation of a turkey live attenuated AMPV-C vaccine derived from an American virus isolate (which is currently commercially available in the USA) was not considered as an option, for fear that its field use might result in the introduction in Europe of an AMPV-C strain adapted to turkeys. The present study was thus implemented in order to assess whether the immunization of ducks with an inactivated adjuvanted antigen derived from a French AMPV-C isolate would help prevent infection by these viruses.


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MATERIALS and METHODS Preparation of immunizing antigens The virus used for antigen preparation was isolate 04268 obtained in the authors’ laboratory. The virus was cloned three times by the limiting dilution in Vero cells, and its assignation to subgroup C was confirmed by RT-PCR amplification and nucleotide sequencing of its SH and G genes. The virus was grown in Vero cells and MEMH so as to obtain a supernatant with a minimal infectious titre of 106.5TCID50/ml. The supernatant was then inactivated by the addition of 1 ‰ of 37 % formaldehyde and mild shaking for one night at 37°C. The supernatant of mock-infected cells was treated similarly to prepare a mock antigen. The antigens were finally adjuvanted by preparation of a water-in-oil emulsion. Each dose of the inactivated adjuvanted AMPV-C antigen contained a virus dose equivalent to at least 105.3TCID50 prior to inactivation. Aliquots were harvested at each production step to allow for further controls. Controls for inactivation and purity Inactivation was tested before adjuvanting by three serial passages in Vero cells. Purity before adjuvanting was assessed as following: Prior inactivation, aliquots of the live AMPV-C virus suspension were neutralized with a monospecific anti-AMPV-C antiserum produced in SPF turkeys, then three serial passages of the neutralized virus suspension were performed in Vero cells, in chicken embryo fibroblast cell cultures and in specific pathogen free 9-day-old embryonated chicken eggs (source Afssa Ploufragan, allantoic fluid route). After inactivation, the absence of mycoplasmas was assessed with a broad spectrum PCR and by culture in the FM4 medium. The absence of fungi was assessed by culture in Sabouraud’s medium. Purity after adjuvanting was assessed by testing the inactivated adjuvanted AMPV-C and control antigens for bacteria according to the European Pharmacopoeia, and by immunizing 14-to-28 day-old SPF Muscovy or White Peking ducklings with two doses of the AMPV-C or mock antigens, followed by a one dose booster injection 3 weeks later. The immunized ducklings were followed clinically, and the sera harvested after the booster injection were tested for the absence of antibodies against 18 bacterial or viral pathogens. Experimental design Two groups of thirty ducklings, either of 3-day-old White Peking ducks without maternal antibodies to AMPV-C, or of day-old specific pathogen free Muscovy ducks (Afssa-Ploufragan), were housed in an A2 containment cell. At day 14, half the birds in each group were transferred to another containment cell and received by the intramuscular route 2 doses per bird of either the AMPV-C immunizing antigen (10 birds) or the control mock antigen (5 birds). The White Peking and SPF Muscovy ducks remaining in the first containment cell were kept as non-immunized controls. Three weeks later, all ducks in both containment cells were submitted to serological testing and the immunized ducklings received a booster injection with one dose per bird using the same antigen and route of immunization as used previously. Two weeks later, all the birds in both containment cells were blood sampled for serological testing, they were then challenged by the intranasal route with 103,5TCID50 per bird of an heterologous French AMPV-C isolate (reference 99178). At days 3, 5, 7 and 11


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post challenge, tracheal swabs were sampled in all birds for qRT-PCR assessment of virus excretion. All birds were humanely killed at 2 weeks post challenge for final serological testing. Assessment of clinical signs, antibody responses and virus replication Clinical signs were registered from day 0 to day 10 post inoculation. Serological testing for AMPV-C antibodies was performed as previously reported (Giraud et al., 1987), using an ELISA antigen derived from French AMPV-C isolate 99178 (Toquin et al., 1999). In such an assay, the positivity threshold was set to an ELISA ratio of 0.200. Quantitation of the number of copies of the SH gene was performed using a SH-gene specific qRT-PCR, as published by Guionie et al., 2007. RESULTS Controls on immunizing antigens No live-virus could be re isolated from the inactivated virus suspension. Neither mycoplasmas nor fungi could be detected. The serological screening following administration of the adjuvanted immunizing antigens did not reveal any antibody to any of the 18 bacteria and viruses tested. Clinical signs No clinical signs were observed in the different groups of ducks prior to the administration of the challenge virus. No clinical signs were observed following the inoculation of the challenge 99178 virus to White Peking ducks, irrespective of their immunization status. Only one or two Muscovy ducks exhibited mild to severe tracheal rales in the different immunized groups. Antibody responses Figure 1 presents the mean and range of ELISA ratios in the different groups at 5 weeks of age (= 3 weeks post priming), 7 weeks of age (= 2 weeks post boost), and 9 weeks of age (= 2 weeks post challenge). No antibodies to AMPV-C were detected in the Muscovy and White Peking non-immunized control groups prior to challenge with the virulent 99178 virus. In both Muscovy and White Peking ducks, the group immunized with AMPV-C exhibited significantly higher levels of antibodies to AMPV-C than the group receiving the mock antigen at 5 weeks of age. However, the mean antibody levels induced in White Peking ducks were very significantly higher than those observed in Muscovy ducks. These antibody levels were not improved by the second injection of the antigens, whereas challenge with the 99178 virus induced a sharp and significant rise of anti-AMPV-C antibodies in the three groups of Muscovy ducks and in the non-immunized group of White Peking ducks. Virus replication post challenge Figure 2 presents the mean and range of the number of RNA copies of the SH gene as detected at 3, 5, 7 and 11 days post challenge from the ducks in the different groups. In Muscovy Ducks, irrespective of the previous immunization schedule, all birds were qRT-PCR positive at 3 and 5 days post inoculation, and expression of the SH gene


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reached a peak (106.5 to 107.0 copies per reaction, no significant difference between groups) at 5 days post inoculation. The number of positive birds and level of SH expression then decreased slightly at day 7 post inoculation (no significant difference in expression levels between the groups) and no qRT-PCR positive Muscovy ducks were finally detected at day 11 post inoculation. In sharp contrast, in White Peking ducks and irrespective of the previous immunization schedule, no qRT-PCR positive bird was detected until day 7 post inoculation, when a single non immunized control proved weakly positive (104.2 copies per reaction). The number of positive birds in this group increased at day 11 post inoculation (5 positives), as did the mean level of SH expression (105.1 copies per reaction). No qRT-PCR positive White Peking ducks were detected in the groups that had previously received either the AMPV-C or the mock antigens. DISCUSSION In this paper we report on attempts to develop and evaluate in Muscovy and White Peking ducks an inactivated adjuvanted AMPV-C antigen. The culture to high titre in Vero cells of the studied AMPV-C isolate proved successful and the inactivation and purity controls proved the inactivated antigens to be suitable for in vivo evaluation. The implemented immunization scheme (2 doses, followed 3 weeks later by one dose, all by the intramuscular route) proved successful to induce specific anti-AMPV-C antibodies, especially in White Peking ducks, however the boost injection did not improve the antibody levels reached after the single prime injection. The challenge at seven weeks of age induced very limited signs in Muscovy ducks and no signs in White Peking ducks. In Muscovy ducks, this is most likely to be related to the age of the birds, as challenge with the 99178 virus in 3-week-old ducklings has been previously shown in the authors’ laboratory to typically induce infectious bronchitis-like rales in 90 % of the inoculated birds (Toquin et al., 2006a). Nevertheless, the infectivity of the challenge virus used in this experiment was clearly demonstrated by the sharp rise in the anti-AMPV-C antibody levels and the high expression of the SH gene in the trachea of all the inoculated Muscovy ducks, irrespective of their immunization scheme. The clinical protection could not be evaluated due to the low level of clinical signs in the different groups. Regarding the protection afforded against infection by the previous immunization, this appeared minimal in Muscovy ducks as the levels and kinetics of AMPV-C SH gene expression proved very similar in the group receiving the AMPV-C inactivated antigen, or the two mock and non-immunized control groups. These results are consistent with previous finding on AMPV inactivated vaccines in turkeys, as these vaccines proved unable to control virus replication at the tracheal level (Cook et al., 1996) when implemented without live priming. The results obtained in White Peking ducks appear somewhat different, but the experimental results make it difficult to conclude regarding protection, as the kinetics of virus excretion appear to be significantly different from that observed in Muscovy ducks. Indeed, the number of qRT-PCR positive White-Peking ducks seemed to be still increasing at day 11 post challenge, a time when the excretion had already ceased in Muscovy ducks. As no qRT-PCR positives were detected at 11 days post inoculation in the White Peking ducks that had been previously immunized with the


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AMPV-C antigen, it might be necessary to repeat the experiment in this species, with an extended period of qRT-PCR study, to determine whether the previous immunization may indeed interfere with virus replication. ACKNOWLEDGEMENTS To Dr Isabelle Kempf for her kind help in checking the antigens for possible mycoplasma contaminants. REFERENCES Bennett, R. S., McComb, B., Shin, H. J., Njenga, M. K., Nagaraja, K. V. and

Halvorson, D. A. (2002). Detection of avian pneumovirus in wild Canada (Branta canadensis) and blue-winged teal (Anas discors) geese. Avian Dis 46(4): 1025-9.

Cook, J. K., Orthel, F., Orbell, S. J., Woods, M. A. and Huggins, M. B. (1996). An experimental turkey rhinotracheitis (TRT) infection in breeding turkeys and the prevention of its clinical effects using live-attenuated and inactivated TRT vaccines. Avian Pathol 25: 231-243.

Cook, J. K. A., Huggins, M. B., Orbell, S. J. and Senne, D. A. (1999). Preliminary antigenic characterization of an avian pneumovirus isolated from commercial turkeys in Colorado, USA. Avian Pathol 28: 607-617.

Cook, J.K., Chesher, J., Orthel, F., Woods, M.A., Orbell, S.J., Baxendale, W., and Huggins M.B. (2000) Avian pneumovirus infection of laying hens: experimental studies. Avian Pathol. 29 : 545-556.

Giraud, P., Toquin, D., Picault, J. P., Guittet, M., L'Hospitalier, R., Kles, V. and Bennejean, G. (1987). Utilisation de la méthode Elisa pour le sérodiagnostic de l'infection par le virus de la rhinotrachéite infectieuse chez la dinde, le poulet et la pintade. Bulletin d'information des laboratoires de Services Vétérinaires 27/28: 65-70.

Guionie, O., Toquin, D., Sellal, E., Bouley, S., Zwingelstein, F., Allee, C., Bougeard, S., Lemiere, S. and Eterradossi, N. (2007). A laboratory evaluation of a quantitative real-time RT-PCR for the detection and identification of the four subgroups of avian metapneumoviruses. J Virol Methods 139(2): 150-8.

Jestin, V., Toquin, D., Le Bras, M. O. and Amenna, N. (2000). The new duck pneumovirus: experimental assessment of the pathogenicity for the respiratory tract of muscovy ducklings. 5th International congress of the European Society for Veterinary Virology, Brescia, Italy.

Senne, D.A., Edson, R.K., Pedersen, J.C. and Panigrahy, B. (1997) Avian pneumovirus update. In Proceediongs of the American Veterinary Medical Association, 134th Annual Conference, Reno, Nevada, USA, p 190.

Toquin, D., Bayon-Auboyer, M. H., Morin, H., Eterradossi, N. and Jestin, V. (1999). Isolation of a pneumovirus from a Musckovy duck. Vet. Rec. 145(23): 680.

Toquin, D., Guionie, O., Allee, C., Morin, Y., Le Coq, L., Zwingelstein, F., V., J. and N., E. (2006a). Compared susceptibility of SPF ducklings and SPF turkeys to the infection by avian metapneumovirus belonging to the four subgroups. 5th International Symposium on Avian Corona- and Pneumoviruses and Complicating Pathogens., Rauischholzhausen, Germany.


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Toquin, D., Guionie, O., Jestin, V., Zwingelstein, F., Allee, C. and Eterradossi, N. (2006b). European and American subgroup C isolates of avian metapneumovirus belong to different genetic lineages. Virus Genes 32(1): 97-103.

Figure 1 : Mean antibody production and range of antibody responses, in Muscovy and White Peking ducks receiving an experimental inactivated AMPV-C antigen, a non-infected mock antigen or no immunization (see in text the immunization schedules). All birds challenged with virulent AMPV-C at 7 weeks of age.


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Figure 2: Mean and range of number (Log10) of copies of the AMPV-C SH gene in tracheal swabs derived from Muscovy and White Peking ducks receiving an experimental inactivated AMPV-C antigen, a non-infected mock antigen or no immunization (see in text immunization schedules), at different times post challenge with virulent AMPV-C at 7 weeks of age. Numbers in white dots indicate the number of birds in each group that proved positive in qRT-PCR. NS: not significant according to the Kruskal Wallis test.


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PERITONITIS DUE TO ORNITHOBACTERIUM RHINOTRACHEALE CO-INFECTION WITH INFECTIOUS BRONCHITIS VIRUS AND ESCHERICHIA

COLI IN LAYING CHICKENS

THACHIL AJ1, VELAYUDHAN2 BT, SHAW3 DP, HALVORSON1 DA and NAGARAJA1 KV

1Department of Veterinary and Biomedical Sciences, University of Minnesota, 1971 Commonwealth Ave, Saint Paul, MN, 55108, USA, 2Center for Molecular Medicine

and Infectious Diseases, Virginia Tech, Blacksburg, VA, 3University of Missouri- Columbia, MO, USA

SUMMARY Respiratory disease conditions caused by co-infections with multiple pathogens are not uncommon in poultry and often results in heavy economic loss to the poultry industry. The objective of this study was to investigate the pathogenesis of ORT infection with exposure to infectious bronchitis virus (IBV) and/or Escherichia coli in laying chickens. In this study, eighty-week old SPF White Leghorn chickens were exposed experimentally to the following combinations: ORT alone; E. coli alone; IBV alone; ORT + E. coli; ORT + IBV; IBV + E. coli or ORT + IBV + E. coli. The clinical signs and pathological changes were evaluated in birds from all groups. Birds exposed to ORT or E. coli alone did not show any overt clinical signs. Birds exposed to IBV, ORT + IBV, IBV + E. coli and ORT + IBV + E. coli revealed clinical signs that varied from droopiness and nasal discharge to mortality. The ORT + IBV + E. coli co-infected birds exhibited gross lesions that included haemorrhagic tracheitis, fibrinous pneumonia, airsacculitis, pericarditis and peritonitis. Histopathological studies of this group revealed distinct pathological changes. Mortality following ORT infection was noticed only in ORT + IBV + E. coli co-infected group. Our results indicate that IBV and E. coli co-infection aggravates ORT pathogenesis in adult laying hens. The results also revealed that ORT persists in tissues like infraorbital sinuses for longer periods without causing much respiratory pathogenesis by itself. The persistence of ORT may predispose birds to develop peritonitis and death when predisposing conditions prevail like IBV and E. coli co-infections. INTRODUCTION Respiratory disease conditions continue to cause heavy economic losses to poultry producers because of the increased mortality rate, medication cost and


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condemnation rate in the processing plant. Ornithobacterium rhinotracheale (ORT) is an emerging respiratory pathogen of turkeys and chicken causing huge economic loss to the poultry industry (van Empel et al., 1996). The affected turkeys show depression, coughing, bloody nasal discharge and mortality (van Empel et al., 1999). ORT causes severe respiratory lesions like tracheitis, airsacculitis, fibrinous pneumonia and pericarditis in turkeys (van Empel et al., 1996, Hafez, 2002). In chickens, there are not many reports of isolation of ORT or cases of severe outbreaks or high mortality as has been observed in turkeys. The clinical disease manifestation by experimental ORT infection in broiler chickens is found to be much less severe than in turkeys and is limited to sneezing, rhinitis [Van Beek et al., 1994) and growth retardation without causing any macroscopic lesions. However, experimental infection with ORT alone and co-infection with Infectious bronchitis virus (IBV) through an intra-airsac route [van Empel et al, 1996) resulted in airsacculitis in broiler chickens. Leghorn chickens have been reported to be least susceptible to ORT infection as compared to broiler chickens and turkeys (Van veen etal, 2000). Though the clinical disease appears less severe in chickens, seroprevalence of ORT has been reported to be as high as 100% in broiler breeder flocks in Southern Brazil [Canal et al. 2003) and in layer flocks in North central regions of United States (Heeder etal, 2001). The objective of this study was to investigate the role of ORT in the pathogenesis of respiratory disease caused by concurrent IBV and E. coli infections.

MATERIALS and METHODS Inoculum Ornithobacterium rhinotracheale (strain 31C, Serotype C) was grown on 5% sheep blood agar (SBA) supplemented with gentamycin (10µg/ml) at 37o C with 5% carbon dioxide for 48 h. The ORT strain 31C was isolated from Leghorn chickens in US from an outbreak of acute respiratory illness. Escherichia coli serotype O 78 isolated from broiler chickens showing peritonitis was grown on Tryptic soya broth for 24 h at 37o C. Infectious bronchitis virus (Arkansas strain) was used in this study. The virus was passaged three times in 10-day-old embryonated eggs by allantoic route. The dose of the inoculum used in birds was 1 ml of 1x 106 mean EID50/ml/bird. Experimental Design One hundred and sixty, eighty-week-old SPF White Leghorn laying hens were used in this study. The birds were kept in individual layer cages on layer feed and water ad libitum. All the birds were tested negative for any respiratory pathogens by bacterial culture of tracheal swabs and for antibodies to ORT by the serum plate agglutination test (Back et al., 1998) and ELISA (Lopes et al., 2000). The birds were randomly divided into groups I through VIII containing twenty birds each. Birds in groups III, V, VI and VII were inoculated intra-tracheally with 1 ml of 1x 106 mean EID50 of IBV/bird. Birds in groups II, IV, VI and VII were aerosolized with one liter each of 2x108 CFU/ml of E.coli serotype O78 using a power sprayer. On 5-day post IBV/E.coli infection, birds in groups I, IV, V and VII were inoculated intra-tracheally with one milliliter of 108 CFU of ORT/bird made in sterile phosphate buffer saline (PBS). Birds in group VIII were inoculated intra-tracheally with sterile PBS and used as sham inoculated controls.


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All the birds were then monitored daily for any clinical signs or mortality. Two birds from each group were euthanized on 2, 4, 7, 14, 21 and 28 days post-ORT-infection and necropsy was conducted. Euthanasia was performed in a carbon dioxide chamber. Gross lesions were recorded in each bird. Tissues were collected in sterile Whirl-pak filter bags for bacteriologic examination or in 10% buffered neutral formalin for immunohistochemistry and histopathology. Samples of trachea, lungs, liver, spleen, kidney, oviduct and bursa were homogenized and inoculated on SBA and MacConkey agar for isolation and identification of ORT and E.coli, respectively. In addition, sterile swabs were collected in PBS from air sac and peritoneum for isolation and identification of ORT and E.coli. After 24 h, suspected colonies were sub-cultured and tested for oxidase and catalase activity. Samples of trachea and lungs were homogenized in PBS containing penicillin-streptomycin and egg-inoculated for isolation of IBV. Infectious bronchitis virus was later confirmed by RT-PCR. Blood samples were collected from treatment and control groups at 0, 7, 14 and 21 days post-ORT infection. The sera were examined for the presence of antibodies against ORT by ELISA. The hematoxylin and eosin (H&E) staining technique was used for histopathological examination. A scoring system (lesion score varied from 1 to 4 depending on the increased severity of the lesions) was used to evaluate histopathological changes in different tissues for analysis and comparison. Trachea, lungs, liver, spleen, kidney and oviduct were subjected to immunohistochemistry for detecting ORT antigens. One way analysis of variance was employed for statistical analysis of the data. A P value of 0.05 was considered as significant. RESULTS Birds infected with IBV alone showed ruffled feathers and droopiness whereas birds infected with IBV + E .coli showed more pronounced clinical signs like nasal discharge and off feed. Two birds in group VII (IBV + E. coli + ORT) and three birds in group VI (IBV + E. coli) succumbed to death on 2 day post-IBV + E. coli infection. Following ORT infection, no mortality was noticed in any groups until 16 days post ORT infection. In the IBV + E. coli + ORT infected group, one bird succumbed to death on 16-days post-ORT infection. Although egg production dropped in the ORT and E. coli alone infected groups when compared with non-infected control birds in group VIII, an abrupt cessation of egg production was noticed in all the birds in the IBV infected groups. Airsacculitis, lung congestion and pericarditis were noticed in most of the birds infected with IBV + E. coli and IBV + E. coli + ORT on 2 and 4 days post- ORT infection during necropsy. Peritonitis was noticed in two birds necropsied from the IBV + E. coli + ORT infected group on 7 and 14 days post-ORT infection. One bird from this group, which died on 16 day-post-ORT infection showed pericarditis, perihepatitis and peritonitis (Figure 1A). In addition, an accumulation of amber colored fluid (Figure 1C), the presence of cheesy deposits (Figure 1D) and thickening of visceral peritoneum was noticed in all the three affected birds.No gross pathological changes were noticed in any of the tissues in birds infected with ORT or E. coli alone. In birds of group I, ORT was isolated from sinus, trachea, airsacs, liver and kidney. In groups IV, V, and VII, ORT was isolated from all the above organs as well as from


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lungs and oviduct. We isolated ORT from air sacs and trachea even on 14 day post-ORT infection and from sinuses up to 28 day post-ORT infection IBV + E. coli + ORT infected group. In 20-50% of all the ORT infected birds we were able to isolate ORT from the infraorbital sinuses on 28 days post-ORT infection. From birds in groups IV, VI and VII, E. coli was isolated from the sinuses and trachea on days 2 and 4 post-ORT-infection. Both ORT and E. coli were isolated from the peritoneal swabs collected from all the three birds having peritonitis. Microscopic lesions were noticed on histopathological examination in the tracheal tissues of birds in group I to VII which showed infiltration of lymphocytes, plasma cells and macrophages in the lamina propria. Lymphoid aggregates were present in severely affected tracheas which caused the mucosa to bulge into the lumen. A few of the glands were dilated and empty and the cilia were disrupted. There were moderate numbers of necrotic cells in the BALT. There were focal areas of fibrin exudation and infiltration of inflammatory cells in the lungs in birds infected with IBV+ ORT and IBV + E.coli+ ORT. Liver sections showed infiltration of portal tracts by varying number of lymphocytes and macrophages and fibrin deposition. The peak histopathologic lesion score was obtained on fourth day post-ORT infection. The overall histopathologic lesion score observed following ORT infection alone (Group I) was found to be statistically significant (P<0.05) when compared with the control group (group VIII). The histopathologic lesion score was numerically higher in ORT groups co-infected with IBV and E. coli (groups IV and V) but was not statistically significant (P>0.05) when compared with the lesion scores in ORT alone infected groups. However, the overall histopathologic lesion score was significantly higher (P<0.001) for the group infected with ORT + IBV + E. coli when compared with any other groups. Birds in the infected and sham-inoculated groups were negative for the presence of ORT antibodies in the serum when tested by ORT ELISA at day 0 post-infection. All birds infected with ORT became seropositive to ORT by ELISA on day 7 of ORT infection. The ORT antibody titers remained high even on 28 day post-ORT infection. DISCUSSION In this study, the authors investigated the pathogenesis of a potential pathogenic isolate of ORT serotype C isolated from a severe field outbreak in adult White Leghorn chickens. ORT is well known for causing acute respiratory clinical signs and lesions in turkeys and it is also attributed as a primary respiratory pathogen in broiler chickens [van Veen et al, 2000). However, the role of ORT as a primary pathogen in laying chickens is not certain. Results of this study demonstrate that ORT infection alone has little effect on adult White Leghorn hens but co-infections with IBV or E. coli can cause severe pathogenesis. In a previous study conducted in 26-day-old leghorn chickens, the microscopic lesions due to ORT infection were restricted to one-day-post-infection only with minimal infiltration of polymorphonuclear granulocytes in the airsacs [ven Empel., et al, 1999). Our results indicated that while experimental infection with ORT alone in adult Leghorn birds produced consistent microscopic lesions in the trachea and lungs, this was not enough to cause any overt clinical signs, gross pathological changes or mortality. The differences found in the current study and the previously


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cited studies may be attributed to the type or virulence of the strain of ORT used, or tolerance to ORT by adult layer chickens. In an experimental study with five-week-old turkeys infected with ORT and E. coli, (Droula et al, 1997) found that E. coli did not appear to enhance the pathogenicity of ORT. No association was also reported in the mortality pattern in colibacillosis affected layer hen flocks in the field due to ORT interaction [Vanderkerchove, et al, 2004). On the contrary, Elgohary et al.1998, noticed an increased severity of lesions associated with ORT in a concomitant E. coli infection in 38-day-old broiler chickens. The overall histopathological lesion score in our study suggests an enhancement of ORT pathogenesis by concomitant E. coli as well as IBV + E. coli co-infection in Leghorn chickens. Experimental infection with IBV (Arkansas strain) alone in nine-day-old leghorn chickens reported no mortality though airsacculitis was present. However, when co-infected with IBV and E. coli, ORT produced severe respiratory lesions and mortality [20] similar to the findings in our study. In broiler chickens, concomitant ORT infection with other respiratory pathogens like Newcastle disease virus (Travers et al., 1996), IBV or E. coli is attributed as the cause for severe respiratory lesions like airsacculitis and pneumonia (odor et al., 1997). Our study demonstrates that ORT co-infection with IBV and IBV + E. coli exacerbate the clinical signs as well as lesions like airsacculitis, tracheitis, pneumonia and pericarditis in Leghorn chickens. Similar findings were reported with experimental co-infections with IBV + ORT in broiler chickens (Franz et al., 1997). When we co-infected with IBV + E. coli+ ORT, the infected birds showed significantly higher clinical sign scores compared with any other groups and also showed peritonitis and mortality over a period of time. These findings validate the lesions including peritonitis reported from prior ORT field outbreaks in commercial adult layer birds Sprenger et al., 2000). In the present study, we isolated ORT from air sacs and trachea even after 14-day-post-infection and from sinuses even on 28-day-post-infection. In a previous study ORT could not be isolated at time points later than 2-day-post-infection from 26-day-old Leghorn chickens (van Empel et al., 1999). The bacterial isolation results we obtained indicated that ORT could persist in adult layer birds for longer periods especially in the infraorbital sinuses without producing visible clinical signs or gross pathological lesions. We suspect that ORT persistence by colonization in the nasal sinuses in adult chickens for longer periods may lead to more severe respiratory disease when the predisposing conditions prevail. Ornithobacterium rhinotracheale antigen detected in the liver and spleen indicated a possible systemic bacterial infection. This suggested that under field conditions this isolate of ORT might become an invasive agent in adult layer chickens. The White Leghorn chickens were found to be not as susceptible to ORT infection as broilers and turkeys (Elgohary et al., 1998). The results of clinical signs, gross lesions, mortality and histopathology suggest that the pathogenesis of ORT either alone or in co-infection was less severe in adult layer chickens when compared with previous reports in broiler chickens as well as in turkeys (Lopes et al., 2002). The pathogenicity indices and clinicopathologic examination in the present study demonstrated that both IBV and E. coli collectively or individually could contribute to ORT pathogenesis and thus increasing the severity of the disease in White Leghorn hens. In laying hens, ORT could persist in tissues for longer periods without causing much respiratory disease and this may predispose layer birds to develop peritonitis


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and death. The chronic role of ORT as a pathogenic agent in adult laying chickens has to be considered during disease diagnosis along with other infectious agents.

REFERENCES Avellenda, G.E., P.Villegas, M. W. Jackwood, and D.J. King. 1994. In vivo evaluation

of the pathogenicity of field isolates of infectious bronchitis virus. Avian Dis. 38:589-597.

Back, A., G. Rajashekara, D.A. Halvorson, and K.V. Nagaraja. 1998. Development of a serum plate agglutination test to detect antibodies to Ornithobacterium rhinotracheale. J. Vet. Diag. Invest. 10:84-86.

Back. A., D.A. Halvorson, and K.V. Nagaraja. 2002. Minimization of pathologic changes in Ornithobacterium rhinotracheale infection in turkeys by temperature sensitive mutant strain. Avian Dis. 46: 177-185.

Canal, C.W., J.A. Leao, D.J. Ferreira, M. Macagnan, C.T. Pippi Salle, and A. Back. 2003. Prevalence of antibodies against Ornithobacterium rhinotracheale in broilers and breeders in Southern Brazil. Avian Dis. 47: 731-737.

Charlton, B. R., A. J. Bermudez, M. Boulianne, R. J. Eckroade, J. S. Jeffrey, L. J. Newman, J. E. Sandeer, and P. S. Wakenell. 1996. Ornithobacterium rhinotracheale Infection. Pages 128-129 in Avian Disease Manua. American Association of Avian Pathologists, University of Pennsylvania, New Bolton Center, Kennett Square, PA.

Droual, R., and R.P. Chin. 1997. Interaction of Ornithobacterium rhinotracheale and Escherichia coli O78, H9 when inoculated into the air sac in turkey poults. Page 11 in Proc. 46th Western Poult. Dis. Conf., Sacramento, CA.

Elgohary, A.A., and M. H. H. Awaad. 1998. Concomitant Ornithobacterium rhinotracheale and E. coli infection in chicken broilers. Vet. Med. J. Giza. 45:65-75.

El-Sukhon, S.N., A. Musa, and M. Al-Attar. 2002. Studies on the bacterial etiology of airsaculitis of broilers in northern and middle Jordan with special reference to Escherichia coli, Ornithobacterium rhinotracheale, and Foretell valium. Avian Dis. 46:605-612.

Franz, G., R. Hein, J. Bricker, P. Walls, E. Odor , M. Salem, and B. Sample. 1997. Experimental studies in broilers with a Delmarva Ornithobacterium rhinotracheale isolate. Pages 46-48 in Proc. 46th Western Poult.Dis. Conf., Sacramento, CA.

Hafez, H. M. 2002. Diagnosis of Ornithobacterium rhinotracheale. International J. Poul. Sci. 19:114-118.

Heeder, C.J., V.C. Lopes, K.V. Nagaraja, D.P. Shaw, and D.A. Halvorson, 2001. Seroprevalence of Ornithobacterium rhinotracheale infection in commercial laying hens in the north central region of the United States. Avian Dis. 45: 1064-1067.Lopes, V.C., A.

Lopes, V.C., G. Rajashekara, A. Back, D. P. Shaw, D.A. Halvorson, and K.V. Nagaraja. 2000. Outer membrane proteins for serological detection of Ornithobacterium rhinotracheale infection in turkeys. Avian Dis. 44: 957-962.

Odor, E.M., M. Salem, C. R. Pope, B. Sample, M. Primm, and M. Murphy. 1997. Isolation and identification of Ornithobacterium rhinotracheale from commercial broiler flocks on the Delmarva Peninsula. Avian Dis. 41:257-260.


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Sprenger, S.J., A. Back, D. P. Shaw, K. V. Nagaraja. D.C. Roepke, and D. A. Halvorson. 1998. Ornithobacterium rhinotracheale infection in turkeys: experimental reproduction of the disease. Avian Dis. 42:154-161.

Sprenger, S.J., D.A. Halvorson, K.V. Nagaraja, , R. Spasojevic, R. S. Dutton, and D. P. Shaw. 2000. Ornithobacterium rhinotracheale infection in commercial laying type chickens. Avian Dis. 44: 725-729.

Travers, A.F. 1996. Concomitant Ornithobacterium rhinotracheale and Newcastle Disease infection in broilers in South Africa. Avian Dis. 40:488-490.

Van Beek, P. N. G. M., P. C. van Empel, G. van der Bosch, P. K. Storm, J. H. Bongers, and J. H. Du Preez. 1994. Respiratory, growth retardation and arthritis in turkeys and broilers caused by a Pasteurella-like organism: Ornithobacterium rhinotracheale or Taxon 28. Tijdsch voor Diergen, 119:99-101.

van Empel, P., M. Vrijenhoek , D. Goovaerts, and H. van der Bosch. (1999). Immunohistochemical and serological investigation of experimental Ornithobacterium rhinotracheale infection in chickens. Avian Pathol. 28:187-193.

van Empel, P, and H. M. Hafez. 1999. Ornithobacterium rhinotracheale a review. Avian Pathol. 28:217-227.

van Empel, P., H. van der Bosch, D. Goovaerts, and P. Storm. 1996. Experimental infection in turkeys and chickens with Ornithobacterium rhinotracheale. Avian Dis. 40:858-864.

van Veen, L.P., P. van Empel, and T. Fabri. 2000. Ornithobacterium rhinotracheale, a primary pathogen in broilers. Avian Dis. 4:896-900.

Vandekerchove, D., P. D. Herdt, H. Laevens, P. Butaye, G. Meulemans, and F. Pasmans. 2004. Significance of interactions between Escherichia coli and respiratory pathogens in layer hen flocks suffering from colibacillosis-associated mortality. Avian Pathol. 33:298-302.

Acknowledgements This work was supported by Midwest Poultry Consortium and University of Minnesota. The authors are thankful to Vanessa Lopes-Berkas for her valuable laboratory help and technical advice.


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A B

C D

Figure 1: Gross lesions and histopathologic changes following exposure to ORT + E. coli + IBV in layer chickens. (A) pericarditis (white arrow head) and airsacculitis (Black arrow head). (B) Histopathology section of lung stained with H & E showing fibrin deposition (Black arrow head) (magnification = 40x). (C) Presence of amber colored fluid in the peritoneum and thickening of the peritoneum (E) Presence of cheesy deposit on the liver and peritoneum ( Black arrow head).


371

ROLE OF UNUSUAL MYCOPLASMAS IN POULTRY – M. LIPOFACIENS AS AN EXAMPLE

LIERZ M

Klinikum Veterinarum, Clinic for Birds, Reptiles, Amphibians and Fish, Justus Liebig University Giessen, Frankfurter Strasse 91-93,

35392 Giessen, Germany

SUMMARY Mycoplasmas are well known pathogens in poultry medicine and of special importance in breeder flocks due to vertical transmission and their pathogenic potential for the embryo and poults. Apart from well known Mycoplasma species, M. lipofaciens (strain ML64), isolated from an egg of a northern goshawk (Accipiter gentilis), was found to be pathogenic for chicken and turkey embryos causing a high mortality. Additionally, horizontal transmission between turkey poults hatched from infected eggs and poults from non-infected controls, was observed in the incubator, which is of importance for spreading the pathogens between flocks. As M. lipofaciens was already isolated from adult chickens and turkey, the above described results raise new concerns about diagnosing mycoplasmas in poultry breeder flocks as M. lipofaciens (strain ML64) might be undiagnosed by using laboratory methods focussing on well known poultry pathogenic Mycoplasma species only. Therefore, a Mycoplasma- genus specific PCR was developed and evaluated using all known avian Mycoplasma reference strains. Additionally a so far unidentified Mycoplasma strain was isolated from different raptors causing false positive results in a M. meleagridis specific PCR which is routinely used in turkey breeder flocks. As host specificity of avian mycoplasmas seem not be very strict, the occurrence of unusual or new mycoplasmas from non-poultry species should be considered when investigating poultry flocks. INTRODUCTION Mycoplasmas are well known pathogens in avian medicine and infections in poultry are mostly accompanied with severe economic problems. They might be a primary pathogen, pave the way for other infections or act as a complicating agent in a disease process. The infection spread by vertical and horizontal means. In the past mycoplasmas appeared to have a restricted host range. This does not seem to be


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true, since several avian mycoplasmas have been isolated from a number of avian hosts including birds of prey (Lierz et al., 2002; Bradbury, 2006). M. lipofaciens was first described in 1983 as a new species isolated from the infraorbital sinus of a healthy chicken (Bradbury et al, 1983). Afterwards M. lipofaciens had only been reported from the chicken, turkey and duck (Bencina et al, 1987). Recently, Lierz et al (2007a) isolated M. lipofaciens from an egg of a Northern Goshawk (Accipiter gentilis). Differentiation was made by immuno binding assay as well as sequencing parts (928bp) of the 16S rRNA gene (GenBank Accession No: DQ653410). Therefore M. lipofaciens seems capable of crossing avian taxonomic divides, and is not specific to a single avian group. Further, this species can be transmitted vertically. In poultry the vertical transmission of mycoplasmas (M. synoviae, M. gallisepticum, M. meleagridis, M. iowae) is well known, causing embryonic death and poorly developed embryos, regularly without causing clinical symptoms in the parent flock. (Yamamoto and Orthmeyer 1966; Carpenter et al, 1981; Lin and Kleven 1982; Glisson and Kleven 1984; Glisson and Kleven 1985). Therefore poultry flocks, especially breeders, are routinely examined for the occurrence of the four well known pathogenic Mycoplasma species by serology and PCR. PATHOGENICITY OF M. LIPOFACIENS FOR CHICKEN AND TURKEY EMBRYOS As M. lipofaciens was demonstrated in several avian species including poultry and seems to be capable of beeing transmitted vertically, the pathogenic potential of a non-poultry strain (ML 64 from a Northern Goshawk) for chicken and turkey embryos was questioned. In a first study Lierz et al. (2007b) demonstrated a high mortality of chicken embryos infected with M. lipofaciens (strain ML64). Major alterations were liver necrosis. In a second study Lierz et al. (2007c) investigated the pathogenicity of this strain in turkey embryos compared to chicken embryos as well as the possibility of a horizontal transmission between poults in the hatcher. Seven days after infection (day 7 of incubation) 12 of 15 infected chicken embryos were dead. Of the 45 turkey embryos, infected at day 8 of incubation via yolk sac with different infectious doses, only 5 hatched. In comparison to the chicken embryos, the turkey embryos died significantly later, mainly immediately prior hatch (19-20 days post infection). Five of six turkey embryos from the negative control group hatched without problems. M. lipofaciens was reisolated from all infected embryos, but not from the control group, which remained also negative in a Mycoplasma- Genus specific PCR (Lierz et al., 2007a). Main macroscopical alterations of the deceased embryos were petechial bleedings, curled toes and dwarfing, comparable to the alterations known from the poultry pathogenic species after vertical transmission. Histologically, a purulent- necrotizing pneumonia was the main alterations in the turkey embryos. Interestingly, the air sacs were without any alterations. Additionally, the lateral spread of the pathogen between poults of the infected and the control group within the hatcher was demonstrated. Last but not least, M. lipofaciens was demonstrated for 5 days in the nose of the study veterinarian, indicating a possible role of humans as a vector for transmission (Lierz et al., 2008c).


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UNIDENTIFIED MYCOPLASMA STRAIN FROM RAPTORS The role of mycoplasmas in poultry is well investigated. In contrast, studies focusing on mycoplasmas in non-poultry avian species are rare or even lacking. In several studies investigating the occurence of mycoplasmas in different raptor populations, Lierz et al. (2008a,b,d,e) demonstrated that more than 90% of the birds are positive for mycoplasmas. However, the four known poultry pathogenic species were not detected (Lierz et al., 2008d). In this study the authors obtained positive results from raptor samples in a PCR which was found to be specific for M. meleagridis when testing nearly all avian and 9 mammalian Mycoplasma reference strains. Using restriction enzyme analysis and sequencing three of the obtained amplificates from raptor samples, the results were identified as false positive. The sequences of the amplificates were identical but demonstrated only a 91% homology to the comparable sequence of M. meleagridis. Although the sequence showed a 98.5% homology to those of M. buteonis, the isolate were antigenetically distinct. Therefore it is likely that this species represent a so far unknown species. As host specificity in avian mycoplasmas seem not be very strict (Bradbury, 2006) and the PCR for the detection of M. meleagridis is routinely used in turkey breeders, the possibility of an introduction of this new species into turkey flocks and the occurrence of false positive results in breeder monitoring should be considered and deserves further attention. DISCUSSION The occurence of mycoplasmas in poultry is always of major concern as usually high economic losses are the results. Therefore poultry flocks, especially breeders, are regular investigated for the occurrence of mycoplasmas, focusing on the four relevant and well investigated species M. gallisepticum, M. iowae, M. meleagridis and M. synoviae. Attention is seldom given to other Mycoplasma species, ignoring the fact that host specificity in avian mycoplasmas seem not be very strict (Bradbury, 2006). Of special importance managing poultry breeder flocks is the vertical transmission of mycoplasmas as they may cause embryonic death or problems in the offspring flocks, but the breeders remain clinically healthy. This is especially true for M. meleagridis and M. iowae. M. lipofaciens was isolated from chicken and turkeys (Bradbury et al, 1983; Bencina et al, 1987) and also from an egg of a northern goshawk (Lierz et al., 2007a) demonstrating the potential to be vertical transmitted and the absence of a strict host specificity. This is underlined by the fact that this species was demonstrated for 5 days in a human nose (Lierz et al., 2008c). This Mycoplasma species is highly pathogenic for chicken and turkey embryos (Lierz et al., 2007b,c). Chicken embryos died significantly earlier than turkey embryos, which died mainly immediate prior or during hatch. Interestingly, the histopathological findings in turkey embryos are mainly located in the lung with a severe purulent to necrotizing pneumonia compared to chicken embryos with the major findings within the liver. This might be the reason why chicken embryos died earlier after infection as they use the liver already during embryonic development. In opposite, the lung is needed directly prior pipping onwards only, when the poult enter the air bulb of the egg and changes to lung breathing. It surprises that the air sac were without histopathological alterations but the same was described for M. iowae in chickens (Bradbury & Mc Carthy, 1983).


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Additionally, the lateral transmission of M. lipofaciens (strain ML64) was clearly demonstrated between turkey poults within the incubator, having great impact of spreading the pathogen within the turkey population. As the symptoms of a M. lipofaciens infection in breeders (healthy adults, dead in shell embryos with petechial bleedings, curled toes, dwarfing) are very similar to those compared to other Mycoplasma infections, investigations into the cause of such clinical signs should be reconsidered. Poultry breeder flocks are monitored to be free for mycoplasmas using serology and PCR (Hafez & Jodas, 1997). Usually the laboratory tests used are specific to a certain Mycoplasma species in particular M. meleagridis, M. iowae, M. gallisepticum or M. synovia. In case of an infection with M. lipofaciens the laboratory tests focussing on these Mycoplasma species will reveal negative results. Therefore, the Mycoplasma- Genus specific PCR (Lierz et al, 2007a), which was evaluated using nearly all known avian Mycoplasma reference strains is recommended to use it in case of a Mycoplasma suspicion in poultry, especially if negative results were obtained in the other investigations. A potentially new Mycoplasma species from raptors caused false positive results in a PCR which is routinely used in turkey breeders to monitor the occurrence of M. meleagridis. This is remarkable as this PCR was thought to be specific when testing nearly all avian and further mammalian Mycoplasma reference strains. Despite the fact that turkey has not yet been identified as a potential host for this species, this possibility should be taken into account. This underlines the importance to consider other mycoplasmas than the known poultry pathogenic species in differential diagnosis or as complicating factors in poultry and the related diagnostic problems this may cause. ACKNOWLEDGEMENT The manuscript is based on studies performed at the Institute for Poultry Diseases, Free University of Berlin. The author is very thankful to the staff of this institution, especially to his mentor, Prof. Dr. H.M. Hafez, for the given opportunities to perform such research and the constant support. REFERENCES Bencina, D., Dorrer, D. & Tadina, T. (1987). Mycoplasma species isolated from six

avian species. Avian Pathology, 16, 653-664 Bradbury, J. M. & McCarthy J. D. (1983). Pathogenicity of Mycoplasma iowae for chick

embryos. Avian Pathology, 12, 483-496 Bradbury, J. M., Forrest, M. & Williams, A. (1983). Mycoplasma lipofaciens, a new

species of avian origin. International Journal of Systematic Bacteriology, 33(2), 329-335

Bradbury, J. M.. (2006). Mycoplasmas: a century and not out. Turkeys, 54, 9-11 Carpenter, T. E., Edson, R. K & Yamamoto, R. (1981). Decreased hatchability of turkey

eggs caused by experimental infection with Mycoplasma meleagridis. Avian Diseases, 25(1), 151-156

Glisson, J. R. & Kleven, S. H. (1984). Mycoplasma gallisepticum vaccination: effects on egg transmission and egg production. Avian Diseases, 28(2), 406-415


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Glisson, J. R. & Kleven, S. H. (1985). Mycoplasma gallisepticum vaccination: further studies on egg transmission and egg production. Avian Diseases, 29(2), 408-415

Hafez H.M. & Jodas, S. (1997). Mykoplasmose. In: Putenkrankheiten: Hafez HM und Jodas S (Hrsg.). Putenkrankheiten. Enke Verlag, Stuttgart, pp. 86-93.

Kotani, H. & Mc Garrity, G. J. (1985). Identification of mycoplasma colonies by immunobinding. Journal of Clinical Microbiology, 23, 783-785

Lierz, M., Schmidt, R. & Runge, M. (2002). Mycoplasma species isolated from falcons in the Middle East. Veterinary Record, 151, 92-93

Lierz, M., Hagen, N., Harcourt-Brown, N., Hernandez-Divers, S.J., Lueschow, D. & Hafez, H.M. (2007a). Prevalence of mycoplasma in eggs from birds of prey using culture and a genus-specific mycoplasma-PCR. Avian Pathology, 36, 145-150

Lierz, M., Stark, R., Brokat, S. & Hafez, H.M. (2007b). Pathogenicity of M. lipofaciens strain ML64, isolated from an egg of a Northern Goshawk (Accipiter gentilis), for chicken embryos. Avian Pathology, 36, 151-153

Lierz, M. Deppenmeier, S., Gruber, A.D., Brokat, S. & Hafez, H.M. (2007c). Pathogenicity of Mycoplasma lipofaciens strain ML64 for turkey embryos. Avian Pathology, 36, 389-393

Lierz, M., Hagen, N., Lueschow, D. & Hafez, H.M. (2008a). Species- Specific Polymerase Chain Reactions for the Detection of Mycoplasma buteonis, Mycoplasma falconis, Mycoplasma gypis and Mycoplasma corogypsi in Captive Birds of Prey. Avian Diseases, 52, 94-99

Lierz, M., Hagen, N., Hernandez-Divers, S.J. & Hafez, H.M. (2008b). Occurrence of mycoplasmas in free-ranging birds of prey in Germany. Journal of Wildlife Diseases, 44, 845-850

Lierz, M., Jansen, A. & Hafez, H.M. (2008c). Avian Mycoplasma lipofaciens Transmission to Veterinarian. Emerging Infectious Diseases, 14 (7), 1161-1163

Lierz, M., Hagen, N., Lueschow, D. & Hafez, H.M. (2008d). Use of polymerase chain reactions for the detection of M. gallisepticum, M. imitans, M. iowae, M. meleagridis and M. synoviae in birds of prey. Avian Pathology, 37, 471-476

Lierz, M., Obon, E., Schink, B., Carbonell, F. & Hafez, H.M. (2008e). The role of mycoplasmas in a conservation project of the Lesser Kestrel (Falco naumanni). Avian Diseases, 52, 641-645

Lin, M. Y. & Kleven, S. H. (1982). Egg transmission of two strains of Mycoplasma gallisepticum in chickens. Avian Diseases, 26(3), 487-495

Yamamoto, R. & Orthmeyer, H. B. (1966). Pathogenicity of Mycoplasma meleagridis for turkey and chicken embryos. Avian Diseases, 10, 268-272


376

IBD EFFICACY DATA IN PRESENCE OF MATERNAL ANTIBODIES IN BROILERS AND LAYERS WITH A NOVEL VECTOR HVT-IBD VACCINE

LE GROS FX

Merial SAS, Lyon Gerland Laboratory, 254 rue Marcel Mérieux, 69007 Lyon, France

SUMMARY Vaccination programmes for Infectious Bursal Disease (IBD) were studied in presence of maternal immunity in broiler and layer type birds. The vaccine take elicited by a turkey herpes virus (HVT)-IBDV commercial vaccine (VAXXITEK® HVT+IBD) administered by the subcutaneous route at the hatchery, was compared in two independent trials to the ones obtained from standard vaccine schedules using oral administrations of attenuated IBD vaccines after 2 weeks of age. The vaccine take was followed by serology and the protections verified by a challenge with a very virulent IBD isolate at the age of 30 or 42 days respectively for the broiler and layer trial. The protection criterion was based on post mortem examination of the bursa of Fabricius 10-11 days post challenge (bursa/body weight ratio). In both trials the serological data indicated an early onset of immunity using the vector vaccine when an immunity gap was evidenced on the groups receiving the standard vaccination schedules after the decay of maternal antibodies. This was confirmed by the challenge results performed during that immunity gap. Poor to no protection was observed in groups receiving the attenuated IBD vaccines when a large and significant protection was evidenced in groups receiving the vector vaccine. It was concluded that opposite to attenuated IBD vaccines, the vector vaccine was not interfered by the IBD maternal immunity. INTRODUCTION Vaccination with attenuated virus is widely used worldwide to control infectious bursal disease in broilers (IBD), an economically important disease of the young chickens (van den Berg, 2000). One of the major problems with attenuated IBD vaccines is their sensitivity to maternally-derived antibodies (MDA) which are always present at the time of vaccination. One way to approach this issue is the use of less attenuated (“intermediate” or “hot / intermediate-plus”) vaccines, but these vaccines can themselves cause a degree of vaccine-induced lesions in the bursa of Fabricius as evidenced in Specific Pathogen free (SPF) birds (Rautenschlein, 2005). Therefore, a ® VAXXITEK is a registered trademark of Merial in the United States of America and elsewhere.


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new vaccine combining excellent safety and efficacy in the presence of IBDV MDA is needed. The widely used Marek’s disease vaccine based upon cell-associated herpes virus of turkeys (HVT) is well-known to be safe and poorly sensitive to MDA interference, which is why it was developed as a vector for IBD (Darteil, 1995; Goutebroze, 2003) and other diseases (Morgan, 1992). The HVT vector virus was generated by inserting an IBDV VP2 gene expression cassette into the HVT genome. The objective of these trials was to verify in field conditions (presence of maternal immunity in broiler and layer type birds) the properties of this novel vaccine as compared to intermediate and “hot” IBD vaccines. Both were recently published separately (Massi, 2008; Le Gros, 2009) and this paper is reviewing them in a synthetic discussion. MATERIALS and METHODS All vaccines used were commercially available and delivered as per manufacturer’s instructions. The HVT vector vaccine (VAXXITEK® HVT+IBD, Merial - later on mentioned as vHVT13 strain) was available as a cell associated frozen vaccine. The donor virus for the VP2 expression cassette in vHVT13 was the IBDV strain of classical virulence F52/70. The first trial was recently reported in details as a field trial (Le Gros; 2009); briefly vHVT13 vaccine was injected subcutaneously at the hatchery in the new born chicks before placement on farm A. The hatch mates placed on farm B received a classical HVT vaccine on the same day. A registered IBD intermediate vaccine was delivered twice to the birds through drinking water on farm B, at the age of 17 and 24 days. Other vaccinations against Infectious Bronchitis and Newcastle diseases were similar on the two sites. On each of the two farms, blood samples were randomly taken from 20 birds at the age of 1, 17, 26, 45, and 56 days. The sera were tested for IBD antibodies using the ELISA kit ProFLOK® Plus IBD Ab test kit (Synbiotics) according to the manufacturer specifications. When necessary the average antibody titres per group were compared using a Student t test. In order to perform an IBD experimental challenge, groups of 15 birds were moved from the hatchery or the farms to the animal room facilities at different times as follows, and summarised in table 1. G.00: before vaccination (30 birds), this group was then divided in two groups of 15 each, receiving or not the infectious challenge (G.01, G.02). G.A: from farm A, on day 14; G.B: from farm B at the age of 25 days (after live IBD vaccination). A group of SPF birds of the same age was also constituted (G.SPF). All the groups, except G.01 were challenged at the age of 30 days with an IBDV very virulent strain (ref: 77165) obtained from Istituto Zooprofilattico Sperimentale della Lombardia e dell’Emilia Romagna (Martin, 2007) The challenge dose was 5.2 log10 EID50 per bird administered via the ocular route via a volume of 50µL. Clinical signs were observed for 10 days post challenge; then the birds were euthanized and necropsied for B/Bw ratio measurement. Due to non-homogeneity of the results variances, these ratios were compared by non- parametric tests: global comparison by one-way analysis of variance using a Kruskal-Wallis test and group to group comparisons by Wilcoxon test with Bonferroni adjustment for multiple comparisons.

® PROFLOK is a registered trademark of Synbiotics Corporation in the United Kingdom and elsewhere.


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The second trial was as well reported in details recently (Massi; 2008); briefly 170 brown commercial pullets were utilized. At 1 day of age, 20 pullets were blood sampled to evaluate the MDA titres. The remaining 150 day-old pullets were split into six groups (G.1 to G.6) each containing 25 birds and placed in isolators as summarized in table 2. The vaccines A and B were both “Intermediate” IBDV vaccines. A commercial dose of these was administered to birds of group 1 and 2, respectively, by oral drop at day 17 and 25. The vaccine C was a “hot” IBDV vaccine (also mentioned as “intermediate-plus”). Birds of group 3 were vaccinated with C once at 17 days of age by oral drop. Vaccination time was chosen in order to simulate the IBD vaccination schemes commonly applied in field conditions. The vaccine D was vHVT13. Chicks of group 4 were injected subcutaneously at 1 day of age with this vaccine. Birds from group 5 and 6 were not vaccinated. Blood samples were taken from twenty 1-day-old pullets to assess the level of IBD MDA. Afterwards, all the groups were blood sampled at the ages of 17, 31, 42 (challenge day) and 53 days. In order to evaluate the impact of vaccination on the bursa of Fabricius, whole bursas were randomly taken from 10 birds per group at 31 days of age, i.e. 6 days after the second vaccination in groups 1 and 2, and 15 and 31 days after vaccination in groups 3 and 4, respectively. At 53 days of age, i.e. 11 days after challenge, whole bursas were randomly taken from all the remaining pullets to evaluate the effects of the challenge in the different groups. Evaluation was based upon B/Bw ratio. All the groups, except group 6, were challenged at the age of 42 days with the same very virulent strain of IBDV and dose than for the first trial, administered by an oral drop. RESULTS During the first trial, after the expected decrease of the maternal immunity a clear serological conversion to IBDV antigen was evidenced in both farms as shown in figure 1. However, the antibody level was significantly higher in farm A than in farm B between the ages of 26 to 45 days. After the experimental challenge, as expected clinical symptoms and mortality were mainly visible in G.SPF: half of the birds were affected and 3 out of 10 died. Only one animal from the control group G.02 displayed clinical signs after challenge. The challenge effect was more obvious on post-mortem examination. The typical bursal oedema was only visible on G.SPF (3 out of 10 birds), but the bursal atrophy was very clear in the control challenged group G.02, opposed to the control unchallenged G.01. The protection afforded by the vaccines could thus be judged based on these comparisons. This is shown in figure 2, illustrating the B/Bw average ratio observed in the 4 broiler groups. G.SPF was not included in the figure as this type of birds (layer type) was not comparable with the broiler type on this criterion. There was a global statistical difference within the 4 groups (p<10-8). G.A was comparable to the unchallenged group G.01 and different from the challenged group G.02, and was thus clearly protected from the challenge. G.B was comparable to the challenged control group G.02 and appeared thus not protected. A threshold value of B/Bw ratio for protection could be defined from the individual values in G.01 and G.02. The score 1.12 ‰ was the lowest value observed in the non challenged birds and only 1/15 birds from the challenged group G.02 was above this value. According to this protection criterion (B/Bw ratio ≥1.12 ‰) 93% of G.A birds (14 out of 15) were protected, compared to 6% in G.B or G.02 (1 out of 15). However in the absence of


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B/Bw evaluation before challenge it cannot be ruled out that the lesions observed in G.B may be due to the vaccine itself and not to the challenge strain. Given the serological observations showing no positive titre at time of challenge we hypothesise that this was not the case. Observations in trial 2 confirmed this hypothesis. Concerning the second trial, before challenge a decline of antibody titres was observed in all groups except in the vHVT13 vaccinated group where antibody titres assessed by IBD Plus ELISA test remained high (>6000) and stable as shown in figure 3 with some statistical indications. The lack of detectable antibody response in the live IBD vaccinated groups on day 42 was probably due to the interference of MDA at the time of vaccination which hampered and/or delayed a correct vaccine take. At day of age MDA showed to be very high as a consequence of the IBD vaccination of parents using inactivated oil emulsified vaccines. After challenge, with both ELISA tests, a marked increase in antibody titres was observed in all groups, except in the vHVT13 vaccinated group which had a less pronounced conversion. These serological observations were largely confirmed by the results of the experimental challenge. In the group of pullets unvaccinated and challenged (G.5), severe clinical signs of IBD such as depression, ruffled feathers and watery diarrhoea were observed in all of the birds and two out of fifteen died. In both groups vaccinated with intermediate IBD vaccines (G.1 and G.2) two out of fifteen birds showed clinical signs of IBD and died. In the group vaccinated with the “intermediate- plus” IBD vaccine (G.3) two out of fifteen birds showed clinical signs of IBD and one of them died. Neither clinical signs of IBD nor mortality were observed in the group vaccinated with vHVT13 (G.4) and in the control group unvaccinated and unchallenged (G.6). At post mortem examination, gross lesions typical of IBD (oedema of bursa of Fabricius, nephritis, haemorrhages in the muscles and on the mucosa of proventriculus) were observed only in the birds of groups 1, 2, 3 and 5 which died following vvIBDV challenge. No lesions were recorded in the group vaccinated with vHVT13 (G.4) and in the group unvaccinated and unchallenged (G.6). The size of bursas of Fabricius taken at 31 days of age (11 days before challenge) and at 53 day (11 days after challenge) is shown in figure 4. No significant differences in B/Bw ratio between the groups were observed before vvIBDV challenge, whereas after challenge this ratio decreased dramatically in the group unvaccinated (G.5) and in all the groups vaccinated with live IBD vaccines (G.1, 2 and 3). On the contrary bursa size was not negatively affected in the unchallenged group (G.6) and only slightly impacted in the vHVT13 group (G.4).


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DISCUSSION These studies aimed to compare the immunogenicity and efficacy of IBD vaccination programmes using two types of IBD vaccines in broilers and layers. Comprehensive and consistent set of results were obtained. The serological observations illustrated the difficulties in managing a vaccine program with attenuated intermediate vaccines in presence of MDA. In the first trial the absence of serological conversion in farm B at the age of 25 days, strongly indicates a failure of the vaccination after the first administration at the age of 17 days and a partial interference of remaining MDA on the second administration at the age of 24 days. The increase of ELISA antibodies detectable at a later age (day 45) indicates also a late vaccine take. Similar and even more critical observations were obtained in the second trial as the decay of MDA in layer type birds is known to be slower than in broilers and thus the interference against live IBD vaccinations stronger. As a mater of fact, no serological conversion was obtained in any of the groups receiving the live IBD vaccine programs including either 2 administrations of intermediate or 1 of an “intermediate-plus” vaccine. In both trials, as a result of this negative interference these birds were likely still relying on passive immunity to face the experimental challenges. Based on B/Bw criteria the protection observed after challenge in both trials in groups receiving the live IBD vaccines was poor. However in the first trial as already mentioned, in absence of B/Bw evaluation before challenge it cannot be ruled out that the lesions observed in G.B may be due to the vaccine itself and not to the challenge strain. Furthermore birds of G.B and most of them in G.02 did not show clinical signs in contrast to birds from G.SPF, as a result of the residual passive protection and/or the well known lower sensitivity to IBD of broiler type compared to layer type birds. Nonetheless, in the second trial where pre-challenge B/Bw observations and clinical signs post challenge were available, the poor protection resulting from IBD live vaccine programs was well established. Despite this negative interference, a late onset of immunity can still arise from these vaccine programs as visible in the first trial, but the birds are then “at risk” for a period of time comprised between the sufficient decay of the passive immunity and the effective induced active immunity which was estimated to be around the age of 40 days in the broiler experiment. This window or immunity gap is a classical observation in attenuated IBD vaccine programmes and is often approached in the field by the use of multiple administrations or more invasive “intermediate-plus” vaccines (van den Berg, 2000). In these studies these standard vaccination schedules were not efficacious enough to cover this immunity gap. In the same conditions of interfering passive immunity, the vector vaccine vHVT13 although delivered at the age of one-day when MDA levels were at their highest level, elicited an active immune response clearly evidenced at the age of 26 days in broilers and 31 days in layers using the PROFLOK Plus IBD Ab test kit. This early vaccine take was further illustrated by the results of the experimental challenges at the age of 30 and 42 days in broilers and layers respectively, There was thus no detectable immunity gap in this vector vaccine programme. These field trial observations were consistent with vaccine development data already reported (Le Gros, 2005; Bublot, 2007) showing an effective vaccine take in presence of MDA, together with an absence of bursal lesions after vaccination. The cell-associated nature of vHVT13, the lack of expression of the IBD VP2 on the


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surface of the infected cells and the mode of replication of the HVT vector, probably all contribute to the ability of the vaccine to overcome MDA. Similar immunogenicity induced by another HVT-vectored IBD vaccine has also been previously reported in SPF birds (Tsukamoto, 2002). In conclusion these trials further confirmed the higher performance on bursal integrity, either as a result of higher safety or efficacy, of a live vector IBD vaccine injected at the hatchery compared to the classical attenuated IBD vaccines delivered at the farm. REFERENCES Bublot, M., Pritchard, N., Le Gros, F. X., and Goutebroze, S. (2007) Use of a

vectored vaccine against Infectious Bursal Disease of chickens in the face of high-titrated maternal derived antibody. J. Comp. Path. 137, S81-S84.

Darteil, R., Bublot, M., Laplace, E., Bouquet, J. F., Audonnet, J. C., Riviere, M. (1995) Herpesvirus of turkey recombinant viruses expressing infectious bursal disease virus (IBDV) VP2 immunogen induce protection against an IBDV virulent challenge in chickens. Virology; 211: 481-490.

Goutebroze, S., Curet, M., Jay, M.L., Roux, C., Le Gros, F.X. (2003) Efficacy of a recombinant HVT-VP2 against Gumboro disease in the presence of maternal antibodies. British poultry Science; 44, 824-825.

Le Gros, F.X., Goutebroze, S., Bublot, M. (2005) Efficacy of and HVT/VP2 vectored vaccine against vvIBD challenge after in ovo injection in presence of IBD maternal antibodies. Proceedings of the 14th World veterinary Poultry Congress, Istanbul, 279.

Le Gros, F.X., Dancer, A., Giacomini, C., Pizzoni, L., Bublot, M., Graziani, M., Prandini, F. (2009) Field efficacy trial of a novel HVT-IBD vector vaccine for one day-old broilers; Vaccine, vol 27 n°4, 22 January 2009, 592-596.

Martin, A.M, Fallacara, F., Barbieri, I., et al. (2007) Genetic and antigenic characterization of infectious bursal disease viruses isolated in Italy during the period 2002-2005. Avian Dis 51(4), 863-72.

Massi, P., Tosi, G., Fiorentini, L.(2008) Experimental challenge trial with a “very virulent” strain of Infectious Bursal Disease virus (vvIBDV) in commercial pullets vaccinated with an IBD vectored vaccine or with three different modified live vaccines , Zootecnica, nov 08, 50-57.

Morgan, R. W., Gelb, J. Jr., Schreurs, C. S., Lutticken, D., Rosenberger, J. K., Sondermeijer, P. J. (1992) Protection of chickens from Newcastle and Marek's diseases with a recombinant herpesvirus of turkeys vaccine expressing the Newcastle disease virus fusion protein. Avian Dis; 36: 858-870.

Rautenschlein, S., Kraemer, C., Vanmarcke, J., and Montiel, E. (2005) Protective efficacy of intermediate and intermediate plus infectious bursal disease virus (IBDV) vaccines against very virulent IBDV in commercial broilers. Avian Dis; 49: 231-237.

Tsukamoto, K., Saito, S., Saeki, S. et al. (2002) Complete, long-lasting protection against lethal infectious bursal disease virus challenge by a single vaccination with an avian herpesvirus vector expressing VP2 antigens. J Virol; 76: 5637-5645.


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Van den Berg, T., Eterradossi, N., Toquin, D., Meulemans, G. (2000) Infectious Bursal Disease (Gumboro disease), Rev. Sci. tech., Aug. 2000; 19(2):509-43.

Animal origin Identification at the farm level

Identification for the experimental challenge IBD Challenge

G.01* No NA

G.02* Yes Farm A: vHVT13

vaccine G.A Yes Commercial

hatchery

Farm B: ML IBD vaccine G.B Yes

SPF quarantine NA G.SPF Yes

NA: not applicable *:G.00 before challenge Table1: Description of the various study groups in the first trial and their relations.

Group Vaccination Vaccine Doses Vaccination timing (days)

vvIBDV challenge at 42 days of age

G.1 Yes A - “Intermediate” 2 17 and 25 Yes G.2 Yes B - “Intermediate” 2 17 and 25 Yes

G.3 Yes C - “Intermediate plus” 1 17 Yes

G.4 Yes D - vHVT13 (Vaxxitek) 1 1 Yes

G.5 No Unvaccinated, challenged - - Yes

G.6 No Unvaccinated, unchallenged - - No

Table 2: Summary of the study groups and treatments in the second trial.


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PROFLOK Plus IBD Ab test kit

0200040006000

80001000012000

0 10 20 30 40 50 60

age in days

aver

age

antib

ody

titre

farm A farm B

Figure 1. Kinetics of ELISA antibodies using the PROFLOK Plus IBD Ab test kit, showing the decrease of the maternal immunity followed by the active immune conversion. Farm A received the vHVT13 vaccine and Farm B the intermediate IBD vaccine. The average titre is shown for each farm and sampling time with its 95% confidence interval. Statistically significant differences were detected at the ages of 26 and 45 days.

0,00

0,50

1,00

1,50

2,00

2,50

3,00

G.01 G.02 G.A G.B

Groups

Burs

a/

body w

eig

ht

rati

o *

1000

a

b b

a

Figure 2. Average bursa/body weight ratio per group 10 days post-experimental challenge, presented with its 95% confidence interval. G.01: unchallenged control; G.02: challenged control; G.A: vHVT013 vaccine; G.B: ML IBD vaccine. a,b: Different letters indicate a statistically significant difference.


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PROFLOK Plus IBD Ab test

0

2000

4000

6000

8000

10000

12000

14000

day 1 day 17 day 31 day 42 day 53

Mea

n E

LIS

A ti

ter

Intermediate vaccine on days 17 + 25 Vaxxi tek SC on day 1Intermediate vaccine on days 17 + 25 Intermediate Plus vaccine on day 17Unvaccinated unchallenged Unvaccinated challenged

challenge

Figure 3: IBDV mean antibody titres detected using the PROFLOK Plus IBD Ab test. There was a statistically significant difference on D31 and D42 within all groups (variance analysis, p<0.01). On D31, G.4 was higher than G.5 (p=0.045) and G.5 higher than the comparable groups G1, 2, 3 and G.6 (p=0.01). On D42, G.4 was higher than the groups G.1, 2 and 3 (p<0.01), themselves higher than G. 5 and 6 (p<0.01).

Average bursa/body weight ratios

012345678

Group 1 Group 2 Group 3 Group 4 Group 5 Group 6

31 days 53 days

Figure 4. Mean B/Bw ratio before and after challenge at 42 days in the second trial, with standard deviation. At 53 days, there was a statistically significant difference within the groups (variance analysis, p<0.01); G.1, 2, 3 and 5 were not different (p=0.52); G.4 and G.6 were significantly different (p=0.01) with G.6>G.4.


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NOVEL HVT/ILT RECOMBINANT VACCINE TO SIMULTANEOUSLY CONTROL INFECTIOUS LARYNGOTRACHEITIS AND

MAREK’S DISEASE IN CHICKENS (INNOVAX®-ILT)

HEIN R

Intervet Schering Plough Animal Health 29160 Intervet Lane, PO Box 318, Millsboro, DE 19966

Infectious Laryngotracheitis (ILT) is an acute viral disease primarily in chickens. Depending on the pathotype it may causes severe respiratory disease with excessive mortality particularly in broilers. In addition, a decrease in egg production may be observed in egg producers.

Currently, ILT is controlled by vaccination using modified live Chicken Embryo Orgin (CEO) and Tissue Culture Orgin (TCO) type vaccines. If properly applied, these vaccines are highly efficacious. However, these live vaccines have been associated with adverse effects and present additional concerns.

• Individual application by eye drop-stress/labor • Short duration of protection ,revaccination in mature birds • Undesirable vaccination reaction • Spread vaccine virus to non-vaccinates, resulting increase of virulence • Latency/persistence of vaccine virus in vaccinated birds

And last but not least, concerns regarding the reactivation of the latent vaccine virus with the increase of virulence causing the common vaccine related ILT breaks. A novel HVT/ILT recombinant vaccine, the Turkey Herpes Virus vector ILT recombinant (rHVT/ILT) INNOVAX®-LT has been developed, which has shown to be safe and not cause the adverse effects observed with the conventional live ILT vaccines. This vaccine was introduced in late 2007 in the US and provides protection against ILTV and Marek’s Disease. The rHVT/ILT construct consists of the HVT FC126 with the glycoprotein gI and gD gene inserts from ILTV. These gene inserts are encoding the immunogenic ILTV viral membranes glycoproteins D and I. The gI and gD genes are with their regulatory elements inserted in the nonessential unique long (UL) region of the HVT genome.


386

This recombinant can be used by the in-ovo and subcutaneous (1 day of age) routes. In areas where virulent MDV is prevalent, the vaccine can be combined with the MDV vaccine SB1 or Rispens (CVI988). The vaccine does not cause respiratory reactions, latency as observed with the live ILT vaccines, and safety is comparable with the parent HVT vaccine strain. Broilers and SPF chickens vaccinated in-ovo or s.c at one day and challenged at around 4 weeks of age by the infraorbital or ocular routes with virulent ILT showed to be fully protected based on ILT clinical respiratory signs. In a duration of immunity study, SPF chickens vaccinated by the s.c route at 1 day of age with the recombinant and challenged with virulent ILT by the ocular route at different ages showed to be fully protected up to 60 weeks of age. In the standard MDV challenge test, SPF chickens were vaccinated s.c at one day of age with the rHVT/ILT and challenged 5 days post-vaccination by the intraabdominal route with vMDV.In that study excellent MD protection, comparable with the conventional HVT inovo vaccinated birds was observed. In addition,a MD shedder trail (“one day of age contact challenge”) in broilers vaccinated by the inovo routewith the rHVT/ILT + Rispens(CVI988) and challenged with the very virulent plus(vv+) MDV was carried out in comparison with the conventional HVT+Rispens(CVI988) in-ovo vaccinated broilers. No statistical difference in MD protection was observed in this severe challenge between birds vaccinated with the rHVT/ILT+Rispens and birds vaccinated with the conventional HVT+Rispens In this challenge model, 1 day-old non vaccinated broilers (shedders) are inoculated intra-abdominally with very virulent plus(vv+)MDV and 10 days after placement of shedders 1 day old broiler vaccinates and unvaccinated contacts are placed with shedders. Vaccinated and unvaccinated survivors are removed and necropsied at 7 weeks of age. Protection was based on MD mortality and macroscopic (gross) lesions. Since the introduction of the rHVT/ILT (INNOVAX®-LT) in the US, over 1 billion broilers and approximately 90 million commercial layers/breeders have been successfully vaccinated with this vaccine In broilers vaccination has been carried out in- ovo and in several cases in combination with MD vaccine SB1. Commercial layers/breeders are vaccinated s.c at 1day of age and in most cases, in combination with the Rispens (CVI988) MD vaccine In summary the advantages of the this novel recombinant HVT/ILT in comparison with the conventional live ILT vaccines are;

No Respiratory Vaccination Reaction /No Negative Effect on Performance No Spread of ILTV-No risk Reversion to Virulence No Latency-No risk or reactivation and Vaccine Related ILT Breaks No interference with IBV and NDV vaccinations

Extended duration of protection (life long after one vaccination) Convenient administration (in ovo or sc at one day of age)

Simultaneous protection against vMDV


387

FINAL REFLECTIONS

JONES RC Chairman of the final discussion session on avian coronaviruses and

metapneumoviruses

AVIAN CORONAVIRUS – INFECTIOUS BRONCHITIS VIRUS Epidemiology Epidemiological information on infectious diseases in recent years is now almost completely based on molecular identification and sequence analysis of viruses and this is the case with IB. The distribution of the well known genotypes of IBV and some of the more recently identified variants in different parts of the world shows that some appear to be very widespread, while others are more localised. With molecular techniques it is now possible to trace the movement of new types with molecular epidemiology but the means of global spread of IBVs is largely unknown. There is no compelling evidence yet that wild birds are important for this. Several reports described the situation in European countries. Longitudinal studies of viruses in IBV outbreaks in Denmark indicated fine amino acid changes in parts of the S1 spike gene. Whether this is due to mutation or selection of particular clones from a mixed viral population is unknown. In Slovenia, four different genotypes were described and interestingly, Italy 02 predominates in the West, while QX does in the East. QX genotype has spread slowly to the ‘extremities’ of Europe, arriving in Spain as late as 2008 (in the UK also), where 4/91 has been displaced by Italy 02. A pan-European survey for 2008 showed that as in a previous study, 793B and Massachusetts types predominated, followed by D274, then QX and Italy 02. Dutch type V1395 re-appeared and 642-1 was detected. Italy 02 was seen only in Mediterranean countries. Is this decline due to the increased use of double vaccination with two different vaccine types or was Italy 02 never a variant of major importance? 4/91 and D274 genotypes have been shown to be present in Jordan and Brazilian strains can be grouped into three unique clusters plus 793B. In both these countries, as in several others, only H120 vaccine is licensed for use and this presents a real dilemma for a rational approach to IBV control. Mainland China was shown to have up to 7 genotypes but different results are obtained depending on whether comparisons are made with S1 or N gene sequences. Taiwan was shown to have two indigenous groups, again different from previously described IBVs.


388

All these presentations serve to illustrate the range of recognised and variant IBV types in different parts of the world, with different countries showing different patterns of prevalence. It reinforces the need for continuous surveillance of types in all regions in order to assist in control strategies. Virus properties and virus-host interactions This section included a range of different topics, covering aspects of replication at vaccine virus molecular and the host cellular levels. In Iran, RFPL was used to differentiate between 793B and homologous vaccines IB88 and 4/91. IN the USA, after routine vaccination with a live IBV vaccine, virus with altered amino acid sequences were found to be detectable in different tissues (see Denmark reference above). Again, is this due to mutation or selection from the original mixed virus population? The significance of minor viral subtypes of the Arkansas vaccine was reviewed: why does the Ark vaccine vary? Does this phenomenon occur with other live IBV vaccines? NF-κB (nuclear factor kappa-light-chain-enhancer of activated B cells) is a protein complex that acts as a transcription factor which has been shown to be important in regulating the immune response to IBV. A study of the role of structural and accessory genes showed that gene 1 of IBV is important in pathogenicity. Sialic acid is necessary for IBV infection of the tracheal epithelium of the chicken. High throughput proteome screening reveals novel host cell-virus interactions including nucleocytoplasmic trafficking. Diagnosis As with most other viral infections, diagnosis of IB is now virtually completely based on molecular methodology. This was reflected in the papers in this session, with none on serology or more traditional methods such as cultivation. However, it should not be forgotten that fur further studies on the virus, such as empirical vaccine production or experimental challenge, live virus is required. Different forms of RT-PCR with further variant identification, including a high resolution melt curve system described here. The value of safe transport of virus on FTA cards was discussed. The establishment and early evaluation of a microarray system for avian viruses including IBV was presented and this is likely to be valuable in the search for poultry viruses including IBV and aMPV in wild avian species. Pathogenesis and immunology Disappointingly, only two papers were presented in this session. In the race to develop new improved vaccines, these aspects of IBV study have not received the attention they deserve. Nonetheless, they are vital to understanding of vaccine formulation and efficacy. One presentation looked at possible differences between a respiratory and an apparent enteric strain of IBV, while a second addressed the acute interferon gamma production in lymphocytes by IBV. Vaccination The appearance of important IBV variants is the biggest problem for IB control worldwide, since the established vaccines may fail in the face of a novel genotype challenge. One approach is to use a combination of two different IBV vaccines (usually a Mass type followed by a 793B type) and this is widely used and has


389

proved very successful in many instances against challenge with viruses quite unrelated to either vaccine. Several papers were presented on this topic, including one attempting to understand the underlying immunological basis for the protocol. The other approach is to tailor vaccine according to the immediate challenge, using S1 or N protein expression or ‘spike swapping’. Both these were covered in this session, together with in ovo delivery of an IBV vaccine. AVIAN CORONAVIRUS - TURKEY CORONAVIRUS In the past, research on turkey coronaviruses (TCov) has been hampered by their inability to be cultured easily and to some extent, still is. However the interesting presentations in this session illustrated the value of molecular techniques in identification, divergence and classification of types, details of replication and pathogenesis and trials with DNA-mediated single protein vaccines. An experimental trial showed that in chicks the enteric virus replicated in the respiratory tract, posing the question as to whether chickens can be reservoirs of TCov. AVIAN METAPNEUMOVIRUS Epidemiology Papers in this section raised concerns about the efficacy of aMPV vaccines. Two separate groups presented evidence from Brazil that there are different subtypes or lineages circulating in the chicken populations that may avoid vaccinal protection. This prompts the question posed of IBV as to whether the virus is mutating or whether immune selection allows variant forms to flourish from the originally diverse population of virus. In Italy it has been shown that outbreaks of disease can occur due to virulent virus derived from vaccine virus. This phenomenon is thought to be caused by initial incomplete vaccine cover in the flock, allowing cycling and exaltation of vaccine among initially unvaccinated individuals. The American subtype C aMPV is more closely related to the human metapneumovirus than are subtypes A or B. Turkeys inoculated with human MPV showed illness which was milder than caused by subtype C virus Virus properties and diagnosis This session comprised several papers relating more to virus properties than diagnosis. Regions of the fusion protein of subtype A aMPV have been shown to be recognised by neutralising antibodies. Studies of this kind are important in formulation of novel vaccines. Evidence was presented of aMPV attachment protein evolution associated with mass live vaccination in Italy. A study of the full length sequences of chicken- and turkey-derived aMPVs revealed no species specific motifs, therefore obviating the need for species-specific vaccines. Reverse genetics was used to develop a positive control virus for RT-nested PCR detection for subtypes A and B. Modification to the SH gene resulted in aMPV producing an altered cytopathic effect (CPE) in Vero cells and this may have implications in viral pathogenesis. In a related study, replacement of the SH gene by a green fluorescent protein (GFP) did not


390

prevent the development of the unusual CPE. After infection of turkeys, in vivo, a GFP-containing aMPV was found to localise in the upper respiratory tract. Finally, generation and biological assessment of recombinant subtype C with different lengths of the G gene were assessed. Vaccination and immunity A comprehensive review of vaccination against avian metapneumoviruses after 30 years indicated that the original empirical vaccines currently used are still effective provided that the recommended dose is delivered accurately to each bird. However, evidence was presented that aMPV evolution is occurring and new variants are appearing that are avoiding vaccine-induced immunity. In another paper on apparent aMPV vaccine failure in turkeys, poor aMPV vaccine performance was considered due to turkey astrovirus persistence preventing a complete immune response. The roles of humoral and cell mediated immunity on aMPV infection was described. Experimental trials testing inactivated oil-adjuvanted aMPV subtype C vaccine vs SPF Muscovy ducks were reported. CONCLUSIONS IBV continues to fascinate and confuse with more and more genotypes being described constantly, presenting a real dilemma for control particularly in countries where only one vaccine is licensed. The long-term answer to IBV control is still uncertain. Other puzzling aspects of IBV include our ignorance of how they spread globally, perhaps mediated by wild birds and their involvement in other disease manifestations including enteric disease and male infertility. There is still much to be learned about pathogenesis and immunity to IBV. Advances are being made in our understanding of turkey coronaviruses, thanks to molecular methods. However their true involvement in disease and the influence of other enteric pathogens need further work. Much has been learned concerning the molecular virology of avian metapneumoviruses and reverse genetics offers a safe effective solution to the problems associated with live vaccine reversion. However, the empirical vaccines can still be very effective provided that each bird receives the correct dose. Evidence that the established vaccines may not be effective against variant field viruses is cause for concern. Again, the international spread of avian metapneumoviruses is poorly understood.


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PARTICIPANT ADDRESS LIST

ABD EL RAHMAN, Sahar Institute of Virology University of Veterinary Medicine Hannover Bünteweg 17 30559 Hannover Germany [email protected]

ALROUSSAN, Dirgam Ahmad Provimi Jordan Airport Road 499 11118 Amman Jordan [email protected]

ASSAYAG Jr., Mario Sérgio University of Sao Paulo Orestes Farina, 94 –Cobertura Concordia-S.C. 89700-000 Brazil [email protected]

ARICIBASI, Merve Lohmann Animal Health Heinz-Lohmann-Str. 4 27472 Cuxhaven Germany [email protected]

AUGUSTINSKI, Karsten Lohmann Animal Health Heinz-Lohmann-Str. 4 27472 Cuxhaven Germany [email protected]

BLOCK, Hermann Gruppenpraxis Meyer-Block-Thien Am Rott 12 D-49843 Uelsen Germany [email protected]

BRANDAO, Paulo Eduardo Dept. Prev. Vet. Med & Animal Health University of Sao Paulo Av. Prof. Dr. Orlando M.Paiva, 87 Sao Paulo 05508-270 Brazil [email protected]

BRITTON, Paul Institute for Animal Health Compton Laboratory, Division of Microbiologyy Compton, Newbury Berks. RG20 7NN United Kingdom [email protected]

BROWN, Paul University of Liverpool Department of Veterinary Pathology Leahurst, Neston South Wirral CH64 7TE United Kingdom [email protected]

CARDOSO, Tereza Christina Laboratory of Animal Virology Sao Paulo State University Rua Clovis Pestana 793 Aracatuba, SP CEP 16050-680 Brazil [email protected]

CATELLI, Elena Dept. Pub. Vet. Health & Animal Pathology Fac. Vet.Med, University of Bologna Via Tolara Di Sopra, 50 40064 Ozzano Emilia Italy [email protected]

CECCHINATO, Mattia Dept Pub. Health, Animal Pathology & Hygiene Fac. Vet. Med, University of Padua Viale dell’Universita, 16 35020 Legnaro (PD) Italy [email protected]














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CHACON, Jorge Luis Ceva-Phylaxia Veterinary Biologicals Co. Ltd. Szallas u. 5 H-1107 Budapest Hungary [email protected]

CHEN, Hui-Wen School of Veterinary Medicine National Taiwan University No. 1 Sec. 4 Roosevelt Rd Taipei 10617 Taiwan [email protected]

CLUBBE, Jayne University of Liverpool Department of Veterinary Pathology Leahurst, Neston South Wirral CH64 7TE United Kingdom [email protected]

COOK, Jane K.A. 138 Hartford Road Huntingdon Cambridgeshire PE29 1XQ United Kingdom [email protected]

CSEREP, Tibor Intervet/Schering-Plough Animal Health (UK) Walton Manor, Walton Milton Keynes MK7 7AJ United Kingdom [email protected]

DE WIT, J.J. GD Deventer Animal Health Service Laboratory Department P.O. Box 9 7400 AA Deventer The Netherlands [email protected]

DOLZ, Roser Centre de Recerca en Sanitat Animal Edifici CreSA, Campus UAB Universitat Autònoma de Barcelona 08193 Bellaterra, Barcelona Spain [email protected]

DRYGIN, Vladimir Federal Centre for Animal Health 600901 Vladimir Russia [email protected]

EL HOUADFI, Mohammed Institut Agronomique et Veterinaire Hassan II BP 6202, Institut Rabat 10101 Morocco [email protected]

EMMOTT, Ed Institute of Molecular and Cellular Biology Garstang of Leeds Garstang Room 8.58 LS2 9JT, Leeds United Kingdom [email protected]

ERIKSSON, Helena National Veterinary Institute Dept. Aniaml Health and Antimicrobioal Strategies SE-75189 Uppsala Sweden [email protected]

ETERRADOSSI, Nicolas French Agency for Food Safety (AFSSA) Avian and Rabbit Virology, Immunology and Parasit. Unit P. O. Box 53 22440 Ploufragan France [email protected]

EVEN-CHEN, Tal Abic Biological Laboratories Ltd. West Industrial Zone, POB 489 Bet-Shemesh 99100 Israel [email protected]

FALCHIERI, Marco University of Liverpool Dep. Of Veterinary Pathology, Jordan building Leahurst Campus Neaston CH64 7TE United Kingdom [email protected]
















393

GANAPATHY, Kannan University of Liverpool Department of Veterinary Pathology, Jordan Building Leahurst, Neston South Wirral CH64 7TE United Kingdom [email protected]

GEERLIGS, Harm Fort Dodge Animal Health Van Houtenlaan 36 1381 CP Weesp The Netherlands [email protected]

GOMAA, Maged Hassan Department of Virology, Faculty of Veterinary. Medicine Kafr EL Sheikh University El Geish street 33516 Kafr EL Sheikh Egypt

HAFEZ, Hafez M. Institute for Poultry Diseases Free University Berlin Königsweg 63 14163 Berlin Germany [email protected]

HAGEN, Stephan Lohmann Animal Health Heinz-Lohmann-Str. 4 27472 Cuxhaven Germany [email protected]

HANDBERG, Kurt National Veterinary Institute Technical University of Denmark Hangoevej 5 Aarhus N, DK-8200 Denmark [email protected]

HEFFELS-REDMANN, Ursula Clinic for Birds, Reptiles, Amphibians & Fish Justus Liebig University Giessen Frankfurter Str. 91 35392 Giessen Germany [email protected]

HEIN, Ruud Intervet/Schering Plough Animal Health Scientific Affairs and Lab. Services Poultry 29160 Intervet Lane, P.O. Box 318 Millsboro, DE 19966 USA [email protected]

HEINE, Sandra Idexx Laboratories Mörikestr. 28/3 71636 Ludwigsburg Germany [email protected]

HEWSON, Kylie The University of Melbourne 250 Princes Highway Werribee 3030 Australia [email protected]

JACKWOOD, Mark College of Veterinary medicine University of Georgia 953 College Station Road Athens, GA 30602 USA [email protected]

JONES, Richard C. University of Liverpool Department of Veterinary Pathology, Jordan Building Leahurst, Neston South Wirral CH64 7TE United Kingdom [email protected]

JUNGBÄCK, Carmen Paul-Ehrlich-Institut Paul-Ehrlich-Straße 51-59 D-63225 Langen Germany [email protected]

KALETA, Erhard F. Clinic for Birds, Reptiles, Amphibians & Fish Justus Liebig University Giessen Frankfurter Str. 91 35392 Giessen Germany [email protected]















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KHALESI, Bahman Poultry Viral Vaccine Dept. Razi Vaccine & Serum Research Institute Hasark, Shahid Beheshti Street Karaj 3197619751 Iran khalesi@[email protected]

KRAPEZ, Uros Inst. for Poultry Health, Vet. Fac. University of Ljubljana Gerbiceva 60 1000 Ljubljana Slovenia [email protected]

KUANA, Suzete Lora Perdigao S.A. Saul Brandalise, 39 Videira 89.560-000 Brazil [email protected]

LE GROS, Francois-Xavier Merial 29, Avenue Tony Garnier BP 7123 F-69348 Lyon Cedex 07 France [email protected]

LIERZ, Michael Clinic for Birds, Reptiles, Amphibians & Fish Justus Liebig University Giessen Frankfurter Str. 91 35392 Giessen Germany [email protected]

LEMIERE, Stéphane Merial 29, Avenue Tony Garnier BP 7123 F-69348 Lyon Cedex 07 France [email protected]

LIN, Fengsheng Intervet/Scgering Plough Animal Health (UK) Walton Manor, Walton Milton Keynes MK7 7AJ United Kingdom [email protected]

LIN, Tsang Long Purdue University ADDL 406 South University Street West Lafayette, IN 47907-2065 USA [email protected]

LÜSCHOW, Dörte Institute for Poultry Diseases Free University Berlin Königsweg 63 14163 Berlin Germany [email protected]

LUPINI, Caterina Dept. Pub. Vet. Health & Animal Pathology Fac. Vet.Med, University of Bologna Via Tolara Di Sopra, 50 40064 Ozzano Emilia Italy [email protected]

MAJDANI, Raheleh Faculty of Veterinary Medicine Urmia University Sero Road, P.O. Box 1177 Urmia, West Azarbaijan Iran [email protected]

MALO, Aris Intervet/Schering Plough Animal health P.O. Box 31 5830 AA Boxmeer The Netherlands [email protected]

MAUREL, Stephan French Agency for Food Safety (AFSSA) Avian and Rabbit Virology, Immunology and Parasit. Unit P. O. Box 53 22440 Ploufragan France [email protected]

MC CRORY, Sarah Institute of Molecular and Cellular Biology University of Leeds Garstang Room 8.58 Leeds LS2 9JT United Kingdom [email protected]

mailto:khalesi@20022002















395

MEIR, Rosie Kimron Veterinary Institute P.O. Box 12 Bet Dagan 50250 Israel [email protected]

MOTITSCHKE, Andreas Paul-Ehrlich-Institut Paul-Ehrlich-Str. 51-95 63225 Langen Germany [email protected]

MONNE, Isabella Istituto Zooprofilattico Spermentale Delle Venezie Viale dell’Universita 10 35020 Legnaro Italy [email protected]

MÜLLER-MOLENAR, Stefan Tierartpraxis MMT Leopoldstr. 116 06366 Köthen Germany [email protected]

NAGARAJA, Kakambi College of Veterinary Medicine University of Minnesota 1971 Commonwealth Avenue Saint Paul, MN 55108 USA [email protected]

NAYLOR, Clive J. University of Liverpool Department of Veterinary Pathology Leahurst, Neston South Wirral CH64 7TE United Kingdom [email protected]

NEUMANN, Ulrich Clinic for Poultry University of Veterinary Medicine Hannover Bünteweg 17 30559 Hannover Germany [email protected]

PESENTE, Patrizia Agricola Tre Valli-Laboratorio Via S. Antonio, 60 37036 S. Martino B.A., Verona Italy [email protected]

PENZES, Zoltan Ceva-Phylaxia Veterinary Biologicals Co. Ltd. Szallas u. 5 H-1107 Budapest Hungary [email protected]

RICCHIZZI, Enrico Dept. Pub. Vet. Health & Animal Pathology Fac. Vet.Med, University of Bologna Via Tolara Di Sopra, 50 40064 Ozzano Emilia Italy [email protected]

RAU, Henriette Paul-Ehrlich-Institut Paul-Ehrlich-Str. 51-59 63225 Langen Germany [email protected]

RAUTENSCHLEIN, Silke Clinic for Poultry University of Veterinary Medicine Hannover Bünteweg 17 30559 Hannover Germany [email protected]

REBESKI, Dierk Lohmann Animal Health Heinz-Lohmann-Str. 4 27472 Cuxhaven GERMANY [email protected]

SAVAGE, Carol University of Liverpool Department of Veterinary Pathology Leahurst, Neston South Wirral CH64 7TE United Kingdom [email protected]
















396

SOOS, Pal Ceva-Phylaxia Veterinary Biologicals Co. Ltd. Szallas u. 5 H-1107 Budapest Hungary [email protected]

SPIES, Sigrid Intervet Deutschland GmbH Feldstr. 1a 85716 Unterschleißheim Germany [email protected]

THACHIL, Anil College of Veterinary Medicine University of Minnesota 1971 Commonwealth Avenue Saint Paul, MN 55108 USA [email protected]

TORO, Haroldo Department of Pathobiology College of Vet. Med., Auburn University 264 Greene Hall Auburn, AL 36849-5519 USA [email protected]

VAN DIJK, Pieter Matthijs Kerkstraat 6 5863 AP Blitterswijck The Netherlands [email protected]

VAN SANTEN, Vicky Department of Pathobiology College of Vet. Med., Auburn University 264 Greene Hall Auburn, AL 36849-5519 USA [email protected]

VERVELDE, Lonneke Faculty of Veterinary Medicine Dept. Infectious Diseases & Immunology Yalelaan 1 3584 CL Utrecht The Netherlands [email protected]

VILLARREAL, Laura Intervet/Schering Plough Animal Health Av. Sir Henry Wellcome, 335 Moinho Velho-Cotia-SP 06714-050 Brazil [email protected]

VOSS, Matthias Lohmann Tierzucht GmbH Veterinärlabor Postfach 460 D-27454 Cuxhaven Germany [email protected]

WEI, Ping Institute for Poultry Science & Health Guangxi University 13 Xiuling Road, Nanning Guangxi 530005 China [email protected]

WEBER, Rita Lohmann Animal Health Vaccine Technical Marketing Heinz-Lohmann-Str. 4 27472 Cuxhaven Germany [email protected]

WIECKERT, Frank Merial GmbH Am Söldnermos 6 D-85399 Halbergmoos Germany [email protected]

WIJMENGA, Willem Fort Dodge Animal Health Benelux BV P.O. Box 14 6290 AA Vaals The Netherlands [email protected]

WU, Ching Ching Dept. of Vet. Pathobiology Purdue University 406 South University Street West Lafayette, IN 47907-1175 USA [email protected]
















397

YAZDANI, Iraj Social welfare Organization of Maragheh Pharmacy Saadi, P.O. Box 55138-77331 Maragheh, East Azarbaijan Iran [email protected]

YU, Qingzhong USDA/ARS Southeast Poultry Research Laboratory 934 College Station Road Athens GA 30605 USA [email protected]

ZANELLA, Antonio University of Milano Instituto Microbiologia Veterinaria Via Gioberti, 19 I-25128 Brescia Italy [email protected]

ZOLNAI, Anna Ceva-Phylaxia Veterinary Biologicals Co. Ltd. Szallas u. 5 H-1107 Budapest Hungary [email protected]






398

AUTHOR INDEX A Ababneh ........................................250 Abd El Rahman .............................100 Abu-Median ...................................125 Aliabad ............................................78 Allée ......................................209, 355 Alroussan.........................................40 Amelot ...........................................355 Ariaans ..........................................142 Armesto ...................................96, 198 Assayag.................................138, 255 Astolphi..........................................245 B Barlič-Maganja.................................13 Barta..............................................225 Bertin .............................................209 Bertran.............................................27 Block................................................38 Bonci .............................................269 Boynton .........................................220 Brandão...................................47, 255 Britton ..............................96, 125, 198 Brown ....................260, 269, 294, 299 C Cao................................................219 Capua..............................................34 Cardoso.........................235, 245, 338 Casais ...........................................198 Catelli ............ 260, 278, 294, 304, 334 Cattoli ..............................................34 Cavanagh ................................96, 198 Cecchinato............. 260, 278, 304, 334 Chacón ..................................138, 251 Chen HW.........................................67 Chen QY..........................................59 Chen YN........................................250 Clubbe ...................................278, 285

Cook ..............................................326 Cowley ...........................................109 D da Silva ..........................................338 de Quadros ....................................114 de Wit.....................142, 158, 176, 188 Devlin.............................................130 Dolz .................................................27 Drago...............................................34 E Easton............................................307 Edworthy........................................307 Emmott ..........................................108 Estevez ..........................................315 Eterradossi.............209, 251, 307, 355 F Fabri ..............................................188 Falchieri .........................................294 Fasolato ...........................................34 Ferrari ............................235, 245, 338 Ferreira ..........................................338 G Gallardo ...........................................88 Galludec.........................................285 Ganapathy .....................................183 Gomaa...........................................225 Gomes ...................................235, 245 Goyal .............................................251 Gray...............................................265 Guedes ..........................................245


399

H Hafez .............................................114 Halvorson ..............................265, 363 Handberg...........................................7 Hein ...............................................385 Herrler ...........................................100 Hewson .........................................130 Hilt. ................................................220 Hirata.............................................235 Hiscox.......................................87,108 Hsieh .............................................250 Huang..............................................67 Hughes ..............................................2 I Ignjatovic .......................................130 Ivo..................................................138 J Jackwood.......................................220 Jansen...........................................142 Joiner...............................................92 Jones......... 2, 109, 183, 269, 285, 387 Jørgensen..........................................7 Jungbäck .......................................118 K Kabell ................................................7 Kaiser ............................................183 Kapczynski ....................................315 Katz ...............................................193 Khalesi....................................151, 167 Khawaldeh.......................................40 Kissinger........................................220 Krapež .............................................13 Krispel ...........................................193 Kuana ..............................................47

L Lamande........................................209 Le Bras ..........................................355 Le Gros..........................................158 Le Men...........................................209 Le-Gros..........................................376 Lemiere..........................................183 Li ......................................................59 Lierz...............................................371 Lin..........................................219, 250 Lupini .....................260, 278, 304, 334 Lüschow.........................................114 Luvizotto ........................235, 245, 338 M Macdonald .......................................87 Maharat..........................................193 Majdani ............................................78 Majó.................................................27 Malo...............................................255 Marandi............................................78 Mardani....................................78, 130 Masoudei ...............................151, 167 Maurel............................................209 McCall............................................220 McCrory ...........................................87 McKinley ........................................220 Meir................................................193 Mo....................................................59 Moghaddampour....................151, 167 Monne..............................................34 Morshedi ..........................................78 Morvan...........................................209 Motitschke......................................118 N Nagaraja ................................265, 363 Naylor ....................260, 269, 278, 285 ...............................294, 299, 304, 334 Ndegwa............................................92 Neumann .......................................100 Noormohammadi ...........................130 Novais............................................245


400

O Ojkic. .............................................225 Olesen ...............................................7 P Paterson ........................................220 Patnayak........................................251 Pedroso. ................................138, 251 Piantino .........................................251 Picault............................................209 Prandini .........................................158 Q Queguiner......................................209 R Rautenschlein................................344 Ravillion.........................................209 Retaux ...........................................209 Revolledo.......................................138 Ricchizzi ................260, 269, 278, 334 Richtzenhain............................47, 255 Rothwell.........................................183 Rubbenstroth .................................344 S Sandri ......................................47, 255 Savage ......................................2, 109 Shaw .............................................363 Silva-Frade ....................................245 Slavec..............................................13 Souza ..............................................47 T Teixeira..................................235, 338 Thachil ...........................................363 Toquin ...................209, 251, 307, 355

Toro ...........................................88, 92 Totanji ..............................................40 Turblin............................................209 V van de Haar ...................................142 van de Sande.........................158, 176 van der Meulen ..............................188 van Ginkel........................................92 van Haarlem ..................................142 van Santen.................................88, 92 Vejarano ........................................138 Velayudhan............................265, 363 Vervelde.........................................142 Vicente...........................................235 Villarreal...................................47, 255 W Wang CH .........................................67 Wang XY..........................................59 Wei P ...............................................59 Wei ZJ..............................................59 Winter ............................................100 Worthington .......................................2 Wu .........................................219, 250 Y Yoo ................................................225 Yu ..................................................315 Z Zorman Rojs ....................................13 Zsak...............................................315 Zwingelstein...................................307

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