STUDIES ON INFECTIOUS BURSAL DISEASE VIRUS (IBDV) dsRNA

EXTRACTED FROM FORMALIN FIXED PARAFFIN EMBEDDED TISSUE

by

MOHAMED MAMDOUH HAMOUD

(Under the Direction of Pedro Villegas and Barry Harmon)

ABSTRACT

In first study, sequencing the hypervariable region of IBDV from formalin fixed paraffin

embedded tissues (FFPET) that originated from chickens experiencing immunosuppression;

located both in the USA and abroad, allowed accurate strain identification of IBDV. This allows

direct correlation between viral identity and tissue lesions. Several new emerging viruses that

don’t group with other known IBDV along with a unique variant virus that had 63 nucleotides

missing from its hypervariable region were identified.

The second project investigated why some positive acute +4 lesions of FFPET yielded no

RT-PCR detectable IBDV RNA. It was hypothesized that different formalin fixation conditions

have negative impact on RNA detection from FFPET. To study this, bursas with high viral loads

and maximum histological IBDV lesion score of 4 were fixed in formalin under various

conditions. Only tissues fixed in formalin with a pH of 7.0, 5 or 10% formaldehyde, storage

temperature of 25°C or less, and kept up to 2 weeks in formalin yielded detectable IBDV RNA

upon extraction. No RNA could be detected from tissues fixed under extreme temperature, pH or

formalin concentrations. Optimal fixation conditions for IHC detection of IBDV were in 10%

formalin, pH 7.0 and 4°C, where maximum intensity of immunostaining was observed.

The third project’s goal was to produce a subunit vaccine against IBDV, without

manipulation of live or killed viral particles, to protect chicken against IBDV infection. The

Edgar strain of IBDV VP2 gene was obtained from frozen bursas, while the hypervariable region

of VP2 was obtained from FFPET. The two genes were cloned into Pichia pastoris and protein

production was confirmed by western blot analysis. Specific pathogen free 1-day old chicks

vaccinated with recombinant VP2 showed best protection against 3 week-old challenge with

Edgar IBDV. There was no morbidity or mortality in the VP2 vaccinated birds compared to 90%

morbidity and 50% mortality in placebo vaccinated birds. Recombinant vaccines don’t totally

protect against viral replication in bursa as confirmed by high histopathological lesion scores and

immunohistochemical detection of IBDV in infected tissue. This was probably due to the high

viral dose and virulence of the challenge virus used.

INDEX WORDS: Infectious bursal disease, Chicken, VP2, formalin fixed paraffin embedded

tissue, immunohistochemistry, Pichia pastoris, recombinant vaccine

production, phylogenetic analysis

STUDIES ON INFECTIOUS BURSAL DISEASE VIRUS (IBDV) dsRNA

EXTRACTED FROM FORMALIN FIXED PARAFFIN EMBEDDED TISSUE.

by

MOHAMED MAMDOUH HAMOUD

B.V.Sc Faculty of Veterinary Medicine, Cairo University, Egypt 1994

M.V.Sc Faculty of Veterinary Medicine, Cairo University, Egypt 1998

A Dissertation Submitted to the Graduate Faculty of The University of Georgia in Partial

Fulfillment of the Requirements for the Degree

DOCTOR OF PHILOSOPHY

ATHENS, GEORGIA

2006

© 2006

Mohamed Mamdouh Hamoud

All Rights Reserved

STUDIES ON INFECTIOUS BURSAL DISEASE VIRUS (IBDV) dsRNA

EXTRACTED FROM FORMALIN FIXED PARAFFIN EMBEDDED TISSUE.

by

MOHAMED MAMDOUH HAMOUD

Major Professor: Pedro Villegas Barry Harmon

Committee: Mark Jackwood

Zhen Fu Bruce LeRoy

Electronic Version Approved: Maureen Grasso Dean of the Graduate School The University of Georgia August 2006

iv

DEDICATION

I would like to dedicate this work to my parents, my wife Rasha Tawfik, my

two wonderful daughters Salma and Yasmin for their unconditional love and

support. I also would like to dedicate this work to my twin brother and two sisters

for helping me be the person I am. It is their love and support that has made this

work possible. They have put up with me and my pursuit for this doctorate for so

long. I hope this dedication can be a simple thank you for all what they have gone

through.

v

ACKNOWLEDGEMENTS

I would like to thank my professor Dr. Pedro Villegas for his support and guidance as a

mentor. I’ve really enjoyed working in his lab and have learned so much along the way. His

humanity and experience have gone a long way with me and has helped me through this. I would

also like to thank Dr. Barry Harmon, for his support and guidance as my co-major advisor. I

really appreciate your help. To my committee members, Dr. Mark Jackwood, Dr. Zhen Fu and

Dr. Bruce LeRoy; I really can not thank them enough. It was always helpful running to you to

seek advice when experiments did not want to work out.

I would like to thank Dr. Thomas P. Brown for believing in me and helping me come to

the Poultry Diagnostic Research Center, he gave me the chance to learn from an excellent

institution specializing in the area of avian diseases. Dr. Mary Pantin-Jackwood, Dr. Ivan

Alvarado, Dr. Francisco Perozo and Mrs. Linda Purvis thank you so much for your help and

assistance.

I would like to acknowledge the good people of PDRC both faculty and staff for their

contributions in me education and research.

vi

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS.............................................................................................................v

LIST OF TABLES....................................................................................................................... viii

LIST OF FIGURES .........................................................................................................................x

CHAPTER

1 INTRODUCTION .........................................................................................................1

Purpose of this study .................................................................................................1

2 LITERATURE REVIEW ..............................................................................................5

Part 1: Infectious Bursal Disease..............................................................................5

Part 2: IBDV Pathogenesis and Immunosuppression.............................................30

Part 3: Pichia pastoris ............................................................................................38

References ...............................................................................................................49

3 Identification of infectious bursal disease viruses from RNA extracted from paraffin

embedded tissue ......................................................................................................84

References ...............................................................................................................98

4 Detection of infectious bursal disease virus from formalin-fixed paraffin-embedded

tissue by immunohistochemistry and real-time reverse transcription-polymerase

chain reaction ........................................................................................................111

References .............................................................................................................125

vii

5 Production of Infectious Bursal Disease (IBD) Viral protein 2 (VP2) and the

hypervariable region of VP2 in Pichia pastoris.....................................................136

References .............................................................................................................159

6 DISCUSSIONS AND CONCLUSIONS ...................................................................178

viii

LIST OF TABLES

Page

Table 3.1: Number of IBDV strains isolated and identified from 2003-2005 according to

geographical origin of the paraffin embedded tissues.................................................107

Table 3.2: Distribution of IBDV strains in the United States of America from 2003-2005........108

Table 3.3: Distribution of IBDV strains in the world during 2003-2005 ....................................110

Table 4.1: Fixation conditions for bursal tissues from IBDV infected and control chickens......130

Table 4.2: Histopathology and IHC mean staining scores of 5 bursas of Fabricius from chickens

infected with IBDV and subjected to different fixation conditions ............................132

Table 4.3: Effect of fixation conditions on extraction of RNA from bursas of Fabricius from

chickens infected with IBDV ......................................................................................134

Table 5.1: Experimental design of immunization experiment. Different vaccines were

administered at 1-day of age with or without Freund’s adjuvant. Challenge was 3

weeks post vaccination. Experimental replicate had the following treatment groups

.....................................................................................................................................167

Table 5.2: Number of morbid and dead birds per total number in different vaccine and challenge

treatments ....................................................................................................................168

Table 5.3: Body weight and relative organ weights of 28 day old SPF chickens vaccinated or non

vaccinated against IBDV, one week post challenge....................................................169

Table 5.4: ELISA GMT at 28 days of age to IBDV following vaccination of 1-day-old SPF

chickens .......................................................................................................................170

ix

Table 5.5: Real-time RT-PCR melting curve detection, histopathological lesion scores and IHC

mean staining scores for bursas of birds vaccinated or non vaccinated at 1-day of age

and challenged with Edgar or 2512 strains of IBDV at 21 days. Sampling was

performed 1 week post challenge. Data for individual bird’s bursa ...........................172

Table 5.6: Mean lymphocyte transformation indices of vaccinated and non-vaccinated birds

challenged or not challenged with 2512 or Edgar IBDV ............................................173

x

LIST OF FIGURES

Page

Figure 3.1: Consensus nucleic acid phylogram of common IBDV vaccines and strains, in

addition to some field strains identified from paraffin embedded tissues deduced from

1000 trees by Neighbor-Joining method. Numbers at the forks indicate the number of

times (as a percentage) the groups consisting of the sequences which are to the right of

that fork occurred after the generation of 1000 trees. P-distance bar in bottom of

figure. Arrows indicate field cases identified in our laboratory which are labeled by

case number. Other isolates are identified with GeneBank accession numbers and trade

name. Samples without accession numbers were also extracted and sequenced in our

laboratory. Only 213 nucleotides were sequenced in case 087 A. ..............................105

Figure 3.2: Consensus amino acid phylogram of common IBDV vaccines and strains, in addition

to some field strains identified from paraffin embedded tissues deduced from 1000

trees by Neighbor-Joining method. Numbers at the forks indicate the number of times

(as a percentage) the groups consisting of the sequences which are to the right of that

fork occurred after the generation of 1000 trees. P-distance bar in bottom of figure.

Arrows indicate field cases identified in our laboratory which are labeled by case

number. Other isolates are identified with GeneBank accession numbers and trade

name. Samples without accession numbers were also extracted and sequenced in our

laboratory. Only 213 amino acids were deduced in isolate 087 A..............................106

xi

Figure 4.1: Photomicrograph of Bursa of Fabricius. (IHC technique specificity control for IBDV

infected and non-infected tissues). 1a: Non infected tissue stained with IHC. 1b: IBDV

infected bursa stained with IHC. All tissues were fixed in 10% buffered formalin at a

temperature of 4°C and pH 7.0. IBDV infected controls show brown positive IHC

staining (indicated by arrow) of cortical intrafollicular lymphocytes. (100X) ..........128

Figure 4.2: : RT-PCR product of amplified VP2 fragment: lane a, molecular size marker; lane b,

amplified VP2 fragment from IBDV control positive; lane c, amplified VP2 fragment

from RNA extracted from a tissue block, lane d, negative RT- PCR control (primers

with water); lane e, negative extraction control from RNA extracted from an

uninfected tissue block ...............................................................................................128

Figure 4.3: Real time RT-PCR melting curve analysis for IBDV amplicons extracted from tissues

fixed in 10% buffered formalin at 4°C and submitted to different fixation durations

(hours). Melting curve peak after 24 hours storage was 83°C, but increased to 84°C

after 48-120 hours of storage, and after 2 weeks of storage, to 86°C, indicating

changes in nucleotide sequences. RT-control is the reverse transcriptase PCR control

for a known IBDV and extraction control is a known paraffin embedded block with

extractable IBDV genetic material. .............................................................................129

Figure 5.1: RT-PCR product of amplified VP2 and pVP2: lane 1, molecular size marker; lane 2,

3, 5 and 6 amplified VP2; lane 4, negative control; lane 8 and 9, amplified pVP2 ....174

xii

Figure 5.2: Western blot analysis of recombinant proteins. Lane 1 Molecular weight marker, two

clones of yeast expressing pVP2 lanes 2 and 3, two clones of yeast expressing VP2

and negative control, yeast transformed with wild-type plasmid. The membrane was

immunoblotted with primary monoclonal antibodies against IBDV (ATCC HB9490)

.....................................................................................................................................174

Figure 5.3. Photomicrographs of bursa from SPF chickens stained with H&E that were Non-

vaccinated, non-challenged control (A); or diluent vaccinated and Edgar IBDV

challenged (B); X100 .................................................................................................175

Figure5.4: Photomicrographs of Thymus from SPF chickens stained with H&E were VP2

vaccinated and Edgar IBDV challenged (A); diluent vaccinated and Edgar IBDV

challenged (B); Non-vaccinated , non-challenged control (C). X100.........................176

Figure 5.5: Photomicrographs of bursas from SPF chickens stained with IHC that were

vaccinated and challenged with diluent (A); Vaccinated with diluent and challenged

with Edgar IBDV (B); Vaccinated with Pichia pastoris control and challenged with

Edgar ..........................................................................................................................177

1

Chapter I

INTRODUCTION

Purpose of the Study

Infectious bursal disease virus (IBDV) is the etiological agent of Gumboro disease

or infectious bursal disease (IBD). IBD is an acute highly contagious viral disease of

young chickens. This virus causes severe immunosuppresion by destroying the B-

lymphocytes precursors found within the bursa of Fabricius, followed by bursal atrophy.

The virus is ubiquitous, highly stable in the environment and has a tendency to persist in

the environment despite thorough cleaning and disinfection, thus IBDV is endemic in

most poultry producing areas of the world. IBD is one of the major economically

important diseases of poultry worldwide despite wide usage of vaccination programs.

Most commercial chickens are exposed to IBDV early in life. In unprotected flocks, the

virus causes mortality and immunosuppression. Although mortality can be quite

significant, the major economic loss is the ability of IBDV to produce

immunosuppression. Immunosuppressed flocks have poor performance which results in

reduced economic return (230)

Controlling infectious bursal disease (IBD) and its associated immune suppression

is critical to the broiler industry. This control is achieved by either vaccinating breeder

hens with conventional live attenuated and/or inactivated IBD vaccines, or by the use of

live IBD vaccines in broiler chicks, layers or young pullets to provide active protection

against IBDV, or a combination of both strategies may be used.

There is significant antigenic, immunogenic, and pathogenic variation between

IBDV strains which determines disease outcome. Some IBDV strains cause an

2

immunosuppressive, subclinical form of disease with less than 5% mortality, while others

can cause a clinical form with up to 100% mortality such as very virulent strains. The

clinical signs and the degree of immunosuppression can also vary significantly.

Vaccination is the primary means for control; and thus most efforts for protection against

IBDV by the commercial poultry industry are focused on developing efficient

vaccination programs. Successful immunization requires reliable IBDV field and vaccine

strain characterization.

The first project was designed to detect of IBDV RNA from formalin fixed

paraffin embedded tissues from field cases that showed signs of IBDV or

immunosuppression. Characterization of extracted RNA allows direct correlation

between viral identity and lesions present in infected tissues. Molecular identification of

the hypervariable region of VP2 allows for strain identification which when aligned with

sequences of vaccine strains will be a tool that can assist veterinarians in choosing which

vaccines to include in their control programs against IBDV. This technology has the

ability to detect newly emerging viruses that have a unique genomic sequence. For

example, one strain of IBDV missing 21 amino acids from the hypervariable region was

able to replicate and produce very severe lesions. Detection of unique viruses that differ

from available vaccine strains can lead to use of these strains for vaccine development in

the future.

Our second goal was to explore optimal RNA extraction from formalin fixed

paraffin embedded tissue as well as refine detection of IBDV by use of

immunohistochemistry. We sought to investigate the effect of various fixing and storage

conditions on RNA extraction and real time RT-PCR identification of IBDV from

3

paraffin embedded tissues as well as the immunohistochemistry (IHC) detection of IBDV

in the infected tissues. Bursa of Fabricius from experimentally exposed chickens with

histopathological lesion scores of +3 or more were always positive for extracted RNA

and identification of IBDV with real time RT-PCR. However, this was not always the

case for bursal tissues that originated from field submissions. Some samples that had

acute +4 lesion scores histologically were negative for IBDV with real time RT-PCR.

This finding occurred more often in samples submitted from outside the United States

than those originating from the United States. The cause of the discordance between

severe histologic lesions and negative RT-PCR results is unknown. We hypothesized that

differences in sample collection and storage methods might be partially responsible. We

assumed that samples are fixed in 10 % buffered formalin and kept at room temperature,

but pH analysis, and microscopic artifacts suggests otherwise as some of the samples

were fixed in 37% formalin with pH 2.4 or were subjected to high temperatures. We

experimentally simulated these field conditions in terms of storage time, storage

temperature, pH of unbuffered formalin, and different concentrations of buffered

formalin to test their effect on RNA extraction from paraffin embedded tissue from a

known positive sample.

Out third aim was to produce a recombinant protein or polypeptide from IBDV

genomic material extracted from formalin fixed paraffin embedded tissue (FFPET)

without actual manipulation of live or killed virus particles. This will allow us to use

archival FFPET as a valuable source of genomic material for production of proteins or

actual viruses using reverse genetics. Our focus is on protein production and not reverse

genetic reproduction of viruses. This method may allow us to produce an efficient subunit

4

vaccine to IBDV in a system that would enable mass production for vaccination of

chickens against IBDV.

We are hopeful our work may lay the foundation for improving current techniques

developed for rapid characterization of IBDV field isolates and determining their

antigenic properties. Production of a subunit vaccine that provides effective protection

against IBDV, will allow differentiation between vaccinated birds and field challenged

birds which may provide another tool to controlling this disease by the commercial

poultry industry.

5

Chapter II

Literature Review

Part 1: Infectious Bursal Disease

Infectious bursal disease virus (IBDV) is the etiological agent of Gumboro disease

or infectious bursal disease (IBD). IBD is an acute and highly contagious viral disease of

young chickens. This virus causes severe immunosuppresion by destroying the B-

lymphocytes precursors found within the bursa of Fabricius, followed by bursal atrophy.

The IBD virus is ubiquitous, highly stable and may persist in the environment despite

thorough cleaning and disinfection, thus IBDV is endemic in most poultry producing

areas of the world. There are two serotypes of IBDV: 1 and 2. All viruses capable of

causing disease in chickens belong to serotype 1, whereas serotype 2 viruses are non-

pathogenic for both chickens and turkeys (112, 157). Chickens are the only avian species

known to be susceptible to clinical disease and lesions produced by IBDV. Turkeys,

ducks and ostriches are susceptible to infection with IBDV but are resistant to its clinical

manifestations (148, 158). IBDV has also been isolated from African black-footed and

Macaroni penguins and have been serologically identified as serotype 2 IBDV (71), and

further confirmed as serotype 2 by molecular identification (115).

IBD is one of the major economically important diseases of poultry worldwide

despite wide usage of vaccination programs. Most commercial chickens get exposed to

IBDV early in life. In unprotected flocks, the virus causes mortality and

immunosuppression. Although mortality can be quite significant, the major economic

6

concern is the ability of IBDV to produce immunosuppression. Immunosuppressed flocks

have poor performance and show reduced economic return (211).

1.1 History:

Infectious bursal disease (IBD) was first reported by Cosgrove in 1957. It was

initially recognized as “avian nephrosis”, and the syndrome became known as “Gumboro

disease” because the first outbreaks occurred in the town of Gumboro, Delaware, USA.

The clinical picture of the syndrome includes white or watery diarrhea, followed by

anorexia, depression, trembling, severe prostration, and death. At necropsy, gross

pathology findings commonly include dehydration, leg and thigh muscles hemorrhages ,

urate deposits in kidneys and enlargement of the bursa of Fabricius (37).

Initially avian nephrosis or Gumboro disease was thought to be caused by the

Gray strain of infectious bronchitis virus (IBV) because of gross changes in the kidney.

This misconception arose because the IBV and IBDV infections were concurrent in many

cases and IBDV was difficult to isolate with the available methods at the time of

discovery (135). In later studies, Winterfield et al (256), succeeded in isolating the

causative agent in embryonating eggs, and later Hitchner proposed the term “infectious

bursal disease” for the disease (85).

In 1972, it was reported that IBDV infections at an early age were

immunosuppressive (1). The recognition of this immunosuppressive capability of IBDV

greatly increased the interest in the control of this disease. The existence of serotype 2

IBDV was reported in 1980 (156).

The Delmarva Peninsula broiler growing area experienced a significant increase

in mortality and higher percentage of condemnations in 1984 and 1985. The clinical

7

syndrome had significant variability, but often was respiratory in nature. Lesions ranged

from moderate to severe, with death usually being attributed to E. coli infection (38).

Rosenberger et al. isolated four isolates designated as A, D, G, and E using vaccinated

sentinel birds (205). These isolates differed from standard strains in that they produced a

very rapid bursal atrophy associated with minimal inflammatory response. The available

killed standard vaccines did not provide complete protection against these four new

Delaware isolates. The Delaware isolates, A, D, G and E were designated as antigenic

variants and killed vaccines were developed, tested and proven effective against them

(205). Currently these and other similar variants are widely distributed in the United

States (220, 221). Snyder et al. first described variant viruses as newly emergent viruses

due to a major antigenic shift within serotype 1 (221, 223). The terminology given to

these newly emergent viruses was “IBDV variants” as they were the result of a major

antigenic shift within serotype 1 (224, 221), while the older serotype 1 viruses discovered

prior to these newly emergent viruses were called standard or classical strains of IBDV.

Acute IBDV outbreaks exhibiting 30% to 60% mortality in broiler and pullet

flocks, respectively, have been commonly reported in Europe since 1987. The first

reports were made by Chettle et al. (32), and van den Berg et al. (243). Some of these

acute outbreaks occurred in broiler flocks where appropriate hygienic and prophylactic

measures had been taken. Although no antigenic drift was detected, these strains of

increased virulence were identified as very virulent IBDV (vvIBDV) strains (242, 243).

The European situation has been dominated for a decade by the emergence of vvIBDV

strains. These strains have now spread all over the world (57). In the Americas, acute

IBD outbreaks due to vvIBDV strains have already been reported in Brazil (46, 93), and

8

the Dominican Republic (5). To date the terminology given to serotype 1 IBDV is

standard or classic, variant, and very virulent IBDV.

1.2 Etiology:

The Birnaviridae family includes three genera: Genus Aquabirnavirus (type

species: infectious pancreatic necrosis virus or IPNV), Genus Avibirnavirus (type

species: Infectious Bursal Disease Virus or IBDV), and Genus Entomobirnavirus (type

species: Drosophilla X virus or DXV) (49). IBDV is a small, non-enveloped virus whose

genome consists of two segmented dsRNA segments (121). These two segments were

designated A and B. Other birnaviruses have been isolated from bivalve mollusks such as

Tellina virus (235), and Oyster Virus (49, 128), and Japanese eels(138). To date, there

has been no report of Birnavirus capable of causing disease in mammals, although dogs

have been found to be carriers for vvIBDV (233).

The virion has a single capsid shell of icosahedral symmetry composed of 32

capsomeres and a diameter of 60 to 70 nm (49, 73, 81, 176). By cryomicroscopy, it was

shown that the IBDV capsid is an icosahedron with a T = 13 lattice composed of trimeric

subunits. The outer face of the particle is composed of 260 trimeric VP2 clusters. Closely

apposed to the inside of this protein layer are 200 Y-shaped trimeric VP3 structures (19,

29).

1.3 Viral genome structure and replication:

The genome of IBDV is formed by two segments of double-stranded RNA

(dsRNA) with the two segments detected by polyacrylamide gel electrophoresis (49,

103). The molecular weight of the double stranded segment A is 2.2 x 106 Da with a

length of approximately 3.2 kb, while the molecular weight of the double stranded

9

segment B is 1.9 x106 Da with a length of approximately 2.8 kb (166, 92).

The larger segment A contains two partially overlapping open reading frames.

The first encodes a nonstructural polypeptide of 17 kDa known as VP5, which is not

essential for replication in vitro but important for virus-induced pathogenicity (171. 172).

The second ORF encodes a 110 kDa polyprotein (NH3-VPX-VP4-VP3-COOH) that is

autoproteolytically cleaved into three polypeptides, VPX or pVP2 (48 kDa), VP3 (32

kDa) and VP4 (28 kDa). pVP2 is further processed to produce a polypeptide known as

VP2 (38 kDa) (4, 91, 119, 163). VPX, VP2, and VP3 are the major structural proteins

that form the virus capsid (19), while VP4 appears to be responsible for the proteolytic

maturation of the polyprotein (116, 119, 139).

Segment B encodes VP1, a 95-kDa protein which is the RNA-dependent RNA

polymerase responsible for the replication of the genome and synthesis of mRNAs (48,

226). VP1 shares a number of primary sequence features with RNA polymerases from

diverse origins (22).

There are direct terminal and inverted repeats at the 5’ and 3’ ends in both genome

segments of IBDV that are likely to contain important signals for replication,

transcription and packaging. It is not known whether virulence variations are due to

mutations in these regions (173). The inverted adjacent repeats at the 3’ terminus on

segments A and 5’ terminus on segment B have the potential to form stem and loop

secondary structures (120), which are involved in the processes of RNA replication,

translation and encapsidation like other RNA viruses such as poliovirus (215).

The synthesis mechanism of both virus-specific ssRNA and dsRNA during

infection with IBDV has not been clearly determined. An RNA-dependent RNA

10

polymerase has been demonstrated in IBDV (225). Genome-linked proteins have been

demonstrated in three different Birnaviruses, (165, 192, 202), indicating that they

replicate their nucleic acid by a strand displacement (semi conservative) mechanism (14,

161,226).

1.4 Viral Proteins:

Four mature viral structural proteins designated VP1, VP2, VP3 and VP4 are

detected in infected cells (9, 47, 176). A non-structural protein designated VP5 has been

identified, the function of this protein is under investigation, and was found to be non-

essential for viral replication in vitro (171, 172). Lombardo et al. reported that VP5 plays

an important role in in vivo virus egression and pathogenesis (146).

During the processing of the polyprotein precursor NH3-pVP2-VP4-VP3-COOH

into pVP2, VP3 and VP4, the existence of two sites, essential for the cleavage of the

VPX-VP4 and VP4-VP3 precursors respectively, has been reported (208). These

sequences are highly conserved among IBDV strains from both serotypes 1 and 2.

VP1, the RNA-dependent RNA polymerase of the virus, is present in small

amounts in the virion, both as a free polypeptide and covalently linked to the two double-

stranded RNA genomic segments as a genome-linked protein (VPg) ( 17, 119, 164). It

plays a key role in the encapsidation of the viral particles (147). Boots et al. reported that

enhanced virulence of vvIBDV is partly determined by its B-segment which encodes VP1

(17), particularly due to the presence of eight conserved amino acid differences (16).

VP2 is the most abundant among IBD viral proteins, accounting for 51% of the

virus proteins of the serotype 1 IBDVs. This protein is the major component of the viral

capsid, and is the host-protective antigen; with most of the neutralizing epitopes

11

occurring in the hypervariable region (amino acids 224–314) of VP2 (17) i.e. VP2

contains the antigenic region responsible for the induction of neutralizing antibodies and

for serotype specificity (60). Rescue of recombinant IBDVs from chimeric A-segments

revealed that amino acids (at positions 253 and 284) within the hypervariable region are

responsible for the cell culture adaptation (18, 20, 140, 169, 247). The transition from the

precursor of VP2 (pVP2) to VP2 involves the cleavage of pVP2 near its C terminus (4).

VP2 has also been identified as an inducer of apoptosis (63).

VP3 which is a structural protein also, accounts for 40% of the virion proteins

(121). VP3 is found only on the inner surfaces of virus-like particles (152). This protein

plays a role in the assembly of viral particles, and packaging of the viral genome (33,

147, 153, 228). Chevalier et al. found out that what actually controls capsid assembly of

Infectious Bursal Disease Virus is the last C-terminal residue of VP3, glutamic acid 257

(33). VP3 is a group-specific antigen that is recognized by non-neutralizing antibodies,

some of which cross-react with both serotypes 1 and 2 (11). It is likely that the outer

subunits in the viral capsid consists of VP2, carrying the dominant neutralizing epitope,

and that the inner trimers consist of protein VP3, as it was shown that the IBDV capsid is

an icosahedron with a T = 13 lattice composed of trimeric subunits. The outer face of the

particle is composed of 260 trimeric VP2 clusters. Closely apposed to the inside of this

protein layer are 200 Y-shaped trimeric VP3 structures (19, 29).

VP4 is the viral protease involved in the processing of the precursor polyprotein

(4). It is a proteolytic enzyme-like protein, which uses a Ser-Lys catalytic dyad to act on

specific substrates and cleavage sites (15). The integrity of VP4 is essential for the

12

proteolytic processing of the polyprotein (51, 116) and either itself, or through proteins

under its control, plays a role in the trans-activation of the synthesis of VP1 (15).

VP5 is a class II membrane protein with a cytoplasmic N-terminus and

extracellular c-terminus domain (146). VP5 was the last IBDV protein identified. This

protein is not essential for IBDV replication in vitro or in vivo (171), however, it plays an

important role in viral pathogenesis as a reverse genetics system with VP5 knockout

mutant did not cause bursal lesions after experimental infections (264). VP5 expression

results in the alteration of cell morphology, the disruption of plasma membrane and

drastic reduction of cell viability. VP5 acts as a death protein, modifying cell membrane

and promoting the release of newly formed IBD virions (146).

1.5 Natural and Experimental Hosts:

For many years, the chicken (Gallus gallus) was considered the only species in

which natural infections occurred (76), where all breeds of chicken are affected. White

Leghorns exhibit the most severe reactions and have the highest mortality rate. Turkeys

(Meleagris gallopavo) may be infected with serotypes 1 and 2 but do not exhibit clinical

signs of the disease (113, 158). There is, however, considerable potential for

immunosuppression or interaction with other diseases under commercial conditions in

turkeys (134). Serotype 2 was originally identified in clinically unaffected adult turkeys

in Ireland (156). Currently chickens and turkeys are considered the natural hosts of the

virus (148). Ducks (Cairina moschata) may develop IBDV infection and antibodies are

detectable by serum virus neutralization, but neither gross nor microscopic lesions have

been observed. Antibodies have been detected in wild birds like weavers (Ploceus

cucullatus), rooks (Corvus frugilegus) and finches (Uraeginthus bengalus) (27, 175) as

13

well as in Antarctic adelaide penguins, but the source of IBDV exposure has not been

defined (65). The IBDV in African black-footed and Macaroni penguins has been

serologically identified as serotype 2 IBDV (71), and was confirmed molecularly as

serotype 2 IBDV by using reverse transcriptase polymerase chain reaction (RT-PCR) and

sequencing (115). Guinea fowl (Numida meleagris) inoculated with the virus did not

develop lesions or antibodies (184, 186). A serotype 1 virus was isolated from two 8-

week-old ostrich chicks (Struthio camelus) that had lymphocyte depletion in the bursa of

Fabricius, spleen, and/or thymus (148). van den Berg experimentally inoculated

pheasants (Phesanus colchicus), partridges, quails, and guinea fowl with vvIBDV and

reported no clinical signs or lesions in these species (240).

1.6 Transmission:

IBDV is highly contagious and the disease may be spread by direct contact

between infected and susceptible flocks. Infected chickens shed IBDV one day after

infection and can transmit the disease for at least 14 days. There are neither experimental

data nor naturally-occurring observations to suggest that IBDV is transmitted vertically

by the transovarian route (148).

Indirect transmission of virus most probably occurs on fomites (feed, clothing and

litter) or through airborne dissemination of virus-laden feathers and poultry house dust

(12); . IBDV is very persistent in the environment of a poultry house. Houses from which

infected birds were removed, were found to harbor infective virus 54 and 122 days later

(12). The lesser mealworms, Alphitobius diaperinus may be reservoir hosts (155, 219).

IBDV has also been isolated from Aedes vexans mosquitoes (89), and antibodies against

IBDV have been detected in rats found on poultry farms (185). No further evidence

14

supports the conclusion that either mosquitoes or rats act as vectors or reservoirs of the

virus. Dogs have been found to be carriers for vvIBDV (233).

1.7 Resistance to chemical and physical agents: Without proper disinfection the

virus can survive on the premises for more than 4 months. The virus is extremely

resistant to disinfection and pH changes between pH 2-12. The virus remains viable

following incubation for 5 hours at 56 °C and 30 min at 60 °C. The infectivity can be

reduced by treatment with 0.5% formalin for 6 hours or 1% formalin for 1 hour (13).

1.8 Clinical forms of IBDV:

The classical form, as described since the early 1960s, is caused by the classic

moderately virulent strains of IBDV. The incubation period of IBD ranges from 2 to 4

days post exposure. One of the earliest signs of the classical infection in a flock is the

tendency for some birds to pick at their own vents. The disease also produces acute onset

of depression, reluctance to move, ruffled feathers, white or watery diarrhea, soiled vent

feathers with urates, trembling, closed eyes and prostration. The feed intake is depressed

but water consumption may be elevated. Severely affected birds become dehydrated and

in terminal stages of the disease have subnormal temperature, after which the bird dies

(37).

The variant form, described initially in the United States, is caused by variant

strains, such as the Delaware variants or GLS strains, which partially resist neutralization

by antibodies against the so-called “classic” or standard strains (220).

The acute and very virulent form, described initially in Europe, and then spread to

Asia, Africa and some countries in Latin America, is caused by hypervirulent strains of

IBDV, and it is characterized by an acute progressive clinical disease, leading to high

15

mortality rates on affected farms. The initial outbreaks in Europe were characterized by

high morbidity usually approaching 100% and mortality reaching 20-30% in broilers and

60% in pullets over a 7-day period (32), (179). Several isolates of vvIBDV caused

mortality rates of 90-100% in 4-week old susceptible leghorn chickens (243)

1.9 Gross lesions:

Chickens which die from IBD infection as a primary cause show dehydration of

the subcutaneous fascia and musculature of the thigh, inguinal and pectoral areas (37,

148). Age-related coagulation disorders coincide with the mortality and lesion severity.

Insignificant changes in birds infected with field IBDV isolates at 17 days were observed

compared to severe effects on birds infected at 42 days (228). Petechial hemorrhages

occur occasionally in the mucosa of the intermediate zone between the proventriculus and

the gizzard, as well as on leg, thigh and breast muscles. There is increased mucus in the

intestine. Kidneys show enlargement and pallor with accumulation of crystalline urate in

tubules (37). Renal lesions were more prominent in early outbreaks in the United States,

perhaps due to co-infection with nephropathogenic strains of avian infectious bronchitis

(Gray strain) (148).

The bursa of Fabricius is the primary target organ following exposure to IBDV

(34). When Cheville studied the bursal weights for 12 days postinfection, he noticed that

by the 2nd or 3rd day the bursa has a gelatinous yellow transudate covering the serosal

surface giving the bursa a creamy color instead of the normal white color. The bursa also

had prominent longitudinal striations on the serosal surface. By the 4th day after the

infection there was a doubling in size and weight of the bursa due to edema and

hyperemia. By the 5th day the bursa returns to normal weight, but it continues to atrophy,

16

and from the 8th day forward it is approximately one-third its original weight (148).

Variant strains isolates have been reported that do not induce an acute inflammatory

response (205, 213). Although one variant strain (IN) was reportedly able to induce such

acute inflammatory response (74). The variant strain’s disease is primarily subclinical

resulting in immunosuppression. Field challenge is likely a result of mild or variant

strains of IBDV, such as Delaware variant E (107, 224).

Spleenomegaly has been documented, with small gray foci uniformly dispersed

through the parenchyma (148, 162). The vvIBDV strains cause greater decrease in thymic

weight index and more severe lesions in cecal tonsils, thymus, spleens, and bone marrow

when compared to moderately pathogenic strains of IBD virus, but the bursal lesions are

similar (148).

1.10 Histopathological lesions:

Histopathological lesions of IBD occur primarily in the lymphoid structures (i.e., cloacal

bursa, spleen, thymus, harderian gland, and cecal tonsil). Infection with standard or

variant serotype 1 IBDV strains can result in death of bursal B lymphocytes.

Degeneration and necrosis of lymphocytes in the medullas of bursal follicles can be

detected within one day of infection, and were soon replaced by heterophils, lymphocytes

associated with karyorrhexis and pyknosis, and hyperplastic stromal histiocytes. By the

third day an inflammatory response with edema, heterophil infiltration, congestion and

hemorrhage were present in infections due to standard strains. At this time the follicles

may be reduced to a necrotic center surrounded by heterophils. From the fourth day after

infection, the acute inflammatory reaction declines, and as necrotic debris were cleared

by phagocytosis, cystic cavities develop in the medullary areas of follicles; necrosis and

17

phagocytosis of heterophils and plasma cells occur; and there may be fibroplasia in

interfollicular connective tissue and the covering epithelium becomes infolded and

irregular (6, 34, 199). Sharma et al (1989) observed that the infection with the variant A

strain did not result in an acute inflammatory response, and follicular lymphoid necrosis

was evident at three days after infection (213). The lack of inflammation associated with

variant infections may be due to the variant viruses killing B-lymphocytes by apoptosis

instead of necrosis (248).

The development of lesions by IBDV in thymus depends on the pathotype of the

virus (94, 231). IBDV-induced cortical thymic lymphocyte depletion is caused by

apoptosis (94). The highly pathogenic vvIBDV strains from Europe and Japan are

associated with severe thymic lymphocyte loss when compared to less pathogenic strains

(231). Although the thymus undergoes marked atrophy and extensive apoptosis of thymic

cells during the acute phase of virus infection, there is no evidence that the virus actually

replicates in T cells (230). Gross and microscopic lesions in the thymus are quickly

resolved and the thymus returns to its normal state within a few days of virus infection

(211).

The spleen may have hyperplasia of stromal histiocytes around the adenoid sheath

arteries in early stages of the infection, and lymphoid necrosis in the germinal follicles

and the periarteriolar lymphoid sheath by the third day (148). The Harderian gland may

also be affected. Normally, this gland is infiltrated and populated with plasma cells as the

chicken ages. Infection with IBDV prevents this infiltration (227). In cecal tonsils, there

may be acute heterophil inflammation, destruction of lymphocytes, and regeneration on

the fifth day after infection (76).

18

Histological lesions in the kidney are nonspecific and probably occur because of

severe dehydration of affected chickens. Lesions observed consisted of large casts of

homogeneous material infiltrated with heterophils, and also glomerular hypercellularity

(76). The liver may have slight perivascular infiltration of monocytes (193).

The vvIBDV strains can cause severe lesions in the cecal tonsils, thymus, and

bone marrow (149). Bursal lesions caused by very virulent strains are even more severe,

and are characterized by cortical lymphocyte necrosis, depletion of medullary

lymphocytes with interstitial inflammation and hyperplasia of epithelial and reticular

cells. The thymus, spleen, liver, and bone marrow are also affected (79). Other very

virulent strains cause severe lysis of heterophil myelocytes with pyknotic nuclei (94).

Thymic cortical lymphocyte necrosis and depletion of lymphocytes are more severe when

caused by vvIBDV strains, than lesions caused by classical virulent strains (95).

1.11 Ultrastructural lesions in the bursa of Fabricius:

Naqi and Millar (174)followed the sequential changes in the surface epithelium of

the bursa of Fabricius of IBDV-infected chicks by scanning electron microscopy. Forty

eight hours post inoculation they observed a reduction in number and size of microvilli

on epithelial cells. There was gradual loss of the button follicles normally seen at the

surface, and by 72 hours, most had involuted. By 96 hours, there were numerous erosions

of the epithelial surface. The surface was intact by day 9 postinoculation, but follicles

were involuted, leaving deep pits.

19

1.12 Diagnosis of IBDV:

1.12.1 Differential diagnosis:

Diagnosis of the clinical forms of IBD is based on typical signs of the disease and

on the histopathological lesions of the bursa of Fabricius. Differential diagnosis should

include velogenic viscerotropic Newcastle disease, chicken infectious anemia, vitamin K

deficiency and mycotoxicosis. In subclinical and immunosuppressive forms of IBD,

Marek’s disease, chicken anemia and mycotoxicosis should be considered (134, 148).

Due to the immunosuppressive nature of IBDV, it may serve as a predisposing factor to

other conditions, such as gangrenous dermatitis, hemorrhagic aplastic anemia syndrome,

inclusion body hepatitis, and respiratory disease. Involvement of IBDV in disease

complexes of these types is usually diagnosed retrospectively by demonstrating persistent

bursal lesions or serologic evidence of previous IBDV infection (206).

1.12.2 Histopathological diagnosis:

Diagnosis by histopathology of the bursa is frequently used, since the lesions

caused by IBDV infection is well characterized in the bursa (34, 199). This approach has

the advantage of giving valuable information about the virulence of the IBDV strain

involved and the possible time when the infection occurred. Virulence of IBDV strain is

measured by a lesion severity score, where a lesion score of 1 represented no lesions, 2

represented mild reduction in overall follicle size, 3 represented moderate reduction in

size of follicles, and 4 represented either necrosis or follicle atrophy. Time of infection is

calculated based on type of inflammatory cells present or type of inflammatory response

seen in tissue.

20

1.12.3 Virus identification:

For virus identification, the agar gel precipitin (AGP) test can be used to detect

IBDV group-specific antigen. The virus-neutralization (VN) test could be used to identify

the virus in embryos or in tissue culture after adaptation to these host systems (206). An

antigen-capture enzyme-linked immunosorbent assay (AC-ELISA) has been described

for detecting and characterizing IBDV isolates (223). Polyclonal antibodies can also be

utilized in the AC-ELISA and may be more effective for general screening of tissue

samples for IBDV (148). Direct and indirect immunofluorescent assays as well as

immunocytochemistry have been shown to be highly reliable for detection of viral

antigens in infected tissues. (206).

Strains of IBDV can be differentiated on the basis of pathotype and antigenic

configuration. There are at least two distinct serotypes and several antigenic subtypes that

can be identified by cross-neutralization and cross challenge tests. Isolates of IBDV may

vary in pathogenicity from being nonpathogenic to virulent (108,122, 148).

1.12.4 Serological diagnosis:

Serologically the AGP test can be used to detect and quantitate antibodies in

convalescent birds. To quantitate the antibodies, usually two fold serum dilutions are

made to obtain an endpoint. The antigen is prepared from bursal homogenates of

experimentally infected birds collected 3-6 days postinfection (206). The virus

neutralization (VN) test is also used for antibody quantitation. The test is routinely

performed in microtiter systems using cell-culture-adapted virus and chicken embryo

fibroblasts. Neutralization titers are expressed as the reciprocal of the highest dilution of

serum that prevents cytopathic effect. The VN test is more sensitive than the AGP test

21

and accordingly should be utilized when antibody titers are low or when quantitation of

antibody is important (206). But for the past decade enzyme-linked immunosorbent assay

(ELISA) has been widely used because it is a sensitive and rapid method. ELISA make it

easy to handle large number of samples and only requires small quantities of serum.

ELISA results do not always correlate with VN test findings. Using serological

techniques it is possible to detect the immunologic response in an outbreak or evaluate

vaccination programs (180).

1.12.5 Molecular diagnosis:

Molecular identification of IBDV has been used to diagnose IBDV. The reverse

transcription-polymerase chain reaction (RT-PCR) allows for the detection of viral RNA

from infected clinical samples (106, 137, 262). Using the RFLP procedure and BstN1 and

Mbo1 restriction enzymes (RE) IBDV were classified into 6 molecular groups (103, 105,

110). IBDV strains that do not match any of the 6 defined molecular groups have been

identified and in the USA 32 new molecular patterns have been found (111). Ikuta et al.

using 6 REs (DraI, SacI, StyI, TaqI and MvaI) classified IBDV into 20 molecular groups.

RT-PCR amplicons were also identified by sequencing (5, 7). Other molecular techniques

include the use of RNA probes (109, 114). Using real-time RT/PCR Sequence homology

or mutations were detected using real-time RT/PCR and four mutation probes. The

melting temperature (Tm) of the mutation probes is an indicator of sequence homology

(102). An in situ RT-PCR was developed to investigate early stages of infection in the

IBDV-infected BF (265). Extraction of fragmented viral genomic RNA from formalin-

fixed paraffin-embedded tissues allowed subsequent sequencing, classification, and when

22

correlated with lesions present in those tissues histologically, allowed identification of

candidate vaccine viruses (188).

1.13 Immunity:

Infectious bursal disease virus is highly infectious and very resistant to

inactivation. Therefore, despite strict hygienic measures, vaccination is unavoidable

under high infection pressure and it is necessary to protect chickens against infection

during the first weeks after hatch. The immunization of breeder flocks is especially

important to confer passive immunity to their progeny (148). Antibodies transmitted from

the hen via the yolk of the egg can protect chicks against early infections with IBDV,

thus protecting against the immunosuppressive effect of the virus (148). Therefore, it is

important to induce high titers of maternally derived antibodies that persist over the

whole laying period in laying breeders, to do so breeder hens are vaccinated two times

once with a live vaccine and once with inactivated oil-emulsified vaccines or vaccinated

twice with inactivated oil-emulsified vaccines (86). After hatching, chickens are

immunized with live vaccines, and because maternal immunity interferes with

vaccination with live vaccines, the major problem with active immunization of young

maternally immune chicks is determining the proper time of vaccination. This

determination is aided by monitoring antibody levels in a breeder flock or its progeny

(242) or by formulas like the one developed by de Wit to determine best time of

vaccination(45). The titers may vary considerably within a flock and revaccinations may

be necessary. It has also to be taken into consideration that vvIBDV will break through

immunity provided by highly attenuated vaccine strains

23

1.14 IBDV Vaccines:

1.14.1 Live vaccines:

It is well known that less attenuated strains may cause lesions in the bursa follicles

and cause immunosuppression even in vaccinated birds. To avoid live vaccine side

effects, a number of alternative vaccines have been produced. Examples include, a

subunit VP2 vaccine produced in yeast and an immune complex vaccine, composed of

the live vaccine virus complexed in vitro with antibodies (167). Satisfactory protection

against IBDV can be achieved by immunization with live or inactivated vaccines.

Classical live vaccines achieve lifelong and broad protection, but posses residual

pathogenicity and a proportional risk of reversion to virulence (241). Many choices of

live vaccines are available that differ in virulence and antigenic diversity. According to

virulence, vaccines are classified as mild, mild intermediate, intermediate, intermediate

plus, or hot. Vaccines that contain Delaware variants are also available (148). Ashraf et al

(3) studied the interactions between two antigenically similar strains of IBDV one being a

mild strain and the other being a pathogenic IBDV strain. They concluded that viral

interference occurs in live chickens, and that the most significant interference occurring

when infection with the mild and field strains occur 24 hr apart. This phenomenon might

be due to competition for host receptor sites or production of cytokine(s), and likely has

practical implications for vaccine usage and protection against IBDV (3).

1.14.2 Inactivated vaccines:

Killed vaccines in oil emulsions stimulate high levels of maternal immunity and

are extensively used in the field (148). Inactivated vaccines and live vaccines made from

variant strains protect chickens from disease caused by either variant or standard strains,

24

whereas inactivated vaccines made from standard strains do not protect, or only partially

protect, against challenge with variant strains (99). To overcome this autogenous killed

vaccines are used by some companies to better prime their breeders (218). Very virulent

strains of IBDV can be controlled adequately under experimental conditions by

vaccination with commercial vaccines prepared from classical attenuated strains (56, 187,

242).

1.14.3 In-ovo vaccines:

In ovo vaccination may provide a way for vaccines to circumvent the effects of

maternal antibody and initiate a primary immune response (64, 69). An “immune

complex” vaccine has been developed, in which the vaccine virus is complexed in vitro

with an optimum amount of antibodies (167). These virus-antibody complex vaccines

seem very promising in protecting against IBDV(72). This new technology utilizes

specific hyperimmune neutralizing antiserum with a vaccine virus under conditions that

are not sufficient to neutralize the vaccine virus but which are sufficient for delaying the

pathological effects of the vaccine alone. This allows chicks to be vaccinated more

effectively in the presence of passive immunity even with a strain that would be too

virulent for use in ovo or at hatching (72, 86, 69).

1.14.4 Molecularly engineered vaccines:

IBDV VP2, expressed in yeast (197, 259), the baculovirus system (52, 151, 196,

217, 238), recombinant NDV vaccines (90) or via transgenic Arabidopsis thaliana plants

(257) have been studied for the use as subunit vaccines. An advantage of this technology

is that a vaccine based on VP2 alone should allow monitoring of the field situation by the

discrimination between antibody induced by vaccine (anti-VP2 only) and that induced by

25

infection (anti-VP2 and VP3) (239). The use of a reverse genetics system could represent

a basis for the genetic attenuation of IBDV strains and for the generation of new

vaccines, although interference of passive immunity would still exist. Therefore,

recombinant viral vaccines expressing the VP2 protein, such as fowl pox virus (8),

herpesvirus of turkey (HVT) (44, 234), or fowl adenovirus (214) might be able to mount

an active immune response, as they are less sensitive to neutralization by anti-IBDV

maternally derived antibodies.

1.15 Antigenic variation of IBDV:

The high mutation rate of RNA viruses and the high selection pressure generated

by intensive vaccination of birds can lead to the emergence of viruses with new

properties allowing them to persist in immune populations. Historically, these mutations

have led to antigenic variation and modifications in virulence of circulating Infectious

Bursal Disease Virus (IBDV) strains. It is therefore essential to identify and characterize

new IBDV isolates as soon as they appear and compare them with previously described

viruses (239, 241). The capsid protein, VP2, is the major host protective immunogen.

Immunization of susceptible chickens with purified VP2 elicits neutralizing antibodies

and confers protection against homologous virulent virus challenge (11, 60). Three panels

of neutralizing monoclonal antibodies (mAbs) have been used in antigen capture enzyme-

linked immunosorbent assays (AC-ELISA) for antigenic characterization of serotype 1

isolates of IBDV (61, 62, 54). Neutralizing mAbs have been shown to bind to VP2 within

a restricted region, called the variable domain, between amino acids 206 and 350, which

is highly hydrophobic but has short hydrophilic regions at each end (7). Since this epitope

was denaturated by SDS, it was determined that is a conformationally-dependent epitope

26

(4). Antigenic epitopes on VP3 protein have also been reported but these antibodies are

not completely neutralizing (4, 59). Antigenic diversity between IBDV serotypes has

been recognized when serotypes 1 and 2 were defined on the basis of their lack of in vitro

cross neutralization (156). Based on studies with monoclonal antibodies, IBDV strains

belonging to serotypes 1 and 2 have been found to not share major neutralizing epitopes

(9, 209). Polyvalent neutralizing antiserotype 1 monoclonal antibodies such as

monoclonal antibodies 1, 6, 7, 8 and 9 (54), monoclonal antibody 8 (222), and

monoclonal antibodies 6F6 and 7C9 (244) have been developed.

Antigenic differences have been shown within serotype 1, and the study of

different strains has led to dividing serotype 1 into six subtypes, differentiation based on

cross neutralization assays using polyclonal sera (108). Studies with monoclonal

antibodies demonstrated the presence of a number of modified neutralizing epitopes

among antigenically variant strains detected in the United States. Based on this evidence,

there may have been an antigenic shift in IBDV viruses in the US (224). There are a

minimum of at least five neutralization epitopes on the standard IBDV strains (e.g. D78

strain) as defined by the monoclonal antibodies 8, 179, B69, R63, and 10. Delaware

viruses have lost the B69 site, GLS viruses lack the B69 and R63 sites but has a

monoclonal 57 reactive site, the DS326 virus lacked the sites for monoclonal antibodies

B69, R63 and 179, the RS593 had the 179, 8 and 67 site only, while the AL2 variant

strain had 8, 57, 67 and 179 sites (222, 223). Thus based on the reactiveness with various

monoclonal antibodies, the IBDV viruses are antigenically grouped as classic or standard,

GLS Variant, RS593 variant, AL2 variant, Y2K Variant DS326 and Delaware type

variants (132, 237)

27

Despite their enhanced pathogenic properties, the vvIBDV strains were

considered to be closely antigenically related to the standard strains such as the Faragher

52/70 strain, based on high cross-neutralization indices (56). Using neutralizing

monoclonal antibodies developed by Snyder to characterize US IBDV variants, van der

Marel studied twelve European isolates of IBDV. He detected no important differences

between the standard strain 52/70 and vvIBDV (246). Similar data was produced by

Oppling et al (187). However, Eterradossi et al. developed nine other monoclonal

antibodies and using these he detected modified binding and neutralizing properties

against French vvIBDV strains (54). All their monoclonal antibodies neutralized most

mild or intermediate vaccines strains, whereas two monoclonal antibodies did not

neutralize a French vvIBDV strain, US variant A, and the European strain Faragher

52/70. Based on their results, they suggested a neutralizing epitope may be altered in the

European vvIBDV strains, causing decreased antibody neutralization. This difference

could be used to differentiate vvIBDV strains (54).

1.16 Molecular basis of IBDV variability:

Sequencing of the VP2 gene of numerous different IBDV strains and escape

mutants confirmed that a hypervariable domain is responsible for antigenic variation

(187, 209, 237) and comparison of this region offers the best option for IBDV typing.

Therefore, the nucleic acid sequencing of genes coding for VP2 and subsequent

deduction of their predicted amino acid sequences is now commonly used for phylogeny.

The amino acid changes between strains are not evenly distributed throughout the open

reading frame but are clustered in certain regions. Most of the changes that occur in VP2

are located between amino acids 239 and 332 (7, 133, 237) whereas van den Berg et al.

28

said this hypervariable region is between amino acids 206 and 350 (245). This highly

variable region falls entirely within those sequences of VP2 identified as the minimum

region required for reaction with virus neutralization monoclonal antibody 5 (60, 261).

Hydrophilicity profiling of this region showed that there are two hydrophilic

peaks at either ends of this region, the larger peak (Major peak A) from amino acids 212

to 224 and the other from amino acids 314 to 324 (Major peak B). These hydrophilic

regions have been shown to be important in binding of neutralizing antibodies and,

hence, are presumed to be a main part of the neutralizing domain (75, 209). It is

interesting that most of amino acid variations in this region fall within these two peaks (7,

133).

Variations in IBDV antigenicity depend on changes in hydrophilic peaks. The

serotype 2 strain 23/82 (209), the North-American antigenic variants A, E, GLS and

DS326 (75, 133, 237), and neutralization resistant escape mutants (209) all exhibit amino

acid changes in these hydrophilic peaks. Only differences in the intervening hydrophobic

domains are found between typical serotype 1 strains (237). A nucleotide sequence

comparison suggested that four amino acid alterations in the VP2 protein of the Delaware

F strain allowed this variant to escape neutralizing antibodies. These amino acids were

located at positions 213, 222, 318, and 323 (75). By restriction enzyme and amino acid

sequence analysis, point mutations have been detected at residues 222, 254 and 323.

Amino acid residues 222 and 254 are consistently mutated in the variant strains (50, 105).

Glycine is present in the standard strains, amino acid residue number 254, whereas the

variants have serine at this position (50, 104).

29

Mundt (2005) showed by reverse genetics systems for IBDV, that amino acid

changes in these hydrophilic peaks was accompanied by loss or acquirement of

monoclonal antibody recognition sites (170). Vakharia et al. used monoclonal antibodies

to correlate antigenic variations with amino acid sequence substitutions in the

hypervariable region of VP2 (237). They found that the amino acid residue glutamine at

position 249 might be involved in the binding of neutralizing antibody B69, which

recognizes epitopes in standard strains. All the variant viruses have lysine instead of

glutamine at this position, and they escape binding with antibody B69. Using a

baculovirus expression system to synthesize all the structural proteins coded for in

segment A of the IBDV genome, Vakharia et al., produced virus like particles (236).

They mapped the antigenic sites by producing chimeric cDNA clones of IBDV using the

variant GLS plasmid as a backbone and inserting fragments from the D78 and Delaware

strains. At least two antigenic reactive sites are present on the surface of IBDV, one falls

between amino acid residues 222 and 249, and the other between 269 and 323.

In highly virulent strains three specific amino acid residues in VP2 have been

reported at position 222 (Ala), 256 (Ile), 294 (Ile) and 299 serine which differ from

classical strains (21). These substitutions are also present in other strains isolated from

other countries such as Bangladesh (98), Brazil (46, 93), China (28), Germany (266),

Israel (195), Japan (142), (262), Malaysia (36, 88), Nigeria (266), Taiwan (144), and

Vietnam (232). Positions 222-223 and 318-324 may be critical for the vvIBDV (55).

These three positions have been identified as “hot spots” for mutations in several escape

mutants resistant to selected neutralizing monoclonal antibodies (209, 244).

The role of VPl in the virulence of IBDV was unknown for a long time, only

30

recently did Liu and Vakharia (2004) by the use of reassortant viruses recovered by use

of reverse genetics system discover the role VP1 plays in the virulence of IBDV in vivo.

They showed virulence was due to the effect of VP1 on viral replication kinetics in vivo

(145). The VP1 sequences of very virulent IBDV strains are genetically distinct from

those of classical virulent or attenuated strains thus, VP1 of vvIBDV constitutes a genetic

lineage distinct from that of classical virulent or attenuated strains and serotype 2 strains

as well (97).

Part 2: IBDV Pathogenesis and Immunosuppression:

2.1 IBDV Pathogenesis:

IBD is characterized by lesions in lymphoid organs, and in chicks older than 3

weeks is characterized by morbidity and in some cases death (213, 251). Chickens that

recover are severely immunosuppressed (1, 25, 84) due to the loss of B cells, especially

the developing B-cell population in the bursa of Fabricius. Only serotype 1 viruses cause

IBD, and pathotypes have been classified in increasing order of virulence as mild,

intermediate, variant virulent (in North America), classical virulent, very virulent (vv) or

hyper-virulent strains (241).

The mature bursa of Fabricius is the main target organ of IBDV as it is the source

for B lymphocytes in avian species. Bursectomized chickens did not develop clinical IBD

despite the presence of infection (79). The severity of the disease is directly related to the

number of susceptible cells present in the bursa of Fabricius; therefore, the highest age

31

susceptibility is between 3 and 6 weeks, when the bursa of Fabricius is at its maximum

development. This age susceptibility is broader in the case of the vvIBDV strains (99,

100, 179).

After oral infection or inhalation, the virus replicates primarily in the lymphocytes

and macrophages of the gut-associated lymphoid tissues. From the gut, the virus is

transported to other tissues by phagocytic cells, most likely resident macrophages (211,

239). By 13 hours post-inoculation (h.p.i.), most bursal follicles are positive for virus and

by 16 h.p.i. a second more pronounced viraemia occurs with secondary replication in

other organs leading to disease and death (168). It is suggested that vvIBDV causes

similar disease signs to those of classical virulent strains with the same incubation time of

4 days but with an exacerbated acute phase (241, 255).

Actively dividing, surface immunoglobulin M-bearing B-cells are lysed by

infection (80, 83, 203), but cells of the monocyte-macrophage lineage can be infected in a

persistent and productive manner, and play a crucial role in dissemination of the virus

(24,96) and in the onset of the disease (127, 131, 212). The exact cause of clinical disease

and death is still unclear but does not seem to be related only to the severity of the lesions

and the bursal damage. Prostration preceding death is very similar to what is observed in

acute coccidiosis, and is reminiscent of a septic shock syndrome (241). The macrophage

could play a specific role in this pathology by exacerbated release of cytokines such as

tumor necrosis factor or interleukin 6 (127). Macrophages are known to be activated by

interferon gamma; increased secretion of interferon as has been demonstrated both in

vitro in chicken embryo cultures and in vivo in chickens after their infection with IBDV

(66, 67).

32

In addition to causing necrosis in the lymphoid cells of the bursa, IBDV also

induces apoptosis (63, 130, 249). Apoptosis is characterized by cell shrinkage and

chromatin condensation and does not generate a local inflammatory response. Induction

of apoptosis in infected cells contributes to the pathogenesis of IBDV in the bursa (118,

183), chicken peripheral blood lymphocytes (248), and in the thymus (94, 230). Virally-

induced apoptosis can occur in cells in the absence of detectable virus (118, 178, 230). A

direct effect of viral proteins like VP2 and VP5 has been implicated in the induction of

apoptosis (63, 263). Apoptotic cells have also been observed in viral antigen-negative

bursal cells, underlining the possible role of immunological mediators in this process

(178, 230). And finally, apoptosis has also been observed in the proventriculus of IBDV

challenged SPF leghorn chickens (189).

2.2 IBDV Immunosuppression:

Most immunosuppression studies have been restricted to vaccine or classical

virulent strains of IBDV (201). Clinical and subclinical infections with IBDV may cause

suppression of both humoral and cellular immune responses (211). The first indication of

damage in the immune system was reported by Helmboldt and Gardner in 1964 (76). In

1970, Cho demonstrated that white leghorn chickens exposed to IBDV at one day of age

were consistently more likely to develop visceral tumors and nerve enlargement by

Marek’s disease virus (35). In 1972 it was reported that IBDV infection at an early age

was immunosuppressive, and severely depressed the antibody response to Newcastle

disease virus (1). IBDV replication in the bursa leads to extensive lymphoid cell

destruction in the follicular medullas and cortices (229). The acute lytic phase of the virus

is associated with a reduction in circulating IgM+ cells (83, 203). IBDV-exposed

33

chickens produce suboptimal levels of antibodies against a number of infectious and

noninfectious antigens (62, 126, 260).

2.2.1 Humoral response to IBDV infection:

Only the primary antibody response is impaired, the secondary responses remain

intact (70, 203,213), and this humoral deficiency may be reversible (211). Although

destruction of Ig-producing B cells may be one of the principal causes of humoral

deficiency, other mechanisms are possible including the adverse effect of IBDV on

antigen-presenting and helper T cell functions (213). A paradox associated with IBDV

infections in chickens is that although there is immunosuppression against many antigens,

the response against IBDV itself is normal, even in 1-day-old susceptible chickens (216).

There appears to be a selective stimulation of the proliferation of B cells committed to

anti-IBDV antibody production (148).

2.2.2 Cellular immune response to IBDV infection:

T-cells are resistant to infection with IBDV (82, 62), and there is no evidence that

the virus actually replicates in thymic lymphocytes (213, 230). However, there is

evidence that in vitro mitogenic proliferation from T cells of IBDV exposed birds is

severely decreased. This mitogenic inhibition is likely mediated by macrophages,

however, how IBDV induces macrophages to exhibit this suppressor effect is not clear

(211). It has been suggested that T cells are induced during an IBDV infection in the

bursa, but phenotypic characterization is still required and (124, 125). Rodenberg et al.

studied changes in the T-cell subsets within the blood, bursa, spleen and thymus after

infection with classical virulent IBDV and suggested that, although the number of

immunoglobulin (Ig)M+ cells in the bursa and spleen decreased significantly, the relative

34

proportions of CD4+ and CD8+ T cells did not change (203). Immunohistochemical

analysis of chicken lymphoid tissues to determine the location of the virus and various

leukocyte subsets showed an influx of T cells into the bursa (177, 251). Bursal CD4+ and

CD8+ αβ1-TCR+ cells increased (251), and some expressed the activation marker CD25

(the interleukin-2 receptor-a) (124). It still remains a matter of contention whether these

cells are a migratory or resident bursal population.

The number of CD4+ and CD8+ T cells increased after infection and were first

detected at the corticomedullary boundary of the bursa, probably reflecting their route of

entry (251). The cortico-medullary boundary is rich in arterioles and venules, which

allow migratory lymphocytes entry into the follicles in response to an appropriate chemo-

attractant(s). Although CD4+ T cells had increased by 7 d.p.i., their location was

restricted to the cortico-medullary boundary and the cortex, even though most IBDV+

cells were present in the medulla. Later, CD8+ T cells were detected throughout the

follicles. The presence of numerous T cells suggests that cell-mediated immunity has a

very important role to play in recovery from an infection with vvIBDV. Poonia and

Charan (198) reported consistently higher titers of IBDV in chickens lacking a fully

functional T-cell-mediated immune system. This increase in bursa T cells appears to be a

consequence of migration and not an artifact due to the loss of B cells or the result of an

expansion of resident T cells (251).

Sharma et al. (211) also detected a dramatic infiltration of T-cells in the bursa

during acute IBDV infection, accompanied by the abrupt drop in the number of IgM+

cells. By the seventh day of infection, the infiltrating cells were predominantly CD8+

lymphocytes. It was suggested that T-cells modulate the infection, limiting viral

35

replication in the bursa in the early phase of the disease. They also promote bursal tissue

damage and delay recovery, possibly through the release of cytokines and cytotoxic

effects (135). Cytotoxic T cells may exacerbate virus-induced cellular destruction by

lysing cells expressing viral antigens. T cells may also promote the production of pro-

inflammatory factors, such as nitric oxide, increasing tissue destruction (211).

2.2.3 Innate immunity

The effect of IBDV on innate immunity is centered in the modulatory effect of

IBDV on macrophage functions. There is evidence that the in vitro phagocytic activity of

these cells is compromised (211).

2.3 Pathogenesis of vvIBDV:

As for vvIBDV, VP2 antigen was most prevalent in the bursa, followed by the

spleen and then the thymus, as reported for classical virus strains (231, 251). However,

the distribution of VP2 antigen within the bursa was different from that with less virulent

IBDV. VP2 antigen was first detected at the cortico-medullary boundary, which probably

corresponds to the site of viral entry. Migratory cells

(e.g. macrophages) probably carry vvIBDV to the bursa from the site of primary

replication (intestine), and once in the medulla, it is able to colonize individual follicles

(101, 255). It is widely accepted that IBDV primarily targets immature B cells located in

the bursal cortex. However, vvIBDV VP2 antigen was mostly present in the medulla, a

site where more mature B cells are found (190, 255); only later was UK661 strain of

IBDV disseminated into the cortex (255).

Rapid depletion of virus from the bursa is associated with distinctive spherical

bodies accumulating in the medulla that resembled highly phagocytosing, activated

36

macrophages (252, 154). Macrophages appear to play a major role in the clearance of

dead (apoptotic and necrotic) cells and cell debris. A large accumulation of cells

expressing the macrophage marker KUL-01 and major histocompatability complex

(MHC) class II invariant chain was present around 3 to 4 d.p.i. (255). The corresponding

decrease in viral load could partly be related to the action of phagocytosing macrophages,

although also to the exhaustion of target cells. Depletion of medullary Bu-1+, IgM+ and

IgG+ cells, prior to their depletion from the cortex, and depletion of IgM+ cells before

IgG+ cells, is consistent with the suggestion that more immature B cells are preferred

targets, although it is important to note that both populations became depleted. This will

have profound consequences both for the immune responses against vvIBDV itself and

the consequent immunosuppression after viral clearance. The extended range of target

lymphocytes could partly explain the increased virulence of vvIBDV isolates (255).

After destruction of the bursal architecture by vvIBDV, Bu-1+ B cells begin to

return to the depleted bursal follicles by 14 d.p.i. However, very few cells with surface

IgM+ or IgG+ are detected with the monoclonal antibodies M1 and G1 (255). There is

also some bursal recovery of the IgM+ cell population at 14 d.p.i. with classical virulent

IBDV (123). The number of IgM+ cells were fewer than the Bu-1+ B cells which could be

due to that IBDV induced conformational changes in the Ig molecules resulting in

alterations in those epitopes recognized by the M1 and G1 monoclonal antibodies. IBDV-

induced changes in chicken IgM giving rise to a monomeric form in the circulation of

convalescent chickens (101). It was recently reported that only B cells expressing a

surface immunoglobulin are able to colonize bursal follicles, whereas those B cells with a

non-productive V-D-J recombination lacking Ig expression are deleted (194). This

37

suggests that the Bu-1+ cells repopulating the bursa are likely to be non-functional and

will be subsequently deleted. The lack of expression of surface Ig on bursal B cells is

consistent with the convalescent chick being severely immunocompromised (101). Lack

of Ig-producing B cells would explain the poor antibody response to secondary antigenic

challenge in chickens recovering from IBDV infection. The situation after vvIBDV

infection seems likely to be far more severe than with less virulent strains ( 25, 77).

An important aspect of the pathogenesis and clinical disease caused by vvIBDV is

the involvement of other lymphoid organs (94). The extensive damage in the thymus

caused by UK661 IBDV strain was far greater than that reported for classical virulent

stains (231). At the height of thymic pathogenesis much of the cortex contained pyknotic

nuclei within large vacuolated spaces, many of which merged. This suggests either that

vvIBDV has a direct apoptotic or necrotic effect on thymocytes (118) or that cell death is

due to bystander activity (254). Lesions in the spleen were less evident and the transient

nature of detectable virus may be related to lymphocyte and macrophage populations

restricting the spread of the IBDV (255). The spleen is the major secondary lymphoid

organ in the chicken where most systemic T-cell and B-cell responses are initiated. The

B-lymphocyte population was most severely affected, suggesting that peripheral B cells

are highly susceptible to vvIBDV. This further supports the notion that immature B cells

are not the only targets for vvIBDV, as both IgM+ and IgG+ B cells decreased. The IgM+

cell population recovered in the spleen, reaching a higher number than before infection,

in association with germinal centers. This is most probably a consequence of expansion

of the peripheral B-cell pool and not due to seeding from the bursa. The IgG+ population

also recovered and was associated with germinal centers, suggesting isotype switching of

38

the IBDV-reactive IgM+ population. The lack of bursal architecture (and probably

function) and the increase in splenic IgM+ and IgG+ cells is consistent with the B-cell

compartment being severely suppressed, while at the same time IBDV-reactive B cells

are present (10, 200). The T-cell compartment within the spleen seemed to be unaffected

after infection (255).

In the thymus, IBDV antigen was only transiently detected at 4 to 7 d.p.i. The

rapid depletion of Bu-1+ medullary B cells probably follows the loss of viral target cells

and restricted viral distribution. Few, if any, Bu-1+, IgM+ or IgG+ cells were detected in

the thymic cortex even though VP2 antigen was detected there. The presence of IBDV

antigen on the surface of numerous, adjacent cortical thymocytes suggests that IBDV is

able to adhere to immature T cells. However, it seems unlikely that IBDV directly infects

and replicates within these cells as Hirai and Calnek were unable to show replication of

IBDV in chicken T cells (80). The proportion of CD4+ and CD8+ T cells in the thymus

altered little following infection.

Part 3: Pichia Pastoris

3.1 Introduction:

Geneticists have learned how to manipulate DNA, to identify, move and place

genes into a variety of organisms that are quite different from the source organism. A

major use for many of these recombinant organisms is to produce proteins. Since many

proteins are of immense commercial value, numerous studies have focused on finding

ways to produce them efficiently and in a functional form. Escherichia coli have been the

choice of researchers for several reasons. It is a single-celled organism that reproduces

39

mainly through asexual reproduction. The organism’s simplicity makes it easy and cheap

to work with. Its food source is simple, and it does not require elaborate facilities for

growth and maintenance. Its rapid growth cycle allows for a quick increase in the

population size of a particular strain, as an Escherichia coli population can double in less

than an hour. In addition to its own circular DNA, Escherichia coli have plasmids that

complement the main segment of DNA. Plasmids are easy to isolate and manipulate. The

plasmid can be removed, a desired gene can be inserted into it, and the plasmid can be

reintroduced into Escherichia coli. The Escherichia coli will then produce the foreign

protein as if the protein were native to the bacteria. In order to express protein, a

promoter from the host needs to be located immediately in front of gene to be expressed.

The promoter regulates when, how much and how often the gene is transcribed. The

simplicity of Escherichia coli makes it a desirable host for production of a foreign

protein; however, it also has its disadvantages as a prokaryote. All prokaryotes do not

have any of the membrane bound organelles found in eukaryotes. In eukaryotes a protein

is often post-translationally modified in different organelles, such as the endoplasmic

reticulum or the Golgi apparatus. Modifications often involve addition of different forms

of glycosylation. Any eukaryotic protein can be mass translated in Escherichia coli , but

many are not quite finished and hence, they are nonfunctional. Escherichia coli will give

the same primary structure, as occurs when that protein is initially produced in its own

cell type. However, the failure to modify that structure often means that the protein will

not form as it would with the presence of certain organelles. Because of the problems

encountered when using a prokaryote to produce eukaryotic products, researchers sought

suitable eukaryotic replacements like mammalian, insect and yeast cells. Of these three

40

eukaryotic expression systems, yeast cells are the most desirable, as they combine the

ease of genetic manipulation and rapid growth characteristics of a prokaryotic organism

with the subcellular machinery for performing post-translational protein modification of

eukaryotic cells (41).

The yeast Saccharomyces cerevisiae has been popular with molecular biologists,

and many proteins have been produced using it as a protein expression system. S.

cerevisiae has several limitations, as its product yields are low, often reaching a

maximum of 1-5 percent of total protein yield. Additionally, the presence of foreign gene

products puts stress on the cells. The production of the protein during the growth phase

hinders growth. Even the use of inducible plasmid promoters to achieve a partial

separation between the growth and protein production phase, has not been effective due

to the instability of the plasmid (23). Besides the difficulties encountered with scaling up

protein production to get better yields, several reports have noted the hyperglycosylation

of secreted glycoproteins which may cause differences in immunogenicity and

diminished activity. Also, many of the secreted proteins of S. cerevisiae are not found

free in the medium, but rather in the periplasmic space. This leads to problems with

purification and further decreases product yield (23).

Pichia pastoris was discovered 30 years ago when Koichi Ogata discovered its

ability to utilize methanol as a sole source of carbon and energy (181). Because methanol

could be inexpensively synthesized from natural gas (methane), there was immediate

interest in exploiting these organisms for the generation of yeast biomass or single-cell

protein (SCP) to be marketed primarily as a high protein animal feed. However, the oil

crisis of the early 1970’s increased the price of methane drastically along with drop of

41

soybean prices rendered this goal unpractical. Use of Pichia pastoris as an alternative

species to S. cerevisiae in molecular biology was appealing, as growth protocols were

similar in both species. Also because P. pastoris has a strong inducible promoter that can

be used for protein production. P. pastoris is capable of generating post-translational

modifications such as proteolytic processing, folding, disulfide bond formation, and

glycosylation that are more similar to human protein modifications than S. cerevisiae was

capable of doing (43). Thus, many proteins that end up as inactive inclusion bodies in

bacterial systems are produced as biologically active molecules in P. pastoris. The P.

pastoris system is also generally regarded as being faster, easier, and less expensive to

use than expression systems derived from higher eukaryotes, such as insect and

mammalian tissue culture cell systems, and usually gives higher expression levels (78).

The production of a functional protein is intimately related to the cellular machinery of

the organism producing the protein. The yeast Pichia pastoris is a useful system for the

expression of milligram-to-gram quantities of proteins for both basic laboratory research

and industrial manufacture. The fermentation can be readily scaled up to meet greater

demands, and parameters influencing protein productivity and activity, such as pH,

aeration and carbon source feed rate, can be controlled (78). Compared with mammalian

cells, Pichia does not require a complex growth medium or culture conditions, is

genetically relatively easy to manipulate, and has a eukaryotic protein synthesis pathway.

Because of these characteristics, some proteins such as G protein-coupled receptors that

cannot be expressed efficiently in bacteria, S. cerevisiae or the insect cell/baculovirus

system, have been successfully produced in functionally active form in P. pastoris (30,

141). Pichia can be grown to very high cell densities using minimal media (250, 253) and

42

integrated vectors help genetic stability of the recombinant elements, even in continuous

and large-scale fermentation processes (204). Simple purification and isolation of a

recombinant protein is facilitated by the fact that P. pastoris does not secrete a lot of its

own proteins (41). Therefore, the powerful genetic techniques available, together with its

economy of use, make P. pastoris a system of choice for heterologous protein expression.

So, what began more than 20 years ago as a program to convert abundant methanol to a

protein source for animal feed has developed into what are today two important

biological tools: a model eukaryote used in cell biology research, and a recombinant

protein production system. Pichia has gained widespread attention as an expression

system because of its ability to express high levels of heterologous proteins. As a result,

recombinant vector construction, methods for transformation, selectable marker

generation, and fermentation methods have been developed to exploit the productive

potential of this system (207).

Research Corporation Technologies (Tucson, AZ, USA) are the current holders of

the patent for the P. pastoris expression system, which they have held since 1993, and the

P. pastoris expression system is available in kit form from Invitrogen Corporation

(Carlsbad, CA, USA).

3.2 Methanol metabolism in Pichia

Methylotrophic yeasts are able to utilize methanol as the sole carbon and energy

source. All such strains identified to date belong to only four genera: Hansenula, Pichia,

Candida and Torulopsis (58). They all share a specific methanol utilization pathway

involving several unique enzymes. The initial reactions take place in specialized

microbodies (the peroxisomes) followed by subsequent metabolic steps in the cytoplasm.

43

Peroxisomes play an indispensable role during growth, as they harbor the three key

enzymes for methanol metabolism, viz. alcohol oxidase, catalase and dihydroxyacetone

synthase. The proliferation of peroxisomes is a reflection of environmental conditions.

When the cells are grown on glucose, very few peroxisomes are present. When grown on

methanol, peroxisomes may take up to 80 percent of the total cell volume. Previous

results clearly show that the alcohol oxidase promoter is both tightly regulated and is a

strong promoter. The production of foreign protein can be repressed until the culture is

saturated with colonies, and then the production of the foreign protein can begin with the

de-repression and induction of the gene. The subsequent reactions of methanol

assimilation and dissimilation are localized in the cytosol. The methanol metabolism is

fully described by Cereghino and Cregg (31, 5, 117), Jahic and Gellissen (68).

3.3 Methanol utilization phenotypes:

There are three phenotypes of P. pastoris host strains with regard to methanol

utilization. The Mut+, or methanol utilization plus phenotype, grow on methanol at the

wild-type rate and require high feeding rates of methanol in large-scale fermentations

(31). The Muts, or methanol utilization slow phenotype, have a disruption in the AOX1

gene. Since the cells must then rely on the weaker AOX2 for methanol metabolism, a

slower growing and slower methanol utilization strain is produced. Muts strains have

been found to be advantageous for production of hepatitis B surface antigen (40). The

Mut-, or methanol utilization minus phenotype, are unable to grow on methanol, since

these strains have both AOX genes deleted. One of the advantages of this phenotype is

that low growth rates may be desirable for production of certain recombinant products

44

(39, 40). Currently, the majority of researchers use the Mut+ phenotype (87, 217),

although some researchers are also using the Muts phenotype (2, 191).

3.4 The Pichia Expression System:

The expression of any foreign gene in P. pastoris comprises three principal steps:

(a) insertion of the gene into an expression vector; (b) introduction of the expression

vector into the P. pastoris host; and (c) examination of potential strains for the expression

of the foreign gene (31, 41, 129)

The majority of heterologous protein production in P. pastoris is based on the fact

that enzymes required for the metabolism of methanol are only present when cells are

grown on methanol (53). This has been the most successful system reported for this

organism (141). The AOX promoters have been the most widely utilized promoters;

however, other promoter options are available for the production of foreign proteins in

Pichia, (31).

3. 4.1 Promoters:

One of the drawbacks with S. cerevisiae was that it did not have a strong

inducible promoter. Pichia pastoris has a strong inducible promoter. This inducible

promoter is related to the fact that P. pastoris is a methylotrophic yeast. The first step in

the utilization of methanol is the oxidation of methanol to formaldehyde and hydrogen

peroxide (136). This step is catalyzed by the enzyme alcohol oxidase. The expression of

this gene is tightly regulated. When the yeast is grown on glucose or ethanol, alcohol

oxidase is not detectable in the cells. However, when the yeast is grown on methanol,

45

alcohol oxidase can make up to thirty-five percent of the total cellular protein. The

control of the amount of alcohol oxidase is largely transcriptional (42).

There are two alcohol oxidase genes: AOX1 and AOX2. The protein coding

regions of the genes are largely homologous, 92 percent and 97 percent at the nucleotide

and amino acid sequence levels respectively (182). The promoters share very little

homology. No mRNA of the two genes is detectable when the yeast is grown in glycerol.

The promoter region for AOX2 has a repressor region that leads to the inhibition of gene

expression, and an activation region that leads to the enhancement of gene expression.

The AOX1 gene promoter probably has a similar mechanism (182).

The AOX1 promoter has been the most widely reported and utilized of all the

available promoters for P. pastoris (141). One of the reasons that the constitutive GAP

(glyceraldehyde 3-phosphate dehydrogenase) promoter has not been widely used is the

belief that constitutive production of foreign proteins in P. pastoris may have cytotoxic

effects (160). However, recent studies have found not only that cytotoxic effects are not

necessarily observed, but also that production levels of a recombinant exo-levanase

(LsdB) using the GAP promoter were similar to those using the AOX1 promoter (160).

Combining the GAP and AOX1 promoters in a strain expressing human granulocyte-

macrophage colony-stimulating factor (hGM-CSF) resulted in a two-fold increase in

production of the recombinant protein (258). By using this combined promoter for

sequential expression of a constitutively produced enzyme required for post-translational

modification, an inducible recombinant protein can be modified to a specified form

within the fermentation process itself (26, 258).

46

The YPT1 [a small GTPase involved in secretion] (210) and PEX8 [a peroxisomal

matrix protein] (143) promoters have not been widely used, probably because of their low

expression levels, since the goal of the majority of researchers using the P. pastoris

expression system is to obtain maximum amounts of expressed foreign proteins (31).

Other promoters include the AOX2 promoter (39), FLD1 promoter (250) and the ICL1

promoter (159).

3.4.2 Selectable markers:

All P. pastoris expression strains are derived from NRRL-Y11 430 (Northern

Regional Research Laboratories, IL, USA), the wild-type strain. One or more auxotrophic

mutants are often present in these strains, allowing for selection during transformation of

strains containing the appropriate selectable marker gene. A number of selectable marker

genes are known for the molecular genetic manipulation of P. pastoris, they include HIS4

(histidinol dehydrogenase gene), ARG4 (argininosuccinate lyase gene), ZeoR (zeocin

resistance gene), Blasticidin S deaminase gene, ADE1-PR-

amidoimidazolesuccinocarboximide synthase, URA3-orotidine 5 -phosphate

decarboxylase and SorR-acetyl-CoA carboxylase (31).

The genetic manipulation of P. pastoris for the production of various

heterologous proteins is simplified by the use of a wide range of selectable markers and

promoters. Choosing the correct markers and promoters is essential for obtaining a high

productivity (150).

3.5 Post-translational modifications:

The post- translational modifications made by P. pastoris are more suitable for

use in humans. The structure of carbohydrate added to secreted proteins is known to be

47

very organism specific. Many proteins secreted from S. cerevisiae have been

demonstrated to be antigenic when introduced into mammals thus; the use of

glycoprotein products synthesized by yeast for therapeutic purposes has been avoided. A

comparison of a S. cerevisiae protein secreted from S. cerevisiae and P. pastoris has

shown distinct differences between N-linked oligosaccharide structures added to proteins

secreted from these yeast. The majority of the N-linked oligosaccharide chains are high

mannose. However, the length of the carbohydrates chains is much shorter in P. pastoris.

Even the longest chains of protein produced in P. pastoris contained only approximately

thirty mannose residues, which is significantly shorter than the 50 to 150 mannose

residue chains typically found on S. cerevisiae glycoproteins. The second major

significant difference between the glycolation by S. cerevisiae and P. pastoris is that

glycans from P. pastoris don't have alpha 1,3-linked mannose residues that are

characteristic of S. cerevisiae (41). The enzyme that makes alpha 1,3 linkages is alpha 1,3

mannosyl transferase and it is undetectable in P. pastoris. It is significant, because the

alpha 1,3 linkages on S. cerevisiae glycans are primarily responsible for the highly

antigenic nature of glycoproteins used for therapeutic products (41)

3.6 Production of heterologous proteins in P. pastoris:

The Pichia expression system has been widely used to produce a variety of

different heterologous proteins (31). P. pastoris grows on a simple mineral media and

does not secrete high amounts of endogenous protein. Therefore the heterologous protein

secreted into the culture is relatively pure and purification is easier to accomplish (58).

Cell growth is particularly important for secreted protein production in bioreactors, since

the concentration of the product in the extracellular medium is roughly proportional to

48

the concentration of cells in the culture in many instances. The yields of proteins

expressed intracellularly in bioreactors are also high, due to the efficiency of the AOX1

promoter(31). Production of large amounts of heterologous proteins in shake-flask

culture is difficult, due to the limitations of volume, oxygen transfer, substrate addition

and an inability to monitor these factors efficiently (150). The use of bioreactors is

preferable, since all of these parameters can be monitored and controlled simultaneously,

allowing more efficient production of the desired heterologous protein. Cereghino et al.

reviewed the use of bioreactors for the production of recombinant proteins from P.

pastoris (30).

Pichia pastoris as a yeast expression system was chosen because it combines the

simplicity of prokaryotic growth requirements, growth potential and ease of manipulation

along with the post-translational modifications of eukaryotes to ensure that the expressed

protein is functional and has the proper conformation.

49

References

1. Allan, W. H., J. T. Faragher, and G. A. Cullen. Immunosuppression by the

infectious bursal agent in chickens immunized against Newcastle disease. Vet

Rec: 511-512. 1972.

2. Aoki, H., M. Nazmul Ahsan, and S. Watabe. Heterologous expression in pichia

pastoris and single-step purification of a cysteine proteinase from northern shrimp.

Protein Expr Purif 31: 213-221. 2003.

3. Ashraf, S., G. Abdel-Alim, M. Q. Al-Natour, and Y. M. Saif. Interference between

mild and pathogenic strains of infectious bursal disease virus in chickens. Avian

Dis. 49: 99-103. 2005.

4. Azad, A. A., M. N. Jagadish, M. A. Brown, and P. J. Hudson. Deletion mapping

and expression in Escherichia coli of the large genomic segment of a Birnavirus.

Virology 161:145-152. 1987.

5. Banda, A., P. Villegas, J. El-Attrache, and C. Estevez. Molecular characterization

of seven field isolates of infectious bursal disease virus obtained from commercial

broiler chickens. Avian Dis 45:620-630. 2001.

6. Barnes, H. J., J. Wheeler, and D. Reed. Serologic evidence of infectious bursal

disease virus infection in Iowa turkeys. Avian Dis 26:560-565. 1982.

7. Bayliss, C. D., U. Spies, K. Shaw, R. W. Peters, A. Papageorgiou, H. Muller, and

M. E. Boursnell. A comparison of the sequences of segment A of four infectious

bursal disease virus strains and identification of a variable region in VP2. J Gen

Virol 71 (Pt 6):1303-1312. 1990.

50

8. Bayliss, C. D., R. W. Peters, J. K. Cook, R. L. Reece, K. Howes, M. M. Binns, and

M. E. Boursnell. A recombinant fowlpox virus that expresses the VP2 antigen of

infectious bursal disease virus induces protection against mortality caused by the

virus. Arch Virol 120:193-205. 1991.

9. Becht, H. Infectious bursal disease virus. Curr Top Microbiol Immunol 90:107-

121. 1980.

10. Becht, H., and H. Muller. Infectious bursal disease-B cell dependent

immunodeficiency syndrome in chickens. Behring Inst Mitt 89: 217-225. 1991.

11. Becht, H., H. Muller, and H. K. Muller. Comparative studies on structural and

antigenic properties of 2 serotypes of infectious bursal disease virus. Journal of

General Virology 69:631-640. 1988.

12. Benton, W., M. S. Cover, and J. K. Rosenberger. Physiochemical properties of the

infectious bursal agent (IBA). Avian Dis 11:438-445. 1967.

13. Benton, W., M. S. Cover, and J. K. Rosenberger. Studies on the transmission of

the infectious bursal agent (IBA) of chickens. Avian Dis:430-438. 1967.

14. Bernard, J. Drosophila X virus RNA polymerase: Tentative model for in vitro

replication of the double-stranded virion RNA. J Virol 33:717-723. 1980.

15. Birghan, C., E. Mundt, and A. Gorbalenya. A non-canonical lon proteinase lacking

the ATPase domain employs the ser-lys catalytic dyad to exercise broad control

over the life cycle of a double-stranded RNA virus. Embo J. 19:114-123. 2000.

16. Boot, H., A. ter Huurne, and B. Peeters. Generation of full-length cDNA of the

two genomic dsRNA segments of infectious bursal disease virus. Journal of

Virological Methods 84:49-58. 2000.

51

17. Boot, H. J., A. J. Hoekman, and A. L. Gielkens. The enhanced virulence of very

virulent infectious bursal disease virus is partly determined by its b-segment. Arch

Virol 150:137-144. 2005.

18. Boot, H. J., A. A. H. M. Ter Huurne, A. J. W. Hoekman, B. P. H. Peeters, and A.

L. J. Gielkens. Rescue of very virulent and mosaic infectious bursal disease virus

from cloned cDNA: VP2 is not the sole determinant of the very virulent

phenotype. J Virol. 74:6701-6711. 2000.

19. Bottcher, B., N. A. Kiselev, V. Y. Stel'Mashchuk, N. A. Perevozchikova, A. V.

Borisov, and R. A. Crowther. Three-dimensional structure of infectious bursal

disease virus determined by electron cryomicroscopy. J Virol 71:325-330. 1997.

20. Brandt, M., K. Yao, M. Liu, R. A. Heckert, and V. N. Vakharia. Molecular

determinants of virulence, cell tropism, and pathogenic phenotype of infectious

bursal disease virus. J Virol 75:11974-11982. 2001.

21. Brown, M. D., P. Green, and M. A. Skinner. VP2 sequences of recent European

very virulent' isolates of infectious bursal disease virus are closely related to each

other but are distinct from those of 'classical' strains. J Gen Virol 75 (Pt 3):675-

680. 1994.

22. Bruenn, J. A. Relationships among the positive strand and double-strand RNA

viruses as viewed through their RNA-dependent RNA polymerases. Nucleic Acids

Res 19:217-226. 1991.

23. Buckholz, R. G., and M. A. Gleeson. Yeast systems for the commercial production

of heterologous proteins. Biotechnology (N Y) 9:1067-1072. 1991.

52

24. Burkhardt, E., and H. Muller. Susceptibility of chicken blood lymphoblasts and

monocytes to infectious bursal disease virus (IBDV). Arch Virol 94:297-303.

1987.

25. Butter, C., T. D. M. Sturman, B. J. G. Baaten, and T. F. Davison. Protection from

infectious bursal disease virus (IBDV)-induced immunosuppression by

immunization with a fowlpox recombinant containing IBDV-VP2. Avian Pathol.

32:597-604. 2003.

26. Callewaert, N., W. Laroy, H. Cadirgi, S. Geysens, X. Saelens, W. Min Jou, and R.

Contreras. Use of hdel-tagged trichoderma reesei mannosyl oligosaccharide 1,2-

alpha-d-mannosidase for n-glycan engineering in pichia pastoris. FEBS Lett

503:173-178. 2001.

27. Campbell, G. Pathogenicity of investigation into evidence of exposure to

infectious bursal disease virus and infectious anaemia virus in wild birds in

Ireland. In: Proceedings II International Symposium on infectious bursal disease

and chicken infectious anaemia. Rauischholzhausen. pp 230 - 235. 2001.

28. Cao, Y. C., W. S. Yeung, M. Law, Y. Z. Bi, F. C. Leung, and B. L. Lim.

Molecular characterization of seven Chinese isolates of infectious bursal disease

virus: Classical, very virulent, and variant strains. Avian Dis 42:340-351. 1998.

29. Caston, J. R., J. L. Martinez-Torrecuadrada, A. Maraver, E. Lombardo, J. F.

Rodriguez, J. I. Casal, and J. L. Carrascosa. C terminus of infectious bursal disease

virus major capsid protein VP2 is involved in definition of the t number for capsid

assembly. J Virol 75:10815-10828. 2001.

53

30. Cereghino, G. P., J. L. Cereghino, C. Ilgen, and J. M. Cregg. Production of

recombinant proteins in fermenter cultures of the yeast pichia pastoris. Curr Opin

Biotechnol 13:329-332. 2002.

31. Cereghino, J. L., and J. M. Cregg. Heterologous protein expression in the

methylotrophic yeast pichia pastoris. FEMS Microbiol Rev 24:45-66. 2000.

32. Chettle, N., J. C. Stuart, and P. J. Wyeth. Outbreak of virulent infectious bursal

disease in East Anglia. Vet Rec 125:271-272. 1989.

33. Chevalier, C., J. Lepault, B. Da Costa, and B. Delmas. The last c-terminal residue

of VP3, glutamic acid 257, controls capsid assembly of infectious bursal disease

virus. J Virol 78:3296-3303. 2004.

34. Cheville, N. F. Studies on the pathogenesis of gumboro disease in the bursa of

fabricius, spleen and thymus of the chicken. Am J Pathol: 527-551. 1967.

35. Cho, B. R. Experimental dual infections of chickens with infectious bursal and

Marek’s disease agents. I. Preliminary observation on the effect of infectious

bursal agent on Marek’s disease. Avian Dis 14:665-675. 1970.

36. Chong, L. K., A. R. Omar, K. Yusoff, M. Hair-Bejo, and I. Aini. Nucleotide

sequence and phylogenetic analysis of a segment of a highly virulent strain of

infectious bursal disease virus. Acta Virol 45:217-226. 2001.

37. Cosgrove, A. An apparently new disease of chickens: Avian nephrosis. Avian Dis:

385-389. 1962.

38. Craig, F. R. Economic impact of respiratory disease. In: 20th National Meeting on

Poultry Health and Condemnations. Ocean City, MD. pp 35-37. 1985.

54

39. Cregg, J., and K. Madden. Development of the methylotrophic yeast, pichia

pastoris, as a host system for the production of foreign proteins. Dev Ind

Microbiol: 23-42. 1988.

40. Cregg, J. M., J. F. Tschopp, and C. Stillman. High-level expression and efficient

assembly of hepatitis b surface antigen in pichia pastoris. BioTechnology: 479-

485. 1987.

41. Cregg, J. M., T. S. Vedvick, and W. C. Raschke. Recent advances in the

expression of foreign genes in pichia pastoris. Biotechnology (N Y) 11:905-910.

1993.

42. Cregg, J. M., K. J. Barringer, A. Y. Hessler, and K. R. Madden. Pichia pastoris as

a host system for transformations. Mol Cell Biol 5:3376-3385. 1985.

43. Cregg, J. M., J. L. Cereghino, J. Shi, and D. R. Higgins. Recombinant protein

expression in pichia pastoris. Mol Biotechnol 16:23-52. 2000.

44. Darteil, R., M. Bublot, E. Laplace, J. F. Bouquet, J. C. Audonnet, and M. Riviere.

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. 1995.

45. de Jong, L. A., S. Grunewald, J. P. Franke, D. R. Uges, and R. Bischoff.

Purification and characterization of the recombinant human dopamine d2s receptor

from pichia pastoris. Protein Expr Purif 33:176-184. 2004.

46. de Wit, J. J. Gumboro disease - the optimal time for vaccination. International

Poultry Production: 19-23. 2003.

55

47. Di Fabio, J., L. I. Rossini, N. Eterradossi, M. D. Toquin, and Y. Gardin. European-

like pathogenic infectious bursal disease viruses in Brazil. Vet Rec 145:203-204.

1999.

48. Dobos, P. Peptide map comparison of the proteins of infectious bursal disease

virus. J Virol 32:1047-1050. 1979.

49. Dobos, P., and T. E. Roberts. The molecular biology of infectious pancreatic

necrosis virus: A review. Can J Microbiol 29:377-384. 1983.

50. Dobos, P., B. J. Hill, R. Hallett, D. T. Kells, H. Becht, and D. Teninges.

Biophysical and biochemical characterization of five animal viruses with

bisegmented double-stranded RNA genomes. J Virol 32:593-605. 1979.

51. Dormitorio, T. V., J. J. Giambrone, and L. W. Duck. Sequence comparisons of the

variable VP2 region of eight infectious bursal disease virus isolates. Avian Dis

41:36-44. 1997.

52. Duncan, R., E. Nagy, P. Krell, and P. Dobos. Synthesis of the infectious pancreatic

necrosis virus polyprotein, detection of a virus-encoded protease, and fine-

structure mapping of genome segment-a coding regions. J Virol. 61:3655-3664.

1987.

53. Dybing, J. K., and D. J. Jackwood. Antigenic and immunogenic properties of

baculovirus-expressed infectious bursal disease viral proteins. Avian Dis 42:80-91.

1998.

54. Egli, T., J. P. van Dijken, M. Veenuis, W. Harder, and A. Fiechter. Methanol

metabolism in yeasts: Regulation of the synthesis of catabolic enzymes. Arch

Microbiol: 115-121. 1980.

56

55. Eterradossi, N., G. Rivallan, D. Toquin, and M. Guittet. Limited antigenic

variation among recent infectious bursal disease virus isolates from France. Arch

Virol 142:2079-2087. 1997.

56. Eterradossi, N., C. Arnauld, D. Toquin, and G. Rivallan. Critical amino acid

changes in VP2 variable domain are associated with typical and atypical

antigenicity in very virulent infectious bursal disease viruses. Arch Virol

143:1627-1636. 1998.

57. Eterradossi, N., J. P. Picault, P. Drouin, M. Guittet, R. Lhospitalier, and G.

Bennejean. Pathogenicity and preliminary antigenic characterization of 6

infectious bursal disease virus-strains isolated in France from acute outbreaks.

Journal of Veterinary Medicine Series B-Zentralblatt Fur Veterinarmedizin Reihe

B-Infectious Diseases and Veterinary Public Health 39:683-691. 1992.

58. Eterradossi, N., C. Arnauld, F. Tekaia, D. Toquin, H. Le Coq, G. Rivallan, M.

Guittet, J. Domenech, T. van den Berg, and M. Skinner. Antigenic and genetic

relationships between European very virulent infectious bursal disease viruses and

an early West African isolate. Avian Pathol. 28:36-46. 1999.

59. Faber, K. N., W. Harder, G. Ab, and M. Veenhuis. Review: Methylotropic yeasts

as factories for the production of foreign proteins. Yeast:1331-1344. 1995.

60. Fahey, K. J., I. J. Odonnell, and T. J. Bagust. Antibody to the 32k structural

protein of infectious bursal disease virus neutralizes viral infectivity invitro and

confers protection on young chickens. Journal of General Virology 66:2693-2702.

1985.

57

61. Fahey, K. J., K. Erny, and J. Crooks. A conformational immunogen on vp-2 of

infectious bursal disease virus that induces virus-neutralizing antibodies that

passively protect chickens. J Gen Virol 70 (Pt 6):1473-1481. 1989.

62. Fahey, K. J., P. McWaters, M. A. Brown, K. Erny, V. J. Murphy, and D. R.

Hewish. Virus-neutralizing and passively protective monoclonal-antibodies to

infectious bursal disease virus of chickens. Avian Dis. 35:365-373. 1991.

63. Faragher, J. T., W. H. Allan, and G. A. Cullen. Immunosuppressive effect of the

infectious bursal agent in the chicken. Nat New Biol 237:118-119. 1972.

64. Faragher, J. T., W. H. Allan, and P. J. Wyeth. Immunosuppressive effect of

infectious bursal agent on vaccination against Newcastle disease. Vet Rec:385-

388. 1974.

65. FernandezArias, A., S. Martinez, and J. Rodriguez. The major antigenic protein of

infectious bursal disease virus, VP2, is an apoptotic inducer. J Virol. 71:8014-

8018. 1997.

66. Gagic, M., C. A. St Hill, and J. M. Sharma. In-ovo vaccination of specific-

pathogen-free chickens with vaccines containing multiple agents. Avian Dis

43:293-301. 1999.

67. Gardner, H., K. Kerry, M. Riddle, S. Brouwer, and L. Gleeson. Poultry virus

infection in antarctic penguins. Nature 387:245. 1997.

68. Gelb, J., C. S. Eidson, and S. H. Kleven Studies on interferon induction by

infectious bursal disease virus (IBDV).1. Interferon-production in chicken-embryo

cell-cultures infected with IBDV. Avian Dis. 23:485-492. 1979.

58

69. Gelb, J., C. S. Eidson, and S. H. Kleven. Interferon-production in embryonating

chicken eggs following inoculation with infectious bursal disease virus. Avian Dis.

23:534-538. 1979.

70. Gellissen, G. Heterologous protein production in methylotrophic yeasts. Appl

Microbiol Biotechnol 54:741-750. 2000.

71. Giambrone, J., T. Dormitorio, and T. Brown. Safety and efficacy of in ovo

administration of infectious bursal disease viral vaccines. Avian Dis. 45:144-148.

2001.

72. Giambrone, J. J., D. L. Ewert, and C. S. Eidson. Effect of infectious bursal disease

virus on immunological responsiveness of chicken. Poultry Sci. 56:1591-1594.

1977.

73. Gough, R., S. Drury, D. d. B. Welchman, J. R. Chitty, and G. E. S. Summerhays.

Isolation of Birnavirus and reovirus-like agents from penguins in the United

Kingdom. Vet Rec.151:422-424(423). 2002.

74. Haddad, E., C. Whitfill, A. Avakian, C. Ricks, P. Andrews, J. Thomas, and P.

Wakenell. Efficacy of a novel infectious bursal disease virus immune complex

vaccine in broiler chickens. Avian Dis. 41:882-889. 1997.

75. Harkness, J. W., D. J. Alexander, M. Pattison, and A. C. Scott. Infectious bursal

disease agent - morphology by negative stain electron-microscopy. Arch

Virol.48:63-73. 1975.

76. Hassan, M., M. AlNatour, L. Ward, and Y. Saif. Pathogenicity, attenuation, and

immunogenicity of infectious bursal disease virus. Avian Dis. 40:567-571. 1996.

59

77. Heine, H. G., M. Haritou, P. Failla, K. Fahey, and A. Azad Sequence analysis and

expression of the host-protective immunogen VP2 of a variant strain of infectious

bursal disease virus which can circumvent vaccination with standard type I strains.

J Gen Virol 72 (Pt 8):1835-1843. 1991.

78. Hemboldt, C. F., and E. Garner. Experimentally induced gumboro disease (IBA).

Avian Dis:561-575. 1964.

79. Higashihara, M., K. Saijo, Y. Fujisaki, and M. Matumoto. Immunosuppressive

effect of infectious bursal disease virus-strains of variable virulence for chickens.

Veterinary Microbiology 26:241-248. 1991.

80. Higgins, D., and J. Cregg. Methods in molecular biology: Pichia protocols.

Humana, Totowa, NJ. 1998.

81. Hiraga, M., T. Nunoya, Y. Otaki, M. Tajima, T. Saito, and T. Nakamura.

Pathogenesis of highly virulent infectious bursal disease virus-infection in intact

and bursectomized chickens.J Vet Med Sci. 56:1057-1063. 1994.

82. Hirai, K., and B. W. Calnek. Invitro replication of infectious bursal disease virus

in established lymphoid-cell lines and chicken lymphocytes-B. Infect Immun

25:964-970. 1979.

83. Hirai, K., Shimakur.S, and E. Kawamoto, Electron-microscope characterization of

infectious bursal disease virus. Avian Dis. 18:467-471. 1974.

84. Hirai, K., K. Kunihiro, and S. Shimakura, Characterization of immunosuppression

in chickens by infectious bursal disease virus. Avian Dis 23:950-965. 1979.

60

85. Hirai, K., T. Funakoshi, T. Nakai, and S. Shimakura, Sequential-changes in the

number of surface immunoglobulin-bearing lymphocytes-b in infectious bursal

disease virus-infected chickens. Avian Dis. 25:484-496. 1981.

86. Hirai, K., Shimakur.S, E. Kawamoto, F. Taguchi, S. T. Kim, C. N. Chang, and Y.

Iritani. Immunodepressive effect of infectious bursal disease virus in chickens.

Avian Dis. 18:50-57. 1974.

87. Hitchner, S. Infectivity of infectious bursal disease virus for embryonating eggs.

Poultry Sci. 49:511-&. 1970.

88. Hofacre, C. L. Personal communication In. 2005.

89. Hohenblum, H., B. Gasser, M. Maurer, N. Borth, and D. Mattanovich. Effects of

gene dosage, promoters, and substrates on unfolded protein stress of recombinant

Pichia pastoris. Biotechnol Bioeng 85:367-375. 2004.

90. Hoque, M., A. Omar, L. Chong, M. Hair-Bejo, and I. Aini. Pathogenicity of sspI-

positive infectious bursal disease virus and molecular characterization of the VP2

hypervariable region. Avian Pathol. 30:369-380. 2001.

91. Howie, R. I., and J. Thorsen. Identification of a strain of infectious bursal disease

virus isolated from mosquitoes. Can J Comp Med 45:315-320. 1981.

92. Huang, Z., S. Elankumaran, A. S. Yunus, and S. K. Samal. A recombinant

Newcastle disease virus (NDV) expressing VP2 protein of infectious bursal

disease virus (IBDV) protects against NDV and IBDV. J Virol 78:10054-10063.

2004.

61

93. Hudson, P. J., N. M. McKern, K. J. Fahey, and A. A. Azad. Predicted sequence of

the host-protective immunogen of infectious bursal disease virus. FEBS Lett

201:143-146. 1986.

94. Hudson, P. J., N. M. McKern, B. E. Power, and A. A. Azad. Genomic structure of

the large RNA segment of infectious bursal disease virus. Nucleic Acids Res

14:5001-5012. 1986.

95. Ikuta, N., J. El-Attrache, P. Villegas, E. M. Garcia, V. R. Lunge, A. S. Fonseca, C.

Oliveira, and E. K. Marques. Molecular characterization of Brazilian infectious

bursal disease viruses. Avian Dis 45:297-306. 2001.

96. Inoue, M., M. Fukuda, and K. Miyano. Thymic lesions in chicken infected with

infectious bursal disease virus. Avian Dis 38:839-846. 1994.

97. Inoue, M., A. Fujita, and K. Maeda. Lysis of myelocytes in chickens infected with

infectious bursal disease virus. Vet Pathol 36:146-151. 1999.

98. Inoue, M., H. Yamamoto, K. Matuo, and H. Hihara. Susceptibility of chicken

monocytic cell-lines to infectious bursal disease virus.J Vet Med Sci. 54:575-577.

1992.

99. Islam, M., K. Zierenberg, and H. Muller. The genome segment b encoding the

RNA-dependent RNA polymerase protein VP1 of very virulent infectious bursal

disease virus (IBDV) is phylogenetically distinct from that of all other IBDV

strains. Arch Virol 146:2481-2492. 2001.

62

100. Islam, M. R., K. Zierenberg, N. Eterradossi, D. Toquin, G. Rivallan, and H.

Muller. Molecular and antigenic characterization of Bangladeshi isolates of

infectious bursal disease virus demonstrate their similarities with recent European,

Asian and African very virulent strains. J Vet Med B Infect Dis Vet Public Health

48:211-221. 2001.

101. Ismail, N. M., and Y. M. Saif. Immunogenicity of infectious bursal disease viruses

in chickens. Avian Dis 35:460-469. 1991.

102. Ismail, N. M., A. M. Fadly, and T. S. Chang. Effect of bursal cell number on the

pathogenesis of infectious bursal disease in chickens. Avian Dis 31:546-555.

1987.

103. Ivanyi, J., and R. Morris Immunodeficiency in the chicken. Iv. An immunological

study of infectious bursal disease. Clin Exp Immunol.:154-165. 1976.

104. Jackwood, D. Rapid identification of the most appropriate vaccine strain to control

infectious bursal disease. http://www.oardc.ohio-

state.edu/IBDV/IBDVLightCylcer.asp In. 2005.

105. Jackwood, D., and R. Jackwood. Infectious bursal disease viruses - molecular

differentiation of antigenic subtypes among serotype-1 viruses. Avian Dis 38:531-

537. 1994.

106. Jackwood, D., and R. Jackwood. Molecular identification of infectious bursal

disease virus strains. Avian Dis. 41:97-104. 1997.

107. Jackwood, D., and S. Sommer. Restriction fragment length polymorphisms in the

VP2 gene of infectious bursal disease viruses. Avian Dis. 41:627-637. 1997.

63

108. Jackwood, D., G. Hanes, and S. Miller. Infectious bursal disease viral RNA

amplification using RT/PCR from bursa tissue following phenol: Chloroform

inactivation of the virus. Avian Dis 40:457-460. 1996.

109. Jackwood, D., R. Jackwood, and S. Sommer. Identification and comparison of

point mutations associated in classic and variant infectious bursal disease viruses.

Virus Res.49:131-137. 1997.

110. Jackwood, D. H., and Y. M. Saif. Antigenic diversity of infectious bursal disease

viruses. Avian Dis 31:766-770. 1987.

111. Jackwood, D. J. Development and characterization of nucleic-acid probes to

infectious bursal disease viruses. Veterinary Microbiology 24:253-260. 1990.

112. Jackwood, D. J., and S. E. Sommer. Genetic heterogeneity in the VP2 gene of

infectious bursal disease viruses detected in commercially reared chickens. Avian

Dis 42:321-339. 1998.

113. Jackwood, D. J., and S. E. Sommer. Virulent vaccine strains of infectious bursal

disease virus not distinguishable from wild-type viruses with marker the use of a

molecular. Avian Dis 46:1030-1032. 2002.

114. Jackwood, D. J., Y. M. Saif, and J. H. Hughes. Characteristics and serologic

studies of 2 serotypes of infectious bursal disease virus in turkeys. Avian Dis.

26:871-882. 1982.

115. Jackwood, D. J., Y. M. Saif, and P. D. Moorhead. Immunogenicity and

antigenicity of infectious bursal disease virus serotypes I and II in chickens. Avian

Dis 29:1184-1194. 1985.

64

116. Jackwood, D. J., D. E. Swayne, and R. J. Fisk. Detection of infectious bursal

disease viruses using insitu hybridization and nonradioactive probes. Avian Dis.

36:154-157. 1992.

117. Jackwood, D. J., S. E. Sommer, R. E. Gough, S. E. Drury, D. d. B. Welchman, J.

R. Chitty, and G. E. S. Summerhays. Sequence analysis of an infectious bursal

disease virus isolated from penguins in the united kingdom. In.

http://www.oardc.ohio-state.edu/IBDV/Penguin%20Page.asp 2004.

118. Jagadish, M. N., V. J. Staton, P. J. Hudson, and A. A. Azad. Birnavirus precursor

polyprotein is processed in Escherichia coli by its own virus-encoded polypeptide.

J Virol 62:1084-1087. 1988.

119. Jahic, M. Process techniques for production of recombinant proteins with pichia

pastoris. PhD thesis. In: Department of Biotechnology. Royal Institute of

Technology, S-10 691 Stockholm, Sweden. 2003.

120. Jungmann, A., H. Nieper, and H. Muller. Apoptosis is induced by infectious bursal

disease virus replication in productively infected cells as well as in antigen-

negative cells in their vicinity. J Gen Virol 82:1107-1115. 2001.

121. Kibenge, F., and V. Dhama. Evidence that virion-associated VP1 of

Avibirnaviruses contains viral RNA sequences. Arch Virol.142:1227-1236. 1997.

122. Kibenge, F., M. Nagarajan, and B. Qian. Determination of the 5' and 3' terminal

noncoding sequences of the bi-segmented genome of the Avibirnavirus infectious

bursal disease virus. Arch Virol 141:1133-1141. 1996.

123. Kibenge, F. S., A. S. Dhillon, and R. G. Russell. Biochemistry and immunology of

infectious bursal disease virus. J Gen Virol 69 (Pt 8):1757-1775. 1988.

65

124. Kibenge, F. S. B., A. S. Dhillon, and R. G. Russell. Identification of serotype-ii

infectious bursal disease virus proteins. Avian Pathol. 17:679-687. 1988.

125. Kim, I., M. Gagic, and J. Sharma. Recovery of antibody-producing ability and

lymphocyte repopulation of bursal follicles in chickens exposed to infectious

bursal disease virus. Avian Dis. 43:401-413. 1999.

126. Kim, I., S. You, H. Kim, H. Yeh, and J. Sharma. Characteristics of bursal t

lymphocytes induced by infectious bursal disease virus. J Virol. 74:8884-8892.

2000.

127. Kim, I. J., and J. M. Sharma. IBDV-induced bursal t lymphocytes inhibit

mitogenic response of normal splenocytes. Vet Immunol Immunopathol 74:47-57.

2000.

128. Kim, I. J., M. Gagic, and J. M. Sharma. Recovery of antibody-producing ability

and lymphocyte repopulation of bursal follicles in chickens exposed to infectious

bursal disease virus. Avian Dis. 43:401-413. 1999.

129. Kim, I. J., K. Karaca, T. L. Pertile, S. A. Erickson, and J. M. Sharma. Enhanced

expression of cytokine genes in spleen macrophages during acute infection with

infectious bursal disease virus in chickens. Vet Immunol Immunopathol 61:331-

341. 1998.

130. Kitamura, S., S. Jung, and S. Suzuki. Seasonal change of infective state of marine

Birnavirus in Japanese pearl oyster Pinctada fucata. Arch Virol 145:2003-2014.

2000.

66

131. Koutz, P., G. R. Davis, C. Stillman, K. Barringer, J. Cregg, and G. Thill.

Structural comparison of the pichia pastoris alcohol oxidase genes. Yeast 5:167-

177. 1989.

132. Lam, K. Morphological evidence of apoptosis in chickens infected with infectious

bursal disease virus. J Comp Pathol. 116:367-377. 1997.

133. Lam, K. Alteration of chicken heterophil and macrophage functions by the

infectious bursal disease virus. Microb Pathog. 25:147-155. 1998.

134. Lamichhane, C. M., S. A. Mengel-Whereat, J. K. Rosenberger, R. S. Hein, R. S.

Resurreccion, and Lovell. Use and interpretation of monoclonal antibodies for the

characterization of infectious bursal disease viruses. In: International Poultry

Exposition. Atlanta, GA. 2001.

135. Lana, D. P., C. E. Beisel, and R. F. Silva. Genetic mechanisms of antigenic

variation in infectious bursal disease virus - analysis of a naturally-occurring

variant virus. Virus Genes 6:247-259. 1992.

136. Lasher, H., and S. Shane. Infectious bursal disease. Worlds Poult Sci J. 50:133-

166. 1994.

137. Lasher, H., and V. Davis. History of infectious bursal disease in the USA - the first

two decades. Avian Dis. 41:11-19. 1997.

138. Ledeboer, A. M., L. Edens, J. Maat, C. Visser, J. W. Bos, C. T. Verrips, Z.

Janowicz, M. Eckart, R. Roggenkamp, and C. P. Hollenberg. Molecular cloning

and characterization of a gene coding for methanol oxidase in hansenula

polymorpha. Nucleic Acids Res 13:3063-3082. 1985.

67

139. Lee, L. H., S. L. Yu, and H. K. Shieh. Detection of infectious bursal disease virus

infection using the polymerase chain reaction. J Virol Methods 40:243-253. 1992.

140. Lee, N., Y. Nomura, and T. Miyazaki. Gill lamellar pillar cell necrosis, a new

Birnavirus disease in Japanese eels. Dis Aquat Organ. 37:13-21. 1999.

141. Lejal, N., B. Da Costa, J. C. Huet, and B. Delmas. Role of ser-652 and lys-692 in

the protease activity of infectious bursal disease virus VP4 and identification of its

substrate cleavage sites. J Gen Virol 81:983-992. 2000.

142. Lim, B. L., Y. C. Cao, T. Yu, and C. W. Mo. Adaptation of very virulent

infectious bursal disease virus to chicken embryonic fibroblasts by site-directed

mutagenesis of residues 279 and 284 of viral coat protein VP2. J Virol. 73:2854-

2862. 1999.

143. Lin Cereghino, G. P., A. J. Sunga, J. Lin Cereghino, and J. M. Cregg. Expression

of foreign genes in the yeast pichia pastoris. Genet Eng (N Y) 23:157-169. 2001.

144. Lin, Z., A. Kato, Y. Otaki, T. Nakamura, E. Sasmaz, and S. Ueda. Sequence

comparisons of a highly virulent infectious bursal disease virus prevalent in Japan.

Avian Dis 37:315-323. 1993.

145. Liu, H., X. Tan, K. A. Russell, M. Veenhuis, and J. M. Cregg. Per3, a gene

required for peroxisome biogenesis in pichia pastoris, encodes a peroxisomal

membrane protein involved in protein import. J Biol Chem 270:10940-10951.

1995.

146. Liu, H. J., P. H. Huang, Y. H. Wu, M. Y. Lin, and M. H. Liao. Molecular

characterization of very virulent infectious bursal disease viruses in Taiwan. Res

Vet Sci 70:139-147. 2001.

68

147. Liu, M., and V. N. Vakharia. VP1 protein of infectious bursal disease virus

modulates the virulence in vivo. Virology 330:62-73. 2004.

148. Lombardo, E., A. Maraver, I. Espinosa, A. Fernandez-Arias, and J. Rodriguez.

VP5, the nonstructural polypeptide of infectious bursal disease virus, accumulates

within the host plasma membrane and induces cell lysis. Virology 277:345-357.

2000.

149. Lombardo, E., A. Maraver, J. R. Cast n, J. Rivera, A. Fernandez-Arias, A.

Serrano, J. L. Carrascosa, and J. F. Rodriguez. VP1, the putative RNA-dependent

RNA polymerase of infectious bursal disease virus, forms complexes with the

capsid protein VP3, leading to efficient encapsidation into virus-like particles. J

Virol 73:6973-6983. 1999.

150. Lukert, P. D., and Y. M. Saif Infectious bursal disease. In: Diseases of

Poultry, 11th edition ed. Y.M. Saif, H.J. Barnes, J.R. Glisson, A.M. Fadly, L.R.

Mc Dougald, D.E. Swayne. Iowa State University Press, Ames, IA. pp 161-179.

2003.

151. Lukert, P. D., J. Leonard, and R. B. Davis. Infectious bursal disease virus - antigen

production and immunity. Am J Vet Res. 36:539-540. 1975.

152. Macauley-Patrick, S., M. L. Fazenda, B. McNeil, and L. M. Harvey. Heterologous

protein production using the pichia pastoris expression system. Yeast 22:249-270.

2005.

69

153. Macreadie, I. G., P. R. Vaughan, A. J. Chapman, N. M. McKern, M. N. Jagadish,

H. G. Heine, C. W. Ward, K. J. Fahey, and A. A. Azad. Passive protection against

infectious bursal disease virus by viral VP2 expressed in yeast. Vaccine 8:549-

552. 1990.

154. Martinez-Torrecuadrada, J., B. Lazaro, J. Rodriguez, and J. Casal. Antigenic

properties and diagnostic potential of baculovirus-expressed infectious bursal

disease virus proteins VPx and VP3. Clin Diag Lab Immunol.7:645-651. 2000.

155. Martinez-Torrecuadrada, J., N. Saubi, A. Pages-Mante, J. Caston, E. Espuna, and

J. Casal. Structure-dependent efficacy of infectious bursal disease virus (IBDV)

recombinant vaccines. Vaccine 21:3342-3350. 2003.

156. Mast, J., B. M. Goddeeris, K. Peeters, F. Vandesande, and L. R. Berghman.

Characterization of chicken monocytes, macrophages and interdigitating cells by

the monoclonal antibody kul01. Vet Immunol Immunopathol 61:343-357. 1998.

157. McAllister, J. C., C. D. Steelman, L. A. Newberry, and J. K. Skeeles. Isolation of

infectious bursal disease virus from the lesser mealworm, alphitobius-diaperinus

(panzer). Poultry Sci. 74:45-49. 1995.

158. McFerran, J. B., M. S. McNulty, F. R. McKillop, T. J. Connor, R. M. McCracken,

D. S. Collins, and G. M. Allan. Isolation and serological studies with infectious

bursal disease viruses from fowl, turkeys and ducks - demonstration of a 2nd

serotype. Avian Pathol. 9:395-404. 1980.

159. McNulty, M., and Y. Saif. Antigenic relationship of non- serotype turkey

infectious bursal disease viruses from the United States and United Kingdom.

Avian Dis. 32:374-375. 1988.

70

160. McNulty, M. S., G. M. Allan, and J. B. McFerran. Isolation of infectious bursal

disease virus from turkeys. Avian Pathol. 8:205-&. 1979.

161. Menendez, J., I. Valdes, and N. Cabrera. The icl1 gene of pichia pastoris,

transcriptional regulation and use of its promoter. Yeast 20:1097-1108. 2003.

162. Menendez, J., L. Hernandez, A. Banguela, and J. Pais. Functional production and

secretion of the gluconoacetobacter diazatrophicus fructose-releasing exo-

levanase (lsdb) in pichia pastoris. Enz Microb Technol:446-452. 2004.

163. Mertens, P. P., P. B. Jamieson, and P. Dobos. In vitro RNA synthesis by infectious

pancreatic necrosis virus-associated RNA polymerase. J Gen Virol 59:47-56.

1982.

164. Morales, O. E., and W. Boclair. Morphometric relations bursa/spleen in infectious

bursal disease. In: 42 nd, Western Poultry Disease Conference. Sacramento, CA.

pp 91-92. 1993.

165. Muller, H., and H. Becht. Biosynthesis of virus-specific proteins in cells infected

with infectious bursal disease virus and their significance as structural elements

for infectious virus and incomplete particles. J Virol 44:384-392. 1982.

166. Muller, H., and R. Nitschke. Molecular weight determination of the two segments

of double-stranded RNA of infectious bursal disease virus, a member of the

Birnavirus group. Med Microbiol Immunol (Berl) 176:113-121. 1987.

167. Muller, H., and R. Nitschke. Genome of infectious bursal disease virus (IBDV) -

electron-microscopic investigations and demonstration of a genome-linked

protein. Zentralbl Bakteriol 267:156-157. 1987.

71

168. Muller, H., and R. Nitschke. The 2 segments of the infectious bursal disease virus

genome are circularized by a 90,000-da protein. Virology 159:174-177. 1987.

169. Muller, H., M. Islam, and R. Raue. Research on infectious bursal disease - the

past, the present and the future. Vet Microbiol. 97:153-165. 2003.

170. Muller, R., I. Kaufer, M. Reinacher, and E. Weiss. Immunofluorescent studies of

early virus propagation after oral infection with infectious bursal disease virus

(IBDV). Zentralbl Veterinarmed B 26:345-352. 1979.

171. Mundt, E. Tissue culture infectivity of different strains of infectious bursal disease

virus is determined by distinct amino acids in VP2. J Gen Virol 80 (Pt 8):2067-

2076. 1999.

172. Mundt, E. Personal communication In. 2005.

173. Mundt, E., J. Beyer, and H. Muller. Identification of a novel viral protein in

infectious bursal disease virus-infected cells. J Gen Virol 76 (Pt 2):437-443. 1995.

174. Mundt, E., B. Kollner, and D. Kretzschmar. VP5 of infectious bursal disease virus

is not essential for viral replication in cell culture. J Virol 71:5647-5651. 1997.

175. Nagarajan, M. M., and F. S. Kibenge. The 5'-terminal 32 base pairs conserved

between genome segments a and b contain a major promoter element of infectious

bursal disease virus. Arch Virol 142:2499-2514. 1997.

176. Naqi, S. A., and D. L. Millar. Morphologic changes in the bursa of fabricius of

chickens after inoculation with infectious bursal disease virus. Am J Vet Res

40:1134-1139. 1979.

72

177. Nawathe, D. R., O. Onunkwo, and I. M. Smith. Serological evidence of infection

with virus of infectious bursal disease in wild and domestic birds in Nigeria. Vet.

Rec 102:444-444. 1978.

178. Nick, H., D. Cursiefen, and H. Becht. Structural and growth-characteristics of

infectious bursal disease virus. J Virol. 18:227-234. 1976.

179. Nieper, H., and H. Muller. Susceptibility of chicken lymphoid cells to infectious

bursal disease virus does not correlate with the presence of specific binding sites. J

Gen Virol 77 (Pt 6):1229-1237. 1996.

180. Nieper, H., J. P. Teifke, A. Jungmann, C. V. Lohr, and H. Muller. Infected and

apoptotic cells in the IBDV-infected bursa of fabricius, studied by double-labeling

techniques. Avian Pathol. 28:279-285. 1999.

181. Nunoya, T., Y. Otaki, M. Tajima, M. Hiraga, and T. Saito. Occurrence of acute

infectious bursal disease with high mortality in Japan and pathogenicity of field

isolates in specific-pathogen-free chickens. Avian Dis 36:597-609. 1992.

182. Odor, E. Elisa serology: Application to IBD in broiler production. In: International

Poultry Symposium-Summit on Infectious Bursal Disease. Athens, GA, U.S.A. pp

104-105. 1995.

183. Ogata, K., H. Nishikawa, and M. Ohsugi. A yeast capable of utilizing methanol.

Agric. Biol. Chem.:1519-1520. 1969.

184. Ohi, H., M. Miura, R. Hiramatsu, and T. Ohmura. The positive and negative cis-

acting elements for methanol regulation in the pichia pastoris aox2 gene. Mol Gen

Genet 243:489-499. 1994.

73

185. Ojeda, F., I. Skardova, M. Guarda, J. Ulloa, and H. Folch. Proliferation and

apoptosis in infection with infectious bursal disease virus: A flow cytometric

study. Avian Dis. 41:312-316. 1997.

186. Okoye, J. O. A. Susceptibility of laboratory rats and guinea-pigs to infectious

bursal disease virus-infection. Acta Vet Brno. 56:167-171. 1987.

187. Okoye, J. O. A., and U. E. Uche. Serological evidence of infectious bursal disease

virus-infection in wild rats. Acta Vet Brno. 55:207-209. 1986.

188. Okoye, J. O. A., and G. C. Okpe. The pathogenicity of an isolate of infectious

bursal disease virus in guinea fowls. Acta Vet Brno. 58:91-96. 1989.

189. Oppling, V., H. Muller, and H. Becht. Heterogeneity of the antigenic site

responsible for the induction of neutralizing antibodies in infectious bursal disease

virus. Arch Virol 119:211-223. 1991.

190. Pantin-Jackwood, M. J., and T. P. Brown. Infectious bursal disease virus and

proventriculitis in broiler chickens. Avian Dis. 47:681-690. 2003.

191. Pantin-Jackwood, M. J., T. P. Brown, and G. R. Huff. Reproduction of

proventriculitis in commercial and specific-pathogen-free broiler chickens. Avian

Dis 49:352-360. 2005.

192. Paramithiotis, E., and M. J. Ratcliffe. Evidence for phenotypic heterogeneity

among b cells emigrating from the bursa of fabricius: A reflection of functional

diversity? Curr Top Microbiol Immunol 212:29-36. 1996.

193. Peng, L., X. Zhong, J. Ou, S. Zheng, J. Liao, L. Wang, and A. Xu. High-level

secretory production of recombinant bovine enterokinase light chain by pichia

pastoris. J Biotechnol 108:185-192. 2004.

74

194. Persson, R. H., and R. D. Macdonald. Evidence that infectious pancreatic necrosis

virus has a genome-linked protein. J Virol 44:437-443. 1982.

195. Peters, G. Histology of gumboro disease. Berl Munch Tierarztl Wochenschr:394-

396. 1967.

196. Pike, K. A., E. Baig, and M. J. Ratcliffe. The avian B-cell receptor complex:

Distinct roles of Ig alpha and Ig beta in B-cell development. Immunol Rev 197:10-

25. 2004.

197. Pitcovski, J., D. Goldberg, B. Z. Levi, D. Di-Castro, A. Azriel, S. Krispel, T.

Maray, and Y. Shaaltiel. Coding region of segment a sequence of a very virulent

isolate of IBDV--comparison with isolates from different countries and virulence.

Avian Dis 42:497-506. 1998.

198. Pitcovski, J., D. Di-Castro, Y. Shaaltiel, A. Azriel, B. Gutter, E. Yarkoni, A.

Michael, S. Krispel, and B. Z. Levi. Insect cell-derived VP2 of infectious bursal

disease virus confers protection against the disease in chickens. Avian Dis 40:753-

761. 1996.

199. Pitcovski, J., B. Gutter, G. Gallili, M. Goldway, B. Perelman, G. Gross, S. Krispel,

M. Barbakov, and A. Michael. Development and large-scale use of recombinant

VP2 vaccine for the prevention of infectious bursal disease of chickens. Vaccine

21:4736-4743. 2003.

200. Poonia, B., and S. Charan. T-cell suppression by cyclosporin-a enhances

infectious bursal disease virus infection in experimentally infected chickens.

Avian Pathol. 30:311-319. 2001.

75

201. Pope, C. R. Lymhoid system, In Avian Histopathology 2nd edition. Ed.C. Riddle

American Association of Avian Pathologists, Kennet Square, PA. 1996.

202. Ramm, H. C., T. J. Wilson, R. L. Boyd, H. A. Ward, K. Mitrangas, and K. J.

Fahey. The effect of infectious bursal disease virus on b-lymphocytes and bursal

stromal components in specific pathogen-free (SPF) white leghorn chickens. Dev

Comp Immunol.15:369-381. 1991.

203. Rautenschlein, S., H. Y. Yeh, and J. M. Sharma. Comparative

immunopathogenesis of mild, intermediate, and virulent strains of classic

infectious bursal disease virus. Avian Dis 47:66-78. 2003.

204. Revet, B., and E. Delain. The drosophila x virus contains a 1-microm double-

stranded RNA circularized by a 67-kd terminal protein: High-resolution

denaturation mapping of its genome. Virology 123:29-44. 1982.

205. Rodenberg, J., J. M. Sharma, S. W. Belzer, R. M. Nordgren, and S. Naqi. Flow

cytometric analysis of b-cell and t-cell subpopulations in specific-pathogen-free

chickens infected with infectious bursal disease virus. Avian Dis. 38:16-21. 1994.

206. Romanos, M. Advances in the use of pichia pastoris for high-level gene

expression. Curr Opin Biotechnol:527-533. 1995.

207. Rosenberger, J. K., and S. S. Cloud. Isolation and characterization of variant

infectious bursal disease viruses. J Am Vet Med Assoc.189:357. 1986.

208. Rosenberger, J. K., Y. M. Saif, and D. J. Jackwood. Infectious bursal disease, In,

A Laboratory Manual for the Isolation and Identification of Avian Pathogens.4th

editioon. Ed.D.E. Swayne, J.R. Glisson, M.W. Jackwood, J.E. Pearson, W.M.

Reed American Association of Avian Pathologists, Kennet Square, PA. 1998.

76

209. Rosenfeld, S. A., D. Nadeau, J. Tirado, G. F. Hollis, R. M. Knabb, and S. Jia.

Production and purification of recombinant hirudin expressed in the

methylotrophic yeast pichia pastoris. Protein Expr Purif 8:476-482. 1996.

210. Sanchez, A., and J. Rodriguez. Proteolytic processing in infectious bursal disease

virus: Identification of the polyprotein cleavage sites by site-directed mutagenesis.

Virology 262:190-199. 1999.

211. Schnitzler, D., F. Bernstein, H. Muller, and H. Becht. The genetic-basis for the

antigenicity of the VP2-protein of the infectious bursal disease virus. Journal of

General Virology 74:1563-1571. 1993.

212. Segev, N., J. Mulholland, and D. Bostein. The yeast gtp-binding ypt1 protein and

a mammalian counterpart are associated with the secretion machinery. Cell:915-

924. 1988.

213. Sharma, J., I. Kim, S. Rautenschlein, and H. Yeh. Infectious bursal disease virus

of chickens: Pathogenesis and immunosuppression. Dev Comp Immunol.24:223-

235. 2000.

214. Sharma, J. M., and L. F. Lee. Effect of infectious bursal disease on natural-killer

cell-activity and mitogenic response of chicken lymphoid-cells - role of adherent

cells in cellular immune suppression. Infect Immun 42:747-754. 1983.

215. Sharma, J. M., J. E. Dohms, and A. L. Metz. Comparative pathogenesis of

serotype-1 and variant serotype-1 isolates of infectious bursal disease virus and

their effect on humoral and cellular immune competence of specific-pathogen-free

chickens. Avian Dis. 33:112-124. 1989.

77

216. Sheppard, M., W. Werner, E. Tsatas, R. McCoy, S. Prowse, and M. Johnson. Fowl

adenovirus recombinant expressing VP2 of infectious bursal disease virus induces

protective immunity against bursal disease. Arch Virol 143:915-930. 1998.

217. Simoes, E. A., and P. Sarnow. An RNA hairpin at the extreme 5' end of the

poliovirus RNA genome modulates viral translation in human cells. J Virol

65:913-921. 1991.

218. Skeeles, J. K., P. D. Lukert, E. V. Debuysscher, O. J. Fletcher, and J. Brown.

Infectious bursal disease viral-infections.2. Relationship of age, complement

levels, virus-neutralizing antibody, clotting, and lesions. Avian Dis. 23:107-117.

1979.

219. Slibinskas, R., D. Samuel, A. Gedvilaite, J. Staniulis, and K. Sasnauskas.

Synthesis of the measles virus nucleoprotein in yeast pichia pastoris and

Saccharomyces cerevisiae. J Biotechnol 107:115-124. 2004.

220. Smith, J. Personal communication In. 2005.

221. Snedeker, C., F. K. Willis, and I. Moulthorp. Some studies on infectious bursal

agent. Avian Dis:519-528. 1967.

222. Snyder, D. B. Changes in the field status of infectious bursal disease virus. Avian

Pathol. 19:419-423. 1990.

223. Snyder, D. B., V. N. Vakharia, and P. K. Savage. Naturally occurring-neutralizing

monoclonal-antibody escape variants define the epidemiology of infectious bursal

disease viruses in the united-states. Arch Virol.127:89-101. 1992.

78

224. Snyder, D. B., F. S. Yancey, and P. K. Savage. A monoclonal antibody-based

agar-gel precipitin test for antigenic assessment of infectious bursal disease

viruses. Avian Pathol. 21:153-157. 1992.

225. Snyder, D. B., D. P. Lana, B. R. Cho, and W. W. Marquardt. Group and strain-

specific neutralization sites of infectious bursal disease virus defined with

monoclonal-antibodies. Avian Dis. 32:527-534. 1988.

226. Snyder, D. B., D. P. Lana, P. K. Savage, F. S. Yancey, S. A. Mengel, and W. W.

Marquardt. Differentiation of infectious bursal disease viruses directly from

infected-tissues with neutralizing monoclonal-antibodies - evidence of a major

antigenic shift in recent field isolates. Avian Dis. 32:535-539. 1988.

227. Spies, U., H. Muller, and H. Becht. RNA-dependent RNA-polymerase activity in

infectious bursal disease virus (IBDV). Zentralbl Bakteriol 267:156-156. 1987.

228. Spies, U., H. Muller, and H. Becht. Properties of RNA polymerase activity

associated with infectious bursal disease virus and characterization of its reaction

products. Virus Research 8:127-140. 1987.

229. Survashe, B. D., I. D. Aitken, and J. R. Powell. The response of the harderian

gland of the fowl to antigen given by occular route. 1. Histological changes. Avian

Pathol:77-93. 1979.

230. Tacken, M., P. Rottier, A. Gielkens, and B. Peeters. Interactions in vivo between

the proteins of infectious bursal disease virus: Capsid protein VP3 interacts with

the RNA-dependent RNA polymerase, VP1. J Gen Virol. 81:209-218. 2000.

79

231. Tanimura, N., and J. Sharma. Appearance of T cells in the bursa of fabricius and

cecal tonsils during the acute phase of infectious bursal disease virus infection in

chickens. Avian Dis. 41:638-645. 1997.

232. Tanimura, N., and J. Sharma. In-situ apoptosis in chickens infected with infectious

bursal disease virus. J Comp Pathol. 118:15-27. 1998.

233. Tanimura, N., K. Tsukamoto, K. Nakamura, M. Narita, and M. Maeda.

Association between pathogenicity of infectious bursal disease virus and viral

antigen distribution detected by immunohistochemistry. Avian Dis 39:9-20. 1995.

234. To, H., T. Yamaguchi, N. T. Nguyen, O. T. Nguyen, S. V. Nguyen, S. Agus, H. J.

Kim, H. Fukushi, and K. Hirai. Sequence comparison of the VP2 variable region

of infectious bursal disease virus isolates from Vietnam. J Vet Med Sci 61:429-

432. 1999.

235. Torrents, D., J. Maldonado, N. Saubi, and E. A. Pages-Mant. Dogs as potential

carriers of infectious bursal disease virus. Avian Pathol 33:205-209. 2004.

236. Tsukamoto, K., C. Kojima, Y. Komori, N. Tanimura, M. Mase, and S.

Yamaguchi. Protection of chickens against very virulent infectious bursal disease

virus (IBDV) and Marek’s disease virus (MDV) with a recombinant MDV

expressing IBDV VP2. Virology 257:352-362. 1999.

237. Underwood, B. O., C. J. Smale, F. Brown, and B. J. Hill. Relationship of a virus

from tellina teniuis to infectious pancreatic necrosis virus. J Gen Virol:93-109.

1977.

238. Vakharia, V. N. Development of recombinant vaccines against infectious bursal

disease virus. Biotech Ann Rev:15-168. 1997.

80

239. Vakharia, V. N., J. He, B. Ahamed, and D. B. Snyder. Molecular basis of

antigenic variation in infectious bursal disease virus. Virus Res 31:265-273. 1994.

240. Vakharia, V. N., D. B. Snyder, D. Lutticken, S. A. Mengelwhereat, P. K. Savage,

G. H. Edwards, and M. A. Goodwin. Active and passive protection against variant

and classic infectious bursal disease virus-strains induced by baculovirus-

expressed structural proteins. Vaccine 12:452-456. 1994.

241. van den Berg, T. B., N. Eterradossi, D. Toquin, and G. Meulemans. Infectious

bursal disease (gumboro disease). In Diseases of Poultry: World Trade and Public

Health Implications, OIE Scientific and Technical Review:509-543. 2000.

242. van den Berg, T. B., A. Ona, D. Morales, and J. F. Rodriguez.Experimental

inoculation of game/ornamental birds with a very virulent strain of IBDV. In:

Proceedings II International Symposium on infectious bursal disease and chicken

infectious anaemia. Rauischholzhausen. pp 236-246. 2001.

243. van den Berg, T. P. Acute infectious bursal disease in poultry: A review. Avian

Pathol. 29:174-194. 2000.

244. van den Berg, T. P., and G. Meulemans. Acute infectious bursal disease in poultry:

Protection afforded by maternally derived antibodies and interference with live

vaccination. Avian Pathology:409-421. 1991.

245. van den Berg, T. P., M. Gonze, and G. Meulemans. Acute infectious bursal

disease in poultry - isolation and characterization of a highly virulent-strain. Avian

Pathol 20:133-143. 1991.

81

246. van den Berg, T. P., M. Gonze, D. Morales, and G. Meulemans. Acute infectious

bursal disease in poultry: Immunological and molecular basis of antigenicity of a

highly virulent strain. Avian Pathol 25:751-768. 1996.

247. van den Berg, T. P., D. Morales, N. Eterradossi, G. Rivallan, D. Toquin, R. Raue,

K. Zierenberg, M. F. Zhang, Y. P. Zhu, C. Q. Wang, H. J. Zheng, X. Wang, G. C.

Chen, B. L. Lim, and H. Muller. Assessment of genetic, antigenic and pathotypic

criteria for the characterization of IBDV strains. Avian Pathol 33:470-476. 2004.

248. Van der Marel, P., D. Snyder, and D. Lutticken. Antigenic characterization of

IBDV field isolates by their reactivity with a panel of monoclonal antibodies.

Dtsch Tierarztl Wochenschr 97:81-83. 1990.

249. van Loon, A. A. W. M., N. de Haas, I. Zeyda, and E. Mundt. Alteration of amino

acids in vp2 of very virulent infectious bursal disease virus results in tissue culture

adaptation and attenuation in chickens. J Gen Virol 83:121-129. 2002.

250. Vasconcelos, A. C., and K. M. Lam. Apoptosis induced by infectious bursal

disease virus. J Gen Virol 75 (Pt 7):1803-1806. 1994.

251. Vasconcelos, A. C., and K. M. Lam. Apoptosis in chicken embryos induced by the

infectious bursal disease virus. J Comp Pathol 112:327-338. 1995.

252. Veenhuis, M., J. P. van Dijken, and W. Harder. The significance of peroxisomes

in the metabolism of one-carbon compounds in yeast. Adv Microb Physiol:1-82.

1983.

253. Vervelde, L., and T. Davison. Comparison of the in situ changes in lymphoid cells

during infection with infectious bursal disease virus in chickens of different ages.

Avian Pathol26:803-821. 1997.

82

254. von Bulow, V., R. Rudolph, and B. Fuchs. Enhanced pathenogicity of chicken

anemia agent (CAA) in dual infections with marek’s disease virus (MDV),

infectious bursal disease virus (IBDV) or reticuloendotheliosis virus (REV).

Zentralbl Veterinarmed B:93-116. 1986.

255. Wegner, G. H. Emerging applications of the methylotrophic yeasts. FEMS

Microbiol Rev 7:279-283. 1990.

256. Welsh, R. M., J. M. McNally, M. A. Brehm, and L. K. Selin. Consequences of

cross-reactive and bystander ctl responses during viral infections. Virology 270:4-

8. 2000.

257. Williams, A., and T. Davison. Enhanced immunopathology induced by very

virulent infectious bursal disease virus. Avian Pathol34:4-14. 2005.

258. Winterfield, R., S. B. Hitchner, G. S. Appleton, and A. Cosgrove. Avian

nephrosis, nephritis and gumboro disease. L&M News Views:103. 1962.

259. Wu, H., N. K. Singh, R. D. Locy, K. Scissum-Gunn, and J. J. Giambrone.

Immunization of chickens with vp2 protein of infectious bursal disease virus

expressed in arabidopsis thaliana. Avian Dis 48:663-668. 2004.

260. Wu, J., J. Lin, L. Chieng, C. Lee, and T. Hsu. Combined use of gap and aox1

promoter to enhance the expression of human granulocyte-macrophage colony-

stimulating factor in pichia pastoris. Enz Microb Technol:453-459. 2003.

261. Wu, P. C., H. Y. Su, L. H. Lee, D. T. Lin, P. C. Yen, and H. J. Liu. Secreted

expression of the vp2 protein of very virulent infectious bursal disease virus in the

methylotrophic yeast pichia pastoris. J Virol Methods 123:221-225. 2005.

83

262. Wyeth, P. J. Effect of infectious bursal disease on response of chickens to s-

typhimurium and escherichia-coli infections. Vet Rec. 96:238-243. 1975.

263. Yamaguchi, T., K. Iwata, M. Kobayashi, M. Ogawa, H. Fukushi, and K. Hirai.

Epitope mapping of capsid proteins vp2 and vp3 of infectious bursal disease virus.

Arch Virol 141:1493-1507. 1996.

264. Yamaguchi, T., M. Ogawa, M. Miyoshi, Y. Inoshima, H. Fukushi, and K. Hirai.

Sequence and phylogenetic analyses of highly virulent infectious bursal disease

virus. Arch Virol 142:1441-1458. 1997.

265. Yao, K., and V. Vakharia. Induction of apoptosis in vitro by the 17-kda

nonstructural protein of infectious bursal disease virus: Possible role in viral

pathogenesis. Virology 285:50-58. 2001.

266. Yao, K., M. Goodwin, and V. Vakharia. Generation of a mutant infectious bursal

disease virus that does not cause bursal lesions. J Virol. 72:2647-2654. 1998.

267. Zhang, M. F., G. M. Huang, and S. L. Qiao. Early stages of infectious bursal

disease virus infection in chickens detected by in situ reverse transcriptase-

polymerase chain reaction. Avian Pathol31:593-597. 2002.

268. Zierenberg, K., H. Nieper, T. P. van den Berg, C. D. Ezeokoli, M. Voss, and H.

Muller. The VP2 variable region of African and German isolates of infectious

bursal disease virus: Comparison with very virulent, "classical" virulent, and

attenuated tissue culture-adapted strains. Arch Virol 145:113-125. 2000.

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Chapter III

Identification of infectious bursal disease viruses from RNA extracted from paraffin

embedded tissue1.

______________________

1Mohamed M. Hamoud and Pedro Villegas accepted in Avian Diseases Reprinted here with permission of publisher, 7/3/2006

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Abstract:

After histopathological screening bursa of Fabricius for the presence of infectious

bursal disease virus (IBDV), the hypervariable region of the VP2 gene for IBDV was

extracted from formalin fixed paraffin embedded tissue blocks. Using real-time RT-PCR

and sequencing, IBDV was identified in 227 different blocks. The ability to identify the

actual virus strain associated with the lesions observed microscopically in the bursa of

Fabricius allowed for direct correlation between viral identity and lesions, which may

help in designing vaccination strategies. Several new emerging viruses that do not group

with other known IBDV in phylogenetic tree analysis were identified, as well as a unique

variant virus that had 63 nucleotides missing from its hypervariable region.

Introduction:

Infectious bursal disease virus (IBDV) is the etiological agent of Gumboro

disease or infectious bursal disease (IBD). IBDV destroys the B lymphocyte precursors

found within the bursa of Fabricius, followed by bursal atrophy in young chickens (32).

In unprotected flocks, the virus may cause mortality and/ or immunosupression (16, 26).

There are two serotypes of IBDV: 1 and 2. All pathogenic viruses belong to serotype 1,

whereas serotype 2 viruses are non-pathogenic for both chickens and turkeys (12, 21).

Very virulent (vv) IBDV strains emerged in Europe in the late eighties, causing up to

60% mortality (4, 30). Strains of vvIBDV have also spread to other continents (1, 7, 28).

The epidemiological origin of vvIBDVs is still unknown (17).

IBDV belongs to the family Birnaviridae (genus Avibirnavirus) which includes

viruses with bisegmented dsRNA genomes (5). Genome segment A has 2 overlapping

open reading frames (ORF) the first encodes a precursor polyprotein in a large ORF, the

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product of which is cleaved by autoproteolysis to yield mature VP2 (outer capsid), VP4

(protease), andVP3 (inner capsid) (15). The other ORF of segment A encodes a non

structural protein,VP5 and is possibly involved in virus release (18, 33). Genome

segment B encodes the virus polymerase,VP1 (20). VP2 is the major host-protective

antigen of IBDV. It contains at least three independent epitopes responsible for the

induction of neutralizing antibody (2), thus the antigenic characterization of IBDV has

been based mainly on the study of the VP2 gene where some amino acid changes have

been shown to be the basis for antigenic variation (29), or have been proposed as putative

markers for vvIBDV (3, 22, 31).

Controlling infectious bursal disease (IBD) and its associated immune

suppression is critical to the broiler industry. This control is achieved by either

vaccinating breeder hens with conventional live attenuated and/or inactivated IBD

vaccines, or by the use of live IBD vaccines in broiler chicks, layers or young pullets to

provide active protection against IBDV, or a combination of both.

Using a histopathology archive can be very important in molecular research.

Bursal tissue specimens that have been preserved as paraffin blocks for many years

represent a historical collection of IBD. Paraffin embedded tissues have proved to be a

valuable source of DNA or RNA for molecular genetic analysis and the identification of

infectious agents like hepatitis C, and polio viruses (8,10, 24). The use of tissues that are

formalin fixed and embedded in paraffin as a source of genetic material may allow a

direct correlation between the IBDV strain present in the tissue and the pathological

lesions visualized by microscopy. Therefore, the information obtained by this technology

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may help refine vaccination programs for IBDV, as well as to detect any new emerging

IBDV strains.

The objective of this study was to utilize RNA extraction from formalin fixed paraffin

embedded tissue for rapid identification of nucleic acids from field strains of IBDV that

are causing actual lesions in commercial chickens, and comparing amino acid sequences

and nucleic acid sequences of the hypervariable region of VP2 of these field strains to

known vaccine and common IBDV strains.

Materials and methods

Tissue and Histopathological evaluation. Bursal tissues mainly from commercial

broiler flocks experiencing IBD-related problems from various regions of the world were

formalin fixed and paraffin embedded. Paraffin-embedded tissue samples were sectioned,

mounted, stained using hematoxylin and eosin (HE), and examined using light

microscopy. All sections of bursa of Fabricius were assigned a lesion score, where a

score of 1 represented no lesions, a score of 2 was defined as mild variation in follicle

size, 3 as moderate variation in follicle size, and 4 as either necrosis or follicle atrophy

(23). Tissues came from chickens with an age range from 14 to 84 days; tissues from

older birds were from broiler breeders.

Viruses: IBDV strains Edgar, Lukert, Variant A, Variant E, GLS, STC kept in our

laboratory were used as reference strains. Commercial IBDV vaccines S706™, SVS-

510™, IBD Blen™ and Bursa Blen™ (Merial Select Inc. Gainesville, GA), Clonevac

D78™ (Intervet Inc. Kenilworth, NJ), Univax™ (Schering-Plough Animal Health,

Millsboro, DE) and Bursine 2™ (Fort Dodge Animal Health, Fort Dodge, IA) were

obtained from each company.

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RNA Extraction. RNA was extracted from formalin-fixed, paraffin-embedded bursas

and examined for IBDV nucleic acid by real-time RT-PCR (23). Sections totaling 50 µm

in thickness were cut from each formalin-fixed, paraffin embedded tissue block,

deparaffinized in HemoDe (Fisher Scientific, Pittsburgh, PA), washed with 100%

ethanol, and digested with 25 ug/ml proteinase K (Sigma Chemical Co., St. Louis, MO)

for 1 hr at 50 C. RNA was extracted using Trizol (Life Technologies, Inc., Gaithersburg,

MD) according to the manufacturer’s recommendations, diluted in 25 µl of 90% dimethyl

sulfoxide, and frozen at -80 C until assayed. RNA from commercial vaccine viruses was

extracted using Trizol (Life Technologies, Inc., Gaithersburg, MD) according to the

manufacture’s recommendations, and then sequenced as mentioned below.

Real-time RT-PCR. Extracted RNA was denatured at 95 C for 5 min and kept on ice.

An RT-PCR was performed using reagents from the Light Cycler RNA Amplification

SYBR Green I Kit (ROCHE Molecular Biochemicals, Indianapolis, IN). The primers

used were designed to amplify a 276-base pair (bp) segment of the IBDV genome shared

by all strains, which represents the hypervariable region of the VP2 gene (primers

synthesized were different from those in cited reference). Amplification and detection of

specific products were also performed using a Light Cycler (ROCHE Molecular

Biochemicals, Indianapolis, IN) according to the manufacturer’s recommendations.

Briefly, RT-PCR was done at 55 C for 30 min., followed by denaturation at 95 C for 30

sec. Fifty five PCR cycles were performed, consisting of denaturation (95 C for 1 sec),

hybridization (55 C for 10 sec), and extension (72 C for 13 sec). A melting curve analysis

was done with an initial denaturation at 95 C. DNA melting was accomplished with an

initial temperature of 65 C for 10 sec and a gradual temperature increase of 0.1 C/sec

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until 95 C was reached. The melting temperature of the expected 276-bp amplicon was

between 82 C and 84 C. This estimated melting temperature was used to confirm the

identity of IBDV specific products obtained using real-time RT-PCR. Additional

confirmation of specific amplification was done using routine gel electrophoretic

techniques of the PCR products on 2% agarose (Sigma Chemical Co., St. Louis, MO),

followed by ethidium bromide staining (23).

Nucleotide sequence analysis: Extracted genomic material that was positive for IBDV

by real-time RT-PCR melting curve analysis was submitted to Integrated Biotech

Laboratories (IBL) at The University of Georgia (Athens, GA) for sequencing. The

nucleotide sequences were retrieved using Chromas program

(http://www.technelysium.com.au/chromas14x.html) and deduced amino acid sequences

were generated using EditSeq™ (DNASTAR inc. Madison, WI). All nucleic and

amino acid sequences were aligned using the clustal W method using the MegAlign™

program (DNASTAR inc. Madison, WI) and sequence distances calculated. Phylogenetic

analyses were conducted using MEGA version 3.0 software

(http://www.megasoftware.net/) by Neighbor-Joining, 1000 bootstrap replicates and p-

distance. This was used to compare the field IBDV strains to common vaccine strains

either sequenced in our lab or available on GenBank, as indicated by accession numbers

written before the strain name.

Selection of sample for evaluation : Tissues (bursa, spleen and thymus) were collected

globally from problem farms with immunosuppressive and performance issues. The

tissues were then fixed in 10% buffered formalin and shipped to a histology lab where the

tissues were processed and put into paraffin blocks. The blocks were cut with a

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microtome and slides were prepared for histological evaluation. The tissues were

evaluated by an avian histopathologist and the tissues with severe acute IBDV lesions

were identified and selected for genomic extraction. Real-time RT-PCR was carried out

on extracted genomic material and IBD was identified in samples by melting curve

analysis. The IBDV genomic material was sequenced and the sequences analyzed.

Results

Histopathological examination. There were 452 paraffin embedded blocks from IBDV

cases submitted to our laboratory from the beginning of January 2003 until the end of

June 2005 for IBDV testing. A majority of those blocks were characterized by the

presence of lesion scores of +3 or +4 (characterized by having severe acute bursitis);

while very few had +1 and +2 lesion scores.

Real time RT-PCR. Melting curve analysis detected that IBDV genomic material was

extracted from 227 of these blocks. The other 225 blocks yielded genomic material that

was negative for IBDV on real time RT-PCR testing, in spite of having severe

histological lesions characteristic of IBDV.

Cases from the United States of America totaled 207, of those 181 (87%)

contained IBDV genomic material. There were 245 international samples, of those only

46 (19%) were positive on real time RT-PCR testing. The extracted genomic material

expressed as location [number of positive cases] represented samples from Alabama [16],

Arkansas [10], Colorado [4], Delaware [22], Georgia [22], Iowa [2], Maryland [1],

Michigan [1], Mississippi [45], North Carolina [19], Ohio [4], Pennsylvania [4], South

Carolina [2], Texas [1], Virginia [13], and unknown domestic origin [15]. While

international samples represented Argentina [4], Brazil [4], Dominican Republic [2], El-

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Salvador [7], Guatemala [4], Mexico [6], Peru [7], Venezuela [12], China [6] and Japan

[1] (Tables 2 and 3).

Some of the submitted samples were autolytic, had chronic lesions, or contained

mild lesions (lesion score ≤ +2) and yielded no detectable genomic material upon

extraction with an exception of one block that had a +2 lesion score and was identified as

a Variant E virus.

Nucleotide and deduced amino acid analysis. The nucleotide sequences of extracted

genomic material from 227 cases were determined across 276 base pairs (bp) of the VP2

hypervariable region. The most dominant viruses present in the U.S.A were the Variant E

viruses while internationally they were the classic strains (Table 1). The geographic

origins of different strains of IBDV are illustrated in Tables 2 and 3. These 227 cases

included 29 classic, 15 Variant A, 160 Variant E, 4 vvIBDV, 7 that could not be

sequenced (Real time melting curve analysis was positive but sequence results - from 2

different independent labs - had too many unknown nucleotides to be properly identified)

and 12 unique viruses that did not fall in any particular group (table 1).

Several virus strains exhibited major differences from other IBDV strains were

identified and therefore are referred here as “unique viruses”. Examples for these unique

viruses included cases identified as 077 and 266. Case 077, in the absence of case 266

falls on a branch of its own (Figure 2), but in the presence of case 266, it clusters with

Nobilis Gumboro 228E™ (Figure 1) with 94.2% similarity in nucleic acid and 94.5% in

deduced amino acid analysis. Case 266 was on a branch of its own on both nucleic and

amino acid phylogenetic trees and did not cluster with any other IBDV strains (figure 1).

Sequence distance data showed that case 266 was closest to Variant E with 96.0%

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similarity in nucleotide sequence but was closest to STC and AF457106 Univax G603

(Schering Plough has G603 under the trade name Bursavac) with 92.3 % similarity in

deduced amino acid analysis.

Another example for unique viruses was case 087 (figure 1). Extraction and

identification of its genomic material revealed that this IBD virus strain lacked 63

nucleotides from its VP2 hypervariable region. In spite of this, the virus was apparently

still capable of replication in the bursa and caused acute follicle necrosis with no

regeneration (+4 lesion score). Nucleic acid analysis indicated that this was 98.1% similar

to Variant A virus while its deduced amino acid analysis concluded it was 95.7% similar

to both Variant A and Variant E viruses.

One Variant E virus was identified from a block containing tissue with +2 lesion

score, and this was an extremely rare result; as all other +2 blocks from experimentally

infected birds and commercial submissions did not yield genomic material for IBDV.

In multivalent vaccines, all the different strains present could not be identified.

Only the viral strain with the highest viral titer was identified in these multivalent

vaccines.

Discussion:

Controlling infectious bursal disease (IBD) and its associated immune

suppression is critical to the broiler industry. This control is achieved by either

vaccinating breeder hens and/or progeny to provide protection against IBDV. The use of

these vaccines may have caused selection pressure on field viruses that has led to the

emergence of variant strains of IBDV (27). It is also speculated that the very virulent

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strains of the IBDV (vvIBDV) may have emerged for the very same reasons that caused

the variant strains to emerge (25).

Because IBD is primarily controlled using vaccination, the failure of vaccines to

generate adequate immunity in young chicks is a major concern. One cause of vaccine

failure can be attributed to genetic drift of field viruses, which has lead to antigenically

divergent IBDV strains known as variants (9, 11). Current variant and classic IBDV

vaccines are able to protect against many of the antigenically divergent IBDV strains (9,

14). However, new antigenic variants of the virus have been reported to break through

maternal and active immunity generated using commercially available vaccines (14).

The purpose of this study was to rapidly identify field isolates of IBDV by using

formalin fixed paraffin embedded tissues as a source of genomic material. This is of

importance because it will allow a direct correlation between the virus strain present and

the lesions produced by this virus in a timely fashion. It will also allow analysis of

foreign strains of IBDV with no importation and biosecurity restriction issues because of

the inactivated nature of the antigen. Therefore, the information obtained by this

technology will help refine vaccination programs for IBDV, as well as detect any new

emerging IBDV strains both domestically and internationally.

Analysis of 452 paraffin embedded tissue blocks for presence of IBDV was based

on histopathological lesion scores for IBDV. These 452 samples were not selected at

random. Instead, they were submitted because IBD was suspected in the flock or on the

farm. This study was therefore biased toward IBDV on farms experiencing immune

suppression-related problems. When the viral genomes identified were from viruses

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associated with disease in commercial poultry operations, they had a greater potential to

be genetic and antigenic mutants (13).

In chickens experimentally infected with IBDV, all paraffin embedded blocks

with bursas having acute +3 or +4 bursitis had extractable genomic material that was

positive for IBDV based on melting curve analysis, indicating high sensitivity of primers

used. Most of the 452 blocks analyzed had +3 or +4 acute lesion scores for IBDV, yet

only 227 were positive for IBDV extractable genomic material. IBDV genomic extraction

rate from samples originating from the U.S.A. was 87%, while that from overseas was

only 19%. A possible explanation is that most of the cases that had acute +3 or +4

histological lesion score for IBDV but were negative on real-time RT-PCR melting curve

analysis for IBDV, were inadequately fixed. This conclusion was based on experimental

simulation of different fixation conditions that could occur in the field, where bursas

positive for extractable IBDV RNA were rendered negative just by changing fixation

conditions. Improving fixation conditions will improve extraction of RNA from paraffin

embedded tissue (6). The seven samples that could not be sequenced were sequenced

repeatedly in 2 different laboratories and yielded similar non specific results. They were

not false positives as they had segments of IBDV sequence, yet they could not be aligned

due to the presence of too many non specific nucleotides (Ns). Also this genomic

material was extracted from bursa’s with specific +4 acute lesions of IBDV.

Nucleic acid and deduced amino acid analysis of extracted IBDV genomic

material has shown that the Variant E strain of IBDV is the predominant strain present in

the U.S.A. as it has also been observed by Jackwood and Sommer-Wagner (13). Classical

strains found in the United States were mainly of vaccine origin. There were no vvIBDV

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strains found in the U.S.A. Internationally, four vvIBDV strains from both Venezuela and

China were identified. Variant A strains were found in both Peru and Venezuela. Variant

E strains were identified in El Salvador, Guatemala, Mexico, Peru and Venezuela.

Internationally, more classical than variant strains were identified, which is similar to

what Lukert and Saif have reported (19).

Phylogenetic analysis shows that serotype 1 viruses grouped together in several

different groups. These groups were identified as Lukert, Edgar, vvIBDV, STC, 2512,

Websters, various classic, Variant A and Variant E groups. Previous knowledge of strain

origin and identity along with location of viruses on phylogenetic trees was the basis

used for grouping these viruses. The Faragher 52/70 (Gallivac) and the 228E strains fell

on different branches and did not cluster with any other classic IBDV strains (Figure 1

and 2). It was noticed that BursaVac™ fell into two different clusters, the first (GenBank

accession number AF498633) being in the STC cluster and the second (AF148075) was

in the Websters cluster. The same observation was seen for Bursine 2™, where the

Bursine 2 sequenced in our laboratory fell in the Lukert group (similar to the Bursine 2

with GenBank accession number AF498631 - not in figure) while Bursine 2 with

accession number AY332561 fell in the group named various classic (Figure 1 and 2).

The explanation for this may be the use of different strains for production of the same

vaccine, different strains are being given the same trade name by different companies and

researchers, or mislabeling occurred and sequences were posted incorrectly on GenBank.

Therefore, extreme caution has to be made when advising which vaccine to use, because

although two vaccines may have the same trade name, they may contain different IBDV

strains which may not cross protect against the same field isolates of IBDV. To overcome

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this problem, it is best if the vaccines that are used for comparison to field isolates be

sequenced by the researchers themselves. Variant viruses clustered into Variant A and

Variant E. Variant E had three different branches, with strain 89/03 on one branch,

RS593 on the second branch and Delaware E on the third branch.

Due to the large number of field strains analyzed in this study, only representative

samples are discussed. Several unique virus strains that did not group with the common

IBDV strains and fell on branches of their own were identified; examples for this are

cases 077 and 266. Sequence distance data showed that case 266 had a peculiar property

of being closest to a Variant E strain (96.0% similarity) when analyzing its nucleotide

sequence but its deduced amino acid sequence distance data concluded it was closest to

STC and Bursavac G603 strains (92.3 % similarity). The reason for the shift from variant

to classic strain based upon deduction of amino acid sequence from a nucleotide

sequence is unknown. There may be a need to include these strains (077 and 266) in

future IBDV vaccines depending on how widespread these viruses become among

chicken flocks, as each of these strains was isolated from one farm only.

Genomic material was isolated and identified from a unique IBD virus strain that

was missing 63 nucleotides from its hypervariable region in the VP2 gene, but was

apparently able to replicate in the bursa and cause lymphocytic damage. This unique

virus was given case number 087. Nucleic acid analysis indicated that the virus in case

087 was 98.1% similar to Variant A virus while its deduced amino acid analysis

concluded it was 95.7% similar to both Variant A and Variant E viruses. VP2 is

important for virus attachment to cell receptors and is the viral protein that is recognized

by the chicken’s immune system (2, 29). Missing these 21 amino acids apparently did not

97

affect the virus’s ability to attach and enter the susceptible B-lymphocytes. Constant

vaccination pressure in the past may have contributed to the emergence of variant strains

and vvIBDV (25, 27). Case 087 may have resulted from random mutations by the virus in

an attempt to escape from the immunoglobulins produced in vaccinated chickens; or

maybe this virus was always there in the population of IBDV viruses and constant

vaccination against variant IBD viruses suppressed other viruses, giving this virus a

chance to infect and propagate in the bursas. Also, the equal base pair distances from

both Variant A and Variant E viruses may provide cross protection against both Variant

A and E viruses, however, this needs to be investigated.

The strain identified from the bursa of Fabricius with a lesion score of +2 needs to

be further investigated to see if it is truly apathogenic, and if the virus possesses any

protective abilities against naturally occurring viruses. In the few bursa of Fabricius

encountered with a lesion score of +2, genomic material for IBDV could not be detected

in any of those blocks, this is likely due to low viral load or because these lesions were

due to stressors other than IBDV; with the exception of this case where there was enough

viral load in the block to extract genomic material. It was identified as a Variant E.

From the large number of samples submitted for identification, most of them had

more than 95% similarity to reference strains, example: case 290 was 100% identical in

nucleic acid to STC and Bursavac G603™, case 1042-1 was most similar to Edgar IBDV

strains, cases 340, 366 and 373 were most similar to AL-1 strain of Variant E.

The inability to identify all strains present in multivalent vaccines was attributed

to the nature of the RT-PCR reaction which amplifies the virus with higher logarithmic

concentration than the other viruses that have lower concentration. So when a sequencing

98

reaction is run and analyzed, only the nucleotides that are most abundant are identified.

This was noticed when sequencing a bivalent vaccine, SVS-510™, where only the

predominant variant portion of the vaccine was identified and not the classic portion. To

overcome this, the seeds of that vaccine were isolated and sequenced independently. This

variant portion of this bivalent vaccine was identified in figure 1 and 2 as SVS-510

(Variant portion) while the classical portion was identified as SVS-510 (Standard

portion).

The research potentials of this technology have allowed quick profiling of IBDV

strains that are causing lesions in the field. This is a diagnostic tool that can assist

veterinarians in choosing which vaccines to use based on the nucleic acid sequence of the

hypervariable region of VP2. This in turn may allow the identification of potentially new

vaccine strains.

As with all molecular techniques that are used to predict the relative similarities

and differences between IBDV strains, determining the actual antigenic differences

among viruses requires testing in-vivo (14).

References:

1. Banda, A., and P. Villegas. Genetic characterization of very virulent Infectious Bursal

Disease viruses from Latin America. Avian Dis. 48:540-549. 2004.

2. Becht, H., H. Muller, and H. Muller. Comparative studies on the structural and

antigenic properties of two serotypes of Infectious Bursal Disease virus. J. Gen.

Virol. :631-640. 1998.

99

3. Brandt, M., K. Yao, M. Liu, R. A. Heckert, and V. N. Vakharia. Molecular

determinants of virulence, cell tropism, and pathogenic phenotype of infectious

bursal disease virus. J. Virol. 75:11974-11982. 2001.

4. Chettle, N., J. C. Stuart, and P. J. Wyeth. Outbreak of virulent infectious bursal disease

in East Anglia. Vet. Rec. 125:271-272. 1989.

5. Dobos, P., L. Berthiaume, J. A. Leong, F. S. Kibenge, H. Muller, and B. Nicholson.

Family birnaviridae. In: Virus taxonomy. Classification and Nomenclature of

viruses, Sixth report on the international committee on Taxonomy of Viruses.

Murphy, F.A., Fauquet, C.M., Bishop, D.H.L., Ghabrial, S.A., Jarvis, A.W.,

Martelli, G.P., Mayo, M.A., Summers, M.D. ed. Arch. Virol. [Suppl. 10]

Springer, Wein, New York. 1995.

6. Hamoud, M. H., T. P. Brown, and M. J. Pantin-Jackwood. Molecular characterization

of viral pathogens: Extraction of genomic RNA from paraffin embedded tissues.

In: 141 AVMA annual convention. CD ROM. Philadelphia, PA. 2004.

7. Hassan, M. K. Very virulent infectious bursal disease virus in Egypt: Epidemiology,

isolation and immunogenicity of classic vaccine. Vet. Res. Commun. 28:347-356.

2004.

8. Hofmann, W.P., V. Dries, E. Herrmann, B. Gartner, S. Zeuzem, and C. Sarrazin.

Comparison of transcription mediated amplification (TMA) and reverse

transcription polymerase chain reaction (RT-PCR) for detection of hepatitis C

virus RNA in liver tissue. J. Clin. Virol. Apr;32(4):289-293. 2005

9. Ismail, N. M., and Y. M. Saif. Immunogenicity of infectious bursal disease viruses in

chickens. Avian Dis 35:460-469. 1991.

100

10. Jackson, D., F. Lewis, G. Taylor, A. Boylston, and P. Quirke. Tissue extraction of

DNA and RNA and analysis by the polymerase chain reaction. J. Clin.

Pathol.:499-504. 990.

11. Jackwood, D., and S. Sommer. Genetic heterogeneity in the VP2 gene of infectious

bursal disease viruses detected in commercially reared chickens. Avian Dis.

42:321-339. 1998.

12. Jackwood, D., Y. Saif, and J. Huges. Characteristics and serological studies of 2

serotypes of infectious bursal disease virus in turkeys. Avian Dis. 26:871-882.

1982.

13. Jackwood, D. J., and S. E. Sommer-Wagner. Molecular epidemiology of infectious

bursal disease viruses: Distribution and genetic analysis of newly emerging

viruses in the United States. Avian Dis. 49:220-226. 2005.

14. Jackwood, D. J., S. E. Sommer, and H. V. Knoblich. Amino acid comparison of

infectious bursal disease viruses placed in the same or different molecular groups

by RT/PCR-RFLP. Avian Dis. 45:330-339. 2001.

15. Kibenge, F. S., A. S. Dhillon, and R. G. Russell. Biochemistry and immunology of

infectious bursal disease virus. J. Gen. Virol. 69 (Pt 8):1757-1775. 1988.

16. Lasher, H., and S. Shane. Infectious bursal disease. World Poul. Sci. J. 50:133-166.

1994.

17. Le Nouen, C., G. Rivallan, D. Toquin, and N. Eterradossi. Significance of the genetic

relationships deduced from partial nucleotide sequencing of infectious bursal

disease virus genome segments A or B. Arch. Virol. 150:313-325. 2005.

101

18. Lombardo, E., A. Maraver, I. Espinosa, A. Fernandez-Arias, and J. F. Rodriguez.

VP5, the nonstructural polypeptide of infectious bursal disease virus, accumulates

within the host plasma membrane and induces cell lysis. Virol. 277:345-357.

2000.

19. Lukert, P. D., and Y. M. Saif. Infectious bursal disease. In: Diseases of

Poultry, 11th edition ed. Y.M. Saif, H.J. Barnes, J.R. Glisson, A.M. Fadly, L.R.

Mc Dougald, D.E. Swayne. Iowa State University Press, Ames, IA. pp 161-179.

2003.

20. Macreadie, I. G., and A. A. Azad. Expression and RNA dependent RNA polymerase

activity of birnavirus VP1 protein in bacteria and yeast. Biochem. Mol. Biol. Int.

30:1169-1178. 1993.

21. McNulty, M., and Y. Saif. Antigenic relationship of non- serotype turkey infectious

bursal disease viruses from the United States and United Kingdom. Avian Dis.

32:374-375. 1988.

22. Mundt, E. Tissue culture infectivity of different strains of infectious bursal

disease virus is determined by distinct amino acids in VP2. J. Gen. Virol. 80 (Pt

8):2067-2076. 1999.

23. Pantin-Jackwood, M. J., T. P. Brown, Y. Kim, and G. R. Huff. Proventriculitis in

broiler chickens: Effects of immunosuppression. Avian Dis. 48:300-316. 2004.

24. Rekand, T., R. Male, A.O. Myking, S.J. Nygaard, J.A. Aarli, L. Haarr. and N.

Langeland. Detection of viral sequences in archival spinal cords from fatal cases

of poliomyelitis in 1951-1952. J Virol Methods. Dec;114(1):91-96 2003.

102

25. Rong, J., T. Cheng, X. Liu, T. Jiang, H. Gu, and G. Zou. Development of

recombinant VP2 vaccine for the prevention of infectious bursal disease of

chickens. Vaccine 23:4844-4851. 2005.

26. Sharma, J. M., I. J. Kim, S. Rautenschlein, and H. Y. Yeh. Infectious bursal disease

virus of chickens: Pathogenesis and immunosuppression. Dev. Comp. Immunol.

24:223-235. 2000.

27. Snyder, D. Changes in the field status of infectious bursal disease virus. Avian Pathol.

19:419-423. 1990.

28. Tan, D. Y., M. Hair-Bejo, A. R. Omar, and I. Aini. Pathogenicity and molecular

analysis of an infectious bursal disease virus isolated from Malaysian village

chickens. Avian Dis. 48:410-416. 2004.

29. Vakharia, V. N., J. He, B. Ahamed, and D. B. Snyder. Molecular basis of antigenic

variation in infectious bursal disease virus. Virus Res. 31:265-273. 1994.

30. van den Berg, T. Acute infectious bursal disease in poultry: A review. Avian

Pathol. 29:175-194. 2000.

31. van Loon, A. A., N. de Haas, I. Zeyda, and E. Mundt. Alteration of amino acids in

VP2 of very virulent infectious bursal disease virus results in tissue culture

adaptation and attenuation in chickens. J. Gen. Virol. 83:121-129. 2002.

32. Wyeth, P. J., and G. A. Cullen. Maternally derived antibody-effect on susceptibility

of chicks to infectious bursal disease. Avian Pathol.:253-260. 1976.

33. Yao, K., and V. N. Vakharia. Induction of apoptosis in vitro by the 17-kda

nonstructural protein of infectious bursal disease virus: Possible role in viral

pathogenesis. Virol. 285:50-58. 2001.

103

Acknowledgment:

The authors would like to thank Dr. Stanley Kleven and Dr. John Glisson for

reviewing this manuscript. Histopathological screening for samples submitted after

August 2004 was done by Dr. Frederick Hoerr.

104

Figure Legend

Figure 3.1: Consensus nucleic acid phylogram of common IBDV vaccines and strains, in

addition to some field strains identified from paraffin embedded tissues deduced from

1000 trees by Neighbor-Joining method. Numbers at the forks indicate the number of

times (as a percentage) the groups consisting of the sequences which are to the right of

that fork occurred after the generation of 1000 trees. P-distance bar in bottom of figure.

Arrows indicate field cases identified in our laboratory which are labeled by case number.

Other isolates are identified with GeneBank accession numbers and trade name. Samples

without accession numbers were also extracted and sequenced in our laboratory. Only

213 nucleotides were sequenced in case 087 A.

Figure 3.2: Consensus amino acid phylogram of common IBDV vaccines and strains, in

addition to some field strains identified from paraffin embedded tissues deduced from

1000 trees by Neighbor-Joining method. Numbers at the forks indicate the number of

times (as a percentage) the groups consisting of the sequences which are to the right of

that fork occurred after the generation of 1000 trees. P-distance bar in bottom of figure.

Arrows indicate field cases identified in our laboratory which are labeled by case number.

Other isolates are identified with GeneBank accession numbers and trade name. Samples

without accession numbers were also extracted and sequenced in our laboratory. Only

213 amino acids were deduced in isolate 087 A.

105

Fig 3.1

106

Fig 3.2

107

Table 3.1: Number of IBDV strains isolated and identified from 2003-2005 according to geographical origin of the paraffin

embedded tissues

2003 2004 2005

Strains of IBDV

U.S.A. International U.S.A. International Unknown

origin

U.S.A. International

Classic 3 12 3 2 2 7

Variant A 3 0 5 0 1 3 3

Variant E 24 0 49 7 9 64 7

vvIBDV 0 1 0 0 0 0 3

Unique* 0 1 4 0 0 5 2

Strains that

could not be

sequenced §

0 1 1 0 5 0 0

Total 30 15 62 9 15 74 22

*Unique: Any virus that did not group with common strains of IBDV or possessed different genomic characteristics.

§Strains that could not be sequenced: The sequence data had too many unknown nucleotides (N’s) that when aligned with

other viruses, it was unidentifiable but melting curve analysis as well as histopathological examination confirmed it was an

IBDV

108

Table 3.2: Distribution of IBDV strains in the United States of America from 2003-2005

Location Classical Variant A Variant E Unique*

Alabama 1 15

Arkansas 7 3

Colorado 1 3

Delaware 1 4 15 2

Georgia 1 1 19 1§

Iowa 1 1

Maryland 1

Michigan 1

Mississippi 1 41 3

North

Carolina

1 18

Ohio 4

Pennsylvania 2 2

South

Carolina

2

Texas 1

Virginia 2 11

Unknown

origin

1 9 5§

*Unique: Any virus that did not group with common strains of IBDV or

possessed different genomic characteristics.

109

§ Strains that could not be sequenced: The sequence data had too many unknown

nucleotides (N’s) that when aligned with other viruses, it was unidentifiable but

melting curve analysis as well as histopathological examination confirmed it was an

IBDV

110

Table 3.3: Distribution of IBDV strains in the world during 2003-2005

Location Classical Variant A Variant E Unique

Argentina 4

Brazil 4

Dominican Republic 2

El-Salvador 2 5

Guatemala 1

Mexico 4 2

Peru 4 1 1 1

Venezuela 2* 2 5 3

China 4+2*

Japan 1

* vvIBDV

111

Chapter IV

Detection of infectious bursal disease virus from formalin-fixed paraffin-embedded

tissue by immunohistochemistry and real-time reverse transcription-polymerase

chain reaction1

______________________________

1Mohamed M. Hamoud, Pedro Villegas and Susan M. Williams Submitted to Journal of Veterinary Diagnostic Investigations

112

Abstract

Formalin fixed paraffin embedded tissues blocks are used routinely to diagnose

the economically important immunosuppressive infectious bursal disease virus (IBDV) in

chickens. Immunohistochemical detection of viruses in tissue blocks has been done with

varying results between laboratories. Extraction of IBDV RNA from tissue blocks allows

IBDV strain identification at a molecular level. This allows correlation between virus

identity and histological lesions present in the tissue. Experimentally RT-PCR detectable

IBDV RNA could always be extracted from tissue blocks with acute +3 or higher

histological lesion scores. However, many blocks from diagnostic field cases did not

yield detectable IBDV RNA, in spite of having severe IBDV histological lesion scores.

The reason for this can be the effect different formalin fixation conditions have on RNA

detection from tissue blocks. To study the effect of various fixation parameters on RNA

extraction and immunohistochemical detection of IBDV, bursas with high viral loads and

maximum histological lesion score of 4 for IBDV were fixed in formalin under various

conditions (different pH levels, temperatures, concentrations of formalin, and fixation

duration). Only tissues fixed in formalin with a pH of 7.0, concentration of 5 or 10%

formaldehyde, storage temperature of 25°C or less, and kept for up to 2 weeks in

formalin yielded detectable IBDV RNA upon extraction. No RNA could be detected

from tissues fixed under extreme temperature, pH or formalin concentrations. Optimal

fixation conditions for IHC detection of IBDV were 10% formalin concentration, pH 7.0

and temperature of 4°C, where maximum intensity of immunostaining was observed.

113

Key words: Infectious Bursal Disease, Chicken, Immunohistochemistry, RNA

extraction, Paraffin embedded tissue, formalin fixation.

114

Introduction

Classical infectious bursal disease (IBD), or Gumboro disease, is an acute, highly

contagious viral disease of young birds characterized by severe necrosis, hypoplasia, or

atrophy in the bursa of Fabricius, mainly observed at 3–6 weeks of age.23 IBD has

worldwide distribution, and the effects of the disease are economically significant to the

commercial poultry industry. Highly virulent strains cause increased mortality rates in

chickens. Survivors of classical IBD are immunosuppressed, which is also the outcome of

IBDV infection with antigenic variant strains. Variant IBDV differ from the classical

IBDV strains in that they usually do not cause mortality and can infect chicken younger

than 3 weeks of age. Variant viruses cause immunosuppression by viral induced

apoptosis of lymphocytes in the bursa. Immunocompromised chickens have an increased

susceptibility to other pathogens, such as Newcastle disease virus, infectious bronchitis

virus, infectious laryngotracheitis virus, Salmonella spp, or to other opportunistic agents

such as Escherichia coli. Furthermore, immunosuppressed birds may not respond

properly to vaccination and may suffer severe or persistent postvaccinal reactions.13

Infectious bursal disease virus (IBDV) is a member of the Birnaviridae family

with a genome consisting of two segments of double-stranded RNA. IBDV is a non-

enveloped icosahedral virus with a diameter of 55-60 nm. The smaller segment of RNA

encodes viral protein (VP) 1, which is the RNA polymerase of the virus. The large

segment encodes a polyprotein which is processed into three structural proteins, VP2,

VP3, and VP4.2,10,19 The large segment also encodes the structural protein VP5 in another

reading frame.15 Of these different genes, VP2 gene encodes for the major antigenic

115

protein and its hypervariable region is what confers variability among different IBDV

strains.

The severity of IBDV infection depends on the virulence of the virus, age of

birds, and level of circulating maternal antibodies. Cytotoxic T cells may exacerbate

virus-induced cellular destruction by lysing cells expressing viral antigens. T cells may

also promote the production of pro-inflammatory factors, such as nitric oxide, increasing

tissue destruction18. Outbreaks with high mortality rates caused by very virulent IBDV

(vvIBDV) strains have been reported in Europe, Asia and Latin America.21 Subclinical

and immunosuppressive forms of the disease are prevalent in the United States, where

reduction of body weight and lack of uniformity may be produced in mild outbreaks.13

Real-time RT-PCR (rt-RT-PCR) has been used to previously to detect IBDV in

tissues of infected chickens that were either frozen or stored in phenol.13,14 IBD viral

RNA has also been detected by rt-RT-PCR from formalin-fixed, paraffin-embedded

bursas.17 Using this technique, formalin fixed paraffin embedded bursa of Fabricius from

chickens experimentally infected with IBDV, and with histological lesion scores of +3 or

more always yielded rt-RT-PCR detectable IBDV RNA. However, diagnostic field cases

did not always yield rt-RT-PCR detectable RNA. Some samples (225 blocks over a 3-

year period) that had acute +4 histological lesion scores were negative for IBDV with rt-

RT-PCR. This finding occurred more often in diagnostic samples submitted from outside

the United States than in those originating from the United States (paper to be published

in Avian Diseases). The reason why some infected samples tested negative by rt-RT-PCR

has yet to be determined. It was recommended that these samples should be fixed in 10 %

buffered formalin and kept at room temperature, but pH analysis of formalin and H&E

116

artifacts indicated that some of the samples were fixed in more concentrated formalin,

formalin with variable pH levels or subjected to high temperatures during formalin

fixation. These observations suggested that formalin fixation conditions might have an

effect on rt-RT-PCR detectable RNA from tissue blocks.

Previous research has proved that formalin fixation of tissues is suitable for

immunohistochemistry because formalin is a relatively mild fixative for preservation of

antigenic reactivity. 22 However, formalin fixation can impair or block the antigenic

reactivity of certain proteins.1,4,16 This impairment may be due to extensive cross-linking

of proteins by formaldehyde, where amino groups become blocked and antibodies to

those amino groups do not participate in IHC staining.20

The objective of this study was to experimentally simulate field conditions in

terms of storage time, temperature, pH of unbuffered formalin, and different

concentrations of buffered formalin to test their effect on IBDV RNA extraction from

paraffin embedded tissue for use in rt-RT-PCR and immunohistochemical staining.

Materials and Methods

Birds and experimental design: A total of 95 specific pathogen free (SPF) chickens were

hatched from fertilized eggsa for this experiment. All birds were housed in Horsfall-

Bauer isolation units in 19 groups of five and maintained under positive pressure. Food

and water were provided ad libitum. At 3 weeks of age, the chickens were inoculated

with IBDV Blen vaccineb which contains the 2512 strain. Inoculation routes of intranasal,

oral and intramuscular administration (1x 104.0 EID 50/route) were given simultaneously

to each infected bird to ensure a high viral load in infected tissue. Five uninfected birds

117

were maintained as uninfected controls. Bursal tissue was collected 5 days post

inoculation and subjected to different fixation protocols as described in the experimental

design shown in Table 1.

Histopathology and immunohistochemistry. A 37% formaldehyde solution was diluted in

either neutral phosphate buffered saline (PBS), distilled water or was used undiluted for

formalin fixation. The pH was adjusted with either 10% sodium hydroxide or 5%

hydrochloric acid. Paraffin-embedded tissues were sectioned, mounted, stained with

hematoxylin and eosin (H&E) by using an automated stainerc, and examined for lesions

by light microscopy. All sections were assigned a lesion severity score. For all samples, a

lesion score of 1 represented no lesions; 2 represented mild reduction in overall follicle

size; 3 represented moderate reduction in follicle size; and 4 represented either necrosis

or follicle atrophy.17 All procedures for immunohistochemistry (IHC) were done at room

temperature. Five-micron thick tissue sections were cut from paraffin-embedded samples

and mounted on charged glass slidesd. Paraffin was melted from slides (10 min at 65°C)

and removed by immersion in Hemo-De e three times (5 min each). Slides were then air

dried. Antigen retrieval was done by incubating slides with 10% proteinase Kf for 5 min

at room temperature. IHC staining was performed with an automated stainerg using a

commercial kith according to the manufacturer’s recommendations. The primary antibody

used was a mouse monoclonal antibody specific to and cross-reactive among all IBDV

strains.i The secondary antibodyh was a peroxidase labeled polymer conjugated to anti-

mouse and anti-rabbit immunoglobulins. After IHC staining, sections were

counterstained with hematoxylin, cover slipped, and examined by light microscopy. The

degree of IBDV infection in each section was scored as follows: - = no infection;

118

+ = minimal infection (<10% of cells present); ++ = moderate infection (>10% and <30%

of cells present); and +++ = intense infection (>30% of cells present).17

RNA extraction and quantification. RNA was extracted from formalin-fixed, paraffin-

embedded bursas and examined for IBDV nucleic acid by rt-RT-PCR.17 Sections totaling

50 µm in thickness were cut from each formalin-fixed, paraffin-embedded tissue block,

deparaffinized in Hemo-Dee, washed with 100% ethanol, and digested with 25 ug/ml

proteinase Kf for 1 hr at 50°C. RNA was extracted using Trizolj according to the

manufacturer’s recommendations, resuspended in 25 µl of 90% dimethyl sulfoxide, and

frozen at -80◦C until assayed.17 Extracted RNA was quantified using eppendorf

biophotometerk and 50-2000 µl UVettel.

Real-time RT-PCR. The RNA extracted from tissue blocks was denatured at 95°C for 5

min and kept on ice. A rt-RT-PCR was performed using reagents from the LightCycler

RNA Amplification SYBR Green I Kitm and the LightCycler machinem according to

manufacturer’s recommendations. The primers (B5 5’ TCTTGGGTATGTGAGGCTTG

and B4 3’ GGATGTGATTGGCTGGGTTA)17 were designed to amplify a 400-base pair

(bp) segment of the hypervariable region of the IBDV VP2 gene (GenBank accession

number DQ355819). Briefly, RT was performed at 55°C for 30 min, followed by

denaturation at 95°C for 30 sec. Fifty-five PCR cycles were performed, consisting of

denaturation (95°C for 1 sec), hybridization (55°C for 10 sec), and extension (72°C for

13 sec). A melting curve analysis was done with an initial denaturation at 95°C (for 1

sec.). DNA melting was accomplished with an initial temperature of 65°C for 10 sec and

a gradual temperature increase of 0.1°C/sec until 95°C was reached. After which, there

was a cooling segment and a gradual temperature decrease of 2°C/ sec till the holding

119

temperature 20°C was reached. The melting temperature of the expected 400-bp

amplicon was between 82°C and 84°C. This estimated melting temperature was used to

confirm the presence of IBDV specific products obtained using real-time RT-PCR.17 A

no template control (DEPC-water) and a positive IBDV control was used in each run.

Electrophoresis of real-time RT-PCR products were run on 1.5% agarose gel and

visualized by ethidium bromide staining, to correlate between melting curve peaks and

size of amplified product. This was done initially to verify the specificity of the melting

curve peaks seen in the rt-RT-PCR reaction printouts.

Results

Histology and Immunohistochemistry. Out of 90 infected birds, 89 bursas had

lesion scores of 4, the remaining bursa had a lesion score of 3. All 5 chickens from the

uninfected negative control group had lesion scores of 1. All of the IBDV uninfected

negative control slides were negative using immunohistochemistry. All 5 IBDV infected

positive control slides demonstrated strong immunoreactivity for IBDV. This

demonstrated the specificity of the IHC technique used (Fig. 1, Table 2).

Histological evaluation of 5 bursas from each treatment group demonstrated that

for IBDV infected bursas fixed in different concentrations of formalin, the optimal

formalin concentration for formalin fixed tissues used for IHC staining was 10% formalin

(Table 2). The use of 5% formalin for formalin fixation produced a light nonspecific

background staining of erythrocytes and fibroblasts. IBDV antigen was not detected in

slides that came from paraffin embedded tissues fixed in 20% or 37% formalin (Table 2).

One area in one of the 5 samples fixed in 37% formalin had lymphoid cells with positive

staining. Light staining of apoptotic bodies, epithelial cytoplasm and vacuoles were

120

present and was interpreted as non-specific background staining. Hematoxylin and eosin

staining showed severe lymphoid depletion, atrophy in follicles, and increase in

interfollicular stroma. Artifacts seen in tissues fixed with 20% or 37% formalin

concentrations included granularity of interplical mucin, and mucin in epithelial cysts.

Tissues fixed in 37% formalin had diffuse eosinophilic staining.

Tissues fixed in 10% buffered formalin at different fixation temperatures showed

variable IHC staining. Fixation at 4°C yielded the best staining. The number of IHC

staining positive cells decreased as the fixation temperature increased (Table 2). H&E

artefacts observed in tissues stored at 50°C included streaming and smudging of nuclei,

loss of follicular architecture and very plump fibroblasts in stroma of atrophied plica.

The pH of formalin used for fixation had the most variable effects on IHC

staining. Optimal staining was observed when infected bursal tissue was fixed at pH 7.0.

At pH 2.0 minimal IHC staining of apoptotic bodies within the center of the follicles and

very rare moderate staining of mononuclear cells was observed in all 5 bursas of this

group. At pH 5.0 there was a complete loss of staining. All tissues fixed in formalin at pH

9.0 and 13.0 exhibited non-specific staining of apoptotic bodies and cellular debris. H&E

staining artifacts at pH 2.0 included tinctorial variations of epithelial cells, while at pH

5.0 there was cell shrinkage. At pH 9.0 there was streaming of epithelial nuclei and at pH

13.0 there were epithelial shrinkage and compressed fibroblasts.

IBDV RNA extraction and real time RT-PCR. The amount of extracted RNA

extracted from tissue blocks ranged from 159.92 – 1194.66 µg/ml. Specificity of rt-RT-

PCR melting curve analysis was verified by running amplicons on a 1.5% agarose gel

stained with ethidium bromide (Fig. 2). Detectable IBDV RNA was extracted from all 5

121

bursas stored in pH 7 as well as 1 of the 5 bursas stored in pH 13,0. All tissues fixed pH

2.0, pH 5.0 and pH 9.0 had a histopathological lesion score of 4 but none yielded

detectable IBDV RNA that was identifiable by rt-RT-PCR melting curve. (Table 3).

Real-time RT-PCR melting curve identifiable RNA was extracted from bursas stored at

temperatures 4°C and 25°C but not from 50°C (Table 3). Real time RT-PCR detectable

IBDV RNA was extracted from bursas of Fabricius stored in 5% or 10% buffered

formalin, but not from those tissues stored in 20% buffered formalin or 37% unbuffered

formalin (Table 3). Storage time for up to 2 weeks had no effect on IBDV RNA

extraction and identification, as melting curve peaks specific for IBDV were present at all

fixation times studied. However, storage times longer than 24 hours increased the melting

peak by 1°C; and after 2 weeks it was increased by 2°C. (Fig. 3).

Discussion

Formalin fixation is a common method for tissue preservation in preparation for

histological evaluation of tissues. For immunohistochemistry, small (10 X 10 X 3 mm)

tissue fragments fixed promptly in neutral buffered formalin for 6-24 hours generally had

adequate cytological preservation and immunolocalization, with minimal antigen

masking.9 Variations in fixation times or conditions cause the majority of false negatives

in immunohistochemistry.7 It was reported that lack of consistency in formalin fixation

protocols among laboratories, influences the outcome of staining in

immunohistochemistry.12 Formaldehyde fixes tissue by reacting primarily with basic

amino acids to form cross-linked “methylene bridges”. Formalin fixation conditions have

a direct effect on number of methylene bridges. 7 The number of methylene bridges and

122

which amino groups are cross-linked in the tissue are important variables in the outcome

of immunohistochemical staining, and could also be an important factor in the ability to

extract rt-RT-PCR detectable RNA from tissue blocks.

Tissues fixed in 20% or 37 % formalin undergo shrinkage10 and may suffer from

alteration of epitope conformation. The IBDV epitopes detected by the monoclonal

antibodies used in this study are conformationally dependent;5 thus fixation conditions

such as formaldehyde concentration or pH that may cause changes in epitope

conformation rendering them undetectable by these monoclonal antibodies. Maximum

tissue fixation, i.e. cross-linking, occurs in the pH range of 4 to 5.5 9, therefore, the

presence of too many cross-links or cross-links of certain amino groups may cause

antigen masking.20 This may be the reason why there was minimal or no immunostaining

in tissues fixed in pH ≤5.0. Inconclusive IHC staining at pH ≥9.0 may also be due to

tissue shrinkage or alteration of reactive epitopes.

Immunohistochemical staining was best in tissues fixed at 4°C. Increased

temperature reduced specific IHC staining, but did not eliminate it. The reduction in

staining may be due to tissue shrinkage at temperatures higher than 4°C.8 This result is in

agreement with Key’s findings; however the mechanism for this effect is unknown. 12

Real-time RT-PCR melting curve analysis of infected and non-infected controls

validated the RNA extraction procedure. Real-time RT-PCR on known IBDV genomic

material was performed as a positive rt-RT-PCR control and on DEPC-water as a

negative rt-RT-PCR control. Extracting genomic RNA for identification of IBDV from

bursas of Fabricius stored at acidic pH (pH 5.0 and pH 2.0) was unsuccessful in all 10

tissue blocks. Previous research which found DNA damage in cells at low pH, 11 may

123

explain the lack of extracted genomic RNA in this experiment. Unbuffered formalin

oxidizes to formic acid and an acidic environment causes degradation of nucleic acids

because the β-glycosidic bonds in the purine bases are hydrolyzed at pH 4.4 Loss of some

amino group cross-links occurs at low pH and may result in loss of RNA from the fixed

tissue.3 Extraction of genomic RNA at a basic pH (pH 9.0, and 13.0) was also

unsuccessful and the reason for this is unknown.

The thermodynamic stability of a nucleic acid duplex strongly depends on the

denaturation temperature of the duplex at atmospheric pressure.6 Denaturation of

genomic RNA may have occurred when tissues were formalin fixed at 50°C for 5 days

and might explain the absence of rt-RT-PCR detectable RNA from these blocks.

At formalin concentrations of 20% or 37 %, the extraction of RNA from tissue

blocks yielded no rt-RT-PCR detectable RNA. This may be due to excessive crosslinking

of amino groups resulting from the acidic nature of formalin at higher concentrations;

indeed, maximum tissue fixation, i.e. cross-linking, occurs in the pH range of 4.0 to 5.5,9

thus hindering genomic RNA extraction.

Fixation time had no effect on RNA extraction or subsequent identification

(Fig.3), but longer fixation times are not recommended due to the higher frequency of

non-reproducible sequence alterations. This is demonstrated by the increase in melting

peak temperature over time. Formalin may cause cross-linking of cytosine nucleotides on

either strand. As a result, in PCR the Taq-DNA polymerase fails to recognize the cytosine

and incorporates an adenine in the place of a guanosine, creating an artificial C-T or G-A

mutation.22

124

In summary, for optimal IBDV RNA extraction and sequence identification,

tissues should be fixed in formalin at a pH of 7.0, concentration of 5% or 10% and stored

at 4°C to 25°C for 7 days at most. Extremes in temperature, pH or formalin concentration

have a negative effect on tissues used for RNA extraction. Longer tissue storage times in

formalin may cause inaccuracies in sequencing results. Optimal conditions for fixation of

tissues infected with IBDV that will be used for IHC evaluation are 10% formalin

concentration, pH 7.0 and temperature of 4°C. For practical purposes, fixation

temperature of 4°C is recommended for both procedures. This can be achieved practically

by placing formalin jars or whirlpaks in a cooler with a cardboard separating the ice from

the samples.

Acknowledgments: The authors would like to thank Dr. Stanley Kleven and Linda

Purvis for reviewing this manuscript.

Sources and manufacturers:

a. Sunrise Farms, Catskill NY

b. IBD-Blen™ a vaccine, Merial Select, Gainesville GA, U.S.A.

c. Leica AutoStainer XL, Nussloch, Germany

d. Superfrost/Plus; Fisher Scientific Pittsburgh, PA, U.S.A.

e. HemoDe Fisher Scientific, Pittsburgh, PA, U.S.A.

f. Proteinase K: DAKO, Carpinteria, CA, U.S.A.

g. Leica ST 5050, Nussloch, Germany

h. DAKO Envision System; DAKO Carpinteria, CA, U.S.A.

i. Monoclonal antibody #HB9490, ATCC, Manassas, VA 20108 U.S.A.

125

j. Trizol: Life Technologies, Inc., Gaithersburg, MD, U.S.A.

k. Eppendorf biophotometer, Eppendorf AG, Hamburg, Germany

l. Uvette, Eppendorf AG, Hamburg, Germany

m. ROCHE Molecular Biochemicals, Indianapolis, IN, U.S.A.

References

1 Altmannsberger M, Osborn M, Schauer A, Weber K: 1981, Antibodies to Different Intermediate Filament Proteins - Cell Type-Specific Markers on Paraffin-Embedded Human-Tissues. Lab Invest 45:427-434.

2 Azad AA, Barrett SA, Fahey KJ: 1985, The characterization and molecular cloning of the double-stranded RNA genome of an Australian strain of infectious bursal disease virus. Virology 143:35-44.

3 Basyuk E, Bertrand E, Journot L: 2000, Alkaline fixation drastically improves the signal of in situ hybridization. Nucleic Acids Res 28:E46.

4 Bonin S, Petrera F, Niccolini B, Stanta G: 2003, PCR analysis in archival postmortem tissues. J Clin Pathol 56:184-186.

5 Coulibaly F, Chevalier C, Gutsche I, et al.: 2005, The birnavirus crystal structure reveals structural relationships among icosahedral viruses. Cell 120:761-772.

6 Dubins DN, Lee A, Macgregor RB, Chalikian TV: 2001, On the stability of double stranded nucleic acids. J Am Chem Soc 123:9254-9259.

7 Farmilo AJ, Stead RH, Atwood KN: 2001, Fixation. In: Immunochemical staining methods, ed. Boenisch T, 3rd ed. pp. 18-22. DAKO Corporation, Carpinteria, CA.

8 Fox CH, Johnson FB, Whiting J, Roller PP: 1985, Formaldehyde Fixation. J Histochem Cytochem 33:845-853.

9 Gustavson KH: 1956, Aldehyde tanning, pp. 244-282. Academic Press, New York. 10 Hudson PJ, McKern NM, Power BE, Azad AA: 1986, Genomic structure of the large

RNA segment of infectious bursal disease virus. Nucleic Acids Res 14:5001-5012.

11 Jolly AJ, Wild CP, Hardie LJ: 2004, Acid and bile salts induce DNA damage in human oesophageal cell lines. Mutagenesis 19:319-324.

12 Key M: 2001, Antigen retrieval. In: Immunochemical staining methods, ed. Boenisch T, 3rd ed. pp. 23-25. DAKO Corporation, Carpinteria, CA.

13 Lukert, P. D., and Y. M. Saif: 2003, Infectious bursal disease. In: Diseases of Poultry, ed. Y.M. Saif, H.J. Barnes, J.R. Glisson, A.M. Fadly, L.R. Mc Dougald, D.E. Swayne, 11th ed. pp 161-179. Iowa State University Press, Ames, IA.

14 Moody A, Sellers S, Bumstead N: 2000, Measuring infectious bursal disease virus RNA in blood by multiplex real-time quantitative RT-PCR. J Virol Methods 85:55-64.

15 Mundt E, Beyer J, Muller H: 1995, Identification of a novel viral protein in infectious bursal disease virus-infected cells. J Gen Virol 76 (Pt 2):437-443.

126

16 Nadji M, Morales AR: 1983, Immunoperoxidase: Part 1 The technique and its pitfalls. Lab Med 14:767-771.

17 Pantin-Jackwood MJ, Brown TP: 2003, Infectious bursal disease virus and proventriculitis in broiler chickens. Avian Dis 47:681-690.

18 Sharma J, Kim I, Rautenschlein S, Yeh H: 2000, Infectious bursal disease virus of chickens: pathogenesis and immunosuppression. Dev Comp Immunol 24:223-235.

19 Spies U, Muller H, Becht H: 1987, Properties of RNA polymerase activity associated with infectious bursal disease virus and characterization of its reaction products. Virus Res 8:127-140.

20 Sternberger LA: 1979, Immunocytochemistry, 2nd ed. John Wiley & Sons, New York.

21 Vandenberg TP, Gonze M, Meulemans G: 1991, Acute Infectious Bursal Disease in Poultry - Isolation and Characterization of a Highly Virulent-Strain. Avian Pathology 20:133-143.

22 Williams C, Ponten F, Moberg C, et al.: 1999, A high frequency of sequence alterations is due to formalin fixation of archival specimens. Am J Pathol 155:1467-1471.

23 Wyeth PJ, Cullen GA: 1978, Susceptibility of chicks to infectious bursal disease (IBD) following vaccination of their parents with live IBD vaccine. Vet Rec 103:281-282.

127

Figure Legend Figure 4.1. Photomicrograph of Bursa of Fabricius. (IHC technique specificity control

for IBDV infected and non-infected tissues). 1a: Non infected tissue stained with IHC.

1b: IBDV infected bursa stained with IHC. All tissues were fixed in 10% buffered

formalin at a temperature of 4°C and pH 7.0. IBDV infected controls show brown

positive IHC staining (indicated by arrow) of cortical intrafollicular lymphocytes. (100X)

Figure 4.2: RT-PCR product of amplified VP2 fragment: lane a, molecular size marker;

lane b, amplified VP2 fragment from IBDV control positive; lane c, amplified VP2

fragment from RNA extracted from a tissue block, lane d, negative RT- PCR control

(primers with water); lane e, negative extraction control from RNA extracted from an

uninfected tissue block

Figure 4.3. Real time RT-PCR melting curve analysis for IBDV amplicons extracted

from tissues fixed in 10% buffered formalin at 4°C and submitted to different fixation

durations (hours). Melting curve peak after 24 hours storage was 83°C, but increased to

84°C after 48-120 hours of storage, and after 2 weeks of storage, to 86°C, indicating

changes in nucleotide sequences. RT-control is the reverse transcriptase PCR control for

a known IBDV and extraction control is a known paraffin embedded block with

extractable IBDV genetic material.

128

Figure 4.1:

Figure 4.2:

129

Figure 4.3:

130

Table 4.1: Fixation conditions for bursal tissues from IBDV infected and control chickens.

Variables Storage conditions Fixation treatment a

5 %

10 %

20 %

Formalin

concentration

4°C for

5 days

37 %

4°C b

25°C

Formalin

temperature

10 % Buffered formalin for

5 days 50°C

2.0

5.0

7.0

9.0

pH

of formalin

4°C, in 10% Non buffered

formalin for 5 days

13.0

1 day

2 days

3 days

4 days

Duration in formalin 4°C, 10 % Buffered formalin

14 days

Infected Control c

10% Buffered

formalin at 4°C No variables

131

Non Infected Control c

10% Buffered

formalin at 4°C No variables

a Five individual bursas were tested per treatment.

b. This group was also used to represent 5 days duration in formalin

c Non infected and infected controls were stored in 10% buffered formalin at 4°C to

validate IHC procedure.

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Table 4.2: Histopathology and IHC mean staining scores of 5 bursas of Fabricius from

chickens infected with IBDV and subjected to different fixation conditions

Variable Treatment Lesion Score a IHC b

5 % 4 ± 0 +

10 % 4 ± 0 +++

20 % 3.2 ± 0.44 -

Formalin

Concentration c

37 % 4 ± 0 -

4°C 4 ± 0 +++

25°C 4 ± 0 ++

Formalin

temperature d 50°C 4 ± 0 +

2 4 ± 0 +

5 4 ± 0 -

7 4 ± 0 +++

9 4 ± 0 +

pH of

formalin e

13 4 ± 0 ±

Non infected

control

10% Buffered

formalin at 4°C 1 ± 0 -

Infected control 10% Buffered

formalin at 4°C 4 ± 0 +++

a Lesion Score: 1= No lesion, 2 = Mild variation in follicle size, 3= Moderate variation in

follicle size, 4 = Either Necrosis or follicle atrophy

b - = No staining; + = Minimal staining, ++ = Moderate Staining, +++ = Severe staining.

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± = inconclusive.

c Samples fixed at 4°C and pH 7

d Samples fixed at 10 % formalin and pH 7

e Samples fixed at 10% formalin and 4°C

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Table 4.3: Effect of fixation conditions on extraction of RNA from bursas of Fabricius

from chickens infected with IBDV

Variable Treatment

Number of real-time RT-PCR

Positive / total number tested

5 % 5/5

10 % 5/5

20 % 0/5

Formalin

concentration a

37 % 0/5

4°C e 5/5

25°C 5/5

Fixation

temperature b 50°C 0/5

2 0/5

5 0/5

7 5/5

9 0/5

pH

of formalin c

13 1/5

1d 5/5

2d 5/5

3d 5/5

4d 5/5

Duration in

formalin d

14d 5/5

135

Non infected

control

10% Buffered

formalin at 4°C 0/5

Extraction control 10% Buffered

formalin at 4°C 5/5

a Samples fixed at 4°C and pH 7

b Samples fixed at 10 % formalin and pH 7

c Samples fixed at 10% formalin and 4°C

d Samples fixed at 4°C % formalin and pH 7

e.This group was also used to represent 5 days duration in formalin

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Chapter V Development of Infectious Bursal Disease (IBD) Viral protein 2 (VP2) and the

hypervariable region of VP2 in Pichia pastoris for the prevention of IBD in chickens. ___________________ Mohamed M. Hamoud , Pedro Villegas 1To be submitted to Vaccine.

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Abstract Infectious bursal disease virus (IBDV) causes immunosuppression in chickens

and is an infectious disease of global economic importance in poultry. Protection against

IBDV is achieved by vaccination with live or inactivated vaccines. Live viruses cause

some bursal damage while killed vaccines are costly to produce. Therefore, the goal of

this study was to produce a subunit vaccine against IBDV, without actual manipulation of

live or killed viral particles, that protects chicken against IBDV infection. The VP2 gene

from the Edgar strain of IBDV was obtained from frozen bursas, while that for the

hypervariable region of VP2 was obtained from formalin fixed paraffin embedded bursal

tissues. The IBDV VP2 gene and the hypervariable region of VP2 were cloned into

Pichia pastoris and protein production was confirmed by western blot analysis. Specific

pathogen free 1-day old chicks vaccinated with recombinant VP2 protein showed the best

protection against 3 week-old challenge with the Edgar strain of IBDV, as there was no

morbidity or mortality in the VP2 vaccinated birds compared to up to 90% morbidity and

50% mortality in placebo vaccinated birds. However, recombinant vaccines did not

totally protect against viral replication in bursa as confirmed by high histopathological

lesion scores and immunohistochemical detection of IBDV in infected tissue. This was

probably due to the high viral dose and high virulence of the challenge virus used.

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Introduction:

Infectious bursal disease virus (IBDV) is the etiological agent of Gumboro

disease or infectious bursal disease (IBD). IBDV destroys the B lymphocyte precursors

found within the bursa of Fabricius, resulting in bursal atrophy in young chickens [1]. In

unprotected flocks, the virus may cause mortality and / or immunosuppression [2]. There

are two serotypes of IBDV: 1 and 2. All pathogenic viruses belong to serotype 1, whereas

serotype 2 viruses are non-pathogenic for both chickens and turkeys [3],[4]. Very virulent

(vv) IBDV strains emerged in Europe in the late eighties, causing up to 60% mortality

[5], [30], [6]. Strains of vvIBDV have also spread to other continents [7], [8].

IBDV belongs to the family Birnaviridae (genus Avibirnavirus) which includes

viruses with bisegmented dsRNA genomes [9]. Genome segment A has 2 overlapping

open reading frames (ORF). The first ORF of segment A encodes a non structural

protein,VP5 and is possibly involved in virus release [10], [11]. The other encodes a

precursor polyprotein in a large ORF, the product of which is cleaved by autoproteolysis

to yield mature VP2 (outer capsid), VP4 (protease), andVP3 (inner capsid) [12]. Genome

segment B encodes the virus polymerase,VP1 [13]. VP2 is the major host-protective

antigen of IBDV. It contains at least three independent epitopes responsible for the

induction of neutralizing antibody [14], thus the antigenic characterization of IBDV has

been based mainly on the study of the VP2 gene where some amino acid changes have

been shown to be the basis for antigenic variation [15], or have been proposed as putative

markers for vvIBDV [16], [17],[18].

Controlling infectious bursal disease (IBD) and its associated immune

suppression is critical to the poultry industry. This control is achieved by passive

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protection of baby chicks through maternal antibodies that come from breeder hens that

have been given conventional live attenuated and/or inactivated IBD vaccines; or by the

use of live IBD vaccines in broiler chicks, layers or young pullets to provide active

protection against IBDV, or by a combination of both.

The IBDV VP2, expressed in yeast [19], [20], the baculovirus system [21]. [22],

[23], [24], recombinant NDV vaccines [25] or via transgenic Arabidopsis thaliana plants

[26] have been studied for the use as subunit vaccines. An advantage of this technology is

that a vaccine based on VP2 alone should allow monitoring of the field situation by the

discrimination between antibody induced by vaccine (anti-VP2 only) and that induced by

infection (anti-VP2 and VP3) [27]. Genetically attenuated viruses generated by the use of

reverse genetics can also be used for the generation of new vaccines, although

interference of maternal antibodies would still exist. Therefore, as recombinant proteins

are less sensitive to neutralization by anti-IBDV maternal antibodies, recombinant viral

vaccines expressing the VP2 protein, such as fowl pox virus [28], herpesvirus of turkey

(HVT) [29], [30], or fowl adenovirus [31] might be able to mount an active immune

response.

Pichia pastoris is a facultative methylotropic yeast which utilizes methanol. The

methanol metabolic pathway of Pichia pastoris involves a unique set of enzymes. In the

first step of this pathway methanol is oxidized to generate formaldehyde and hydrogen

peroxide which is then decomposed to water and molecular oxygen by catalase. The

oxidation is carried out by two alcohol oxidase genes AOX1 and AOX2 [32]. AOX1 is

the more active alcohol oxidase and may reach as much as 30% of the total protein in the

cell when cultured under growth-limiting rates of methanol. This gene’s promoter is

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utilized for expression of heterologous genes. The expression of heterologous proteins in

Pichia pastoris is fast, simple and inexpensive. Strong aerobic growth allows culturing at

high cell densities. High levels of foreign protein expression have been shown for this

vector [33].

The objective of this study was to produce a recombinant protein or polypeptide from

IBDV genomic material extracted from formalin fixed paraffin embedded tissue blocks

without actual manipulation of live or killed virus particles. Proof of principal of this

technique could provide use of archival tissue as a valuable source of genomic material

for production of proteins or actual viruses using reverse genetics. Here we are aiming for

protein production. Another objective was to produce an efficient recombinant vaccine in

a system that allows mass production.

2. Materials and methods

2.1. Isolation of the virus

The Edgar strain of IBDV was isolated and characterized in our laboratory. The

virus was passed in specific pathogen free chicks several times. The bursa of Fabricius

(bursa) was collected from clinically ill birds then ground and homogenized. The cells

were frozen and thawed three times and centrifuged at 12,000 × g, 4 °C for 10 min. The

supernatant was collected and viral particles were pelleted by centrifugation at 96,400 × g

for 2 h. The pellet was resuspended in Tris- ethylenediaminetetraacetic acid buffer and

kept at -20 °C until use.

2.2. Isolation of IBDV RNA

The Edgar IBD virus was incubated for 3 h in a solution containing 0.01M Tris,

0.01M NaCl, 0.01M EDTA, 0.5% SDS (w/v) and 50 µg/ml proteinase K. IBDV RNA

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was separated by electrophoresis on a 1.0 % agarose gel. The two RNA fragments of the

virus, with sizes of 3.4 and 2.9 kb, were visualized by ethidium bromide staining. The

large fragment was agarose gel purified and extracted by use of gel extraction kit

(QIAquick, Qiagen Sciences, Valencia, CA). This was the source of genomic material for

VP2.

RNA was extracted from formalin-fixed, paraffin-embedded bursas and examined

for IBDV nucleic acid by real-time RT-PCR [34]. Sections totalling 50 µm in thickness

were cut from each formalin-fixed, paraffin embedded tissue block, deparaffinized in

HemoDe (Fisher Scientific, Pittsburgh, PA), washed with 100% ethanol, and digested

with 25 µg/ml proteinase K (Sigma Chemical Co., St. Louis, MO) for 1 hr at 50 °C. RNA

was extracted using Trizol (Life Technologies, Inc., Gaithersburg, MD) according to the

manufacturer’s recommendations, diluted in 25 µl of 90% dimethyl sulfoxide, and frozen

at -80 °C until assayed by RT-PCR. This was the source of genomic material for the

hypervariable region of VP2 called partial VP2 (pVP2).

2.3. Cloning of VP2 and pVP2 in TOP10 cells and GS-115 Pichia pastoris cells.

Synthetic oligonucleotides (Invitrogen, Carlsbad, CA) corresponding to the 5’ and

3’ conserved ends of VP2 which included restriction sites EcoR1 and Not1, respectively,

as well as an ATG start codon on 3’ end and a factor XA cleavage site on the 5’ end were

used for the production of cDNA by RT-PCR. Template dsRNA (5 µl) was heated at

97 °C for 5 min and immediately transferred to ice. RT-PCR master mix was then

prepared as follows: 1 µl (10 µM) of sense and anti-sense primers, 25 µl of 2X Reaction

Mix (a buffer containing 0.4 mM of each dNTP, 3.2 mM MgSO4), and 2 µl SuperScript®

III/ Platinum® Taq mix were added to 16 µl autoclaved distilled water. The template

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RNA and RT-PCR mixture were gently mixed to a final volume of 50 µl. The synthesis

of cDNA was done at 55 °C for 30 min. The PCR scheme was 40 cycles of 94 °C for 15

sec, 55 °C for 30 sec and 68 °C for 100 sec followed by a final extension cycle of 68 °C

for 5 min. The amplified segment was separated on a 1.2% agarose gel and the

appropriate band was excised and purified.

The 2 fragments, corresponding to VP2 and pVP2 of the Edgar strain of IBDV

were cleaved by EcoR1 and Not1 restriction enzymes that were introduced by the

oligonucleotides during synthesis. The isolated fragments were mixed with 1 µl pCR 2.1

vector and according to manufacturer’s recommendation were cloned into TOP10

Escherechia coli cells. After 24 hour growth on Luria-Bertani (LB) plates containing

kanamycin (50µg/ml) white colonies were picked up and grown in 5 ml of LB kanamycin

broth (50 µg/ml) at 37 °C in a shaker incubator set at 225 rpm. Minipreps were performed

using Qiaprep® Spin kit (Qiagen Sciences, Valencia, CA), and cloned inserts were double

digested with EcoR1 and Not1 and visualized on an ethidium bromide gel. Inserts were

then ligated to pPICZ A plasmids provided in EasySelect™ Pichia expression kit

(Invitrogen, Carlsbad, CA) which had also been digested with the EcoR1 and Not1

restriction enzymes. The ligated DNA construct was transformed into Escherechia coli

TOP10 cells and white colonies that grew on low salt LB plates containing zeocin 25 µg/

ml were isolated. Plasmid DNA was extracted and analyzed for the presence of VP2 or

pVP2 using restriction enzymes. Several cDNA clones corresponding to VP2 were

isolated and sequenced. DNA sequences were determined by the dideoxy chain

termination method. A plasmid harbouring VP2 or pVP2 with the correct sequence was

cloned into P. pastoris after point mutations were corrected using a site directed

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mutagenesis kit (Stratagene, La Jolla, CA). Pichia transformation was accomplished by

electroporation of GS115 Pichia pastoris cells pre-treated with 0.1 M lithium acetate and

10mM dithiothreitol [35] using a Gene Pulser Xcell total Electroporation System (Bio-

Rad, Hercules, CA) according manufacturer’s settings for yeast (Saccharomyces

cerevisiae). Briefly, following characterization, the plasmid was linearized to promote

integration into the AOX1 locus, and introduced into GS115 cells; the P. pastoris host

strain, by electroporation transformation.

2.4. Screening for transformed colonies expressing VP2 and pVP2

Putative multi-copy recombinants were selected by plating the transformation mix

on increasing concentrations of Zeocin. Transformed cells were grown on YPDS plates

containing 100, 200, 500, 1000 and 2000 μg/ml Zeocin and incubated at 30 °C for 2 days.

Colonies that grew in presence of higher concentrations of zeocin were selected and then

checked for methanol utilization phenotype by replicate plating of colonies on Minimal

Dextrose with histidine (MDH) agar plates, Minimal Methanol with histidine (MMH)

agar plates. Strains GS115 Albumin (MutS) and GS115/pPICZ/lacZ (Mut+) were also

plated on MMH and MDH plates as controls for MutS and Mut+growth. Further

verification of transformed Mut+ colonies was carried out by PCR analysis of Pichia

integrants (VP2 and pVP2). To do this we developed a simple genomic DNA isolation

protocol, where 5-10 colonies were re-suspended in 50 µl of autoclaved distilled water

and subjected to one freeze thaw cycle by placing the PCR tube containing the colony in

liquid Nitrogen for one minute followed by incubation at 37 °C for 5 min. and then use of

the Qiaprep® Spin miniprep kit according to manufacturer’s recommendation (Qiagen,

Valencia CA).

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Positive colonies were grown in 100 ml minimal glycerol medium in 500 ml

baffled flacks to an OD of 1.5 (usually 1 day). At this point, VP2 production was induced

by adding 500 µl methanol every day. The optimal period of induction was determined

by collecting samples every day after induction started. Following induction, cells were

broken by vortex with glass beads (500 µm), centrifuged, and the supernatant contained

the soluble VP2.

2.5. Western blot analysis

Protein extraction and Western blot analyses were performed using standard

procedures. The membranes were immunoblotted with primary monoclonal antibodies

against IBDV HB9490 (ATCC, Manassas, VA) at 1:500 dilution and secondary antibody

horseradish peroxidase-conjugated goat anti-mouse IgG at a 1:8000 dilution (Jackson

Lab, West Grove, PA).

2.6. Biofermentation

The fermentor was a New Brunswick Bioflo 3000 fermentor (New Brunswick

Scientific Co., Inc. Edison, NJ) with a working volume of 5.0 L (total volume 7.5 L). The

fermentation process was scaled up from shaker flasks to 5.0 L fermentation in three

stages. The main objectives were: (a) achieving a high mass of active yeast cells at the

end of the glycerol-fed-batch stage, (b) adapting to methanol as an inducer as well as a

metabolic and anabolic carbon source, and (c) efficiently preserving the VP2 protein

accumulated during the final stages of the fermentation. For that purpose 1mM

phenylmethanesulfonyl fluoride (PMSF) (Sigma, St. Louis. MO) was added to breaking

buffer when breaking the yeast cells with acid-washed glass beads (Sigma, St. Louis.

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MO). This step was done according to EasySelect™ Pichia Expression Kit Version G

122701 25-0172 (Invitrogen, Valencia, CA).

2.7. Vaccine and challenge study:

Studies in isolation units

Two hundred and seventy specific pathogen free (SPF) light breed chicks were

grown in Horsfal-Bauer isolation units from 1 day of age. They were divided into 54

groups of 5 birds each. Each treatment consisted of 2 groups (replicates) of five birds

each. Experimental design is shown in Table 1. Vaccines used were prepared with or

without complete Freund’s adjuvant (Sigma, St. Louis, MO). Vaccines were administered

by injecting 0.1 ml of vaccine per bird intra-muscularly at one day of age with the

exception of the 2512 vaccinated group that was vaccinated orally with 1 X 103.5 EID50 at

one day of age. Maternal antibody level at 1-day of age was determined by ELISA. Birds

were challenged at 3 weeks of age with oral inoculation of either 0.1 ml breaking buffer

(diluent), 0.1 ml containing 103 EID50 Edgar or 0.1 ml containing 1.2 X 104.5 EID50 of

2512 strains of IBDV, or not challenged at all for non-vaccinated non-challenged

controls. Surviving birds were euthanized one week post challenge.

2.8. Evaluation of vaccine protection against challenge

2.8.1 Sample collection and processing

After vaccinating the birds and placing them in isolation units, daily morbidity

and mortality was recorded. One week post challenge birds were examined, weighed,

bled, euthanatized by cervical dislocation, and necropsied. Bursa, spleen, and thymus

were collected from each bird, weighed, and a portion of each was fixed immediately by

immersion in 10% neutral buffered formalin for 24 hr. Tissues were then processed and

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embedded in paraffin using routine histologic techniques. Relative organ weights were

obtained using the formula [relative organ weight = (organ weight/body weight) × 100]

[36].

2.8.2. ELISA

Blood was collected from the wing vein at 2, and 3 weeks post vaccination and 1

week post challenge. An IBD antibody test kit (IDEXX, FlockChek IBD, Westbrook,

Me) was used to determine antibody titres according to manufacturer’s instruction.

2.8.3 Histopathology.

Paraffin-embedded tissues were sectioned, mounted, stained with hematoxylin

and eosin (H&E) by using an automated stainer (Leica AutoStainer XL, Nussloch,

Germany) , and examined for lesions by light microscopy. All sections were assigned a

lesion severity score. For all tissues, a lesion score of 1 represented no lesions, 2

represented mild variation in follicle size, 3 represented moderate variation in size of

follicles, and 4 represented either necrosis or follicle atrophy [34].

2.8.4 Immunohistochemistry (IHC)

All procedures were done at room temperature. Tissue sections were cut (5 μm)

from paraffin-embedded samples and mounted on charged glass slides (Superfrost/Plus;

Fisher Scientific Pittsburgh, PA, U.S.A.). Paraffin was melted from slides (10 min at

65 °C) and removed by immersion in Hemo-De three times (5 min each). Slides were

then air dried and antigen retrieval performed by digestion with 10% proteinase K

(DAKO, Carpinteria, CA, U.S.A.) for 5 min to expose antigenic target sites. IHC staining

was performed with an automated stainer (Leica ST 5050, Nussloch, Germany) with a

nonbiotin peroxidase kit (DAKO Envision System; DAKO Carpinteria, CA, U.S.A.)

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according to the manufacturer’s recommendations. The primary antibody used was a

mouse monoclonal antibody specific to and cross-reactive for all IBDVs (ATCC no.

HB9490). The secondary antibody was a peroxidase labelled polymer conjugated to anti-

mouse and anti-rabbit immunoglobulins. After IHC staining, sections were

counterstained with hematoxylin, air dried, cover slipped, and examined by light

microscopy. Intensity of IBDV staining in each section was scored as follows:

- = no staining, + = minimal staining (<10% of cells present), ++ = moderate staining

(>10% and <30% of cells present), and +++ = extensive staining (>30% of cells present)

[34].

2.8.5 IBDV RNA evaluation

2.8.5.1 RNA extraction.

RNA was extracted from formalin-fixed, paraffin-embedded bursas and examined

for IBDV nucleic acid by real-time RT-PCR (Pantin-Jackwood and Brown, 2003).

Sections totalling 50 µm in thickness were cut from each formalin-fixed, paraffin

embedded tissue block, deparaffinized in HemoDe (Fisher Scientific, Pittsburgh, PA,

U.S.A.), washed with 100% ethanol, and digested with 25 µg/ml proteinase K (Sigma

Chemical Co.) for 1 hr at 50 °C. RNA was extracted using Trizol (Life Technologies,

Inc., Gaithersburg, MD, U.S.A.) according to the manufacturer’s recommendations,

diluted in 25 µl of 90% dimethyl sulfoxide, and frozen at -80 °C until assayed [34].

2.8.5.2 Real time RT-PCR.

Extracted RNA was denatured at 95 °C for 5 min and kept on ice. An RT-PCR was

performed using reagents from the Light Cycler RNA Amplification SYBR Green I Kit

(ROCHE Molecular Biochemicals, Indianapolis, IN, U.S.A.). The primers used were

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designed to amplify a 400-base pair (bp) segment of the IBDV genome shared by all

strains, which flanks a hypervariable region of the VP2 gene. Amplification and detection

of specific products was also performed using a Light Cycler (ROCHE Molecular

Biochemicals, Indianapolis, IN, U.S.A.) according to the manufacturer’s

recommendations. Briefly, RT was done at 55 °C for 30 min, followed by denaturation at

95 °C for 30 sec. Fifty five PCR cycles were performed, consisting of denaturation

(95 °C for 1 sec), hybridization (55 °C for 10 sec), and extension (72 °C for 13 sec). A

melting curve analysis was done with an initial denaturation at 95 °C. DNA melting was

accomplished with an initial temperature of 65 °C for 10 sec and a gradual temperature

increase of 0.1 °C/sec until 95 °C was reached. The melting temperature of the expected

400-bp amplicon was between 82 °C and 84 °C. This estimated melting temperature was

used to confirm the identity of IBDV specific products obtained using real-time RT-PCR

[34].

2.8.6 Lymphoblastogenesis assay

Lymphocytes used for lymphoblastogenesis assay were isolated from peripheral

blood using Histopaque®-1077 (Sigma-Aldrich, St. Louis, MO) and prepared for plating

according to procedure described by Hassan et al. [37] using cold RPMI 1640 medium

and 10% foetal bovine serum (FBS) (Sigma-Aldrich, St. Louis, MO). The viability of the

lymphocytes was determined by staining with a 0.4% solution of trypan blue. The

concentration of viable lymphocytes was adjusted to 1 x 107 cells/ml by adding RPMI

1640 medium with 10% FBS as required. Tests were performed in 96-well flat-bottomed

tissue culture plates. To each of four wells in the first row, 150 µl RPMI containing 10%

FBS was added (media control). To each of four wells in the second row, 50 µl RPMI

containing 10% FBS and 100 µl lymphocyte suspension (1 x 106 cells) was added (cell

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control). To each of four wells in the third row, 100 µl lymphocyte suspension and 50 µl

of 10 mg/ml Con A (Sigma, St Louis, MI, USA) was added (stimulated cells #1). To each

of four wells in the fourth row, 100 µl lymphocyte suspension and 50 µl of 5 mg/ml

PWM (Sigma) was added (stimulated cells #2). Finally, to each of four wells in the fifth

row, 100 µl lymphocyte suspension and 50 µl of 10 mg/ml IBDV antigen was added. The

plates were covered and incubated at 37 °C in a humidified 5% CO2 atmosphere. The

evaluation of lymphocyte blastogenesis by dye reduction was carried out according to the

method described by Gogal et al. [38]. After 16-18 hr of culture, 20 µl of the Alamar blue

dye was added to each well of one plate and was returned to the incubator. Twenty-four

hours after the dye was added, the absorbance at 570 nm and 600 nm was measured with

a microplate reader. The dye when added is in an oxidized (blue colour) form that is

reduced (red colour) as cells proliferate. The 570 nm absorbance measures the reduced

form and the 600 nm measures the oxidized form. Because there is some degree of

overlap between the two absorbencies, it is necessary to subtract the 600 nm absorbance

from the 570 nm absorbance to obtain the true absorbance (specific absorbance), which

reflects the specific level of proliferation. The specific absorbance of unstimulated cells

(in media alone) was subtracted from the specific absorbance of cells incubated with the

mitogens to yield a ∆-specific absorbance.

The lymphocyte stimulation index was expressed as: Stimulation index (SI) = (C2

– C1) / (C3 – C1), where C1 = concentration of spontaneously reduced Alamar blue in

the medium, C2 = concentration of reduced Alamar blue in the medium of stimulated

cells, and C3 = concentration of reduced Alamar Blue in the medium of unstimulated

control cells [39].

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2.9 Statistical analysis

The body weight gain, relative bursal, spleen and thymus weights were analysed

using analysis of variance and comparisons with a control using Dunnet’s method by use

of JMP 4.0.2 (SAS Institute, Cary, NC).

3. Results 3.1. Cloning of pVP2 and VP2 in yeast

RNA was isolated from an Edgar strain of IBDV. Oligonucleotides corresponding

to both ends of VP2 were designed and synthesized. The open reading frame of VP2 and

the hypervariable region of VP2 were amplified by RT-PCR followed by PCR (Fig. 1).

The 1452 bp and the 510 bp fragments were cloned in E. coli and following amplification

of the plasmid, transfected into GS 115 Pichia pastoris cells.

3.2. Screening for transformed colonies

Transformation of Pichia pastoris without lithium acetate and dithiothreitol was

very poor. Only after adding these two chemicals to GS-115 cells prior to transformation,

were there enough transformed colonies generated for evaluation of transformation

efficacy. The ability of the transformed yeast to grow in presence of zeocin; along with

growth on both MMH and MDH plates followed by performing PCR on yeast genomic

material for the presence of VP2 and pVP2 inserts allowed selection of which colonies to

choose for expression.

3.3. Expression of pVP2 and VP2

VP2 and pVP2 were produced in yeast cells following induction with methanol.

Cells were disrupted by acid-washed glass beads and centrifuged (10,000 rpm, 10 min,

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and 4 °C). The protein was found in the supernatant and analyzed by Western blot. A

band with the molecular weight of VP2 and another of the molecular weight pVP2 were

seen when the membrane was immunoblotted with primary monoclonal antibodies

against IBDV HB9490 at 1:500 dilution and secondary antibody horseradish peroxidase-

conjugated goat anti-mouse IgG at a 1:8000 dilution (Fig. 2).

3.4 Immunization studies – efficacy of VP2 and pVP2 to protect against challenge with

Edgar and 2512 strains of IBDV.

3.4.1. Morbidity and mortality:

Birds vaccinated wild type yeast, diluent or the hypervariable region peptide

started showing depression, ruffled feathers, and trembling 2 days post challenge with the

Edgar IBDV. Severe prostration and death of morbid birds started on day 3 post

challenge (Table 2). Birds challenged with diluent or the 2512 strain of IBDV showed no

symptoms of disease and looked apparently healthy. Birds vaccinated with VP2 and

challenged with the Edgar IBDV were also apparently healthy and showed no signs of

sickness and no mortality was observed (Table 2).

3.4.2. Body and organ relative weights and lesions:

No significant body weight differences were seen between control and

experimental groups for body weight gain (Table 3). There was a significant decrease in

relative bursa of Fabricius weights among the Edgar challenged groups compared to the

2512 vaccinated and non vaccinated non infected controls. Among the birds challenged

with the Edgar strain of IBDV, the VP2 vaccinated birds had the highest bursal relative

weights (Table 3). All birds challenged with the Edgar strain of IBDV suffered from +4

severe subacute bursitis with severe lymphocytic depletion with some exceptions in the

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VP2 and pVP2 vaccinated groups where some birds had acute lesions instead as indicated

by presence of neutrophils in the medulla of the bursal follicles (Table 4). Birds

vaccinated with Pichia control or diluent that died 3-4 days post challenge with Edgar

IBDV had severe bursal haemorrhage (Fig 3). Dead birds in the group vaccinated with

the hypervariable region of VP2 showed oedematous swelling of bursas but had no

haemorrhages. Birds that died on 5th day post challenge with the Edgar IBDV had no

haemorrhages and their bursas appeared to be of normal size. The 2512 strain used for

vaccination and challenge was very mild as the bursas showed very mild or no bursitis

and had relative weights comparable to the non- vaccinated non-infected controls (data

not shown).

The thymus relative weights of birds challenged with the Edgar strain of IBDV,

regardless of vaccine given, were significantly lower in groups not challenged with this

strain of virus, with the exception of the VP2 with no adjuvant vaccinated group. The

VP2 with the adjuvant vaccinated group had significantly higher relative thymus weights

compared to the other Edgar IBDV challenged groups; but was significantly less than

non-vaccinated non infected control group ( Table 3). This was confirmed by the

microscopic observation that some areas of the thymus had moderate to severe depletion

of thymic lymphocytes (Fig. 4).

There was spleenomegaly in birds vaccinated with VP2 or VP2 and challenged

with the Edgar IBD virus (Table 3). In the enlarged spleens, microscopically there were

more lymphocytes and swollen macrophages around the sheathed arteries. There was also

more red blood cells seen in the splenic red pulp that seen in spleens of the non-infected

non-vaccinated control group.

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3.4.2 ELISA

Some of the extra SPF chicks that were purchased were sacrificed and

serologically tested for IBDV. They were seronegative at 1-day of age for antibodies to

IBDV as determined by ELISA. All birds had no detectable antibody titres at 2 weeks

post vaccination. At 3 weeks post vaccination, no antibody titres were found in the

groups vaccinated with Pichia controls, the hypervariable region of VP2, diluent, or non-

vaccinated. The only groups at 3 weeks post vaccination with detectable antibody titres

were those vaccinated with VP2 and they had antibody levels ranging from 59 to 250 for

birds given the vaccine without adjuvant and 113 to 455 for birds given the vaccine with

adjuvant, while the group vaccinated with the live 2512 virus had titres ranging from 684

to 1013. ELISA GMT antibody titres for 4-week old birds, one week post challenge are

presented in Table 5. The low antibody titres, seen one week post challenge with the

Edgar IBDV strain; in the VP2 vaccinated group compared to the Pichia control and

diluent vaccinated groups, suggest that there were antibodies at time of challenge that

neutralized some of the challenge Edgar IBDV, opposed to the Pichia pastoris and the

diluent control vaccine groups that had higher titres one week post challenge suggesting

that there were no antibodies to neutralize the challenge virus.

3.4.3 Immunohistochemistry and real-time RT-PCR detection of IBDV.

The Edgar IBDV challenged birds were all positive for IBDV when using IHC

with variable degrees of staining as shown in Table 5. The bursas of birds vaccinated

with VP2 mixed with Freund’s adjuvant and challenged with Edgar strain had the least

staining intensities, suggesting low viral load in tissues. This was confirmed by the

154

inability to detect any IBDV RNA in genomic material extracted from formalin fixed

paraffin embedded tissue blocks using real-time RT-PCR (the paraffin blocks used for

extraction were the same ones used in making the slides for IHC). The bursas of birds

vaccinated with VP2 without Freund’s adjuvant and challenged with Edgar IBDV had

slightly more intensity than the previously mentioned group but much less when

compared to the Pichia pastoris control and the diluent control groups. They also had

detectable IBDV RNA in genomic material extracted from the same paraffin embedded

tissue blocks (Table 4). The low intensity of IHC staining along with the absence of

detectable IBDV RNA by real-time PCR in some of the VP2 vaccinated groups,

compared to the very intense staining seen in non-vaccinated Edgar IBDV challenged

groups, suggest that some virus neutralization occurred at time of challenge and hence the

lower antibody responses detected. The birds vaccinated with the hypervariable region of

VP2 and challenged with the Edgar IBDV showed similar IHC staining intensities and

real-time RT-PCR positive for IBDV just like the Pichia pastoris control and diluent

control groups when challenged with a similar virus. This suggests that the hypervariable

region of VP2 subunit vaccine did not provided protection against IBDV challenge.

Pichia pastoris and diluent controls groups that were non-challenged with IBDV viruses

had no IHC staining and were negative for real-time RT-PCR detectable IBDV RNA.

Immunohistochemistry could not detect IBDV in all of the 2512 IBDV vaccinated or

challenged birds with only one exception where mild staining of the bursa was observed

in one of the diluent vaccinated 2512 challenged birds. However, real-time RT-PCR

could detect the 2512 virus in the diluent control and non vaccinated 2512 challenged

groups. The 2512 vaccinated and 2512 challenged group had no real-time RT-PCR

155

detectable RNA, no IBDV IHC staining, and +1 lesion scores suggesting that the

antibodies generated by using the live virus completely neutralized the challenge virus.

The VP2 vaccinated group also had no detectable IBDV RNA suggesting that the virus

was neutralized at time of challenge (data not shown).

3.4.4. Lymphoblastogenesis assay

The stimulation indices of the different treatments are presented in Table 6. Lymphocytes

stimulated with the T-cell specific mitogen (Concanavalin A) were not significantly

different from each other with the exception of one group which was vaccinated with the

Pichia control and challenged with Edgar, that was significantly higher than the rest. The

lymphocytes of birds vaccinated with VP2 and adjuvant as well as those vaccinated with

the Pichia control vaccines had a significantly higher response to the B-cell specific

pokeweed mitogen, while non-vaccinated 2515 IBDV challenged birds had significantly

lower response to the IBDV antigen, when compared to other treatment groups.

Comparing the lymphocyte stimulation indices of birds given the vaccines with or

without adjuvant revealed a very slight increase in stimulation indices of the ones that

had been given the vaccine with adjuvant although the difference was not significant.

4. Discussion

Infectious bursal disease virus is highly infectious and very resistant to

inactivation. Thus, despite strict hygienic measures, vaccination is unavoidable under

high infection pressure and it is necessary to protect chickens against infection during the

first weeks after hatch. After hatching, chickens are immunized with live vaccines, and

because maternal immunity interferes with vaccination with live vaccines, the major

problem with active immunization of young maternally immune chicks is determining the

156

proper time of vaccination. It is well known that less attenuated strains (“hot vaccines”)

may cause lesions in the bursa follicles and, thus, immunosuppression even in vaccinated

birds. To avoid live vaccine side effects [40], a Pichia pastoris expression system

expressing IBDV VP2 or the hypervariable region of VP2 has been developed. Pichia

pastoris as a yeast expression system was chosen because it combines the simplicity of

prokaryotic growth requirements, growth potential and ease of manipulation along with

the post-translational modifications of eukaryotes to ensure that the expressed protein is

functional and has the proper conformation. Using this expression system both VP2 and

the hyper variable region of VP2 were expressed. Ethidium bromide gels of RT-PCR

amplicons showed that the primers succeeded in amplifying VP2 and pVP2 of Edgar

IBDV. Colonies screened for ability to grow in presence of high levels of zeocin,

methanol utilization phenotype and presence of genomic inserts were selected for

biofermentation. Intra-cellular expression of VP2 and pVP2 of Edgar in Pichia pastoris

was confirmed by immunoblotting with primary monoclonal antibodies.

Vaccination with recombinant VP2 in combination with Freund’s adjuvant gave

best results in terms of protection parameters. Birds vaccinated with recombinant VP2

mixed in Freund’s adjuvant showed no morbidity or mortality. This was also observed

the VP2 vaccine without adjuvant, but the difference between the two VP2 vaccinated

groups with or without adjuvant could be observed in the intensity of IHC staining and

ability to detect IBDV RNA from infected tissue. Low intensity of IHC staining of bursas

of Fabricius of the VP2 with adjuvant vaccinated group that was challenged with Edgar

IBDV indicates the presence of IBDV in tissue; however, the absence of real-time RT-

PCR detectable RNA suggests low viral load in tissue. This low viral load was still able

157

to cause severe damage to bursal tissue. This can be attributed to the high virulence of the

challenge dose, as the Edgar IBDV used for challenge was serially passed in chickens

several times prior to use in this experiment. Wild type Pichia pastoris, diluent or the

hypervariable region of VP2 vaccinated birds had 30 to 90 % morbidity and 20 to 60 %

mortality; indicating the virulence of the Edgar IBDV strain used. Also, that the yeast and

breaking buffer used to make the VP2 vaccine did not protect against challenge with

Edgar IBDV. Thus, the protection against morbidity and mortality observed in the VP2

vaccinated group is due to vaccination with VP2. This protection maybe due to T-

lymphocytes, as indicated by increase in relative thymus weight (Table 3), and that

cortical depletion was minimal in the VP2 vaccinated group compared to the other

treatment groups that were Edgar IBDV challenged (Fig 4). The effect of IBDV infection

on cell-mediated immune competence is transient and less obvious than its effect on

humoral responses [41]. Some in vitro cellular functions have been found to be

compromised in IBDV-infected chickens, with lymphoid cells of chickens exposed to

IBDV responding poorly to mitogens [42]. While others reported that maximal

suppression of cell-mediated immune responses occurred at 6 weeks post infection [43],

in our study, mitogen stimulation of peripheral blood lymphocytes of vaccinated and non

vaccinated chickens could not verify the role that T-cells have on IBDV vaccination or

challenge, as there were no significant differences between stimulation indices of

lymphocytes of different treatments when stimulated with T-cell mitogen (Concanavilin

A) with the exception of the Pichia control vaccinated group challenged with the Edgar

IBDV. This may be because the Pichia pastoris homogenate acts like an adjuvant and

stimulated the antigen presenting cells in particularly the macrophages to secrete

158

cytokines that stimulate the T-cell mediated immune system. The lymphocytes of birds

vaccinated with VP2 and adjuvant as well as those vaccinated with the Pichia control

vaccines had significantly higher stimulation indices when stimulated with the B-cell

specific pokeweed mitogen, compared to other treatments indicating the role of B

lymphocytes in protecting against the Edgar IBDV. This could also be due to the

previously mentioned probability of innate immune stimulation and the role that

macrophages have in enhancing the humoral immune system. The use of a lower

infective dose or less virulent virus for challenge in the future might be important to help

identify the role of T-lymphocytes in protecting against IBDV infections.

The low geometric mean titres of antibodies against IBDV at 2 and 3 weeks are

not unusual when recombinant vaccines are used. Previous researchers have reported that

titres of 150 – 400 are common and that these low titres are fully protective against

controlled challenge and exposure to IBDV [19]. The antibody titres against IBDV one

week post challenge (Table 4) are typical of those seen in IBDV challenge studies [44],

as non-vaccinated groups had higher GMT titres, than the vaccinated groups. This may

be due to that the protected birds have antibodies that neutralize some of the challenge

virus thus leaving the bird with less virus particles available for infection opposed to the

unprotected birds that get the whole challenge dose. Generally antibodies of recombinant

protein vaccines have lower GMT than that seen with live virus vaccines [19], as live

virus vaccines produce polyclonal antibodies that react to both VP2 and VP3 that is

present in the IDEXX ELISA plates, whereas the VP2 vaccines produce antibodies only

for VP2 and thus get a lower reaction with the IDEXX ELISA plates. Anti-VP3

antibodies do not confer protection against IBDV infection [45]. The low level of anti-

159

VP2 antibodies or the ability to detect VP3 genomic material by RT-PCR can be used to

differentiate between vaccinated and infected flocks.

In summary, recombinant vaccines expressing VP2 and not just the hypervariable

region of VP2 are needed to protect against IBDV infections. The use of paraffin

embedded tissue as a source of genomic material for protein production in yeast is

feasible. However, it highly recommended for conformationally dependant proteins; to

fully reconstruct the whole genome before expressing it in yeast expression system.

Using these recombinant vaccines will help combat and control this disease in the future,

without the side effects seen with live viruses. Pichia pastoris expressed antigenic

proteins can be mass produced at very high levels; to achieve comparable antigenic levels

in inactivated vaccines, sacrificing a lot of SPF chicken embryos will be required.

Specific pathogen free embryos might not be available in the future for veterinary use; as

shortages of available SPF embryos have been reported. This is due to the high demand

for these embryos for the use in production of human therapeutics. A demand that will

increase drastically with the emerging of new human disease epidemics or pandemics that

threatens the human race.

References:

[1] Wyeth PJ, Cullen GA. Maternally derived antibody-effect on susceptibility of

chicks to infectious bursal disease. Avian Pathol 1976(5):253-60.

[2] Sharma J, Kim I, Rautenschlein S, Yeh H. Infectious bursal disease virus of

chickens: pathogenesis and immunosuppression. Dev Comp Immunol.2000;24(2-3):223-

35.

160

[3] Jackwood D, Saif Y, Huges J. Characteristics and serological studies of 2

serotypes of infectious bursal disease virus in turkeys. Avian Dis 1982;26(4):871-82.

[4] McNulty M, Saif Y. Antigenic relationship of non- serotype turkey infectious

bursal disease viruses from the United States and United Kingdom. Avian Dis.

1988;32(2):374-75.

[5] Chettle N, Stuart JC, Wyeth PJ. Outbreak of virulent infectious bursal disease in

East Anglia. Vet Rec 1989;125(10):271-2.

[6] van den Berg T. Acute infectious bursal disease in poultry: a review. Avian Pathol

2000;29(3):175-94.

[7] Banda A, Villegas P. Genetic characterization of very virulent infectious bursal

disease viruses from Latin America. Avian Dis 2004;48(3):540-9.

[8] Hassan MK. Very virulent infectious bursal disease virus in Egypt: epidemiology,

isolation and immunogenicity of classic vaccine. Vet Res Commun 2004;28(4):347-56.

[9] Dobos P, Berthiaume L, Leong JA, Kibenge FS, Muller H, Nicholson B.L.

Family Birnaviridae. In Murphy, FA, Fauquet, CM, Bishop, DHL, Ghabrial, SA, Jarvis,

AW. Martelli, GP, Mayo, MA, Summers, MD. Editors Virus taxonomy. Classification

and Nomenclature of viruses, Sixth Report on the International Committee on Taxonomy

of Viruses. Arch Virol [Suppl 10] Wein New York: Springer, 1995. pp 240-244

[10] Lombardo E, Maraver A, Espinosa I, Fernandez-Arias A, Rodriguez JF. VP5, the

nonstructural polypeptide of infectious bursal disease virus, accumulates within the host

plasma membrane and induces cell lysis. Virology 2000;277(2):345-57.

161

[11] Yao K, Vakharia VN. Induction of apoptosis in vitro by the 17-kDa nonstructural

protein of infectious bursal disease virus: possible role in viral pathogenesis. Virology

2001;285(1):50-8.

[12] Kibenge FS, Dhillon AS, Russell RG. Biochemistry and immunology of

infectious bursal disease virus. J Gen Virol 1988;69 (Pt 8):1757-75.

[13] Macreadie IG, Azad AA. Expression and RNA dependent RNA polymerase

activity of birnavirus VP1 protein in bacteria and yeast. Biochem Mol Biol Int

1993;30(6):1169-78.

[14] Becht H, Muller H, Muller H. Comparative studies on the structural and antigenic

properties of two serotypes of infectious bursal disease virus. J Gen Virol 1998(69):631-

40.

[15] Vakharia VN, He J, Ahamed B, Snyder DB. Molecular basis of antigenic

variation in infectious bursal disease virus. Virus Res 1994;31(2):265-73.

[16] Brandt M, Yao K, Liu M, Heckert RA, Vakharia VN. Molecular determinants of

virulence, cell tropism, and pathogenic phenotype of infectious bursal disease virus. J

Virol 2001;75(24):11974-82.

[17] Mundt E. Tissue culture infectivity of different strains of infectious bursal disease

virus is determined by distinct amino acids in VP2. J Gen Virol 1999;80 (Pt 8):2067-76.

[18] van Loon AA, de Haas N, Zeyda I, Mundt E. Alteration of amino acids in VP2 of

very virulent infectious bursal disease virus results in tissue culture adaptation and

attenuation in chickens. J Gen Virol 2002;83(Pt 1):121-9.

162

[19] Pitcovski J, Gutter B, Gallili G, Goldway M, Perelman B, Gross G, et al.

Development and large-scale use of recombinant VP2 vaccine for the prevention of

infectious bursal disease of chickens. Vaccine 2003;21(32):4736-43.

[20] Wu PC, Su HY, Lee LH, Lin DT, Yen PC, Liu HJ. Secreted expression of the

VP2 protein of very virulent infectious bursal disease virus in the methylotrophic yeast

Pichia pastoris. J Virol Methods 2005;123(2):221-5.

[21] Pitcovski J, Di-Castro D, Shaaltiel Y, Azriel A, Gutter B, Yarkoni E, et al. Insect

cell-derived VP2 of infectious bursal disease virus confers protection against the disease

in chickens. Avian Dis 1996;40(4):753-61.

[22] Dybing JK, Jackwood DJ. Antigenic and immunogenic properties of baculovirus-

expressed infectious bursal disease viral proteins. Avian Dis 1998;42(1):80-91.

[23] Macreadie IG, Vaughan PR, Chapman AJ, McKern NM, Jagadish MN, Heine

HG, et al. Passive protection against infectious bursal disease virus by viral VP2

expressed in yeast. Vaccine 1990;8(6):549-52.

[24] Vakharia VN, Snyder DB, Lutticken D, Mengelwhereat SA, Savage PK, Edwards

GH, et al. Active and Passive Protection against Variant and Classic Infectious Bursal

Disease Virus-Strains Induced by Baculovirus-Expressed Structural Proteins. Vaccine

1994;12(5):452-56.

[25] Huang Z, Elankumaran S, Yunus AS, Samal SK. A recombinant Newcastle

disease virus (NDV) expressing VP2 protein of infectious bursal disease virus (IBDV)

protects against NDV and IBDV. J Virol 2004;78(18):10054-63.

163

[26] Wu H, Singh NK, Locy RD, Scissum-Gunn K, Giambrone JJ. Immunization of

chickens with VP2 protein of infectious bursal disease virus expressed in Arabidopsis

thaliana. Avian Dis 2004;48(3):663-8.

[27] van den Berg TB, Eterradossi N, Toquin D, Meulemans G. Infectious Bursal

Disease (Gumboro Disease). In Diseases of Poultry: World Trade and Public Health

Implications, OIE Scientific and Technical Review 2000:509-43.

[28] Bayliss CD, Peters RW, Cook JK, Reece RL, Howes K, Binns MM, et al. A

recombinant fowlpox virus that expresses the VP2 antigen of infectious bursal disease

virus induces protection against mortality caused by the virus. Arch Virol 1991;120(3-

4):193-205.

[29] Darteil R, Bublot M, Laplace E, Bouquet JF, Audonnet JC, Riviere M.

Herpesvirus of turkey recombinant viruses expressing infectious bursal disease virus

(IBDV) VP2 immunogen induce protection against an IBDV virulent challenge in

chickens. Virology 1995;211(2):481-90.

[30] Tsukamoto K, Kojima C, Komori Y, Tanimura N, Mase M, Yamaguchi S.

Protection of chickens against very virulent infectious bursal disease virus (IBDV) and

Marek's disease virus (MDV) with a recombinant MDV expressing IBDV VP2. Virology

1999;257(2):352-62.

[31] Sheppard M, Werner W, Tsatas E, McCoy R, Prowse S, Johnson M. Fowl

adenovirus recombinant expressing VP2 of infectious bursal disease virus induces

protective immunity against bursal disease. Arch Virol 1998;143(5):915-30.

[32] Cregg JM, Barringer KJ, Hessler AY, Madden KR. Pichia pastoris as a host

system for transformations. Mol Cell Biol 1985;5(12):3376-85.

164

[33] Clare JJ, Rayment FB, Ballantine SP, Sreekrishna K, Romanos MA. High-level

expression of tetanus toxin fragment C in Pichia pastoris strains containing multiple

tandem integrations of the gene. Biotechnology (N Y) 1991;9(5):455-60.

[34] Pantin-Jackwood MJ, Brown TP. Infectious bursal disease virus and

proventriculitis in broiler chickens. Avian Dis. 2003;47(3):681-90.

[35] Wu S, Letchworth GJ. High efficiency transformation by electroporation of

Pichia pastoris pretreated with lithium acetate and dithiothreitol. Biotechniques

2004;36(1):152-4.

[36] Pantin-Jackwood MJ, Brown TP, Kim Y, Huff GR. Proventriculitis in broiler

chickens: effects of immunosuppression. Avian Dis 2004;48(2):300-16.

[37] Hassan M, Afify M, Aly M. Susceptibility of vaccinated and unvaccinated

Egyptian chickens to very virulent infectious bursal disease virus. Avian Pathol.

2002;31(2):149-56.

[38] Gogal RM, Jr., Ahmed SA, Larsen CT. Analysis of avian lymphocyte

proliferation by a new, simple, nonradioactive assay (lympho-pro). Avian Dis

1997;41(3):714-25.

[39] Carpenter CR, Arnold SR, Bose HR, Jr. Suppression of mitogen-stimulated

blastogenic response during reticuloendotheliosis virus-induced tumorigenesis:

investigation into the mechanism of action of the suppressor. J Immunol 1978

(120):1313-20.

[40] Muller H, Islam M, Raue R. Research on infectious bursal disease - the past, the

present and the future. Vet. Microb. 2003;97(1-2):153-65.

165

[41] Lukert, P. D., and Y. M. Saif Infectious bursal disease. In: Diseases of

Poultry, 11th edition ed. Y.M. Saif, H.J. Barnes, J.R. Glisson, A.M. Fadly, L.R. Mc

Dougald, D.E. Swayne. Iowa State University Press, Ames, IA. pp 161-179. 2003.

[42] Sharma JM, Lee LF. Effect of Infectious Bursal Disease on Natural-Killer Cell-

Activity and Mitogenic Response of Chicken Lymphoid-Cells - Role of Adherent Cells in

Cellular Immune Suppression. Infect Immun. 1983;42(2):747-54.

[43] Sivanandan V, Maheswaran SK. Immune profile of infectious bursal disease. III.

Effect of infectious bursal disease virus on the lymphocyte responses to phytomitogens

and on mixed lymphocyte reaction of chickens. Avian Dis 1981;25(1):112-20.

[44] Haddad E, Whitfill C, Avakian A, Ricks C, Andrews P, Thoma J, et al. Efficacy

of a novel infectious bursal disease virus immune complex vaccine in broiler chickens.

Avian Dis. 1997;41(4):882-89.

[45] Pitcovski J, Levi BZ, Maray T, Di-Castro D, Safadi A, Krispel S, et al. Failure of

viral protein 3 of infectious bursal disease virus produced in prokaryotic and eukaryotic

expression systems to protect chickens against the disease. Avian Dis 1999;43(1):8-15.

166

Figure legends: Fig .5.1. RT-PCR product of amplified VP2 and pVP2: lane 1, molecular size marker; lane 2, 3, 5 and 6 amplified VP2; lane 4, negative control; lane 8 and 9, amplified pVP2 Fig. 5.2. Western blot analysis of recombinant proteins. Lane 1 Molecular weight marker, lanes 2 and 3 two clones of yeast expressing pVP2, Lanes 4 and 5 two clones of yeast expressing VP2 and lanes 6 and 7 negative controls (yeast transformed with wild-type plasmid). The membrane was immunoblotted with primary monoclonal antibodies against IBDV (ATCC HB9490) Fig: 5.3. Photomicrographs of bursas from SPF chickens stained with IHC that were vaccinated and challenged with diluent (A); Vaccinated with diluent and challenged with Edgar IBDV (B); Vaccinated with Pichia pastoris control and challenged with Edgar IBDV (C); non vaccinated non infected control (D); pVP2 vaccinated and Edgar IBDV challenged (E) and VP2 vaccinated and Edgar IBDV challenged (F). X100 Fig: 5.4. Photomicrographs of Thymus from SPF chickens stained with H&E were VP2 vaccinated and Edgar IBDV challenged (A); diluent vaccinated and Edgar IBDV challenged (B); Non-vaccinated , non-challenged control (C). X100 Fig: 5.5. Photomicrographs of bursa from SPF chickens stained with H&E that were non-vaccinated , non-challenged control (A). or diluent vaccinated and Edgar IBDV challenged (B); X100

167

Table 5.1: Experimental design of immunization experiment. Different vaccines were administered at 1-day of age with or without Freund’s adjuvant. Challenge was 3 weeks post vaccination. Experimental replicates had the following treatment groups. Vaccine Pichia control VP2 in pichia pVP2 in pichia Diluent 2512 None none

No

adjuvant

+ + + + + + + + + + + + + + +

Adjuvant

+ + + + + + + + + + + + - - -

Challenge diluent Edgar 2512 diluent Edgar 2512 diluent Edgar 2512 diluent Edgar 2512 2512 2512 none

+: Group contained 5 SPF birds -: Not done ( as live viruses are administered with adjuvants in commercial poultry).

168

Table 5.2: Number of morbid and dead birds per total number in different vaccine and challenge treatments

Vaccine Pichia control VP2 in pichia pVP2 in pichia Diluent 2512 none noneChallenge diluent Edgar 2512 diluent Edgar 2512 diluent Edgar 2512 diluent Edgar 2512 2512 none none

No adjuvant

* 3/10 * * * * * 3/10 * * 9/10 * * * * Morbidity

Adjuvant

* 5/10 * * * * * 3/10 * * 5/10 * n/a n/a n/a

No adjuvant

* 2/10 * * * * * 3/10 * * 6/10 * * * * Mortality

Adjuvant

* 4/10 * * * * * 2/10 * * 3/10 * n/a n/a n/a

* = 0/10 n/a = not applicable

169

Table 5.3: Body weight and relative organ weights of 28 day old SPF chickens vaccinated or non-vaccinated against IBDV, one week post challenge. Values ± standard deviations within a column that are followed by the same superscript letter are not significantly different (P < 0.05).

Vaccine Challenge Body weight Bursa relative weight thymus relative weight Spleen relative weight

VP2 Edgar 273.03 ± 38.32 0.409 ± 0.23b 0.479 ± 0.15a 0.337 ± 0.05c

VP2 + adjuvant Edgar 270.29 ± 22.29 0.282 ± 0.21c 0.396 ± 0.07b 0.303 ± 0.08b

pVP2 Edgar 259.35 ± 29.02 0.197 ± 0.16d 0.227 ± 0.08d 0.325 ± 0.06bc

pVP2 + adjuvant Edgar 259.05 ± 25.24 0.177 ± 0.07d 0.262 ± 0.07d 0.315 ± 0.06b

Pichia Edgar 259.89 ± 51.89 0.162 ± 0.04d 0.271 ± 0.03d 0.242 ± 0.04a

Pichia + adjuvant Edgar 274.70 ± 32.26 0.186 ± 0.11d 0.297 ± 0.05c 0.241 ± 0.08a

Diluent Edgar 241.88 ± 57.27 0.230 ± 0.15c 0.306 ± 0.10c 0.299 ± 0.05a

Diluent + adjuvant Edgar 264.78 ± 20.24 0.114 ± 0.03d 0.250 ± 0.07d 0.234 ± 0.05a

Pichia Diluent 316.60 ± 36.37 0.625 ± 0.22a 0.657 ± 0.15a 0.225 ± 0.06a

Pichia + adjuvant Diluent 309.53 ± 40.77 0.545 ± 0.08a 0.541 ± 0.06a 0.225 ± 0.05a

2512 2512 306.23 ± 41.90 0.423 ± 0.09a 0.595 ± 0.13a 0.180 ± 0.06a

none 2515 271.74 ± 21.60 0.567 ± 0.12a 0.429 ± 0.13a 0.225 ± 0.05a

none none 301.90 ± 21.73 0.616 ± 0.09a 0.553 ± 0.12a 0.229 ± 0.04a

170

Table 5.4: Real-time RT-PCR melting curve detection, histopathological lesion scores and IHC mean staining scores for bursas of birds vaccinated or non vaccinated at 1-day of age and challenged with Edgar or 2512 strains of IBDV at 21 days. Sampling was performed 1 week post challenge. Data for individual bird’s bursa.

Vaccine without Freund’s adjuvant Vaccine with Freund’s adjuvant Vaccine Challenge RT-PCR A Lesion

score B IHC C RT-PCR Lesion

score IHC

VP2 Edgar IBDV

+ + + +

4 4N 4N 4N

+ ++ ++ ++

- - - -

4 4N 4 4

+ + + +

pVP2 Edgar IBDV

- + + +

4 4N 4N 4N

+ +++ +++ +++

- + + +

4 4N 4 4

+ +++ +++ +++

Pichia control

Edgar IBDV

+ + + +

4 4 4 4

++ +++ ++

+++

+ + + +

4 4 4 4

+ +++ +++ +++

Diluent Edgar IBDV

+ + + +

4 4 4 4

+++ +++ +++ +++

+ + + +

4 4 4 4

+ +++ +++ +++

Pichia control

diluent - - - -

1 1 1 1

- - - -

- - - -

1 1 1 1

- - - -

2512 2512 - - - -

1 1 2 1

- - - -

N/A N/A N/A

171

none 2512 + + + +

2 2 2 2

- - - -

N/A N/A N/A

Diluent Diluent - - - -

1 2 1 1

- - - -

N/A N/A N/A

none none - - - -

1 1 1 1

- - - -

N/A N/A N/A

A - = Negative; + = Positive

B Lesion Score: 1= No lesion, 2 = Mild variation in follicle size, 3= Moderate variation in follicle size, 4 = Either Necrosis or follicle atrophy, N = acute necrosis C - = No staining; + = Minimal staining, ++ = Moderate Staining, +++ = Intense staining.

172

Table 5.5: ELISA GMT at 28 days of age to IBDV following vaccination of 1-day-old

SPF chickens

Vaccine without

Freund’s adjuvant

Vaccine with Freund’s

adjuvant

Vaccine

IBDV Challenge

strain A IBDV GM titer IBDV GM titer

VP2 Edgar 436 576

pVP2 Edgar 1825 1884

Pichia control Edgar 2540 2679

Diluent vaccine Edgar 2505 2364

2512 2512 884 N/A

None 2512 57 N/A

Pichia control Diluent 22 1

None None 1 N/A

A Chickens were challenged at 21 days of age with IBDV

N/A = not applicable

173

Table 5.6: Mean lymphocyte transformation indices of vaccinated and non-vaccinated birds challenged or not challenged with 2512 or Edgar IBDV

Vaccine Challenge Con A PWM IBDV

VP2 Edgar 1.13± 0.22 a 0.99.± 0.15 a 0.68 ± 0.21 a

VP2+adj Edgar 1.17± 0.26 a 1.30.± 0.43 b 0.71 ± 0.43 a

pVP2 Edgar 0.99± 0.21 a 1.03.± 0.21 a 0.68 ± 0.38 a

pVP2 +adj Edgar 1.01 ±.0.12 a 1.05.± 0.11 a 0.75 ± 0.31 a

Pichia control Edgar 1.25 ± 0.29 b 1.27.± 0.25 b 0.88 ± 0.17 a

2512 2512 1.03 ± 0.01 a 1.00.± 0.02 a 0.76 ± 0.23 a

None 2512 C 1.19 ± 0.37 a 1.09.± 0.22 a 0.32 ± 0.14 b

None none 1.02 ± 0.01 a 0.98 ± 0.02 a 0.78 ± 0.20 a

Values represent the mean of five chicken per treatment; Con A = Concanavalin A; PWM = pokeweed mitogen. IBDV= Infectious bursal disease virus.Values within a column followed by the same superscript letter are not significantly different (P < 0.05).

174

Fig .5.1. RT-PCR product of amplified VP2 and pVP2: lane 1, molecular size marker;

lane 2, 3, 5 and 6 amplified VP2; lane 4, negative control; lane 8 and 9, amplified pVP2

Fig. 5.2. Western blot analysis of recombinant proteins. Lane 1 Molecular weight marker, two clones of yeast expressing pVP2 lanes 2 and 3, two clones of yeast expressing VP2 and negative control, yeast transformed with wild-type plasmid. The membrane was immunoblotted with primary monoclonal antibodies against IBDV (ATCC HB9490)

175

Fig 5.3

A BA B

Fig: 5.3. Photomicrographs of bursa from SPF chickens stained with H&E that were Non-vaccinated, non-challenged control (A); or diluent vaccinated and Edgar IBDV challenged (B); X100

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Fig 5.4:

Fig: 5.4. Photomicrographs of Thymus from SPF chickens stained with H&E were VP2 vaccinated and Edgar IBDV challenged (A); diluent vaccinated and Edgar IBDV challenged (B); Non-vaccinated , non-challenged control (C). X100

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Fig 5.5

Fig 5.5. Photomicrographs of bursas from SPF chickens stained with IHC that were vaccinated and challenged with diluent (A); Vaccinated with diluent and challenged with Edgar IBDV (B); Vaccinated with Pichia pastoris control and challenged with Edgar IBDV (C); non vaccinated non infected control (D); pVP2 vaccinated and Edgar IBDV challenged (E) and VP2 vaccinated and Edgar IBDV challenged (F). X100

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CHAPTER VI

DISCUSSION AND CONCLUSIONS

Identification of infectious bursal disease viruses from RNA extracted from paraffin

embedded tissue.

Formalin fixed paraffin embedded tissues used as a source of genomic material to

rapidly identify field isolates of IBDV by RT-PCR and sequencing allowed a direct

correlation between the virus strain present and the lesions produced by this virus, in a

timely fashion. This also allowed analysis of foreign strains of IBDV with no importation

and biosecurity restriction issues because of the inactivated nature of the antigen.

Therefore, the information obtained by this technology will help refine vaccination

programs against IBDV, as well as to detect new emerging IBDV strains both

domestically and internationally.

Fixation conditions can have a tremendous impact on the success of RNA

extraction, as properly handled samples generate good results while improperly handled

samples can generate average results at best. In chickens experimentally infected with

IBDV, all paraffin embedded blocks with bursas having acute +3 or +4 bursitis had

extractable genomic material that was positive for IBDV based on melting curve analysis,

indicating high sensitivity of the primers used. Most of the 452 blocks analyzed had +3 or

+4 acute lesion scores for IBDV, yet only 227 were positive for IBDV extractable

genomic material. IBDV genomic extraction rate from samples originating from the

U.S.A. was 87%, while that from overseas was only 19%. The effects of different

fixation conditions on tissues with higher than normal viral load was evaluated. These

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changes in temperature, pH and concentration of formalin were enough to render a

positive IBDV formalin fixed paraffin embedded tissue block into a negative one.

Nucleic acid and deduced amino acid analysis of extracted IBDV genomic

material has shown that the Variant E strain of IBDV is the predominant strain present in

the U.S.A. The few classical strains found in samples from the United States were

mainly of vaccine origin. There were no vvIBDV strains found in the U.S.A.

Internationally, four vvIBDV strains from both Venezuela and China were identified.

Variant A strains were found in both Peru and Venezuela. Variant E strains were

identified in El Salvador, Guatemala, Mexico, Peru and Venezuela. Internationally, more

classical than variant strains were identified, this is similar to what Lukert and Saif have

reported.

Phylogenetic analysis showed that serotype 1 viruses grouped together in several

different groups. Previous knowledge of strain origin and identity along with location of

viruses on phylogenetic trees was the basis used for grouping these viruses. All variant

viruses sequenced were either Variant A or Variant E. Variant E had three different

branches.

Several unique virus strains that did not group with the common IBDV strains and

fell on branches of their own were identified. Genomic material was isolated and

identified from a unique IBD virus strain that was missing 63 nucleotides from its

hypervariable region in the VP2 gene, yet the virus was apparently able to replicate in the

bursa and cause +4 acute lymphocytic damage. VP2 is important for virus attachment to

cell receptors and is the viral protein that is recognized by the chicken’s immune system.

Missing these 21 amino acids apparently did not affect the virus’s ability to attach and

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enter the susceptible B-lymphocytes. Constant vaccination pressure in the past may have

contributed to the emergence of variant strains and vvIBDV. This unique case missing

the 21 amino acids may have resulted from random mutations by the virus in an attempt

to escape from the immunoglobulins produced in vaccinated chickens; or maybe this

virus was always there in the population of IBDV viruses, and constant vaccination

against variant IBD viruses suppressed other viruses, giving this virus favorable

propagation conditions.

The inability to identify all strains present in multivalent vaccines was attributed

to the nature of the RT-PCR reaction which amplifies the virus with higher logarithmic

concentration than the other viruses that have lower concentration. So when a sequencing

reaction is run and analyzed, only the nucleotides that are most abundant are identified.

This was noticed when sequencing a bivalent vaccine, SVS-510™, where only the

predominant variant portion of the vaccine was identified and not the classic portion. To

overcome this, the seeds of that vaccine were isolated and sequenced independently. This

variant portion of this bivalent vaccine was identified in figure 1 and 2 as SVS-510

(Variant portion) while the classical portion was identified as SVS-510 (Standard

portion).

To investigate why samples with +4 acute lesion scores of IBDV yielded no

detectable RNA, bursas with high vial loads were subjected to different fixation

conditions. Extracting genomic RNA for identification of IBDV from bursas of Fabricius

stored at acidic pH (pH 5.0 and pH 2.0) was unsuccessful. Previous research which found

DNA damage in cells at low pH, may explain the lack of extracted genomic RNA in this

experiment. Unbuffered formalin oxidizes to formic acid and an acidic environment

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causes degradation of nucleic acids because the β-glycosidic bonds in the purine bases

are hydrolyzed at pH 4. Loss of some amino group cross-links occurs at low pH and may

result in loss of RNA from the fixed tissue. Extraction of genomic RNA at a basic pH

(pH 9.0, and 13.0) was also unsuccessful and the reason for this is unknown.

The thermodynamic stability of a nucleic acid duplex strongly depends on the

denaturation temperature of the duplex at atmospheric pressure. Denaturation of genomic

RNA may have occurred when tissues were formalin fixed at 50°C for 5 days and maybe

the reason why there were no rt-RT-PCR detectable RNA extracted from these blocks.

At formalin concentrations of 20% or 37 %, the extraction of RNA from FFPET

blocks yielded no rt-RT-PCR detectable RNA. This may be due to excessive crosslinking

of amino groups resulting from the acidic nature of formalin at higher concentrations;

indeed, maximum tissue fixation, i.e. cross-linking, occurs in the pH range of 4.0 to 5.5,

thus hindering genomic RNA extraction.

Fixation time had no effect on RNA extraction or subsequent identification , but

longer fixation times are not recommended due to the higher frequency of non-

reproducible sequence alterations. This is demonstrated by the increase in melting peak

temperature over time. Formalin may cause cross-linking of cytosine nucleotides on

either strand. As a result, in PCR the Taq-DNA polymerase fails to recognize the cytosine

and incorporates an adenine in the place of a guanosine, creating an artificial C-T or G-A

mutation.

Real-time RT-PCR as diagnostic tool for IBDV is not always available, so

another important tool for identifying IBDV is immunohistochemistry. We had the

hypothesis that just as fixation conditions have a negative impact on RNA used for

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identifying IBDV in formalin fixed paraffin embedded tissues; these same adverse

fixation conditions may also impact IHC identification negatively. Over the years

formalin fixation of tissues has been used for tissue preservation and for histological

evaluation of tissues. Some have shown that small (10 X 10 X 3 mm) tissue fragments

fixed promptly in neutral buffered formalin for 6-24 hours generally had adequate

cytological preservation and immunolocalization, with minimal antigen masking.

Variations in fixation times or conditions cause the majority of false negatives in

immunohistochemistry, as lack of consistency in formalin fixation protocols (formalin

concentration, pH and fixation time) among laboratories influences the outcome of

staining in immunohistochemistry. Formaldehyde fixatives fix tissue by reacting

primarily with basic amino acids to form cross-linked “methylene bridges”. Formalin

fixation conditions have a direct effect on number of methylene brigdes. The number of

methylene bridges and which amino groups are cross-linked in the tissue are important

variables in the outcome of immunohistochemical staining, and could also be an

important factor on the ability to extract rt-RT-PCR detectable RNA from FFPET.

Tissues fixed in 20% or 37 % formalin undergo shrinkage and may suffer from

alteration of epitope conformation. The IBDV epitopes detected by the monoclonal

antibodies used in this study are conformationally dependent; thus fixation conditions

such as formaldehyde concentration or pH that may cause changes in epitope

conformation rendering them undetectable by these monoclonal antibodies. Maximum

tissue fixation, i.e. cross-linking, occurs in the pH range of 4 to 5.5, therefore, the

presence of too many cross-links or cross-links of certain amino groups may cause

antigen masking. This may be the reason why there was minimal or no immunostaining

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in tissues fixed in a pH <7.0. Inconclusive IHC staining at pH >7.0 may also be due to

tissue shrinkage or alteration of reactive epitopes.

Immunohistochemical staining intensity was highest in tissues fixed at 4 °C.

Increased temperature reduced specific IHC staining, but did not eliminate it. The

reduction in staining intensity may be due to tissue shrinkage at temperatures higher than

4 °C. This result is in agreement with other researchers findings; however the mechanism

for this effect is unknown.

One of the ideas we had when planning for these experiments, was that we could

use the genomic material we extracted from infected bursal tissues for strain

identification, to produce recombinant proteins that provide protection against IBDV in

susceptible flocks. We had the vision of getting field samples from flocks suffering from

immunosuppression; identify IBDV histopathologically and molecularly identify the

strain causing the problem and provide the grower with a tailor made vaccine to control

his IBDV problem. This proof of concept is possible as we were able to identify IBDV

strains by sequencing, and the genomic material extracted from paraffin embedded tissue

was successfully expressed in Pichia pastoris yeast expression system.

Using the information and genomic material gathered from this technology we can

use Pichia pastoris to express various proteins specific for each strain of IBDV

identified; thus, generating a recombinant protein vaccine library that can provide

specific protection against a particular IBDV strains. Once a recombinant IBDV VP2

library is established, very specific vaccines can be provided rapidly to prevent economic

losses caused by IBDV infections. This rapid screening and identification process allows

for processing relatively large number of samples in a very short time. This combined

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with the easy of processing, allows for extensive epidemiological survey to be carried

out. This helps in finding out what types of virus are present in a certain location at a

certain time. This technology has allowed identification of unique viruses, and newly

emerging viruses. Depending on how wide spread these newly emerging viruses become;

we will be able to determine what vaccines are needed to be produced in order to protect

flocks against particular strains of IBDV. Infectious bursal disease virus is highly

infectious and very resistant to inactivation. Therefore, despite strict hygienic measures,

vaccination is unavoidable under high infection pressure and it is necessary to protect

chickens against infection during the first weeks after hatch. After hatching, chickens are

immunized with live vaccines, and because maternal immunity interferes with

vaccination with live vaccines, the major problem with active immunization of young

maternally immune chicks is determining the proper time of vaccination. It is well known

that less attenuated strains (“hot vaccines”) may cause lesions in the bursa follicles and,

thus, immunosuppression even in vaccinated birds. To avoid live vaccine side effects, a

subunit VP2 vaccine was produced in Pichia pastoris that is a yeast expression system. It

combines the simplicity of prokaryotic growth requirements, growth potential and ease of

manipulation along with the post-translational modifications of eukaryotes to ensure that

the expressed protein is functional and has the proper conformation. Using this

expression system we expressed both VP2 and the hyper variable region of VP2. Using

these recombinant vaccines will help combat and control this disease in the future,

without the side effects seen with live viruses. Pichia pastoris expressed antigenic

proteins can be mass produced at very high levels; to achieve comparable antigenic levels

in inactivated killed vaccines, sacrificing a lot of SPF chicken embryos will be required.

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Specific pathogen free embryos might not be available in the future for veterinary use; as

shortages of available SPF embryos have been reported. This is due to the high demand

for these embryos for the use in production of human therapeutics. This demand may

increase drastically with the emerging of new human disease epidemics or pandemics that

threatens the human race.

Extracting genomic material from formalin fixed tissue has been done, unlocking

a historical archive. Expressing these extracted genes and using these generated

sequences to look back will help identify viruses, their biology and their evolution.

Researchers have reconstructed the human influenza virus that caused the 1918 epidemic

and found it to be of avian origin. Using this technology to identify IBDV strains can also

be done and may give an insight on how the virus is changing over time. Also rapid

identification of newly emerging viruses and how predominant they are is important for

veterinarians, to allow them more information in controlling a complex disease.

In summary, the research potentials of extracting RNA from formalin fixed

paraffin embedded tissue has allowed rapid accurate profiling of IBDV strains that are

causing lesions in the field. This is a diagnostic tool that has assisted veterinarians in

choosing which vaccines to use based on the nucleic acid sequence of the hypervariable

region of VP2 of IBDV viruses present on their farms. Another potential is the

identification of potentially new vaccine strains. As with all molecular techniques that are

used to predict the relative similarities and differences between IBDV strains,

determining the actual antigenic differences among viruses requires testing in-vivo. For

optimal IBDV RNA extraction and sequence identification, tissues should be fixed in

formalin at a pH of 7.0, concentration of 5% or 10% and stored at 25 °C or lower

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temperature. Extremes in temperature, pH or formalin concentration have a negative

effect on tissues used for RNA extraction. Optimal conditions for fixation of tissues

infected with IBDV that will be used for IHC evaluation are 10% formalin, pH 7.0 and

temperature of 4 °C. For practical purposes, fixation temperature of 4 °C is recommended

for both procedures. Also recombinant vaccines expressing VP2 and not just the

hypervariable region of VP2 are needed to protect against IBDV infections. Use of

paraffin embedded tissue as a source of genomic material for protein production in yeast

is feasible. But it is highly recommended for conformationally dependant proteins to fully

construct the whole genome before expressing in yeast expression system.