Comparsion of methods used to evaluate infectious Bursal Disease Virus

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Comparsion of methods used to evaluate infectious Bursal Disease Virus
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COMPARISON OF METHODS USED TO EVALUATE INFECTIOUS
BURSAL DISEASE VIRUS



By



JOHN STEVEN ELYAR


























A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE
UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE
REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2005




)>














ACKNOWLEDGMENTS

I would like to express my deepest appreciation to Dr. Gary Butcher, my major

professor, for giving me the opportunity and advice to run this project. In

addition, I would like to gratefully thank Dr. Eric Hessket and Ana Zometa for

invaluable advice on hatching, rearing, challenging, and blood collection, not to

mention valuable assistance during necropsy days.

I want to also thank many of the great co-workers and personal friends, who

made my work seem easier: Dr. Diane Hulse, Dr. Amy Stone, Mr. James

Coleman, MS, Mr. Clifford, and Dr. Francesco Origgi.

In terms of thanks for key reagents, I would also like to thank Dr. Carlos H.

Romero, Dr. Jim Lowenthal, and Dr. Kirk Klassing, along with Intervet, Inc.

Many eternal thanks go to Mrs. Sally O'Connell for her amazing

professionalism and skill in helping me endless times.

My final thanks is for my family, who have strongly and endlessly supported

me in every step of this work: Frank Elyar and Anna K. Elyar.














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TABLE OF CONTENTS

ACKNOWLEDGMENTS................................................................ii

ABSTRACT..............................................................................vii

CHAPTER

1 AVIAN HUMORAL IMMUNITY ....................................................1

Central organs of chicken humoral immunity ......................................1

Peripheral organs of the chicken lymphoid system................................3

The bf: organ ofb-cell expansion and immunoglobulin
diversity..................................................... ........................6

Avian immunoglobulin isotypes and their peripheral roles....................10

Bursal restoration......................................................................11

Ontogeny of chicken b-lymphocytes............................................... 13

Immunocompetence of the embryo and the newly-hatched chick ..............15


Immunopathogenesis caused by infectious chicken viruses.................. 18

Virus effects on chicken antibody repertoire........ .................. .........22

Goals of IBDV studies..............................................................23

2 IBDV LITERATURE REVIEW....................................................26

IBDV introduction.....................................................................26

Evolution Of IBDV .................................................................27

IBDV Antigenic Variation...........................................................28




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Virulent IBDV Infection............. ...............................................30

Variant IBDV Infection........................................................... 32

IBDV Classification.................................................... ..............33

IBDV Molecular Characteristics......................................................35

IBDV Vaccination..................................................................39

IBDV Decontamination........................................................... 44

IBDV Infection........................................................................45

IBDV-Induced Immunosuppression..............................................49

IBDV Replication .....................................................................62

Clinical And Subclinical IBDV Infections...................................... 63

IBDV Pathogenesis...................................................................65

Diagnosis Of IBDV ...................................................................72

3 CHALLENGE OF SPAFAS LAYER AND MATERNALLY IMMUNE
BROILER CHICKENS ON DAY 16 OR 18 WITH VERY VIRULENT IBDV
ALAN LABORATORIES-2 OR DELMARVA VARIANT E
ISOLATES............................................................................75

Introduction...........................................................................75

Project design...........................................................................77

Materials and methods................................................................78

Results.............................................................................80

Discussion ..........................................................................88




4 CHALLENGE OF SPAFAS LAYER AND MATERNALLY IMMUNE
BROILER CHICKENS ON DAY 16 OR 18 WITH A NEWLY ISOLATED
IBDV STRAIN DESIGNATED IBDV-R...........................................111




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Introduction ......... ............... ... ...... .......... ........................... .... 111

Project design ......... ............... ............................... ...............112

Materials and methods .................................. ......................... 112

Results .............................................................................115

Discussion .......................................................................120

5 AVIAN CELLULAR IMMUNITY............................................... 129

Central organs of chicken cellular immunity................................. 129

Oncogenic avian viruses......................................................... 136

6 HISTORY OF MAREK'S DISEASE .............................................141

Introduction...... ..........................................................................141

Biology of the MDV group........................................................149

Virus-cell interaction............. ................................................. 156

Pathogenesis of MDV infection..................................................160

Consequences of infection with MDV ............................................170

Clinical MD ......................................................................... 172

Immune rsponses to MDV infection........................................... 189

Innate immune responses .......................................................... 191

Concluding remarks.............................................................. 191

7 CREATION OF PLASMID DNA CONSTRUCTS ENCODING SEROTYPE-
1 MAREK'S DISEASE VIRUS (MDV-1) AND HERPESVIRUS OF
TURKEY'S (HVT) GLYCOPROTEINS.................................... .......202

Introduction ...........................................................................202

Project design......................................................................204

Materials and methods...............................................................204




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R esults ..................................................................................217

Discussion ........................................................................... 218

8 CYTOKINE LITERATURE REVIEW..........................................235

Introduction.......... .............................................................235

Introduction to interferons......................................................... 239

Biological effects of interferons...................................................247

Molecular stimuli for IFN production.........................................250

General interleukin-2 background ................................................264

11-2 background .....................................................................266

Avian cytokine introduction...........................................................270

9 PLASMID DNA MOLECULES EXPRESSING AVIAN

Cytokines....... .........................................................................273

Introduction................. .............................................................273

Project design...................................... .................................274

Materials and methods............................................................. 274

Results.......... ...........................................................................282

Discussion........ .........................................................................283

References.......... ...........................................................................299

Biographical sketch.....................................................................349












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Abstract of Dissertation Presented to the Graduate School of the University of
Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of
Philosophy

COMPARISON OF METHODS USED TO EVALUATE INFECTIOUS
BURSAL DISEASE VIRUS

BY

John Steven Elyar

May 2005

Chairman: Gary Butcher
Major Department: Veterinary Medicince

This dissertation reports the development and validation of molecular cloning

for in vitro MDV-1 and HVT glycoproteins, along with indirect in vivo

expression in mice with MDV-1 gB and in vivo fate of this construct after 12

post-injection. Additionally, two avian cytokines were cloned and validated. In

vitro expression was shown in mammalian cells. Additionally, this dissertation

investigated whether recommended standards of IBDV pathology methods,

including gross bursa scoring and bursa/body ratio using two standardized variant

IBDV strains: AL-2 and DVE. Similar research was also conducted using a

newly discovered, uncharacterized strain called IBDV-R.
















vii

















CHAPTER 1
AVIAN HUMORAL IMMUNOLOGY

Central Organs of Chicken Humoral Immunity

Introduction

Over years of investigation, the immune system of the chicken has provided an

invaluable model for studying basic immunology. Birds and mammals evolved

from common reptilian ancestors more than 200 million years ago and have

inherited many common immunological systems. However, they also developed

a number of different and, in the case of birds, remarkable strategies. A key

feature of research on the chicken immune system has been the seminal

contributions it has made toward the development of fundamental concepts in

immunology (Davison, 2003).

Graft versus host responses and the key role of lymphocytes in adaptive

immunity were first described in work with chicken embryos and chickens. Most

notably, the bursa of Fabricius provided the first substantive evidence that there

are two major lineages oflymphocytes. Bursal-derived lymphocytes make

antibodies while thymus-derived lymphocytes are involved in cell-mediated

immune responses. Gene conversion, the mechanism used by the chicken to

produce its antibody repertoire, was first described in the chicken and requires the



1





2


(MHC) was the first non-mammalian MHC to be sequenced. Louis Pasteur

developed the first attenuated vaccine against a chicken pathogen, fowl cholera.

In addition, the first vaccine against an infectious cancer agent, Marek's disease

virus (MDV), was developed for the chicken. Lastly, evidence that widespread

and intensive vaccination can lead to increased virulence with some pathogens,

such as MDV and infectious bursal disease virus (IBDV), was first described in

chicken populations (Davison, 2003).

The Bursa of Fabricius

The bursa of Fabricius (BF) mucosa has 11-13 longitudinal folds covered by

specialized follicular epithelium, which forms the raised follicular pad, and

columnar or pseudostratified interfollicular epithelium. The underlying

connective tissue contains 8000-12000 lymphoid (bursal) follicles separated from

each other by delicate connective tissue (Olah and Glick, 1978). Each bursal

follicle has an outer cortex containing densely packed lymphocytes and an inner

medulla, which contains loosely packed lymphocytes and reticular cells. The

cortex is separated from the medulla by a single layer of cuboidal epithelial cells

resting on a basement lamina, which is continuous with the basal cell layer of the

interfollicular epithelium. Small blood vessels are present in the cortex but not

medulla. A diffuse collection of lymphocytes just dorsal to the opening of the

bursal duct contains numerous thymus-dependent T cells, indicating the BF also

functions as a secondary lymphoid organ (Odend'had and Breazile, 1980). Active

bursal duct ligation experiments (Dolfi et al. 1989) provide further evidence of its

secondary role as part of the gut-associated lymphoid tissue (GALT).





3


Peripheral Organs of the Chicken Lymphoid System

Introduction

Secondary lymphoid organs provide the indispensable microenvironment

where the complex interactions among cells, antigens, and cytokines required for

immune responses can occur. Because of the absence of well-developed lymph

nodes in most avian species, including chickens, the chicken spleen has a

dominant role in the generation of immune responses. This seems particularly the

case in the late embryonic and neonatal stage, when lymphoid organs, such as the

cecal tonsils and the Meckel's diverticulum, are not yet present. A typical feature

of chicken spleen is the well-developed ellipsoids, otherwise known as

Schweiger-Seidel sheaths. These ellipsoids consist of a fine network of ellipsoid-

associated reticular cells (EARC) and reticular fibers that surround the penicillary

capillaries and contain macrophages and some lymphocytes. The ellipsoids

together with the surrounding peri-ellipsoid lymphoid sheath (PELS) and

macrophages are considered as the functional analogue of the mammalian

marginal zone (Jeurissen, 1993).

By immunohistochemical staining specific subpopulations of T-cells, B-cells,

macrophage, and EARC were identified early in the development of chicken

spleen (Mast et al. 1998). However, the characteristic structures of the spleen,

such as the PALS and the ellipsoids with their surrounding ring of macrophages,

were only formed around embryonic day (ED) 20. These structures and

especially the microfold (M) cell compartment, i.e., the PELS, gradually matured

during the first week post hatch (Mast and Goddeeris, 1998). This implies,





4


assuming a strong relationship between structural organization and function, that

the immune function of the late embryonic and neonatal spleen may not entirely

be developed.

Spleen

White pulp and red pulp comprise about 80% of splenic tissue (Michael and

Hodges, 1974). They are not sharply distinct from each other in the chicken

spleen. White pulp consists of periarterial sheaths (periarterial lymphoid sheaths,

PALS) surrounding medium and small branches of central splenic arteries that

contain small, T-dependent lymphocytes. Germinal centers (B-dependent tissue)

are often located adjacent to central arteries within these T-dependent sheaths.

Penicillar arterioles at the periphery of the white pulp give rise to capillaries,

which become sheathed with reticular cells forming ellipsoids (Payne, 1979).

These vessels have high endothelial cells, thick basement laminae, and intimate

association with reticular cells. Ellipsoidal cells, peri-ellipsoid B-cell sheaths, and

surrounding macrophages form a complex considered to be the functional

equivalent of the marginal zone in the mammalian spleen (Jeurissen et al. 1992).

Red pulp is a loose spongy tissue with chords of reticular cells located between

venous sinuses that contain lymphocytes, macrophages, granulocytes, and plasma

cells. The relationship of T- and B-dependent areas to blood vessels in the

chicken spleen (Cheville and Beard, 1972), and blood flow from the central artery

through the periarterial lymphoid sheath, the periarteriolar reticular sheath, and

red pulp into the venous sinus of the turkey, which is identical to that in the

chicken, have been described (Cheville and Sato, 1977).





5


Gut-Associated Lymphoid Tissue (GALT)

Cecal tonsils contain dense masses of small lymphocytes and large numbers of

immature and mature plasma cells. Lymphoid tissue with a similar histological

structure to cecal tonsils is also found in the distal region of each cecum about 3

cm from the ileo-cecal junction (del Cacho et al. 1993). Peyer's patches, located

in the small intestinal mucosa, are structurally similar to cecal tonsils. Epithelium

covering Peyer's patches contains numerous lymphocytes, few, if any, goblet

cells, and lacks a continuous basal lamina. Subjacent to the epithelium is a heavy

B-dependent lymphocytic infiltration. A dense core of T-dependent lymphoid

tissue containing B-dependent lymphoid follicles lies deeper in the lamina propria

(Hoshi and Mori, 1973; Befus et al. 1980). Peyer's patches in chickens share

several characteristics with mammalian Peyer's patches including a specialized

lympho-epithelium, presence of M-cells, follicular structure, active particle

uptake, ontogenic development, and age-associated involution. The majority of

intraepithelial lymphocytes in the intestine are T cells (Lawn et al. 1988).

Lymphoid aggregates in the urodeum and proctodeum are also part of the GALT.

Head-Associated Lymphoid Tissue (HALT)

HALT is found in the Harderian (paraocular) and paranasal glands, lachrymal

and lateral nasal ducts, and conjunctival lymphoid tissue (CALT) (Bang and

Bang, 1968; Fix and Arp, 1991). The Harderian gland (HG) has large numbers of

plasma cells in subepithelial connective tissue. Testosterone treatment does not

inhibit HG development, which suggests that this lymphoid organ is relatively BF





6


independent (Kittner and Olah, 1980). Stromal elements of the HG may produce

secretions that influence proliferation and differentiation of plasma cells (Scott et

al. 1993).

Bronchial-Associated Lymphoid Tissue (BALT)

Bronchial epithelium overlying lymphoid tissue is primarily squamous and

non-ciliated at day 1 and week 1, becoming progressively more columnar and

ciliated with age. It does not contain M cells (Fagerland and Arp, 1993).

Occasional lymphoid nodules can be found in the lung as isolated foci not

associated with primary bronchi.

Mural Nodules

Mural lymphoid nodules are closely associated with lymph vessels. They are

circular, elongated, or oval, non-encapsulated, and contain diffuse lymphoid

tissue within which are usually found three or four germinal centers (Biggs, 1957;

Payne, 1979).

The BF: Organ ofB-cell Expansion and Immunoglobulin Diversity

Antibody Genes in Chickens

The cluster of genes encoding the chicken Ig light chain has only a single copy

of the functional variable light (VL) and joining light (JL) genes. Hence, diversity

due to VLJL joining can only be introduced through inaccuracies during the

process of recombination, rather than by selection of different combinations. The

effects of V-J rearrangement on the Ig repertoire are minimal (McCormack et al,

1989b). Like in the Ig heavy chain locus, the presence of a single functional VH

and JH genes means that little diversity can be generated through VnDJn





7


rearrangement. Although there are 16 D genes between the VH and JH regions,

these have very similar sequences, except for one D segment that is not much

used, so these do not introduce much diversity during the rearrangement process

(Reynaud et al. 1989).

However, in both the heavy and light chain Ig loci there are clusters of

pseudogenes upstream of the single functional heavy and light V genes. There are

80 pseudogenes upstream of the function VH gene and 25 pseudogenes in the case

of the VL gene (Reynaud et al. 1987). These pseudogenes (WV) lack leader

sequences but are critical for the generation of chicken antibody diversity.

Following VLJL rearrangement, a process called gene conversion replaces VL

sequences with pseudogene sequences (WVL). Likewise the heavy chain VH

sequences are replaced with WVH sequences. The process occurs after V-J

rearrangement (Weill and Reynaud, 1987) and has been described in some detail

(McCormack and Thompson, 1990).

In summary, an enormous amount of diversity can be generated because (1)

there is substantial diversity in the hypervariable regions of the donor WV genes,

(2) gene conversion events can accumulate within single function VL or VH genes

and (3) different donor WVL or yVH can donate sequence to the respective

functional VL or VH gene (Ratcliffe and Paramithiotis E, 1990). It seems that

birds rely solely on gene conversion for generating an antibody repertoire equal in

an immunocompetent mammal. Interestingly, it has also been observed that gene

conversion is not limited to birds. Gene conversion has also been shown to occur





8


in rabbits (Becker and Knight, 1990) and in sheep, though neither of these species

appears to rely upon gene conversion as the sole means of generating its antibody

repertoires.

Antibody Gene Conversion

The striking fact about gene conversion in the chicken is that it only occurs in

the BF. For instance, if the bursa should be destroyed early in development (60

hours), then those chicks that hatch produce some non-specific IgM but are

unable to mount a specific antibody response when exposed to an antigen.

Therefore, they do not have an antibody repertoire and are incapable of eliciting

typical antibody responses or isotype switching to produce IgG. However, if the

bursa is removed much later in incubation, but before 18 days when the B cells

have begun to migrate from the bursa into the peripheral lymphoid tissues, then

the hatched chickens lack circulating immunoglobulins and likewise are incapable

of eliciting an antibody response (Davison, 2003).

Prebursal B-cell Development

Cells committed to the B-cell lineage, as determined by the presence of surface

B-cell markers, Ig gene rearrangements, and surface immunoglobulin (sIg)

expression, have been identified in extrabursal compartments of the developing

embryo (Benatar et al. 1991; Reynaud et al. 1992) demonstrating that the bursa is

not required for Ig gene rearrangement, although there is the likelihood that the

bursal microenvironment is required for high rate V gene conversion.

In contrast to most mammalian models of B-cell development, the

rearrangement of chicken Ig genes is restricted to a short window of time during





9


embryonic life (until days 8 and 14). These cells have already undergone Ig gene

rearrangement, probably in the embryonic spleen and bone marrow. They express

IgM on the surface (Ratcliffe, 1989). In addition, following DJ rearrangement,

either H or L chain loci complete rearrangement in a random order (Benatar et al.

1992), in contrast to mammalian V genes where H chain rearrangement typically

precedes that of L chain.

In mammals, an appreciable proportion of mature B-cells have both Ig loci

rearranged; only one locus is productively rearranged, resulting in a monospecific

B-cell. In chickens, few mature B-cells contain non-productive V gene

rearrangements, suggesting a distinct mechanism for allelic exclusion (Ratcliffe

and Jacobsen, 1994).

Bursal B-Cell Development

The BF plays a key role in avian B-cell development and antibody

diversification (Paramithiotis et al. 1996). Following colonization by a small

number of B-cell precursors during embryonic life, cells expressing surface

immunoglobulin undergo rapid proliferation, such that by about 2 months of age

there are approximately 10,000 follicles in the bursa, with each containing about

105 B-cells (Olah and Glick, 1978). Seeding B-cells from the bursa into the

periphery begins at about the time of hatch, and continues until the bird has

reached an age of approximately 4-6 months, at which time the bursa begins to

atrophy.





10


Postbursal B-Cell Development

B-cells that have migrated from the bursa to the periphery include those cells,

which have the potential to respond to antigen and subsequently go on to secrete

immunoglobulins (Ig). In addition, the postbursal B-cell compartment includes

the capacity for self-renewal since the bursa undergoes functional involution by

about 6 months of age. Recent data have demonstrated that function B-cell

heterogeneity established in the bursa is reflected in discrete populations ofB-

cells in the periphery, although the physiological basis for this heterogeneity

remains speculative (Paramithiotis and Ratcliffe, 1993, 1996).

Avian Immunoglobulin Isotypes and Their Peripheral Roles

Chicken Immunoglobulin Class M (IgM)

Chicken IgM is the first antibody observed after primary immunization of

chickens and the high molecular weight form of serum can be reduced to heavy

chains and light chains predicting a pentameric structure similar to mammalian

IgM. This notion is supported by the amino acid sequence of the g-chain, which

maintains key amino acids required for pentamer assembly and binding to J-chain,

despite overall homology to the mammalian p of 28-36%. IgM is found on the

surface of most chicken B cells (Kincade and Cooper, 1971) and can transduce

signals to the B-cell cytoplasm (Ratcliffe and Tkalec, 1990).

Chicken Immunoglobulin Class G (IgG)

Chicken IgG is functionally homologous to mammalian IgG in that it

participates in the recall response to antigen. However, analysis of structure and

sequence of chicken IgG has demonstrated that, evolutionarily, it is as similar to





11


mammalian IgE as it is to IgG. This has led to the suggestion that the chicken

molecule is the evolutionary ancestor to both IgE and IgG in mammals (Parvari et

al. 1988).

Chicken Immunoglobulin Class A (IgA)

In mammals, IgA is the primary isotype produced in the mucosal immune

system. In external secretions, IgA exists in a dimeric or tetrameric form of IgA

monomers joined by a J chain, whereas serum IgA is monomeric. Cloning of the

cDNA of Ca from a chicken Harderian cDNA library demonstrated that the Ca

chain is divided into four Ig domain, three of which have 32-41% homology to

human Ca. In mammalian species, a-heavy chains have three Ca Ig domains and

a hinge region between Cal and Ca2. This hinge region may have resulted from

deletions during evolution from a Ca2 Ig domain in the primordial Ca gene, which

has been more conserved in chickens (Mansikka, 1992).

Bursal Restoration in Chickens

Three somatic mechanisms are known to diversify the limited germ-line

repertoire of the chicken immunoglobulin genes: gene hyperconversion (Reynaud

et al. 1987; Weill and Reynaud, 1987; and McCormack et al. 1991), V-J flexible

joining (McCormack et al. 1989a,b) and somatic point mutations (Parvari et al.

1990). Gene hyperconversion, the major generator of antibody diversity in

chickens, starts around 15-17 days of incubation, after immature B-cell

progenitors migrate to the bursa. The bursal microenvironment has been shown

to provide an essential milieu for selecting and amplifying B-cells with productive

antibody gene rearrangement and promoting the antibody repertoire expansion





12


(McCormack et al. 1989b). During the gene hyperconversion process, blocks of

DNA sequence are transferred from pseudo-V regions to the recombined variable

regions of the immunoglobulin genes, resulting in the production of mature B-

cells that are competent to form a functional humoral immune system in the adult

bird (Masteller et al. 1997). These B-cells with diversified immunoglobulin

receptors begin to leave the bursa and populate the secondary lymphoid organs

around the time of hatching; however, the hyperconversion process continues

until the bursa involutes at sexual maturity (Masteller et al. 1995). Damage or

lack of this conversion process induces immunosuppression due to the decreased

diversity of immunoglobulin receptors and the lack of responding B-cell clones

seeded to the peripheral lymphoid tissues.

Severity of the bursal lesions may be varied from transitional to

irreversible depending on the pathogenicity of the virus strains. In cases in which

the damage is reversible, the histological regeneration of the bursa is well

documented and partial or full restoration of the humoral immune functions was

also demonstrated (Edwards et al. 1982 and Kim et al. 1999). However, no direct

evidence has been described concerning the functional restoration of bursal B-cell

activity following the histological regeneration.

There are reports that the duration of immunosuppression and restoration

of the humoral immune response seem to be correlated with the histological

regeneration of the bursa. Edwards et al. (1982) investigated the relationship

between the bursal damage and the depression of humoral immune response to

Brucella abortus in specific pathogen free (SPF) chickens caused by IBDV and





13


suggested that chickens are unlikely to be fully immunocompetent until

approximately 50% of the bursa is fully repopulated. Kim et al. (1999) showed in

SPF chickens that the antibody responses to Newcastle Disease virus (NDV) were

compromised only during the first 6 weeks of IBDV exposure and the recovery of

bursal morphology coincided with the normal levels of antibody. On the other

hand, Giambrone (1979) found permanently depressed immune responses to NDV

in adult, 42-week-old chickens that had been infected early in their life and

suffered irreversible bursal damage. These findings suggest that histological

regeneration of the bursa is necessary for the resumption of a normal antibody

response.

Ontogeny of Chicken B-Lymphocytes

Early Precursor Ontogeny

Early hematopoiesis in the chicken begins in the yolk sac on embryonic day

(ED) 12 and probably plays an important role in embryonic erythropoiesis

(Martin et al. 1978). It is unlikely that cells originating in the yolk sac participate

in the generation of lymphoid precursors. Intra-embryonic hematopoiesis begins

on ED4. Hematopoietic stem cells in the early embryo localize first to intra-aortic

cell clusters and at a later stage to para-aortic mesenchyme, ventrally of the aorta

(Dieterlen-Lievre and Martin, 1981; Cormier and Dieterlen-Lievre, 1988). At

present, the stem cells can only be identified functionally. When transferred into

irradiated hosts, cells from para-aortic mesenchyme are able to generate both B-

and T-cells of the donor type. During normal development, these stem cells seed

the primary lymphoid organs and thus generate the various lymphocyte





14


populations. Bone marrow is also seeded by the stem cells and, after hatching,

becomes a major site ofhematopoiesis. Its role in lymphopoiesis, however, is less

clear in the chicken than in mammals, especially during the embryonic period

(Toivanen and Toivanen, 1973).

Chicken B-lymphocyte Ontogeny

The BF develops as an outgrowth of cloacal epithelium and is seeded between

ED8 and ED14 by stem cells originating in the para-aortic area (Toivanen and

Toivanen, 1973; Houssaint et al. 1976). In irradiated hosts, these stem cells are

capable of reconstituting the entire B cell lineage. These cells can first be found

in embryonic spleen, from which they migrate to bursa and give rise to lymphoid

follicles. The rearrangement ofIg genes occurs before entry into the bursa, e.g.,

in yolk sac, spleen, blood and bone marrow (Ratcliffe et al. 1986; Weill et al.

1986; Mansikka et al. 1990a). Unlike mammalian species, however,

rearrangement in chickens does not generate significant diversity in the Ig genes.

Because the chicken has only one functional V and J gene segment in both the

heavy and light chain locus, each of the B-cells precursors expresses practically

identical immunoglobulins (Reynaud et al. 1985). It has been suggested that

when expressed on cell surface, this prototype immunoglobulin molecule can bind

to a yet unknown self-ligand, triggering proliferation and further differentiation

(Masteller and Thompson, 1994).

The BF has an essential role in B-cell development because it is the site of

immunoglobulin gene diversification and, in bursectomized animals, only

oligoclonal antibodies are observed (Weill et al. 1986; Reynaud et al. 1987;





15


Mansikka et al. 1990b). The precursors entering BF give rise to lymphoid

follicles that start with only a few precursors but after proliferation each contain

approximately 100,000 cells (Pink et al. 1985). Within these follicles, the

developing B-cells undergo gene conversion, a process in which parts of

nonfunctional pseudogenes are copied into the rearranged immunoglobulin gene

(Weill et al. 1986; Reynaud et al. 1987). The heterogeneity of the developing B-

cells within the follicles increases from ED 15 onwards, and almost all of the

immunoglobulin gene diversity in the chicken is due to gene conversion.

Although rearrangement is clearly independent of primary lymphoid organs, gene

conversion takes place only in the BF.

Similar to mammals, the primary lymphoid organs of chickens are sites of

extensive cell death. In the BF, it has been estimated that only 5% of the total cell

numbers survive to form the mature B-cell population (Motyka and Reynolds,

1991). It has been reported the bursal cells undergoing apoptotic cell death down-

modulate the expression of surface immunoglobulin (Paramithiotis et al. 1995). It

is thus probable that one reason leading to cell death is inability to express a

function immunoglobulin. Another possible reason may include expression of

auto-reactive antigen receptor, but details of the B-cell repertoire selection are

poorly understood. The minority of cells which survive start to migrate out of BF

around hatching.

Immunocompetence of the Embryo and the Newly-Hatched Chick





16


Non-Specific Immune Defenses

Although no specific markers for natural killer (NK) cells have been identified

in the chicken, numerous reports state that cells possessing NK-like activity do

exist (Fleischer, 1980; Leibold et al. 1980; Sharma and Okazaki, 1981; Chai and

Lillehoj, 1988). These cells have been isolated from the intestine, BF, spleen,

thymus, and peripheral blood. NK-like activity increases with age and does not

reach adult levels until approximately 6 weeks post hatching, depending upon the

genetic lineage (Lillehoj and Chai, 1988). Yamada and Hayami (1983) reported

that a-fetoprotein in chicken amniotic fluid stimulated suppressor cells which

then reduced NK activity. In another report, injection ofthymulin caused a

reduction in NK activity in chickens that were infected with MDV (Quere et al.

1989). NK cell activity may be in the resistance to MDV (Sharma, 1981), a

disease commonly acquired in the early post hatching period.

Cells of the monocytes-macrophage lineage form early in the development of

the embryo around day 3 (Dieterlen-Lievre, 1989) and exhibit enough function to

respond to some bacterial pathogens during the second week of incubation

(Klasing, 1991). The availability of specific antibody and/or complement can be

a limiting factor in early embryonic macrophage responsiveness, and the rapid

immunologic response to certain pathogens immediately post hatching has been

associated with an increase in complement availability (Powell, 1987; Klasing,

1991).





17


Embryo Vaccination

Chicken embryo vaccination is unique as it is the first widespread commercial

use in any species of prenatal vaccination. The concept was initially devised by

Sharma and Burmester (1982) to protect chicks from virulent MDV exposure that

occurred too early for adequate protection by conventional at-hatch vaccination.

Vaccination of 18-day-old embryos with HVT protected 80-90% of chicks from

challenge to virulent MDV at 3 days post hatching compared with 16-22% of

chicks vaccinated at hatch with HVT. No deleterious effect on hatching was

observed in these trials. Timing of vaccination was critical because embryos

inoculated with HVT prior to ED16 sustained extensive embryonic and

extraembryonic tissue damage (Longenecker et al. 1975; Sharma, 1987). Embryo

vaccination also has the additive benefits of ensuring the precise delivery of

vaccines to each individual and of labor savings via automation of the delivery

system. Experimental vaccination in chickens has been successful for infectious

bronchitis virus (Wakenell and Sharma, 1986) and IBDV (Sharma, 1984) alone or

in combination with HVT, and HE, NDV in turkeys (Ahmad and Sharma, 1993).

The commercial use of embryo-vaccination for protection against MDV (Sharma,

1987) is widespread, with 75-80% of all commercial broilers being vaccinated

embryonically.

Maternal Antibodies

IgM and IgA are located in the amniotic fluid; thus swallowing by the embryo

corresponds to colostrums ingestion in mammals (Kowalczyk et al. 1985),

although minimal transfer occurs. IgG is found in the yolk and begins to be





18


absorbed in the late stages of embryonic development until shortly after hatch

(Powell, 1987). Failure of absorption can affect transfer of maternal immunity

and results in an immunocompromised chick. Chick IgG half-life is

approximately two times that of the adult bird in order to compensate for the time

it takes to fully absorb the yolk. Serum IgA appears at approximately 10 days old

and IgM at 4 days old. The amount of antibody transferred from hen to chick can

vary with the age of the hen and the point of time in lay, and also with the titer

level in the hen's serum. Increasing a hen's serum titer will not necessarily

stimulate a corresponding degree of increase in titer in the embryo (Kowalczyk et

al. 1985). Although maternal antibodies provide variable degrees of protection

against pathologic organisms (Powell, 1987), interference with certain embryonic

or at-hatch vaccines can be substantial. Of particular importance to the

commercial industry is IBDV infection in which vaccination of hens results in

transfer of high levels of maternal antibodies to their progeny (Wyeth, 1975; Naqi

et al. 1983). Although these antibodies are fairly effective in protecting the chick

until approximately 21 days post hatching, interference with initial vaccines will

often completely prevent development of active immunity; therefore, predicting

the timing of IBDV vaccination can be difficult (Solano et al. 1986).

Immunopathogenesis Caused by Infectious Chicken Viruses

Circovirus

Chicken infectious anemia (CIA) is caused by a circovirus. CIA virus is

known to occur worldwide. Only a single serotype has been recognized. The

icosahedral virions contain a circular minus sense single-stranded DNA (ssDNA).





19


Disease occurs in chicks hatched to breeder hens that are infected with CIA virus

after they come into lay; the virus is transmitted vertically. At 2-3 weeks of age,

chicks show anemia, bone marrow aplasia, and atrophy of the thymus, bursa, and

spleen (Lucio et al. 1990; Cloud et al. 1992a). Antibody production remains

unchanged (Goodwin et al. 1992). In vitro proliferation is increased in the spleen,

and decreased in peripheral blood (Cloud et al. 1992b). T-cell function is altered

by a decrease in CTL numbers in the spleen and thymus (Cloud et al. 1992a;

Bounous et al. 1995). Non-specific immunity is also affected by CIA virus

infection. NK cell numbers are reduced in acute infection. Nitric oxide

production is reduced resulting in decreased phagocytosis, bactericidal activity,

and Fc expression (McConnell et al. 1993a,b). Cytokine production is also

affected by infection. IL-1 is reduced throughout infection (McConnell et al.

1993a,b). IL-2 is reduced during acute infection (Adair et al. 1991). Interferon

production is elevated early and decreased late during infection (McConnell et al.

1993a,b). CIA virus infection causes reduced responses to vaccines, including

decreased protection by MDV, NDV, and ILT vaccines (Box et al. 1988; Otaki et

al. 1989; Cloud et al. 1992a). Disease is often most severe in chicks that are

superinfected with CIA virus and other viruses, such as reoviruses and

adenoviruses. Dual infections with immunosuppressive viruses, such as

reticuloendotheliosis virus (REV), virulent MDV, or IBDV, also enhance the

severity of CIA infection, resulting in higher mortalities and more persistent

anemia (Lucio et al. 1990).





20


Retroviruses

The leucosis/sarcoma group of diseases comprises a variety of transmissible

benign and malignant neoplasms of chickens caused by members of a genus

Oncornaviridae of the family Retroviridae. These avian viruses are characterized

as retroviruses by possession of an enzyme, reverse transcriptase (RT). RT

directs the synthesis of proviral DNA from the RNA virus itself (Coffin, 1992).

Economic losses from the leucosis/sarcoma virus group are due to two reasons:

firstly from mortality and secondly from subclinical infection resulting in

decreased egg production and quality (Gavora et al. 1987).

Exogenous, non-defective avian retroviruses cause tumors only in birds that

are congenitally infected and have a persistent viremia. Avian leukosis occurs in

chickens 14 to 30 weeks of age. Clinical signs are nonspecific. The comb may be

pale, shriveled, and occasionally cyanotic. Tumors may be present for some time

before clinical illness is recognized, though with the onset of the first signs the

course may be rapid. Tumors are usually present in the liver, spleen, and bursa

and may occur in other internal organs. Microscopically, the lesions are focal,

multi-centric aggregates of lymphoblasts with B-cell markers. They may secrete

large amount of IgM, but their capacity to differentiate into IgG-, IgA-, or IgE-

producing cells is arrested. The primary target cells are post-stem cells in the

bursa, within which the transformed cells invade blood vessels and metastasize

hematogenously. Bursectomy, even up to 5 months of age, abrogates the

development of lymphoid leucosis (Payne et al. 1991).





21


Hemorrhagic Enteritis Virus

HEV causes splenomagaly. B-cell functions are altered by reduced numbers

due to lytic infection (Suresh and Sharma, 1995, 1996). T-cell functions may be

altered, however research results are variable in terms of T-cell numbers. Suresh

and Sharma (1995) reported that T-cell counts are unchanged. Nagaraja (1982,

1985) reported that T-cell numbers are reduced in acute infection. HEV infection

results in reduced NDV vaccine efficacy (Nagaraja et al. 1985). Secondary

infections also results in increased incidence of colibacillosis (van den Hurk et al.

1994).

Reoviruses

Reovirus infection results in transient bursal atrophy and transient atrophy of

the thymus (Montgomery et al. 1986). Splenomagaly is also reported to occur

during infection (Kerr and Olson, 1969; Tang et al. 1987a,b). B-cell function is

altered due to reduced antibody production (Rosenberger et al. 1989). T-cell

numbers remain unchanged in the spleen but are reduced in peripheral blood

during acute infection (Pertile et al. 1995). In vitro proliferation remains reduced

during acute infection (Rosenberger et al. 1989; Montgomery et al. 1986; Sharma

et al. 1994). Non-specific immunity is affected by increased number of

macrophages in the spleen (Kerr and Olson, 1969; Pertile et al. 1995) and

increased in peripheral blood samples (Sharma et al. 1994). Nitric oxide

production remains unchanged (Pertile et al. 1995). Cytokine production is

altered during infection in many ways. IL-2 is reduced, but normal following

macrophage removal (Pertile et al. 1996). Interferon production is enhanced by





22


attenuated virus (Ellis et al. 1983a,b). Reovirus infection results in reduced MDV

vaccine efficacy (Rosenberger et al. 1989).

Herpesviruses

Marek's disease virus (MDV) is a pathogenic alpha herpesvirus of chickens

(Biggs et al. 1965; Churchill et al. 1969). MDV is a cell-associated virus with

lymphotropic properties similar to gamma herpesviruses (Buckmaster et al. 1988).

MDV is the prototype virus of pathogenic chicken herpesvirus and is designated

as serotype-1. Serotype-2 herpesviruses are apathogenic in chickens and

serotype-3 herpesviruses are pathogenic in turkeys (HVT) (Biggs et al. 1972,

Kawamura et al. 1969). Infection, pathogenesis, pathology, and disease signs are

discussed much further in Chapter 6.

Birnavirus

Immunosuppression due to IBDV infection is described in detail in Chapter 2.

Viral Effects on Chicken Antibody Repertoire

Generating the antibody repertoire in a burst of activity in the young animal is

not without risk. Any virus that targets and destroys bursal cells will have a

devastating effect on antibody-dependent immune responses. One such virus,

IBDV, results of infecting neonatal chicks causes no clinical disease but destroys

B-cells in the bursal follicles leaving the chick incapable of mounting an antibody

response to other viruses, although paradoxically there is a good response to

IBDV itself. This insidious virus leaves the chick vulnerable to opportunistic

infections, and unprotected by subsequent vaccinations. So relying on the





23


generation of the antibody repertoire in a single location over a relatively short

time span is not without its hazards and represents one of the risky immunological

strategies that birds have adopted (Payne et al. 1991)

Goals of IBDV Studies

This research project was designed to answer questions of significance to the

poultry industry in regards to subclinical IBD in the US and worldwide. Prior

attempts to directly correlate IBDV-specific antibody levels with protection have

often not provided consistent results. A standard recommendation by the

"Infectious Bursal Disease Manual" states the best way to evaluate subclinical

classic and variant infection is by bursa/body (B/B) weight ratio. As discussed in

Chapter 2, in the US, all IBDV field isolates are of classic or variant classification

with the variant strains predominating. However, excluding the US, worldwide

IBDV infections are classified as virulent or very virulent. Bursas rarely become

edematous and inflamed after variant IBDV infection; whereas, bursas usually

become edematous after IBDV infection with virulent and very virulent classical

strains. This period of edema is followed by bursal atrophy. Currently, the

USDA recommends the use of bursa/body weight ratios to evaluate subclinical

classic or variant IBDV infection. However, it is believed that this system does

not include the potential for subclinical classical IBDV infection or variant IBDV

infection where bursal edema may occur in some cases. Therefore, this system

can only accurately determine whether birds are experiencing subclinical





24


infection after the earlier stage of edema. Thus, the bursa/body weight ratios

would need to be evaluated at more than 8 days following infection rather than the

4 or 5 days, as is often done now.

As previously discussed, current broiler breeder vaccination programs, which

include a variant in the inactivated product, are capable of protection to chicks via

maternal antibody against variant IBDV. It was proposed that newly emerging

variant IBDV might be able to escape maternal antibody-mediated protection. In

addition to evaluating two characterized IBDV strains, AL-2 and Delaware

Variant E, a newly isolated IBDV strain, termed IBDV R was also studied. This

trial would also permit an evaluation of current recommendations from the

USDA, including B/B weight ratio and gross bursa scoring and determine if they

are an accurate measurement of protection against subclinical variant IBDV

infection.

In addition, results of this project have potential application to broader issues

in the poultry industry. For example, 1) allow for more accurate assessment for

subclinical effects of IBDV challenge with newly emerging variants, (2) increase

accuracy of measuring subclinical effects of infection and may aid in better

evaluation of IBDV virulence, thereby (3) potentially aid research in the

development of new IBDV vaccines.

By creating more effective IBDV vaccines, the significance of clinical and

subclinical flock infections may be decreased. This means less costs due to

better-feed conversion, body weights, egg production, etc. It would also decrease

the incidence ofimmunosuppression, morbidity, and mortality. Chicks would





25


have adequate immune responses in reference to other microbes, thereby reducing

the potential of opportunistic infections and vaccine interference. Finally, this

could in turn, spark new interest in re-evaluating current clinical and subclinical

infections with other avian pathogens.













CHAPTER 2
IBDV LITERATURE REVIEW


IBDV Introduction

In the Delmarva Peninsula of Delaware, infectious bursal disease (IBD) was

initially recognized as a syndrome of chickens in 1957 and Cosgrove

subsequently identified IBDV as the causal agent in broiler flocks (1962). The

viral etiology of infectious pancreatic necrosis (IPNV) of fish was recognized in

1960. In 1973, it was noted that IBDV and IPNV had similar and distinctive

morphologies. Their assignment to a new viral family was initiated by the

recognition in the late 1970s that the genome of each virus consisted of two pieces

of dsRNA (Mueller et al. 1979) with unique biophysical characteristics (Dobos et

al. 1979), but it was not until 1984 that the family was officially designated

Birnaviridae (Dobos, 1979). IBDV is classified in the Avibirnavirus genus within

the family Birnaviridae (Murphy and Johnson et al. 1995). Other members of

Birnaviridae include IPNV as an Aquabirnavirus, tellina and oyster viruses of

mollusks, and Drosophila X virus of fruit flies (Drosophilia melanogaster) as an

Entromobirnavirus. The taxonomic relationship of other Birnaviruses and IBDV

is based on morphology, dsRNA, and similarity of capsid proteins as denoted by

analytical untracentrifugation and polyacrylamide gel electrophoresis (Dobos et

al. 1979). Furthermore, morphological and physiochemical similarities between



26





27


IPNV and IBDV, especially regarding polypeptide profiles and electrophoretic

mobility of RNA segments, have also been indicated (Todd and McNulty et al.

1979).

Interestingly, it was hypothesized that the initial outbreaks of IBDV in the US

arose from mutation of an Aquabimavirus, such as IPNV, capable of infecting

marine species, including menhaden (Brevoortia tyrannus). This fish was

commonly used to manufacture fishmeal and was incorporated into broiler diets

fed in the Delmarva area during the 1950s and 1960s (Lasher et al. 1997). It was

suggested that the relatively heat-resistant aquabirnavirus may have survived

incomplete processing of meal. A subsequent study by Dobos et al. (1979)

confirmed a close relationship only between the three aquabirnaviruses (IPNV,

tellinavirus, and oyster virus of mulluscs), which can be differentiated from

IBDV, and Drosophila virus of fruit flies. This conclusion was based on cross-

neutralization tests and tryptic peptide analysis of 125I-labeled viral proteins. This

evidence tends to disfavor an etiological relationship between IBDV and IPNV.

However, the true ancestor of IBDV has not yet been identified. The possibility

of introduction of IBDV from an insect reservoir should also be considered

(Howie and Thorsen, 1981).

Evolution of IBDV

In addition, from its original identification in 1962, IBDV has evolved from

relatively mild to highly virulent pathotypes and to antigenic variants. In 1983,

diagnosticians in the Delmarva area documented an increase in plant downgrades

despite the use of conventional live IBD vaccines. A multidisciplinary team





28


initiated extensive investigations involving a review of flock records, serological

data, and pathology of affected flocks operated by nine integrators in three

contiguous states. Sentinel chickens immunized against conventional type 1

IBDV were placed among problem flocks for 9-day periods during the growing

cycle. These birds yielded four variants of type 1 virus (Rosenberger and Gelb,

1978, Rosenberger and Cloud, 1989). Changes in the epitope of the variant

Serotype 1 viruses from the conventional strain were demonstrated by Snyder et

al. (1988) applying monoclonal antibody analysis. Yamaguchi et al.

demonstrated that rather than a genetic recombination event; a genetic re-

assortment might play an important role in the emergence of highly virulent

IBDV (1997).

IBDV Antigenic Variation

Antigenic diversity among IBDV isolates has been recognized since 1981,

when serotypes 1 and 2 were defined on the basis of their lack of in vitro cross-

neutralization (McFerran et al. 1980). Further antigenic differences have been

demonstrated with serotype 1 since 1984, and the study of North American IBDV

isolates causing little mortality but marked immunosuppression (Rosenberger and

Gelb, 1976) has led to dividing serotype 1 into six subtypes, which were

originally differentiated by cross neutralization assays using polyclonal sera

(Jackwood and Saif, 1987). Studies based on monoclonal antibodies subsequently

demonstrated a growing number of modified neutralizing epitopes in the more

recent serotype 1 isolates from the US (Snyder et al. 1992), which were

designated as "variant" IBDV. It was, hence, suggested the North American





29


classic IBDV isolates might have been affected by an antigenic drift resulting in

variant IBDV strains (Snyder et al. 1988). The continual shifts in antigenic

components within field IBDV populations may lead to the emergence of new

variants and strains with enhanced virulence or which have altered host or tissue

specificity (Van der Berg et al. 1990). Intensive vaccination in some areas of the

US may have influenced antigenic properties of field IBDV. The epidemiology

ofIBDV in the US has been defined since then by the natural occurrence of

variant virus able to escape the effects of neutralizing monoclonal antibodies

(Snyder et al. 1992).

The structural basis for such antigenic variations has been traced to a

hypervariable antigenic domain on VP2, which is highly conformation dependent

and elicits virus-neutralizing (VN) antibodies (Azad et al. 1985). This

hypervariable region has been recently shown to include two highly hydrophilic

amino-acid domains (212-224 and 314-324) (Schnitzler et al. 1993 and Vakharia

et al. 1994). Amino-acid changes in one or both regions lead respectively to the

emergence either of an antigenically variant serotype 1 strain (Heine et al. 1991;

Jackwood and Jackwood, 1994;Schnitzler et al. 1993), or of a new serotype

(Schnitzler et al. 1993). Recently, variant serotype 1 IBDV strains have been

isolated from vaccinated flocks on Delaware's Delmarva Peninsula (Rodriguez-

Chavez et al. 2002). These strains are able to infect vaccinated chickens in the

presence of high antibody levels against IBDV (Rodriguez-Chavez et al. 2002).

Further antigenic variation was discovered in the Delmarva region a few years

later (Snyder et al. 1988).





30


Virulent IBDV Infection

Since the mid-1980s, highly pathogenic IBDV strains designated very virulent

(vvIBDV) have been reported in many European, African, and Asian countries.

The emergence of vvBDVs significantly increased the economic impact of the

disease. In France, mortality rates up to 60% were described in 1989 in broiler

and pullet flocks, despite vaccination practices (Eterradossi et al. 1992).

Mortality rates from 30% to 70% in SPF chickens were reported in Japan

(Nunoya et al. 1992). The vvIBDV strains were reported to break through high

levels of maternal antibodies in commercial flocks, causing from 60% to 100%

mortality in chickens and producing lesions typical of IBDV (Cho and Edgar,

1969). Such newly emerging strains were characterized as serotype 1 viruses but

were shown to cause IBD in the presence of high levels of antibodies that were

protective against classic serotype 1 strains (Cho et al. 1969; Chettle and Wyeth,

1989; Van der Berg et al. 1990).

To date, vvIBDV has yet to be reported from North America or Australia.

Contrary to the situation in the US with variant IBDVs, the vvIBDV European

strains were reported to be antigenically similar to other serotype 1 classic strains

but very different in virulence (Van der Berg et al. 2000). Additionally, a

vvIBDV isolate from the UK was characterized by Chettle and Wyeth (1989),

who confirmed that spontaneous enhancement of virulence had occurred without

any major alteration in antigenic structure. Recently, several sequence studies

were conducted to identify the molecular basis of antigenicity and differences in

genomic segments coding the major protective epitopes of vvBDVs (Brown et al.





31


1994; Brown and Skinner, 1996; Lin, et al. 1993; Vakharia et al. 1994). Studies

with Japanese vvIBDVs indicated that they were different from all conventional

classic and variant strains of IBDV studied (Lin, et al. 1993). In a comparison of

a limited number of IBDVs, some nucleotide sequence differences were

correlated with virulence (Lin et al. 1993; Nakamura et al. 1994). Ture et al.

(1993) characterized five wIBDV isolates by RT-PCR and RFLP techniques and

compared with the US Serotype 1 (classic and variant) and serotype 2 viruses.

When the PCR products treated with restriction fragment length polymorphism

(RFLP), similarities and differences from the American classic and variant

Serotype 1 strains were shown, and some common digestion products were

unique for vvIBDVs. However, variations in RFLP patterns are not necessarily

an indication of antigenic variation or immunogenicity of variant and wIBDV.

Such variation must be determined by in vitro cross-neutralization assays and in

vivo experimental challenge. With the appearance of these highly virulent and

variant IBDVs and the guarantee of newer strains evolving in the future, the

possibility of current vaccination protocols becoming obsolete is a major concern

for the poultry industry.

The marked increase in the number of recorded acute IBD cases since 1988 in

several European countries (Chettle and Wyeth, 1989, Eterradossi et al. 1992;

Van der Berg TP et al. 1990) has raised the question of a possible similar

antigenic evolution of European vvIBDV strains. The recent wIBDV isolates

obtained in Europe have been shown to be significantly more pathogenic than the

Faragher 52/70 strain (Eterradossi et al. 1992; Van der Berg et al. 1990), which is





32


widely used as the European reference for pathogenic serotype 1 strains. In spite

of their enhanced pathogenicity, these vvIBDVs are considered to be still closely

antigenically related to the reference strain on the basis of high in vitro cross-

neutralization indices (Eterradossi et al. 1992) and of the lack of antigenic

differences in studies based on monoclonal antibodies (Van der Berg et al. 2000;

Van der Marel et al. 1990). Sequence determinations seem so far to support such

antigenic analysis since the amino-acid changes that have been evidenced in the

VP2 hypervariable region of the vvIBDVs have not been demonstrated to clearly

influence antigenicity (Brown et al. 1994; Lin et al. 1993).

As vvIBDVs are not adapted to cell-culture, their antigenic characterization

has mainly been performed in assays, such as antigen capture studies (Van der

Marel et al. 1990). Using neutralizing monoclonal antibodies that had been

previously developed to characterize US variant IBDV, Van der Marel et al.

studied 12 European isolates of IBDV, four of which were from France (1990): no

important antigenic differences could be noted among strain F52/70 and the

recent European isolates.

Variant IBDV Infection

The continual shifts in antigenic components within field IBDV populations

may lead to the emergence of new variants and strains with enhanced virulence or

which have altered host or tissue specificity (Van der Berg et al. 1990). Intensive

vaccination in some areas of the US may have influenced antigenic properties of





33


field IBDV. European viruses responsible for vvIBDV, in contrast, have

increased pathogenicity without demonstrating antigenic shifts (Snyder et al.

1988).

IBDV Classification

McFerran et al. were the first to report antigenic variations among IBDV

isolates of European origin (1978). They presented evidence for the presence of

two serotypes designated 1 and 2, and showed only 30% relatedness between

several strains of serotype 1 and the designated prototype of that serotype.

Similar results were observed in the US, and the American serotypes were

designated II and I. Later studies indicated the relatedness of the European and

American isolates of the second serotype and use of the Arabic numeral 1 and 2 to

describe the two serotypes of IBDV was proposed. Antigenic relatedness of only

33% between two strains of serotype 2 was reported, indicating an antigenic

diversity similar to that of serotype 1 viruses. The two IBDV serotypes can be

differentiated by virus neutralization tests.

The first isolates of serotype-2 originated from turkeys and it was thought that

this serotype was host specific. However, later studies showed that viruses of

serotype-2 could be isolated from chickens, and antibodies to serotype-2 IBDVs

are common in both chickens and turkeys. Chickens are the only avian species

known to be susceptible to clinical disease and characteristic lesions caused by

IBDV. Turkeys, ducks and ostriches are susceptible to infection with IBDV but

are resistant to clinical disease (Giambrone et al. 1978). In addition,

immunization against serotype-2 does not protect against serotype-1. The reverse





34


situation cannot be tested because there are no virulent serotype-2 viruses

available for challenge. Therefore, all viruses capable of causing disease in

chickens belong to serotype-1; serotype-2 viruses may infect chickens and turkeys

and are non-pathogenic for both species.

Variant and vvIBDV isolates of serotype-1 were previously described.

Vaccine strains available at the time they were isolated did not protect against the

variants, which were antigenically different from the standard serotype-1 isolates.

Jackwood and Saif (1987) conducted a cross-neutralization study of 8 serotype-1

commercial vaccine strains, 5 serotype-1 field strains, and 2 serotype-2 field

strains. Six subtypes were studied. Van der Marel et al. using monoclonal

antibodies suggested that a major antigenic shift in serotype-1 viruses had

occurred in the field (1990).

Several techniques have been developed in order to molecularly characterize

and analyze variant strains of IBDV. By using a reverse transcriptase/polymerase

chain reaction (RT-PCR), cDNA from several variant strains have been

characterized by restriction fragment length polymorphisms (RFLP) (Jackwood

and Nielsen, 1997). These techniques can provide profiles based on small

variations of DNA. More specifically, nucleotide sequencing can be performed

on cDNA produced by RT-PCR of IBDV RNA. However, it should be noted that

changes in viral RNA sequences do not necessarily mean changes in antigenicity.

Another technique is to characterize IBDV variants by reactivity with a panel of

neutralizing monoclonal antibodies (Vakharia et al. 1994). In addition, ELISAs





35


have been developed utilizing the VP2 protein antigen, which proved invaluable

in predicting the percentage of protection against classic or variant IBDV strains

in vaccinated flocks (Jackwood et al, 1999).

IBDV Molecular Characteristics

IBD is caused by non-enveloped virions classified as members of the family

Birnaviridae (Montgomery et al. 1986). The genome consists of two linear,

double-stranded RNA. The virions contain no lipid. The double stranded RNA

genome is comprised of two segments: Segment A that is approximately 3.4

kilobases (kB), and Segment B which is approximately 2.9 kb. The larger open

reading frame codes a long polypeptide represented as N-VPX-VP4-VP3-C (Azad

et al. 1985). The precursor polyprotein is processed by a series of post-

translational proteolytic cleavage steps to yield mature virion proteins, most of

which are non-glycosylated (Hudson et al. 1986, Azad et al. 1985). VPX is

further processed by VP4, the viral protease, to produce VP2.

VP2 is considered to be the major host-protective immunogen, and at least two

neutralizing epitopes were found to be located on this peptide (Azad et al. 1985;

Becht et al. 1988, Fahey et al. 1989). Antibodies to these epitopes were found to

passively protect chickens. VP2 determines serotype specificity and is

responsible for eliciting protective antibody, the epitopes being highly

conformation-dependent. VP2 is the only viral encoded protein that may be

glycosylated. Therefore, VP2 is of major interest in the development of new

vaccines against IBDV. Since VP2 is credited with eliciting protective immunity

in chickens, much effort has been directed toward using VP2 as a vaccine.





36


Subunit vaccines containing VP2 and live recombinant vectored viral vaccines

containing VP2 insert alone or in combination with other viral polypeptides have

been developed (Bayliss et al. 1991; Fahey et al. 1989; Vakharia et al. 1994).

Most of these vaccines elicit a significant anti-IBDV antibody response with

variable, often sub-optimal, levels of protection against challenge with virulent

IBDV.

Vaccines containing live replicating or inactivated IBDV continue to be the

best choice for immunizing commercial flocks. A number of such vaccines are

available in the market. The major immunodominant epitopes responsible for

eliciting host protective antibodies against IBDV have been mapped to a 145-aa

polypeptide that is located within the major virus capsid protein VP2 (Fahey et al.

1989; Heine et al. 1991). This region is comprised of a central core of

hydrophobic amino acid residues flanked on either end by two hydrophilic

regions. Mutations in this hypervariable coding region are thought to be

responsible for the evolution of antigenically variant and virulent serotype 1 virus

strains (Oppling et al. 1991; Vakharia et al. 1994). RFLP profiles ofRT-PCR

products of the hypervariable region of wild-type IBDV strains suggest there is a

relatively high degree of genetic heterogeneity in the hypervariable region of VP2

(Jackwood and Sommer, 1998). Nucleotide sequences determined for the

hypervariable-coding region of recent field isolates of IBDV suggest that there is

continuing evolution of the conformational epitopes formed by this polypeptide

(Cao et al. 1998; Dormitorio et al. 1997; Islam et al. 2001). These data suggest





37


the hypervariable region of VP2 may have a high mutation rate that could affect

the antigenicity and pathogenicity of viruses passaged for laboratory studies and

vaccine preparations.

Birnaviruses are cytolytic viruses, but the molecular mechanism(s) employed

for virus egress is, as yet, unknown. Most of the knowledge on virus release

mechanisms derives from studies on enveloped viruses that bud from the plasma

membrane (Garoffet al. 1998). In contrast, non-enveloped viruses have long

been thought to be released following cell lysis. It is thought that either the viral

gene expression or the formation and accumulation of virus particles induce

changes in membrane permeability, eventually leading to cell lysis. However,

data from different virus-cell systems suggest that expression of a single viral

protein may be responsible for cell lysis (Carrascoet al. 1996). Several such

proteins have been identified, i.e., the 2B proteins of poliovirus and

coxsachievirus, the rotavirus NSP4, and the adenovirus E3-11.6K. Death proteins

have been implicated in the alteration and eventual disruption of the host cell

plasma membrane permeability (Aldabe et al. 1996; Tollefson et al. 1996, van

Kuppeveld et al. 1997).

VP5 is a protein whose sequence overlaps that ofVP2. Immunofluorescence

analyses showed that upon expression VP5 accumulates within the plasma

membrane. Expression of VP5 was shown to be highly cytotoxic. Induction of

VP5 expression resulted in the alteration of cell morphology, the disruption of the

plasma membrane, and a drastic reduction of cell viability. Blocking its transport

to the membrane with Brefeldin A prevented vP5-induced cytoxicity. These







results suggest that VP5 plays an important role in the release of the IBDV

progeny from infected cells (Lombardo et al. 2000). However, results using a

virus mutant lacking VP5 were replication competent in cell culture, which

suggests the VP5 is not required for productive replication of IBDV (Mundt et al.

1997).

VP4 is the viral protease which is responsible for self-processing of the

polyprotein, but the exact locations of the cleavage sites are unknown (Azad et al.

1987, Jagadish et al. 1988). VP3 is a minor structural protein. The smaller

segment B encodes a single gene product VP1 that is presumed to be the viral

RNA polymerase. The presence of the VP1-VP3 complex in IBDV-infected cells

was confirmed by co-immunoprecipitation studies. Kinetic analyses showed that

the complex of VP1 and VP3 is formed in the cytoplasm and eventually is

released into the cell-culture medium, indicating that VP1-VP3 complexes are

present in mature virions. In IBDV-infected cells, VP1 was present in two forms

of 90 and 95kDa. Whereas, VP3 initially interacted with both the 90 and 95kDa

proteins, later it interacted exclusively with the 95kDa protein both in infected

cells and in the culture supernatant. These results suggest that the VP1-VP3

complex is involved in replication and packaging of the IBDV genome.

The dsRNA of the IBDV genome has two segments, as shown by

polyacrylamide gel electrophoresis. Jackwood and Jackwood (1994) and Becht et

al. reported that the two segments of five serotype-1 viruses migrated similarly

when co-electrophoresed. The RNA segments from serotype-2 viruses migrated





39


similarly but were different from serotype-1 IBDV when co-electrophoresed,

suggesting that RNA migration patterns could be used to differentiate IBDV

isolates that differ serotypically.

IBDV Vaccination

IBDV causes considerable economic loss in the poultry industry by inducing

severe clinical signs, high mortality (50%), and immunosuppression in chickens

because bursal B-cells are targets for IBDV infection resulting in B-cell depletion.

Most IBD has been controlled by live IBDV vaccines based on strains of

intermediate virulence (Ismail and Saif, 1991). However, it is difficult to protect

field chickens with maternal antibodies induced by live IBDV vaccination (Ismail

and Saif, 1991; Tsukamoto et al. 1995b). In addition, live vaccines induce

moderate bursal atrophy (Muskett and Reed, 1985). Currently, the disease is

prevented by application of an inactivated vaccine in breeder chicken flocks, after

chickens are primed with attenuated live IBDV vaccine. This has kept economic

losses caused by IBD to a minimum.

Vaccination of poultry flocks, especially parent flocks, is often performed with

the intention of protecting the progeny via maternal antibodies during the first

weeks of life. The efficacy of this vaccination schedule, more precisely described

as induction of indirect protection, is normally proven by challenging the chickens

(Jungback and Finkler, 1996). IBDV-specific antibodies transmitted from the

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

resultant protection against the immunosuppressive effects of the virus. Maternal





40


antibody will normally protect chicks against infection for 1-3 weeks, including

the boosting of immunity in breeder flocks with oil-adjuvanted inactivated

vaccines.

The major problem with active immunization of young maternally immune

chicks is determining the proper time of vaccination. This varies with levels of

maternal antibody, route of vaccination, and virulence of the vaccine virus.

Environmental stressors and management may be factors to consider when

developing a vaccination program that will be effective for a flock. Results

suggest that serological determination of the optimum vaccination time for each

flock is required to effectively control highly virulent IBDV in the field. The

optimum vaccination timing could be approximated by titration of the maternal

IBDV antibodies of day-old chicks by ELISA (Tsukamoto, et al. 1995b). ELISA

has the advantage of being a rapid test with the results easily entered into

computer software programs. With these programs, one can establish an antibody

profile on breeder flocks that will indicate the flock immunity level and provide

information for developing proper immunization programs for both breeder flocks

and their progeny.

Another potential problem with active immunization exists due to antibody

interference. A negative feedback loop of antibodies on B-cells can be explained

in molecular terms. B-cells posses FcyRII receptors capable of binding

antibodies. If these antibodies bind to an antigen that is also bound to a B-cell

receptor, the two receptors become cross-linked. This cross-linking draws the

two receptors close together. As a result of this aggregation and lack of co-





41


stimulation, their signal transduction molecules interact and a critical tyrosine

residue is phosphorylated, preventing calcium influx and thus cellular activation.

Thus, this pathway is a feedback mechanism capable of controlling B-cell

responses, whereby B-cell activation is also regulated by antibody but prevents

uncontrolled B-cell responses. Therefore, the presence of maternal antibody in a

newborn chick effectively delays the onset ofimmunoglobulin synthesis through

this negative feedback mechanism. IgM appears about 5 days following exposure

to a disease organism and will disappear in 10-12 days. IgG is detectable 5 days

following exposure, peaks at 21-25 days and then slowly decreases. Thus, if

chick-produced antibody titers are needed, one should collect sera after 21-25

days. This creates difficulty in interpreting vaccination programs. IgA appears 5

days following exposure and peaks similar to IgG (Homer et al. 1992).

Serum samples from day-old chicks contained maternal anti-IBDV antibodies,

which declined to undetectable levels by four weeks of age (Armstrong, et al.

1981). Inducing maternal antibody in progeny from vaccinated breeders prevents

early infection with IBDV and diminishes problems associated with

immunosuppression. The level of IBDV-specific maternal antibody in the

circulation of day-old layer strain chickens was found to be on average, 45% of

the antibody titer in their respective dam, while the minimum ELISA titer which

protected against a challenge of 1000CIDso of virus was 1:400. Note that this

reported titer was determined in a homologous classic IBDV challenge system.

Maternal antibody was found to disappear from the circulation of these crossbred

chickens with a half-life of 6.7 days (Fahey et al, 1987). Furthermore, attenuated





42


live vaccines have been used successfully in commercial chicken flocks after

maternal antibody fades (Nakamura et al, 1993). Thus, a newer generation of

IBDV vaccines, safer and more efficacious, must be studied.

There are many choices of live vaccines available, based on virulence and

antigenic diversity. The most virulent vaccine has been discontinued in the

marketplace. Presently available in the US are IBDV vaccines of intermediate

virulence and high attenuation, including some cell culture-adapted variant

strains. The full impact of the use of variant strain vaccines is still being studied.

Highly virulent, intermediate, and avirulent strains have been shown to break

through maternal VN antibody titers of 1:500, 1:250, and less than 1:100,

respectively. Intermediate strains vary in their virulence and can induce bursal

atrophy and immunosuppression in day-old and 3 week old SPF chickens. If

maternal VN antibody titers are less than 1:1000, chicks may be vaccinated by

injection with avirulent strains of virus. The vaccine virus replicates in the

thymus, spleen, and bursa where it persists for 2 weeks. Once the maternal

antibody is catabolized, there is an ensuing primary antibody response to the

persisting vaccine virus.

Oil-adjuvant, killed-virus vaccines are commonly used to boost and prolong

immunity in breeder flocks, but they are not practical or desirable for inducing a

primary response in young chickens. Oil-adjuvant vaccines are most effective in

chickens that have been primed with live virus, either in the form of active

vaccination or field exposure to the IBDV. Oil-adjuvant vaccines presently may





43


contain both standard and variant strains of IBDV. Antibody profiling of breeder

flocks is advised to assess effectiveness of vaccination and persistence of

antibody titers.

A universal vaccination program cannot be offered because of the variability in

maternal immunity, and existing operational conditions. If very high levels of

maternal antibody are achieved and the field challenge is reduced, then

vaccination of broilers may not be needed. Vaccination timing with attenuated

and intermediate vaccines varies from as early as 1 to 3 weeks. If broilers are

vaccinated at 1 day of age, the IBDV vaccine can be given by injection. Priming

of breeder replacement chickens may be necessary and many producers vaccinate

with live vaccine at 10-14 weeks of age. Killed oil-adjuvant vaccines are

commonly administered at 16-18 weeks. Revaccination of breeders may be

required if antibody profiling should indicate the need.

In addition to conventional vaccines, there are several viral vector systems,

including retrovirus, poxvirus, herpesvirus, adenovirus, and adeno-associated

virus, which are useful for gene therapy and recombinant vaccines, as well as for

in vitro expression systems. Research on viral vector-based polyvalent vaccines

is especially important in the poultry industry to control several important

infectious diseases. Three live vaccine-based viral vectors of chickens lead this

research field: (1) MDV (Parcells et al. 1994), (2) HVT (Morgan et al. 1992), and

(3) fowlpox virus (Bayliss et al. 1991). Both MDV and HVT vectors are

developed for the induction of long-term protective immunity in chickens because

both vectors are herpesviruses, whereas the FPV vector is used to quickly induce





44


protective immunity in chickens. Such viral vector-based recombinant vaccines

are safe for chickens and have no risk of producing antigenic/pathogenic variants

because of subunit-type vaccines. Their low vaccine efficacy, however, is

currently a hindrance for practical use in the field when compared to commercial

live vaccines (Ismail and Saif 1991; Tsukamoto et al. 1995b). For example, viral

vector-based recombinant vaccines poorly protected against the formation of

gross bursa lesions when challenged with vvIBDV, although they provided

protection against the development of clinical signs and mortality (Bayliss et al.

1991; Darteil et al. 1995; Tsukamoto et al. 1999b). Further studies are required,

not only to develop safe and highly efficacious recombinant vaccines, but also to

know how to use them effectively.

IBDV Decontamination

Studies have indicated that IBDV is very stable. Benton et al. found that

IBDV resisted treatment with ether and chloroform, was inactivated at pH 12 but

unaffected by pH 2, and was still viable after 5 hours at 560C. The virus was

unaffected by exposure to 0.5% phenol and 0.125% thimerosal for 1 hour at 300C.

There was a marked reduction in virus infectivity when exposed to 0.5% formalin

for 6 hours. The virus was also treated with various concentrations of three

disinfectants (an iodine complex, a phenolic derivative, a quaternary ammonium

compound) for a period of 2 minutes at 230C. Only the iodine complex had any

deleterious effects. Landgrafet al. found that the virus survived 600C but not

700C for 30 minutes (1967). Certainly, the hardy nature of this virus is one

reason for its long, persistent survival in poultry houses even when thorough





45


cleaning and disinfection procedures are followed. In addition, the agent is

relatively refractory to ultraviolet irradiation and photodynamic inactivation

consistent with dsRNA viruses. Virus can remain viable for up to 60 days in

poultry house litter (Petek et al. 1973).

IBDV is a highly contagious viral infection of breeder, broiler, and layer

chickens. Contamination of chickens by IBDV generally takes place on farms.

Hence, the use of disinfectants is necessary. However, there is a problem related

to IBDV having strong resistance toward the effects of most disinfectants. Only

chlorines and aldehyde-containing disinfectants are effect against IBDV. Because

chlorines have oxidizing action that results in metal corrosion, repeated

disinfection of chicken houses with chlorines should be avoided. The aldehydes,

especially formaldehyde, commonly used to disinfect chicken houses, have been

evaluated in the US by the Department of Labor for their harmful effects on the

human body. No disinfectant effective against IBDV and safe to the human body

is presently available (Shirai, et al. 1994).

IBDV Infection

In vivo and in vitro studies have shown that the target cell for IBDV is an IgM-

bearing B-cell (Ivanyi and Morris, 1976). Within hours of exposure, IBDV-

containing cells appear in the bursa and the virus spreads rapidly through the

bursal follicles. Virulent serotype 1 strains of IBDV have a selective tropism for

chicken B-cells and cause marked necrosis of lymphoid follicles within the bursa.

Virus replication leads to extensive lymphoid cell destruction in the medullary

and the cortical regions of the follicles (Tanimura and Sharma, 1998). Previous





46


reports showed that a virulent strain of IBDV was propagated in B-cells bearing

surface IgM (sIgM), which exist in the bursa (Hirai and Calnek, 1979; Nakai and

Hirai, 1981). However, IBDV infection in susceptible host cells has not been

completely studied at the level of virus attachment. In addition, the cellular

receptor, which bound IBDV in the course of the infection, has not been

identified.

The first step in virus infection is the attachment of IBDV to a specific

receptor on the surface of susceptible host cells. The distribution of a virus

receptor is a major determinant of the cell and tissue tropism of the virus (Bass

and Greenberg, 1992; Haywood, 1994) and the site of pathology associated with

infection (Racaniello, 1990; Ubol and Griffin, 1991). Therefore, it is important to

study the virus infection at the level of virus binding for understanding the virus-

host cell interactions and pathogenesis of the virus disease. Additionally, such

interaction can be exploited for the development of effective IBDV vaccines.

Ogawa M et al. (1998) used a flow cytometric virus-binding assay that directly

visualizes the binding of IBDV to its target cells in a study. A chicken B-

lymphoblastoid cell line, highly permissive for IBDV infection, bound

significantly high levels of the virus. Another B-lymphoblastoid cell line bound

low levels of the virus, although such cells were non-permissive to IBDV

infection. No virus binding was detected in non-permissive T-lymphoblastoid

cell lines. In the binding assay to heterogeneous cell populations of chicken

lymphocytes, IBDV bound to 94% cells in the lymphocytes prepared from the

bursa, 37% cells in those prepared from the spleen, 3% cells in those prepared





47


from the thymus, and 21% cells in those prepared from peripheral blood. Most of

the cells, which bound the virus, were lymphocytes bearing sIgM. Additionally,

binding of IBDV to permissive B-cells was affected by treatment of the cells with

proteases and N-glycosylation inhibitors. These findings may indicate that IBDV

host range is mainly controlled by the presence of a virus receptor composed of

N-glycosylated protein associated. Such viral protein appears to be associated

with the subtle differentiation stage of B-lymphocyte maturation, represented

mostly by sIgM-bearing cells.

Most of the virus-binding cells observed in the study by Ogawa et al. (1998)

were sIgM-bearing B-cells. Results from another study also reported that virulent

IBDV infected sIgM-bearing B cells, however, infection was inhibited by anti-

IgM antibody (Hirai and Calnek, 1979). These findings suggested that an IBDV

receptor was specifically present on the sIgM-bearing B-lymphocyte, indicating

the possibility that the sIgM molecule may be the receptor for virulent IBDV

attachment. However, in this study, bound virus particles were also observed in

the sIgM-negative cells even though their number was small. Interestingly, the

binding of the virus to sIgM-bearing B-cells was not inhibited by anti-IgM

antibody. These additional findings may show that IBDV does not solely utilize

the sIgM molecule as the virus receptor.

The function of the viral receptor used by IBDV to infect the target cells

appears to depend on a molecule associated with the subtle differentiation stages

of B-lymphocytes represented mostly by sIgM-positive cells. Previously, several

B cell-specific surface antigens other than sIgM were reported (Olson and Ewert,





48


1990), which appeared to closely parallel the expression of the sIgM molecule on

the bursal lymphocytes. The possibility should be considered that these

molecules might have served as IBDV receptors or even co-receptors.

Furthermore, a virulent IBDV infection was observed only in cell lines of sIgM-

bearing cells, but not in cell lines of sIgM-negative cells (Hirai and Calnek, 1979).

This result may indicate that the sIgM molecule is important for processes that

occur after virus attachment, such as penetration, un-coating, etc.

Recently, Mueller et al. (1986) reported that IBDV bound to proteins with

molecular masses of 40 and 45kDa expressed on chicken embryo fibroblast cells

(CEFs) and chicken lymphocytes by using various overlay protein blotting assay.

However, it is unclear whether virulent IBDV also bound to the same molecules

because CEF cell-adapted strains of IBDV were used in this study. Generally,

field isolates (virulent strains) of IBDV propagate in chicken lymphocytes but not

in CEFs (Lukert and Davis, 1974). With successive in vitro passages, however,

the virus becomes progressively adapted to growth in CEFs (Izawa et al. 1978;

Yamaguchi et al. 1996). Adaptation of IBDV by serial passage in CEFs

presumably results from selection of variants that are better adapted for

replication in CEFs and, conversely, less well adapted for replication in their

natural hosts. For these reasons, the virus overlay protein-blotting assay, by using

virulent IBDV and permissive B-cells, may be needed to determine the cellular

receptor for the virulent IBDV infection in vivo. Furthermore, virus-binding

assays using the virulent IBDV and CEFs may be needed to determine whether

CEFs have the receptor molecules for the virulent IBDV.





49


Within hours of exposure, virus-containing cells appear in the bursa and the

virus spreads rapidly through the bursal follicles. Virus replication leads to

extensive lymphoid cell destruction in the medullary and the cortical regions of

the follicles (Tanimura and Sharma, 1997). The cellular destructive process may

be accentuated by apoptosis of virus-free bystander cells (Tanimura and Sharma,

1998). Although there is no detectable reduction in circulating immunoglobulins

(Giambrone et al. 1977; Kim et al. 1999), the acute lytic phase of the virus is

associated with a reduction in circulating sIgM-bearing B-cells (Hirai et al. 1981;

Rodenberg et al. 1994).

Although the thymus undergoes marked atrophy and extensive apoptosis of

thymic B-cells during the acute phase of virus infection, there is no evidence that

the virus actually replicates in thymic T-cells (Tanimura and Sharma, 1998;

Sharma et al. 1989). T-cells have been shown to be resistant to infection with

IBDV (Hirai and Calnek, 1979). Furthermore, gross and microscopic lesions in

the thymus are quickly overcome and the thymus returns to its normal states

within a few days of virus infection.

IBDV-induced Immunosuppression

General Immunosuppression

Variation ofB cells bearing surface immunoglobulins M and G (sIgM and

sIgG) was studied in the spleen and peripheral blood of chickens infected with

IBDV. The proportion of surface immunoglobulin-bearing B-cells and sIgM and

sIgG-bearing B-cells in chickens infected at one day of age decreased from week

one post infection (pi) onward and was significantly lower at 8 weeks pi (Hirai, et





50


al. 1981). Chicks infected with IBDV had normal levels of serum IgM and IgG,

but significantly lower levels of IgA when compared to uninfected control birds

(Giambrone, et al. 1977). Passages of live IBDV vaccines in chickens have been

shown to increase the virulence (Muskett et al. 1985).

Chickens infected with IBDV develop reduced humoral and cellular immune

responses and respond poorly to routinely used vaccines. Although T-cells do not

serve as targets for IBDV replication (Tanimura and Sharma, 1998; Sharma et al.

1989), cellular responses of virus-exposed birds are compromised (Confer et al.

1981; Kim et al. 1998). T-cell mitogenic responses of peripheral blood

lymphocytes and splenocytes are significantly reduced following IBDV infection

(Sivanandan and Maheswaran, 1980). IBDV-infected chickens become deficient

in the production of optimum levels of antibodies against diverse antigens,

partially because of the destruction of B-cells (Ivanyi and Morris, 1976;

Giambrone et al. 1977). Interestingly, IBDV has been shown to reduce only

primary antibody responses; secondary antibody responses are spared (Hirai et al.

1981; Rodenberg et al. 1994; Kim et al. 1999). Notably, most studies on the

effect of IBDV on humoral immunity have been limited to the first 5-7 weeks of

virus exposure. However, the depression of antibody titers against diverse

antigens following IBDV inoculation suggests compromise of both local and

systemic immune function, a finding of importance to the broiler industry (Dohms

and Jaeger, 1988).

It is possible that IBDV-induced T-cell immunity will enhance viral lesions.

For example, cytotoxic T-lymphocytes (CTL) may exasperate virus-induced





51


cellular destruction by lysing cells expressing viral antigens. T-cells may also

promote the production of inflammatory factors that may accentuate tissue

destruction. Nitric oxide (NO) produced by macrophages activated by T-cell

cytokines (e.g. IFN-y) may promote cellular destruction.

The combined effects of IBDV-induced immature B-cell lysis and T-cell

impairment results in immunosuppressive effects most pronounced if virus

exposure occurs within the first 2-3 weeks following hatch (Allan et al. 1972). In

commercial chicken flocks, immunosuppression may be clinically manifested in a

number of ways. In general, the flock performance is reduced. Specifically,

immunosuppressed flocks tend to experience an increased incidence of secondary

infections, poor feed conversion, reduced protective response to commonly used

vaccines, and an increased rate of carcass condemnation at the processing plant.

Immunosuppression may accompany overt clinical or subclinical outbreaks of

IBDV. Commercial chicken flocks commonly experience recurring losses due to

IBDV-induced immunosuppression despite widely used vaccination programs.

Specifically, exposure to IBDV impairs the response to vaccines administered

after IBD-induced immunosuppression. Increased susceptibility to respiratory

viruses, including Newcastle disease (Faragher et al. 1974) and avian infectious

bronchitis (Pejkovski et al. 1979), leads to depression in egg laying strain flocks.

Immunosuppressed breeder flocks may undergo a decline in egg production and

hatchability following exposure to viral pathogens. Reduced egg numbers and

hatchability diminish chick yield per breeder. Performance of broiler progeny

from immunosuppressed parent flocks is adversely affected due to relatively low





52


maternal antibody transfer (Lucio and Hitchner, 1979). Infection with IBDV may

exert a profound impact on the profitability of an integrated broiler operation by

reducing efficiency and return from both parent and commercial generations.

The first published description of the immunosuppressive effect of IBDV in

the chicken demonstrated a diminished antibody response to Newcastle disease

vaccination (Faragher et al. 1972). The immunosuppressive properties of IBDV

were quantified in chicks vaccinated against Newcastle disease using attenuated

and inactivated products at various ages (Hirai et al. 1974). These authors

showed that immunosuppression was more severe in 6-week-old SPF chickens

than in 4-week-old birds. This observation is inconsistent with subsequent studies

and field observations, which confirm that age of infection is directly related to

the degree of immunosuppression. Ivanyi and Morris (1976) showed a 50%

reduction in antibody response to human serum albumen and sheep red blood

cells when chickens were infected a 1-day-old. In contrast, there was no

immunosuppressive response following IBDV infection at 3 weeks of age,

although severe clinical disease with 50% mortality was observed. Infection at

either 1-day-old or 21 days resulted in follicular atrophy of the bursa. The authors

concluded that bursal progenitors of B cells were targets of the IBDV and

peripheral B cells were not affected. Similar results were obtained by Giambrone

et al. (1977) following intraocular infection of 1-day-old and 21-day-old SPF

chicks with 0.06mL of highly pathogenic IBDV containing 106ELD50/mL. The

infection (vaccinated) with IBVD at 1-day-old showed an impaired response to

28-day bovine serum albumin. In contrast, chicks infected at 21 days of age





53


showed an antibody response similar to controls. Infection with IBDV at either

age did not affect skin graft rejection, a measure ofthymus-dependent response.

Effect of IBDV on Humoral Immunity

Although B-cell destruction is most pronounced in the bursa, evidence of viral

replication and associated cellular destruction can also be found in several

secondary lymphoid organs, including cecal tonsils and spleen (Ivanyi and

Morris, 1976; Hirai et al. 1979, 1981). The cytolytic effect of IBDV on B cells

leads to a dramatic reduction in circulating IgM-positive B-cells (Hirai et al.

1981; Kim et al. 1999). IBDV-exposed chickens produce sub-optimal levels of

antibodies against a number of infectious and non-infectious antigens (Kim et al.

1999; Faragher et al. 1972). Only the primary antibody responses are impaired;

the secondary responses remain intact (Giambrone et al. 1977; Sharma et al.

1989).

Recent studies indicate that IBDV-induced humoral deficiency is reversible.

Chickens were exposed to IM-IBDV at 3 weeks of age. At 3, 5, 7, 12, or 17

weeks pi, groups of virus-infected and control birds were inoculated

subcutaneously with 200gg of Tetanus toxoid (TTX) and 1501pG of Brucella

abortus in Freund's incomplete adjuvant. At 10 days after antigenic stimulation,

chickens were examined for levels of anti-TTX and anti-B. abortus antibodies

(Kim et al. 1994). Until 7 weeks pi, the antibody levels were significantly lower

in virus-exposed birds than in control birds. However, the antibody levels against

B. abortus and TTX had returned to normal levels at 12 and 17 weeks pi,

respectively.





54


Interestingly, chronology of the restoration of antibody production was

associated with morphologic restoration of the normal architecture of bursal

follicles (Kim et al. 2000). Although destruction of Ig-producing B-cells may be

one of the principal causes of humoral deficiency, other possible mechanism(s)

needs to be examined. For example, possible adverse effects of IBDV on antigen

presenting cells, such as macrophages and bursal follicular dendritic cells, and

helper T-cell functions remain to be investigated.

Effect of IBDV on Cellular Immunity

T-cell immunity plays an important role in defense against IBDV. This idea is

substantiated by recent observations that replication of IBDV in the bursa was

accompanied by a dramatic infiltration ofT-cells into this organ (Kim et al.

1999). In IBDV-infected chickens, there was an increase in the numbers of

intrabursal T-cells, while the bursa of uninfected chickens had very few resident

T-cells (Kim et al. 1999). Initially, bursal T-cells were detected by

immunohistochemistry at 1-day pi (Sivanandan and Maheswaran, 1980). Such T-

cells were subsequently shown to persist for several weeks (Kim et al. 1999). The

infiltrating T-cells were closely associated with the foci of viral antigen in bursal

follicles. The majority of IBDV-induced bursal T-lymphocytes were T-cell

receptor 2-expressing (TCR2+) oa/-T cells, and a few were TCRI+ y/8 T-cells

(Sivanandan and Maheswaran, 1980).

In a study by Kim I et al. (2000), SPF chickens were exposed to a pathogenic

strain of IBDV. The virus rapidly destroyed B-cells in the bursa. Extensive viral

replication was accompanied by an infiltration ofT-cells in the bursa. Flow





55


cytometric analysis of single-cell suspension of bursal cells was mostly T-cells

with a minority being B-cells (7%). After virus infection, the numbers of bursal

T-cells expressing activation markers Ia and CD25 were significantly increased.

In addition, IBDV-induced bursal T-cells produced elevated levels of IL-6-like

factor and NO-inducing factor in vitro. Spleen and bursal cells of IBDV-infected

chickens had up-regulated IFN-y gene expression in comparison with virus-free

chickens. In IBDV-infected chickens, bursal T-cells proliferated in vitro upon

stimulation with purified IBDV in a dose-dependent manner, whereas virus-

specific T-cell expansion was not detected in the spleen. Cyclosporin A

treatment, which reduced the number of circulating T-cells and compromised T-

cell mitogenesis, increased viral burden in the bursa of IBDV-infected chickens.

These results suggest that intrabursal T-cells and T-cell mediated responses may

be important in viral clearance and promoting recovery from infection.

Although the data on the effect of IBDV on antigen-specific T-cell functions

are controversial (Giambrone et al. 1977), there is convincing evidence that in

vitro mitogenic proliferation of T-cells of IBDV-exposed birds is significantly

compromised. T-cells in the spleen, as well as in the peripheral circulation, were

affected (Confer et al. 1981; Kim et al. 1998). The mitogenic inhibition occurred

early, during the first 3-5 days of virus exposure. Subsequently, the mitogenic

response of T-cells returned to normal levels. During the period of mitogenic

inhibition, T-cells of IBDV-infected chickens also failed to secrete IL-2 upon in

vitro stimulation with mitogens (Sharma and Frederickson, 1987; Kim et al.

1998). Previous cell fractionation studies (Sharma and Lee, 1983) and more





56


recent studies with enriched T-cell populations (Kim et al. 1998) have shown that

adherent cells, most probably macrophages, mediate mitogenic inhibition in

splenocyte suspensions. Pan-purified T-cells from spleens of IBDV-exposed

chickens were responsive to T-cell mitogens; addition of adherent cells from

spleens of virus-exposed but not from virus-free chickens inhibited mitogenesis of

the sorted T-cells. The relevance of in vitro mitogenic inhibition of T-cells to the

in vivo role of T-cells in the pathogenesis of IBDV in chickens is yet not known

Exactly how IBDV induces macrophages to exhibit suppressor effect(s) needs

to be further investigated. Because the inhibitory effect(s) can be transferred by

conditioned medium, apparently macrophages secrete soluble products with

suppressive activities. These products have not been identified. Recently, Kim et

al. (1998) have shown by RT-PCR that during the acute phase of infection with

IBDV, spleen macrophages exhibited a marked enhancement of expression of a

number of cytokine genes. These included type I IFN, chicken myelomonocytic

growth factor, an avian homologue of mammalian IL-8 (Barker et al. 1993; Leutz

et al. 1989). The elevated gene expression by macrophages coincided with in

vitro inhibition of T-cells mitogenic response of spleen cells. Further, mitogen-

stimulated cultures of spleens of IBDV-exposed chickens had elevated nitric

oxide (NO) concentrations in the supernatant. It can be speculated that T-cell

cytokines, such as IFN-y, stimulated macrophages to produce NO, which may

have inhibited mitogen-induced T-cell proliferation (Pertile et al. 1995; Evans,

1995).





57


The direct immunosuppressive effect of IBDV on T-cells has yet to be clearly

identified. Spleen cells from IBDV-exposed chickens produced IFN-y (Kim et al.

2002). Assuming that T-cells were the principal producers of IFN-y, this

observation provides circumstantial evidence that the virus modulates T-cell

function. How this modulation affects the cellular immune competence of the

bird remains to be established. It has been suggested that the virus can also cause

depression of cell-mediated immunity; however, this has been less well

characterized. Other investigators have reported decreased response to

herpesvirus of turkeys vaccination (Sharma, 1984), decreased mitogenic response

of cultured lymphocytes (Confer et al. 1981; Sharma and Lee, 1983), and the

sporadic occurrence of histopathologic lesions in the thymus (Cheville, 1967).

In a study by Rodenberg et al. (1994), using immunofluorescence, there was an

appreciable decline from control levels in the percentage of lymphocytes

expressing sIgM in the spleen and bursa of infected chickens. However, the

relative proportions of T-lymphocytes expressing CD4 and CD8 molecules in

peripheral blood and spleen remained unchanged following infection. Also, in

their study, the absolute number of T-cells per unit sample were not reported.

Therefore, it is possible that an equal reduction of all subpopulations occurred that

would not have been detected by CD4:CD8 ratio. However, the proportional

values obtained generally agreed with previously established levels for normal

chickens given the influences of protocol variation and genetic factors (Hala et al.

1992; Lillehoj et al. 1988).





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As noted above, T-cells that infiltrate the bursa during the acute phase of the

disease inhibited in vitro mitogenic response of normal spleen cells (Kim et al.

1998). Possible suppressive effects of these cells on the immune functions of the

chicken are not known and seem unlikely because the suppressor T-cells were

most pronounced in the bursa. Spleens from IBDV-exposed chickens did not

have an appreciable proportion of suppressor T-cells at the time when bursal T-

cells had well pronounce suppressor activity (Kim et al. 2002).

Effect of IBDV on Innate Immunity

IBDV modulates macrophage functions. There is indirect evidence that the in

vitro phagocytic activity of these cells may be compromised (Lam, 1998). As

noted above, macrophages from IBDV-exposed chickens had up-regulated

cytokine gene expression and produced elevated levels of NO (Kim et al. 1998).

Macrophages are important cells of the immune system and the altered functions

of these cells may influence normal immune responsiveness and inflammation in

chickens (Evans, 1995). Earlier data suggest that natural killer (NK) cell activity

in chickens of two genetic backgrounds remained unaffected by exposure to

virulent IBDV (Sharma and Lee, 1983). However, further studies are needed to

molecularly characterize chicken NK activation.

IBDV-Induced Apoptosis

Programmed cell death or apoptosis is an active type of cell death that is

characterized by nuclear fragmentation and cellular breakdown into apoptotic

vesicles. Unlike necrosis, there is no release of cellular contents in the

interstitium and consequently no inflammation surrounding the dead cells





59


(Rosenberger et al. 1989). This sort of "cellular self-destruction" is usually

initiated by physiological stimuli, but pathological stimuli, such as IBDV, can

also be the triggering factor.

IBDV is a known immunosuppressive agent of chickens (Ivanyi and Morris,

1976; Kaufer and Weiss, 1980; Kim et al. 1999; Confer et al. 1981; Kim et al.

1998), the mechanism of which is not well understood. It has been determined

that the virus causes direct cytopathic effects on the immature B-cells resulting in

severe bursal necrosis, lymphoid depletion, and subsequent immunosuppression.

It has also been reported that the infected bursa undergoes a very rapid and

extensive atrophy with little or no inflammatory responses (Rosenberger and

Cloud, 1989). Immunosuppression without severe inflammatory response of the

bursa is an unexplained phenomenon. This suggests the possible involvement of

apoptotic processes in the pathogenesis of IBD.

In a report by Vasconcelos and Lam (1994), heparinized blood was taken from

white Leghorn chickens free of antibodies against IBDV, to harvest PBLs, which

were divided into 3 groups. One group received IBDV serotype 1, a second group

received hydrocortisone, and a third group received RPMI 1640 medium only.

Each sample was counted and the apoptotic and necrotic indices were measured

as described by Cohen et al. (1992). In an additional experiment, the cells were

lysed and DNA was extracted and precipitated. Aliquots of DNA were

electrophoresed.

DNA extracted from IBDV infected lymphocytes showed an intense laddering

pattern in agarose gel electrophoresis. IBDV-infected PBLs had significantly





60


higher apoptotic and necrotic indices than did control lymphocytes. Electron

micrographs of the IBDV-infected PBLs showed typical aspects of apoptosis,

such as peripheral condensation of chromatin, blebbing of the plasma membrane,

fragmentation of the nucleus and of the cell, leading to the formation of apoptotic

bodies. These finding indicated that IBDV, in addition to causing necrosis in

avian lymphocytes, could induce apoptosis (Vasconcelos and Lam, 1994).

The induction of apoptosis in IBDV-infected chicken peripheral blood

lymphocytes has been reported (Vasconcelos and Lam, 1994). Apoptotic cell

death was also observed in vitro in IBDV-infected Vero cells and CEFs (Tham

and Moon, 1996). IBDV infection of susceptible chickens resulted in the

induction ofapoptosis of cells in the bursa, (Ojeda et al. 1997; Tanimura and

Sharma, 1997) as well as in the thymus (Inoue et al. 1994; Tanimura and Sharma,

1997).

Two IBDV proteins have been suspected to play a role in the induction of

apoptosis. Fernandez-Arias et al. (1997) showed that the structural protein VP2

induced apoptotic cell death of mammalian cells but not in CEFs. A VP5-deletion

mutant IBDV strain also induced apoptosis in a reduced number of infected CEFs

compared with the parental strain; this mutant strain replicated more slowly than

the parental strain (Yao et al. 1998). Results of previous studies indicated a

correlation between virus replication and apoptosis of bural cells. The

involvement of indirect mechanisms was suggested by Inoue et a (1994) since

apoptosis was observed in T-cells of the thymus of infected chickens, whereas

IBDV antigens were found mainly in infikrated B-cells or in reticular cels





61


Furthermore, Tanimura and Sharma (1997) investigated sections of IBDV-

infected bursas and demonstrated apoptotic cells in not only antigen-positive but

also antigen-negative bursal follicles.

Jungmann et al. (2001) studied the kinetics of IBDV replication and induction

of apoptosis in vitro and in vivo. After infection of CEFs with IBDV, the

proportion of apoptotic cells increased from 5.8% at 4 hours pi to 64.5% at 48

hours pi. The proportion of apoptotic cells correlated with IBDV replication.

UV-inactivated IBDV particles did not induce apoptosis. Double labeling

revealed that primarily in the early stages after infection, the majority of antigen-

expressing cells were not apoptotic; double-labeled cells appeared more

frequently at later times. Remarkably, apoptotic cells were frequently located in

the vicinity of antigen-expressing cells. This indicated that cells replicating IBDV

might release an apoptosis-inducing factor(s). Since IFN production has been

demonstrated after IBDV infection, IFN was considered to be one of several

factors. However, supernatants of infected CEFs in which virus infectivity had

been neutralized were not sufficient to induce apoptosis. Similar results were

observed in the infected bursas: early after infection, most of the cells either

showed virus antigens or were apoptotic. Again, double-labeled cells appeared

more frequently late after infection. This suggests that indirect mechanisms might

also be involved in the induction of apoptosis in vivo, contributing to the rapid

deletion of cells in the IBDV-infected bursa (Jungmann et al. 2001).

Yao and Vakharia (2001) reported that the NS protein of IBDV alone is

capable of inducing apoptosis in cell culture. Transfection of a chicken B-cell





62


line and CEFs with a plasmid DNA, containing the NS protein gene under the

control of the immediate-early promoter-enhancer region of human

cytomegliovirus, induced apoptosis in both cell lines. Apoptotic changes, such as

chromatin condensation, DNA fragmentation, and the appearance of apoptotic

nuclear bodies, were observed in cell cultures 48 hours pi. This demonstrated that

the mutant virus is closely associated with its yield from the supernatant;

approximately 30-fold lower than the wild-type due to increased cell association,

indicating a deficiency in lysis of virus-infected cells. Taken together, these

results indicate that the NS protein of IBDV is highly cytotoxic, which brings

about the release of the viral progeny from cells, and thus play an important role

in viral pathogenesis.

IBDV Replication

Many IBDV strains replicate in both chicken and mammalian cell lines;

however, highly pathogenic strains are often difficult to cultivate. Both viruses

produce cytopathic effects 1-2 days after inoculation. Biraviruses replicate in

the cytoplasm without greatly depressing cellular RNA or protein synthesis. The

viral mRNA is transcribed by a virion-associated transcriptase (Kibenge et al.

1988).

Replication involves the synthesis by the virion RNA-dependent RNA

polymerase of two genome length mRNAs, one from each of the genome

segments (Macdonald, 1980). Viral RNA is transcribed by a semi-conservative

strand displacement mechanism (Spies et al. 1987). Segment A mRNA is

translated to a polyprotein that is cleaved to form (5' to 3') the pre-VP2, VP4, and





63


VP3 proteins. Pre-VPS is later processed by a slow maturation cleavage to

produce VP2 (Becht et al. 1988). The mRNA from segment B is translated to

form VP1 (MacDonald and Dobos, 1981). Virus particles assemble and

accumulate in the cytoplasm. IBDV is transmitted horizontally and there is no

evidence that IBDV is transmitted through the egg (Kibenge et al. 1988).

Clinical and Subclinical IBDV Infections

Classical IBD is characterized by acute onset, relatively high morbidity, and

low flock mortality in 3-6-week-old broilers or replacement pullets, resulting in

significant clinical signs (Hanson et al. 1967). Clinical signs usually appear after

an incubation period of 2 to 4 days and are associated with acute disease,

including anorexia, depression, ruffled feathers, diarrhea, prostration, and death.

Birds are disinclined to move and peck at their vents and pericloacal feathers are

stained with urates (Landgrafet al. 1967). Feed intake is depressed but water

consumption may be elevated. Terminally, birds may show sternal or lateral

recumbency with coarse tremor (Appleton et al. 1963). The short duration of

clinical signs and the mortality patterns are considered to be of diagnostic

significance for IBD. Affected flocks show depression for 5-7 days during which

mortality increases rapidly for the first two days then declines sharply as clinical

normality returns (Parkhurst, 1964). Such clinical signs occur usually in chicks

infected after 3 weeks of age when passively acquired IBDV-specific maternal

antibodies fade. The incidence of mortality is highly variable ranging from 1000/

to negligible. Lesions include bursal atrophy, dehydration, and darkened

discoloration of pectoral muscles (Cosgrove, 1962). Often hemorrhages may be





64


present in the thigh and pectoral muscles and the bursa (Hitchner, 2004). Atrophy

of the bursa is the most prominent gross lesion found in chickens suffering from

acute IBD. Detection of virus neutralizing (VN) antibodies to IBDV can be

accomplished by ELISA. Early studies attempted to correlate ELISA titers with

VN antibodies and suggested the results were indicative of protection from IBDV

(Briggs et al, 1986; Whetzel and Jackwood, 1995). However, a current ELISA kit

has been produced to highly correlate (99%) with VN antibodies against IBDV

VP2 subunit antigen (Jackwood and Sommer, 1998). Birds that survive the acute

phase of the disease clear the virus and recover from clinical disease.

The subclinical form of disease occurs generally in chickens less than 3 weeks

of age and results in immunosuppression. This is an important point since

immunosuppression results in the presence of passively acquired maternal

antibodies produced by conventional vaccination protocols. IBDV-induced

immunosuppression, including inhibition ofB- and T-cell functions in subclinical

infection, is usually overcome weeks later. However, a variety of field infections,

especially of the respiratory system, may follow immunosuppression caused by

IBD (Faragher et al. 1974). The specific clinical manifestations will reflect the

type and the severity of primary viral and protozoal agents and secondary

bacterial infection, including E. coli (Rosenberger and Gelb, 1978).

Birds that succumb to the infection are dehydrated, with darkened

discoloration of pectoral muscles. Frequently, hemorrhages are present in the

thigh and pectoral muscles. There is increased mucus in the intestine, and renal





65


changes may be prominent in birds that die or are in advanced stages of the

disease (Cosgrove, 1962). Such lesions are most probably a consequence of

severe dehydration.

In fully susceptible flocks, the disease appears suddenly and there is high

morbidity, possibly approaching 100%. Mortality usually begins day 3 pi and

will peak and recede within a period of 5-7 days. Striking features of this disease

are the sudden and high morbidity, spiking death curve, and rapid flock recovery

(Parkhurst, 1964). Initial outbreaks on poultry farms are usually the most acute.

Recurrent outbreaks in succeeding broods are less severe and frequently go

undetected. Many infections are silent, owing to age of birds (less than 3 weeks),

infection with avirulent field strains, or infection in presence of IBDV-specific

maternal antibodies in progeny.

IBDV Pathogenesis

Field IBDVs exhibit different degrees of pathogenicity in chickens. Vaccine

viruses also have varying pathogenic potential in chickens. All breeds of chicken

are affected. It was observed by many that white Leghorns exhibited the most

severe reactions and had the highest percentage mortality. However, Meroz

(1966) found no difference in mortality between heavy and light breeds in a

survey of 700 outbreaks of the disease.

The period of greatest susceptibility is between 3 and 6 weeks of age.

Susceptible chickens younger than 3 weeks do not exhibit clinical signs but have

subclinical infections that are economically important because the result can be

severe immunosuppression of the chicken (Allan et al. 1972). The incubation





66


period is very short and clinical signs of the disease are seen in 2-3 days and

histologic evidence of infection can be detected in the bursa within 24 hours

(Hemboldt and Garner, 1964). Mueller et al. using immunofluorescence

techniques, observed infected gut-associated macrophages and lymphoid cells

within 4-5 hours after oral exposure to IBDV (1986). Virus-infected cells were

present in the bursa by 11 hours after oral exposure and 6 hours after direct

application of virus to the bursa.

It is noteworthy that bursectomized chicks do not show clinical signs following

infection with pathogenic strains of IBDV. Due to the absence of host cells, virus

multipliation is inhibited, although IBDV can be re-isolated from spleen, thymus,

and liver up to 5 days after infection in bursectomized chicks. The concentration

of virus is only 103 of the level in non-bursectomized, infected controls (Kaufer

and Weiss, 1980).

Recent observations provide new information on the pathogenesis of IBDV

and the mechanism of recovery from acute infection. It is important to note that a

healthy bursal follicle consists ofB-cells (85-95%), T-cells (<4%), and other non-

lymphoid cells (Ewert et al. 1984; Palojoki et al. 1992). In a study by Sharma et

al. (1989), 3-week-old SPF chickens were inoculated with virulent IBDV. During

the acute phase of the infection, the phenotype of the cells that populated bursal

follicles was examined. As expected, the number of sIgM-positive B-cells

dropped precipitously as the virus replicated within bursal follicles. However, the

appearance of viral antigen in the bursa was accompanied by a dramatic

infiltration of T-cells in and around the site of virus replication. Infiltrating T-





67


cells were first detected at 1-day pi and persisted until at least 12 weeks pi,

although the viral antigen had disappeared by 3 weeks pi. Flow cytometry

performed on single bursa cell suspensions at intervals after virus exposure

demonstrated that the highest numbers ofintra-bursal T-cells were present at 7

days pi. At peak accumulation, 65% of the bursal cells were T-cells and 7% had

sIgM expression. Although CD4-positive and CD8-positive lymphocytes were

roughly in equal proportions during the first 7 days pi, CDS-positive T-cells

became predominant thereafter.

Starting at 5 weeks pi, signs of bursal recovery were noted. Bursal follicles

that had been depleted of lymphocytes during the acute phase of the disease began

to be filled with sIgM-positive lymphoid cells. By 12 weeks pi, almost all bursal

follicles had been replete with sIgM-positive B-cells and the morphology of the

bursa had returned to the pre-infection state. However, mechanisms of bursal

recovery need to be further investigated. Because there is a dramatic influx ofT-

cells at the site of viral replication, one can speculate that the infiltrating T-cells

may be involved in limiting viral spread and thus initiating the recovery process.

T-cells seem to be important for normal development of the bursa and the

maturation of B-cells in the embryo. Furthermore, studies have shown that

selectively induced immunodeficiency in the T-cell system promoted virus

persistence in the bursa. However, more data are needed to confirm this

observation.

The lytic effect of IBDV is most prominent in the B-cells in the bursa (Hirai et

al. 1981; Rodenberg et al. 1994). During the acute phase of IBDV infection,




68


chickens experience severe bursal atrophy characterized by necrosis and depletion

of lymphoid cells, cyst formation in bursal follicles, and infiltration of

inflammatory cells. The bursal atrophy may be associated with sudden death

within 3-5 days of the virus exposure. The pathogenesis ofIBDV in chickens

appears to be influenced by the age at which virus exposure occurs (Kim et al.

1999). Immunosuppression induced by IBDV was most pronounced in chickens

younger than 3 weeks of age although clinical disease was most pronounced if

virus exposure occurred after 3 weeks of age. This immunosuppressive effect of

IBDV was first recognized by Faragher et al. (1972). The immunoglobulin class

of IBDV-specific antibodies in serum was found to be IgG when determined by

ELISA (Hoshi, et al. 1995).

The bursa appears to be the primary target organ of the virus. Cheville made a

detailed study of bursal weights for 12 days pi. (1967). It is important that the

sequence of changes be understood when examining birds for diagnosis. By day

2 or 3 pi, the bursa has a gelatinous yellowish transudate covering the surface.

Longitudinal striations on the surface become prominent, and the normal white

color resembles a cream color. As the transudate begins to disappear, the bursa

returns to its normal size becoming gray during the period of atrophy. By day 3

pi, the bursa begins to increase in size and weight because of edema and

hyperemia. By the day 4, the bursa may double its normal weight and then it

begins to recede in size. By day 5, it may return to normal bursa weight, but the

bursa continues to atrophy. Upon and after day 8, it can shrink to approximately

one-third of its original weight.





69


IBDV-infected bursas often show necrotic foci. In addition, petechial

hemorrhages may be found on the mucosal surface. Occasionally, extensive

hemorrhaging throughout the entire bursa has been observed. However, in these

cases, chickens may void blood in their droppings. The spleen may also enlarge

slightly and often have small gray foci uniformly dispersed on the surface

(Rinaldi et al. 1972). Additionally, hemorrhages may be observed in the mucosa

at the junction of the proventriculus and gizzard.

Under natural conditions, the most common mode of infection appears to be

via the oral route. From the gut, the virus is transported to other tissues by

phagocytic cells, most likely resident macrophages. Although viral antigen has

been detected in liver and kidney within the first few hours of infection, extensive

viral replication takes place primarily in the bursa (Muller et al. 1979).

In vivo and in vitro studies have shown that the target cell is an IgM-bearing B-

cell (Ivanyi and Morris, 1976; Kaufer and Weiss, 1980). Within hours of

exposure, virus-containing cells appear in the bursa and the virus spreads rapidly

through the bursal follicles. Virus replication leads to extensive lymphoid cell

destruction in the medullary and the cortical regions of the follicles (Tanimura

and Sharma, 1997). The cellular destructive process may be accentuated by

apoptosis of virus-free bystander cells (Tanimura and Sharma, 1998). The acute

lytic phase of infection is associated with a reduction in circulating sIgM-positive

B-cells (Hirai et al. 1981; Rodenberg et al. 1994), although there is no detectable

reduction in circulating immunoglobulins (Giambrone et al. 1977; Kim et al.

1999).





70


T-cells are resistant to infection with IBDV (Hirai et al. 1979). Although the

thymus undergoes marked atrophy and extensive apoptosis ofthymocytes during

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

replicates in thymic cells (Tanimura and Sharma, 1998). Gross and microscopic

lesions in the thymus are quickly overcome and the thymus returns to its normal

state within a few days of virus infection.

Recent observations provide new information on the pathogenesis of IBDV

and the mechanism of recovery from acute infection (Sharma et al, 1994). Three-

week-old SPF chickens were inoculated with virulent IBDV. During the acute

phase of the infection, the phenotype of the cells that populated bursal follicles

was examined. As expected, the number of sIgM-positive B-cells dropped

precipitously as the virus replicated within bursal follicles. However, the

appearance of viral antigen in the bursa was accompanied by a dramatic

infiltration of T-cells in and around the site of virus replication. The infiltrating

T-cells were first detected at 1-day pi and persisted until at least 12 weeks pi,

although the viral antigen had disappeared by 3 weeks pi. Flow cytometric

analyses of single bursa cell suspensions at intervals after virus exposure revealed

that the highest numbers of intra-bursal T-cell were present at 7 days pi. At peak

accumulation, 65% of the bursal cells were T-cells and 7% had sIgM expression.

Although CD4+ and CD8+ cells were roughly in equal proportions during the

first 7 days pi, CD8+ cells became predominant thereafter. At 5 weeks pi, signs

of bursal recovery were noted. Bursal follicles that had been depleted of

lymphocytes during the acute phase of the disease began to be replaced with





71


sIgM-bearing B-lymphocytes. By 7 week pi, about 40% of bursal follicles had

been repopulated with lymphocytes. By 12 weeks pi, almost all bursal follicles

had been replete with IgM-positive B-cells and the morphology of the bursa had

returned to the pre-infection state. The mechanism of bursal recovery needs to be

investigated. Because there is a dramatic influx ofT-cells at the site of viral

replication, it can be speculated that the infiltrating T-cells may be involved in

limiting viral spread and thus initiating the recovery process.

T-cells seem to be important for normal development of the bursa and the

maturation of B-cells in the embryo (Hirota and Bito, 1978). Data from a

preliminary experiment in which chickens were treated with cyclosporin A (CsA)

before exposure to IBDV support this possibility (Kim et al. 2000). CsA

treatment inhibits transcription of the genes encoding a number of cytokines and

selectively suppresses T-cell function by inhibiting IL-2 receptor expression and

blocking IL-2-mediated signal transduction (Nowak et al. 1982; Zenke et al.

1993). Virus prevalence in the bursa was compared in chickens with or without

CsA treatment. The CsA-treated chickens had lower numbers of T-cells

infiltrating the bursal follicles and higher levels of viral antigen than the CsA-free

chickens. These results indicate that selective immunodeficiency in the T-cell

system promoted virus persistence in the bursa. However, additional research in

this area is needed to further support this observation.

On the contrary, it is also a possibility that IBDV-induced T-cells may enhance

viral lesions. For example, CTL may exasperate virus-induced cellular

destruction by lysing cells expressing viral antigens. T-cells may also promote





72


the production of inflammatory factors that may accentuate tissue destruction.

NO produced by macrophages activated by T-cell cytokines, such as IFN-y, may

promote cellular destruction. Chickens treated with L-NAME (NO synthetase

inhibitor) before exposure to IBDV had much less bursal necrosis and lower

levels of viral antigen than the untreated virus-exposed chickens (Yeh et al. 2002).

Clearly, additional studies are needed to examine the role of T-cells in IBDV

pathogenesis.

Sharma et al. (1994) examined the characteristics of IBDV-induced bursal T-

cells. At 7 days pi, when the majority of the lymphocytes in the bursa were

expected to be T-cells, single cell suspensions of the bursal tissue were prepared

and the cells were examined by a number of assays. The results revealed that: (a)

bursal T-cells had elevated surface expression of MHC class I and IL-2

receptors; (b) bursal cells had elevated expression of cytokines, such as IFN-y and

IL-6-like factor; (c) bursal T cells from the IBDV-infected chickens proliferated

when stimulated in vitro with purified IBDV; and, (d) bursal T-cells inhibited

mitogenic response of normal, histocompatible splenocytes in a dose-dependent

manner. The mitogenic inhibition was mediated by CD4+ T-cells, as well as by

the conditioned medium of such cells.

Diagnosis of IBDV

Isolation of IBDV can be accomplished from bursal tissue obtained during the

acute stage of infection (Rosenberger and Gelb, 1978). The suggested procedure

involves pooling inflamed bursae from birds. An organ homogenate comprising

20:80% weight:volume, oftryptose phosphate broth is treated with antibiotics and





73


centrifuged. Virus can be propagated in 10-day embryonated SPF eggs inoculated

via the chorioallantoic membrane (Hitchner, 1970). Conventional type 1 IBDV is

embryo-lethal in 3-5 days, producing vascular congestion and subcutaneous

hemorrhages. In contrast, US type 1 variants produce stunting of embryos on the

seventh day after infection. Affected embryos are edematous and show

splenomagaly and hepatic necrosis. Embryonic hemorrhage and death are not

observed following inoculation of SPF eggs with variants of type 1 IBDV.

Chicken embryo bursal and kidney cells (Lukert and Davis, 1974) can be used

to propagate IBDV, but adaptation is required to grow virus on CEFs (McNulty et

al. 1986). The IBDV can be identified by electron microscopy (McFerran et al.

1978) or direct immunofluorescence (Snyder et al. 1984). IBDV antigen can be

demonstrated in formalin-fixed and paraffin-embedded preparations of bursal

tissue. Joensson and Engstrom (1986) showed that pretreatment with trypsin or

pronase before fixing in Bouin's solution enhanced subsequent detection of IBDV

by indirect immunoperoxidase and immunofluorescence staining. Snyder et al.

(1984) used monoclonal antibodies to identify IBDV in tissues.

Antibodies to IBDV can be detected using a number of serological procedures.

The agar gel diffusion precipitin test (AGDP) was the original qualitative method

to detect antibody. Bursal homogenate is used as the antigen to demonstrate

antibody 7 days after infection (Rosenberger and Cloud, 1989). Commercial

AGDP kits can be used for serological screening. The system can be used as a

quantitative gel diffusion precipitin test as described by Cullen and Wyeth (1975).

The method used extensively in the UK during the 1970s and 1980s correlates





74


with data obtained from serum virus neutralization and ELISA and has been

applied to evaluate immunity in breeder flocks (Wyeth and Chettle, 1982). Box et

al. (1988) compared the results of QAGDP, ELISA, and serum virus

neutralization to quantify antibody levels to IBDV.

The constant virus serum dilution neutralization test has been used extensively

for research, serological surveys and flock surveillance. The microtiter system

has replaced inoculation of SPF eggs as the neutralization procedure of choice.

Rosenberger and Cloud (1989) described a method, which uses CEFs and IBDV

adapted to the cell culture system. Neutralization titers represent the reciprocal of

the specific serum dilution, which inhibits cytopathology. The serum virus

neutralization procedure is extremely sensitive (Weisman and Hitchner, 1978) and

is sufficiently specific to differentiate among serotypes of IBDV (Chin et al.

1984).

Subclinical IBD infection of flocks with variable maternal antibody protection

and infection with variant type 1 strains may be difficult to diagnose without

recourse to serology, histopathology, and isolation and identification of the

pathogen.













CHAPTER 3
CHALLENGE OF SPAFAS LAYER AND MATERNALLY IMMUNE
BROILER CHICKENS ON DAY 16 OR 18 WITH VERY VIRULENT IBDV
ALAN LABORATORIES-2 OR DELMARVA VARIANT E ISOLATES

Introduction

IBDV causes considerable economic loss in the poultry industry by inducing

severe clinical signs, high mortality (50%), and immunosuppression in chickens

because bursal B-cells are targets for IBDV infection resulting in B-cell depletion.

The subclinical form of disease occurs generally in chickens less than 3 weeks of

age and results in immunosuppression. This is an important point since

immunosuppression results in the presence of passively acquired maternal

antibodies produced by conventional vaccination protocols. IBDV-induced

immunosuppression, including inhibition of B- and T-cell functions in subclinical

infection, is usually overcome weeks later. However, a variety of field infections,

especially of the respiratory system, may follow immunosuppression caused by

IBD (Faragher et al. 1974). The specific clinical manifestations will reflect the

type and the severity of primary viral and protozoan agents and secondary

bacterial infection, including E. coli (Rosenberger and Gelb, 1976).

Most IBD has been controlled by live IBDV vaccines based on strains of

intermediate virulence (Ismail and Saif, 1991). However, it is difficult to protect

field chickens with maternal antibodies induced by live IBDV vaccination (Ismail

and Saif, 1991; Tsukamoto et al. 1995b). In addition, live vaccines induce




75





76


moderate bursal atrophy (Muskett et al. 1985), and the antigenic or pathogenic

characters are not stable. Currently, the disease is prevented by application of an

inactivated vaccine in breeder chicken flocks, after chickens are primed with

attenuated live IBDV vaccine. This has minimized economic losses caused by

IBD.

Vaccination of poultry flocks, especially parent flocks, is often performed with

the intention of protecting the progeny via maternal antibodies during the first

weeks of life. The efficacy of this vaccination schedule, more precisely described

as induction of indirect protection, is normally proven by challenging the chickens

(Jungback and Finkler, 1996). IBDV-specific antibodies transmitted from the

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

resultant protection against the immunosuppressive effects of the virus. Maternal

antibody will normally protect chicks against infection for 1-3 weeks, but by

boosting the immunity in breeder flocks with oil-adjuvanted inactivated vaccines,

passive immunity may be extended to 4 or 5 weeks.

The major problem with active immunization of young maternally immune

chicks is determining the proper time of vaccination. This varies with levels of

maternal antibody, route of vaccination, and virulence of the vaccine virus.

Environmental stresses and management may be factors to consider when

developing a vaccination program that will be effective. Results suggest that

serological determination of the optimum vaccination time for each flock is

required to effectively control highly virulent IBDV in the field. The optimum

vaccination timing could be approximated by titration of the maternal IBDV





77


antibodies of day-old chicks by ELISA (Tsukamoto, et al. 1995b). Since

subclinical IBD infection of flocks with variable maternal antibody protection and

infection with variant type 1 strains may be difficult to diagnose, use of serology,

isolation, and identification of the pathogen is important.

This chapter is related to the many problems that exist for IBDV vaccination

protocols. In terms of vaccination, newly emerging very virulent IBDV isolates

continue to escape vaccine-induced protection. Vaccination of dams to transfer

maternal IBDV-specific antibodies to chicks can also interfere with vaccination

schedules of progeny. Additionally, the USDA recommends evaluation of IBDV-

induced pathology be performed by B/B weight ratios. This recommendation is

because IBDV infection causes bursal atrophy. However, such bursal atrophy

follows an initial period of bursal edema. Therefore, this experiment was

performed to determine whether current standards to evaluate subclinical IBDV

infection, including gross bursa scoring and B/B weight ratio, are as accurate

measurements of virulence as bursal histopathology.

Project Design

SPAFAS layer and maternally immune broiler chickens were challenged on

Day 16 or 18 with very virulent IBDV; chicks received one of either very virulent

IBDV strains, Alan Laboratories-2 (AL-2) or Delmarva Variant E (DVE). Gross

bursa scoring, B/B weight ratios, and histopathology 1-week post infection (pi)

measured subclinical affected after challenge on either Days 23 or 25. Statistical





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methods analyzed whether a difference exists in B/B weight ratios among

chickens from 3-commercial producers on and between challenge days. Animal

design and time line of treatment are shown in Tables 3-1 and 3-2.

Materials and Methods

Experimental Animals

SPAFAS Leghorn and maternally immune broiler eggs were obtained from 3-

commercial producers: T, C, and M.

Incubation and Hatching Conditions

Incubation of all eggs to place for 16 days under the following conditions:

99.50F and 60% relative humidity. Subsequently, all eggs were hatched under the

following conditions: 98.50F and 65% relative humidity. All day-old chicks

were neck-banded with color-coded and numbered system to facilitate

identification. All chickens were raised under a stringent biosecurity program,

including limited room entry, mandatory showering before and after entrance, and

controlled management conditions, including temperature, feed, and water. Day-

old chicks were placed into constantly lighted (for 3 days), pre-heated (880F)

rooms. Each room contained a single chicken cage battery. On Day 6, room

temperature was gradually lowered to 780F to enhance feed consumption. Chicks

were allowed to drink fresh water and eat "starter feed" ad libitum.

Location of IBDV Research

All IBDV research was conducted at the University of Florida, College of

Veterinary Medicine, Poultry Medicine Laboratory in Building 177 and

accompanying chicken battery rooms within the College of Veterinary Medicine





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at the University of Florida in Gainesville, Florida. Chickens receiving different

treatments were housed in order, as shown in Table 3-3. This animal project was

approved by IACUC (#C315).

Animal Room Preparation

Buildings with separate rooms were prepared by disinfection before chick

placement. Rooms and battery cages were disinfected using Environ One-Stroke

(1:256) and heated to 1200F for a period of 72 hours. A subsequent bleaching

(1:2) process was performed with reheating the room to 1200F for 72 hours. A

final disinfecting process was performed on sealed rooms by para-formaldehyde

fumigation. This was performed by combing potassium permanganate (KMnO4)

and 10% formalin to create gaseous para-formaldehyde.

Experimental IBDV Challenge

Two strains of IBDV, including AL-2 and DVE isolates, provided by Intervet,

Inc., Millsboro, Delaware. Challenge virus was diluted in tryptose-phosphate

broth according to the manufacturer. Challenge was performed by oro/nasal route

with 103.5EIDso of either IBDV isolate.

Measurement of Subclinical Effects of IBDV Challenge

B/B weight ratio

Upon necropsy 1-week pi, total chicken and bursa weights were measured to

calculate average B/B weight ratios, where the B/B weight ratio = (Bursa

weight/total Body weight) x 100.





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Gross bursa scoring

A simple gross bursa scoring system was applied according to the following

scale: 0 (normal), 1 (edematous), and 2 (atrophic).

Histopathology

Briefly, bursas were stored in 10%-buffered formalin and cut into 5um

sections, embedded in paraffin, and stained with hematoxylin and eosin for

subsequent microscopic evaluation of histopathology.

Statistical analysis

B/B weight ratios were measured and average and standard deviation was

calculated. To determine whether a significant difference existed among ratios

between commercial broilers and SPF Leghorns receiving identical treatment,

statistical analysis involved the use of the Kruskal-Wallis one-way nonparametric

ANOVA. The rejection value of this test was 0.050, therefore, any p value less

than 0.05 was considered to be a significant difference. To determine whether a

significant difference existed among identical IBDV strain treatment groups

between Days 16 and 18, a two-sample T-test was performed. This test was

performed at a level of 95% certainty for differentiation, therefore, any p value

less than 0.05 was considered to be a significant difference. B/B weight ratios

were also compared to gross bursa scoring and histopathology.

Results

B/B Weight Ratios

One week after respective challenge, all chickens were necropsied. B/B

weight ratios were calculated for broilers and SPAFAS layers challenged at the





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age of 16 or 18 days with either AL-2 or Del Variant E isolates ofIBDV.

Unchallenged controls were compared with challenged chicks of identical age and

commercial source. A summary of B/B weight ratios is shown in Figure 3-1.

In terms of IBDV AL-2 challenge on Day 16, maternally immune broilers

from companies T, C, and M had B/B weight ratios of 0.141, 0.094, and 0.102,

respectively. SPAFAS layers that received identical IBDV AL-2 challenge had

an average B/B weight ratio of 0.189. Within this treatment group, there were

three groups in which the means were not significantly different from one

another: T and M, M and C, and SPAFAS layers. SPAFAS layers displayed

ratios statistically higher than those of commercial broilers (p<0.0001).

In terms of IBDV DVE challenge on Day 16, maternally immune broilers from

companies T, C, and M had B/B weight ratios of 0.184, 0.148, and 0.163,

respectively. SPAFAS layers that received identical IBDV DVE challenge had an

average B/B weight ratio of 0.147. There are no significant pair wise differences

among either group of commercial broilers or SPAFAS layers (p=0.1662).

One-week post SHAM challenge on Day 16, maternally immune commercial

broilers and SPAFAS layers were treated identically as same age challenged

chickens. B/B weight ratios were calculated. Leghorns from T, C, and M had

average B/B weight ratios of 0.277, 0.293, and 0.285, respectively. SPAFAS

layers that received identical SHAM challenge had B/B weight ratio of 0.701.

There were two groups in which the means were not significantly different from

one another: T, C, and M and SPAFAS (p<0.0001).





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In addition to the chickens challenged on day 16 of age, other groups of

commercial Leghorn and SPAFAS layer chickens were challenged on day 18 of

age with either AL-2 or DVE IBDV isolates to be compared with unchallenged

chicks of identical age.

B/B weight ratios were calculated for AL-2 challenged Leghorns and layers at

day 18 of age. Commercial Leghorns from companies T, C, and M had average

B/B weight ratios of 0.132, 0.122, and 0.102, respectively. SPAFAS layers

challenged with AL-2 variant virus had an average B/B weight ratio of 0.157.

There are three groups in which the means are not significantly different from one

another: T and C, C and M, and T and SPAFAS (p<0.0001).

B/B weight ratios of 18 day-old DVE challenged broilers and layers also had

similar results as 18 day-old Leghorns challenged with AL-2 IBDV. Broilers from

companies T, C, and M had average B/B weight ratios of 0.184, 0.209, and 0.143,

respectively. In addition to DVE challenged broilers, SPAFAS layers challenged

with the same variant virus demonstrated an average B/B weight ratio of 0.152.

There were two groups in which the means were significantly different from one

another: T, C, and SPAFAS and M and SPAFAS (p=0.0044).

One-week post SHAM challenge on Day 18, maternally immune commercial

Leghorns and SPAFAS layers were treated identically as same age challenged

chickens. B/B weight ratios were calculated. Leghorns from T, C, and M had

average B/B weight ratios of 0.266g, 0.285g, 0.28 g, respectively. SPAFAS





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layers that received identical SHAM challenge had B/B weight ratio of 0.671g.

There were two groups in which the means were not significantly different from

one another: T, C, and M and SPAFAS (p<0.0001).

Broilers from company T challenged with IBDV AL-2 on either day 16 or 18,

B/B weight ratios were not significantly different (p=0.4902). Broilers from

company C challenged with IBDV AL-2 on either day 16 or 18, B/B weight ratios

were not significantly different (p=0.3319). Broilers from company M challenged

with IBDV AL-2 on either day 16 or 18, B/B weight ratios were not significantly

different (p=0.9438). In terms of SPAFAS Leghorns challenged with IBDV AL-2

on either day 16 or 18, B/B weight ratios were not significantly different

(p=0.0796).

Broilers from company T challenged with IBDV DVE on either day 16 or 18,

B/B weight ratios were not significantly different (p=0.9967). Broilers from

company C challenged with IBDV DVE on either day 16 or 18, B/B weight ratios

were significantly different (p=0.0172). Broilers from company M challenged

with IBDV DVE on either day 16 or 18, B/B weight ratios were not significantly

different (p=0.2183). SPAFAS Leghorns challenged with IBDV DVE on either

day 16 or 18, B/B weight ratios were not significantly different (p=0.7026).

Broilers from company T SHAM-challenged on either day 16 or 18, B/B

weight ratios were not significantly different (p=0.4961). Broilers from company

C SHAM-challenged on either day 16 or 18, B/B weight ratios were not

significantly different (p=0.5934). Broilers from company M SHAM-challenged





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on either day 16 or 18, B/B weight ratios were not significantly different

(p=0.6883). SPAFAS Leghorns SHAM-challenged on either day 16 or 18, B/B

weight ratios were not significantly different (p=0.6536).

Body Weight Averages

A summary of average body weight results is shown in Figure 3-2. Necropsy

results of commercial Leghorns from T, C, and M challenged with IBDV AL-2 on

day 16 had average total body weights of 587.8g, 667.8g, and 659.4g,

respectively. Identically challenged SPAFAS layers had average total body

weight of 214.5g.

Necropsy results of commercial Leghorns from T, C, and M challenged with

IBDV DVE on day 16 had average total body weights of 579.4g, 638.0g, and

653.5g, respectively. Identically challenged SPAFAS layers had an average total

body weight of 224.6g.

Necropsy results of commercial Leghorns from T, C, and M SHAM-

challenged on day 16 had average total body weights of 535.0g, 597.9g, and

590.7g, respectively. Identically challenged SPAFAS layers had an average total

body weight of 204.6g.

Necropsy results of commercial broilers from T, C, and M challenged with

IBDV AL-2 on day 18 had average total body weights of 609.2g, 641.6g, and

689.6g, respectively. Identically challenged SPAFAS layers had average total

body weight of 217.9g.

Necropsy results of commercial broilers from T, C, and M challenged with

IBDV DVE on day 18 had average total body weights of 626.4g, 673.5g, and





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678.7g, respectively. Identically challenged SPAFAS layers had average total

body weight of 229.1g.

Necropsy results of commercial broilers from T, C, and M SHAM-challenged

on day 18 had average total body weights of 641.2g, 695.7g, and 642.8g,

respectively. Identically challenged SPAFAS layers had an average total body

weight of 239.6g

Bursa Weight Averages

A summary of bursa weights is shown in Figure 3-3. In terms of IBDV AL-2

challenge on Day 16, maternally immune Leghorns from companies T, C, and M

had average bursa weights of 0.802g, 0.614g, and 0.680g, respectively. SPAFAS

layers that received identical IBDV AL-2 challenge had an average bursa weight

of 0.403g.

IBDV DVE challenge upon 16 days of age resulted in maternally immune

Leghorns from companies T, C, and M having an average bursa weight of 1.03g,

0.940g, and 1.04g, respectively. SPAFAS layers that received identical IBDV

DVE challenge had bursa weight of 0.325g.

One-week post SHAM challenge on Day 16, maternally immune commercial

Leghorns and SPAFAS layers were treated identically as same age challenged

chickens.

Total bursa weights were measured. Commercial Leghorns from T, C, and M had

an average bursa weight of 01.44g, 1.73g, and 1.67g, respectively. SPAFAS

layers that received identical SHAM challenge had an average bursa weight of

1.41g.





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IBDV AL-2 challenge upon 18 days of age resulted in maternally immune

Leghorns from companies T, C, and M having an average total bursa weight of

0.794g, 0.659g, and 0.689.6g, respectively. SPAFAS layers that received

identical IBDV AL-2 challenge had an average bursa weight of 0.346g.

IBDV DVE challenge upon 18 days of age resulted in maternally immune

Leghorns from companies T, C, and M having an average total bursa weight of

1.14g, 1.25g, and 0.952g, respectively. SPAFAS layers that received identical

IBDV DVE challenge had an average bursa weight of 0.350g.

One-week post SHAM challenge on Day 18, maternally immune commercial

Leghorns and SPAFAS layers were treated identically as same age challenged

chickens. Total bursa weights were measured. Commercial Leghorns from T, C,

and M had an average bursa weight of 1.68g, 1.98g, and 1.79g, respectively.

SPAFAS layers that received identical SHAM challenge had an average bursa

weight of 1.61g.

Gross Bursa Scoring

Upon necropsy, gross bursa scoring was performed based on the following

scale: (0) normal, (1) edematous, and (2) atrophy. Results are summarized in

Tables 3-4 and 3-5. IBDV AL-2 challenge on Day 16 resulted in maternally

immune Leghorns from company T having 13/55 (23.6%) with a score of 1 and

42/55 (76.4%) with a score of 2. Company C had 2/51 (3.9%) with a score of 1

and 49/51 (96.1%) with a score of 2. Company M had 4/50 (8.0%) with a score

of 1 and 46/50 (92%) with a score of 2. All SPAFAS layers had (12/12) with a

score 1(100%).





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IBDV DVE on Day 16 resulted in maternally immune Leghorns from

company T having 25/55 (45.5%) with a score of 1 and 29/59 (49.2%) with a

score of 2. Company C

had 13/51 (25.5%) with a score of 1 and 38/51 (74.5%) with a score of 2.

Company M had 15/53 (28.3%) with a score of 1 and 38/53 (71.7%) with a score

of 2. All SPAFAS layers had 11/11 with a score of 2 (100%).

On Day 16, SHAM-challenged Leghorns and layers resulted in maternally

immune Leghorns from company T having no edematous or atrophied bursas and

57/57 (100%) with a score of 0. Company C (0/51) and Company M (52/52) had

identical score 0 results (100%). SPAFAS layers also had normal bursas by gross

bursa scoring (11/11) with a score of 0.

On Day 18, IBDV AL-2 challenged maternally immune Leghorns from

company T had 8/56 (14.3%) with a score of 1 and 48/56 (85.7%) with a score of

2. Company C had 1/52 (1.9%) with a score of 1 and 51/52 (98.1%) with a score

of 2. Company M had 1/52 (1.9%) with a score of 1 and 51/52 (98.1%) with a

score of 2. All SPAFAS layers (10/10) has a score of 2 (100%).

On Day 18, IBDV DVE challenged maternally immune Leghorns from

company T had 26/56 (46.4%) with a score of 1 and 30/56 (53.6%) with a score

of 2. Company C had 22/51 (43.1%) with a score of 1 and 29/51 (56.9%/) with a

score of 2. Company M had 9/52 (17.3%) with a score of 1 and 43/52 (82.7%)

with a score of 2. All SPAFAS layers (10/10) has gross bursal score of 2 (100%0/).

SHAM-challenged Leghorns and layers on Day 18, maternally immune

Leghorns from company T had no edematous or atrophied bursas (0/54).





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Company C (0/52) and Company M (0/51) had identical results. SPAFAS layers

also had normal bursas by gross bursa scoring (0/12). A summary of gross bursa

scoring is shown in Tables 3-4 and 3-5. Gross bursa images of normal, IBDV-

induced edema, and IBDV-induced atrophy are shown in Figure 3-4.

Histopathology Results

Histopathology slides were performed on all IBDV-infected and SHAM-

challenged chicken bursas. The only chickens that did not show any signs of

bursal change were SHAM-challenged. Normal bursas from uninfected chickens

are shown in Figure 3-5. Results of infected chickens resulted in the induction of

bursal changes but to differing degrees, as shown in Figures 3-6 through 3-10.

Discussion

B/B Weight Ratios

One week following challenge on either Day 16 or Day 18, necropsy of

chickens was conducted. B/B weight ratios were calculated for commercial

Leghors and SPAFAS layers challenged with either AL-2 or DVE isolates of

IBDV. Such measurements were compared to unchallenged controls of identical

age and from the same commercial producer.

In terms of IBDV AL-2 challenge on Day 16, there was no statistical

difference in B/B weight ratios among maternally immune broilers from all

companies. A significant difference in B/B weight ratios was found between

commercial broilers and SPAFAS Leghorns. This difference was expected due to

size differences between broiler and layer chickens. Commercial Leghorns from

companies T, C, and M had average B/B weight ratios of 1.96-, 3.12-, and 2.79-





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times less than those of respective unchallenged controls, respectively. These

results indicate moderate to severe bursal atrophy induced by IBDV AL-2

infection in maternally immune broilers. This is in agreement with in vivo and in

vitro studies demonstrating that the target cell for IBDV is an IgM-bearing B-cell

(Ivanyi and Morris, 1976). Virulent serotype 1 strains of IBDV have a selective

tropism for chicken B-cells and cause marked necrosis of lymphoid follicles

within the bursa. Virus replication leads to extensive lymphoid cell destruction in

the medullary and the cortical regions of the follicles (Tanimura and Sharma,

1998), thereby decreasing B/B weight ratios in infected chickens.

These results also suggest that commercial IBDV vaccination protocols of

dams do not provide chicks with adequate maternally transferred immunity

against IBDV AL-2 challenge. SPAFAS layers that received identical IBDV AL-

2 challenge had B/B weight ratios demonstrating severe bursal atrophy. B/B

weight ratios from challenged layers were 3.71-times lower than those from

unchallenged controls, demonstrating complete lack of protection against IBDV

AL-2 challenge. Results of severe bursal atrophy from challenged SPAFAS

layers were expected since these controls lacked IBDV-specific maternal

immunity.

On Day 16 of age, IBDV DVE challenge resulted in no statistical differences

in B/B weight ratios among maternally immune broilers from all companies and

layers. Commercial broilers from companies T, C, and M demonstrated B/B

weight ratios of 1.51-, 1.98-, and 1.75-times less than those of respective

unchallenged controls, respectively. These results indicate that moderate to





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severe bursal atrophy was induced by IBDV DVE infection. Additionally, these

results suggest that commercial IBDV vaccination protocols of dams do not

provide chicks with adequate maternally transferred immunity. SPAFAS layers

that received identical IBDV DVE challenge had B/B weight ratios demonstrating

severe bursal atrophy. B/B weight ratios from challenged layers were 4.77-times

lower than those from unchallenged controls, demonstrating complete lack of

protection against IBDV DVE challenge. Results of B/B weight ratios from

challenged layers were expected since SPAFAS layers were used as controls

without IBDV-specific maternal immunity.

On Day 18 of age, IBDV AL-2 challenge resulted in no statistical differences

in B/B weight ratios among maternally immune broilers from all broiler

companies and SPAFAS layers. Commercial Leghorns from companies T, C, and

M demonstrated B/B weight ratios of 2.02-, 2.34-, and 2.76-times less than those

of respective unchallenged controls, respectively. These results indicate moderate

to severe bursal atrophy was induced by IBDV AL-2 infection. Additionally,

these results suggest that commercial IBDV vaccination protocols of dams do not

provide chicks with adequate maternally transferred immunity. SPAFAS layers

that received identical IBDV AL-2 challenge had B/B weight ratios demonstrating

severe bursal atrophy. B/B weight ratios from challenged layers were 4.27-times

lower than those from unchallenged controls, demonstrating complete lack of

protection against IBDV AL-2 challenge. However, results from challenged

layers are expected since SPAFAS layers were used as controls without IBDV-

specific maternal immunity.





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On Day 18 of age, IBDV DVE challenge resulted in no statistical differences

in B/B weight ratios among maternally immune broilers from all companies and

SPAFAS layers. Commercial broilers from companies T, C, and M demonstrated

B/B weight ratios of 1.45-, 1.36-, and 1.97-times less than those of respective

unchallenged controls, respectively. These results indicate moderate to severe

bursal atrophy was induced by IBDV DVE infection. These results also suggest

that commercial IBDV vaccination protocols of dams do not provide chicks with

adequate maternally transferred immunity. SPAFAS layers that received identical

IBDV DVE challenge had B/B weight ratios demonstrating severe bursal atrophy.

B/B weight ratios from challenged layers were 4.41-times lower than those from

unchallenged controls, demonstrating complete lack of protection against IBDV

DVE challenge. As mentioned above, results from challenged layers are expected

since SPAFAS layers were used as controls without IBDV-specific maternal

immunity.

Comments Related to Day 16 Versus Day 18 IBDV Challenge

In addition to comparing maternally immune broilers receiving identical

treatments, B/B weight ratios were also compared among chickens receiving

identical challenge on Days 16 and 18. This was done with the understanding that

the half-life of maternal antibodies in the commercial broiler is 3.5 days and by 16

to 18 days, these titers would be declining but still critical in protecting against

late subclinical IBD. Average B/B weight ratios of broilers from company T

challenged with IBDV AL-2 on either Day 16 or 18 was not significantly

different (p=0.4902). B/B weight ratios of broilers from company C challenged





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with IBDV AL-2 on either day 16 or 18 were not significantly different

(p=0.3319). B/B weight of broilers from company M challenged with IBDV AL-

2 on either day 16 or 18, B/B weight ratios were not significantly different

(p=0.9438). In terms of SPAFAS Leghorns challenged with IBDV AL-2 on

either day 16 or 18, B/B weight ratios were not significantly different (p=0.0796).

B/B weight ratios of broilers from company T challenged with IBDV DVE on

either day 16 or 18, B/B weight ratios were not significantly different (p=0.9967).

B/B weight of broilers from company C challenged with IBDV DVE on either

day 16 or 18 were significantly different (p=0.0172), thereby demonstrating a

decrease in B/B weight ratios from Days 16 to 18. B/B weight ratios of broilers

from company M challenged with IBDV DVE on either day 16 or 18 were not

significantly different (p=0.2183). SPAFAS Leghorns challenged with IBDV

DVE on either day 16 or 18 had average B/B weight ratios that were not

significantly different (p=0.7026).

B/B weight ratios of broilers from company T SHAM-challenged on either day

16 or 18 was not significantly different (p=0.4961). B/B weight ratios of broilers

from company C SHAM-challenged on either day 16 or 18 were not significantly

different (p=0.5934). B/B weight ratios of broilers from company M SHAM-

challenged on either day 16 or 18 were not significantly different (p=0.6883).

SPAFAS Leghorns SHAM-challenged on either day 16 or 18 had average B/B

weight ratios that were not significantly different (p=0.6536). Therefore, these





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results show that only challenged chickens have decreased B/B weight ratios in

comparison to unchallenged controls. The decrease in B/B weight ratios was

shown to occur on either challenge day.

Body Weight

Chicken body weight averages, as shown in Figure 3-2, were needed to

compare and evaluate chicken growth among commercial broiler stains of

chickens between Day 16 and Day 18 of challenge. Body weights indicated that

chickens were in the projected range for their respective strain of broiler.

Bursa Weight

Chicken bursa weight averages, as shown in Figure 3-3, were needed to

compare and evaluate bursal development and damage among commercial broiler

strains of chickens between Day 16 and Day 18 of challenge. Bursas

demonstrating edema weighed more than bursas that were atrophied by viral

infection. These results further support the decrease in B/B weight ratios in

challenged chickens.

Gross Bursal Scoring

All IBDV AL-2 or DVE challenged broilers and layers demonstrated changes

in gross bursal scoring. However, it should be noted that even edematous-staged

bursas (Score 1) displayed moderate histopathologic changes in the bursa. In

addition, edema can increase the B/B weight ratio by the presence of a heavier

bursa, thereby yielding false negative results by B/B weight ratio testing.

Upon necropsy, gross bursa scoring was performed based on the following

scale: (0) normal, (1) edematous, and (2) atrophy. In terms of IBDV AL-2