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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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Elyar, John Steven
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vii, 349 leaves : ill. ; 29 cm.

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Antibodies ( jstor )
B lymphocytes ( jstor )
Cells ( jstor )
Chickens ( jstor )
Diseases ( jstor )
Infections ( jstor )
Infectious bursal disease virus ( jstor )
Interferons ( jstor )
Lymphocytes ( jstor )
Vaccinations ( jstor )
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bibliography ( marcgt )
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non-fiction ( marcgt )

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Thesis (Ph. D.)--University of Florida, 2005.
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Includes bibliographical references.
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Printout.
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Vita.
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by John Steven Elyar.

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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.














ii














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




iii








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




iv








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




v








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












vi








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).





58


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





78


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





79


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





81


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).





82


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





83


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





84


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





85


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%).





87

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).





88


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-





89


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





90


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.





91


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





92


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





93


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




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FILES


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

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.
n

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 of b-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 EBDV studies 23
2 IBDV LITERATURE REVIEW 26
IBDV introduction 26
Evolution Of IBDV 27
iii
EBDV Antigenic Variation
28

Virulent EBDV 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
IV

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
v

Results 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
vi

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 of lymphocytes. 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
l

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 Fabricáis
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 of B-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. Flence, 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 V»DJh

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 (i|/V) 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 (vj/Vl). Likewise the heavy chain Vh
sequences are replaced with \)/Vh 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 v[/V genes,
(2) gene conversion events can accumulate within single function Vl or Vh genes
and (3) different donor v/Vl or v|/Vji 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 (slg)
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
10s 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 of B-
cells in the periphery, although the physiological basis for this heterogeneity
remains speculative (Paramithiotis and Ratcliffe, 1993, 1996).
Avian Immunoglobulin Isotvpes and Their Peripheral Roles
Chicken Immunoglobulin Class M (IgMI
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 p-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 C« 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 CaIg 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-Lvmphocvtes
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 of hematopoiesis. 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-lvmphocyte Ontogeny
The BF develops as an outgrowth of cloacal epithelium and is seeded between
ED8 and ED 14 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 of Ig 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 of thymulin 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 ED 16 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 bv 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
Oncomaviridae 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 Vims
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).
Reo viruses
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
USD A 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 EBDV 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 of immunosuppression, 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
Bimaviridae (Dobos, 1979). IBDV is classified in the Avibimavirus genus within
the family Bimaviridae (Murphy and Johnson et al. 1995). Other members of
Bimaviridae include IPNV as an Aquabimavirus, tellina and oyster viruses of
mollusks, and Drosophila X vims of fruit flies (Drosophilia melanogaster) as an
Entromobimavirus. The taxonomic relationship of other Bimaviruses 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 EBDV, 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 aquabimavirus may have survived
incomplete processing of meal. A subsequent study by Dobos et al. (1979)
confirmed a close relationship only between the three aquabimaviruses (IPNV,
tellinavirus, and oyster virus of mulluscs), which can be differentiated from
IBDV, and Drosophila vims 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
of IBDV 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
(wIBDV) have been reported in many European, African, and Asian countries.
The emergence of wIBDVs 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 wIBDV 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, wIBDV has yet to be reported from North America or Australia.
Contrary to the situation in the US with variant IBD Vs, the wIBDV 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
wIBDV 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 wIBDVs (Brown et al.

31
1994; Brown and Skinner, 1996; Lin, et al. 1993; Vakharia et al. 1994). Studies
with Japanese wEBDVs 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 EBDVs, 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 wEBDVs. However, variations in RFLP patterns are not necessarily
an indication of antigenic variation or immunogenicity of variant and wDBDV.
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 EBD 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 wEBDV 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 wIBDVs 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 wIBDVs have not been demonstrated to clearly
influence antigenicity (Brown et al. 1994; Lin et al. 1993).
As wIBDVs 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 wIBDV, 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 wIBDV 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 EBDV. 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 DBDV 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, ELIS As

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
Bimaviridae (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-EBDV 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 of RT-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.
Bimaviruses 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 (Garoff et 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 of VP2. 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 EBDV 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 of immunoglobulin 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 IOOOCID50 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 EBDV 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 Uve virus, either in the form of active
vaccination or field exposure to the EBDV. 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 wIBDV, 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 56°C. The virus was
unaffected by exposure to 0.5% phenol and 0.125% thimerosal for 1 hour at 30°C.
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 23 °C. Only the iodine complex had any
deleterious effects. Landgraf et al. found that the virus survived 60°C but not
70°C 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 EBDV. 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).
EBDV 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 (slgM), 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 EBDV 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 slgM. 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 slgM-bearing cells.
Most of the virus-binding cells observed in the study by Ogawa et al. (1998)
were slgM-bearing B-cells. Results from another study also reported that virulent
IBDV infected slgM-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 slgM-bearing B-lymphocyte, indicating
the possibility that the slgM molecule may be the receptor for virulent IBDV
attachment. However, in this study, bound virus particles were also observed in
the slgM-negative cells even though their number was small. Interestingly, the
binding of the virus to slgM-bearing B-cells was not inhibited by anti-IgM
antibody. These additional findings may show that IBDV does not solely utilize
the slgM 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 slgM-positive cells. Previously, several
B cell-specific surface antigens other than slgM were reported (Olson and Ewert,

48
1990), which appeared to closely parallel the expression of the slgM 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 EBDV infection was observed only in cell lines of slgM-
bearing cells, but not in cell lines of slgM-negative cells (Hirai and Calnek, 1979).
This result may indicate that the slgM 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 slgM-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 of B cells bearing surface immunoglobulins M and G (slgM and
slgG) was studied in the spleen and peripheral blood of chickens infected with
IBDV. The proportion of surface immunoglobulin-bearing B-cells and slgM and
slgG-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 EBDV 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 EBDV-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
EBDV. Commercial chicken flocks commonly experience recurring losses due to
EBDV-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 of thymus-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 200(ig of Tetanus toxoid (TTX) and 150|iG of Brucella
abortus in Freund’s incomplete adjuvant. At 10 days after antigenic stimulation,
chickens were examined for levels of anti-TTX and anti-5, 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 of T-cells into this organ (Kim et al.
1999). In EBDV-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 EBDV-induced bursal T-lymphocytes were T-cell
receptor 2-expressing (TCR2+) a/(3-T cells, and a few were TCR1+ y/8 T-cells
(Sivanandan and Maheswaran, 1980).
In a study by Kim IJ 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 of T-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 la 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 EL-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 IIFN, 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 (Perfile 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 slgM 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).

58
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 IBP V 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.
EBDV-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
i

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 of apoptosis 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-ddetion
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 bursal cells The
involvement of indirect mechanisms was suggested by Inoue et al. (1994) since
apoptosis was observed in T-cells of the thymus of infected chickens, whereas
IBDV antigens were found mainly in infiltrated B-cefls or in reticular cells

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. Bimaviruses 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 (Landgraf et 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 EBD. 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 100%
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 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 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 Gamer, 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 vims to the bursa.
It is noteworthy that bursectomized chicks do not show clinical signs following
infection with pathogenic strains of EBDV. Due to the absence of host cells, vims
multipliation is inhibited, although EBDV can be re-isolated from spleen, thymus,
and liver up to 5 days after infection in bursectomized chicks. The concentration
of vims is only 10'3 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 of B-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 slgM-positive B-cells
dropped precipitously as the vims 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 vims 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 of intra-bursal T-cells were present at 7
days pi. At peak accumulation, 65% of the bursal cells were T-cells and 7% had
slgM expression. Although CD4-positive and CD8-positive lymphocytes were
roughly in equal proportions during the first 7 days pi, CD8-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 slgM-positive lymphoid cells. By 12 weeks pi, almost all bursal
follicles had been replete with slgM-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 of T-
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 EBDV 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 of IBDV 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 slgM-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 of thymocytes 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 slgM-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 slgM 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
slgM-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 of T-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 DL-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-ffee
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 II and EL-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, of tryptose phosphate broth is treated with antibiotics and

73
centrifuged. Vims 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 vims 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 Bourn’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). flBDV-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

78
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:
99.5°F and 60% relative humidity. Subsequently, all eggs were hatched under the
following conditions: 98.5°F 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 (88°F)
rooms. Each room contained a single chicken cage battery. On Day 6, room
temperature was gradually lowered to 78°F 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

79
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 120°F for a period of 72 hours. A subsequent bleaching
(1:2) process was performed with reheating the room to 120°F for 72 hours. A
final disinfecting process was performed on sealed rooms by para-formaldehyde
fumigation. This was performed by combing potassium permanganate (KMn04)
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 5EID5o 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.

80
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 5 pm
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

81
age of 16 or 18 days with either AL-2 or Del Variant E isolates of IBDV.
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 (pO.OOOl).
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 (pO.OOOl).

82
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, 0.28lg, respectively. SPAFAS

83
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 EBDV 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

84
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

85
678.7g, respectively. Identically challenged SPAFAS layers had average total
body weight of 229. lg.
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 -4 lg.

86
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 1.61 g.
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%).

87
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, EBDV 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%).
SHAM-challenged Leghorns and layers on Day 18, maternally immune
Leghorns from company T had no edematous or atrophied bursas (0/54).

88
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
Flistopathology 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
Leghorns and SPAFAS layers challenged with either AL-2 or DVE isolates of
EBDV. 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-

89
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 w 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

90
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 EBDV-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.

91
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

92
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

93
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

94
challenge on Day 16, maternally immune broilers from all 3 commercial
producers received a gross bursa score of either 1 or 2. No broilers received a
score of 0. Gross bursa scoring from company T had 3.24-times as many Score 2
than Score 1. Company C had 24.6-times as many Score 2 than Score 1.
Company M had 11.5-times as may Score 2 and Score 1. All SPAFAS layers had
(12/12) with a Score 2 bursas (100%).
IBDV Delaware Variant E challenge on Day 16 resulted in maternally immune
broilers from all 3 commercial producers having gross bursa scores of ether 1 or
2. Broilers from company T had 1.0-times more Score 1 than Score 2. Broilers
from company C had 2.9-times more Score 2 than Score 1. Broilers from
company M had 2.5-times as many Score 2 than Score 1. All SPAFAS layers
(11/11) had with a score of 2 (100%).
SHAM-challenged Leghorns and layers on Day 16 resulted in scores from
company T having 1.0-times as many edematous or atrophied bursas and 57/57
(100%) with a score of 0. Company C (51/51) 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.
IBDV AL-2 challenge on Day 18 resulted in maternally immune broilers from
company T having 6.0-times more Score 2 than Score 1. Broilers from company
C had 51.6-times more Score 2 than Score 1. Broilers from company M had 51.6-
times as many Score 2 than Score 1. All SPAFAS layers (10/10) has a score of 2
(100%).

95
IBDV DVE challenge on Day 18 resulted in maternally immune broilers from
company T having 1 16-times more Score 2 than Score 1. Broilers from company
C had 1.3-times more Score 2 and Score 1. Broilers from company M had 4.8-
times more Score 2 than Score 1. All SPAFAS layers (10/10) have gross bursal
score of 2 (100%).
SHAM-challenged Leghorns and layers on Day 18 resulted in maternally
immune Leghorns from company T having no edematous or atrophied bursas
(0/54). Company C (0/52) and Company M (0/51) also had no edematous or
atrophied bursas. SPAFAS layers also had normal bursas by gross bursa scoring
(0/12). Summaries of gross bursa scoring are 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. In this study, gross bursa scoring detected infected bursas
based on edema from chickens that displayed normal B/B weight ratios.
Histopathologv
Histopathology demonstrated that all IBDV-challenged chickens displayed
signs of bursal change. These severe changes were noted in infected chickens
with a gross bursa score of 2 (atrophy). The remaining infected chickens with a
gross bursa score of 1 (edema) showed moderate bursal changes. Therefore, all
IBDV challenged SPAFAS layers and maternally vaccinated broiler chickens
from 3 commercial poultry companies had notable bursal pathologic change.
Changes in bursal histopathology detected infected bursas that were not noted by
B/B weight ratios and gross bursal scoring. Therefore, it was determined that the

96
B/B weight ratio system, as recommended by the USD A, and gross bursa scoring
are not as accurate a measurement of IBDV-induced bursal pathology as
compared to histopathologic study.

97
Table 3-1: Animal Design for IBDV Experiments
Total Number
Blood
Total Number
Age of
BIB Weight3
IBDV
of Chickens
Collection
of Chickens
Challenge
Challenge
Challenge
7 days Plb
Group
Placed
(Numbers)
Challenged
(Days)
Virus
Dose
# of birds
100
0
50
16
AL2
10 35
50
I
50
18
AL2
icF5
50
100
n
50
16
Del Variant
10 3 5
50
I
50
18
Del Variant
10 35
50
150
50
N/A
16
Neg Controls
N/A
50
I
N/A
18
Neg Controls
N/A
50
100
0
50
16
AL2
10 3.5
50
50
18
AL2
10 35
50
100
0
50
16
Del Variant
10 35
50
50
18
Del Variant
10 35
50
150
50
N/A
16
Neg Controls
N/A
50
N/A
18
Neg Controls
N/A
50
M
100
n
50
16
AL2
10 35
50
50
18
AL2
10 35
50
M
100
n
50
16
Del Variant
10 3 5
50
50
18
Del Variant
10 35
50
M
150
50
N/A
16
Neg Controls
N/A
50
N/A
18
Neg Controls
N/A
50
10
16
AL2
10
10
10
18
AL2
10 3 5
10
SPAFAS
70
10
10
16
Del Variant
10 35
10
10
18
Del Variant
10 35
10
10
16
Neg Controls
N/A
10
10
18
Neg Controls
N/A
10
aNumber of chickens from which BIB Weight Ratios were calculated.
bPost-infection

98
Table 3-2. Timeline for Study IBDV
Egg Hatch
Challenge 18d
Term, dl 8
Set Egg
s
Challen^
ie 16d
Term, d
16
Day -
21
Day
16
Day
29
Day 0
Day 18
Day 31

99
Table 3-3. Room & Cage Groupings for IBDVQ03
Room #2 - 160 birds
Day 16 with AL-2
T- 16
T- 17
T- 17
MM
C- 17
mm
M - 17
EmpT
EmpT
SPAFAS -10
Room #3-160 birds
Day 18 with AL-2
T- 16
T- 17
T- 17
C-16
C-17
C-17
M-l 6
1-17
EmpT
EmpT
SPAFAS-10
Room #4-160 birds
Day 16 with Variant E
T- 16
T- 17
T- 17
C-16
c-«
^M~
M- 16
fr-17
M - 17
EmpT
EmpT
SPAFAS -10
Room #5-160 birds
Day 18 with Variant E
T-16
T-17
T- 17
£-17
C-17
M - 16
M - 17
M - 17
EmpT
EmpT
SPAFAS - 10
Room #6-210 birds
Day 16 (Ctrol)
T- 16
T- 17
T- 17
C-16
C-17
WK—
â– 
M- 17
M - 17
T- 25 (17d)
T- 25 (17d)
SPAFAS -10

Table 3-4. Gross Bursa Scoring Results
Age at
Challenge
(Days)
Virus
Strain
Commercial
Chicken
Source
Total
Number
of
Normal
Bursas
(0)
Total
Number of
Edematous
Bursas
(1)
Total
Number of
Atrophied
Bursas
(2)
Total
Number
of
Chickens
16
AL-2
T
0
13
42
55
C
0
49
2
51
M
0
4
46
50
SPAFAS
0
0
12
12
Delmarva
Variant E
T
0
26
29
59
C
0
13
38
51
M
0
15
38
53
SPAFAS
0
0
11
11
SHAM
Challenge
T
54
0
0
57
C
52
0
0
51
M
51
0
0
52
SPAFAS
11
0
0
11

101
Table 3-5. Gross Bursa Scoring Results
Age at
Challenge
(Days)
Virus
Strain
Commercial
Chicken
Source
Total
Number
of
Normal
Bursas
(0)
Total
Number
of
Edematou
s Bursas
(1)
Total
Number of
Atrophied
Bursas
(2)
Total
Number
of
Chickens
18
AL-2
T
0
8
48
56
C
0
1
51
52
M
0
1
51
52
SPAFAS
0
0
11
11
Delmarva
Variant E
T
0
26
30
56
C
0
22
28
50
M
0
9
43
52
SPAFAS
0
0
10
10
SHAM
Challenge
T
54
0
0
54
C
52
0
0
52
M
51
0
0
51
SPAFAS
12
0
0
12

102
Figure 3-1. Bursa/Body Weight Ratios for Chicks Challenged with either IBDV
AL-2 or D'VE compared to SHAM Chicks on either Day 16 or Day 18.

103
Day 16 Day 18
Figure 3-2. Total body weight averages from chicks challenged with either
IBDV AL-2 or D'VE compared to SHAM chicks on either Day 16 or 18.
I

(grams)
104
Day 16
Day 18
â–¡ T
â–¡ C
â–¡ M
â–¡SPAFAS
Figure 3-3. Total bursa weight averages for chicks challenged with either IBDV
AS-2 or D'VE compared to SHAM chicks on either Day 16 or Day 18.

105
Figure 3-4. Gross visual observation of normally healthy (NH), severe atrophy
(SA), and edematous (E) bursas.

106
Figure 3-5. Uninfected bursa showing healthy infrastructure (Magnification at
100X).

107
Figure 3-6. Onset of IBDV-induced bursal edems (Magnification at 100X).
Early edema noted by a black arrow.

108
Figure 3-7. Significant cellular infiltration in bursal follicles due to
IBDV infection (Magnification at 100X).

109
Figure 3-8. Edematous Bursa due to IBDV-infection along with partial
follicular destruction (Magnification at 100X). Follicular destruction is
noted by a black arrow.

110
Figure 3-9. Destruction of bursal follicles due to infection with
IBDV (Magnification at 100X). Destroyed follicles are noted by
black arrows.

CHAPTER 4
CHALLENGE OF SPAFAS LAYER AND MATERNALLY IMMUNE
BROILER CHICKENS ON DAY 16 OR 18 WITH A NEWLY ISOLATED
IBDV STRAIN DESIGNATED IBDV-R
Introduction
Since the mid-1980s, highly pathogenic IBDV strains designated very
virulent (wIBDV) have been reported in many European, African, and Asian
countries. The emergence of wIBDVs 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 specific-pathogen-free (SPF) chickens
were reported in Japan (Nunoya et al. 1992). The wIBDV strains were reported
to break through high levels of maternal antibodies in commercial flocks, causing
up to 60% to 100% mortality rates in chickens and producing lesions typical of
IBDV (Chai et al. 2001). 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 (Chai et al. 2001;
Chettle and Wyeth et al. 1989).
To date, wIBDV has yet to be reported from North America or Australia.
Contrary to the situation in the US with variant IBD Vs, the wIBDV European
strains were reported to be antigenically similar to other serotype 1 classic strains
but very different in virulence (Zierenberg et al. 2000).
ill

112
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, 1990). Intensive
vaccination in some areas of the US may have influenced antigenic properties of
field IBDV. European viruses responsible for wIBDV, in contrast, have
increased pathogenicity without demonstrating antigenic shifts (Snyder et al.
1990). To this end, SPAFAS layers and maternally immune broilers were
challenged with a newly isolated strain called IBDV-R.
Project Design
SPF Leghorn and maternally immune broiler eggs from 3-commercial
producers were be incubated, hatched, as described in Chapter 2. Chicks were
challenged only on Day 18 and organized into treatments groups as described in
Table 6. Chicks received ocular challenge with IBDV R at either 1:10 or 1:100
dilutions. Evaluate subclinical infection of SPF Leghorn and maternally immune
broilers with a newly isolated IBDV variant, IBDV-R. Challenge SPF and
maternally immune broiler chickens with EBDV isolate. Broilers from 3-
commercial producers were challenged on Day 18 with either 1:10 or 1:100
dilutions of IBDV R isolate. Chickens were evaluated for subclinical effects 1-
week pc (Day 25) using identical parameters as described in Chapter 3.
Materials and Methods
Experimental Animals
Specific-pathogen-free (SPF) Leghorn and maternally immune broiler eggs
were obtained from 3-commercial producers: T, C, and M.

113
Incubation and Hatching Conditions
Incubation of all eggs to place for 16 days were subjected to the following
conditions: 99.5°F and 60% relative humidity. Subsequently, all eggs were
hatched under the following conditions: 98.5°F and 65% relative humidity. All
day-old chicks were neck-banded with a 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
(88°F) rooms. Each room contained a single chicken cage battery. On Day 6,
room temperature was gradually lowered to 78°F to enhance feed consumption.
Chicks were allowed to drink fresh water and eat “starter feed” ad lib.
Location of EBDV 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
at the University of Florida in Gainesville, Florida. Chickens receiving different
treatments were housed in order, as shown in Table 7 and 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 120°F for a period of 72 hours. A subsequent bleaching

114
(1:2) process was performed with reheating the room to 120°F for 72 hours. A
final disinfecting process was performed on sealed rooms by para-formaldehyde
fumigation. This was performed by combing potassium permanganate (KMn04)
and 10% formalin to create gaseous para-formaldehyde.
Experimental IBDV Challenge
IBDV-R was 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 5EID5o 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.
Gross bursa scoring
A simple gross bursa scoring system was applied according to the following
scale: 0 (normal), 1 (edematous), and 2 (atrophic).
Histopathologv
Briefly, bursas were stored in 10%-buffered formalin and cut into 5 pm
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 were
calculated. To determine whether a significant difference existed among ratios

115
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 were 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 compared to gross bursa scoring and histopathology.
Results
B/B Weight Ratios
One week after respective challenge, necropsy of all chickens was performed.
Average bursa to total body weight ratios were calculated upon necropsy of
commercial broilers and SPAFAS layers challenged at the age of 18 days with
either a 1:10 or 1:100 dilution of IBDV-R. Unchallenged controls were compared
with challenged chicks of identical age. Summary of B/B weight ratios is shown
in Figure 4-1.
On Day 18, another group of commercially obtained Leghorn and SPAFAS
layer chicks were challenged with recently isolated IBDV-R strain. All chickens
were challenged with either a 1:10 or 1:100 dose of virus. Leghorns challenged
with 1:10 IBDV-R from companies T, C, and M had average B/B weight ratios of
0.136g, 0.0.134, and 0.159, respectively. Leghorns challenged with 1:100 IBDV-
R from companies T, C, and M had average B/B weight ratios of 0.203, 0.239,

116
and 0.218, respectively. SPAFAS layer chicks challenged with either a 1:10 or
1:100 dilution had average B/B weight ratios of 0.182 and 0.221, respectively. In
addition, the correlation between virus dilution and bursal atrophy can be noticed.
However, even broilers challenged with 1:100 did display bursal atrophy in
comparison to unchallenged controls.
One-week post SHAM challenge on Day 18, maternally immune commercial
broilers and SPAFAS layers were treated identically as same age challenged
chickens. B/B weight ratios were calculated. Broilers from T, C, and M had B/B
weight ratios of 0.266, 0.285, 0.281, respectively. SPAFAS layers that received
identical SHAM challenge had B/B weight ratio of 0.671.
Body Weight Averages
A summary of body weight results is shown in Figure 4-2. Necropsy results of
commercial Leghorns from T, C, and M challenged with either dilution of IBDV-
R on day 18. Necropsy results of commercial Leghorns from T, C, and M
SHAM-challenged on day 18 demonstrated 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.
On Day 18, another group of commercially obtained Leghorn and SPAFAS
layer chicks were challenged with recently isolated IBDV-R strain. All chickens
were challenged with either a 1:10 or 1:100 dose of virus. Broilers challenged
with 1:10 IBDV-R from companies T, C, and M had average body weights of
623.lg, 652.8g, and 803.6g, respectively. Broilers challenged with 1:100 IBDV-
R from companies T, C, and M had average body weights of 631,6g, 613.2g, and

117
851 .Og, respectively. Among broilers from companies T, C, and M and SPAFAS
layers that received a 1:10 IBDV-R challenge, there were no significant pair wise
differences among the means (p=0.2944). Among broilers from companies T, C,
and M and SPAFAS layers that received a 1:100 IBDV-R challenge, there were
no significant pair wise differences among the means (p=0.1141). SPAFAS layer
chicks challenged with either a 1:10 or 1:100 dilution had average body weights
of 209.7g and 219.9g, respectively.
One week post SHAM challenge on Day 18, maternally immune commercial
broilers and SPAFAS layers were treated identically as same age challenged
chickens. Average body weights were calculated. Broilers form T, C, and M had
average body weights of 0.266, 0.285, and 0.281, respectively. SPAFAS layers
that received identical SHAM challenge had an average body weight of 0.671.
1:10 dilution of IBDV-R versus SHAM-treated chickens resulted in significant
differences in average body weights between challenged and unchallenged
broilers from companies T, C, and M (p=<0.0001, pO.0001, p=0.0089,
respectively). 1:100 dilution of IBDV-R versus SHAM-treated chickens resulted
in significant differences in average body weights between challenged and
unchallenged broilers from companies T, C, and M (P=0.0066, p=0.0179, and
p<0.0001, respectively). SPAFAS layers challenged with either a 1:10 or 1:100
dilution of IBDV-R had average body weights that were significantly different
from unchallenged SPAFAS layers (p<0.0001 and p<0.0001, respectively).
In terms of average body weights among challenge with either 1:10 or 1:100
dilution of IBDV-R, broilers from companies T and C had significant differences

118
(p=0.0137 and p<0.0001, respectively). A significant difference was not found
among company M broilers (p=0.0748). However, it should be noted that this p
value is close to (p=0.3464).
Bursa Weight Averages
Summary of bursa weight results is shown in Figure 4-3. 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
average bursa weights of 1 68g, 1 98g, and 1,79g, respectively. SPAFAS layers
that received identical SHAM challenge had an average bursa weight of 1.61 g.
On Day 18, another group of commercially obtained Leghorn and
SPAFAS layer chicks were challenged with recently isolated IBDV-R strain. All
chickens were challenged with either a 1:10 or 1:100 dose of virus. Leghorns
challenged with 1:10 IBDV-R from companies T, C, and M had average bursa
weights of 0.845g, 0.866g, and 1.30g, respectively. Leghorns challenged with
1:100 IBDV-R from companies T, C, and M had average bursa weights of 1,27g,
1.48g, and 1.81 g, respectively. SPAFAS layer chicks challenged with either a
1:10 or 1:100 dilution had average bursa weights of 0.390g and 0.482g,
respectively.
Gross Bursa Scoring
Upon necropsy, gross bursa scoring was performed based on the following
scale: (0) normal, (1) edematous, and (2) atrophy. Summary of gross bursa
scoring is shown in Table 8. In terms of SHAM-challenged Leghorns and layers

119
on Day 16, maternally immune Leghorns from company T had no edematous or
atrophied bursas (0/50). Company C (0/52) and Company M (0/51) had identical
results. SPAFAS layers also had normal bursas by gross bursa scoring (0/12).
On Day 18, another group of commercially obtained Leghorns and SPAFAS
layers were challenged with recently isolated IBDV-R strain. All chickens were
challenged with either a 1:10 or 1:100 dose of virus. Leghorns challenged with
1:10 IBDV-R from company T had 2/16 (12.5%) edematous and 14/16 (87.5%)
atrophied bursas. Company C had 3/16 (18.75%) edematous and 13/16 (81.25%)
atrophied bursas. Company M had had 4/9 (44.44%) edematous and 5/9
(55.56%) atrophied bursas. SPAFAS layers had 3/5 (60%) edematous and 2/5
(40%) atrophied bursas.
Leghorns challenged with 1:100 IBDV-R from company T had 8/14 (57.14%)
edematous and 6/14 (42.86%) atrophied bursas. Company C Leghorns had 11/15
(73.33%) edematous and 4/15 (26.66%) atrophied bursas. Company M had 2/9
(22.22%) edematous and 7/9 (77.78%) atrophied bursas. SPAFAS layers had 2/4
(50%) edematous and 2/2 (100%) atrophied bursas.
Histopathology
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 animals. Normal bursas from uninfected
chickens are shown in Figure 4-1. Results of infected chickens all induced bursal
changes but to differing degrees, as shown in Figures 3-6 through 3-9.

120
Discussion
B/B Weight Ratios
One week after challenge on Day 18 with IBDV-R, necropsy of all chickens
was performed. Bursa to total body weight ratios were calculated for commercial
Leghorns and SPAFAS layers. Such measurements were compared to
unchallenged controls of identical age, as shown in Figure 4-1. These results are
similar to those from other in vivo and in vitro studies (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
EBDV 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.
Body Weight
Chicken body weight averages, as shown in Figure 4-2, were needed to
compare and evaluate chicken growth among the commercial strains of broilers
on Day 18 challenge.
Bursa Weight
Chicken bursa weight averages, as shown in Figure 4-3, were needed to
compare and evaluate bursal development and damage among commercial broiler
strains on Day 18 challenge.

121
Gross Bursal Scoring
All IBDV-R challenged broilers and layers displayed changes related to gross
bursal scoring. However, it should be noted that some bursas had edematous
(Score 1) lesions found on histopathologic study, as shown in the following
section. Edema would actually increase the B/B weight ratio due to the presence
of a heavier and enlarged bursa. Most bursas had undergone the classical atrophy
following IBDV challenge, however, in individual birds; there were some bursas
with edema. Summary of gross bursal scoring is shown in Table 4-3. Gross
bursal scoring results demonstrated infected bursas that were not detected by B/B
weight ratios.
Histopathologv
Histopathology showed that all IBDV-challenged chickens displayed signs of
bursal change. Many of these severe changes were noted in all infected chickens
with a gross bursa score of 2 (atrophy). The remaining infected chickens with a
gross bursa score of 1 (edema) had bursal changes. Therefore, all IBDV
challenged SPAFAS Leghorns and maternally vaccinated broiler chickens from 3
commercial poultry companies showed notable bursal pathology as observed from
gross morphology and histopathologic study. Therefore, it can be concluded that
the B/B weight ratio system, as recommended by the USD A, and gross bursa
scoring are not as accurate and consistent measurements of IBDV-induced bursal
pathology. As these techniques have been used as the standard of evaluating
vaccine efficacy against IBDV challenge, clearly this system needs to be updated.
This system had assumed that the strains that were classified as variants would

122
undergo atrophy following challenge of susceptible chickens. This work
demonstrated that in a percentage of the experimental birds, there is an
enlargement of the bursa as had been associated only with strains of IBDV
classified as classical-types in the past.

123
Table 4-1: Animal Design for IBDV Experiments
Number of Number of Number of Challenge B/B Weight11
IBDV Birds Birds for Birds Age Challenge Challenge 7 days Plc
Group Placed Collection3 Challenged (Days) Virus Dose # of birds
T
40
0
20
18
New variant
1:10
20
20
New variant
one:hundred
20
C
40
0
20
18
New variant
1:10
20
20
New variant
one:hundred
20
M
40
0
20
18
New variant
1:10
20
20
New variant
one:hundred
20
SPAFAS
20
0
10
18
New variant
1:10
20
10
New variant
one:hundred
20
aBlood Collection
bBursa to body weight ratio
cPost-infection

124
Table 4-2. Room and Cage Groupings for IBDV
Room #8 - 260 birds
Day 18 (Ctrol)
T- 16
T- 17
T- 17
C- 16
C- 17
C- 17
M- 17
^B
C-25 (Id)
C-25
B
Empty
M - 25 (17d)
SPAFAS -10
Room #7-140 birds
Day 18 (New Variant, IBDV 4 Titration)

Table 4-3. Gross Bursa Scoring Results
Age at
Challenge
(Days)
Virus
Strain
Commercial
Chicken
Source
Total
Numbe
r of
Normal
Bursas
(0)
Total
Number of
Edematous
Bursas
(1)
Total
Number of
Atrophied
Bursas
(2)
Total
Number
of
Chickens
18
IBDV-4
@ 1:10
T
0
2
14
16
C
0
3
13
16
M
0
4
5
9
SPAFAS
0
4
2
6
IBDV-R
@ 1:100
T
0
8
6
14
C
0
11
4
15
M
0
2
7
9
SPAFAS
0
2
2
4
SHAM
Challenge
T
54
0
0
54
C
52
0
0
52
M
51
0
0
51
SPAFAS
12
0
0
12

126
IBDV 1:10 IBDV 1:100 SHAM
â–¡ T
ac
DM
â–¡SPAFAS
Figure 4-1. Bursa/Body Weight Ratios for chicks challenged on Day 18
with IBDV-R

127
900 it
Figure 4-2. Total body weight averages from chicks
challenged with either dose of IBDV-R on Day 18.

128
IBDV-R IBDV-R SHAM
(1:10) (1:100)
â–¡ T
â–¡ C
â–¡ M
â–¡ SPAFAS
Figure 4-3. Total bursa weight averages for chicks
challenged with either dose of IBDV-R compared to
SHAM chicks on Day 18.

CHAPTER 5
AVIAN CELLULAR IMMUNITY
Central Organs of Chicken Cellular Immunity
Thymus
Thymic lobes are divided into lobules by fine, connective tissue. Lobules have
a distinct outer cortex composed of densely packed small lymphocytes and an
inner medulla containing less densely packed lymphocytes, reticular cells, and
islands of epithelial cells called Hassall’s corpuscles, variable numbers of
granulocytes are commonly found in these corpuscles (Hudson and Payne, 1973).
T-cell Receptor Repertoire Ontogeny
Chicken T cells have been well characterized using polyclonal and monoclonal
antibody raised against functionally important T cell molecules, such as T-cell
receptors. Three major lineages of chicken T cells have been defined, TCR1,
TCR2, and TCR3, named for their ontogenetic order (Chen et al. 1990).
Ontogeny of T-cell Subsets
The developing chicken thymus exhibits three periods of receptivity to
thymocytes precursor influx, followed by three waves of differentiation of all
three T cell lineages. Chicken yS T cells (TCR1+) are detected at day 11 of
embryogenesis (El 1), and peak in relative cell numbers (30%) at El5, a(3 T cells
expressing V01 (TCR2+) appear at E15, and predominate in the thymus by E17-
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130
18, whereas a(3 T cells expressing VP2 (TCR3+) develop around E18 and are
maintained as a smaller subset (Mueller et al. 1989; Sowder et al. 1988; Lahti et
al. 1991).
Based on developmental parallels with mammals, chicken ap T cell
maturation involves the same positive and negative selection mechanisms as in
mammals (Cooper and Losch, 1991). Chicken ap T cells pass slowly through the
thymic cortex, gradually increase surface TCR expression, and exhibit the same
pattern of CD4/CD8 expression during thymic maturation as for mammalian aP T
cells. In contrast, chicken y8 T cells do not appear to undergo intrathymic
selection, as they migrate quickly from the thymus without clonal expansion,
immediately express high levels of surface TCR and are resistant to receptor
modulation.
In contrast to reports that some mammalian T cells may be of extrathymic
origin, most early chicken T cell development appears to be thymus-dependent
(Mueller et al. 1989; Dunon and Imhof, 1996). Experimental evidence includes
data from chick-quail chimeras (Buey et al. 1989), and from chimeras of congenie
chicken strains differing in the ov alloantigen T cell marker (Dunon et al. 1993a,
b).
The waves of Vpi T cells vary in their homing preferences to peripheral
organs: the first wave preferentially migrates to the spleen, the second wave to
the spleen and intestine, and the third wave and later T cells to the spleen (Dunon

131
et al. 1993a,b; Dunon and Imhof, 1996). Homing of Vpi and V(32 T cells subsets
to the intestine differs markedly given the relative absence of Vp2 T cells in the
intestine (Dunon et al. 1993a).
Ontogeny of TCR Gene Repertoires
Despite low homologies to their mammalian homologues, genes for all four
chicken TCR loci have been cloned. The genomic organizations of all four loci
are not completely known; however, conserved features of mammalian and avian
TCR genes include consensus amino acids for structural requirements, overall
genomic organization, although chicken loci encode fewer gene segments), and
location of TCR 5d genes within the TCR a locus (Kubota et al. 1994). Somatic
diversity of rearranged TCR genes appears to be generated through combinatorial
and junctional mechanisms, as in mammalian TCR genes, rather than by gene
conversion as in avian immunoglobulin genes (McCormack and Thompson, 1990;
Lahti et al. 1991).
The embryonic thymus Vpi repertoire of all three waves of T cell
development shows no evidence for preferential Vpi or jp usage or selection for
CDR3 lengths, and therefore no preselection of Vpi T cells for colonization into
spleen or intestine (Dunon et al. 1993a,b; Dunon and Imhof, 1996).
Distribution of aP and y8 TCR Populations
Migration of T cell subsets from the thymus to the periphery occurs in the
same order as their development, i.e. y8 T cells appear in spleen around E15-17,
Vpl+ aP T cells appear around E19, and VP2+ aP T cells appear at 3 days post
hatching (Sowder et al. 1988; Coltey et al. 1989). Dunon and coworkers

132
(1993a,b; 1996) have shown that emigration of y8 and Vpi T cell subsets into
spleen and intestine occurs in waves, following each wave of thymic T cell
development.
After migration from the thymus, most Vpl+ a(3 T cells express CD4 and are
localized in the splenic periarteriolar sheath and intestinal lamina propria.
Chicken ap T cells are responsive to T cell mitogens, are capable of graft-versus-
host alloreactivity, and CD4+ cells secrete lymphokines (Lahti et al. 1991).
Thymectomy at hatch results in only a small decrease in the numbers of aP T
cells in the periphery, owing to T cell migration from the thymus by the time of
hatching.
Splenic y8 T cells in chickens are localized to the sinusoids of the red pulp and
remain dispersed (Buey et al. 1989). y8 T cells predominate in the intestinal
epithelium, but are minor populations or absent in lamina propria, Peyer’s
patches, cecal tonsils, and normal skin (Buey et al. 1998). Thymic y8 T cells
enter the intestinal epithelium at all levels of the villi just after hatching, and
persist longer than in the spleen due to an apparent higher capacity for self¬
renewal (Dunon et al. 1993a,b). Although y8 T cells in the thymus and blood are
CD4-CD8-, about two-thirds of the yS T cells in the spleen and intestine express
CD8. Intestinal CD8+ cells are further divided into CD8aa and CD8aP subsets
(Tregaskes et al. 1995).
Avian aP T cells respond to in vitro stimulation via mitogens and TCR-cross
linking, but y8 T cells (CD8+) respond only in the presence of CD4+ T cells or
soluble factors derived from them (Arstila et al. 1993; Kasahara et al. 1993), and

133
have an activated phenotype based on size and MHC class II expression (Hala et
al. 1984). Only CD4+ cell (Vf31 and V02) are capable of inducing GVH lesions
in chick embryos, and y5 T cells are recruited by the alloreactive aP T cells (Tsuji
et al. 1996; Sowder et al. 1988). These data suggest mutual regulatory functions
between a.p and yS T cells in avian immune responses (Kasahara et al. 1993).
Function of T-cell Subsets
Chicken Vpi and VP2 T cell subpopulations appear to have some distinct
differences in ontogeny and tissue distribution and in function. Monoclonal
antibody depletion experiments revealed that Vpi+ T cells are essential for IgA
production and normal mucosal antibody responses (Sowder et al. 1996).
Functional differences between the Vpi and VP2 TCR repertoires are also
revealed by GVH reactions using various strains differing in MHC class II alleles
The avian immune system is characterized by a higher frequency of y8 T cells
than mouse and human, and may reach 30-50% of peripheral blood lymphocytes
(Sowder et al. 1988). Peripheral blood y8 T cells also respond to androgens,
resulting in an expanded population in males (Arstila et al. 1993).
T-Cell Development
It is currently held that practically all T cell development in the chicken is
thymus-dependent (Dunon et al. 1993a,b). Thus, unlike in the mouse, in which
extrathymic T cell populations have been described, in chicken all T cells,
including the intestinal intraepithelial T cells, originate in thymus from which
they emigrate to form the peripheral cell pool. Thymus is seeded in three waves
of blood-borne precursors (Jotereau et al. 1980). The first wave starts to arrive in

134
thymus on ED6, the second on ED 12 and the third on EDI 8. Each of these waves
gives rise to all T cell subsets. The first T cell receptor-expressing thymocytes are
y8 TCR+, first observed on ED 12, a|3 TCR+ thymocytes expressing Vj3l family
TCR genes develop next, on ED 15 and VJ32 TCR+ thymocytes on ED 18 (Buey et
al. 1990). The mature T cells probably migrate to periphery in distinct waves, as
well. The first peripheral TCR+ cells can be observed in spleen, on EDI5. These
cells express the y8 TCR; a(3 T cells appear 3 days later.
The genes encoding the TCR chains in the chicken have been described, and
their main characteristics resemble the mammalian genes (Tjoelker et al. 1990;
Kubota et al. 1994). Thus, they consist of multiple gene segments, which can, on
the basis of sequence comparison, be divided into families. As in mammals, and
unlike in the avian immunoglobulin genes, diversity of the TCR genes results
mainly from rearrangement and junctional variation. The mechanism determining
the segregation of thymocytes to either y8 or aP T cell lineage is unclear. It has
been suggested the aP thymocytes may represent those cells, which fail to express
y5 TCR. In these cells the TCR 8 locus, situated within the a locus, is deleted
during rearrangement. Support for this scenario is provided by studies of chicken
aP T cell lines, in which 8 locus was shown to be deleted from both alleles
(Kubota et al. 1994). In contrast, the TCR yg genes have been reported to be
frequently rearranged and transcribed in aP T cell lines (Sowder et al. 1988),
which argues against the model of transcriptional silencing of y genes being the
determinative factor.

135
Studies in mouse have shown that during the maturational process within the
thymus T cell are subjected to negative and positive selection (Nossal, 1984).
These events result in the death of most of the thymocytes and modify the
repertoire of the surviving cells to imprint MHC restriction and self-tolerance.
Data obtained in the chicken suggest that aP T cells develop through a similar
pathway (George and Cooper, 1990; Banbura et al. 1991). They pass from
CD4/CD8 double-negative stage to a double-positive stage in the thymic cortex,
and thereafter one of the co-receptors is down modulated. The aP TCR is first
expressed at low levels and at a certain stage the cells are susceptible to TCR
down-modulation triggered by receptor cross-linking with mAb. It is less clear
what kind of selection, if any, the y8 TCR+ thymocytes are subject to (George and
Cooper, 1990; Banbura et al. 1991). They express high levels of TCR
immediately and appear to pass quickly through the thymus. Cross-linking of y5
TCR has no effect on it expression, and cyclosporin A treatment, which arrests
the development of aP T cells, does not affect y5 T cell development. After
thymic maturation both aP and y8 T cells emigrate to all lymphoid tissues in the
periphery. An interesting exception is the homing pattern of aP T cells
expressing TCR VP2-gene family (Lahti et al. 1991). These cells are very rare or
absent in the intestinal epithelium and perhaps because of this cannot provide help
for antibody responses of the IgA class (Cihak et al. 1991). The reasons for this
difference between the two aP T cell lineages are not known.

136
Conclusion
Although a significant part of what is known about the ontogeny of the
immune system has been obtained from studies in the chicken, many aspects
remain unclear. Lack of suitable markers for the Hematopoietic stem cells is a
major obstacle to understanding the early events of lymphopoiesis. B cell
precursors entering the bursa have already rearranged their immunoglobulin
genes, but for T cell precursors, the degree of lineage commitment prior to
reaching thymus remains a matter of speculation. The developmental pathways
within the primary lymphoid organs and the formation of mature lymphocyte
repertoire remain areas for further study. The development of y5 T cells is of
considerable interest because the chicken is a species with a very high frequency
of these cells. The recent characterization of genes encoding the TCR chains
and the growing list of molecules defined by monoclonal antibodies with
undoubtedly bring new insights. In this way, the chicken continues to contribute
to the understanding of developmental immunology in general.
Oncogenic Avian Viruses
Retroviruses
Avian Leukosis Virus (ALV) and Rous Sarcoma Virus (RSV)
The ALV and RSV of chickens belong to the family of retroviruses and are
divided in five subgroups A, B, C, D, and E based on differences in the
glycoproteins present in the viral envelope. These glycoproteins consist of a
spike known as gp35 and a knob known as gp85. The gp85 is different for each

137
subgroup and is important for the interaction with the cellular receptors. The
absence of the receptor for a specific subgroup will result in cellular resistance to
the virus of that subgroup.
The viruses of subgroups A, B, C, and D all belong to the exogenous virus
groups, while subgroup E is an endogenous virus. The A and B subgroups of
ALV are most commonly encountered in commercial poultry populations
(Calnek, 1968) wile subgroups C and D have only been reported in a few
countries (Sandelin et al. 1974). Complete or defective subgroup E virus is
present in virtually all chickens but is often not completely expressed (Crittenden
and Astrin, 1981). The genetic information of endogenous virus is located on at
least 12 different loci (ev genes). The importance of the ev loci is not clear and
chickens have been produced free of ev genes (line 0) (Astrin et al. 1980). In the
last 5 years, the subgroup J virus has emerged as a major player.
The RNA genome of the ALV has been well characterized and consists of
three genes. These are from the 5’to 3’end: gag, pol, and env. At both ends a
short unique sequence with a small terminal repeat is found. After infection of
the cells, the genomic RNA is transcribed in double-stranded DNA by the reverse
transcriptase. The provirus is flanked at both ends by a large terminal repeat
(LTR), which is formed by the combination of the unique sequences at the 5’ and
3’ end of the genome. The mechanisms responsible for the synthesis, integration,
and transcription of the provirus have been reviewed recently (Hughes, 1983).

138
RSV and the other acutely transforming viruses on the other hand carry an one
gene, which is absent in the avian leukosis viruses. The viral one genes originate
from cellular genes (Muller and Verma, 1984).
The sre gene of RSV is the best-studied v-onc gene of avian leukosis viruses.
«
Most of the acutely transforming viruses do not have a complete genome; often
the gag and/or env gene is lacking or incomplete (Hayman, 1984). These viruses
need an ALV as helper virus in order to produce infectious virus. The mechanism
of transformation has been the focus of intensive study and has been recently
reviewed (Muller and Verma, 1984; Hayman and Buey, 1984). The
transformation by v-one is apparently caused by excessive production of the gene
product in the case of RSV a protein kinase. The provirus of the slowly
transforming ALV can activate c-one genes by the LTR, which acts as a promoter
sequence. Lymphoid leukosis virus (LLV) in tumor cells seems often to be
integrated next to or near the c-myc gene (Hayward et al. 1981).
The transmission of ALV is important for the understanding of the
pathogenesis and the immune responses. The exogenous virus can spread either
horizontally or congenitally. The former will result in a transient viremia and
antibody development; these chickens seldom develop lymphoid leukosis (LL).
Congenital infection will cause a chronic viremia and tolerance to the exogenous
virus, and LL is a common consequence (Witter et al. 1981). In addition to LL,
infection with ALV, RSV, or the defective, acutely transforming viruses can
result in the development of a wide range of tumors, which are reviewed in detail
by Purchase and Payne (1987). The pathogenesis of LL is not completely

139
elucidated, but transformed cells develop in the bursa of Fabricáis and consist of
immature B cells (Calnek, 1968). The transformation in the bursa is a slow
process and bursectomy up to several weeks of age can prevent the development
of tumors in other organs. Even inoculation with autologous transformed cells
from the bursa of Fabricius after bursectomy will not cause the development of
metastatic tumors when done before 13 weeks of age (Fadly et al. 1981).
Reticuloendotheliosis Virus (REV)
The REV belong also the retroviruses but can be distinguished from the
ALV/RSV group by orphological, immunological, and structural criteria (Witter
and Glass, 1984). It has been suggested the REV are closely related to
mammalian retroviruses (Barbacid et al. 1979). The genomic structure of
nondefective REV (ndREV) is similar to ALV and consists of the gag-pol-env
genes. The acutely transforming T strain lack parts of the gag-pol sequence and a
part of env gene, but it carries an onc-gene (v-rel). The cellular sequences related
to v-rel are present in normal DNA of avian species (Chen et al. 1981). The
pathogenesis of infection with ndREV has similarities with ALV; congenital
transmission will result in chronic viremia in the absence of antibody
development (Witter et al. 1981). Under these conditions or after infection of
immunologically incompetent animals by a number of disease signs can develop
such as a runting syndrome, lymphomas, and sarcomas (Witter et al. 1981; Li et
al. 1983).
Herpesvirus

140
Marek’s disease virus (MDV)
Vaccination, pathogenesis, and pathology will be discussed in the following
chapter for avian herpesviruses, including pathogenic serotype 1 and apathogenic
serotype 2 MDV and herpesvirus of turkeys (HVT).

CHAPTER 6
HISTORY OF MAREK’S DISEASE
Introduction
The Disease and Its Definition
Marek’s Disease (MD) was the term chosen in 1960 for a common
lymphoproliferative disease principally of the domestic chicken (Gallus
domesticus), which is now known to be caused by a cell-associated alpha
herpesvirus. The lymphoproliferative process in MD can involve most organs and
tissues including peripheral nerves. The most commonly affected organs and
tissues are peripheral nerves, iris, gonads, spleen, heart, lung, liver, and skin.
Cytolytic changes and atherosclerosis can be a manifestation of MD virus (MDV)
infection as can a clinical syndrome described as transient paralysis. MD can
occur at any age from a few weeks old, but is most common between 2 and 6
months of age. The incidence of disease in a flock before vaccination was
introduced would vary between a few percent of a flock to as high as 60% in
exceptional circumstances. However, 20-30% was typical (Weiss and Biggs,
1972).
Involvement of MD in organs and tissues in addition to peripheral nerves
occurs in more severe outbreaks of disease. In general, involvement of tissues
and organs, other than peripheral nerves, increases with reducing age at which
disease occurs and increasing incidence of disease. MD occurs in all poultry-
141

142
producing countries and, before vaccination was introduced for its control, was
responsible for serious economic loss to the poultry industries throughout the
world. Infection with MDV is ubiquitous in the domestic chicken and is also
present in jungle fowl (Gallus gallus) (Weiss and Biggs 1972).
First Description of MD
Dr. Joseph Marek, an eminent veterinary clinician and pathologist and
Professor and Head of the Department of Veterinary Medicine at the Royal
Hungarian Veterinary School in Budapest, published a paper entitled “Multiple
Nervenentzuendung (Polyneuritis) bei Huehnem” (Marek, 1907). In this paper,
Marek describes a disease in four adult cockerels, which were affected by
paralysis of the legs and wings. In detailed examination of the cockerel, Marek
noted thickening of the sacral plexuses and spinal routes that were infiltrated by
mononuclear cells. He further described the disease as a “neuritis interstitialis” or
a “polyneuritis”. A study of his description leaves little doubt that this was the
first published account of what was to be later called MD (Marek, 1907). These
early descriptions of MD suggested that the pathological changes only occurred in
the central and peripheral nervous systems. The disease was known to
poultrymen as “fowl paralysis” or “range paralysis”.
Recognition of the Lvmphoproliferative Nature of MD
The next landmark was the seminal studies by Pappenheimer et al. (1926).
These researchers made a detailed study of the epidemiology and distribution of
the disease and concluded that it was present in varying prevalence in most states
of the US. However, the most important observation was that of 60 cases studied

143
in detail; they found that in addition to lesions in nerves and the central nervous
system, 10% of the chickens had lymphoid tumors. Although these principally
involved the ovary, tumors were also seen in the liver, kidneys, lungs, adrenals,
and muscle. From clinical and research findings, they considered these tumors to
be a manifestation of the disease. Thus, they considered the terms previously
used to describe the disease as inappropriate. The outstanding pathological
features of the disease were lymphoid cell infiltration of peripheral nerves and
lymphomatous growths in other tissues and organs frequently accompanied by
inflammatory lesions. This suggested the term neurolymphomatosis gallinarum
for the disease and used “visceral lymphomatosis” and “visceral lymphomata” to
describe the visceral tumors.
The significance of these findings was the recognition that the disease was not
one of the nervous system alone but was a lymphoproliferative process. Infection
resulted in lesions in peripheral nerves, which were essentially similar in nature to
the visceral tumors that occurred in a proportion of cases.
Changes in MD through the Years
The disease described by Marek (1907) only involved the nervous system,
which he first saw in the US in 1914. By 1922, the disease was present in most
parts of the US and it appeared to have become more severe. Outbreaks involving
up to 20% of a flock with a significant proportion of affected birds having
lymphoid tumors in organs and tissues were common (Pappenheimer et al. 1926).
A high incidence of “visceral lymphomatosis”, in addition to neural
involvement, was reported in the 1950s in the USA in young chickens between 8

144
and 10 weeks of age (Benton and Cover, 1957). This condition became widely
known a “acute leukosis”. A similar condition was reported in Great Britain
(Biggs et al. 1965). In one study of the disease in the field and transmission
experiments, it was concluded that it was a form of MD and that it would more
appropriately be called acute MD than “acute leukosis” (Biggs et al. 1965;
Purchase and Biggs, 1967). The features of acute MD were the explosive nature
of outbreaks, the young age at which the disease could first appear with high
mortality of 30% or more and the high incidence of visceral lymphoid and neural
tumors. This form of MD was clearly more severe than the disease present in the
1920s, 1930s, and 1940s, which Biggs (1967) described as classical MD to
differentiate it from the acute form. Acute MD became the predominant form by
the 1960s in most countries that had a well-developed poultry industry.
After the introduction and widespread use of vaccines in the early 1970s, MD
was at first well controlled, although vaccines do not protect against
superinfection with wild type virus (Churchill et al. 1969; Witter et al. 1976).
However, strategies and type of vaccine had to be modified in response to
increasing reports of disease appearing in vaccinated chickens during the late
1970s. A similar situation again arose in the 1990s. In both cases, it was shown
that changes in the virulence of the circulating field virus had occurred (Nazerian
et al. 1996). Although the disease was appearing commonly in vaccinated flocks,
some outbreaks had unusual clinical and pathological aspects, such as a high
incidence of ocular lesions (Ficken et al. 1991), early mortality without visceral

145
lymphomata resulting from severe cytolytic changes in lymphoid organs, or an
encephalitis which sometimes lead to transient paralysis (Barrow and Venugopal,
1999; Nazerian et al. 1996).
Experimental Transmission of MD
Earlier publications show that there was widespread belief that MD was
infectious. Attempts to transmit the disease stated with the early studies of and
continued over the years until unequivocal transmission was achieved in the
1960s (Sevoian et al. 1963; Sevoian and Chamberlain, 1963; Biggs and Payne,
1967).
These early studies were often inconclusive and were not accepted as
providing irrefutable evidence of the infectious nature of the disease. Close
examination of these reports showed, for example, the criteria used for diagnosis
in the transmission experiments were not satisfactory, and lymphoid leukosis and
even other manifestations of avian leukosis were confused with MD (Johnson,
1934). In these studies, there was always an incidence of the disease in control
groups since there were not adequate isolation facilities, for an infection we now
know to be highly contagious, to prevent spread from the experimental to the
control chickens. In many reports there was also inadequate detail presented or
the number of chickens used was too small to provide convincing evidence of
transmission. It is probable that MD was transmitted by some of these authors
because an examination, in the light of current knowledge, of the more notable
reports of the period (Pappenheimer et al. 1926) suggests that the disease was
transmissible. Belief in this view was not possible at the time because of either a

146
relatively low incidence of MD in experimentally treated chickens or high
incidence in the controls. For example, Pappenheimer et al (1926), who made the
first extensive and well-documented report on transmission of MD, produced 25%
incidence of MD in inoculated chickens as compared with only 6% in the
controls.
Convincing evidence for the transmissibility of MD came from the studies of
Sevoian et al. (1963) and Biggs and Payne (1967). Three factors led to this
success. The availability of secured isolation facilities and highly susceptible
lines of chicken and the use of living cells as the inoculum still needs to be
investigated. Biggs and Payne used a crude homogenate of an ovarian lymphoid
tumor to initiate their studies, but whole blood for subsequent passage.
Isolation of A Cell-Associated Herpesvirus
Because the behavior of MD in the field suggested that it was an infectious
disease and that its cause was likely to be a virus, attempts were made to isolate a
virus since the early days of research on the disease, but without success. It was
not until cell culture techniques became available, the right choice of cell type
chosen for primary isolation and the recognition of the cell-associated nature of
the causative agent that its isolation and identification were successful. An avidly
cell-associated herpesvirus was isolated from blood and from tumor cells of
chickens with MD in chick kidney cells (CKC) (Churchill and Biggs, 1967) and
in duck embryo fibroblasts (DEF) (Nazerian et al. 1968, Solomon et al. 1968). In
each cell type the inocula produced a cytopathic effect which had characteristics
of changes produced by herpesviruses. Inclusion bodies were present in the areas

147
of cytopathic effect and herpes-type virus particles were seen in infected cell
cultures. The cytopathic effect was in the form of plaques and these could be
used to quantify the dose in the inoculum. Following these reports, publications
from several areas of the US reported the isolation of a herpesvirus from field
cases of MD and from recognized isolates of the causative agent of the disease
(Ahmed and Schidlovsky, 1968; Bankowski et al. 1969; Eidson et al. 1969).
Evidence that a Herpesvirus Is the Etiological Agent of MD
Whether the herpesvirus was the etiological agent of MD could not initially be
answered. Koch’s postulates are often difficult to fulfill for viruses due to the
difficulty of guaranteeing the purity of cell cultures. In the case of a cell-
associated virus it is impossible. In addition, because herpesviruses are common
and widespread in animal populations, it was suggested that the herpesvirus
isolated from cases of MD could be a common virus infection of the domestic
chicken that was co-isolated as a passenger with the causative agent.
Explanation for the Contagious Nature of MD
The question of how a disease caused by a cell-associated agent could be
highly contagious was answered by the demonstration by Calnek and his co¬
workers that antigens of the herpesvirus were consistently present in the
superficial layers of the feather follicle epithelium. They also demonstrated
enveloped herpesvirus particles were present at this site (Calnek and Hitchner,
1969) and that they also showed that cell-free extracts of skin were infectious
both for cultured cells and the chicken (Calnek et al. 1970a,b). These
observations not only provided further evidence that the herpesvirus is the

148
causative agent of MD, but also provided an explanation of how the virus could
spread. Poultry dust and litter, which contain desquaminated epithelial cells, have
been shown to be infectious (Beasley et al. 1970, Jarajda and Klimes 1970,
Carrozza et al. 1973) and remain infectious for over a year (Witter et al. 1968;
Carrozza et al. 1973; Hlozanek et al. 1973). These observations provided a
satisfactory explanation as to why and how the etiological agent of MD is highly
contagious.
Conclusive Evidence for the Herpesvirus Etiology of MD
Conclusive evidence for the herpesvirus etiology of MD was the
demonstration that an attenuated pathogenic herpesvirus and a herpesvirus
isolated from turkeys closely related antigenically to the chicken herpesvirus were
both capable of protecting chickens from MD (Churchill et al. 1969b, Okazaki et
al. 1970).
Morphology
Subdivision of the herpesvirus family into subfamilies is based on
biological properties that in general coincide with genome structure; these
subfamilies are alpha-, beta-, and gamma-herpesviruses. MDV is a strongly cell-
associated herpesvirus with lymphotropic properties similar to those of gamma
herpesviruses. However, its molecular structure and genomic organization are
similar to those of alpha herpesviruses. Therefore, it is currently classified as an
alpha herpesvirus. There have been many ultrastructural studies describing the
morphology and morphogenesis of MDV and HVT, and these have been reviewed
by Payne et al. (1976), Kato and Hirai (1985), and Schat et al. 2001). These

149
studies have shown essentially similar results and, in general, viruses of all three
serotypes have characteristics typical of other herpesviruses. Most studies have
used infected cell cultures because the presence of particles is infrequent in the
chicken in tissues other than the feather follicle epithelium (Calnek et al. 1970a,b;
Cauchy et al. 1979). Cell culture studies have almost exclusively been with
serotype-1 MDV or HVT infections.
Biology of the MDV Group
Introduction
Antigenically related viruses varying in pathogenicity from highly oncogenic
to non-oncogenic have been isolated from domestic chickens, and together with
an antigenically related virus, isolated from turkeys (HVT), they form the MDV
group.
Historical Aspects of MDV Isolates
A characteristic of the early isolates was that they varied in their pathogenicity
and oncogenicity. For example, HPRS-B14 produced mainly neural lesions with
only 18% of 451 cases with visceral tumors in a susceptible line of chicken and
only 3.3% out of 61 cases in a resistant line of chicken, and none out of 118 cases
in another resistant line (Biggs et al. 1967). HPRS016 produced a higher
incidence of disease and proportion of visceral tumors in the same strains of
chicken: 85% of 42 and 64% of 44 cases of MD in susceptible and resistant lines
of chicken, respectively (Purchase and Biggs, 1967). The JM and GA strains
were similar to the HPRS-16 strain (Sevoian et al. 1963; Eidson et al. 1969), and
the Conn-A and HPRS-17 strains to the HPRS-B14 strain (Chomiak et al. 1967;

150
Purchase and Biggs, 1967). Classical and acute MD were terms introduced to
describe these two types of disease (Biggs et al. 1965). The terms were also used
to describe strains of the causative agent that produced these two types of MD;
however, it was recognized that there may be strains in the field with all grades of
virulence between “severe acute and mild chronic forms” (Purchase and Biggs,
1967). Interestingly, the disease produced by most isolates fit neatly until the late
1970s into one or other of these types. In the late 1970s, an acute type of disease
appeared in HVT-vaccinated flocks. The characteristic of viruses isolated from
such flocks (Md/5, Md/11, ALA-8, RB1B) was that they produced more severe
disease and a significant incidence of disease in HVT-vaccinated experimental
chickens (Stephens et al. 1980; Eidson et al. 1981; Schat et al. 1982). In order to
control disease in the field, a change in vaccine strategy was made and bivalent
and trivalent vaccines were introduced. During the 1990s, MD was seen at an
increased incidence in flocks vaccinated under the new regimes. This suggested
that a further change in virulence had occurred. A series of isolates was made and
a proportion was found to produce, under standard test conditions, a higher
incidence of MD in bivalently vaccinated (HVT+SB-1) chickens than those
isolated during the 1980s (Nazerian et al. 1996). These isolates are also
characterized by their ability to cause early mortality. Isolates with a similar
property or other novel properties have been made during the 1990s from the
USA, Europe, and Japan (Nazerian et al. 1977).
In addition to the isolation of pathogenic strains of varying virulence, isolates
were made that were found to be non-oncogenic and in some cases apathogenic.

151
Examples of early isolates are HPRS-24 and -27 (Biggs and Milne, 1971), HN
(Cho and Kenzy, 1972) and SB-1 (Schat and Calnek, 1978). In a study in which
isolates from several flocks with varying incidence of MD were categorized, it
was found that “apathogenic” MDV was widespread in the poultry population
(Biggs and Milne, 1971; Jackson et al. 1976). These viruses, which may have a
very low level of pathogenicity but no oncogenicity, are most frequently referred
to as non-oncogenic MDVs.
Witter et al. (1970) isolated from turkeys an apparently identical virus, which
they showed to be antigenically related to MDV. It was classified as a
herpesvirus of turkeys (HVT) and the strain was designated FC-126. It was cell-
associated and non-pathogenic for turkeys and chickens. This was to become the
first widely used commercial vaccine in chickens to control losses associated with
MD.
Classification of the MD Group
Initially, the MD viruses were classified as either classical or acute MDV
(Biggs et al. 1965) based on the type of disease they produced. HVT was added
to the group because it was shown to be antigenically related to MDV (Witter et
al. 1970), and this was followed by apathogenic MDV which was also shown to
be antigenically related to pathogenic MDV (Biggs and Milne, 1971). As time
progressed, the classification of pathogenic MDVs into classical and acute
became inadequate due to the isolation of viruses of greater virulence.

152
Serotype
It has been shown that HVT is antigenically related to MDV. Antibody to
HVT could be distinguished from antibody to MDV by the distribution of
antigens in HVT-infected cell cultures detected by immunofluorescence (Witter et
al. 1970; Purchase et al. 1971). Von Bulow et al. (1975) found no difference in
the distribution of MDV-induced and HVT-induced antigens regardless of
whether homologous or heterologous antisera were used. This could have been
due to the use of different cell types. However, they did find a difference between
HVT and MDV in quantitative immunofluorescence studies.
In a series of studies using immunofluorescence, immunodiffusion and
neutralization, it was found that there were antigenic differences between
pathogenic and apathogenic MDV and between both of these and HVT (von
Bulow et al. 1975). Based on these results, von Bulow et al. (1975) suggested
that the virus strains they examined HPRS-1, JM, and GA acute MDV and their
attenuated derivatives, HPRS-B14 and VC classical MDV, HPRS-24 and -27
apathogenic MDV and FC-126 HVT) could be divided into three serotypes: (1)
pathogenic strains of MDV and their attenuated derivatives (MDV-1), (2)
apathogenic strains of MDV (MDV-2), and (3) apathogenic HVT.
The division of the MDV group into three serotypes has been supported by
subsequent studies (von Bulow et al. 1975; Schat and Calnek 1978; King et al.
1981; Ikuta et al. 1982, Lee et al. 1983) and by the demonstration that genomes of
viruses of the three serotypes differ in their restriction endonuclease digestion
patterns (Hirai et al. 1979, Ross et al. 1983,Camp et al. 1991). Monoclonal

153
antibodies aided in the serotyping of viruses (Ikuta et al. 1982; Lee et al. 1983).
The classification of MDV group viruses into three serotypes is now widely
accepted.
MDV-1 Pathotvpes
The pathotype is of considerable importance in determining the outcome of
MDV infection. Marked differences exist among serotype-1 virus strains in their
pathogenic potential, with an apparent evolutionary shift toward higher virulence,
broader host spectrum, and more varied lesion response in recent years, perhaps
related to selective pressure associated with the use of MD vaccines (Witter,
1997). In comparisons involving viruses that would now be classified (Witter,
1997) as mild (mMDV), or virulent (vMDV) strains, few differences were found
between pathotype and pattern of infection during the early cytolytic period,
which is 3-6 days post-infection (dpi) (Calnek et al. 1978; Fabricant et al. 1977;
Smith and Calnek, 1974). Although higher virulence strains differed from lower
virulence strains by showing a significant increase in virus titers in tissues and
blood after the early infection period. In more recent studies involving vMDV,
very virulent (wMDV) and very virulent plus (w+MDV) strains of viruses were
found. Calnek et al. (1998) determined that the two higher-virulence pathotypes
(wMDV and w+MDV) resulted in higher virus-isolation rates from splenocytes
at 4-8 dpi than were found with the lower-virulence pathotype (vMDV).
Furthermore, although all three pathotypes induced similar levels of early (4-5
dpi) cytolytic infection, there were marked differences at 7-8 dpi when the
w+MDV strains had significantly higher levels of cytolytic infection in the bursa

154
of Fabricáis and thymus than were seen in chickens infected with vMDV or
wMDV strains. This indicated a persistence of cytolytic infection beyond the
time when a switch to latency would normally occur, a feature attributed to loss of
immune competence. Also, a correlation was found to exist between virulence
and atrophy of the bursa and thymus. Higher-virulence pathotypes caused more
severe immunosuppression as measured by organ weights and histopathological
evidence of necrosis and atrophy. Additionally, lymphoid organs of chickens
infected with w+MDVs showed little recovery between 8 and 14 dpi, whereas
those of birds infected with vMDVs evidenced a return to normal by 14 dpi.
Chickens given wMDVs were intermediate in their recovery rate. All of these
observations suggested a correlation between immunosuppressive potential and
virulence, at least in the case of some of the more highly pathogenic strains of
virus. This agrees with reports by others in which newer isolates of MDV were
associated with unusually high virulence based on severe early cytolytic infection
and exceptionally marked necrobiotic changes in lymphoid organs, high early
mortality, severe and persistent atrophy of the bursa of Fabricáis and thymus, and
high mortality at 9-11 dpi (Witter, 1997). The reasons for the exceptional
virulence of some MDV isolates remains to be determined, although Barrow and
Venugopal (1999) found some changes in amino acids within the meq and ICP4
proteins that were conserved in three very virulent European MDV isolates,
suggesting that they may be involved in their high pathogenicity.
The possible association between high virulence and high immunosuppressive
potential raises interesting questions (Calnek et al. 1998) including: (1) do

155
w+MDVs have a special propensity for lymphoid organs above and beyond that
of wMDVs or vMDVs (2) Do they infect a higher proportion of B and/or T-cells;
and (3) do they more frequently induce a cytolytic versus latent infection in target
lymphocytes. This effect is due to inherent differences in viral replication events
or because of differences in responsiveness to factors that govern virus
replication, such as the cytokines shown to affect the establishment or
maintenance of latency? Immunofluorescence data failed to support the idea that
the w+MDVs infect more lymphocytes during the early period (Calnek et al.
1998). Rather, it seems likely that the failure to enter latency with the
accompanying continued destruction of lymphocytes accounts for the more severe
atrophy of bursa and thymus in birds infected with the more virulent MDVs. This
enhanced immunosuppressive activity could be because the w+MDVs infect
specific subsets of cells involved in effecting the switch to latency, thus reducing
the required immune factors. Or, cells infected with the more virulent viruses
may fail to display the surface marker required for interaction with cytokines
known to help modulate virus replication and thus latency. In any case, the severe
destruction of the bursa and thymus results in overwhelming immunosuppression,
a significant factor in the incidence of MD. The importance of immune
competence was apparent in early studies in which neonatal thymectomy
enhanced the tumorogenicity of low-virulence strains of MDV (Calnek et al.
1977) and increased the incidence of lymphomas in genetically resistant fowl
(Payne et al. 1992).

156
Recombination and Mutation in MDV
There has been ample opportunity for recombination amongst the three
serotypes of MDV group of viruses. Superinfection with field virus of chickens
vaccinated with an attenuated serotype-and with serotype-3 HVT virus has been
reported (Churchill et al. 1969b; Purchase et al. 1971), and clearly is a widespread
phenomenon. Viruses of serotype-1 and -2 have been isolated from single pens
of chickens, from the same chicken in sequential studies (Biggs, 1972; Jackson et
al. 1976) and from the same tissues in dually infected chickens and from a cell
line in culture (Cho and Kenzy, 1972). Despite these observations, there has been
no report of spontaneous recombination between the three serotypes.
However, spontaneous mutation apparently does occur. All three serotypes
alter in their biological and molecular properties on passage in cell culture
(Churchill et al. 1969a; Witter and Offenbecker, 1979; Hirai et al. 1981). A
temperature-sensitive mutant of MDV and a phosphonoacetate mutant of HVT
have also been described (Lee et al. 1978; Witter and Offenbecker, 1979).
Changes in the virulence of MD over the years also provide support for the view
that mutation under suitable conditions is part of the biology of the MDV group.
Virus-Cell Interaction
Introduction
There are four virus-cell interactions recognized in oncogenic MDV infection:
productive, restrictively productive or abortive, and two forms of non-productive,
latent and transforming. Four phases of MDV infection in vivo can be delineated:
1) early productive-restrictive virus infection causing primarily degenerative

157
changes, 2) latent infection, 3) a second phase of cytolytic, productive-restrictive
infection associated with permanent immunosuppression, and 4) a proliferative
phase involving nonproductively infected lymphoid cells that may or may not
progress to lymphoma formation. Lymphoproliferative changes constituting the
ultimate response in the disease may progress to tumor development, although
regression of lesions commonly does occur either before or after T-cell
lymphomas are apparent (Biggs, 1966).
The composition of MDV-induced lymphomas is complex, consisting of a
mixture of neoplastic, inflammatory, and immunologically active cells. Both T-
and B-cells are present, although the former predominate. The usual target cells
for transformation in vivo are activated CD4+/CD8- T-cells. If experimental
conditions of infection are modified, it is possible to show that a variety of T-cell
subsets, including CD4+/CD8-, CD4-/CD8+, and CD4-/CD8- cells, are
transformable in vitro. It is possible that MD tumors are of clonal origin, but
studies have not yet confirmed this notion. However, there is evidence to suggest
that immune responses of the host may be directed against the early virologic
events or the later proliferative phase, and that an effective response at either
stage might reduce the chance of disease. In the case of MDV infection, no
published information on apparent cytokine regulation was found (Biggs, 1966).
Productive Infection
In productive infection, virus replication is complete and enveloped virus is
produced and released, resulting in death of the cell. In the chicken, this only
occurs in the feather follicle epithelium where large numbers of fully infectious

158
virus particles are produced (Calnek et al. 1970a,b). Full replication of the virus
occurs variably in cell culture. In some virus-cell culture systems, virus is
released into the supernatant and fully infectious virus can also be harvested by
disruption of cultured cells (Calnek et al. 1970b; von Bulow et al. 1975; Cho,
1977).
Restrictivelv Productive Infection (Abortive)
Restrictively productive or abortive infections are those where there is no fully
infectious virus released from the cell or tissue, but expression of the viral
genome may range from the production of virus-specific antigen to, in rare cases,
enveloped intracellular virions. In the latter case, it is a cytolytic infection. The
presence of nucleocapsids, mainly in the cell nucleus, and fewer intracellular
enveloped virions is seen in cytolytic cell culture infections. In infected chickens,
enveloped virions are not seen except in the feather follicle epithelium or
occasionally in other cells and tissue when young antibody-free chicks are
infected (Frazier and Biggs, 1972). Cytolytic infection also occurs in a number of
tissues and organs in chickens infected with w- and w+MDV. Otherwise,
nucleocapsids have only been seen in lymphoid cells (Calnek et al. 1970a,b).
Most of the viral genome is transcribed in infected chick embryo fibroblasts in
culture (Nazerian, 1971).
Latent Infection
Latent infection can only be detected by hybridization with viral DNA probes
or methods that activate the viral genome. For example, in vitro cultivation of
latently infected cells results in the expression of gene products and virus particles

159
(Calnek et al. 1981). Latent infection has been reported in T lymphocytes and is
present in chickens infected with either serotype-1, -2, or -3 viruses. These cells
contain less than five copies of viral DNA and do not express viral antigen
(Calnek et al. 1984).
Transforming Infection
Transforming infection differs from latent infection by the transcription and
expression of a number of genes resulting in the presence of virus-associated non-
structural proteins and transcripts in lymphoma cells and lymphoblastoid cell lines
developed from MD lymphomas. A number of viral transcripts mapping in the
inverted repeats flanking the long and short unique regions and to a lesser extent,
the adjacent unique sequences have been described (Sugaya et al. 1990). Genes
that have been described as expressed in lymphoma cells and lymphoblastoid cell
lines are present in the same region of the genome (Biggs, 1997). For example, a
gene found in the EcoRI Q fragment called meq, an acronym for Marek’s Eco Q,
is highly expressed in lymphoblastoid tumors (Kost et al. 1993), and an MDV-
specific phosphorylated proteins, pp38, has been found in lymphoma cells and
cells of a lymphoblastoid cell line (Nakajima et al. 1987). Transcripts which map
antisense to the homologue of herpes simplex virus infected cell protein 4 (ICP4)
gene have also been found in lymphomas and lymphoblastoid cell lines (Cantello
et al. 1994; Li et al. 1994). These also map to the same region of the genome.
Cells of lymphoma and of lymphoblastoid cell lines contain at least 10-20 copies
of viral DNA per cell, which is highly methylated (Ross et al. 1982; Kanamori et
al. 1987).

160
Conclusion
MD is the response of the chicken to infection with an alpha herpesvirus. It
forms a continual threat to the poultry industry requiring effective vaccines and
strategies to keep it under control. The virus appears to be continuously evolving,
thus requiring new vaccine strategies and possibly in the future more effective
vaccines. It is a disease of particular interest because infection with MDV can
result in a cytolytic and/or proliferative response of lymphoid cells resulting in
lymphoma, both of which can be fatal. The availability of well-studied and
documented experimental animal systems, together with oncogenic and non-
oncogenic viruses the MDV group, provide unparalleled opportunities for
understanding the oncogenesis of this herpesvirus infection and for the increased
understanding of the neoplastic process.
Pathogenesis of MDV Infection
Introduction
The term pathogenesis is simply defined as the development of disease. Many
uncomplicated infectious diseases proceed in a direct fashion from infection to a
specific pathologic expression. However, other diseases are relatively complex in
terms of pathogenesis: MD most certainly belongs in the category.
Pattern of Infection in Susceptible Chickens Infected with Oncogenic (Serotvpe-
1) Virus
Infection with MDV leading to lymphoma formation can be generally divided
into four phases: (1) early cytolytic, (2) latent, (3) late cytolytic, and (4)
transforming. Although essentially sequential, these are not necessarily discrete

161
phases. A sharp line demarcates the first two stages, and latent infection in
certain cell types is a prerequisite to transformation, but both transforming and
latent infections may exist intermixed with cytolytic infections in different cell
populations as lymphomas developing in the later stages. A permanent
immunosuppression develops concurrent with the phase-3 cytolytic infection.
The earliest event, i.e., the initial infection of the chicken, occurs via the
respiratory tract following inhalation of infectious cell-free MDV from a
contaminated environment. The role of the lung as a site of primary infection is
not entirely clear, since evidence of infection of parenchymal cells as an initial
event has been meager at best (Adldinger and Calnek, 1973; Phillips and Biggs,
1972; Purchase, 1970). On the other hand, the lung is the site from which
phagocytic cells are presumed to pick up the virus and carry it to lymphoid organs
such as the bursa of Fabricius (bursa), the thymus, and the spleen. At those sites,
splenic elipsoid-associated reticulum cells in blood vessel walls appear to be
involved in the access to lymphocytes for MDV (Jeurissen et al. 1992). A
productive, cytolytic, infection in those organs ensues and is particularly evident
between 3 and 6 dpi. This necrotizing infection provokes an acute inflammation
in which there is an influx of many cell types including macrophages, thymus-
derived (T) and bursa-derived (B) lymphocytes, and various granulocytes. Both
uncommitted and immunologically committed lymphocytes are present (Hudson
and Payne, 1973).
At 7-8 dpi, or sometimes slightly later, the infection in lymphoid organs
switches from productive to latent and a “viremia”, i.e., latent infection of

162
peripheral blood lymphocytes (PBL), can be detected. There is a transient
immunosuppression at about 7 dpi, apparently attributable to macrophage
functions (Lee et al. 1978), and at the same time, a transient hyperplasia may be
seen in the spleen. It seems likely that latency results from extrinsic factors
associated with the immune responses that become evident at about 6-7 dpi
(Higgins and Calnek, 1976), although the lymphocyte subset might able be
influential, given the fact that most cytolytically infected cells are B-cells whereas
most latently infected cells are T-cells. The immune-response association with
latency induction is compelling. Various cytokines, including interferon, are
influential in the matter of latency (Buscaglia and Calnek, 1988; Volpini et al.
1995, 1996), and immunocompetence is a requirement for latency to develop and
to be maintained (Volpini et al. 1995). Latency develops in embryonicly
bursectomized chickens (Schat et al. 1981), so it can be presumed that cell-
mediated immune functions are those of importance. Omar and Schat (1996)
reported that cytotoxic T lymphocytes (CTLs) directed against early viral antigens
are detectable by 6 dpi. It should be noted that the recrudescence of MDV
cytolytic infection correlates with a concurrent permanent immunosuppression.
In addition to specific immune responses, nonspecific immunity factors may also
be involved in the switch to latency. Nitric oxide, produced by macrophages, and
recombinant gamma interferon were able to inhibit the replication of MDV in
vitro and in vivo (Xing and Schat, 2000).
Infected PBL probably are the disseminators of virus to a large variety of
tissues of epithelial origin where, by the end of the second week, a second phase

163
of productive cytolytic infection is evident based on immunofluorescence tests
and histopathology. No extracellular virus can be detected in the blood and so it
must be concluded that virus spread is cell-to-cell. The question of virus
receptors has not been resolved. Cell-free virus binds to, but does not easily
penetrate, target cells in vitro (Adldinger and Calnek, 1971); on the other hand,
intracellular bridges appear to be an effective means for cell-to-cell spread of
MDV (Phillips and Biggs, 1972).
Organs found to have infected non-lymphoid cells include the kidney, adrenal
gland, proventriculus, uropygial gland, lung, liver, peripheral nerves, gonad,
esophagus, crop, Harder’s gland, and feather follicle (Adldinger and Calnek,
1973; Calnek and Hitchner, 1969). Probably there are few tissues truly spared.
Organs that develop cytolytic infections in epithelial-derived tissues generally
have infiltrating cells, including inflammatory cells and immunologically
committed lymphocytes. Proliferating lymphoblasts are seen in some of the
infected tissues, but the pleomorphic nature of the infiltrations suggests that
neoplastically transformed lymphocytes are intermixed with non-transformed
cells. A few of the infiltrating cells, including some that are neoplastically
transformed, may show evidence of productive cytolytic infection; others are
most certainly carrying viral genome based on the ability to secure virus or
otherwise demonstrate the presence of MDV genome.
Of all of the cytolytically infected tissues, only the feather follicle epithelium
(FFE) produces virus that is infectious in the cell-free state (Calnek et al. 1970a).
Although other tissues may contain a few enveloped virus particles detected by

164
electron microscopy, no infectious cell-free virus can be obtained from disrupted
cells. Thus, the feather follicle is unique in its epizootiologic role as the origin of
virus that can spread from bird to bird. Virus is generally associated with
desquamated, keratinized FFE cells, which are shed along with molted feathers
and as dander, thus contaminating the environment to complete the infection cycle
(Beasley et al. 1970; Calnek et al. 1970a). Latent infections of PBL and splenic
lymphocytes, and productive infection of the FFE, are lifelong events regardless
of the consequences of infection, i.e., whether or not the bird develops MD
(Witter et al. 1971).
Beginning as early as 12-14 days, to as late as several weeks or even months
after infection, microscopic and gross lesions may be seen in one or more of a
large number of sites including lymphoid organs, visceral organs, muscle, skin,
the eye, peripheral nerves, and brain. Even earlier, at 8-10 dpi, there may be an
acute degenerative disease or a transient paralysis under the right conditions.
Lymphoma development after 2 weeks or more is generally coupled with a
permanent immunosuppression affecting both humoral and cell-mediated
immunity (CMI). The degree of immunosuppression can vary and may be so
severe as to permit lethal infections with other organisms in the absence of any
lymphomatous lesions normally associated with MD. Immunosuppression
generally accompanies the second wave of cytolytic infection but it is not known
which is cause and which is effect, assuming they are related. Although
lymphomatous lesions may regress (Sharma et al. 1973), they often progress to
massive size, thus compromising the function to the organ(s) in which they

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develop and resulting in paralysis (if nerves are involved), blindness (if eyes are
affected), cachexia (particularly with visceral involvement), and death. In some
cases, the development of disease can be very rapid and may cause death before
clinical signs are prominent. Lymphomas can be diffuse or discrete in character.
Target Cells
Target cells for infection with oncogenic strains of MDV are of considerable
interest because of their importance to the pathogenesis of the disease. After it
was determined that the neoplastically transformed cells in lymphomas are T-
cells, it became important to learn the steps in pathogenesis that determine how
and why they are singled out, and the mechanism(s) by which they are selectively
transformed.
Target for Cytolytic Infection
Shek et al. (1983) first determined that B-lymphocytes are the primary targets
for the early cytolytic infection in lymphoid organs. This finding was consistent
with a report by Schat et al. (1981) showing that embryonic bursectomy obviated
the early cytolytic infection phase following exposure to MDV and found that B-
lymphocytes were the major target for in vitro infection of splenocytes. Burgess
and Davison (1999) reported that most spleen cells expressing the viral gene
product pp38 during the 4-6 dpi period were in aggregates of B-cells surrounding
sheathed capillaries. A few infected T-cells could also be detected during the
early infection period (Calnek et al. 1984). Those expressing pp38 were of both
CD4+ and CD8+ phenotypes (Burgess and Davison, 1999). Undoubtedly, these
were activated T-cells since resting T-cells are refractory to infection with MDV

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(Calnek et al. 1984). The extent to which activated T-cells support productive
infection with MDV is not known, but they participate, to at least a minor extent,
in the early cytolytic infection phase and probably to a greater extent as they
emerge in larger numbers during the last part of this phase just before latency
occurs. Aside from lymphocytes, a number of epithelial cells can be targets for
cytolytic infection, as noted above.
Targets for Latent Infection
In contrast to the early cytolytic infection target cells, the predominant
lymphocyte targets for latent infection are activated T-cells, mainly of the CD4+
phenotype, with only a minor population of latently infected B-cells (Calnek et al.
1984; Morimura et al. 1998; Shek et al. 1983). Probably, latently infected
lymphocytes are those that become infected during the cytolytic phase when virus
can easily spread from cell to cell, and then are prevented from completing the
replicative cycle by intrinsic and/or extrinsic factors. The reason for the marked
predominance of T-cells rather than B-cells during the latent phase is not known,
but it could be that B-cells are quickly lost because of their short life span
compared to longer-lived T-cells, or that the latent state in B-cells in not as well
induced and/or controlled by external influences as it is in T-cells, or that T-cells
are intrinsically less likely to support a productive infection. There is some
reason to suspect that the latter is at least of some importance; during the second
wave of cytolytic infection that is responsible for the biphasic pattern of MDV
infection. Latently infected, transformable T-cells apparently coexist with
productively infected lymphocytes in lymphoid organs in chickens that are «

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severely immunosuppressed. Although epithelial cells also are particularly prone
to cytolytic infection, little work has been done to determine if they may also
become latently infected. Pepose et al. (1981) reported that latent infections can
be detected in satellite cells and non-myelinating Schwann cells of peripheral
nerves and spinal ganglia.
Target for Transformation
The heterogenous makeup of lymphomas made characterization of
transformation targets difficult until continuous cell lines of MD tumor cells
provided uniform populations for study. Akiyama et al. (1973) first reported the
development of MD tumor cell lines and described two cell lines, which they
characterized as T-cells. Other lines subsequently examined were invariably
identified as T-cells, including the original lines developed by Akiyama et al.
(1973), Nazerian and Sharma 1975, and Nazerian et al. (1977). A variety of other
antigens were subsequently found on cultured MD cells including so-called
Marek’s disease tumor-associated surface antigen (MATSA) (Matsuda et al.
1976; Witter et al. 1975), now known to be a marker associated with normal
activated T-cells (McColl et al. 1987), chicken fetal antigens (Murthy et al. 1979),
and heterophil and Forssman antigens (Ikuta et al. 1981a,b). The significance of
the latter antigens in unknown, but because MATSA seems to be a marker
associated with activated T-cells, it is reasonable to expect that transformed cells
would express it.
Through the use of monoclonal antibodies, Schat et al. (1982b, 1991) were
able to characterize a large number of MD cell lines and found that the vast

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majority were la-bearing (activated) CD4+ cells that expressed either T-cell
receptor (TCR) a(3-l (TCR2+) or a(3-2 (TCR3+). Only a single line, of 45 tested,
failed to express CD4 antigen and all were negative for CD8 expression. It could
be deduced that the usual transformation targets in MD are a very specific subset,
i.e., activated T helper cells. However, it was also learned that although the usual
transformation targets are CD4+ T-cells, this is apparently due more to the
sequence of pathogenetic events than to a uniqueness of the subset as MDV-
transformation target cells. Calnek et al. (1989b) tested the hypothesis that
activation of T-cells is of crucial importance in lymphomagenesis because it
provides an abundance of transformation targets. They inoculated alloantigens
along with MDV in the wing-web or in the pectoral muscle of young chickens
with the anticipation that the additional stimulation to the immune response would
increase the number of activated T-cells and thus enhance tumor development.
Indeed, it did, because tumors developed at the site of inoculation in many of the
test birds. Unexpectedly, MD lymphoclastoid tumor cell lines could be
established from lymphocytes harvested from the early (4-6 dpi) inflammatory
lesions and, surprisingly, these were found to represent a large number of
combinations when tested for the presence of CD4, CD8, and TCR markers. Of
56 lines examined, only 12 (21%) were CD4+, whereas 25 (45%) were CD8+ and
the remained 19 (34%) were double negative, i.e., neither CD4+ nor CD8+. Both
TCR2+ and TCR3+ markers were found, but with no obvious relationship to the
CD4+ or CD8+ status of the lines. Probably, the alloantigens accelerated the
immune response resulting in a more mixed population of T-cell subsets being

169
present at the site of the cytolytic-phase infection before latency developed. The
lack of uniqueness of CD4+ cells as transformation targets were suggested by the
Schat et al. (1991) study in which a single cell line was found to be double
negative (neither CD4+ nor CD8+), a point rendered deficient for CD4+ and
CD8+ lymphocytes.
Three prerequisites for transformation of a given population of cells have been
noted (Calnek et al. 1998): (1) they must be available at the time of, and at the
site of, active infection; (2) they must be able to restrict viral replication, either
intrinsically or because they are responsive to extrinsic inhibitors; and (3) they
must be responsive to viral gene capable of effecting transformation either
directly or indirectly. The foregoing observations suggest that these conditions
are met by a variety of activated T-cells (albeit most often a CD4+ subset) but not
B-cells in chickens.
Although T-cells are apparently the invariable transformation target in
chickens, both B- and T-cell lymphomas have been observed in turkeys infected
with. Interestingly, infection of turkeys with oncogenic MDV strains produced
little or no evidence of cytolytic infection in the various lymphoid organs during
the period when acute necrotizing infection of these organs occurs in chickens
(Elmubarak et al. 1982), so the events leading to lymphoma formation are clearly
different. The absence of productive infection in the bursa suggests that B-cells
are either not a favored early target, or they are intrinsically unable to support
virus replication. The ability of MDV to transform both B and T-cells, on the
other hand, indicates that neither of these cell types is refractory to infection.

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Pattern of Virus Infection with Non-oncogenic MDV and HVT
There are three types of non-oncogenic virus that can be compared with
oncogenic serotype-1 MDV: attenuated serotype-1 MDV, e.g., HPRS-16 att
(Churchill et al. 1969b), naturally non-oncogenic serotype-2 MDV, e g., strains
like SB-1 (Schat and Calnek, 1978), and serotype-2 HVT strains such as FC126
(Witter et al. 1970). These all have a general pattern of tissue and organ infection
somewhat similar to that of oncogenic serotype-1 MDV (Calnek et al. 1979;
Fabricant et al. 1982; Payne and Rennie, 1976), but there are significant
differences. The most prominent difference is a complete absence of productive,
cytolytic infection in lymphoid organs or other tissue in chickens infected with
HVT or attenuated MDV, even though virus can be isolated from latently infected
cells in lymphoid organs or blood. In contrast, serotype-2 viruses cause at least a
low level of cytolytic infection of lymphoid organs during the early infection
period, but as with oncogenic viruses, the infection then becomes latent (Calnek et
al. 1979; Lin et al. 1991). Lin et al. (1991) found that productive infection with
the ML-6 strain of serotype-2 virus was restricted to lymphoid organs with a
single exception of a minor involvement of the lung at 3 dpi and the FFE from the
14th day onward. Serotype-2 viruses spread horizontally (Lin et al. 1991; Schat
and Calnek, 1978), but neither attenuated MDV nor HVT do so, even though viral
antigen can be detected for a limited period in the feather follicles of chickens
infected with the latter (Fabricant et al. 1982).
Consequences of Infection with MDV

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Cellular Level
Infection at the cellular level can result in at least four different responses:
cytolysis, latency, proliferation, and transformation. In general, it should be noted
that the cell type, the immune-competency of the host coupled with its immune
status, the virus strain, and vaccinal immunity all could influence the outcome of
disease.
The importance of the cell type has already been discussed. For instance, B-
cells appear to be particularly prone to cytolytic infection whereas activated T-
cells are most often latently infected. With both B and T-cells, this apparent
difference is dictated partly, though perhaps not entirely, by the phase of infection
(prior to or after immune responses) and the immunocompetence status and
immune response(s) of the host. Proliferation, as a response to infection, has been
observed in vitro. Two examples are a study by Spencer (1970), who observed
focal proliferation in cultured duck embryo fibroblasts following infection with
MDV, and one by Schat (1991) in which in vitro MDV infection of activated T-
cells resulted in prolonged proliferation but not transformation. Finally, as
already noted, only T-cells are susceptible to transformation in chickens.
Host Level
The initial description, in 1907, of what we now call MD made no mention of
many of the responses we presently attribute to MDV infection; rather, it
consisted of a detailed description of pathology classified as a “polyneuritis”
(Marek, 1907). It was only later that it was learned that visceral lymphomas and
eye lesions could be associated with the disease (Pappenheimer et al. 1926), and

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that nerve lesions previously described as “polyneuritis” could be complex and
involve both neoplastic and inflammatory components (Pappenheimer et al.
1926). The disease is now known to be very complex and to have many
additional potential consequences, including apparently unrelated conditions that
result from inflammatory/degenerative lesions, e.g., an early-mortality syndrome,
a transient paralysis syndrome, and severe lymphoid organ atrophy with
immunosuppression and the attendant consequences (Witter, 1997). Furthermore,
MDV has been found to be a cause of atherosclerosis in chickens (Fabricant et al.
1978).
Clinical MD
Inflammatory and Necrotizing Disease
The inflammatory responses and necrotizing effects of cytolytic infection in
lymphoid or other tissues are, by themselves, important aspects of the disease in
addition to their obvious role in the cascade of events leading to lymphomas.
Even though we now tend to consider the lymphomagenic nature of MD as being
the primary concern, our present understanding of the disease recognizes that
inflammation by itself may have serious consequences, and that the degenerative
lesions that derive from productive infection are themselves sometimes the major
effect of infection by MDV. This is particularly true with certain strains of virus,
or under conditions that favor inflammatory or degenerative lesion development.
Transient Paralysis
Some strains of chickens infected with certain strains of serotype-1 MDV may
develop lesions of the central nervous system (CNS) that cause a syndrome called

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transient paralysis (TP) (Kenzy et al. 1973). This condition is characterized by
flaccid paralysis, often first involving the neck and then becoming more
generalized, as shown in Figure 6-1. It occurs suddenly in young chickens at
about 8-12 dpi and is generally followed by full recovery within 1-2 days.
Prineas and Wright (1972) described histopathological lesions including mild,
diffuse cellular infiltration, mild interfolial meningitis, swelling and proliferation
of the capillary endothelium and mild perivascular cuffing by mononuclear cells;
these lesions were particularly evident in the cerebellum. Based on studies by
Komegay et al. (1983a,b) and by Swayne et al. (1988, 1989), it is not evident that
the clinical syndrome is the result of vasogenic brain edema. When edema is
resolved, the clinical signs disappear. The edema is secondary to a vasculitis
responsible for altered permeability of blood vessels. The pathogenesis of the
vasculitis is not known, although it has been shown that there is B-cell
involvement, given the need for a humoral immune response (Parker and
Schierman, 1983), and that the genotype is important since the condition is
governed by genes within, or closely linked to, the MHC (Coleman and
Schierman, 1980). Resistance to TP is dominant and appears to be independent of
genetically controlled resistance to lymphomas or neural lesions characteristic of
MD. However, TP-susceptible birds that recover from the transient disease often
succumb a few weeks later from lymphomas and/or the more usual neural form of
MD (Kenzy et al. 1973). Recent reports by Witter et al. (1999) and Gimeno et al.

174
(1999) described a more acute form of TP in which chicks infected with a highly
virulent strain of MDV failed to recover but instead succumbed within 1-3 days
after the onset of clinical signs.
Ocular Disease
Ocular lesions consisting of pupil irregularities and loss of pigmentation of the
iris (“gray eye”) have long been associated with MD. Although proliferating
lymphoblasts may be found in ocular lesions (Smith et al. 1974), suggesting a
neoplastic element, often there is only evidence of inflammation in the form of
edema and cellular infiltration of the iris, cornea, retina, pectin, uveal tract, and
other structures (Smith et al. 1974), as shown in Figure 6-2. The result is
blindness, a serious problem in its own right. As in the case of the vasculitis
associated with transient paralysis, the pathogenesis of these inflammatory eye
lesions is not known.
Degenerative Lesions. Immunosuppression, and Earlv-Mortalitv Syndrome
Severe systemic disease can result from the necrotizing infection of the two
major immunity-related organs, the bursa and thymus, and other tissues including
bone marrow (Jakowski et al. 1970; Sharma et al. 1980). The absence of
protective maternal antibodies and infection with the more highly virulent strains
of MDV enhances this non-neoplastic response, which has been called “early-
mortality syndrome” (EMS) (Sharma et al. 1980). The consequences were
reported to be aplasia of bone marrow (causing aplastic anemia) and marked
atrophy of the bursa, as shown in Figures 6-3 and 6-4, and thymus with an
attendant depression of immunity (Jakowski et al. 1970), although it is important

175
to remember that some of the MDV strains studied may have contained chicken
anemia virus as a contaminant, which could cloud the interpretation of certain of
the described consequences. An extravascular form of hemolytic anemia caused
by high-virulence MDV isolates also has been described. With certain virus
isolates, high mortality may occur after infection at a young age (Goodwin et al.
1995; Ross et al. 1996; Nazerian et al. 1996). Based on persistence of early
cytolytic infection and the degree of atrophy of the bursa and thymus, Calnek et
al. (1998) determined that there is relationship between the immunosuppressive
potential and the pathotypes of MDV isolates, but that is could be due to: (1) a
particular propensity of high-virulence viruses to target lymphoid organs, above
and beyond that of lower-virulence pathotypes; (2) infectivity for an expanded
range of target lymphocyte subsets; (3) a greater likelihood that infection with the
more virulent viruses results in cytolytic rather than latent infection of some
lymphocyte subsets; or (4) differences in responsiveness to factors governing
virus replication. Regardless, it is clear that certain isolates of MDV, when
infecting genetically susceptible chickens in the absence of maternal antibody or
vaccinal immunity, cause a virtual collapse of the immune system, which in turn
is undoubtedly a major factor in EMS (Witter et al. 1999).
The immunosuppression associated with MD could be directly related to loss
of effector cell populations due to productive infection, but apoptosis and/or
down-regulation of CD8 (Morimura et al. 1995), or immunosuppression by
certain MDV-encoded proteins (Morimura et al. 1996) could also be involved.

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Nerve Lesions
The pathogenesis of neural lesions in MD is somewhat confusing since there is
no published evidence of early virus infection that could serve as an incident in
peripheral nerves. Viral antigen and/or virions have occasionally been observed
in lymphocytes and Schwann cells of affected peripheral nerves (Adldinger and
Calnek 1973), but this was after gross lesions had developed. Inflamed siatic
plexus and nerve from MDV-infected chicken is shown in Figure 6-5. Most of
the available information on the development of neural lesions in MD is derived
from histopathological and ultrastructural examinations of peripheral nerves.
Ubertini and Calnek (1970) described the pathologic changes in peripheral
nerves as being of two main types. One, called A-type, is essentially neoplastic in
character with proliferation of lymphoid cells, sometimes with demyelination and
Schwann cell proliferation, as shown in Figure 6-6. B-type lesions, on the other
hand, are inflammatory in nature being characterized by edema and infiltration
with small lymphocytes and plasma cells, as shown in Figure 6-7.
A chronological study of ultrastructural changes in the peripheral nerves of
MDV-infected chickens (Lawn and Payne, 1979) provides valuable insight into
the sequence of events. Cellular infiltration of nerves occurred as early as five
dpi, with progression to proliferative A-type lesions by 3 weeks. Demyelination
was evident at 4 weeks followed by the appearance of B-type lesion in some
birds. It is interesting that the earliest nerve lesions coincided with the period of
peak cytolytic infection of lymphoid organs, but the absence of virions or other
evidence of MDV infection of peripheral nerves at that time raises the question of

177
why the initial infiltrations occurred. No satisfactory explanation has been put
forth, although Lawn and Payne (1979) pointed out that it is unlikely that auto¬
sensitization (as the result of demyelination) is involved because the damage
follows rather than precedes the early infiltration. Because demyelination that
occurs later in the evolution of nerve lesions was associated with the infiltration
of the basement membrane of Schwann cells by lymphocytes and macrophages, it
was concluded that the demyelination is likely the result of an allergic
sensitization to normal nerve antigens (Lawn and Payne, 1979). Payne (1979)
suggested that at some point, probably during a later stage of nerve infiltration, an
autosensitization to myelin occurs, provoking a progressive cell-mediated primary
demyelination. This conclusion is consistent with observation of others who
noted a similarity between MD nerve lesions and experimental allergic neuritis
and the Landry-Guillain-Barre’ syndrome (Fugimoto and Okada, 1977; Ichijo et
al. 1981; Lampert et al. 1977). The possibility of an autoimmune component for
MD is further supported by studies in which immune complexes were found in
the kidneys of MDV-infected chickens and quail (Kaul and Pradhan, 1991). It is
believed that the inflammatory B-type lesions develop following some repair and
remyelination (Payne, 1979). This presumes that either there is regression of
neoplastic elements or that the earlier proliferative lesions did not contain
transformed cells. It is not uncommon in MD for there to be mixtures of A- and
B-type lesions in peripheral nerves supporting the idea that the pathologic
scenario is complex.

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Lymphomas
Lymphomagenesis is the consequence of MDV infection that is most
commonly associated with MD. Lymphomatous tumors can appear as early as
12-14 dpi following infection of young, genetically susceptible chickens by a
highly virulent strain of MDV, or they may not appear until several weeks or
months after infection with less virulent viruses or under circumstances in which
there are moderating factors such as older age at infection, vaccinal immunity,
maternal antibodies, genetic resistance, etc. Lymphomas may occur in almost any
visceral organ and also in various other sites including muscle, peripheral nerves,
the eye, and skin. Lymphoid tumor in muscle tissue is shown in Figure 6-8.
Most MD lymphomas consist of a variable mixture of inflammatory cells,
immunologically committed and uncommitted lymphocytes, macrophages,
plasma cells, and neoplastically transformed T lymphocytes (Hudson and Payne,
1973; Powell et al. 1974; Payne and Rennie, 1975). Cellular infiltration is shown
in lung and liver samples in Figures 6-9 and 6-10, respectively. Those in visceral
organs may be mostly proliferative in character, but in other tissues, particularly,
skin, eye, and nerves, inflammatory lesions may predominate in some cases.
MDV infection in transformed tumor cells in largely nonproductive, although
occasional cells may “turn on” and show evidence of viral antigen and virions.
The selective involvement of various tissues is affected to some extent by both the
virus strains and the genotype of the host, but the reasons for this are unknown. It
could be that certain virus strains have an affinity for particular organs and the
ensuing infection there could attract transformed (or potentially transformable) T-

179
cells to the site with local lymphomas a the consequence. This appeared to be the
case when lymphomas were induced at the site of inoculation in the local-lesion
model (Calnek et al. 1989b). Also, as reported by Shearman and Longenecker
(1981), transformed T-cells may home to a particular site because of an organ-
specific metastasis-associated antigen.
The pathogenesis of the lymphomatous lesions remains somewhat enigmatic.
Much is known about some of the factors that affect the incidence of tumors, but
the exact mechanism(s) by which some of these operate is not well defined.
Essentially, it has been determined that a specific cascade of events leading to
infection of different populations of lymphocytes is critical to lymphomagenesis.
It begins with the infection of B-lymphocytes responsible for transformation of T-
cells. The two latter stages are influenced by the first. One factor seemingly not
involved in determining the probability of lymphoma development is not
susceptibility to infection per se. Although the incidence of tumors is influenced
by host genotype, age at infection, and virus strain, the intensity of the early
cytolytic infection is largely unaffected by these variables (Calnek
et al. 1998b; Lee, 1979). Of course, the early infection phase involves mostly B-
cells and, as Calnek et al. (1998) pointed out, there could be differences in the rate
of infection of activated T-cells associated with genotype, virus strain, etc.
The importance of the early cytolytic infection cannot be overstated. It is
undoubtedly this necrotizing infection and the ensuing inflammatory response that
stimulates the activation of T-cells infiltrating the site of active productive
infection. The greater the inflammation/immune response, the larger the number

180
of activated T-cells that might be expected to be present and therefore exposed to
the productively infected B-cells at the site. Indeed, enhancement of the
inflammatory/immune response at a given site of MDV infection increases the
probability of tumors (Calnek et al. 1989b). Conversely, the reduction or absence
of early cytolytic infection, whether due to maternal antibodies (Calnek, 1972;
Payne and Rennie 1975), vaccination (Calnek et al. 1983), virus serotype (Calnek
et al. 1979), virus attenuation (Calnek et al. 1983), or the absence of key early
target organs (Schat et al. 1981) correlates with the absence or reduced incidence
of lymphomas. Many of the modifying factors that affect tumor incidence are not
absolute in their effects. Vaccinal immunity is a good example since vaccination
usually does not totally eliminate MD, thus the more transformable targets that
become infected, the greater the likelihood that a successful transformation will
occur. Vaccination, as an example of a modifying factor, simply lowers the odds
by reducing the level of active infection which, in turn, has two key effects: (1) it
reduces the inflammatory response and thus keeps the number of activated T-cells
low, and (2) it eliminates foci of active infection where activated T-cells that are
already present could be infected.
Given the assumption that transformation is virus-directed, the most obvious
conclusion is that one or more “oncogenes” provoke transformation either directly
or indirectly. Approaches to identify various genes that could contribute to
oncogenicity of MDV have been described (Kawamura et al. 1991). Perhaps the
most interesting of these is meq, an MDV-encoded oncogene, which has
transactivating activity and whose structure resembles that of the jun/fos

181
oncogene family. Interference with the expression of several candidate
oncogenes, including meq, ICP4, and a 1.8kb gene family, but antisense
sequences appears to alter the transformed phenotype of a MD cell line,
suggesting that they are all involved in the maintenance of transformation.
Unfortunately, it is often difficult to take a direct approach in testing the
significance of specific candidate oncogenes. For instance, the lack of
oncogenicity of attenuated derivatives could be attributed to the inability of these
viruses to infect lymphocytes efficiently (Schat et al. 1982a), and the significance
of the apparent loss of oncogenicity in an MDV deletion mutant lacking a
functional meq coding region is tempered by the fact that the mutant virus
induced infection of chickens less efficiently than the oncogenic parent virus.
A role for a chromosomal aberration in the pathogenesis of lymphomas was
suggested by the discovery of an extra G-band on one homologue of the short arm
of chromosome-1 in several MD lymphoblastoid cell lines (Macera and Bloom,
1981; Moore et al. 1993). This extra band was due to the amplification of
genomic DNA, but the potential of its significance was diminished by several
subsequent observations: (1) it was infrequently seen in MD cell lines established
from MD local lesions (Moore et al. 1994); (2) the site of the aberration was not a
site of MDV integration (Delecluse and Hammerschimidt, 1993); and (3) there
was not a consistent correlation in MD cell lines between the occurrence of the
aberration and features usually associated with neoplastic transformation such as
colony formation in soft agar and transplantability (Moore et al. 1994).

182
Regardless of the molecular basis, transformation is a relatively rare event in
infected T-cells. If latently infected, activated T-cells all have the potential of
becoming transformed, and given the knowledge that large numbers of activated
T-cells are infected during the period of latency, then it must be concluded that
the transformation event itself is a very infrequent consequence of infection. This
is supported when the question of clonality of tumors is considered. Delacluse
and coworkers (Delecluse and Hammerschimidt 1993) proposed that MD tumors
are clonal in origin based on their finding that MDV DNA was randomly
integrated into the genomes of lymphoma cells but the pattern of integration sites
in the cells of a given lymphoma was consistent. On the other hand, multiple
tumors in the same bird can be different, i.e., they may represent different T-cell
phenotypes (Schat et al. 1991), and some cell lines established from individual
local MD lesions that contained transformed cells of more than one phenotype
(Calnek et al. 1998b).
Atherosclerosis
Fabricant and Fabricant (1999) made the highly interesting observation that
cell cultures infected with a feline herpesvirus contained both intracellular and
extracellular cholesterol crystals, and they noted that such viral infections might
have a role in degenerative vascular disease. Fabricant and Fabricant (1999)
hypothesized a possible causal relationship between herpesvirus infection and
human atherosclerosis. She and her coworkers’ subsequently tested the
hypothesis using a low-virulence strain of MDV and genetically susceptible, SPF
chickens (Fabricant and Fabricant, 1999). They found that the lesions induced by

183
MDV included proliferative and fatty-proliferative changes in aortic, coronary,
celiac, gastric, and mesenteric arteries. These changes were described as being
strikingly similar to those seen in occlusive atherosclerosis in humans (Minick et
al. 1979).
The pathogenesis of the lesion is of considerable interest from the viewpoint of
comparative medicine. Although supplemental cholesterol in the diet appeared to
enhance the severity of arterial lesions, the requirement for MDV infection as the
principal incident was proven by the failure of a high cholesterol intake alone to
produce atherosclerotic lesions, and more importantly, by the effectiveness of MD
vaccines in preventing atherosclerosis in MDV-challenged chickens (Fabricant
and Fabricant, 1999). Finally, it was shown that lipid metabolism was altered in
arterial smooth muscle cells infected by MDV in vitro] there was an accumulation
of phospholipids, free fatty acid, cholesterol, and cholesterol esters (Hajjar et al.
1985). Hajjar et al. (1986) detected similar alterations in cholesterol/cholesteryl
ester metabolism in vivo during early stages of the disease. In spite of these
observations about the association between the MDV herpesvirus and
atherosclerosis, the exact mechanism of the lesion evolution is presently
unknown. In view of the known capacity of MDV to induce proliferation in
certain other cells (Spencer 1969), it is interesting to speculate that a similar
response of infected smooth muscle cells, coupled with the known alteration in
lipid metabolism, might be a significant aspect of the pathogenesis of the lesion.

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Factors Affecting Pathogenesis
The incidence of MD following exposure to MDV can be highly variable.
Many factors are known to influence the likelihood that a given chicken will
develop pathologic lesions and clinical disease after infection. Virus-related
factors include both the serotype and the pathotype. Host factors include
genotype, age, and sex. Modifying factors related to the host are those associated
with its immune status, such as maternal antibody, immunosuppression from other
conditions, and vaccinal immunity.
Maternal Antibody
Jakowski et al. (1970) reported that MDV-infected SPF chicks developed an
acute disease characterized by severe hematopoietic destruction and lymphoid
organ degeneration, apparently due to the absence of MDV maternal antibody;
however, it is also possible that maternal antibody against chicken infectious
anemia virus (CIAV) may have been absent as well and that CIAV could have
contributed to the damage reported. A degree of protective efficacy against MD
mortality was attributed to the presence of maternal antibodies by several groups
(Ball et al. 1971; Chubb and Churchill, 1969; Spencer and Robertson, 1972).
Subsequently, passively acquired antibody, showing that its effect was to
markedly suppress the initial productive virus infection and acute inflammation
seen in lymphoid organs, became of importance. The ameliorative effect of
maternally derived protection is likely due to virus-neutralizing antibodies
directed against gB (Omar and Schat, 1996).

185
Vaccinal Immunity
Vaccinal immunity induced by any of the three vaccine serotypes has the
effect of markedly reducing the level of early cytolytic infection (Hitchner and
Calnek, 1980; Schat et al. 1982b; Smith and Calnek 1974), thus incriminating
antiviral immunity as a very significant factor in the protective efficacy of MD
vaccines. Based on cell-mediated cytotoxicity assays, it has been shown that
there are common CMI epitopes on cells infected with oncogenic or vaccine
viruses (Omar and Schat, 1996). The significance of antiviral immunity in
protection against lymphomas was confirmed by the efficacy of inactivated viral
antigens (Murthy and Calnek, 1979) and the ability of recombinant fowl pox virus
expressing the gB of MDV to induce protection against MD (Nazerian et al. 1992)
probably though the stimulation of CTLs. Also, although immunization does not
prevent infection, the level of latent infection and productive infection of the FFE
is low. Lee et al. (1999) reported that the MDV re-isolation rates from vaccinated
chickens were significantly lower than those of non-vaccinated chickens, and that
CD4+ T-cells appeared to be the preferential subset for MDV infection in both
groups, Omar and Schat (1997) identified a population of CD8+ TCRabl, but not
CD8+ TCRab2 nor CD4+, T-cells that were involved in antiviral immunity in
chickens vaccinated with serotype-2 SB-1 virus. Morimura et al. (1998)
confirmed that CD8+ T-cells are largely responsible for the control of viral
infection, and offered the hypothesis that these cells, in conjunction with other
effector mechanisms including NK cells, NK T-cells, double-negative T-cells, and
normal T-cells destroy the remaining infected cells and/or transformed cells. The

186
possibility that there is a direct effect on immune responses on transformed cells
is debatable, particularly since no unique tumor-associated antigens have been
found. On the other hand, it is very reasonable to expect that only the antiviral
activity of vaccines would be enough to reduce the incidence of lymphomas by
interrupting the normal cascade of pathogenetic events. The low level of latent T-
cell infection in vaccinated birds is evidence supporting this concept since this is
governed by the extent of T-cell activation and infection provoked by the
cytolytic/inflammatory phase. Three types of live vaccines are currently used to
protect commercial chickens against MD: (1) attenuated serotype 1 (MDV
HPRS-16/ATT and CVI0988), (2) naturally non-oncogenic MDV-2 (SB-1), and
(3) HVT-3 (FC-126)
Vaccination with attenuated MDV-1, non-oncogenic serotype 2 (e g., SB-1),
and/or serotype 3 (HVT) strains has been used successfully to prevent MD over
the past three decades, and represent one of the few cancer-preventing vaccines in
any species. Although vaccination reduces viral replication sharply during the
first cytolytic phase of infection (Calnek et al. 1981; Schat et al. 1982b) and
prevents tumorogenesis, it does not prevent the establishment of infection and
latency (Calnek et al. 1981). The mechanisms of vaccine-induced immunity are
only partially elucidated. It is clear that vaccine-induced immune responses
follow a pattern similar to that seen with pathogenic strains regarding
development of both virus-neutralizing antibodies (Onuma et al. 1975; Ikuta et al.
1984) and virus-specific CTL (Omar and Schat, 1996; Omar et al. 1998). In
addition, vaccination with HVT, SB-1, and especially HVT+SB-1, increases NK

187
cell activity (Uni et al. 1994). Studies using CVI988-vaccinated chickens
depleted of either CD4+ or CD8+ T-cells prior to challenge with the oncogenic
Md-5 strain strongly support the hypothesis that CD8+ CTL are needed for
antiviral immunity. Titers of Md-5 were significantly higher in vaccinated CD8-
depleted birds than in intact birds at the onset of latency (Morimura et al. 1998).
Interestingly, tumor formation or mortality did not occur in any of the vaccinated
CD4- or CD8-depleted chickens.
Vaccination may also reduce MDV-induced immunosuppression. For
example, vaccination with HVT prevented the impairment of phytohemagglutinin
(PHA)-stimulation of T-cells usually seen in unvaccinated, MDV-challenged
birds (Lee, 1979). This impairment had originally been attributed to “suppressor”
macrophages, but as discussed earlier, the suppressor cells may actually function
through the production of NO, which is beneficial in vaccinated birds by reducing
viral replication and allowing for more efficient T-cell responses.
The question of whether or not vaccination induces anti-tumor immune
responses has not been settled. It is likely that the reduction in virus load by
antiviral immune responses in combination with the absence of damage to the
lymphoid organs translates into a decreased level of MDV-positive target cells for
transformation. In addition, it is plausible that the intact, sensitized immune
system is able to maintain the latent state of the virus or eliminate cells in which
virus becomes reactivated, thus preventing the second lytic phase of infection.

188
Immunocompetence
MDV is, itself, immunosuppressive, and as discussed above, there may be a
relationship between immunosuppressive potential and virulence of certain MDV
isolates. It has not yet been determined whether the correlation among permanent
immunosuppression, late cytolytic infection, and lymphoma development is one
of cause or effect. Regardless, it is important to note that immunosuppression
from other conditions can affect the incidence of MD. Powell (1987) observed an
increase in MD incidence in chickens experimentally immunosuppressed with
betamethasone or corticosteroids. Subsequently, Buscagia et al. (1988) found that
either experimentally induced immunosuppression or natural incompetence based
on young age resulted in prolonged and more widespread early cytolytic infection,
thus interfering with the establishment of latency. Furthermore, immuno¬
suppression after latency had developed caused the reappearance of cytolytic
infection in the spleen suggesting that immunocompetence is also required for the
maintenance of latency. So, it appears that immunocompetence is very significant
in the pathogenesis of MD, and that a vicious cycle can be initiated in which
incompetence and failure to entry latency causes a continuation of cytolytic
infection, thus inducing even more destruction to the immune system and further
immunosuppression (Buscagia et al. 1988).
Conclusions
Clearly, the pathogenesis of MD is complex with many influential factors and
with many possible expressions of pathology. Probably, the most complex of the
varied potential outcomes of MDV infection is lymphomagenesis in which a very

189
complicated and specific cascade of events must occur. Viral pathotype and host
genotype are of critical importance in determining the likelihood of lymphoma
development and these affect the influence imparted by other modifying factors
such as age, maternal antibody, and vaccinal immunity. The underlying
mechanisms by which many of these various factors affect the events comprising
the pathogenesis of the disease are still the subject of investigation. Some of the
specific questions that remain to be investigated are:
(1) What are the exact relationships among permanent immunosuppression,
recrudescence of cytolytic infection, and lymphomagenesis, i.e., which comes
first and how does one influence the other?
(2) Why is transformation such a rare event in latently infected T-cells and
exactly what is the mechanism by which transformation is effected?
(3) Why are more virulent strains of virus more immunosuppressive and is this a
key factor in their enhanced virulence?
(4) How do transformed cells escape immunosuppression?
These, and other questions, will assure that the subject of pathogenesis will be of
continuing interest.
Immune Responses to MDV Infection
Introduction
MD, a herpesvirus-induced lymphomatous disease in chickens, has attracted
the interest of immunologists since MDV was isolated in 1968 and vaccines
became available shortly afterwards (Witter et al. 1979). Infection of chickens
with MDV is characterized by several distinct phases in which innate and

190
acquired immune responses play important roles. The first phase is characterized
by the replication of MDV in lymphoid cells, which are mostly B-lymphocytes.
The consequence of the lytic infection is that T lymphocytes become activated.
Only activated, but not resting, T-cells can be infected with MDV. Xing and
Schat (2000) hypothesized recently that a viral homologue of EL-8 (Vil-8) may be
involved in attracting the activated T-cells to the infected B-cells to facilitate the
transfer of virus. Intimate contact between B and T-cells is important for the
transfer of infectious virus from cell to cell (Kaleta, 1977), because MDV is
highly cell-associated (Schat et al. 1982b). Temporal immunosuppression of the
humoral immune responses is often one of the consequences of the lytic infection
in B-lymphocytes. During the second phase, a latent infection is normally
established in these activated T-cells, although some activated CD4+ and CD8+
T-cells may undergo a lytic infection (Baigent et al. 1979). The actual process of
establishment and maintenance of latency has not been elucidated, but it is likely
that a complex set of interactions, including immune responses and specific
cellular and viral genes, is responsible. The lytic and latent phases occur in all
MDV-infected chickens. Tumors may develop during the third phase of the
infection depending on the genetic resistance and immune status of the host
(Bacon et al. 2000), and the virulence of the virus strain. A secondary lytic
infection with subsequent immunosuppression often precedes the development of
tumors. The pathogenesis of MD can be modified by several immunologically

191
based factors such as the presence of maternal antibodies, vaccination status,
genetic resistance, stress level in the bird, and the presence of immunosuppressive
agents interfering with innate and acquired immune responses.
Innate Immune Responses
Introduction
Infection with MDV results in the activation of innate (nonspecific) and
acquired (specific) immune responses. The innate responses include activation of
macrophages, NK cells, and soluble factors such as cytokines, nitric oxide (NO),
and other soluble mediators. These responses are characterized by rapid
activation after infection, lack of specific antigen recognition, and lack of
memory. However, the distinction among innate and acquired responses in not
always clearly delineated. For example, interferon-gamma is an important
cytokine mechanism of CTL. Cytokines and soluble mediators, macrophages,
and NK cells are important in MDV-infected chickens (Schat et al. 1987).
Concluding Remarks
The next chapter details the cloning of MDV-1 glycorotein B (gB), gC, and gD
and HVT gC and gD. Plasmid DNA constructs will be confirmed by restriction
enzyme digest analysis. In vitro expression of such glycoproteins was confirmed
by I-IFA of transfected mammalian cells. MDV-1 gB was evaluated for indirect
in vivo expression in mice. Additionally, the in vivo fate of the MDV-1 gB
construct in various immune tissues and site of i.m. injection in chickens.

192
Figure 6-l.Immunodepressed MDV-infected chicken resulting in paralysis.

193
Figure 6-2. Gross observation of ocular effects of MDV infection. Left eye
is normal and right eye is effected.

194
Figure 6-3. Degeneration, necrosis, and cyst formation of MDV-infected
chicken bursa (Magnification at 100X).

195
Figure 6-4. Gross observation of uninfected (left) and MDV-
infection (right) chicken bursas.

196
Figure 6-5. Gross observation of the sciatic plexus and nerve from an
MDV-infected chicken.

197
Figure 6-6. Demyelination of nerves in MDV-infected chicken
(Magnification at 100X).

198
Figure 6-7. Cellular infiltration of nerve from MDV-infected chicken
(Magnification at 100X).

199
Figure 6-8. Gross observation of MDV-induced lymphoid tumor in the
pectoral major muscle.

200
Figure 6-9. MDV-infected large lymphoblastic cells from chicken lung
(Magnification at 100X). Lymphoblastic cells are noted with black arrows.

201
Figure 6-10. Lymphoid cell infiltration of MDV-infected chicken liver
(Magnification at 100X). Infiltrating lymphoid cells are noted with black
arrows.

CHAPTER 7
CREATION OF PLASMID DNA CONSTRUCTS ENCODING SEROTYPE-1
MAREK’S DISEASE VIRUS (MDV-1) AND HERPESVIRUS OF TURKEY’S
(HVT) GLYCOPROTEINS
Introduction
Project Background
The main purpose of this study was to evaluate in vivo murine protein
expression indirectly. Furthermore, the initial fate (after 12 hours) of a naked
DNA construct encoding Marek’s disease virus-glycoprotein B (MDV-gB) in
tissue sections of MDV-gB-injected three-week-old leghorn chickens.
Additionally, this study involved the creation of plasmid DNA molecules
expressing MDV-1 gC and gD and HVT gC and gD.
Background on Monoclonal Antibody Production
The technique of producing hybridomas required culture of myeloma cells
lines that grow in normal culture medium but not in a defined “selection” medium
because they lack functional genes required for DNA synthesis in this selection
medium. Fusing normal cells to these defective myeloma fusion partners
provided the necessary genes from the normal cells, so that only the somatic cells
hybrids grew in the selection medium. Moreover, genes from the myeloma cell
made such hybrids immortal. Inducing defects in nucleotide synthesis pathways
created cell lines that were used as fusion partners. Normal animal cells
synthesize purine nucleotides and thymidylate, both precursors of DNA, by a de
202

203
novo pathway required tetrahydrofolate. Antifolate drugs, such as aminopterin,
block activation of tetrahydrofolate, thereby inhibiting the synthesis of purines
and thereby preventing DNA synthesis via the de novo pathway. Amonopterin-
treated cells were able to use a salvage pathway in which purine is synthesized
from exogenously supplied hypoxanthine using the enzyme hypoxanthine-
guanine phosphoriboslytransferase (HGPRT), and thymidylate is synthesized
from thymidine using the enzyme thymidine kinase (TK). Therefore, cells grew
normally in the presence of aminopterin if the culture medium is also
supplemented with hypoxanthine and thymidine (called HAT medium). Myeloma
cell lines were made defective in HGPRT or TK by mutagenesis followed by
selection in media containing substrates for the enzymes that yield lethal products.
Only HGPRT- or TK- cells survived under these selection conditions. Such
HGPRT- or TK- myeloma cells could not use the salvage pathway and will,
therefore, died in HAT medium. When normal B-cells were fused to HGPRT- or
TK- cells, the B-cells provided the necessary enzymes, so that the hybrids
synthesize DNA and grow in HAT medium.
After fusion of B-cells with myeloma cells in PEG, the cells were allowed to
grow in selective HAT medium in 96-well plates. After the second feeding,
supernatants were screened by I-IFA of transfected cells. Whichever wells
yielded positive results were then subcloned by limiting dilution. By microscope,
one had to investigate these wells for growth that appears to be of clonal origin.

204
Such well supernatants were further screened by I-IFA. At least three subcloning
steps of positively reacting wells were needed to “weed-out” any contaminating
B-cells of different specificity.
Project Design
Initial Fate of Injected pTarget/MDV-1 gB in Chickens
Test chickens were injected i.m. with 500pg MDV-gB in sterile IX phosphate-
buffered solution (PBS). Negative control chickens were injected i.m. with sterile
IX PBS. This project investigated the in vivo fate of MDV-gB DNA in muscle,
spleen, bursa, thymus, and cecal tonsil tissues by intramuscular (i.m.) injection at
12 hours post injection.
Indirect In Vivo Expression Of MDV-1 gB In Mice
BALB/C mice were i.m. injected with pTarget/MDV-1 gB at 250pg and
boosted with the construct. Tail-bled mice that tested positive for anti-MDV-1 gB
sera were chosen for the generation monoclonal antibodies.
Plasmid DNA Constructs of MDV-1 and HVT Glycoproteins
pTarget was the DNA vector used to clone MDV-1 and HVT glycoproteins,
gC and gD. After restriction enzyme analysis confirming proper gene orientation,
in vitro expression was performed after transfection by electroporation.
Materials and Methods
Mice
8-week old female BALB/C mice were ordered. Upon arrival, mice were
raised in cages within a negative pressure room. Mice were fed ad lib a
commercial mouse chow.

205
Chickens
Leghorn chicken eggs were incubated and hatched in Serengeti and Safari
incubator models, respectively. 21 chicks were subsequently transferred into a
Horrfoll-Bauer type isolator operated under negative pressure and filled with high
efficiency particulate air (HEPA) filters. Chickens were feed ad libitum. They
were also allowed to drink water ad libitim.
MDV-1 and HYT Virus Culture
105 primary CEFs were plated in DMEM with 5% FBS and cultured for about
3 days until contact inhibition. Primary CEFs were trypsinized to establish
secondary CEF cultures. After about 3 days of culture, either MDV-1 RB1B or
HVT inocula were added to the cells in culture. After 30 minutes, media and non¬
cell-bound virus were removed and new DMEM with 5% FBS was added to
CEFs. Plates were monitored for virus-induced cytopathic effect. Infected
secondary CEFs were removed from culture plates by scrapping method. CEFs
were collected in 50mL screw-capped tubes and spun at lOOOrpm for 10 minutes
at 4°C. The supernatant was removed and the pellet saved. The pellet was rinsed
with 9mL of sterile IX PBS and spun again at lOOOrpm for 10 minutes at 4°C.
TE buffer at pH 7.5 was added to the pellet to give a total volume of 15mL.
Gentile resuspension was done using the Bélico Tissue Homogenizer and
homogenized 35 times. The homogenized suspension was centrifuged at
3000rpm for 20 minutes at 4°C. At this point, cell debris was pelleted and the
virus was in the supernatant. The supernatant was collected into a 50mL Sarstedt
tube. In the hood, 5mL of 25% Sucrose/TE was placed into a small Beckman

206
Ultracentrifuge tube with which viral suspension was carefully added. The
sample was ultracentrifuged at 28,000rpm for 1.5 hours at 4°C. The supernatant
was removed and the tube well drained. The viral pellet was dissolved with 2mL
of cold TE buffer pH 7.4-7.6. The sample was vortexed gently and pipetted to
ensure complete suspension. During this procedure, pipette tips were cut in half
so as not to shear viral DNA. To this sample, 200uL of 10% Triton and 5uL of 2-
beta-mercaptoethanol was added(in hood only). The sample was vortexed gently
and placed on ice for 10 minutes while mixing occasionally. The sample was
placed through another 25% sucrose cushion. Samples were then pelleted before
adding 500uL of cold TE buffer pH 7.4 - 7.6. The pellet was completely
resuspended by gently vortexing and pipetting. If necessary, leave at 4°C to soak
the pellet and resuspended again. At that point, cells were treated with Trizol
Reagent to isolate DNA. After the Trizol DNA isolation procedure, viral DNA
was precipitated with 2uL of glycogen, 1/3 volume of 3M NaAcetate, and 3 times
volume of ice cold ethanol. DNA was precipitated overnight at -80°C. DNA was
pelleted by centrifugation at 4,500rpm for 20 minutes at 4°C. DNA was then
washed with 3 times with a 3-time volume of 70% ethanol. The cleaned DNA
pellet was drained well and allowed to dry in the laminar flow cabinet. When
completely dry, the DNA was dissolved in 200-250uL of TE buffer at pH 7.4 -
7.6.
Transformation of Competent Bacteria
Individual blue colonies of JM109 E. coli were picked into 5mL of 2X YT
broth in Sarstedt 30mL screw-cap tube and shaken overnight at 37C at 275rpm.

207
25mL of 2X YT broth was placed in a 125mL Erlenmeyer flask to which was
added 500uL of overnight culture. New cultures were shaken at 275rpm for 1.5
hours. Cells are best when absorbance OD600 is between 0.3 - 0.4. Cells were
spun at 2000rpm for 10 minutes at 4°C. Tubes were drained completely before
addition to pellet of bacteria 2.4mL of TSS. The bacterial pellet was resuspended
carefully so that no frothing occured. Freeze at -70C in aliquots of 0.5-lmL.
Agar/2X YT for Bacterial Growth
Approximately 1L of mQ water was placed in a 4-liter Erlenmeyer flask and
stir with a Teflon bar. 16g of Bactotrypton, lOg of Bacto-Yeast Extract, 5g of
NaCl, and 15g of Bacto-Agar were added. The contents were stirred for a few
minutes to wet and dissolve the ingredients. Not all of the components dissolved
completely, which is normal before autoclaving. Contents were poured into a 2L
cylinder and brought up to a volume of 1 5L. Contents were poured back into an
Erlenmeyer flask and the cylinder was rinsed with 500mL of mQ water.
Therefore, the Erlenmeyer flask contained a total of 2L. Sterilization by
autoclaving the flask in fluid cycle for 20 minutes was performed.
Ampicillin Solution
Two grams of ampicillin (Sigma, Inc.) were dissolved in 20mL of mQ water
and sterilized by filtration through a 0.2um mini-filter directly into sterile 30mL
Sarstedt tubes. This stock was stored in the refrigerator.

208
X-Gal Solution
250 mg of X-gal (S-bromo-4-chloro-3-indolyl-B-D-Galactopuranoside,
molecular weight 408.6) was dissolved into lmL of dimethylformamide. This
stock was stored in the refrigerator.
Blue/White Selection for Agar Plates
2X YT agar was melted in a microwave oven. Agar was cooled at room
temperature and equilibrated at 50°C in water bath. For a stock agar sample of
lOOmL volume, addition of lOOuL of ampicilin (lOOmg/mL), lOuL of X-gal
(250mg/mL), and 12.5uL of IPTG (200mg/mL) was performed. Agar was
poured onto bacteriological 100mm plates in a laminar flow cabinet and allowed
to dry for half an hour. Selection plates were sealed and stored at 4°C until use.
Amplification and Cloning of MDV-1 and HVT Glycoproteins
PCR was performed on isolated DNA from either MDV-1 RB1B or HVT.
PCR was set for 35 cycles (99°C, 60°C, 72°C) and allowed a final extension cycle
of 72°C for 1 hour. Samples were then refrigerated. Oligonucleotide primers
were designed for subsequent site-specific cloning into the pTarget vector. The
following primers were used to amplify specific MDV-1 and HVT glycoproteins:
forward primer (5’atgctaaacgatggagttgtg) and reverse primer (5’taggactcg-
gagaatactagag) for MDV-1 gC; forward primer (5’atgctagcgcagcttcggtatcggtag)
and reverse primer (5’atggtacagtacattacatcgcaatc) for MDV-1 gD; forward primer
(5’atgctagctaccatattacgcatcttatcg) and reverse primer (5’attggtaccagtt-
gtcctggactggaatc) for HVT gC; and, forward primer (5’tggctagcaaat-
actgagttgaactta) and reverse primer (5’taggtaccacgtactggctataaacg) for HVT gD.

209
pTarget/MDV-1 gB Injection of Mice
Enough of this epitope to inject into mice was prepared. The immunogen can
be isolated, synthesized, or recombinantly produced or encoded. For this
question, injection of DNA encoding antigen is described. Young female mice
(about 8-weeks old) were chosen because they are the best antibody producers. A
humoral immune response must be in the mice. For this project, 250pg of DNA
construct per i.m. injection. After waiting 10 days, mice were tail-bled for
antiserum and evaluated in an indirect immunofluorescence assay (UFA) using
antigen-transfected COS-7 cells. After no humoral response occurred on I-IFA
(data not shown), immunization of mice was performed again 2 weeks later.
Because no antibody response was detected (data not shown), it was decided that
the injection dose should be increased to 500|ig per injection. Upon 10 days after
injection, serum was collected and reacted in UFA. A positive response resulted
with immunized mouse serum in reference to controls, which included positive
serum on transfected and untransfected cells and negative serum on transfected
and untransfected cells. At this point, confirmation of anti-MDV-1 gB sera in
mice was detected. To increase the amount of B-cell producing such specific
antibodies, along with antibody affinity maturation, mice were boosted again 2
weeks later. Similar testing of mouse serum in UFA yielded even strong positive
results.
pTarget/MDV-1 gB Injection of Chickens
Leghorn chickens were injected into the left pectoralis major muscle about
0.25 inch deep with 500|ig of MDV-1 gB construct. The injected construct

210
solution in IX PBS also contained trypan blue in order to specifically mark the
site of i.m. injection.
Transfection of Mammalian Cells with pTarget/MDV-1 aB
Transfection of COS-7 cells with MDV-1 gB was performed by
electroporation. Briefly, COS-7 cells were cultured in DMEM/BSA (5%) for 2
days. Upon typsinization, 10xl06 COS-7 cells were resuspended in sterile IX
PBS. 5p.g of MDV-1 gB naked DNA were added to the cells. Electroporation
was performed at .6kV for 1 second. Transfected cells were then added to 8-well
chamber slides containing DMEM/BSA (5%) and cultured for 36 hours.
I-EFA of MDV-1 gB
MDV-1 gB transfected COS-7 cells were evaluated using MDV-1 gB
polyclonal chicken anti-sera as primary reagent in triple-sandwich, indirect
immunofluorescence assays. Subsequent I-IFA was performed with mouse anti-
MDV-1 gB serum.
Transfection of Mammalian Cells with MDV-1 and HVT Glycoproteins
Each MDV glycoprotein-encoded construct will be tested for in vitro
expression in mammalian cell lines. Briefly, 4xl06 cells in sterile IX PBS were
electroporated at 0.6kV with 3|ig of DNA construct. Transfected cells are plated
onto chamber slides containing DMEM and 5%FBS.
Transfection and I-IFA of other MDV-1 and HVT Glycoproteins
COS-7 cells were transfected with either MDV-1 or HVT Glycoproteins, as
described for MDV-1 gB. Cells transfected with DNA constructs, based on the
pTarget vector, also contained G418, a selectable antibiotic. After 2 days of

211
culture at 39°C, cells were fixed with ice cold methanol for 5 minutes. Primary
anti-sera for each MDV-1 or HVT glycoprotein, either polyclonal or monoclonal,
were diluted in PBS/3%BSA at 1:10 for overnight incubation of slides. Upon
washing slides 3 times with mQH20 and PBS/3%BSA, secondary reagents (i.e.
anti-mouse or anti-chicken antibodies) were allowed to react for 45 minutes.
After an identical washing procedure was performed, tertiary reagent was added
for 45 minutes, biotinylated-streptavidin. A final wash was performed before
addition of PBS/DABCO solution. Expression of viral glycoproteins was
evaluated using a fluorescent microscope [Zeiss, Inc],
Required Media for Monoclonal Antibody Production (Products Available at
Gibco. Inc)
DMEM with 10% serum supplement
This media was needed for culture of HL-4 myeloma cells. Note: 400-500mL
of this media is more than sufficient for a single fusion.
“Re-filtered DMEM “
This media was needed for various washing steps. Note; 300-400mL of this
media is more than enough for a single fusion.
Fusion Media #1: DMEM with 25% HL-4 condition media, 20% serum
supplement
IX HAT
This media was needed for seeding the newly fused cells and for performing
the first 2-3 feedings of cells in the 96-well plates. Note: 200mL of this media is
more than enough for a single fusion.

212
Fusion media #2
Fusion Media #2 was composed of DMEM with 25% HL-4 conditioned
media, 20% serum supplement, and IX HT. This media was needed for
maintaining 96-well plates and transferring antibody positive 96-well plate
cultures into 24-well plates and feeding the 24-well plates. Note: 150-200mL of
this media is more than enough for a single fisión.
Fusion media #3: DMEM with 15% serum supplement and 0.5X HT
This media was needed for maintenance of 24-well plate cultures and initial
transfer from 24-well plates to 6-well plates. Note: I00-150mL of this media is
more than enough for a single fusion.
Growth media
Growth media was composed of DMEM with 15% serum supplement. This
media was needed for maintaining macroscopic colonies in 6-well plates for
freezing.
Washing of HL-4 Myeloma cells
HL-4 myeloma cells were expanded in DMEM with 10% serum supplement.
Before fusion with splenocytes, HL-4 cells were washed in “Refiltered” DMEM
without serum supplementation.
Processing of Mouse Spleen
A mouse was anesthetized in a 50mL conical tube containing Metofane. The
right eye was removed and the head was gently squeezed to drain about 1 mL
blood into a serum tube. The sera were then screened for antibody production.
The spleen was aseptically excised and then placed into a sterile plate containing

213
“Re-filtered DMEM”. The spleen was punctured with a 19 gauge needle and as
many splenocytes as possible were squeezed from the capsule. Note: this was
completed upon significant color change of the capsule (from red to whitish).
This splenocyte “soup” was drawn through the following size needles 2-times in
this order: 20 gauge, 23 gauge, and then a 25 gauge needle. After washing and
extracting spleen cells, the splenocyte soup was divided into 2-50mL conical
tubes and centrifuged at 2,000rpm for 10 minutes. After centrifugation, the pellet
of each conical tube was resuspended in the same volume of fresh “Re-filtered
DMEM”. In a 1.5mL microfuge tube, O.lmL splenocyte soup was mixed with
0.4mL 3.0% glacial acetic acid. Note: the acetate lyses RBCs with only
mononuclear cells remaining. After counting the splenocytes, the amount of HL-
4 cells needed for a 7:1 dilution of splenocytes to HL-4 cells was determined.
Cells were added into a conical tube and centrifuged at 2,000rpm for 8 minutes.
Note: HL-4 cells are white and splenocytes red. Very carefully, the supernatant
was decanted into a waste beaker. Briefly, the pellet was drained onto sterile
gauze. Note: immediately proceed to the fusion step.
Performing Hvbridoma Fusion
A water bath was set for exactly 37°C. After this temperature was reached, 2-
conical tubes were placed individually containing PEG1500 (l.lmL) and “Re¬
filtered DMEM with antibiotic/antimycotic” (30. lmL) into the water bath. Both
of these tubes were warmed for at least 10 minutes to ensure exact temperature of
reagents. A timer was set for exactly 6.5 minutes. The timing began as soon as
the first reagent was added. To the splenocytes/HL-4 pellet, lmL 50% PEG1500

214
was added with a glass pipet. This was a time-sensitive step in that this lmL PEG
was to be gently added over the course of only one minute. As soon as possible,
30mL “Re-filtered DMEM with antibiotic/antimycotic” was pipetted. The PEG-
treated splenocyte/myeloma cells were diluted with “Re-filtered DMEM with
antimicrobiotics”. Note: this media should only be added at a rate of lmL/every
10 seconds over a period of 10 mimites. This suspension was centrifuged at
2,000rpm for 8 minutes. The supernatant was carefully decanted into the waste.
Inverting the tube over sterile gauze drained the pellet. The pellet was
resuspended in 35-50mL “Fusion Media #1”. 9.6mL of these cells were seeded
into 96-well flat-bottom plates using lOOpL/well. Note: this step has to be
performed using large orifice pipet tips on a 12-channel pipet. After seeding
cells, an additional lOOpL/well of “Fusion Media #1” was added. Therefore, each
well contained 200pL of suspended cells that should form monolayer-like
colonies. Note: the amount ^“Fusion Media #1” should be determined such that
the final cell concentration is about 2.0 x 105 cells/well. 96-well plates were
placed into incubator @ 37°C. Note: cells should usually form colonies on top of
each other in about 3-5 days.
Care and Feeding of Potential Hybridoma Cells
The first feeding usually occurred between 3-5 days. 175pL of media was
removed from each well very slowly using a small orifice pipette tip. Note: do
not disturb cells. 200pL of Fusion Media # 1 was added to each well using a
large orifice pipet tip. Plates were placed back into the 37°C incubator. Note: be
aware ofpotential fibroblast growth since this may be a common event. About 3-

215
5 days later, colony formation checked upon second feeding. Note: the larger
cells should be hybridoma cells. Depending upon how well the hybridoma cells
grew, the addition of either Fusion Media #1 or Fusion Media #2 was be chosen.
Note: in general, continually re-feed plates with Fusion Media #1 until it is gone.
Then begin re-seeding with Fusion Media #2. However, if upon the second
feeding you see a good amount of hybridomas, then you can proceed to re-feeding
with Fusion Media #2. By day 7 postfusion, notice macroscopic colony formation
appeared. Primary antibody screening was usually performed 10-14 days
postfusion. Note: the 96-well plate supernatants should be screened once the
media changes to a yellow color, along with macroscopic colony formation, and
the 96-well plates should be fed a minimum of 2-3 times before supernatants are
removed for primary antibody screening. Removal of supernatants for screening
was done with a dedicated tissue culture pipetteman using small orifice, aerosol
pipette tips. Note: usually 100pL is a sufficient volume for screening. Once the
supernatants have been removed, the well should be re-feed with Fusion Media #1
or #2, depending upon the culture growth characteristics. For the 96-well plates,
positive samples from primary screening were immediately transferred to 24-well
plates. This transfer was done using a sterile cotton-plugged Pasteur pipette. To
set-up these 24-well plates, l-1.5mL Fusion Media #2/well (maximum of
2.5mL/well) was added. Then a small amount of this media was pipetted (large
orifice) from each well to help re-suspend the hybridoma colonies. Note:
carefidly pipet “up and down ” several times to resuspend the hybridomas in the
96-well plate. In addition, make sure that you transfer all of the potential

216
hybridoma cells into the 24-well plate. After mixing cells in a 24-well plate, a
small amount was added back into the original 96-well plate well. Note: the 96-
well and 24-well plates should be placed into different incubators in case of
incubator failure that may residí in destroying the samples. The cells in the 24-
well plate were monitored for growth and fed as needed. In general, all the
Fusion Media #2 was used. When this reagent was gone, cells were re-feed with
Fusion Media #3. Note: when the cells become confluent and the media changes
color to yellow, the supernatants shoidd be subjected to secondary antibody
screening assays. Usually, 0.5-1. OmL of supernatant is removed from these
larger plates. Positive results from the secondary antibody screening was the
beginning of the third cell plate transfer into 6-well plates. As with the transfer
from 96-well to 24-well plates, the transfer to 6-well plates was similar. 7mL of
Fusion Media #3 was added into each well of a 6-well plate. The samples were
gently pipetted “up and down” in 24-well plates and transferred to a 6-well plate.
After the transfer, the original 24-well plate was re-seeded with 1.0-1.5mL (as a
back-up). Cells in the 6-well plates were monitored for subsequent growth. Note:
when the cells reach log phase and appear confluent, a vial should be
frozen in liquid nitrogen in “Growth Media”. A second set of cells shoidd be
saved on the next day. After the 2-vials from the 6-well plate macroscopic
cultures were frozen and supernatants collected, the cells, including all back-up
plates, may be discontinued.

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Results
Indirect in vivo expression of MDV-1 gB
Upon injection and boosting of mice with MDV-1 gB construct, mice were
able to produce polyclonal anti-MDV-1 gB serum.
Initial Fate of Injected pTarget/MDV-1 gB in Chickens
In order to evaluate plasmid construct fate upon i.m. injection, the sensitivity
of MDV-1 gB PCR was investigated. Results demonstrate a sensitivity of lOOfg.
12 hours after infection chicken tissue samples were collected and plasmid DNA
was extracted, including the site of muscle injection, bursa of Fabricius, spleen,
thymus, and cecal tonsils. Results demonstrated that at 12 hours only muscle
tissue contained the MDV-1 gB construct.
MDV-1 aC
Restriction analysis confirmed proper orientation of gC insert into pTarget. In
vitro expression was confirmed by I-IFA.
HVT gC
Restriction analysis confirmed proper orientation of gC insert into pTarget. In
vitro expression was confirmed by I-IFA.
MDV-1 gD
Restriction analysis confirmed proper orientation of gC insert into pTarget. In
vitro expression was confirmed by I-IFA.
HVT eD
Restriction analysis confirmed proper orientation of gC insert into pTarget. In
vitro expression was confirmed by I-EFA.

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Discussion
Indirect in vivo expression of MDV-1 gB
The results of this project demonstrated, indirectly, that in vivo expression of
MDV-1 gB occurred. Polyclonal antiserum generated in injected mice is
evidence that the protein must have been produced in vivo. The production of
such antiserum was confirmed by positive I-IFA results on MDV-1 gB transfected
mammalian cells. Therefore, in vivo expression of MDV-1 gB can occur in mice.
This is the first study directly reporting in vivo expression of MDVv-1 gB in mice
by I-IFA.
Initial Fate of Injected pTarget/MDV-1 gB in Chickens
Results of the initial fate of construct injected chickens demonstrated the
presence of the construct only at the site of muscle injection. Other immune
tissues, such as bursa of Fabricius, thymus, and cecal tonsils, did not yield
positive PCR results, despite the sensitivity of the assay being lOOfg. It should be
noted that in vivo fate of this molecule may have yielded different results if this
study was extended to include other time points, such as 48 or 72 hours.
Plasmid DNA Constructs Expressing MDV-1 and HVT Glycoproteins gC and gD
As all glycoprotein constructs were confirmed for proper insert orientation, in
vitro expression was demonstrated by I-IFA on transfected mammalian cells.
Further investigation should be performed to evaluate whether in vivo expression
of such constructs exists in chickens.

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Bglll 834?
MDV-1 gB
Figure 7-1. Plasmid map of MDV-1 gB-encoding DNA construct.

220
Figure 7-2. I-IFA of MDV-1 gB-transfected COS-7 cells using anti-MDV-
1 gB sera from immunized mice (Magnification at 200X).

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Plasmid amount
3.0kb
2.0kb
lOOng lOng lng lOOpg lOpg lpg lOOfg* lOfg
Figure 7-3. MDV-1 gB (RB1B) PCR sensitivity of lOOfg.

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M, S, T, CT, B N Pos (10ng)
3.0kb
2.0kb
M=pectoralis major
S=spleen
T=thymus
CT=cecal tonsils
B=bursa
Figure 7-4. PCR results showing the fate of MDV-1 gB in chicken
tissues by PCR (12hr after injection).

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CMVp/e
SV40 p(A)
Figure 7-5. Plasmid map of MDV-1 gC-encoding DNA construct.

224
StuI, EcoRI, and Asp718
3.0kb
2.0kb
1.6kb
Figure 7-6. Restriction enzyme digest confirming the
integrity of pTarget:gCl.

225
Figure 7-7. In vitro I-IFA of MDV-1 gC with chicken polyclonal
antiserum (Magnification at 200X).

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CMVp/e
SV40 p(A)
Figure 7-8. Plasmid map of HVT gC-encoding DNA construct.

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BamHI
EcoRV
1.6kb
Figure 7-9. Restriction enzyme digest for integrity of HVT gC construct.

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Figure 7-10. I-EFA of gC3-transfeted COS-7 cells using anti-MDV
sera from immunized mice (Magnification at 200X).

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CMVp/e
Figure 7-11. Plasmid map of MDV-1 gD-encoding DNA construct.

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Sall=linearize
EcoRI^drop-out
BglII=orientation
Figure 7-12. Restriction enzyme digests for integrity of serotype 1 MDV gD

Figure 7-13. I-IFA of MDV-1 gD with chicken polyclonal antiserum
(Magnification at 200X).

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CMVp/e
Figure 7-14. Plasmid map of HVT gD-encoding DNA construct.

233
EcoRI
Ndel
Figure 7-15. Restriction enzyme digests for integrity of pTarget:gD3 construct.

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Figure 7-16. I-IFA of HVT gD using polyclonal
chicken antisera (Magnification at 200X).

CHAPTER 8
CYTOKINE LITERATURE REVIEW
Introduction
General Cytokine Background
The term cytokine, or immunocytokines, was used initially to separate a group
of immunomodulatory proteins that modulate the proliferation and bioactivities of
non-immune cells (Thomas and Balkwill, 1991). Some cytokines are produced
by a rather limited number of different-cell types while others are produced by
almost the entire spectrum of known cell types. The initial concept of "one
producer cell -one cytokine -one target-cell" has been negated for almost every
cytokine investigated more closely. Today the term cytokine is used as a generic
name for a diverse group of soluble proteins and peptides, which act as humoral
regulators at nano- to picomolar concentrations and which, either under normal or
pathological conditions, modulate the functional activities of individual cells and
tissues. These proteins also mediate interactions between cells directly and
regulate processes taking place in the extracellular environment. Many growth
factors and cytokines act as cellular survival factors by preventing programmed
cell death. In many respects the biological activities of cytokines resemble those
of classical hormones produced in specialized glandular tissues. Some cytokines
also behave like classical hormones in that they act at a systemic level, affecting,
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236
for example, biological phenomena, such as inflammation, septic shock, and acute
phase reaction, wound healing, and the neuroimmune network (Kennedy and
Jones, 1991).
In general cytokines act on a wider spectrum of target-cells than hormones.
Perhaps the major feature distinguishing cytokines from mediators regarded
generally as hormones is the fact that, unlike hormones, cytokines are not
produced by specialized cells which are organized in specialized glands, i. e. there
is not a single organ source for these mediators. The fact that cytokines are
secreted proteins also means that the sites of their expression do not necessarily
predict the sites at which they exert their biological function (Arai et al. 1990). In
the more restricted sense cytokines comprise interleukins, initially thought to be
produced exclusively by leukocytes, lymphokines, initially thought to be
produced exclusively by lymphocytes, monokines, initially thought to be
produced exclusively by monocytes, interferons, initially thought to be involved
in antiviral responses, colony stimulating factors, initially thought to support the
growth of cells in semisolid media, chemokine, thought to be involved in
chemotaxis, and a variety of other proteins (Hall and Rao, 1992). Almost all
cytokines are pleiotropic effectors showing multiple biological activities. In
addition, multiple cytokines often have overlapping activities and a single cell
frequently interacts with multiple cytokines with seemingly identical responses
(cross-talk). One of the consequences of this functional overlap is the observation
that one factor may frequently functionally replace another factor altogether or at
least partially compensate for the lack of another factor (Ono et al.2003). Since

237
most cytokines have ubiquitous biological activities, their physiologic
significance as normal regulators of physiology is often difficult to assess
(Atamas, 2002).
TH1 vs. TH2 Cytokine Profiles
The termTHi cytokines and TH2 cytokines refers to the patterns of cytokines
secreted by two different subpopulations of murine CD4(+) T-cells that determine
the outcome of an antigenic response toward humoral or cell-mediated immunity.
Numerous cells other than T-cells expressing CD4 have been shown to be capable
of producing TH1 cytokines and TH2 cytokines. These cells include CD8 (+) T-
cells, monocytes, natural killer cells, B-cells, eosinophils, mast-cells, basophils,
and other cells (Janeway et al. 1988). Type-1 cytokines include EL-2, EFN-gamma,
EL-12, and TNF-beta, while Type-2 cytokines include EL-4, IL-5, EL-6, IL-10, and
IL-13. Type-1 helper cells (TEE 1), but not type-2 helper cells (TEE2), secrete EL-2,
EFN-gamma, and TNF-beta, whereas TEE2 cells, but not TEE 1 cells, express EL-4,
EL-5, EL-6, and EL-10. Murine TEE2 cells, but not TH1 cells, also express P600,
the human counterpart of which has been identified as EL-13 (Clerici and Shearer,
1993). A novel cytokine inducing the synthesis of EFTSi-gamma in TH1 cells has
been identified recently as EGEF (and renamed EL-18). The molecular
mechanisms underlying the evolution of these two different-cell types from
common precursors are still not completely known. Studies with transgenic mice
carrying null mutations of the EL-4 gene have shown that EL-4 plays an important
role in the establishment of a functional TH2 immune responses (Romagnani,
1991).

238
The different patterns of cytokine secretion correspond with different functions
as immune effectors (Cherwinski et al.1987). TH1 cells promote cell-mediated
effector responses. TH2 cells are mainly helper cells that influence B-cell
development and augment humoral responses such as the secretion of antibodies,
predominantly of IgE, by B-cells. Both types of TH cells influence each other by
the Cytokines they secrete; IFN-gamma, for example, can down-regulate TH2
clones while TH2 cytokines, such as IL-10, can suppress TH1 functions. IFN-
gamma has been shown also to inhibit the proliferation of murine TH2 cells but
not that of TH1 helper T-lymphocyte clones (Kawakami and Parker, 1992). It
thus appears that these functional subsets are mutually antagonistic such that the
decision of which subset predominates within an infection may determine also its
outcome. Murine IL-12 has been shown recently to alter differentiation of a
subset of CD4 (+) cells and to be involved in the induction of protective immunity
against intracellular parasitic infections in mice. IL-12 appears to prevent
deleterious TH2 T-cell responses and to promote curative TH1 responses in an
IFN-gamma dependent fashion during murine leishmaniasis. Another exogenous
factor also influencing the development of undifferentiated CD4 (+) T-cells
towards either the TH1 or TH2 phenotype is TGF-beta. A third subset of memory
cells, designated THO cells and believed to be precursor cells that develop into
either TH1 or TH2 cells, can produce all Cytokines found to be secreted either by
TH1 or TH2 cells at low levels (Gajewski and Fitch, 1988). It appears that

239
different types of helper cell populations resembling those observed in mice are
found also in humans (Rooney et al., 1994). However, the differences in cytokine
expression seem to be quantitative rather than qualitative.
Introduction to Interferons
Interferon Background
By definition, interferons are proteins that, at least in homologous cells, elicit
a virus-unspecific antiviral activity. This activity requires new synthesis of RNA
and proteins and is not observed in the presence of suitable RNA and protein
synthesis inhibitors. Apart from their antiviral activities interferons also possess
anti-proliferative and immunomodulating activities and influence the metabolism,
growth and differentiation of cells in many different ways (Borden, 1992).
General actions and activities of interferons
The three main human interferons are known as IFN-alpha, IFN-beta, and
IFN-gamma. IFN-alpha and IFN-beta are called also Type-1 interferon. Bovine
TP-1 (trophoblast protein-1) is also a type-1 interferon. IFN-delta is classified
also as a type-1 interferon. IFN-gamma has been designated Type-2 interferon.
Another protein, called originally IFN-beta-2, is not an interferon but is identical
with IL-6. Some older names of interferons such as leukocyte interferon (IFN-
alpha), fibroblast interferon (IFN-beta) and immune interferon (IFN-y) are still in
use. These names are derived from the main producers and from the typical
bioactivity and still reflect the concept of a typical producer cell. This concept
has been discarded now because it is known that a plethora of different-cell types
is capable of producing interferons (Garbe and Krasagakis, 1993).

240
Interferons are a heterogeneous group of proteins with some similar biological
activities that are distinguished from each other by many different physical and
immunochemical properties. They are also encoded by different structural genes.
Most interferons are multifunctional proteins with bioactivities that are strictly
species-specific. These substances are synthesized following the activation of the
immune system. Two different single genes encode the human interferons IFN-
beta and IFN-gamma while human IFN-alpha constitutes a family of at least 23
different genes (Tanaka and Taniguchi, 1992).
Although there are some indications of constitutive interferon synthesis by
some cell types interferons are generally inducible proteins. Their synthesis