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THESIS

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This is to certify that the

thesis entitled

Experimental Bovine Viral Diarrhea
Virus Infection in Swine

presented by

Paul Harold Walz

has been accepted towards fulﬁllment
of the requirements for

M.S. degree in L.A.C.SJ

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Major professor

 

Date Alex 9. 1997

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MSU to An Afﬁrmative Action/Ewe! Opportunity lnetltuton
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EXPERIMENTAL BOVINE VIRAL DIARRHEA VIRUS INFECTION IN SWINE
By

Paul Harold Walz

A THESIS

Submitted to
Michigan State University
in partial fulﬁllment of the requirements
for the degree of

MASTER OF SCIENCE

Department of Large Animal Clinical Sciences

1997

ABSTRACT
EXPERIMENTAL BOVINE VIRAL DIARRHEA VIRUS INFECTION IN SWINE
By

Paul Harold Walz

The objective of this study was to determine if swine may serve as an alternative
animal model of BVDV transplacental infection. Preliminary in vitro and in vivo studies
with type I and type II BVDV were performed, and a type I isolate was selected for
further study in pregnant gilts. Eight pregnant gilts were divided into two groups of four
pigs each in an isolation facility. The control group was intranasally sham-inoculated;
and the principal group was intranasally inoculated with 107 TCIDso with the type I
BVDV isolate on day 65 of gestation. Clinical parameters were measured daily, and
serum and buffy coat samples were tested throughout the study period for BVDV and
antibody to BVDV. On day 110 of gestation, a caesarean section was performed. Serum
was obtained for virus isolation and antibody determination from all piglets, and all
experimental animals were euthanized. Selected tissues were collected for virus isolation
and histopathology. Bovine Viral diarrhea virus was isolated between day 5 and 7 after
infection and seroconversion was demonstrated in the gilts from the infected group.
Bovine viral diarrhea Virus was not isolated from serum samples from any fetuses from
the infected gilts, and BVDV was isolated from the spleen of one fetus from the infected
group. Viremia was estabished in the pregnant gilts, however transplacental infection at

day 65 of gestation in the pig was not consistently demonstrated.

It gives me great pleasure to dedicate this work
to Lloyd and Marion Walz,
my father and my mother,
for their unfailing love, support, and encouragement.

iii

ACKNOWLEDGMENTS

I would like to extend my thanks and gratitude to my graduate committee members,
Drs. John C. Baker, Roger K. Maes, Thomas P. Mullaney, David J. Sprecher, and John
Kaneene for their guidance and assistance. I would especially like to extend my sincere
and heartfelt gratitude to Dr. John C. Baker, my mentor and major professor, for his
guidance, friendship, and assistance above and beyond the work entailed within this
project.

I would also like to thank Dan Taylor, Lori Kunze, Cunquin Han, and Cheri Benson in
the Virology section of the Animal Health Diagnostic Laboratory for their assistance in
my training in the laboratory methods during the project.

I would like to thank Sherri Lenneman, Victoria Hoelzer-Maddox, and Dr. Lana
Kaiser for their assistance in preparing the manuscript.

Funding for this project was provided by a grant through the Agriculture Industry

Initiative and the Michigan Animal Experiment Station.

iv

TABLE OF CONTENTS

LIST OF TABLES ........................................................................................................... viii
INTRODUCTION ................................................................................................................ 1
LITERATURE REVIEW ..................................................................................................... 4
History of the Virus ........................................................................................................ 4
The Virus ....................................................................................................................... 6
Taxonomy ................................................................................................................. 6
Morphology .............................................................................................................. 6
Molecular biology .................................................................................................... 7
Biolype ..................................................................................................................... 8
Antigenic diversity ................................................................................................... 8
Genotype .................................................................................................................. 9
Clinical Manifestations of Bovine Viral Diarrhea Virus Infection in Cattle ............... 10
BVDV infection in immunocompetent, nonpregnant cattle .................................... 10

B VDVinfection in immunocompetent, pregnant cattle (fetal infection) ................ 14

B VDV infection in immunotolerant cattle .............................................................. l6
Ruminant Pestivirus Infection in Swine ...................................................................... 19
Relationship between ruminant pestiviruses and HC V .......................................... 19
Border Disease Virus Infections in Swine ................................................................... 21
Natural infections .................................................................................................. 21
Experimental infections ......................................................................................... 2 1
Clinical manifestations .......................................................................................... 22
Pathologic ﬁndings ................................................................................................ 22
Bovine Viral Diarrhea Virus Infections in Swine ........................................................ 23
Natural infections .................................................................................................. 23
Experimental infections ......................................................................................... 25
Clinical manifestations .......................................................................................... 28
Pathologicfindings ................................................................................................ 30
Diagnosis ............................................................................................................... 3 l
Porcine Fetal Immunology ........................................................................................... 31
Comparison of bovine and porcine placentation ................................................... 31
Development of humoral immune system ............................................................... 32
Specific immunotolerance induced by other viruses .............................................. 32
LITERATURE REVIEW - SUMMARY ........................................................................... 34

HYPOTHESES .................................................................................................................. 36

MATERIALS AND METHODS ....................................................................................... 37
Experimental Animals ................................................................................................. 37
Viral Isolates ................................................................................................................ 38
Cell Culture .................................................................................................................. 38
Determination of Viral Genotype ................................................................................. 39
Detection of BVDV Antigen by the Immunoperoxidase Monolayer Assay ................ 40
Preparation of Viral and Sham Inocula ........................................................................ 41
Determination of Viral Titer ........................................................................................ 42
Inoculation of Pigs ....................................................................................................... 43
Virus Isolation .............................................................................................................. 43
Serologic Testing ......................................................................................................... 45
Pathologic Examination ............................................................................................... 45
Experimental Design: In Vitro Selection of Isolates ................................................... 46
Experimental Design: In Vivo Selection of Isolates .................................................... 46
Experimental Design: Type I and Type II BVDV Dose Titration Study .................... 47
Experimental Design: Type I BVDV Infection in Pregnant Gilts ............................... 48
Statistical Analysis ....................................................................................................... 50

RESULTS .......................................................................................................................... 51
In Vitro Selection of Isolates ........................................................................................ 51
In Vivo Selection of Isolates ......................................................................................... 51

Clinical parameters ............................................................................................... 5 l
Serology ................................................................................................................. 52
Virus isolation ........................................................................................................ 52
Pathologic examination ......................................................................................... 52
Type I and Type II Dose Titration Study ..................................................................... 52
Clinical parameters ............................................................................................... 53
Serology ................................................................................................................. 53
Virus isolation ........................................................................................................ 53
Pathologic examination ......................................................................................... 54
Type I BVDV Infection in Pregnant Gilts .................................................................... 54
Clinical parameters: gilts ..................................................................................... 55
Serology: gilts ....................................................................................................... 55
Virus isolation: gilts .............................................................................................. 55
Pathologic examination: gilts ............................................................................... 56
Serology: fetuses ................................................................................................... 56
Virus isolation: fetuses .......................................................................................... 56
Pathologic examination: fetuses ........................................................................... 56
Conclusions ............................................................................................................ 56
DISCUSSION .................................................................................................................... 62

vi

SUMMARY AND CONCLUSIONS ................................................................................ 71

APPENDIX A .................................................................................................................... 73
APPENDIX B .................................................................................................................... 74
APPENDIX C .................................................................................................................... 76
LIST OF REFERENCES ................................................................................................... 78

vii

TABLE 1.

TABLE 2.

TABLE 3.

TABLE 4.

TABLE 5.

LIST OF TABLES

Characteristics of BVDV isolates used for in vitro study ............................... 57
Virus isolation results for in vivo selection of the BVDV isolates .................. 58
Virus isolation results for serum and buffy coat

samples from feeder age swine in type I and type II

dose titration study .......................................................................................... 59
Virus isolation results for tissue specimens from

feeder age swine in type I and type 11 close titration

study ................................................................................................................ 6O

Reciprocal antibody titers in pregnant gilts following
infection with type I BVDV on day 65 of gestation ........................................ 61

viii

INTRODUCTION

The disease of cattle that would become known as bovine viral diarrhea was first
described in 1946 (Olafson, 1946; Childs, 1946). The virus, which would be redundantly
named bovine Viral diarrhea virus (BVDV), was discovered seven years later. The 50
years following the first report brought into View other clinicopathologic entities
associated with the virus such as persistent infection, mucosal disease,
immunosuppression, hemorrhagic syndrome, and reproductive inefﬁciency (Baker, 1995).
In the past 15 years, there has been a vast increase in the knowledge of the molecular
biology of this virus. The pathogenesis of mucosal disease has been determined
(Brownlie, 1984; Bolin, 1985), and the complete genomes for the BVDV isolates
designated NADL, SD-l, and 031053 have been sequenced (Collett, 1988; Deng, 1992;
DeMoerlooze, 1993). The transmission of the virus by persistently infected carriers and
intrauterine infection has been elucidated with the addition of epidemiologic
investigations in the study of BVDV (McClurkin, 1984; Meyling, 1990).

In spite of the advances that have been made in our understanding of this complex
virus and its associated diseases, BVDV continues to be an important pathogen of cattle
and control has not been achieved. Economic losses created by infection with BVDV are

substantial. Estimated losses in Denmark amount to 17 million US. dollars per year

2

(Houe, 1993). In addition, recent outbreaks of severe clinical disease in
immunocompetent cattle have further demonstrated that control has not been attained
(Carman, 1994; David, 1994; Drake, 1994; Pellerin, 1994). These recent outbreaks were
characterized by severe, often fatal disease, and have been associated with the type II
genotype of BVDV (Ridpath, 1994).

Methods to control BVDV infection center around vaccination of susceptible animals
and the identiﬁcation and elimination of persistently infected carrier animals. Persistently
infected carrier animals are the major source of virus spread within and among cattle
herds. Persistently infected carriers can only be created by in utero infection of
susceptible, pregnant cattle with a noncytopathic isolate of BVDV prior to the
development of fetal immunocompetence (McClurkin, 1984). Therefore, a logical
solution to the problem of persistent infection is to prevent occurrence of transplacental
infection. Prevention may be achieved by vaccination of cattle prior to breeding in order
to elicit immunity against BVDV exposure and thus prevent transplacental infection.
Identiﬁcation and elimination of persistently infected carrier animals within the herd that
are a source of exposure to pregnant cattle is another method of preventing transplacental
infection. Strict biosecurity measures to prevent further introduction of persistently
infected carriers into herds is considered an additional preventive measure. Biosecurity
measures include isolation and testing of new additions to the herd (Ames, 1990).

More than 140 federally licensed vaccines for BVDV are available in the United States
(Bolin, 1995). The majority of these vaccines were licensed prior to a complete

understanding of the pathogenesis of BVDV infection. In addition, these vaccines have

3

not been tested to determine if they are efﬁcacious in preventing transplacental infection
with BVDV. Prevention of cell-associated Viremia, with subsequent transplacental
infection, is the realistic measure of efﬁcacy for vaccines. In Europe, a commercially
available vaccine was recently determined to be efﬁcacious in preventing transplacental
infection with BVDV after challenge exposure (Brownlie, 1995). However, experiments
utilizing pregnant cattle in vaccine efﬁcacy trials are difﬁcult and expensive to perform.
The purpose of the present study was to develop an alternative animal model of BVDV
transplacental infection utilizing swine in order to develop a means to economically and
efﬁciently test commercially available vaccines, and new-generation vaccines against

BVDV infection.

LITERATURE REVIEW

History of the Virus

It has been more than 50 years since the discovery of an acute, infectious, and
transmissible disease of cattle that would later become known as bovine viral diarrhea
(Olafson, 1946). The initial outbreak involved 6 dairy herds in New York state with
clinical signs of gastroenteritis with severe diarrhea. Other clinical characteristics of
affected cattle included elevated body temperatures, salivation, nasal discharge,
depression, anorexia, dehydration, and abortions during the outbreak and extending three
months beyond. Leukopenia was a characteristic laboratory ﬁnding. Postmortem
examination revealed ulceration of the mucous membranes of the oropharynx and
esophagus, and hemorrhages in the gastrointestinal tract, subcutaneous tissues,
epicardium, and vaginal mucosa. Experimental reproduction of this novel disease was
also reported. Oral inoculation with fecal material, subcutaneous injection of blood
collected from febrile cases, and subcutaneous injection of splenic emulsions from
deceased cases all proved to be infective to susceptible cattle (Olafson, 1946). A very

9

similar disease syndrome, which was termed “X disease,’ was reported from Canada
during the same year (Childs, 1946). This report included bloody diarrhea as a clinical

Sign, in addition to those mentioned previously. Acute and subacute forms of the disease

were also described in cattle affected by “X” disease.

5

A viral etiology for this new transmissible disease of cattle was not demonstrated until
seven years after the initial reports (Baker, 1954). Early descriptions of the virus
indicated that two forms existed, one causing cell vacuolation and death and the other
producing no obvious pathology in cell monolayers (Lee, 1957; Gillespie, 1960). This
formed the ﬁrst subclassiﬁcation of BVDV, cytopathic and noncytopathic BVDV, which
later was designated as the biotype Of the virus.

The Virus subsequently became associated with a sporadically occun'ing enteric
disease characterized by low morbidity and high mortality that was termed mucosal
disease (Ramsey, 1953). Bovine viral diarrhea virus isolates associated with mucosal
disease were found to belong to the cytopathic biotype (Gillespie, 1961). This would set
the stage for the most widely studied aspect of BVDV infection, the pathogenesis of
mucosal disease. Nearly 35 years passed before the pathogenesis of mucosal disease was
determined (Brownlie, 1984; Bolin, 1985). Cattle persistently infected with
noncytopathic BVDV develop mucosal disease after superinfection with cytopathic
BVDV (discussed later). The persistent infection established in cattle was determined to
be a result of transplacental infection with the noncytopathic biotype of BVDV during the
ﬁrst 100 days of gestation (Liess, 1984; McClurkin, 1984).

The past 20 years have brought about a rapid increase in the knowledge of the
molecular biology and epidemiology of BVDV. Viral isolates have been sequenced
(Collett, 1988; Deng, 1992; DeMoerlooze, 1993) and the organization of the viral
genome has been determined (Collett, 1992). The role of persistently infected carriers in

the transmission of the virus has been deﬁned (Houe, 1995).

The Virus

Taxonomy. Bovine viral diarrhea virus is the prototype Virus of the genus Pestivirus
within the family Flaviviridae (Wengler, 1991). The genus Pestivirus is comprised of
three viruses of veterinary importance, which were grouped together based upon serologic
ﬁndings suggesting that they were related (Darbyshire, 1960; Plant, 1973). The three
viruses are BVDV, hog cholera virus (which is also known as classical swine fever virus),
and border disease virus of Sheep and goats. In 1978, BVDV and the other pestiviruses
received status as a genus and were classiﬁed together within the family Togaviridae
(Porterﬁeld, 1978). The pestiviruses, however, were considered an oddity in that family
because they do not require arthropods for transmission. This unusual feature, combined
with modern molecular biological techniques that allowed for nucleotide sequence
determinations, gene expression strategies, and the determination of genome
organization, was the stimulus for reclassiﬁcation within the family Flaviviridae (Collett,
1988). In 1991, the pestiviruses received ofﬁcial status as a member genus of the family
Flaviviridae (Wengler, I991). The family Flaviviridae contains three genera: Pestivirus,
Flavivirus, and an unnamed genus that contains human hepatitis C virus. Yellow fever
virus is the prototype Virus for the genus Flavivirus, and human hepatitis C virus is
considered the prototype virus for the unnamed genus.

Morphology. The morphology of BVDV has been variably described. This is due to
inherent characteristics of the virus that make puriﬁcation and visualization of virions
difﬁcult. The density of the virion is similar to that of subcellular components and the

fragility of mature virions makes physical separation difﬁcult (Ohmann, 1990). Early

7

reports described enveloped particles of variable size from 35 to 100 nm (Hermodson,
1962; Hafez, 1968; Maess, 1970; Horzinek, 1973; Stott, 1974; Scott, 1977). The ﬁndings
of more recent reports have indicated that BVDV is an enveloped virus measuring
approximately 40-60 nm in diameter (Chu, 1984; Ward, 1984). Underlying the envelope
is the nucleocapsid, which measures approximately 27-30 nm in diameter and possesses
icosahedral symmetry (Chasey, 1981; Ohmann, 1982, Ohmann, 1988).

Molecular biology. Recent advances have been made in the understanding of the
molecular biology of BVDV and the other pestiviruses and are contained in several
review articles (Donis, 1995; Meyers, 1996). The Viral genome of BVDV is composed of
a single stranded RNA molecule of positive sense polarity. The complete genome has
been sequenced for multiple isolates and is approximately 12.5 kilobases in length
(Collett, 1988; Deng, 1992; DeMoerlooze, 1993). The BVDV genome can be subdivided
into three regions, which consist of a large open reading frame that is ﬂanked by a 5’
untranslated region and a 3’ untranslated region. The 5’ untranslated region is
approximately 385 nucleotides in length and lacks the S’methyl guanosine cap that is
typical for eucaryotic and some Viral RNA molecules (Brock, 1992). The 5’ untranslated
region has a complex secondary structure similar to picornaviruses. Because of the
complex secondary structure, this region has been determined to be an internal ribosomal
entry Site for the initiation of translation (Pellerin, 1994; Poole, 1996). The open reading
frame is approximately 11,700 nucleotides in length, which correlates into 3898 codons
(Collett, 1988). The open reading frame is translated into a polyprotein that is co- and

posttranslationally cleaved. A total of 13 polypeptides have been found in BVDV-

8

infected cells, and can be divided into structural and nonstructural proteins (Akkina,
1991; Collett, 1988; Donis, 1987; Donis, 1991). Three structural proteins (E0, E1, E2)
are encoded by the 5’ end of the open reading frame, while the remaining nonstructural
proteins are encoded by the 3’ end. The structural glycoprotein E2, which was previously
titled gp53, is the target for neutralizing antibody production by the humoral immune
system (Donis, 1988; Xue, 1990). Genomic sequence analysis has determined
hypervariable regions within the RNA encoding for E2, and this variability is responsible
for antigenic diversity (Bolin, 1991; Donis, 1991; Paton, 1992).

Biotype. Bovine Viral diarrhea virus exists as one of two possible biotypes,
noncytopathic or cytopathic. This classiﬁcation is based upon the effect of viral infection
of cell monolayers in tissue culture. Cytopathic BVDV infection results in cell
vacuolation and death with 24 to 48 hours following inoculation. In contrast,
noncytopathic BVDV infection induces no overt effects on either the appearance or
survival of cells in culture (Bolin, 1990; Deregt, 1995). Although biotype does not imply
virulence in the host, the classiﬁcation of biotype does have clinical implications that
extend beyond cell culture. The noncytopathic biotype is considered the predominant
biotype in nature, while the cytopathic biotype is rare (Bolin, 1990). Both biotypes have
been reported to cause acute infection, but only noncytopathic isolates are involved in the
establishment of persistent infection (Baker, 1987; Deregt, I995). The pathogenesis of
mucosal disease has been determined and involves both biotypes (discussed later).

Antigenic diversity. Although BVDV isolates appear to represent a single serotype

(Dubovi, 1992), considerable antigenic diversity exists among isolates of BVDV based

9

upon cross neutralization studies utilizing polyclonal and/or monoclonal antibodies
(Castrucci, 1975; Bolin, 1988; Wenswoort, 1989). Mutational events are considered to
be the origin of antigenic diversity (Paton, I995). Viruses composed of RNA are highly
susceptible to mutations in the genetic code because of poor proofreading during
replication (Dubovi, 1992). Some mutational events occur in the viral genome encoding
for the structural protein E2, and three hypervariable regions have been identiﬁed. (Bolin,
1991; Donis, 1991, Paton, 1992). The importance of mutations in the structural
glycoprotein E2 lies in the fact that this protein is the target for neutralizing antibody
production by the humoral immune system. Antigenic diversity may have important
consequences for vaccination, potentially limiting the protective spectrum offered by
vaccines carrying only one isolate. Suspected vaccine failure with BVDV has been
reported as a result of antigenic variation (Harkness, 1987; Meyling, 1987; Bolin, 1991).
Genotype. Bovine viral diarrhea virus isolates may be subdivided based upon
differences in the nucleotide sequence. Speciﬁc differences in the viral nucleic acid
sequences are the basis for genotyping. Several different procedures are used in
genotyping isolates of BVDV. These include analysis of nucleic acid sequences from the
5’ UTR and E2 segments of the Viral genome (Ridpath, 1994; Pellerin, 1994), and
polymerase chain reaction ampliﬁcation of the 5’ UTR followed by restriction
endonuclease analysis of the ampliﬁcation product (Harpin, 1995). Two distinct
genotypes of BVDV exist, which are classiﬁed as type I and type II (Ridpath, 1994).
Analysis and comparison of nucleic acid sequences from type I and type II genotypes has

revealed greater than 30% dissimilarity (Ridpath, 1995). This dissimilarity is

10

concentrated in the regions of the viral genome encoding for E2 and the 5’ UTR. The
differences in nucleic acid sequences between type I and type II BVDV have been
reported to be as great as the differences in nucleic acid sequences between type I BVDV

and hog cholera virus (Bolin, 1996).

Clinical Manifestations of BVDV Infection in Cattle

Bovine viral diarrhea virus infection in cattle may result in a wide spectrum of clinical
manifestations ranging from subclinical to fatal disease. Clinical manifestations are
dependent upon the interplay of host factors, environmental stress levels, and viral
factors. Host factors inﬂuencing the outcome of viral infection include immune status,
pregnancy status, gestational age of the fetus at the time of infection, and whether the host
is immunotolerant or immunocompetent. Viral factors include biotypic variation,
genotypic variation, and antigenic diversity.

The clinical manifestations of BVDV infection in cattle have been reviewed (Baker,
1995), and can be subdivided into three categories: BVDV infection in immuno-
competent, nonpregnant cattle; BVDV infection in immunocompetent, pregnant cattle
(fetal infection); and BVDV infection in immunotolerant cattle.

B VDV infection in immunocompetent, nonpregnant cattle. Multiple clinical forms
of BVDV infection in immunocompetent cattle exist, including subclinical, clinical
(BVD), severe BVD, and a hemorrhagic syndrome. The majority Of BVDV infections in
immunocompetent and seronegative cattle are subclinical (Ames, 1986). If observed

closely, cattle undergoing a subclinical infection may develop pyrexia, a mild leukopenia,

11
and a decrease in milk production (Moerman, 1994). The source of BVDV is usually a

persistently infected animal, and exposed cattle develop BVDV-speciﬁc neutralizing
antibody (Moerman, 1994).

Bovine Viral diarrhea (BVD) is the term used to describe the clinical form of BVDV
infection in immunocompetent cattle. Clinical signs of BVD include diarrhea,
depression, oculonasal discharge, anorexia, decreased milk production, oral ulcerations,
pyrexia, and mild leukopenia (Brownlie, 1987; Perdrizet, 1987). BVDV can be isolated
from serum and/or peripheral blood leukocytes from 4 to 5 days after infection until 15
days after infection (Brownlie, 1987). In addition, BVD can be diagnosed by the
presence of neutralizing antibody to BVDV 2 to 4 weeks after infection (Moerman,
1994)

Outbreaks of severe, peracute BVDV infections in immunocompetent cattle have
recently been reported in the United States, Canada, and the United Kingdom involving
beef, dairy, and veal operations (Pellerin, 1994; Drake, 1994; Carman, 1994; David,
1994). In Quebec, mortality due to BVDV in veal operations was estimated at 25% of
143,000 calves for 1993 (Pellerin, 1994). The outbreak in Ontario involved 150 dairy,
600 beef, and 100 veal herds with mortality of up to 50% in some herds (Carman, 1994).
Analysis of BVDV isolates from outbreaks in Canada and the United States has
conﬁrmed that they were of the type II genotype of BVDV. Clinical manifestations of
severe, peracute BVD include diarrhea, pyrexia, decreased milk production, and oral
ulcerations in some cases (Drake, 1994; Carman, 1994). In addition, concurrent diseases

such as pneumonia and abortion were frequently reported in herds experiencing outbreaks

l2
(Drake, 1994; David, 1994). Prior to these described type II BVDV outbreaks, there was

a tendency to depreciate the importance of BVDV infection in immunocompetent cattle.
The reports Of these severe BVDV infections clearly demonstrate that some isolates of
BVDV can cause severe life-threatening disease in immunocompetent cattle.

Infection of nonimmune, immunocompetent cattle with some noncytopathic isolates of
type II BVDV has been associated with a hemorrhagic disorder characterized by
thrombocytopenia, hemorrhage, leukopenia, pyrexia, diarrhea, and death (Bolin, 1992;
Rebhun, 1989; Corapi, 1989; Corapi, 1990; Bezek, 1994). Thrombocytopenia associated
with BVDV has been reported under natural conditions in adult cattle and veal calves
(Rebhun, 1989; Corapi, 1990). Clinical signs of BVDV-induced thrombocytopenia
include bloody diarrhea, epistaxis, petechial and ecchymotic hemorrhages on mucous
membranes, and prolonged bleeding from injection sites (Rebhun, I989). BVDV-
induced thrombocytopenia has also been experimentally reproduced (Corapi, 1989;
Corapi, 1990; Bolin, I992; Bezek, 1994). Platelet decline may begin as early as day 3
aﬁer inoculation with maximum platelet count depression occurring between day 9 and
17 after inoculation (Bolin, 1992; Corapi, I989). Bovine viral diarrhea virus association
with peripheral platelets has been reported to occur beginning as early as day 4 after
inoculation as indicated by the isolation of BVDV from puriﬁed platelet preparations
(Bolin, 1992; Walz, 1996) or by the demonstration of BVDV antigen in association with
platelet preparations by immunoﬂuorescent antibody testing (Corapi, 1990; Bezek, 1994).
In addition, the presence of BVDV antigen following experimental type II BVDV

challenge has been detected in bone marrow megakaryocytes by immunohistochemistry

13
and by immunoﬂuorescent antibody staining (Marshall, 1996; Corapi, 1990; Walz, 1996).

The mechanism by which some isolates of type II BVDV induce thrombocytopenia has
yet to be deﬁnitively elucidated.

Immunosuppression is another feature of BVDV infection in immunocompetent cattle.
As a result, BVDV has the potential to enhance disease by other pathogens or make the
host more susceptible to opportunistic infections. A transient leukopenia may occur
during BVDV infection, and this leukopenia is characterized by decreased absolute
numbers of B and T lymphocytes (Bolin, 1985). Bovine Viral diarrhea virus has been
reported to enhance the pathogenicity of bovine herpesvirus-l (BHV -1), actinomycosis,
papular stomatitis, enteritis caused by Salmonella spp., colibacillosis, metritis, and
mastitis (Ames, 1987; Bohac, 1980; Greig, 1981; Potgetier, 1977). Experimental
evidence of the immunosuppressive role of BVDV in respiratory disease was
demonstrated in calves by showing that a BVDV infection established prior to challenge
with BHV-l resulted in dissemination of BHV-l infection in the lungs as compared to
controls just infected with BHV-l (Potgetier, 1984). In addition, synergism in respiratory
disease between BVDV and Pasteurella haemolytica has been experimentally
demonstrated (Potgetier, 1984).

Venereal infections have also been identiﬁed following BVDV infection in
nonimmune, immunocompetent cattle. Decreased conception rates as a result of
fertilization failure have been reported in association with BVDV infection in
nonimmune, immunocompetent cattle (Houe, 1993; McGowan, 1993). Bovine viral

diarrhea Virus infection in seronegative, immunocompetent bulls often results in

14
infertility and poor semen quality (Harkness, 1987; Paton, 1989). In addition, semen

quality may remain inferior for as long as 76 days following acute infection with BVDV
(Paton, 1989).

BVDVinfection in'immunocompetent, pregnant cattle (fetal infection). The clinical
signs of disease associated with BVDV infection in the dam are similar to those
encountered for acute infection in nonpregnant cattle. Since 70 to 90% of infections are
subclinical (Ames, 1990), it is common that clinical signs of BVDV infection in the dam
are absent. The lack of clinical signs does not exclude the possibility of transplacental
infection. In fact, transplacental infection with BVDV is a reliable event following
infection of pregnant cattle, with estimates of 100% efﬁciency (Duffell, 1985). The
outcome of transplacental infection is fetal infection. Clinical manifestations following
fetal infection are primarily dependent upon the gestational age of the fetus at the time
BVDV transplacental infection occurs. Transplacental fetal infection may result in early
embryonic death, abortion, mummiﬁcation, congenital defects, stillbirths, normal calves
born seropositive to BVDV, and the birth of calves immunotolerant to, and persistently
infected with BVDV.

Reduced pregnancy rates as a result of early embyonic death have been reported as a
result of BVDV infection in cattle after insemination (McGowan, 1993). When
introduced into the genital tract at the time of insemination, BVDV has been reported to
persist in the uterine environment up to 53 days (Archbald, 1977). In addition, early

stages of pregnancy are very sensitive to BVDV infection. Bovine Viral diarrhea virus

15

infection between day 29 and 41 of gestation resulted in fetal death and resorption in four
of four seronegative cows (Carlsson, 1989).

Abortion associated with BVDV usually occurs as a result of infection between 50 and
100 days of gestation (Cassaro, 1971; Kahrs, 1973), but it has also been reported to
happen during the last trimester (Bolin, 1990). Expulsion of the fetus may occur from
several days to several months after fetal infection and fetal death (Kahrs, 1973).
Abortions have been a feature of the recent outbreaks of severe BVD associated with the
type II genotype (Drake, 1994; Carman, 1994). Late-term abortions, although
uncommon, have been reported (Harkness, 1988; Lohr, 1983), and it has been speculated
that this may represent the abortion of a persistently infected calf (Harkness, 1988).

Congenital defects may result if transplacental infection of the fetus takes place
between approximately 100 and 150 days of gestation (Duffel, 1985). This gestational
age range corresponds to the stage of organogenesis of the nervous system. Potential
congenital defects associated with BVDV infection include retinal atrophy, cataracts,
microencephalopathy, hydrocephalus, and cerebellar hypoplasia (Baker, 1995).

Calves infected with BVDV following the development of the immunocompetence
can appear normal at birth, but they will be seropositive to BVDV (Duffel, 1985).

Infection with noncytopathic strains Of BVDV before the development of
immunocompetence by the fetus may result in the birth of calves that are immunotolerant
and persistently infected with BVDV (McClurkin, 1984). The development of bovine

fetal immunocompetence is generally thought to occur between 100 and 125 days of

16

gestation. Cattle that are persistently infected with BVDV are at risk for development of
mucosal disease.

BVDV infection in immunotolerant cattle. Cattle that are immunotolerant to, and
persistently infected with BVDV are viremic and continuosly shed the virus (Barber,
1985). Persistently infected cattle are the primary source of virus for transmission to
susceptible cattle. In addition, persistently infected breeding females will produce
persistently infected Offspring, thus providing a means to perpetuate the disease within a
herd (McClurkin, 1979; Straver, 1983). Persistently infected cattle have decreased
survival rates, as evidenced by reports documenting a greater than 50% mortality rate in
the ﬁrst year of life (Duffell, 1985; Houe, 1993). This mortality may be due in part to the
fact that persistently infected animals may have an impaired immune response, which
makes them more susceptible to opportunistic pathogens (Roberts, 1988). This is
supported by data documenting that persistently infected cattle have lower percentages of
functional B cells (Johnson, 1973; Muscoplat, 1973) However, the fact remains that
some persistently infected animals remain healthy and can exist in a herd undetected for
prolonged periods of time. A study evaluating the fate of 34 persistently infected animals
reported that 44% of these calves remained clinically normal until slaughter (Houe,
1993)

Persistently infected cattle are at risk of developing mucosal disease. Mucosal disease
occurs when cattle that are persistently infected with a noncytopathic isolate of BVDV
become “superinfected” with a cytopathic isolate of BVDV (Brownlie, 1984; Bolin,

1985). There are multiple clinical forms of mucosal disease, with the variations being

17

due to the antigenic relationship between the persistently infecting noncytopathic isolate
and the superinfecting cytopathic isolate (Brownlie, 1991). At one end of the spectrum is
acute, fatal mucosal disease, in which the superinfecting cytopathic isolate is homologous
to the persistently infecting noncytopathic isolate. The other end of the spectrum is
mucosal disease with recovery, in which the superinfecting cytopathic isolate is
heterologous to the persistently infecting noncytopathic isolate. The animal mounts an
immune response to the heterologous superinfecting cytopathic isolate and recovers. The
multiple clinical forms Of mucosal disease have been reviewed and can be divided into
acute, fatal mucosal disease; chronic mucosal disease; chronic mucosal disease with
recovery; and delayed onset mucosal disease (Baker, 1995 ; Bolin, 1995).

Acute fatal mucosal disease occurs when the superinfecting cytopathic isolate shares
close antigenic homology to the persistently infecting noncytopathic isolate. Monoclonal
antibody analysis of noncytOpathic-cytopathic pairs of BVDV from outbreaks of acute
mucosal disease has been used to assess the antigenic similarity between them (Corapi,
1988; Howard, 1987). The cytopathic isolate may arise de novo by a mutational event
from the noncytopathic isolate in the persistently infected animal (Meyers, 1991; Meyers,
1992; Qi, 1992; Tautz, 1994). The mutational event affects the viral biotype but does not
affect the viral antigenicity. The clinical ﬁndings associated with acute mucosal disease
include pyrexia, depression, anorexia, decreased milk production, profuse diarrhea,
ptyalism, mucopurulent oculonasal discharge, and oral erosions (Baker, 1990; Brownlie,

1985; Nagele, 1984; Radostits, 1988). Oral erosions are present on the lips, gingival

l8

margins, tongue, dental pad, and the hard palate. Acute mucosal disease is considered to
be 100% fatal.

Chronic mucosal disease and mucosal disease with recovery are forms of mucosal
disease in which the superinfecting cytopathic isolate is heterologous to the persistently
infecting noncytopathic isolate. The likely source of the superinfecting cytopathic isolate
is external, rather than a mutational event in the persistently infecting noncytopathic
isolate. The clinical manifestations of chronic mucosal disease are anorexia, weight loss,
diarrhea, chronic bloat, alopecia, erosive lesions on the mouth and skin, and lameness
(Baker, 1995). The lameness may develop as a result of laminitis, interdigital necrosis
secondary to erosive lesions, and hoof deformities. Mucosal disease with recovery occurs
when the persistently infected animal mounts an immune response to the superinfecting,
heterologous BVDV isolate, and clears the superinfection (Edwards, 1991). The animal,
however, remains persistently infected with the noncytopathic biotype.

Delayed onset mucosal disease occurs when an antigenically heterologous cytopathic
isolate infects a persistently infected animal (Bolin, 1995). The origin of the
superinfecting cytopathic isolate is external, such as a modiﬁed-live vaccine (Ridpath,
1995). Neutralizing antibody is produced by the perisistently infected animal and the
superinfecting cytopathic isolate is destroyed. However, RNA from the superinfecting
cytopathic isolate may recombine with RNA from the resident, persistently infecting
noncytopathic isolate. This, in effect, creates a new cytopathic isolate that is antigenically
identical to the persistently infecting noncytopathic isolate (Ridpath, 1995). Clinical

signs of acute mucosal disease follow. Mucosal disease does not occur within 2 to 4

19

weeks after the initial infection with the cytopathic isolate, but occurs several weeks to

months later.

Ruminant Pestivirus Infections in Swine

The discovery in 1960 of a close antigenic relationship between bovine viral diarrhea
virus (BVDV) and hog cholera virus (HCV) (Darbyshire, 1960) prompted investigations
of BVDV infection in swine. Early studies focused on the use of intentional BVDV
infection as a means to immunize pigs against subsequent HCV infection (Beckenhauer,
1961; Atkinson, 1962; Simonyi, 1967; Baker, 1969). With the implementation of the
HCV eradication program in the United States in 1962, it became clear that BVDV
infection in pigs could potentially interfere with the screening of swine herds for HCV.
With the achievement of eradication in the United States in 1976, attention has shifted to
surveillance. In the United States and in other countries declared free of HCV, it has
become imperative to differentiate BVDV infection in pigs from HCV infection in order
to maintain the HCV-free status. At the present time, BVDV infection in swine has
become important, as several Scandinavian countries begin the process of eradicating
BVDV. It is important to assess the role of swine as a potential resevoir of BVDV and an
obstacle in BVDV eradication programs.

Relationship between ruminant pestiviruses and HC V. Bovine viral diarrhea virus is
a positive-sense single-stranded RNA virus and is the prototype virus of the genus
pestivirus within the family F laviviridae. The genus Pestivirus is comprised of three

viruses of veterinary importance, which were grouped together based upon serologic

 

20
ﬁndings, suggesting that they were related (Darbyshire, 1962; Plant, 1973). The three

viruses are BVDV, hog cholera virus (HCV) which is also known as classical swine fever
virus, and border disease virus (BDV) of sheep and goats. The antigenic relationship
between BVDV and HCV was ﬁrst reported based upon a line of identity between BVDV
and HCV in agar gel irnmunodiffusion tests (Darbyshire, 1960; Darbyshire, 1962). This
relationship was further supported by cross-neutralization studies (Kumagai, 1962), cross
reactivity in immunoﬂuorescence tests between BVDV and HCV using the same
antibody-ﬂuorescein conjugates (Mengeling, 1963), agar double diffusion studies
utilizing soluble antigen of BVDV and HCV (Gutekunst, 1963), and complement ﬁxation
tests (Gutekunst, 1964). The relationship of BDV to BVDV and HCV was discovered
later (Hamilton, 1972; Osburn, 1973; Wenswoort, 1989). In addition, pestiviruses
possess the ability to replicate in cells from heterologous species (Roche, 1994; Moennig,
1990). Bovine Viral diarrhea virus, BDV, and HCV have been reported to replicate in
cells of porcine, bovine, and mine origin (Roche, 1994). With the potential that BVDV
or BDV infections in pigs could be confused with HCV in the eradication program, it
became important that differences between the three viruses be characterized.
Monoclonal antibodies were developed and genetic analysis of BVDV and HCV isolates
has been performed (Edwards, 1988; Wenswoort, 1989). Bovine Viral diarrhea virus and
BDV recovered from pigs can now be differentiated from HCV by the use of monoclonal
antibodies (Wenswoort, 1989; Zhou, 1989) and polymerase chain reaction ampliﬁcation

of genomic segments (Hertig, 1991; Katz, 1993; Harding, 1996).

21

Border Disease Virus Infection in Swine

Border disease was ﬁrst described in 1959 as a clinical disease affecting sheep
(Hughes, 1959). The name border disease reflects the fact that the disease was ﬁrst
described in the border counties between England and Wales. Synonyms of border
disease include hairy shaker disease or fuzzy-lamb syndrome because some border
disease lambs show signs of trembling and a hairy (as opposed to wool), rough coat. A
viral etiology was demonstrated in 1973 (Hamilton, 1973).

Natural infections. Natural infections of pigs with BDV have been reported (V annier,
1988; Roehe, 1992). The use of a pseudorabies vaccine that had been contaminated with
BDV was the source of one outbreak that also involved transplacental infections
(Vannier, 1988). The isolate recovered during this outbreak was determined to be BDV
based upon comparative serology with the highest neutralizing antibody titers being
present against BDV.

Experimental infections. Border disease Virus infection in pigs has been
experimentally reproduced (Wrathall, 1978; Vannier; 1988; Leforban, 1990; Chappuis,
1984). All studies on experimental BDV infection in pigs have been done in pregnant
swine in order to demonstrate transplacental infection. Three sows were infected by
intramuscular inoculation at 34 days of gestation with an emulsion of tissues recovered
from lambs infected with border disease Virus (Wrathall, 1978). Although BDV and
antibody to BDV were not recovered from fetuses, congenital defects such as cerebellar

hypoplasia were recognized. In another study, two pregnant sows were intramuscularly

 

22

inoculated between days 25 and 29 of gestation with the reference strain of BDV
designated as the Aveyron isolate (Leforban, 1990). Transplacental infection with BDV
occurred as evidenced by the birth of persistently infected piglets with BDV.
Experimental reproduction of the natural outbreak involving a contaminated pseudorabies
vaccine with BDV was performed by inoculating sows on day 25 of gestation by
intramuscular injection of the vaccine. Transplacental infection as evidenced by the birth
of persistently infected piglets was demonstrated in 2 of 2 sows infected (V annier, 1988).

Clinical manifestations. Natural infections of pigs with BDV have been associated
with repeat breeding, stillbirths, and mummiﬁed fetuses at birth (V annier, 1988). Most
cases of BDV infection in the dam are not apparent and the only evidence of infection is
the development of neutralizing antibodies to BDV following infection (Vannier, 1988;
Leforban, 1990). In the piglets born from sows naturally infected with BDV, eyelid
edema, locomotor disorders, diarrhea, and arthritis were reported (Vannier, 1988).
Mortality for the ﬁrst 2 days of life in litters of congenitally infected piglets varied ﬁ'om
30% to 70% (Vannier, 1988).

Following experimental infection of pregnant sows with BDV, clinical signs of disease
in the sows were not evident (Wrathall, 1978, Leforban, 1990). However, piglets born
from experimentally infected sows with BDV demonstrate eyelid edema, pyrexia,
diarrhea, and stunting, as well as an increased perinatal mortality (Leforban, 1990).

Pathologic ﬁndings. Lesions following BDV infection in pigs have been primarily
observed in the offspring of sows infected during gestation. Hemorrhagic lesions

involving lymph nodes and other lymphoid tissues have been reported in piglets that died

23
in the ﬁrst few days of life (Chappuis, 1984; Leforban, I990; Vannier, 1988). A

meningocoele in l piglet and cerebellar hypoplasia in 9 piglets out of a total of 19 piglets
was reported following experimental BDV infection in 3 pregnant sows at day 34 of

gestation (Wrathall, 1978).

Bovine Viral Diarrhea Virus Infection in Swine

Natural infections. In the cattle population, BVDV has a worldwide distribution, with
serum antibody prevalence in cattle ranging from 50 to 90% (Ernst, 1983). Bovine viral
diarrhea virus infection in swine has been reported to occur under natural conditions as
evidenced by the presence of speciﬁc antibodies to BVDV (Holm Jensen, 1985; Loken,
1991). Serological surveys have also conﬁrmed the worldwide distribution of BVDV in
swine. The ﬁrst report suggesting natural BVDV infection in pigs was from Australia
(Flynn, 1964). A serological test, the gel diffusion precipitin test, was employed in the
diagnosis of HCV in an outbreak in New South Wales. The validity of the test was
questioned when pigs were positive without clinical signs or pathologic lesions of HCV,
thus suggesting possible natural BVDV infection. Natural BVDV infection of pigs has
been conﬁmred by other serologic surveys. A study of breeding pigs in Germany
indicated that 42% of pigs possessed neutralizing antibodies against BVDV with
reciprocal titers ranging from 5 to 640 (Liess, 1976). A study conducted in the Republic
of Ireland showed that 27.8% of pigs in contact with cattle possessed neutralizing
antibodies to BVDV, as compared to 4% of pigs without contact (Lenihan, 1977).

Serological surveys performed on pigs in the United States have shown higher

24
neutralizing antibody titers to BVDV when compared to HCV, indicating natural

infection with BVDV (Femelius, 1973; Carbrey, 1976). A study performed in the
Netherlands indicated that a range of 15-20% of the pig population possessed antibodies
to BVDV (Terpstra, 1988). A serological survey performed on 2,996 Danish pigs
indicated that 6.4% possessed antibodies to BVDV (Holm Jensen, 1985).

The ﬁrst reported isolation of BVDV from naturally infected pigs came 12 years after
it was conﬁrmed that BVDV and HCV were serologically related (Femelius, 1973). The
source of BVDV in this case was suggested to be a calf that was in direct contact with
pigs. The virus was determined to be an isolate of BVDV based upon
immunoﬂuorescence and the production of bovine viral diarrhea and death in susceptible
calves. Close contact between cattle and pigs was also the focus of another report in
which natural infection of pigs with BVDV was documented (Paton, 1992). In this
particular report, transplacental infection was reported with the birth of persistently
viremic piglets. Other reports have documented natural infections of pigs with BVDV,
without direct contact between cattle and pigs (Carbrey, 1976; Stewart, 1971; Terpstra,
1988). Contaminated personnel and equipment was considered the source of infection of
pigs with BVDV on a farm in Indiana where intensive production methods were
employed and pigs and cattle were separated (Carbrey, 1976). The isolate of BVDV in
this particular case was originally considered to be an isolate of HCV. Only through the
experimental infection of calves with the presence of severe disease was it revealed that
the isolate obtained from this outbreak was an isolate of BVDV. The feeding of bovine

offal to pigs has also been a source of natural infection with BVDV (Carbrey, 1976;

25
Stewart, 1971). Other origins of BVDV infection in pigs include the feeding of BVDV-

contaminated whey and milk products to pigs (Terpstra, 1988) and the use of BVDV-
contarninated vaccines (Wenswoort, 1988). Modiﬁed-live vaccines for HCV
(Wenswoort, 198 8) that were contaminated with BVDV were responsible for outbreaks in
pig herds and resulted in transplacental infection.

Transplacental infection of swine with BVDV has been reported under natural
conditions, with the clinical signs in piglets closely resembling transplacental infections
with HCV (Terpstra, 1988). The sources of virus for infection of pigs were close contact
with persistently infected calves (Terpstra, 1988; Paton, 1992) or the use of vaccines
contaminated with BVDV (Wenswoort, 1988). The transplacental infections were
suspected by the presence of viremic piglets or the isolation of BVDV in tissues from
preweaned piglets. The differentiation of BVDV from HCV was achieved by
determining reactivity patterns to monoclonal antibodies speciﬁc to BVDV and/or HCV
(Terpstra, 1988; Wenswoort, 1988; Paton, 1992). When a persistently infected calf was
in contact with 6 breeding gilts, transplacental infection with BVDV was conﬁned to l
litter of piglets with 5 of 7 piglets positive for BVDV by virus isolation at birth (Paton,
1992). In sows receiving contaminated vaccine, an examination of 232 piglets from 31
litters revealed that 135 piglets were positive for BVDV by immunoﬂuorescent antibody
testing Of tissues taken at necropsy (Wenswoort, 1988).

Experimental infections. Bovine viral diarrhea virus infection has also been
experimentally reproduced in pigs. Early studies utilized BVDV infection in swine as a

means of immunizing pigs against HCV challenge (Sheffy, 1961; Beckenhauer, 1961;

26
Langer, 1963; Baker, 1963; Tamoglia, 1965; Simonyi, 1967). Different BVDV isolates

were tested, but all BVDV isolates were administered by parenteral administration
(intravenous, intramuscular, subcutaneous). The results of the early experiments
indicated that BVDV infection in swine had some value in protecting pigs from severe
disease following challenge with HCV (Beckenhauer, 1961; Sheffy, 1961). In contrast, a
later experiment using the same viral isolate demonstrated that no protection was offered
as all pigs vaccinated with BVDV died following HCV challenge (Simonyi, I967). The
discrepancy between these early experiments may have been due in part to the preparation
of the inocula, as the actual viral titers of the inocula were unknown. Another experiment
demonstrated that BVDV infection of swine as a vaccination would not meet the criteria
set by the United States Department of Agriculture for licensure because of variability in
protection against HCV challenge and the threat of BVDV transmission to cattle
(Tamoglia, 1965).

Further experimental infections of BVDV in swine utilizing different BVDV isolates
and routes of inoculation were performed in order to obtain additional information on the
immunologic, pathologic, serologic, and virologic responses of pigs infected with BVDV
and to further deﬁne the taxonomic relationship between HCV and BVDV. An early
experimental study documented the infection of pigs with BVDV by exposure of pigs to a
calf (Snowdon, 1968). The report also cited that feeding of tissue from a calf infected
with BVDV resulted in infection and seroconversion. Serologic responses following
experimental infection were further characterized. Neutralizing antibodies following

experimental intranasal BVDV infection in weaned pigs may appear as early as 14 days

27

after infection (Stewart, 1971), but is consistently present at 3 to 4 weeks after inoculation
(Stewart, 1971; Dahle, 1993). Isolation of BVDV following experimental inoculation
was achieved as evidenced by the isolation of BVDV from peripheral blood leukocytes
between days 4 and 17 after inoculation (Castrucci, 1975; Dahle, 1991; Femelius, 1973).
Bovine viral diarrhea virus was isolated from feces and nasal mucus, thus suggesting that
pigs may be involved in the transmission of BVDV to cattle (Femelius, 1973). Intranasal
inoculations of pigs with BVDV were performed to simulate natural infections. Bovine
viral diarrhea virus was isolated most consistently at 7 days after inoculation from tissues
collected at postmortem examination following intranasal inoculation (Stewart, 1971;
Phillip, 1972).

Transplacental infection with BVDV in pigs has been experimentally reproduced
(Stewart, 1980; Dahle, 1987; Leforban, 1990; Paton, 1994). Results following
experimental infection have been quite variable based upon differences in routes of
inoculation, dose of inoculum, isolates used, and the stage Of gestation when inoculation
is performed. Routes of inoculation have included intranasal (Stewart, 1980; Dahle,
1987; Paton, 1994), intramuscular (Leforban, 1990), and intrauterine (Paton, 1994). The
titers of infecting inoculum have ranged from 105‘5 to 107'5 TCIDso. Of 5 isolates
administered by intranasal inoculation between 25 and 55 days of gestation,
transplacental infection with BVDV was achieved with only 1 isolate given on day 46 of
gestation (Stewart, 1980). The isolate causing transplacental infection was originally
isolated from a naturally infected pig and passaged once in a calf. Isolates NY-l and

NADL, which are common reference strains, did not cause transplacental infection.

28
Another study utilizing the NADL isolate also failed to Show transplacental infection

following intramuscular inoculation on day 29 of gestation (Leforban, 1990).
Transplacental infection by intranasal inoculation was not achieved in a study using two
pregnant gilts infected on days 34 and 79 of gestation (Dahle, 1987). The isolate used
was a ﬁeld strain Obtained from a persistently infected heifer from a herd in which there
was a histrory of BVDV-associated abortions. Transplacental infection was reported to
occur at day 37 of gestation following intranasal inoculation and days 35 and 42 of
gestation following intrauterine inoculation of a BVDV strain originally isolated from a
pig (Paton, 1994). Transplacental infection was documented by the presence of viremic
piglets at birth or the presence of neutralizing antibodies to BVDV in precolostral serum
samples obtained from piglets at birth. In addition, transplacental infection occurred
following intrauterine inoculation at day 27 of gestation with an isolate obtained from the
persistently infected steer that was thought to be the index case of the outbreak involving
natural infection of pigs with BVDV (Paton, 1992).

Clinical manifestations. Bovine viral diarrhea Virus infection in immunocompetent,
nonpregnant pigs causes subclinical to mild clinical signs. Most cases of natural infection
are not apparent and the only evidence of infection is the development of neutralizing
antibodies to BVDV following infection (Femelius, 1973). Experimental inoculation of
nonpregnant pigs has also indicated that BVDV infection in swine is subclinical (Phillip,
1972; Femelius, 1973; Dahle, 1991) or causes only mild clinical disease with mild
anorexia and elevated rectal temperatures being reported (Stewart, 1971). Leukopenia is

the only laboratory abnormality reported following infection with BVDV in swine

29
(Stewart, 1971; Carbrey, 1976). These studies have also conﬁrmed that the virus

recovered following experimental infection in swine has retained its virulence for calves
(Stewart, 1971; Femelius, 1973).

Clinical manifestations following infection of pregnant swine have been varied based
upon differences in routes of inoculation and isolates of BVDV used. Bovine viral
diarrhea virus infection in the pregnant pig usually results in subclinical disease, which is
similar to what is reported for infection of nonpregnant pigs (Stewart, 1980). Some cases
of natural BVDV infections on swine farms have been associated with breeding problems
such as poor conception, small litters, and abortions (Terpstra, 1988).

Clinical manifestations following BVDV infection in pregnant swine have been
reported in newborn piglets that have acquired the infection in utero. These clinical signs
are indistinguishable from signs of congenital HCV infection, which include elevated
neonatal mortality, the birth of weak piglets with tremors, and skin hemorrhages (Dahle,
I990). Neurological signs consisting of clonic spasms or tremors have also been reported
at birth following congenital BVDV infection (Stewart, 1977; Terpstra, 1988). Viremic
piglets may also appear normal at birth (Paton, 1994). Clinical manifestations of
congenital BVDV infection usually develop 2 to 3 weeks after birth and include rough
hair coat, growth retardation, wasting, hypertherrnia, diarrhea, petechiae in the skin and
mucus membranes, blue eartips, and conjunctivitis (Terpstra, 1988). In addition, elevated
mortality rates in piglets congenitally infected with BVDV have been reported

(Wenswoort, 1988; Paton, 1994). Anemia and leukopenia have been reported as

30
laboratory ﬁndings associated with congenital BVDV infection (Terpstra, 1988; Paton,

1992)

Pathologic ﬁndings. When BVDV infection occurs in immunocompetent pigs, very
mild or no pathologic lesions have been observed (Stewart, 1971; Carbrey, 1976; Dahle,
1991). Hyperemia of the small intestine was reported as a lesion in a sentinal pig in close
contact with BVDV-infected calves (Stewart, 1971).

Lesions have, however, been reported in congenitally infected piglets born after
transplacental infection and are indistinguishable from congenital HCV infection.
Petechial hemorrhages are the most consistent gross pathologic lesion following prenatal
infection and have been reported in lymph nodes, mucus membranes, epicardium,
kidneys, small intestine, the epiglottis, and the lung (Terpstra, 1988; Paton, 1992; Paton,
1994). Frank hemorrhage into the pericardial sac has also been reported (Paton, 1992).
Other gross lesions that have been reported include anasarca, chronic gastroenteritis,
polyarthritis, necrotic tonsillitis, atrophy of the thymus, generalized emaciation,
generalized paleness (anemia), icterus, generalized edema, ascites, and hypertrophy or
ulceration of the small intestinal mucosa (Terpstra, 1988; Paton, 1992).

Histopathologic lesions have been reported in piglets with congenital BVDV infection.
Meningitis was identiﬁed in 8 of 12 fetuses following experimental transplacental
infection and was characterized by lymphocytes, histiocytes, and adventitial cell
accumulations in the choroid plexus and leptomeninges (Stewart, 1980). Meningitis,

myocarditis, hepatic necrosis, nephritis, widespread hemorrhages, and lymphoid depletion

31
of the germinal centers Of lymph nodes have also been reported (Paton, 1992; Paton,

1994).

Diagnosis. Traditional methods of diagnosis of BVDV infection in cattle can also be
used in the diagnosis of BVDV infection in swine. Virus isolation and serology are the
standard methods for diagnosing BVDV infections and are employed by most diagnostic
laboratories (Dubovi, 1996).

Virus isolation may be performed on precolostral serum samples, buffy coat
preparations, and tissues collected during postmortem examination. Tissues most
suitable for virus isolation for BVDV include tonsil, lung, spleen, and lymph nodes.
Following the isolation, differentiation of BVDV from HCV can be performed using
monoclonal antibody analysis (Zhou, 1989; Edwards, 1991) or polymerase chain reaction

using primers speciﬁc to BVDV or HCV genomes (Katz, 1993; Harding, 1996).

Porcine Fetal Immunology

Two of the possible outcomes following Pestivirus transplacental infection are the
birth of persistently infected offspring or the birth of offspring that are seropositive.
These outcomes are dependent on the stage of development of the immune system of the
fetus at the time infection occurs. Therefore, it is important to understand the
development of the porcine fetal immune response in planning experimental
transplacental infection in pigs with BVDV.

Comparison of bovine and porcine placentation. Swine possess a diffuse,

noninvasive epitheliochorial form of placentation (Jainudeen, 1987). In the early stages

32

of gestation, as many as six layers separate the fetal and maternal bloodstrearns. As
pregnancy advances, fetal capillaries penetrate the trophoblast cell layer and bring fetal
and maternal vascular beds closer. Maternal antibodies, however, do not cross the
placenta in the pig (Trebichavsdy, 1996).

Cattle possess a cotyledonary, noninvasive epitheliochorial form of placentation
(Jainudeen, 1987). The same six layers are present between fetal and maternal circulation
that is present in the porcine placentation, and maternal antibodies do not cross the
placenta in the cow.

Development of the humoral immune system. Porcine fetal pre-B cells are capable of
differentiation into B-cells in the liver by day 35 of gestation (Trebichavsky, 1996). The
fetal B cell humoral immune response as evidenced by the secretion of immunoglobulins
was Observed in liver cells Of 44-day-Old pig fetuses when stimulated in vivo for 6 days
with polyclonal B mitogen derived from Nocardia opaca and administered by
intramuscular injection (Sterzl, 1977). The production of speciﬁc anti-ﬂagellin
antibodies has been shown in 55-day-old pig fetuses when immunized in vivo with
ﬂagellin in incomplete Freund’s adjuvant (Tlaskalova-Hogenova, 1994). Based upon
these studies, it appears pig fetuses become immunocompetent beginning between days
45 and 55 of gestation.

Specific immunotolerance induced by other viruses. Porcine parvovirus is a small,
nonenveloped DNA Virus of the family Parvoviridae that causes reproductive failure of
swine characterized by embryonic and fetal infection and death, usually in the absence of

clinical signs in the dam (Mengeling, 1986). Serological examination of fetuses

33

following experimental in utero infection at different gestational ages suggests that
immunocompetence for porcine parvovirus develops before day 70 of gestation (Nielsen,
1991)

Hog cholera Virus is an enveloped RNA virus of the family F laviviridae, and is related
to BVDV as described in previous sections. Experiments with the Glentorf strain of
HCV have shown that the outcome of transplacental infection is dependent upon the
gestational ages of the fetuses when infection occurs (Meyer, 1981). Inoculation prior to
day 70 of gestation results in abortion or the birth of stillborn fetuses with typical lesions
of HCV. Persistent infections of the litters were exclusively observed when sows had
been inoculated between days 68 and 88 of gestation. Other studies on experimental
transplacental infection Of HCV have supported the concept that the generation of
persistently infected piglets occurs after the stage of immunocompetence of the fetus
(Van Oirschot, 1979; Trautwein, 1986). The mechanism of immunotolerance that
develops during the latter parts of gestation upon exposure to HCV is undetermined.

Porcine reproductive and respiratory syndrome virus is an enveloped RNA virus and a
member of the family Arteriviridae. Under experimental conditions, the virus has been
associated with late-term abortions and the birth of persistently infected piglets following
infection of pregnant gilts at day 90 of gestation (Christianson, 1992; Mengeling, 1994).
The generation of persistently infected piglets beyond the gestational age where

immunocompetence is supposedly achieved is currently under investigation.

LITERATURE REVIEW - SUMMARY

Bovine viral diarrhea virus is an important viral pathogen of cattle and is responsible
for multiple clinicopathologic entities. Infection of pregnant cattle with noncytopathic
BVDV at gestational ages prior to the development of fetal immunocompetence may
result in the birth of calves that are immunotolerant to and persistently infected with
BVDV. Persistently infected offspring are viremic and at risk for the development of
mucosal disease, and they continuously shed Virus. Thus, persistently infected animals
are the primary means by which BVDV is perpetuated within the cattle population.

Methods to control BVDV infections in cattle center around vaccination of susceptible
animals, and the identiﬁcation and removal of persistently infected carriers. More than
140 federally liscenced vaccines are available for BVDV in the United States. The
majority of these vaccines were licensed prior to a complete understanding of the
pathogenesis of BVDV infection. In addition, these vaccines, when applied before
breeding to female cattle, have not been tested to determine whether or not they are
efﬁcacious in preventing transplacental BVDV infection. Prevention of transplacental
infection, in order to prevent the birth of persistently infected carrier offspring, should be
the realistic measure of efﬁcacy for vaccines licensed for BVDV. Vaccine efficacy trials

utilizing pregnant cattle are difﬁcult and expensive to perform.

34

35

Based on a review of the literature, there is strong evidence to support that pigs may
provide an alternative model to cattle for the study of transplacental BVDV infection.
Evidence supporting this includes the fact that pigs can naturally, and under experimental
conditions, be infected with BVDV. In addition, transplacental infection of the fetus has
been reported in pregnant pigs under both natural and experimental conditions.

Developing a model of transplacental infection of pigs has several advantages in the
study of BVDV vaccine efﬁcacy: unlike cattle, it is easy to obtain pigs that are
seronegative to BVDV; the cost of purchasing pregnant gilts is much lower than the cost
of purchasing pregnant heifers; the gestational period of swine is 114 days as opposed to
270 days for cattle, which allows experiments to proceed more rapidly; and, average litter
size in swine is 10, which allows for more replicates, as Opposed to the Single births of
cattle.

The purpose Of the present study was to develop an alternative animal model of BVDV
transplacental infection utilizing swine. This model could then be used to economically
and efﬁciently test commercially available vaccines and new generation vaccines against

BVDV infection.

HYPOTHESES

The overall goal of this project was to develop an alternative animal model of
BVDV transplacental infection utilizing swine in order to develop a means to
economically and efﬁciently test commercially available vaccines and new-generation
vaccines against BVDV. The following hypotheses were tested:

1) Type I and type II BVDV isolates can replicate in porcine turbinate cell
cultures.

2) Feeder age swine can be infected with type I and type II BVDV and
become Viremic.

3) Transplacental BVDV infections can be experimentally produced in

pregnant gilts.

36

MATERIALS AND METHODS

Experimental Animals

F orty-two two-month-Old crossbred pigs obtained from the Michigan State University
Swine Teaching and Research Farm were used in studies to select the BVDV isolate and
dose of virus to be used in experimental infection studies. All pigs were determined to be
free of BVDV by virus isolation on serum and buffy coat samples prior to inoculation
(described below). In addition, all pigs were determined to be free of neutralizing
antibodies to BVDV by serum virus neutralization tests prior to inoculation (described
below). The pigs were housed in isolation rooms at the Michigan State University
Animal Containment Facility and were fed a swine grower ration during the experimental
studies.

Eight crossbred postpubertal gilts, also obtained from the Michigan State University
Swine Teaching and Research Farm, were used for transplacental infection studies. The
sample size of eight animals was determined based upon the minimal sample size needed
to achieve statistical signiﬁcance using the chi-square of independence test with the
power set at 80% and the p value set at 0.05. Estrus was synchronized by the inclusion in
the feed of 0.25 mg melengesterol acetateal per head per day for 7 days. Five days after

the withdrawal of the feed containing the melengesterol acetate, all gilts were exposed to

 

' The Upjohn Co., Kalamazoo, MI
37

38
a crossbred boar for a three day period. Pregnancy was conﬁrmed 40 days later through

the use of scanning transabdominal ultrasonography. Prior to breeding and inoculation,
pregnant gilts were determined to be free of BVDV by virus isolation on serum and buffy
coat samples and seronegative to BVDV by serum virus neutralization. The pregnant
gilts were housed in isolation rooms at the Michigan State University Animal

Containment Facility and were fed a swine gestation ration.

Viral Isolates

Fourteen isolates of BVDV were used in experimental studies, and the source, biotype,
and genotype are summarized in Table 1. Eleven of the isolates were submissions from
ﬁeld cases involving cattle, while the remaining three isolates consisted of BVDV 890,

the prototypical type II isolate, and two isolates obtained from pigs.

Cell Culture

Porcine turbinate (PT) cellsb grown in Eagle’s minimal essential media (EMEM)c
containing 10% fetal equine serum (FES)d, L-glutaminee, penicillin G“, and streptomycinc
were used for the in vitro selection of isolates. Bovine turbinate (BT) cellsf grown in
EMEM containing 10% FES, L-glutamine, penicillin, and streptomycin were used to

propagate Viral isolates used in all portions of this study, for Virus isolation procedures

 

b Dr. Kanitz, Animal Disease Diagnostic Laboratory, Purdue University
° JRH Biosciences, Lenexa, KS

d Sigma Chemical Co., St. Louis, MO

° Gibco BRL, Life Technologies, Grand Island, NY

f National Veterinary Services Laboratory, Ames, IA

39

from specimens collected, and for virus neutralization procedures. Prior to use, cells and

media were determined to be free of BVDV and Mycoplasma spp.

Determination of Viral Genotype

Genotyping of the 11 ﬁeld submissions was performed using the nested reverse
transcription-polymerase chain reaction, as previously described (Ridpath, 1994). Flasks
of BT cells were inoculated with each of the 11 isolates, and were incubated for 48 hours
at 37°C in humidiﬁed air containing 5% C02. Total RNA from BVDV-infected BT cells
was isolated by acid guandinium thiocyanate / phenol / chloroform extraction as adapted
for tissue culture cells (Chromczynski, 1987; Kingston, 1993). Segregation of the 11
BVDV isolates into genotypes was performed using a nested PCR test that ampliﬁed
sequences from the 5’ untranslated region. The ﬁrst round of PCR ampliﬁcation was
performed to differentiate BVDV from other pestiviruses. All BVDV isolates would be
ampliﬁed. The second round of PCR amplication was performed to differentiate type I
BVDV from type II BVDV. Only type II BVDV would be ampliﬁed with the second
round of PCR ampliﬁcation. The PCR primers, reagents, and ampliﬁcation conditions

have previously been described in detail (Ridpath, 1994).

40

Detection of BVDV Antigen by the Immunoperoxidase Monolayer

Assay (IPMA)

Viral infectivity was determined by an immunoperoxidase monolayer assay as
previously described (Meyling,1988). Ninety-six-well microtiter plates‘3 were seeded
with bovine turbinate cells in EMEM containing 10% FES, L-glutamine, penicillin G,
and streptomycin and incubated for 48 hours at 37°C in humidiﬁed air containing 5%
C02. After the media was decanted and replaced with fresh EMEM containing F ES, L-
glutamine, and antibiotics, each well was inoculated with 25 ul of sample. The
inoculated plates were incubated for 72 hours at 37°C in humidiﬁed air containing 5%
C02. The plates were then drained and rinsed twice with 0.01 M phosphate-buffered
saline (PBS). Cells were ﬁxed with 35% acetone in PBS containing 0.02% bovine serum
albumin (BSA)h and incubated at room temperature for 10 minutes. The plates were then
dried. One hundred microliters of polyclonal BVDV antiserum’ diluted 1:90 in binding
buffer (PBS containing 0.05% Tween 20 and 2.95% NaCl) was added to each well and
incubated for 30 minutes at room temperature. Plates were drained and rinsed with wash
buffer (PBS containing 0.05% Tween 20), and 50 ul of protein G-horseradish peroxidasej
diluted 121000 with binding buffer was added to each well and incubated for 15 minutes
at room temperature. Plates were then drained and washed twice with wash buffer, and
100 pl of freshly prepared substrate chromogen solution (3-arnino-9-ethylcarbazolei

dissolved in N,N-dimethyl-formamidej in sodium acetate with hydrogen peroxide) was

 

3 Becton Dickinson & Co., Lincoln Park, NJ

" Sigma Chemical Co., St. Louis, MO

’ National Veterinary Services Laboratory, Ames, IA
JZymed Laboratories lnc., San Francisco, CA

41
added to each well and incubated in the dark for 1 hour at room temperature. The plates

were drained and rinsed twice in tap water. Cells positive for BVDV had a reddish-

brown staining cytoplasm upon examination by light microscopy.

Preparation of Viral and Sham Inocula

Bovine viral diarrhea virus isolates to be used to inoculate pigs were prepared by 2
passages in bovine turbinate cells. Tissue culture ﬂasks were seeded with bovine
turbinate cells in EMEM containing 10% FES, glutamine, and antibiotics, as previously
described. After 48 hours incubation at 37°C in humidiﬁed air containing 5% C02, the
EMEM containing PBS was decanted and replaced by EMEM without serum. The ﬂasks
were inoculated with each BVDV isolate at a multiplicity of infection of 0.1. The viral
inoculum was adsorbed for 60 minutes on a shaker at room temperature. After
adsorption, the viral inoculum was decanted and replaced with EMEM containing FES,
glutamine, and antibiotics. The ﬂasks were then incubated for an additional 5 days at
37°C in humidiﬁed air containing 5% C02. The infected tissue culture ﬂasks were then
subjected to three freeze/thaw cycles to promote cell rupture and release of virus into the
media. The third cell culture passage of virus was used for animal inoculations. All viral
stocks were stored at -80°C.

The sham inoculum was prepared identical to the viral inoculum except for the
exclusion of the viral isolates. A tissue culture ﬂask was seeded with bovine turbinate
cells in EMEM containing 10% FES, glutamine, and antibiotics, as previously described.

After 48 hours incubation at 37°C in humidiﬁed air containing 5% C02, the EMEM

42
containing PBS was decanted and replaced by EMEM without serum, followed by

incubation for 60 minutes on a Shaker at room temperature. The sham inoculum was then
decanted and replaced with EMEM containing F ES, glutamine, and antibiotics. The ﬂask
was incubated for an additional 5 days at 37°C in humidiﬁed air containing 5% C02.
The sham tissue culture ﬂask was then subjected to three freeze/thaw cycles to promote

cell rupture. The sham inoculum was stored at -80°C.

Determination of Viral Titer

The viral titer of all BVDV inocula was determined according to previously described
methods (Carbrey, 1971). Serial 10-fold dilutions of each Viral isolate were made in
EMEM without serum. Each viral isolate was tested at 100 to 10'7 dilutions. Viral
titrations were performed in 96-well microtiter plates". Microtiter plates were seeded
with bovine turbinate cells in EMEM containing 10% F ES, glutamine, and antibiotics as
previously described, and incubated for 48 hours at 37°C in humidiﬁed air containing 5%
C02. After the media was decanted and replaced with fresh EMEM containing FES,
glutamine, and antibiotics, twelve wells were inoculated with 25 ul of sample for each
dilution. The inoculated plates were incubated for 72 hours at 37°C in humidiﬁed air
containing 5% C02. Following incubation, the presence of viral antigen was
demonstrated by the immunoperoxidase monolayer assay as previously described. The
calculation of the viral titer was determined by previously described methods (Karber,

1931).

 

" Becton Dickinson & Co., Lincoln Park, NJ

43

Inoculation of Pigs

The viral stocks and sham inoculum were rapidly thawed in 37°C waterbath, and
dilutions were made using EMEM containing 10% FES, glutamine, and antibiotics.
Feeder age pigs and pregnant gilts were inoculated by intranasal instillation with a
cannula'. The viral stocks were diluted such that the total volume for instillation was 1.0

ml per nostril. The same volume of sham inocula was used.

Virus Isolation

Whole blood was collected by jugular venipuncture for virus isolation in Vacutainer®
tubes"1 containing EDTA. The whole blood was processed in order to obtain buffy coat
preparations. The whole blood was centifuged for 15 minutes at 2200 x g and the plasma
was decanted with a sterile pipet. The buffy coat layer was removed from the underlying
red cells using a sterile pipet, and diluted in 1.0 ml of EMEM with 10% FES. Following
three freeze/thaw cycles, the buffy coat preparations were centrifuged for 10 minutes at
1500 x g. The supernatant was decanted and ﬁltered through a sterile 0.45 um ﬁlter unit“.
The buffy coat preparations were stored at -80°C.

Serum was collected by jugular venipuncture for serology and virus isolation in
Vacutainer® clot tubes°. The clot tubes were centrifuged 15 minutes at 2200 x g and the

serum was decanted with a sterile pipet. The serum samples were stored at ~80°C.

 

’ Pﬁzer lnc., Animal Health Division, New York, NY
m Becton Dickinson & Co., Lincoln Park, NJ

" Millipore Corporation, Bedford, MA

° Becton Dickinson & Co., Lincoln Park, NJ

44

Tissues collected during postmortem examination were processed prior to virus
isolation. Approximately 0.5 gram of tissue was diluted in 4 ml of EMEM with 10% PBS
in a sterile mortar. Sterile sand was added and the tissue was homogenized by grinding
with a pestle. The homogenized tissue sample was transferred into conical tubes and
centrifuged for 30 minutes at 2300 x g. The supernatant was decanted and ﬁltered
through a 0.45 pm ﬁlter unit. Following processing, tissue preparations were stored at
-80°C.

Virus isolation was performed on serum, buffy coat preparations, and tissue
preparations. Roller tubes” were seeded with BT cells in EMEM containing 10% FES,
glutamine, and antibiotics; and incubated for 2 days at 37°C in humidiﬁed air containing
5% C02. Following the 2 days of incubation, the roller tubes were inoculated. The
media was decanted and 600 pl of EMEM without PBS and 400 pl of sample were added.
The roller tubes were incubated for 1 hour at room temperature in a roller drum.
Following the 1 hour adsorption, the samples were decanted and the roller tubes were
washed with 1.0 m1 of EMEM without F ES. A total of 3.0 ml of EMEM containing 10%
FES, glutamine, and antibiotics was added to the roller tubes, which were then incubated
for an additional 7 days in a roller drum at 37°C at 95% humidity in 5% C02. The roller
tubes were subjected to three freeze/thaw cycles and 25 ul of sample was inoculated in
duplicate into wells on 96-well microtiter plates containing monolayers of BT cells in

EMEM with 10% FES. After 3 days of incubation at 37°C in humidiﬁed air containing

 

P Coming lnc., Corning, NY

45
5% C02, the BT cells were stained for BVDV antigen by the IPMA as previously

described.

Serologic Testing

Serology for BVDV was performed by a microtiter serum virus neutralization
procedure according to previously described methods (Carbrey, 1971). Sera obtained
from all experimental animals were tested for neutralizing antibodies to type I and type II
BVDV. For type I antibody determinations, 500 TCIDso of cytopathic Singer isolateq of
BVDV was used. For type 11 antibody determination, 500 TCID50 of MSU-AHDL
#cp1080626 isolater of BVDV was used. Sera were inactivated for 30 minutes in a 56°C
waterbath. The virus neutralization test was set up in 96-well microtiter plates. Serial
two-fold dilutions, ranging from 1:4 to 1:4,096, were made for each serum sample.
Bovine turbinate cells were used as the indicator cells at a dilution of 15,000 cells per
well. Each test included a back titration of the virus and a positive and negative serum
control. The antibody titer was read as the highest dilution with complete inhibition of

cytopathic effect.

Pathologic Examination

Feeder age pigs and pregnant gilts were euthanized by barbiturates overdose by

intravenous injection into the external/intemal jugular veins or anterior vena cava.

 

‘1 National Veterinary Services Laboratory, Ames, IA
rAnimal Health Diagnostic Laboratory, Virology Section, E. Lansing, MI
’ Vortech Pharmaceuticals, Detroit, MI

46

Fetuses were euthanized by barbiturate overdose by intracardiac injection. A gross
postmortem examination was performed on all pigs and fetuses, and gross lesions were
recorded. Tissues collected at postmortem examination were ﬁxed in 10% neutral-
buffered formalin, embeddid in parafﬁn, sectioned at 6 pm, and H & E stained for

histological examination.

Experimental Design: In Vitro Selection of Isolates

Fourteen isolates of BVDV were evaluated for their ability to establish infection in
porcine turbinate cell cultures. The source, biotype, and genotype for the 14 isolates used
in this study are described in Table 1. Viral infectivity in porcine turbinate cells was
determined by the immunoperoxidase monolayer assay, as previously described. A total
of 5 isolates were selected for further study based upon the ability to infect porcine

turbinate cell monolayers.

Experimental Design: In Vivo Selection of Isolates

To further select isolates to use for in vivo studies, three noncytopathic type I BVDV
isolates and 1 noncytopathic type II BVDV isolate were selected based upon the results of
the in vitro study. A ﬁfth untyped noncytopathic isolate, which is a BVDV isolate
obtained from a pig, was also used in this study.

Two days prior to inoculations, ten 2-month-old BVDV-seronegative crossbred pigs
were placed in an animal isolation facility, randomly allocated into 5 groups of 2 pigs

each, with each group housed in a separate room. The 5 groups of pigs each received a

47
different viral isolate as follows: group A, isolate #1330478-1 (type I); group B, isolate

#1322634 (type I); group C, isolate #MPVK-66 (untyped); group D, isolate #1321478
(type I); and group B, isolate #890 (type II).

The dose of Virus for all intranasal inoculations was 107 TCIDso. One pig in each
group received a single inoculation on day 0, and the other pig in each group received 2
inoculations given on day 0 and day 1. All pigs were examined daily, and rectal
temperature, appetite, and fecal consistency were recorded. Serum and whole blood
samples were collected at days 0, 3, 5, and 7 after inoculation for virus isolation. Serum
collected at days 0 and 7 after inoculation was tested for BVDV-speciﬁc neutralizing
antibody by virus neutralization. On day 7 after inoculation, all pigs were euthanized, and
a postmortem examination was performed. Tonsil, bronchial lymph node, ileum, lung,
spleen, and mesenteric lymph node were collected for virus isolation. In addition to the
six tissues listed above, brain, esophagus, heart, stomach, multiple sections of small and
large intestine, liver, kidney, adrenal gland, and skeletal muscle were collected for

histopathologic examination.

Experimental Design: Type I and Type II BVDV Dose Titration Study
Following the in vivo study of viral isolates, a type I (1330478-1) and type H (890)
BVDV isolate were selected for the dose titration studies.
For dose titration studies of type I and type II BVDV, sixteen 2-month-old crossbred
pigs were placed in an animal isolation facility, randomly allocated into 4 groups of 4

pigs each, and housed in separate isolation rooms. The 4 groups of pigs in both the type I

48

and type II dose titration were intranasally inoculated as follows: group A, control (sham-
inoculated); group B, 103 TCIDso; group C, 105 TCIDso; and group D, 107 TCIDso.

All pigs were examined daily, and rectal temperature, appetite, and fecal consistency
were recorded. Serum and whole blood samples were collected at days 0, 3, 5, and 7 after
inoculation for virus isolation. Serum collected at days 0 and 7 after inoculation was used
for antibody determination by virus neutralization. On day 7 after inoculation, all pigs
were euthanized, and a postmortem examination was performed. Tonsil, bronchial lymph
node, ileum, lung, spleen, and mesenteric lymph node were collected for virus isolation.
In addition to the six tissues listed above, brain, esophagus, heart, stomach, multiple
sections of small and large intestine, liver, kidney, adrenal gland, and skeletal muscle

were collected for histopathologic examination.

Experimental Design: Type I BVDV Infection in Pregnant Gilts

Eight pregnant gilts were placed in an animal isolation facility at day 62 of gestation,
randomly allocated into 2 groups of 4 pigs each, and housed in separate rooms. The
control group was intranasally sham-inoculated while the infected group was inoculated
intranasally with 107 TCIDso of the type I BVDV with the dose and genotype being
selected from the results of the previously described dose titration study in feeder pigs.
All pigs were inoculated at day 65 of gestation. All pigs were examined once daily.
Body temperature, appetite, and fecal consistency were recorded. Serum and whole blood
were collected at days 0, 3, 5, 7, 10, 14, 21, 28, 35, 42, and 50 after inoculation for virus

isolation. Serum collected from pregnant gilts on days 0, 7, 14, 21, 28, 35, 42, and 50

49

after inoculation was tested for antibodies to BVDV by virus neutralization. On day 50
after inoculation (day 110 of gestation), a caesarean section was performed. All gilts
were anesthetized with a combination of 2.0 mg/kg xylazine (Rompun’m) and 5 mg/kg
tiletamine-zolazapam (Telazol®"). When the gilts became recumbant, 20 ml of 2%
lidocaine" was subcutaneously administered along the ventral-lateral abdominal wall
beginning at the ﬂank and extending cranially. Following incision of the skin and
abdominal musculature, the uterus was exposed and opened at the distal extremity of the
left uterine horn. Fetuses were removed and identiﬁed by position with a letter
designation beginning from the left uterine horn to the body of the uterus to the right
uterine horn. Serum samples were collected from the umbilical vein for virus isolation
and determination of antibody titer to BVDV by virus neutralization. After antemortem
sampling had been completed, gilts and fetuses were euthanized. A postmortem
examination was performed on all fetuses and gilts. Tonsil, bronchial lymph node, ileum,
lung, spleen, and mesenteric lymph node were collected from the pregnant gilts for virus
isolation. In addition to the six tissues listed above, thymus was also collected for virus
isolation from the fetuses. In addition to the tissues just mentioned, brain, esophagus,
heart, stomach, multiple sections of small and large intestine, liver, kidney, adrenal gland,

and skeletal muscle were collected from fetuses and gilts for histopathologic examination.

 

‘ Bayer Corporation, Shawnee, KS
“ Fort Dodge Laboratories, Fort Dodge, IA
" Butler Co., Dublin, OH

50

Statistical Analysis

A Fisher’s Exact test was used to evaluate virus isolation and virus neutralization data.

Signiﬁcance was set at 0.05 and power was set at 80%.

RESULTS

In Vitro Selection of Isolates

The results of the genotyping of BVDV isolates are presented in Table 1. Four isolates
were type I, 8 isolates were type II, and 2 isolates were untyped. In porcine turbinate (PT)
cell culture, 8 of the 14 isolates grew, as evidenced by demonstration of viral antigen by
the IPMA test and results are summarized in Table 1. Three noncytopathic type I isolates,
one noncytopathic type II isolate, and the noncytopathic untyped isolate that represented
an isolate Of BVDV obtained from a pig were selected based upon their demonstrated

ability to grow in PT cells for further in vivo evaluation.

In Vivo Selection of Isolates

Clinical parameters. The clinical parameters measured in the 5 groups Of feeder age
pigs infected with the BVDV isolates selected from in vitro evaluation were within the
normal limits throughout the study period. Results of rectal temperature measurements
are presented in Appendix 1. Appetite remained normal and the fecal consistency was

unchanged throughout the study period.

51

52
Serology. Neutralizing antibody titers to type I and type II BVDV from serum

obtained at days 0 and 7 after inoculation were negative (less than 4) from all
experimental animals.

Virus isolation. Results of Virus isolation for serum, buffy coat preparations, and
tissues collected at postmortem examination on day 7 after inoculation are presented in
Table 2. BVDV was isolated from all tissues in pigs from groups A (BVDV 1330478-1),
B (BVDV 1322634), C (BVDV MPVK-66), and E (BVDV 890). In group D (BVDV
1322478), however, BVDV was isolated from both pigs from antemortem samples
collected on day 5 after inoculation, yet BVDV was isolated from tissues in only 1 of the
2 pigs. No difference was noted in the virus isolation results between pigs that received
one dose or two doses of the inoculum.

Pathologic examination. No lesions were noted on gross or histopathologic

examination.

Type I and Type II Dose Titration Study

The type I isolate BVDV 1330478-1 and the type II isolate BVDV 890 were selected
for further study in a dose titration study based upon virus isolation results and genotype.
The type I isolate BVDV 1330478-1 was selected over the other 2 type I isolates because
of a more consistent isolation of virus from serum and buffy coat samples. BVDV 890
was selected for further study because it belonged to the type II genotype. In addition, the

comparison between 1 inoculation and 2 inoculations on successive days demonstrated no

53

beneﬁcial response in terms of virus recovery. Therefore, a single inoculation was
selected for further studies.

Clinical parameters. Rectal temperature measurements remained within normal
limits among controls (group A, sham) and infected groups (group B, 103; group C, 105;
and group D, 107 TCIDso) for both type I ( BVDV 1330478-1) and type II (BVDV 890)
genotypes in dose titration studies. The rectal temperature measurements are presented in
Appendix 2. Appetite and fecal consistency remained unchanged throughout the study
period.

Serology. Neutralizing antibody titers to type I and type H BVDV from serum
obtained at days 0 and 7 after inoculation were < 4 from all pigs.

Virus isolation. The results of virus isolation on serum and buffy coat preparations
are presented in Table 3. For isolate #1330478-1 (type I isolate), BVDV was not isolated
from any blood samples in groups A (controls) and B (103 TCIDso). Bovine viral diarrhea
virus was isolated from the serum of 1 pig by day 5 after inoculation in group C (105
TCIDSO). On day 7 after inoculation in group C, 3 of 4 pigs were positive for BVDV,
which was statistically signiﬁcant (p S .05, chi-square test) when compared to groups A
and B. In group D (107 TCIDso), one pig was positive for BVDV by as early as day 3
after inoculation. Virus was recovered from serum in 2 of 4 pigs by day 5 after
inoculation and from serum and buffy coat preparations from 3 of 4 pigs by day 7 after
inoculation. These results for group D were statistically signiﬁcant (p S .05, chi-square
test) when compared to groups A and B. For isolate BVDV 890 (type II isolate), BVDV

was not isolated from antemortem samples.

54

The results of virus isolation on tissues collected at postmortem examination are
presented in Table 4. For isolate #1330478-1 (type I isolate), BVDV was not isolated
from tissues collected at postmortem examination from groups A (controls) and B (103
TCIDso). Bovine viral diarrhea virus was isolated from all tissues collected in groups C
(105 TCID50) and D (107 TCID50), which was signiﬁcant (p S .05, chi-square test) as
compared to groups A and B.

For isolate BVDV 890 (type II isolate), BVDV was not isolated from tissues collected
at postmortem examination from groups A (controls), B (103 TCIDSO), and C (105
TCIDso). Bovine viral diarrhea virus was isolated from 5 of 6 tissues collected in group
D, which was signiﬁcant (p S .05, chi-square test) as compared to groups A, B, and C.
BVDV was isolated in only 1 of 4 pigs from the lung, which was not signiﬁcant (p S .05,
chi-square test) when compared to groups A, B, and C.

Pathologic examination. NO lesions were noted on gross or histopathologic

examination.

Type I BVDV Infection in Pregnant Gilts

The type I isolate BVDV 1330478-1 was selected for further study in pregnant gilts
because of the demonstration of Viremia in serum and buffy coat preparations, as
compared to the type II isolate BVDV 890 in which Virus was not recovered in serum and
buffy coat preparations. In addition, the type I isolate was selected for further study over

the type II isolate based upon the generation of Viremia at a dose of 105 TCIDso, as

55

compared to the type II isolate in which virus could only be recovered in pigs receiving
107 TCIDso.

The dose of 107 TCIDso of the type I isolate was selected for further study in pregnant
gilts over the dose of 105 TCIDso based upon a signiﬁcantly earlier isolation of virus from
serum and buffy coat preparations in the 107 TCIDso group.

Clinical parameters: gilts. Following intranasal inoculation on day 65 of gestation
with the type I isolate of BVDV (BVDV 1330478-1), rectal temperature measurements
were within normal limits in the control and infected groups. The rectal temperature
measurements are presented in Appendix 3. Appetite and fecal consistency remained
unchanged throughout the study period.

Serology: gilts. None of the pigs possessed serum antibodies to BVDV before and
after breeding while still at the Michigan State University Swine Farm. All group A pigs
remained seronegative to BVDV throughout the experiment, whereas group B pigs
displayed seroconversion to BVDV (Table 5), with seroconversion being deﬁned as a
four-fold rise in titer or a change from undetectable to detectable levels of antibody.
Neutralizing antibodies to type I BVDV were ﬁrst detected by day 14 after inoculation in
group B. However, neutralizing antibodies to type II BVDV were not detectable until day
21 after inoculation, and seroconversion as demonstrated by type 11 antibody titers was
only demonstrated in 3 of 4 gilts.

Virus isolation: gilts. Bovine viral diarrhea virus was not isolated from antemortem
or postmortem samples from gilts in the control group. Bovine viral diarrhea virus was

isolated from antemortem samples in group B between days 5 and 7 after inoculation.

56

The pig numbers and samples that were positive for BVDV by virus isolation are as
follows: pig #77, day 5 buffy coat and day 7 buffy coat; pig #80, day 7 serum; pig #81,
day 5 buffy coat, day 7 serum, and day 7 buffy coat; and pig #83, day 7 serum and day 7
buffy coat. Bovine viral diarrhea virus was not isolated from postmortem samples from
gilts in group B.

Pathologic examination: gilts. No lesions were noted on gross or histopathologic
examination of the gilts.

Serology: fetuses. Antibody was not detected in fetuses from gilts in groups A or B.

Virus isolation: fetuses. Bovine Viral diarrhea virus was not isolated in fetuses from
gilts in group A. From the infected group, BVDV was only isolated from the spleen of
one fetus from pig #80.

Pathologic examination: fetuses. A total of 76 fetuses were delivered by caesarean
section on day 110 of gestation. Thirty—three fetuses were derived from the control group
and 43 fetuses from the infected group. All fetuses appeared normal and at term with the
exception of the presence of a mummiﬁed fetus in pig #77 (Group B). No lesions were
noted on gross or histopathologic examination of the fetuses.

Conclusion. Viremia was demonstrated in the pregnant gilts following intranasal
inoculation of 107 TCIDSO of type I BVDV; however, transplacental infection was not
demonstrated, as evidenced by the inability to isolate Virus or demonstrate antibodies to

BVDV in serum samples obtained from piglets at birth.

57

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58

Table 2. Virus isolation results for in viva selection of the BVDV
isolates

GROUPS (Isolates)
Group A Group B Group C Group D Group E

Sample 11330478-12 113226342 SMPVK-662 11322478! 1890!

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Serum 0/2 0/2 0/2 0/2 0/2
dayO
Buffycoat 0/2 0/2 0/2 0/2 0/2
dayO
Serum 0/2 0/2 0/2 0/2 0/2
day3
Buffycoat 0/2 0/2 0/2 0/2 0/2
day3
Serum 2/2 1/2 1/2 2/2 0/2
dayS
Buffycoat 2/2 2/2 1/2 2/2 0/2
day5
Serum 2/2 2/2 1/2 1/2 0/2
day7
Buffycoat 2/2 1/2 0/2 0/2 0/2
dax7
Spleen 2/2 2/2 2/2 1/2 2/2
Tonsil 2/2 2/2 2/2 1/2 2/2
Mesenteric 2/2 2/2 2/2 1/2 2/2
lymphnode
Lung 2/2 2/2 2/2 1/2 2/2
Bronchial 2/2 2/2 2/2 1/2 2/2
lymphnode .
Ileum 2/2 2/2 2/2 1/2 2/2
Key:

All pigs were inoculated by intranasal instillation

Dose of virus was 107 TCIDSO for inoculation

Tissues were obtained on day 7 after inoculation

Cells under group contain # of pigs virus positive / # of pigs in group

One pig in each group was inoculated on day 0, and the other pig in each group
was inoculated on days 0 and 1

59

Table 3. Virus isolation results for serum and buffy coat samples
from feeder age swine in type I and type II dose titration study

Dose titration study: Type I BVDV (isolate #1330478-1)

 

 

 

 

 
   

 

 

 

 

 

 

 

 

 

 

 

 

 

I Genotype / Group designation / Dose
Sample/Day Type I Group A Type I Group B Type I Group C Type I Group D
(control) (103 TCID502 10s TCID50

serum, day 0 __0/_4-—r—_W—
buffycoat,day0 0/4 0/4 0/4 0/4

serum,day3 0/4 0/4 0/4 0/4
buffycoat,day3 0/4 0/4 0/4 1/4

serum,dayS 0/4 0/4 1/4 2/4*
buffycoat,day5 0/4 0/4 0/4 1/4

serum,day7 0/4 0/4 3/4* 3/4*
buffycoat,day7 0/4 0/4 2/4* 3/4*

 

 

Dose titration study: Type II BVDV (isolate # 890)

I Genotype / Group designation / Dose I

 

 

 

 

 

  

  

 

 

 

 

 

 

 

 

 

 

 

 

 

 

m/Day Type II Grout: A Type II Group B Type I I Group C Type II Group D
(control) (103 TCIDSO) (105 TCIDQJ 107 TCIDso
serum,dayO 0/4 0/4 0/4 0/4
buffycoat,day0 0/4 0/4 0/4 0/4
serum,day3 0/4 0/4 0/4 0/4
buffycoat,day3 0/4 0/4 0/4 0/4
serum,dayS 0/4 0/4 0/4 0/4
buffycoat,day5 0/4 0/4 0/4 0/4
serum,day7 0/4 0/4 0/4 0/4
buffycoat,day7 0/4 0/4 0/4 0/4
Key:

* = signiﬁcance at p s .05 as compared to the control group A (within row comparison)

Group A was sham-inoculated; group B was inoculated with 103 TCIDSO, C with 105 TCIDso, and D with
107 TCID50 of BVDV on day 0
Cells under groups contain # of pigs virus positive / # of pigs in group

60

Table 4. Virus isolation results for tissue specimens from feeder age swine
in type I and type II dose titration study

Dose titration study: g Type I BVDV (isolate #1330478-1)

Genotype / Group designation / Dose

 

 

 

 

 

 

 

 

Postmortem Type I Group A Type I Group B Type I Group C Type I Group D
Sample (control) glo3 TCID502 10’ TCID50 lo7 TCIDSO
Spleen 0/4 0/4 4/4* 4/4*
Tonsil 0/4 0/4 4/4* 4/4*

Mesenteric 0/4 0/4 4/4* 4/4*

lymph node
Ileum 0/4 0/4 3/4* 4/4*

Bronchial 0/4 0/4 4/4* 4/4*

lymph node
Lung 0/4 0/4 4/4* 4/4*

 

 

 

 

 

 

 

Dose titration study: Type II BVDV (isolate # 890)

Genotype / Group designation / Dose I

r

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Postmortem Type II Group A Type II Group B Type II Group C Type II Group D
Sample ﬂ (control) (103 TCIDSO) (lo5 TCIDSO) lo7 TCID50
Spleen 0/4 0/4 0/4 2/4*
Tonsil 0/4 0/4 0/4 3/4*

Mesenteric 0/4 0/4 0/4 3/4"

lymph node
Ileum 0/4 0/4 0/4 2/4*

Bronchial 0/4 0/4 0/4 2/4*

lymmnode
Lung 0/4 0/4 0/4 1/4

 

 

Key:

"' = signiﬁcance at p S .05 as compared to the control group A (within row comparison)

Group A was sham-inoculated; group B was inoculated with 103 TCID50, C with 105 TCIDso, and D with
107 T0050 of BVDV on day 0
Tissue samples were collected on day 7 after inoculation
Cells under groups contain # of pigs virus positive / # of pigs in group

61

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DISCUSSION

Bovine viral diarrhea virus possessess the ability to replicate in porcine cells as
demonstrated by the results from the present study. In the in vitro selection of isolates
study, 8 of 14 BVDV isolates grew in porcine turbinate (PT) cell culture. Segregation of
these 8 isolates into genotypes revealed 4 type I, 2 type II, and 2 untyped isolates. All 6
of the isolates in which viral antigen could not be demonstrated in PT cell culture were
type II BVDV. Previous experimental in vitro studies were performed utilizing untyped
isolates of BVDV, so the present study gives the ﬁrst comparison of type I and type II
BVDV replication in porcine cell culture. Previous studies have also demonstrated that
some BVDV isolates possess the ability to replicate in cells from heterologous host
species including pigs, sheep, goats, hamsters, and human cell lines (Femelius, 1969;
Liess, 1990; Moennig, 1990; Roehe, 1994; Bolin, 1994). However, the ability of BVDV
isolates to replicate in cells of porcine origin is diminished when compared to the ability
to replicate in cells of bovine origin (Liess, 1990; Roehe, 1994). For example, of 35
BVDV isolates examined for replication in a porcine kidney cell line (PK-15), only 2
isolates grew efﬁciently (Moennig, 1990). In another study examining the ability of 26
pestiviruses to grow in bovine turbinate (BT), sheep choroid plexus (SCP), and PK-15
cells, only 2 of 8 BVDV isolates replicated in PK-lS cells (Roche, 1994). In addition, the

2 isolates replicated to a much lower titer when compared to replication in bovine

62

 

63

turbinate (BT) cells. This is in contrast to hog cholera virus isolates, in which 11 of 12
isolates replicated in BT cells. This inability of BVDV isolates to replicate in porcine cell
culture does not exclude the possibility of these isolates to infect pigs. (Roche, 1994;
Paton, 1994).

The absence of clinical signs of disease following experimental infection in the
feeder age pigs and the pregnant gilts in the present study is similar to previous reports
(Femelius, 1973; Stewart, 1980; Stewart, 1971). Bovine viral diarrhea virus infection is
subclinical in immunocompetent, seronegative pigs. These previous reports utilized
isolates of BVDV in which the genotype was undetermined. Some type 11 isolates of
BVDV have been associated with severe, clinical disease in immunocompetent cattle
(Drake, 1994; Carman, 1994; Pellerin, 1994). The type II isolate (BVDV 890) used in the
present study has been previously reported to cause clinical disease and thrombocytopenia
in calves following experimental intranasal inoculation (Bolin, 1992; Walz, 1996). The
present report is the ﬁrst comparative study of type I and type II BVDV infection in
feeder age swine, and both type I and type II BVDV infections failed to produce clinical
signs in pigs.

Seroconversion to BVDV as evidenced by the presence of detectable levels of serum
neutralizing antibodies was not observed in the in viva selection of isolates study and the
type I and type II BVDV dose titration studies. This can be explained by the experimental
design, because the feeder age pigs were euthanized on day 7 after inoculation. Previous
experiments on BVDV infection in feeder pigs have demonstrated that serum neutralizing

antibodies ﬁrst appear at day 14 after inoculation (Stewart, 1971; Phillip, 1972). At day 7

64

after inoculation, it is too early in the infection to detect a neutralizing antibody response
to BVDV infection. This is similar to a report in cattle in which seroconversion generally
occurs 2 to 4 weeks after infection (Baker, 1995).

The in vivo selection of isolates study was conducted in an uncontrolled manner in
order to initially screen and select isolates for further in vivo controlled studies in feeder
pigs. Bovine viral diarrhea virus was isolated from serum and buffy coat preparations
between days 5 and 7 after inoculation from 4 of the 5 groups of feeder age swine in the
in vivo selection of isolates study. This is comparable to previous experimental reports in
which virus was isolated in peripheral blood leukocytes between days 4 and 17 after
inoculation (Dahle, 1991; Femelius, 1973). As shown in Table 2, BVDV was isolated
from all serum and buffy coat samples taken from pigs infected with the type I isolate
(BVDV 1330478-1) between day 5 and 7 after inoculation. Virus was recovered less
frequently from the other type 1 isolates and the untyped isolate (BVDV MPVK-66),
which was originally isolated from a pig. In contrast, BVDV was not isolated from serum
or buﬂy coat samples in group B (type H isolate, BVDV 890). Although only one type II
isolate was used and the experiment was uncontrolled, it appears that the type I isolates of
BVDV are better at establishing infection in pigs. This is based on the virus isolation
results which demonstrated that BVDV was more frequently recovered from serum and
buffy coat preparations from pigs receiving the type I isolates than from the type II
isolate. The type I isolate BVDV 1330478-1 was selected for further study in feeder pigs
over the other type I isolates and the untyped isolate because of the more consistent

isolation of virus from serum and buffy coat preparations. The type II isolate, BVDV

65

890, was selected for further study in feeder pigs to allow for a comparison between type
I and type II BVDV infection in pigs.

Virus isolation procedures were performed on both serum and buffy coat preparations
in the present study. From the results of the in vivo selection of isolates study and the
type I and type II BVDV dose titration studies, it appears that serum is a better sample
than buffy coat preparations in isolating BVDV. This is in contrast to previous reports
indicating that mononuclear cells in the buffy coat obtained from whole blood are the best
sample for virus isolation procedures for submissions from cattle (Brock, 1995; Bezek,
1996; Dubovi, 1996). The decreased recovery of BVDV from porcine buffy coat
preparations may be due to the variability in the harvesting of mononuclear cells ﬁ'om the
buffy coats. Hypotonic lysis of red blood cells with recovery of the leukocytes is the
method for buffy coat preparations from bovine whole blood. Hypotonic lysis procedures
on porcine whole blood result in lysis of leukocytes as well as red blood cells (Walz,
personal observation). Therefore, centrifugation and pipetting of the buffy coat layer was
used in this experiment with variable success.

Based upon the results of the type I and type II BVDV dose titration study, it appears
that the type I BVDV isolate established infection more consistently than the type II
isolate. In the type I BVDV dose titration study, BVDV was isolated signiﬁcantly from
serum and buffy coat samples from groups C (105) and D (107) of feeder pigs receiving
the type I isolate as compared to groups A (controls) and B (1 03). In addition, BVDV was
signiﬁcantly isolated from all tissues collected in group C (105) and group D (107)

receiving the type I isolate, as compared to groups A (controls) and B (103). In the type II

66

dose titration study, BVDV was isolated only from tissues collected during the
postmortem examination. Bovine viral diarrhea virus was signiﬁcantly isolated from 5 of
6 tissues in group D (107) as compared to groups A (controls), B (103), and C (105)
receiving the type II isolate. Based upon these results and the results from the in vivo
selection of isolates study, it appears that there is an increased ability for type 1 isolates to
establish infection in pigs over type 11 isolates. This difference in ability of type I and
type II BVDV to establish infection in pigs may be due to the same differences between
type I and type II BVDV in establishing infection in porcine turbinate cell culture.
Comparison of nucleic acid sequences from type I and type II BVDV revealed greater
than a 30% dissimilarity (Ridpath, 1995). This dissimilarity is concentrated in the regions
of the BVDV genome encoding for the structural glycoprotein E2 and the S’untranslated
region. The signiﬁcance of differences in the sequence encoding for the glycoprotein E2
is the fact that this glycoprotein is responsible for the viral attachment to cells (Xue,
1993). Bovine viral diarrhea virus enters cells by receptor-mediated endocytosis
(Boulanger, 1992). The triggering of endocytosis occurs as a result of the interaction
between the viral glycoprotein E2 and a 50 kilodalton receptor (Xue, 1993). This 50
kilodalton receptor has been identiﬁed in porcine cell culture through the use of an anti-
idiotype antibody to BVDV glycoprotein E2 (Xue, 1996). The difference in nucleotide
sequence of E2 between type I and type II BVDV isolates may have an effect on the
glycoprotein structure, and may account for the inability of type 11 isolates to attach to
porcine cells in vivo and in vitro. The signiﬁcance of differences in the S’untranslated

region sequence is the fact that the internal ribosome entry site element lies within this

67

region. The internal ribosome entry site element is responsible for the initiation of
translation of the viral RNA (Poole, 1995). The virus may be able to penetrate
susceptible porcine cells, but porcine ribosomes may be unable to bind to this internal
ribosome entry site element, and the infection would thus become abortive.

In the present study, the pregnant gilts were infected at day 65 of gestation in order to
demonstrate transplacental infection by measuring two possible outcomes. The potential
outcomes measured were fetal serum neutralizing antibody titers and the presence of virus
in sera and tissues obtained from fetuses. Previous experiments involving BVDV
infection in pregnant gilts have used earlier gestation ages, different routes of inoculation,
and different isolates, in order to demonstrate persistently viremic piglets (Stewart, 1980;
Paton, 1994), but the results from these studies have been inconsistent with respect to the
demonstration of transplacental infection. The stage of gestation when the porcine fetus
becomes immunocompetent occurs between days 45 and 55 of gestation (Tlaskalova-
Hoganova, 1994). To our knowledge, only one study exists in which pregnant gilts were
infected with BVDV beyond the development of fetal immunocompetence (Dahle, 1987).
Although transplacental infection did not occur, the study has several weaknesses. Only
1 sow was infected with BVDV beyond the development of fetal immunocompetence at
day 79 of gestation, and there were no control animals. The present report is the ﬁrst
controlled study of BVDV infection in pregnant gilts beyond the development of fetal
immunocompetence.

An additional reason for the choice of inoculation on day 65 of gestation came from a

review of the literature on hog cholera virus infection. Following hog cholera virus

68
infection, transplacental infection of pregnant gilts with the birth of persistently infected

piglets occurs exclusively between days 68 and 88 of gestation (Liess, 1984). Other
studies of experimental transplacental infection of hog cholera virus have supported the
concept that the generation of persistently infected piglets occurs after the stage of
immunocompetence of the fetus (Van Oirschot, 1979; Trautwein, 1986). The evidence
that the related pestivirus, hog cholera virus, caused persistent infection of piglets beyond
the development of fetal immunocompetence ﬁuther supported the choice of day 65 of
gestation for inoculation of pregnant gilts in the present study. Virus isolation was
performed on sera and tissues obtained from fetuses in order to demonstrate
transplacental infection by the birth of viremic piglets.

Viremia was induced in the pregnant gilts following inoculation at day 65 of gestation
with the type I BVDV isolate, but transplacental infection was not demonstrated with the
exception of one fetus out of 43 fetuses from the 4 gilts in the infected group. Bovine
viral diarrhea virus was isolated from serum and buffy coat samples from all gilts in the
infected group between days 5 and 7 after inoculation. This is consistent with a previous
report of Viremia being demonstrated on day 7 after inoculation with 4 of 5 isolates
following experimental intranasal inoculation of pregnant gilts (Stewart, 1980). In the
present study, seroconversion was also demonstrated in all gilts from the infected group.

Seroconversion was demonstrated in pregnant gilts following infection with the type I
isolate of BVDV. Detectable serum neutralizing antibodies to type I BVDV appeared at
day 14 after inoculation in 2 of 4 gilts and all gilts were seropositive by day 21 after

inoculation in the infected group. This is similar to previous studies with detectable

69

serum neutralizing antibody ﬁrst appearing around day 14 after inoculation (Stewart,
1980). The comparative serology titers between type I and type H BVDV demonstrate
appreciable differences in the levels of neutralizing antibodies to type I and type H BVDV
(refer to Table 5). Seroconversion to type II BVDV was demonstrated in only 3 of 4 gilts.
In addition, the presence of detectable serum neutralizing antibody to type II BVDV was
delayed to day 21 after inoculation, and the levels of neutralizing antibody to type II
BVDV were less when compared to type I neutralizing antibodies. Type I and type 11
antibody determinations in calves following vaccination with a modiﬁed live vaccine or
inactivated vaccine containing type I BVDV has demonstrated lower antibody titers to
type II BVDV than to type I BVDV (Fulton, 1995).

The ideal alternative animal model for transplacental infection with BVDV infection
would be one that produces transplacental infection in 100% of the infected group
animals. It has been previously reported that BVDV transplacental infection in cattle
occurs at a relatively high frequency, with estimates of 100% efﬁciency (Duffell, 1985).
Inoculating pregnant gilts at day 65 of gestation with the type I isolate 1330478-1 did not
consistently produce transplacental infection, and thus would not be recommended as an
alternative animal model of transplacental BVDV infection.

Different experimental conditions could have been used in this experiment to
potentially demonstrate transplacental infection. Inoculation of the pregnant gilts at an
earlier gestational age (days 35 to 45) of the fetuses might have resulted in transplacental
infection. A caesarean section could be performed 3 weeks aﬁer experimental

inoculation in order to collect sera and tissues from fetuses for virus isolation. Previous

70

studies on BVDV infection in pregnant gilts at earlier gestational ages, however, have
been inconsistent in the demonstration of transplacental infection (Paton, 1994; Stewart,
1980). Thus, it remains questionable if swine can serve as an alternative animal model to

study transplacental infection with BVDV.

SUMMARY AND CONCLUSIONS

Fourteen BVDV isolates were examined for their ability to replicate in porcine cells.
Segregation of the isolates into genotypes revealed that 4 of 5 type I isolates replicated in
porcine turbinate cells, as compared to only 1 of 9 type 11 isolates.

Feeder age swine are susceptible to experimental type I and type II BVDV infection.
The present study demonstrates that a detectable Viremia can be induced in feeder age
swine following intranasal inoculation with a dose of 105 TCIDso of a type I isolate of
BVDV. In contrast, a detectable Viremia could not be induced in feeder age swine
following intranasal inoculation with a type II isolate of BVDV. Bovine viral diarrhea
virus was isolated from tissues collected at postmortem examination from pigs infected
with a dose of 105 TCIDso of a type I isolate of BVDV, whereas BVDV could only be
isolated from tissues from pigs receiving a dose of 107 TCIDso of the type II isolate of
BVDV. From these experiments, it appears that type I BVDV isolates are able to infect
pigs more effectively than type II BVDV isolates.

Swine did not serve as an alternative animal model for BVDV transplacental infection
under the conditions described in the present study. Following experimental intranasal
inoculation of pregnant gilts at day 65 of gestation with type I BVDV, Viremia was
established, but transplacental BVDV infection was not demonstrated. The development

of porcine fetal immunocompetence occurs between days 45 and 55 of gestation. The

7]

72

related pestivirus, hog cholera virus, is capable of inducing transplacental infection with
the generation of persistently infected piglets beyond the gestational age when porcine
fetal immunocompetence occurs. Based on the results of this study, BVDV does not
appear to possess this ability to induce persistent infections beyond the development of
fetal immunocompetence, nor did it appear that BVDV fetal infection occurred because

of an absence of neutralizing antibody response in the fetal pigs.

APPENDICES

APPENDIX A

Results of rectal temperatures measured for in vivo selection of isolates

 

 

 

 

 

 

 

 

 

 

 

 

Group A Group B Group C Group D Group E

pig pig pig pig pig pig pig pig pig pig

#1 #2 #3 #4 #5 #6 #7 #8 #92 #93
day -2 102.0 101.4 102.0 102.8 103.0 102.4 102.2 102.4 102.0 103.4
day -1 102.4 102.2 102.8 103.0 102.8 102.2 102.8 102.8 101.6 101.8
day 0 101.7 104.0 102.4 102.4 103.0 102.2 102.4 101.4 102.0 102.0
day +1 101.4 103.0 103.0 103.2 103.0 102.6 102.8 102.4 102.6 102.0
day +2 102.6 102.4 103.4 103.6 103.4 102.6 101.8 101.2 101.8 102.0
day +3 103.0 102.6 103.0 103.6 102.6 102.8 102.6 102.0 102.0 102.4
day +4 102.6 102.6 102.6 103.2 101.4 102.4 101.6 102.4 102.6 102.2
day +5 101.6 102.0 103.4 102.8 101.6 102.0 102.0 102.0 101.6 102.0
day +6 102.6 102.0 103.6 103.8 101.8 101.4 101.4 101.4 102.0 101.2
day +7 102.2 102.0 102.4 102.6 102.6 102.0 102.4 102.0 102.6 101.8

 

 

 

 

 

 

 

 

 

 

 

Key: All temperatures expressed in °F.

Group A inoculated with BVDV 1330478-1; group B with BVDV 1322634;

group C with BVDV MPVK-66; group D with BVDV 1322478; and group B with

BVDV 890
Dose of virus was 107 TCID50 for inoculation

73

 

APPENDIX B

Results of rectal temperature measurements for dose titration study of type I and
type II BVDV

Type I isolate: BVDV 1330478-1

 

 

 

 

 

 

 

 

 

 

 

 

 

Group A Group B

pig #1 pig #ZL pig #3 pig #4 i #5 i #6 pig #7 i #8
day -2 103.0 103.8 103.4 103.6 '— 104.8 103.6 102.8 103.6
day -1 103.0 102.6 103.6 102.4 102.6 102.6 102.4 103.2
day 0 102.8 102.6 102.8 102.8 103.0 102.6 101.2 102.4
day +1 103.0 102.6 101.6 101.0 103.4 102.6 102.8 102.0
day +2 102.2 103.0 102.0 101.0 102.4 101.8 102.0 102.2
day +3 102.4 102.2 100.2 99.0 102.0 101.8 102.0 101.8
day +4 101.2 102.2 101.3 100.0 102.0 102.0 102.2 102.0
day +5 101.4 100.6 100.0 100.8 102.4 102.6 102.0 101.8
day +6 100.8 100.6 101.4 101.0 102.4 102.0 101.8 102.2
day +7 102.2 102.4 102.0 101.8 102.4 102.6 102.6 102.2

 

 

 

 

 

 

 

 

 

 

 

- -. .i#l4 n-lm

102.8 ND ND
102.8 103.0 103.4

102.8 101.8 102.6
101.6 102.6 102.0
102.0 102.6 102.8
102.6 102.2 103.2
102.2 102.0 102.8
102.6 102.6 102.8
102.4 102.4 102.6
102.6 102.8 103.0

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Key: All temperatures expressed in °F.
Group A was sham-inoculated; groups B, C, and D, were inoculated with 103, 105, and 107 TCID50 of
BVDV, respectively, on day 0.

74

75

Results of rectal temperature measurements for dose titration study of type I and

type 11 BVDV

Type II isolate: BVDV 890

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Group A Group B

day -2
day-l
day 0 . . . . . . . .
day +1 103.8 104.2 104.2 103.6 102.4 102.2 103.0 102.4
day +2 103.8 104.0 103.2 103.4 102.0 102.2 103.0 102.0
day +3 103.8 104.0 102.6 102.8 101.8 102.4 102.0 103.4
day +4 103.2 104.0 102.2 102.6 102.2 102.0 102.4 102.4
day +5 103.4 103.6 102.2 103.0 102.0 102.6 102.2 102.8
day +6 102.4 103.6 102.8 102.6 102.2 102.4 102.8 102.4
day +7 102.8 102.2 103.2 102.8 102.6 102.4 102.4 102.6

_ Grog) C

P18 P18 Pig P18 P18

__ #100 #101 #102 #10_#104

- my 103.4 103.2 104.0 105.0 103.8
day -1 103.8 103.6 104.0 103.2 102.8 102.5 102.0 103.8
day 0 104.0 103.0 104.6 103.6 103.8 104.6 103.2 103.6
day +1 103.2 103.0 103.0 103.4 104.6 103.2 103.0 102.6
day +2 103.2 102.6 103.0 103.2 104.6 104.0 102.6 103.2
day +3 102.8 103.2 102.8 103.0 104.4 105.4 103.0 103.2
day +4 102.0 102.0 102.6 103.0 103.8 103.0 103.0 104.4
day +5 103.2 103.2 102.8 103.6 104.4 104.0 103.6 104.4
day +6 103.4 103.2 102.2 103.6 103.6 104.4 103.4 102.8
day +7 103.0 102.8 102.8 103.2 103.6 103.8 103.0 103.0

 

Key: All temperatures expressed in °F.
Group A was sham-inoculated; groups B, C, and D, were inoculated with 103, 105, and 107 TCIDso of
BVDV, respectively, on day 0.

 

 

 

APPENDIX C

Rectal temperature measurements for pregnant gilts infected with BVDV on day 65 of
gestation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Group A (controls) Group B (infected)
pig #76 pig #78 pig #79 pig #82 pig #77 pig #80 pig #81

day -2 ==101.8 102.0 WWW—“rm— 102.8 102.6 101.8
day -1 101.8 103.0 103.0 101.8 102.6 102.6 101.8 101.8

day 0 102.0 102.6 102.8 101.6 101.8 102.4 101.8 102.8
day +1 102.6 101.8 101.8 103.0 101.8 102.0 101.8 102.6
day +2 101.6 101.8 101.8 102.8 102.8 102.8 102.4 102.8
day +3 101.8 101.8 102.0 102.6 102.6 102.6 102.4 102.6
day +4 102.4 102.4 101.8 102.4 102.4 102.4 102.0 102.4
day +5 102.4 102.4 101.6 102.0 102.0 102.0 102.2 102.0
day +6 102.0 102.0 103.0 102.2 102.4 102.2 102.8 102.8
day +7 102.2 102.2 102.8 102.6 102.4 102.6 102.6 102.6
day +8 101.8 102.6 102.6 102.6 102.0 102.6 102.4 102.4
day +9 102.2 102.6 101.8 103.0 102.2 103.0 102.0 102.0
day +10 102.2 103.0 101.6 102.2 101.6 102.2 102.6 102.2
day +11 101.4 102.8 102.2 102.6 102.2 102.6 102.6 102.6
day +12 101.8 102.0 101.6 102.4 102.6 102.2 103.0 102.6
day +13 101.8 102.4 102.2 102.0 102.4 102.6 102.8 103.0
day +14 102.0 102.6 102.6 102.0 101.8 102.4 102.0 102.2
day +15 102.0 101.8 102.4 101.6 102.0 102.0 102.4 102.6
day +16 102.2 101.6 102.0 101.8 102.0 102.0 102.6 101.8
day +17 102.6 101.6 102.0 102.0 102.2 101.6 102.4 101.8
day +18 102.6 101.8 101.6 101.8 103.0 101.6 101.8 101.8
day +19 102.4 101.8 101.8 101.8 102.8 101.6 102.0 102.0
day +20 102.0 101.8 102.0 101.8 102.8 102.6 102.0 103.0
day +21 101.6 101.8 102.6 102.0 102.6 101.8 102.2 102.8
day +22 101.6 102.0 102.6 103.0 101.8 101.8 102.4 102.0
day +23 102.6 103.0 102.4 102.8 101.8 101.4 101.6 103.0
day +24 101.8 102.8 101.6 102.8 101.4 102.2 101.6 102.8
day +25 101.8 102.8 101.6 102.6 102.2 102.0 103.2 102.8

 

 

 

 

 

 

 

 

 

 

 

76

 

'r‘

Rectal temperature measurements for pregnant gilts infected with BVDV on day 65 of

77

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

gestation
Group A (controls) Group B (infected)
pig #76 pig #78 pig #79 pig #82 pig #77 pig #80 pig #81 pig #83
day +26 101.4 102.6 103.2 102.0 102.0 102.8 102.8 102.6
day +27 102.2 102.0 102.8 102.0 102.0 102.6 102.6 102.0
day +28 102.0 102.0 102.6 102.2 102.0 102.6 102.6 102.0
day +29 102.8 102.0 102.6 102.0 102.6 101.8 102.0 102.2
day +30 102.6 102.6 102.0 101.8 101.8 101.8 102.0 102.0
day +31 102.6 101.8 102.0 101.6 101.8 101.4 101.8 101.8
day +32 102.2 101.8 101.8 101.6 101.6 102.2 101.6 102.0
day +33 102.2 102.0 101.6 102.4 103.0 102.2 102.4 102.6
day +34 102.0 102.2 103.0 102.2 102.0 102.0 102.2 101.8
day +35 101.8 101.8 102.0 102.2 101.2 101.8 102.2 101.8
day +36 101.6 101.6 101.2 101.8 101.6 101.6 101.8 101.6
day +37 101.6 101.8 101.6 101.8 101.8 101.6 102.0 102.6
day +38 102.4 101.2 102.0 101.8 101.8 102.4 102.0 102.6
day +39 102.2 101.2 101.8 102.0 102.0 102.2 102.6 102.0
day +40 102.2 101.6 101.8 102.2 102.2 102.2 101.8 102.0
day +41 101.8 102.0 102.0 101.8 101.8 101.8 101.8 101.8
day +42 101.8 101.8 102.2 101.6 101.6 102.2 101.6 103.0
day +43 101.6 102.0 103.2 101.6 102.4 101.8 102.0 102.0
day +44 101.8 101.8 102.8 102.4 102.2 101.6 101.8 101.2
day +45 103.0 102.6 102.6 102.2 102.2 101.6 102.0 101.6

 

 

 

 

 

 

 

 

 

 

Key: All temperatures expressed in °F.
Group A was sham-inoculated; group B was inoculated with 107 TCIDSO of BVDV isolate #1330478-1 on day 0.

 

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