"Michsgm’a Mate
University

 

——"‘ __—

This is to certify that the

dissertation entitled

COMPARATIVE VIRULENCE OF PORCINE ROTAVIRUS

SEROTYPES 1 AND 2 IN GNOTOBIOTIC PIGS

presented by

JAMES EDWARD COLLINS

has been accepted towards fulfillment
of the requirements for

Ph'D° degreein Pathology

 

 

Major professor

Date July 2, 1986

MSU is an Affirmative Action/Equal Opportunity Institution 0—12771

 

 

 

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COMPARATIVE VIRULENCE OF PORCINE ROTAVIRUS
SEROTYPES 1 AND 2 IN GNOTOBIOTIC PIGS

BY

James Edward Collins

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Department of Pathology

1986

 

 

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928. 8134

ABSTRACT
COMPARATIVE VIRULENCE OF PORCINE ROTAVIRUS
SEROTYPES 1 AND 2 IN GNOTOBIOTIC PIGS
BY
JAMES EDWARD COLLINS

The purpose of the present study was to compare
serotypes 1 (Ohio State University strain) and 2 (South
Dakota State University strain) of porcine rotavirus to
determine if specific clinical signs, mortality rates,
virus shedding patterns, virus antigen distribution, or
lesion distribution and severity were related to the
serotype of rotavirus with which pigs were infected.

Forty hysterotomy derived gnotobiotic pigs were
inoculated orally at 3 days of age with 2 ml of
homogenate containing 105 pig-infectious dose50 of either
the Ohio State University (OSU) strain or the South
Dakota State University (SDSU) strain of porcine
rotavirus. Controls were inoculated with media only.
Mortality, clinical signs, and body weights were
monitored daily. Five pigs in each of the virus
inoculated groups and 4 control pigs were killed at 24
and 168 hours after inoculation. Four pigs in each of

the 3 experimental groups (OSU, SDSU, and control) were

 

killed
collect
immunof
examin;
A
strain
throug
SDSU 5
hours
obserV
OSU st
did h<
strah
but d
given
immun
given
after
duode
and j
and j
flexi
COIIIp.‘
Althl

the

killed 72 hours after inoculation. Specimens were
collected at necropsy for serum chemistry, histologic,
immunofluorescent, and scanning electron microscopic
examinations.

All of the pigs inoculated with the OSU or SDSU
strains survived. Control pigs remained healthy
throughout the study. Pigs inoculated with the OSU or
SDSU strains developed diarrhea 19-48 hours and 24-54
hours after inoculation, respectively. Vomiting was
observed in 5 of 14 (36%) of the pigs infected with the
OSU strain whereas pigs inoculated with the SDSU strain
did not vomit. Pigs inoculated with the OSU or SDSU
strains had reduced weight gain compared to control pigs,
but there was no difference in weight gain between pigs
given the OSU or SDSU strains. Results of
immunofluorescent examinations were similar for pigs
given either rotavirus strain except that at 24 hours
after inoculation, viral antigen was detected at the
duodenal-jejunal flexure in 5/5 pigs given the OSU strain
and in 1/5 pigs given the SDSU strain. Villous atrophy
and fusion were more severe at the duodenal—jejunal
flexure of pigs inoculated with the SDSU strain when
compared to pigs inoculated with the OSU strain.
Although the OSU and SDSU strains did show differences in
the occurrence of vomiting and distribution of Villous
atrophy, these strains were equally virulent for

3-day—old gnotobiotic pigs.

 

 

 

Dedicated, with love, to my wife, Barbara,

and our sons, Brian and Daniel.

iv

 

 

 

Depar
Adalb
to pu

fashi

Dr. 8
Dr. E
SUPPC
stud}
Thong

C0mmi

assi

Rita

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Dak

ACKNOWLEDGEMENTS

I wish to express my gratitude to the faculty in the
Department of Pathology, particularly Drs. Robert Leader,
Adalbert Koestner, and Janver Krehbiel, for allowing me
to pursue a doctorate degree in a rather non-traditional
fashion.

I wish to express my sincere appreciation to
Dr. Stuart Sleight, my major professor, and to
Dr. David Benfield, my research advisor, for their
support, guidance, and friendship during my course of
study. I also wish to thank Drs. Roger Maes,

Thomas Mullaney, and Glenn Waxler, members of my guidance
committee, for their helpful suggestions.

Special thanks to Dr. Mahlon Vorhies, Chairman,
Department of Veterinary Science, South Dakota State
University, for providing financial and personal support
for this and several other research projects.

I wish to acknowledge Julie Duimstra for technical
assistance with electron microscopic studies and
Rita Miller for typing this dissertation.

I also wish to thank the many faculty and staff
members in the Department of Veterinary Science at South

Dakota State University who contributed to this project.

 

sons,

under:

 

My deepest appreciation to my wife, Barbara, and our
sons, Brian and Daniel, for their support, love, and

understanding throughout my studies.

vi

 

 

 

LIST I
LIST
INTRO

LITER

MATE

TABLE OF CONTENTS
Page
LIST OF TABLES. . . . . . . . . . . . . . . . . . . . ix
LIST OF FIGURES . . . . . . . . . . . . . . . . . . . xi
INTRODUCTION. 1
LITERATURE REVIEW . 4
Classification . . . . . . . 4
Rotavirus Serotypes and Subgroups . . . . . 4
. . . . . g
9

Rotavirus Electropherotypes
Classification Systems. .

Atypical Rotaviruses . . . . .
Physicochemical Properties . . . . . . . . . . . 12
Cultivation in Cell Culture. . . . . . . . . . . l3
Morphogenesis. . . . . . . . . . . . . . . l7
Epidemiology . 20
Clinical Signs . . . . . . . . . . . . . . . . . 23
Clinical Pathology . . . . . . . . . . . . . . . 28
Pathophysiology. . . . . . . . . . . . . 30
Lesions and Pathogenesis . . . . . . . . . . . . 31
Gross Lesions . . . . . . . . . . . . 31
Histopathologic Lesions . . . . . . . . . . 32
Scanning Electron Microscopy. . . . . . . . 36
Transmission Electron Microscopy. . . . . . 37

Immunofluorescence and Location of

Viral Antigens . . . . . . . . . . . . . 37
Viral Shedding. . . . . . . . . . . . . . . 39
Immunology . . . . . . . . . . . . . . 39
Humoral Immunity. . . . . . . . . . . . . . 39
Cell- mediated Immunity. . . . . . . . . . . 41
Immunization. . . . . . . . . . . . . . . 43
Diagnosis. . . . . . 46
Enzyme— linked Immunosorbent Assays. . . . . 46
Electron Microscopy . . . . . . . . . . . . 48
Immunofluorescence. . . . . . . . . . . . 49
Other Diagnostic Techniques . . . . . . . . 50
Treatment and Control. . . . . . . . . . . . . . 50
MATERIALS AND METHODS . . . . . . . . . . . . . . . . 53
Gnotobiotic Pigs . . . . . . . . . . . . . . . . 53
Virus. . . . . . . . . . . . . . . . . . . 54

 

 

 

RESUl

DISC
SUI-ll?
BIBI

VITI‘

 

Virus Infectivity Titrations . . . . .

Experimental Design. . -

Clinical Signs and Body Weight .

Clinical Pathology . . .

Necropsy .

Preparation of Fluorescein- conjugated
Gamma Globulin. .

Examination by Immunofluorescence.

Light Microscopy .

Scanning Electron Microscopy

Electron Microscopic Examination of
Feces . . . .

Statistical Analysis

RESULTS .

Clinical Signs

Body Weight. .

Clinical Pathology .

Gross Lesions.

Light and Scanning Electron Microscopic
Changes . . . . . . . . . . .

Immunofluorescence .

Electron Microscopic Examination of Feces.

DISCUSSION.
SUMMARY .
BIBLIOGRAPHY.
VITA.

viii

 

Table

LIST OF TABLES

Table Page
1 Mean body weights (g 1 SD) for gnotobiotic
pigs infected with the OSU or SDSU strains

of porcine rotavirus . . . . . . . 69
2 Serum chemistry and hematocrit values for

gnotobiotic pigs 24 hours after inoculation
with the OSU or SDSU strains of porcine
rotavirus. . . . . . . . 71

3 Serum chemistry and hematocrit values for
gnotobiotic pigs 72 hours after inoculation
with the OSU or SDSU strains of porcine
rotavirus. . . . . . 72

4 Serum chemistry and hematocrit values for
gnotobiotic pigs 168 hours after inoculation
with the OSU or SDSU strains of porcine
rotavirus. . . . . . . . 73

5 Villous length (pm) in the small intestines
of gnotobiotic pigs infected with the OSU
or SDSU strains of porcine rotavirus . . . . 75

6 Crypt depths (um) in the small intestines of
gnotobiotic pigs infected with the OSU or
SDSU strains of porcine rotavirus. . . . . . 85

7 Light microscopic findings in the small
intestine of gnotobiotic pigs 24 hours after
inoculation with the OSU or SDSU strain of
porcine rotavirus. . . . . . . . 86

8 Light microscopic findings in the small
intestine of gnotobiotic pigs 72 hours after
inoculation with the OSU or SDSU strain of
porcine rotavirus. . . . . . . . . . . . . . 87

9 Light microscopic findings in the small
intestine of gnotobiotic pigs 168 hours after
inoculation with the OSU or SDSU strain of
porcine rotavirus. . . . . . . . . . . . . . 88

ix

 

 

 

 

Table
10

ll

12

Table
10

11

12

Page

Immunofluorescence in the small intestine
of gnotobiotic piglets infected with the
SDSU strain of porcine rotavirus . . . . . . 99

Immunofluorescence in the small intestine of
gnotobiotic piglets infected with the OSU
strain of porcine rotavirus. . . . . . . . 100

Results of direct electron microscopic
examination of feces or cecal content from

pigs infected with the SDSU or OSU strain

of porcine rotavirus . . . . . . . . . . 102

 

 

 

it...

 

 

Figur

 

\l

 

Figures

1

10

 

LIST OF FIGURES

Scanning electron micrograph of the
UJ of a pig 24 hours after inoculation
with the OSU strain of porcine rotavirus

Section of small intestine from the MI
of a pig 72 hours after inoculation
with the OSU strain of porcine rotavirus

Section of small intestine from the DU
of a pig 72 hours after inoculation
with the SDSU strain of porcine rotavirus.

Section of small intestine from the DU
of a pig 168 hours after inoculation
with the SDSU strain of porcine rotavirus.

Scanning electron micrograph of the DU of
a pig 168 hours after inoculation with the
SDSU strain of porcine rotavirus .

Section of small intestine from the DU of
a pig 168 hours after inoculation with the
OSU strain of porcine rotavirus.

Scanning electron micrograph of the DU of
a pig 168 hours after inoculation with the
OSU strain of porcine rotavirus.

Villous fusion in a section of small
intestine from the LJ of a pig 72 hours
after inoculation with the SDSU strain
of porcine rotavirus . . . . . .

Scanning electron micrograph of small
intestine from the LJ of a pig 168 hours
after inoculation with the SDSU strain
of porcine rotavirus

Villous fusion in a section of UJ from

a pig 168 hours after inoculation with
the OSU strain of porcine rotavirus.

xi

 

Page

77

78

79

80

81

82

83

89

9O

91

 

Figur

ll

 

 

 

 

12

l3

14

IS

Figures

11

12

l3

14

15

Most of the absorptive cells lining
villi in the LI of a control '

24 hours after inoculation with media
are vacuolated (+4) . . . .

Immature cuboidal to low columnar
absorptive cells line an atrophic
villus in the UJ of a pig 24 hours
after inoculation with the OSU strain
of porcine rotavirus.

Duodenum of a control pig killed at
post- -inocu1ation hour 24. .

Scanning electron micrograph of the
duodenum of a control pig at
post—inoculation hour 24. .

Scanning electron micrograph of the lower
ileum of a control pig at post- inoculation
hour 24 . . . .

Page

92

94

95

96

97

,3

 

 

 

 

 

 

 

 

 

 

 

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INTRODUCTION

Rotaviruses were first identified in the feces of a

 

healthy vervet monkey and in intestinal washings from
sheep and cattle at a slaughterhouse (Malherbe and
Strickland-Cholmley, 1967). However, it was not until
Mebus et a1 (1969) associated a rotavirus with diarrhea
in calves in Nebraska that the significance of these
viruses was recognized. Rotaviruses are now considered
important causes of neonatal gastroenteritis in many

mammalian and avian species (Mebus et al, 1971b; Bishop

 

et al, 1973; Bohl et a1, 1978; McNulty et al, 1978). In
children, rotavirus is the single most important
etiologic agent of acute infantile gastroenteritis
throughout the world and has been estimated to account
for one million diarrheal deaths annually (Vesikari et
al, 1983).

Initially, all viruses with rotavirus morphology
were thought to possess a common group antigen located on
the inner capsid layer of the virus (Woode et al, 1976a;
Yolken et al, 1978a). Recently, several rotaviruses
which are morphologically similar to but antigenically
distinct from the conventional rotaviruses have been

described. These viruses have been referred to as

 

 

parar
virus
(Snod
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typic
the p
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Serow
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1980

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pararotaviruses (Bohl et a1, 1982), rotavirus-like
viruses (Rodger et a1, 1982), and atypical rotaviruses
(Snodgrass et a1, 1984). To distinguish among these
isolates, Pedley et a1 (1983) proposed a scheme in which

typical and atypical rotaviruses are classified based on

 

the presence of distinctive group specific antigens into
groups designated A, B, C, and recently D and E (Pedley
et al, 1986).

Group A rotaviruses have been further classified
based on serotype and subgroup specific antigens.
Serotype specific antigens are located on the outer
capsid layer of the virus (Bridger, 1978), and are
detected by plaque reduction and fluorescent focus

neutralization assays (Sato et a1, 1982; Bohl et a1,

 

1984; Hoshino et a1, 1984). Subgroup specific antigens
are located on the inner capsid layer (Woode et al,
1976a) and are detected by enzyme—linked immunosorbent,
immune-adherence hemagglutination, and complement
fixation assays (Struker et a1, 1979; Zissis and Lambert,
1980). By using these assays, multiple serotypes and
subgroups of mammalian rotaviruses have been identified.
Recent reports have indicated differences in
clinical signs, viral antigen distribution, and lesion
severity for different rotavirus serotypes, subgroups,
and groups. An avian group A rotavirus, studied by
McNulty et a1 (1983) had a predilection for the duodenal

mucosa whereas a non—group A avian rotavirus replicated

 

 

 

 

 

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best in the mid-small intestine. In gnotobiotic pigs, a
group B porcine rotavirus caused mild clinical signs and
lesions (Theil et a1, 1985) whereas a group A porcine
rotavirus (Ohio State University strain) caused severe
diarrhea and extensive intestinal damage (Theil et a1,
1980). Four strains of rotavirus isolated from diarrheic
calves caused different amounts of Villous atrophy when
inoculated into ligated intestinal loops in colostrum
deprived calves (Carpio et al, 1981b). Differences in
the clinical symptoms associated with infection by two
subgroups of rotavirus in humans have been reported by
some investigators (Uhnoo and Svensson, 1986), but these
findings are in disagreement with the findings of others
(White et a1, 1984).

Two serotypes of porcine rotavirus have been
identified by plaque reduction neutralization tests,
cross protection studies in gnotobiotic piglets, and
electrophoresis of rotaviral double-stranded RNA (Bohl et
a1, 1984; Hoshino et a1, 1984). The purpose of the
present study was to determine if specific clinical
signs, mortality rates, virus shedding patterns, and

lesion distribution and severity were related to the

 

 

serotype of porcine rotavirus with which gnotobiotic pigs

were infected.

 

simi

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See]
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LITERATURE REVIEW

Classification

Rotaviruses derive their name from the Latin word
"rota" meaning wheel, which they resemble in appearance
(Flewett et a1, 1974). Initially, they were referred to
as "orbivirus", ”duovirus”, "reovirus-like agent" and
"infantile gastroenteritis virus" (Mebus et al, 1971b;
Bishop et a1, 1973; Kapikian et a1, 1975).

Rotaviruses, reoviruses, and orbiviruses have many
similarities. They are approximately the same size (65
to 75 nm), have double-shelled capsids, have similar
buoyant density in cesium chloride and possess a
double-stranded RNA genome (Palmer et a1, 1977).
Rotaviruses can be distinguished from orbiviruses and
reoviruses by the presence of 11 segments of the RNA
genome; orbiviruses and reoviruses possess only 10
segments (Palmer et a1, 1977). On the basis of these
similarities and differences, rotaviruses are classified
with reoviruses and orbiviruses within the family
reoviridae (Matthews, 1979).

Rotavirus Serotvpes and Subgroups
Most rotaviruses from mammalian and avian species

share a common antigen that is associated with the inner

 

 

 

 

 

 

 

capsi
antig
locat
rotat
adapt
and 1
devei
rota1
seve:
and .

I983

 

 

capsid layer (Woode et al, 1976a). Serotype specific
antigens are distinct from the common antigen and are
located on the outer capsid (Bridger, 1978). When
rotaviruses from several mammalian species became
adapted to replicate in cell culture, plaque reduction
and fluorescent focus neutralization assays were
developed for the antigenic comparison of different
rotaviruses. These assays led to the recognition of
several serotypes of rotaviruses from different mammalian
and avian species (Sato et a1, 1982; Murakami et a1,
1983; Bohl et a1, 1984; Hoshino et a1, 1984).

In addition to neutralization assays, rotaviruses
have been studied by enzyme-linked immunoassays,
complement fixation, immune—adherence hemagglutination,
and other techniques (Struker et a1, 1979; Zissis and
Lambert, 1980). The use of these assays led to much
confusion in the literature about the serologic
classification of rotaviruses because it was assumed that
the above techniques gave identical results to
conventional neutralization assays. It has been shown,
however, that antigens detected by enzyme immunoassays,
immune—adherence hemagglutination, and complement
fixation are distinct from those detected by plaque
reduction or fluorescent focus neutralization (Kapikian
et a1, 1981).

”Serotypes" as determined by neutralization tests

depend on differences in polypeptides located on the

 

 

outer capsi
enzyme-1mm
hmmne-adh
difference
et al, 19%
proposed
designati
"subgroup
differenc
fixation
et al, 1
A c
and dive
rotaviri
1984).
from se
Studied
antiger
basis t
neutra
antlse
IOtavj
human
serot-
an eq
serOt

hoshj

outer capsid whereas ”serotypes" as determined by
enzyme-immunoassays, complement fixation, and
immune-adherence hemagglutination are based on
differences in the major inner capsid polypeptide (Kalica
et a1, 1981). To avoid confusion, Kapikian et a1 (1981)
proposed that the term "serotype" be reserved for
designation of neutralization specificity, and the term
"subgroup" be used in place of serotype, to indicate
differences detected by enzyme immunoassays, complement
fixation, and immune-adherence hemagglutination (Kapikian
et a1, 1981).

A comprehensive study of the serotypic similarity
and diversity of human and other mammalian and avian
rotaviruses has been reported recently (Hoshino et a1,
1984). Sixteen different strains of rotavirus derived
from seven mammalian species and two avian species were
studied by plaque reduction neutralization. Seven
antigenically distinct serotypes were established on the
basis of a greater than 20-fold difference between
neutralizing titers of homologous and heterologous
antiserum. In this study, three strains of porcine
rotavirus (Gottfried, SB-lA and SB—Z), and one strain of
human rotavirus (St. Thomas No. 4) were of the same
serotype designated type 4. Porcine rotavirus (OSU) and
an equine rotavirus (H—l) made up a possible fifth
serotype. The serotype 4 rotaviruses as described by

Hoshino et a1 (1984) represent an example of shared

 

 

 

 

serotype
and thos
The
OSU stre
subgroup
Strain E
I, wher:
sungOU'
example
are of
specifi
Bc
rotavir
with dj
identi:
rossa
electr.
Ohio 8
as the
Strain
SerOty
rotavi
t0 se1
HoshiI

Et al

in thl

serotype specificity between rotaviruses of human origin
and those from other animals.

The Gottfried strain of porcine rotavirus and the
OSU strain of porcine rotavirus were found to have
subgroup II and subgroup I specificities, respectively.
Strain SB-Z of porcine rotavirus was found to be subgroup
I, whereas the Gottfried strain, as mentioned above, was
subgroup II. This pair of isolates represents the first
example of two rotaviruses from one animal species that
are of the same serotype, but differ in subgroup
specificity.

Bohl et a1 (1984) studied seven strains of
rotaviruses isolated from intestinal contents of piglets
with diarrhea. Two serotypes of porcine rotavirus were
identified by plaque reduction neutralization tests,
cross—protection studies on gnotobiotic piglets, and
electrophoresis of rotaviral double-stranded RNA. The
Ohio State University (OSU) porcine isolate was suggested
as the prototype serotype 1 rotavirus. The Gottfried (G)
strain porcine rotavirus was suggested as the prototype
serotype 2 rotavirus. Serotypes l and 2 porcine
rotaviruses as described by Bohl et a1 (1984) correspond
to serotypes 5 and 4, respectively, as described by
Hoshino et a1 (1984). The classification system of Bohl
et a1 (1984) for porcine rotavirus strains will be used

in the remainder of this dissertation.

 

 

It
systems
based c
polyacr
(1981)
electrt
segmen
are dm
7-9 as
Differ
within
la, 11
classi
conduc

I
rotavi
subgrc
rOtavi
t0 sul
10 an<
Subgn
apply
Elecp
Of my
disti

1982,

Rotavirus Electropherotypes

 

In addition to subgroup and serotype classification
systems, there has been an effort to classify rotaviruses
based on the mobility of their RNA segments during
polyacrylamide gel electrophoresis. Lourenco et a1
(1981) have proposed a scheme for rotaviral
electropherotype classification in which the 11 RNA
segments are divided into four groups. RNA segments 1—4
are denoted as group I, bands 5 and 6 as group II, bands
7-9 as group III, and bands 10 and 11 as group IV.
Differences in the relative migration of RNA segments
within a group are designated by a small letter (example
Ia, IIa, IIIb, IVc). This system allows rapid
classification of RNA patterns when standard methods of
conducting the test are employed.

The migration of RNA segments 10 and 11 of human
rotavirus strains has been shown to correlate with
subgroup specificity (Kalica et al, 1981). Human
rotaviruses with slow migrating segments 10 and 11 belong
to subgroup I, whereas those with fast migrating segments
10 and 11 belong to subgroup II. Such a correlation of
subgroup specificity with RNA electropherotype does not
apply to animal rotaviruses. Comparison of RNA
electropherotypes has proved useful in the identification
of new groups of rotaviruses that are antigenically
distinct from the classical rotaviruses (Bohl et a1,

1982; Bridger et a1, 1982; Snodgrass et a1, 1984).

 

Classii

A
rotavi:
scheme
Graham
serolo
crypto
distin
source
need u
that r
at leg

or 9 a

from
Commc
Capsi
a1
micrt

indl.‘

Classification Systems

 

A unified serological classification for
rotaviruses, similar to the universal classification
scheme for influenza virus, has not yet been achieved.
Graham and Estes (1985) recently have proposed a
serologic classification system which utilizes a
cryptogram to convey the information necessary to
distinguish and identify new rotavirus isolates from any
source. However, this classification system may already
need modification. Greenberg et a1 (1983) demonstrated
that neutralization of a simian rotavirus is mediated by
at least two gene products of rotavirus; those of gene 8
or 9 and gene 4. Rotavirus genes 8 or 9 code for the
major neutralization antigen (VP7) whereas gene 4 codes
for a minor neutralization antigen (VP3). Hoshino et al
(1984) suggested that future systems of rotaviral
classification may need to indicate two, instead of one,
distinct serotypic specificities for each isolate of
rotavirus.

Atypical Rotaviruses

 

Initially, all viruses with rotavirus morphology
from different animal species were thought to possess a
common group antigen or antigens located in the inner
capsid layer of the virus (Woode et al, 1976a; Yolken et
a1, 1978). In 1980, Saif et a1 detected, by electron
microscopy, virus particles that were morphologically

indistinguishable from rotaviruses but which did not

 

 

cross-1
rotavii
infecti
particl
seroty]
finding
type.
this ”i
(Bohl
R
which
antige
referr
rotavi
rotavi
A
became
antlge
t0 eac
a Clas
Virus;
SChEme
Class:
subcli
SPEci;
Additj

SpeCi:

10

cross—react with antisera to porcine, bovine, or human
rotaviruses or to reovirus type 3. Furthermore, previous
infection of gnotobiotic pigs with the rotavirus-like
particles failed to protect them from challenge with
serotype 1 porcine rotavirus (strain OSU). These
findings suggested the presence of a ”new" rotavirus
type. The pathogenesis of rotaviral diarrhea caused by
this "new rotavirus” was detailed in a subsequent report
(Bohl et a1, 1982).

Recently, several additional atypical rotaviruses
which lack the conventional rotavirus group specific
antigen have been described. These viruses have been
referred to as pararotaviruses (Bohl et a1, 1982),
rotavirus-like viruses (Rodger et a1, 1982), and atypical
rotaviruses (Snodgrass et a1, 1984).

As these viruses were studied in greater detail, it
became evident that several atypical rotaviruses were
antigenically distinct from and biochemically unrelated
to each other. Therefore, it became necessary to develop
a classification scheme to distinguish among these
viruses. Pedley et a1 (1983) proposed a classification
scheme in which typical and atypical rotaviruses were
classified in the genus as rotaviruses but further
subclassified, based on the presence of distinctive group
specific antigens into groups designated A, B, and C.
Additional atypical rotaviruses that do not possess group

specific antigen characteristic of groups A, B, or C have

 

 

been 1
groups
Pedley

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rotav:
for g
enzym
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antis
is re

rotav

also

(Pedl
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Where

doubi

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al
idem
f8ce
Davi

in t

 

11

been identified indicating the need for additional
groups (Snodgrass et a1, 1984; Vonderfecht et a1, 1984;
Pedley et a1, 1986).

Detection of samples which are positive for
rotavirus particles by electron microscopy but negative
for group A rotavirus in fluorescent antibody or
enzyme—linked immunoassay tests is suggestive of a
nongroup A rotavirus. But further testing, using
antisera which reacts with only the subgroup in question,
is required to confirm the presence of atypical
rotaviruses (Pedley et a1, 1983).

Electrophoresis of viral RNA in polyacrylamide gel
also is helpful in distinguishing rotavirus groups
(Pedley et a1, 1983). The major difference in the RNA
electrophoretic profiles of groups A, B, and C is that
group A rotavirus bands 7, 8, and 9 migrate as a triplet,
whereas in groups B and C, this triplet is replaced by a
doublet.

At present, the importance of atypical rotaviruses
in diarrheal diseases of mammalian and avian species is
unknown. It appears that the prevalence of atypical
rotaviruses in cattle and swine is quite low (Chasey et
a1, 1984; Theil et a1, 1985). Atypical rotavirus are
identified more frequently than group A rotaviruses in
feces from diarrheic turkey poults and lambs (Chasey and

Davies, 1984; Saif et a1, 1985) so they are of importance

in those species.

 

 

 

I
of sex
and Sc
agree:
does 1
gener.
types
Negat
viewe
smoot
”fuzz
almos
IOtav
diame
C8811
aPPT<
Chlo:
al, j

been
Rota

solv

Physicochemical Properties

 

The structure of the rotavirion has been the subject
of several investigations (Martin et a1, 1975; Stammond
and Schoub, 1977). Although there is no general

agreement about the number of capsomeres, the rotavirion

 

does have a definite capsomere structure. It is
generally agreed that rotaviruses have two main particle
types: single— and double—shelled (see Morphogenesis).
Negatively stained double—shelled virus particles, as
viewed with the transmission electron microscope, have a
smooth outer layer. In contrast, the orbiviruses have a
"fuzzy" outer layer, and the outer capsid of reovirus is
almost featureless (Palmer et a1, 1977). Double-shelled
rotavirus particles are approximately 70 to 75 nm in
diameter and have a buoyant density of 1.36 g/ml in
cesium chloride. Single—shelled particles are
approximately 65 nm in diameter with a density in cesium
chloride of 1.38 g/ml (Bridger and Woode, 1976; Rodger et
a1, 1975).

The physicochemical properties of rotaviruses have
been discussed in detail (Flewett and Wood, 1978a).
Rotaviruses are stable to non—ionic detergents, lipid
solvents, heat, extremes of pH and high salt
concentrations. Such stability is central to the
survival of rotaviruses in the intestinal tract. Simian
rotavirus SA—ll is rapidly inactivated when heated at

50°C in the presence of 2 M MgClZ, but is stabilized by

 

heatin
also s
1981).
rotavi
from t
rendei

l
disin
and c
(752)
reduc
organ
wides
(I977
conte
disit
rota;

Cond:

habi
adap
SA—l
asyu
res;
rOte

Stic

13

heating in 2 M MgSO4 (Estes et a1, 1979). Calcium ions
also stabilize rotavirus infectivity (Shirley et a1,
1981). Chelators such as EDTA may be used to destabilize
rotavirions by removing divalent cations such as calcium
from the outer coat of the virus particle, thereby

rendering them more permeable (Cohen, 1977).

 

Rotaviruses are resistant to many chemical
disinfectants and antiseptics commonly used in research
and chemical laboratories (Sattar et a1, 1983). Ethanol
(752) and formaldehyde (3.7%) are somewhat effective in
reducing the amount of virus even in the presence of
organic matter, but these chemicals are impractical for
widespread use in a farm environment. Snodgrass et a1

(1977) found a rotavirus purified from the intestinal

 

contents of a lamb resistant to iodine-based
disinfectants. Because of the resistant nature of
rotaviruses, it is unlikely that disinfection under farm
conditions will be completely successful.

Cultivation in Cell Culture

 

Rotaviruses grow extremely well in their natural
habitat, the intestinal tract, but have been difficult to
adapt to cell culture. The first rotaviruses cultured,
SA—ll and the ”0" agent, were isolated from an
asymptomatic vervet monkey and slaughterhouse waste,
respectively. These viruses were not recognized as
rotaviruses until much later (Malherbe and

Stickland—Cholmley, 1967). Interest in the cultivation

 

 

of rot
virus

SUCCEE

19713}

the

isol
hung
Cont
Feh‘

to}

 

14

of rotaviruses intensified when neonatal calf diarrhea
virus (NCDV), a cause of bovine calf diarrhea, was
successfully isolated in cell culture (Mebus et al,
1971a).

Excluding bovine rotavirus, serial propagation of
these viruses in cell culture proved difficult (Woode et
al, 1976b; McNulty et al, 1976a). Porcine rotavirus was
successfully passaged in cell culture by treatment of
viral suspensions with pancreatin prior to inoculation
onto cell monolayers (Theil et al, 1977). In a
subsequent paper, Theil et a1 (1980) reported that
rotavirus infectivity for porcine kidney cells (PK—15)
was enhanced by incorporation of pancreatic
endopeptidases into the cell culture maintenance medium.
These important findings were widely applied and led to
the successful cultivation of rotaviruses derived from
many mammalian and avian species (McNulty et a1, 1979;
Wyatt et a1, 1980; Tajima et a1, 1984; Makabe et a1,
1985). It has not proved possible, however, to culture
all rotaviruses by these methods.

Rotaviruses vary in their requirement for trypsin in
the culture medium. The SA-ll and bovine rotavirus
isolates (NCDV) were cultured without trypsin whereas
human rotavirus has only been isolated with culture media
containing trypsin (Wyatt et al, 1980; Sato et a1, 1981).
Feline, canine, and simian rotavirus strains were found

to be less dependent upon trypsin than human, bovine,

 

 

 

 

 

porcin
1981).
not re
strait
cell <

virus

 

15

porcine, chicken, and turkey rotaviruses (Hoshino et a1,
1981). Although trypsin treatment of bovine rotavirus is
not required for the isolation of some bovine rotavirus
strains, it can effectively enhance viral replication in
cell culture, thus producing high titer stocks of the
virus (Clark et a1, 1979).

The molecular basis for the proteolytic enhancement
of rotavirus infectivity has been studied (Espejo et a1,
1981; Estes et al 1981). Trypsin cleaves a major
non—glycosylated polypeptide of the outer capsid (VP3)
yielding two polypeptides with molecular weights of
approximately 60,000 and 28,000. It is proposed that the
latter polypeptides contain hydrophobic regions that,
once exposed, aid in viral penetration into the host
cell.

Primary kidney cell cultures from a variety of
species have been the cell—type most frequently used to
culture rotaviruses. At present, a number of cell lines
are being used successfully, including MDBK (Madin-Darby
bovine kidney), PK-lS (porcine kidney), BSC—l (green
monkey kidney), LLC—MK2 (Rhesus monkey kidney), CV-l
(green monkey kidney), and MA—104 (Rhesus monkey kidney)
(Wyatt and James, 1982). Primary cells appear to support
virus growth more efficiently than continuous cell lines
and therefore are more sensitive for rotavirus isolation
(Ward et a1, 1984).

Rotavirus-specific antibodies are prevalent in the

 

 

 

 

 

 

sera c
1981),
use of
Rotavj
serum
inter:
or an‘
part,

in se

cultu
tryps
diarr
inocr
(Welt
deter

hours

VaCUt
incl

cres

smal
The
of C
bovi

iDOC

l6

sera of domestic and laboratory animals (Sato et a1,
1981), so for cultivation investigators must avoid the
use of adult animal sera containing rotavirus antibody.
Rotavirus antibodies have been detected in fetal bovine
serum and in purified serum albumin preparations and may
interfere with diagnostic assays for rotavirus antigens
or antibodies (Offit et a1, 1984). This finding, in
part, explains why human rotavirus has been isolated only
in serum-free media.

Cytopathic effect (CPE) caused by rotavirus in cell
culture depends on the cell-type used, the presence of
trypsin, and the strain of rotavirus. Neonatal calf
diarrhea virus produces an easily discernible CPE when
inoculated at high concentrations onto cell cultures
(Welch and Twiehaus, 1973). Cytopathic effect is
detectable by 24 hours after inoculation, and by 48
hours, some areas of the cell culture may be devoid of
cells. Infected cells may appear granular and finely
vacuolated, and contain single or multiple eosinophilic
inclusions. Some cells become thin, elongated, and
crescent-shaped and remain attached to the monolayer only
by a single process. Those that completely detach are
small, rounded or elongated and have pyknotic nuclei.

The cytopathic effect has been produced in a wide variety
of cell culture systems (Fernelius et a1, 1972). Another
bovine rotavirus strain produces similar CPE when

inoculated onto MDBK kidney cells, except a larger

 

 

 

 

 

portior

inclusi
II
rotavi1
culture
produc:
the de‘
Plaque
medium
other

inocul
a plaq
for as
widely
rotavi

I984),

l7

portion of the monolayer is destroyed, and cytoplasmic
inclusions are uncommon (McNulty et a1, 1977).

In contrast to neonatal calf diarrhea virus, porcine
rotavirus strains produce minimal to moderate CPE in cell
culture unless the strain has been passaged several times
13 yipgg. The detection of rotavirus strains that
produce minimal CPE in cell culture has been improved by
the development of plaque assays (Bohl et a1, 1984).
Plaque assays utilize pancreatin or trypsin in an overlay
medium. In this system, porcine rotavirus, as well as
other rotavirus strains, will produce plaques after
inoculation onto MA-104 cells. By use of this technique,
a plaque reduction neutralization test has been developed
for assaying neutralizing antibody and has been used
widely for the detection of antigenic diversity among
rotavirus strains (Bohl et a1, 1984; Hoshino et a1,
1984).

Morphogenesis

The morphogenesis of rotavirus has been studied by
electron microscopic examination of cell cultures
(Chasey, 1977; Saif et a1, 1978; Quan and Doane, 1983;
Suzuki et a1, 1984) and intestinal epithelial cells
(Chasey, 1977; Pearson and McNulty, 1979). The
morphogenetic stages of rotaviruses isolated from
different animal species is similar both in giggg and 13
yiyg. Because of these similarities as well as the early

unsuccessful attempts to isolate human rotavirus strains,

 

 

 

tissue-cultua
model system
(Quan and Do
Study 0
culture reve
virions ente
(b) in some
into lysosou
found first
as 6 hours a
cytoplasmic
viroplasm b'
these part1
enveIOpe, w
thus formin
lysis.
DeSpit
morPhogene:
the replic.
(1985) hay
104 cells
PTEtreatme
CYtoplasm
dissolutic
abSence of
Virions we

V.
lrug att.‘

18

tissue-culture-adapted simian rotavirus SA-ll became a
model system for the study of rotavirus morphogenesis
(Quan and Doane, 1983).

Study of the morphogenesis of SA-ll virus in cell
culture revealed the following sequence of events: (a)
virions enter the cell by receptor—mediated endocytosis;
(b) in some instances, the virus is taken up directly
into lysosomes, but in others, the virus particles are
found first in endosomes and then lysosomes; (c) as early
as 6 hours after infection, immature virus can be seen in
cytoplasmic aggregates of viroplasm and at the edge of
viroplasm budding into rough endoplasmic reticulum; (d)
these particles become enveloped during budding, but the
envelope, which is not required for infectivity, is lost,
thus forming a mature virion which is released by cell
lysis.

Despite the many reports about rotavirus
morphogenesis, some aspects concerning early stages of
the replication cycle are controversial. Suzuki et a1
(1985) have found that the mode of viral entry into MA
104 cells is affected by trypsin. With trypsin
pretreatment, viral nucleoids passed directly into the
cytoplasm within 5 minutes after inoculation through
dissolution of viral capsid and cell membrane. In the
absence of trypsin, phagocytosis occurred in which
virions were sequestered into lysosomes 20 minutes after

virus attachment to cell membranes. After sequestration,

 

 

 

 

 

 

uncoating of
but it did n
Therefore, S
phagocytosis
The cap
least 5 proi
1W2, VP6) m;
(VP3 and VP
Estes et a1
identified
inclusions
believed ti
assembled f
Outer sheli
Process (P.
multivesic
morPhogene
(SuzUki 8t
Multi
identifiec
Petrie et
SiHSIE-sh.
cells, bu
Particles
NOQde, 19
CYtOplaSK

a1. 1981,

l9

uncoating of rotavirus virions within lysosomes was seen,
but it did not result in the release of the viral genome.
Therefore, Suzuki et a1 (1985) discounted the theory that
phagocytosis is related to viral replication.

The capsid of simian rotavirus SA-ll consists of at
least 5 protein classes. Three of these proteins (VPl,
VP2, VP6) make up the inner layer, whereas the other two
(VP3 and VP7) form the outer layer (Espejo et a1, 1981;
Estes et a1, 1981). Immunocytochemistry studies have
identified VP2 and VP6 associated with viroplasmic
inclusions (Petrie et a1, 1982). Therefore, it is
believed that the core and inner capsid layer are
assembled in the viroplasmic inclusions and that the
outer shell glycoprotein is added during the budding
process (Petrie et a1, 1982). Tubules, fibrils, and
multivesicular bodies have been associated with rotavirus
morphogenesis, but their origin and function are unknown
(Suzuki et a1, 1984).

Multiple rotaviral particle types have been
identified (Saif et a1, 1978; Pearson and McNulty, 1979,
Petrie et al, 1981). Double—shelled particles and
single—shelled particles are released from infected
cells, but only the smooth double—shelled rotavirus
particles have been found to be infectious (Bridger and
Woode, 1976). Two additional particle types in
cytoplasmic organelles have been described by Petrie et

a1, 1981. The first is a subviral particle which is the

 

 

 

 

 

 

uncoated vir
enveloped pa
Enveloped pe
released frc
in negative?
examined wii
(1972) did .
negatively

monkey, but

contaminati

Serolc
antibodies
mammalian 2
et al, 197%
Bohl et 31
samples an
serologica
IOtaviruS.
also has b
31, 1984;
antibody t
common in
1958)» bin
pIEValenC,
Ohio SWin

indiCate

 

 

20

uncoated virion seen within lysosomes. The second is an
enveloped particle seen in the endoplasmic reticulum.
Enveloped particles lose their envelopes prior to being
released from the endoplasmic reticulum so are not found
in negatively stained preparations of fecal material when
examined with an electron microscope. E13 and Lecatsas
(1972) did observe enveloped virus particles in
negatively stained preparations from a healthy vervet
monkey, but this is unusual and may have represented
contamination by another virus.
Epidemiology

Serologic studies on the prevalence of rotavirus
antibodies indicate a widespread distribution in many
mammalian and avian species (Petri et a1, 1978; McNulty
et a1, 1978; Bohl et a1, 1984; Bridger and Brown, 1985).
Bohl et a1 (1984) found that 94% of 274 porcine serum
samples and 1002 of 75 herds in the United States were
serologically positive to the OSU strain of porcine
rotavirus. A high prevalence of antibodies to rotavirus
also has been found in swine in other countries (Utera et
a1, 1984; Bridger and Brown, 1985). The prevalence of
antibody to the atypical rotaviruses (groups B and C) is
common in pigs in the United Kingdom.(Bridger and Brown,
1958), but Theil et a1 (1985) found a much lower
prevalence (23%) of antibody to group B rotaviruses in
Ohio swine sera. Results of serologic surveys also

indicate that some serotypes of human rotavirus are more

 

 

 

 

prevalent th
least two se
distinguishe
comparing tl
rotavirus t}

There a
many infect:
is more pre‘
et a1, 1976
is not know
suggested a
the and Shi
feces on er
at high or
humidity,
rotavirus :
DEER Sugge:
be a refle-
numbers,

TO as
transmissi
Comparison
diffErent

infectiOn

21

prevalent than_others (Yolken et a1, 1978). Although at
least two serotypes of porcine rotavirus have been
distinguished (Bohl et a1, 1984), epidemiologic surveys
comparing the prevalence of infection by porcine
rotavirus types 1 and 2 have not been reported.

There are seasonal variations in the prevalence of
many infectious diseases. Rotaviral diarrhea in people
is more prevalent in the fall and winter months (Kapikian
et a1, 1976). Although the cause of the seasonal pattern
is not known, low relative humidity in the homes has been
suggested as a factor influencing rotavirus survival.

Moe and Shirley (1982) found that rotaviruses in

feces on environmental surfaces survive for days or weeks
at high or low relative humidity, but not intermediate
humidity. A seasonal increase in the prevalence of
rotavirus infections in animals during the winter has
been suggested (Woode and Bridger, 1975a), but this may
be a reflection of seasonal variation in livestock
numbers.

To assess the importance of interspecies
transmission of rotaviruses and to make antigenic
comparisons between rotavirus strains isolated from
different species, numerous experimental cross—species
infection studies have been done (Mebus et a1, 1977;
Tzipori et a1, 1980; Bridger and Brown, 1984). Some
strains of rotavirus isolated from children with diarrhea

readily infected pigs (Middleton et a1, 1975). Woode and

 

 

 

 

Bridger (197
infection by
investigate:
rotavirus i:
Many 0
infection b‘
recognition
discrepanci
human or ot
animals may
in these e)
have shown
infect pig:
porcine ro'
PP-l) that
rotavirus
relationsh
because, i
r0tavirus
CTOSS-neut
81.1ng
of rotavi]
et 81 (19:
are SUSCe‘
rotavirus
Alth

aanSt ev

22

Bridger (1975a) reported that calves were refractory to
infection by a strain of human rotavirus, but other
investigators successfully infected calves with a
rotavirus isolated from people (Mebus et a1, 1977).

Many of the investigations of cross-species
infection by rotaviruses were completed prior to the
recognition of multiple rotavirus serotypes. Thus,
discrepancies in the literature concerning the ability of
human or other animal rotaviruses to infect experimental
animals may be related to the use of different serotypes
in these experiments. Bridger and Brown (1984) recently
have shown that the ability of bovine rotaviruses to
infect pigs depends on the antigenic relationship to
porcine rotaviruses. Only a bovine rotavirus (strain
PP-l) that was antigenically closely related to porcine
rotavirus caused disease. However, a close antigenic
relationship does not imply similarities in Virulence
because, in the same report, another strain of bovine
rotavirus (CP—l) which was indistinguishable from PP-l by
cross-neutralization tests, was not pathogenic for pigs.

Subgroup characteristics also influence the ability
of rotaviruses to infect atypical host species. Zissis
et a1 (1983) have shown that colostrum-deprived piglets
are susceptible to several human—origin strains of
rotavirus, except those belonging to subgroup I.

Although rotaviruses are widely distributed in

almost every species, there is no evidence that under

 

 

 

 

natural cone
Cross-specie
has enabled
models for ‘

Simult.
electropher'
detected (8
Bohl et a1,
such "mixed
known prOpe
occur. Th1
variation 5
continued p
antigenic \
rotaviruses
(Gething 9
among rota‘

reported r

The c
described
gnotobioti

23

natural conditions, animal rotaviruses infect people.

Cross-species infection is important, however, because it

has enabled the development of many laboratory animal

models for the study of rotavirus infections.
Simultaneous infection by more than one

electropherotype of human or porcine rotavirus has been

 

detected (Spencer et a1, 1983; Rodriguez et a1, 1983;
Bohl et a1, 1984). It has been postulated that during
such "mixed” infections, genetic reasssortment, a well
known property of rotaviruses (Greenberg et a1, 1981) may
occur. This genetic reassortment could lead to antigenic
variation among rotaviruses, thus enabling their
continued persistence within a population. Constant
antigenic variation by influenza viruses which, like
rotaviruses, have a segmented genome, is well recognized

I!

(Gething et a1 1980), and evidence of antigenic drift"
among rotavirus isolates from human neonates has been
reported recently (Coulson et a1, 1985).
Clinical Signs

The clinical signs of rotavirus infection have been
described in conventional, colostrum~deprived, and
gnotobiotic animals of many species (Theil et a1, 1978;
McAdaragh et a1, 1980; Johnson et a1, 1983; McNulty et
a1, 1983; Gillespie et a1, 1984). The first clinical
signs, anorexia and reluctance to move, are seen in
gnotobiotic piglets within 12—36 hours after inoculation

(Crouch and Wood, 1978; Theil et a1, 1978). Similar

 

 

incdbation 1
species inc:
is usually :
which time '
Vomiti
has been re
McAdaragh e
1980). Var
used to exp
the discrep
vomiting 11
been repor-
subgroup o
isolate ca
1 rotaviru
81, l978b)
that clini
severe in
I rOtavin
difference
among Chi

Ther
mucosa is

1977),

24

incubation periods have been reported in several other
species including people (Yolken et al, 1978b). Anorexia
is usually resolved by 72 hours after inoculation at
which time piglets resume eating.

Vomiting by piglets infected with porcine rotavirus
has been reported inconsistently (Theil et a1, 1978;
McAdaragh et a1, 1980; Torres-Medina and Underdahl,
1980). Variations in the serotypes of porcine rotavirus
used to experimentally infect the piglets might explain
the discrepancies in reports of vomiting. For example,
vomiting in people with rotaviral gastroenteritis has
been reported to be influenced by the serotype or
subgroup of rotavirus involved. A Type 2 human rotavirus
isolate caused vomiting in 9 of 16 patients whereas Type
1 rotavirus caused vomiting in only 3 of 11 (Yolken et
a1, l978b). Uhnoo and Svensson (1986) recently reported
that clinical symptoms (vomiting and diarrhea) were more
severe in people infected with subgroup II than subgroup
I rotaviruses. However, White et a1 (1984) found no
differences in the occurrence of fever or vomiting
among children shedding different subgroups of rotavirus.

There has been no clear evidence that the gastric
mucosa is infected by rotavirus (Pearson and McNulty,
1977). Apparently, injury to the small bowel mucosa is
sufficient to initiate a reflex resulting in vomiting.
Vomiting is usually not a feature of field cases of

porcine rotavirus infection (Bohl et al, 1978) a finding

 

 

that is some
from transm.
seen in rot
1977) but h
in human in
Rotavi
acute loss
days follow
inoculatior
expected we
Mortai
variable.
contents 0;
exPeriment
did not co
(Woode et
rotavirus
reported (
affected b
management
(1978) for
Conditions
increased
p188. It
Well fed,
Subc

(Cr00ch a.

that is sometimes useful in differentiating this disease
from transmissible gastroenteritis. Fever is usually not
seen in rotaviral infection in pigs (Pearson and McNulty,
1977) but has been associated with rotaviral infections
in human infants and young children (Carr et al, 1976).

Rotavirus infection in gnotobiotic pigs may cause
acute loss of body weight (lo-40%) over the first 2—3
days followed by rapid recovery. At 5—7 days after
inoculation, pigs may still be below (20-432) their
expected weight (Woode, 1979).

Mortality rates caused by porcine rotavirus are
variable. When gnotobiotic piglets were inoculated with
contents of the same vial of porcine rotavirus in two
experiments, mortality rates varied from 0-13Z and this
did not correlate with the dose of Virus inoculated
(Woode et al, 1976b). In field cases of porcine
rotavirus infection, mortality rates of 7-152 were
reported (Bohl et al, 1978). Mortality rates are
affected by environmental, nutritional, and other
management factors. For example, Crouch and Woode
(1978) found that, under natural or experimental
conditions, a drop of 10 to 20°C in ambient temperature
increased the mortality associated with the disease in
pigs. It appears that if pigs are kept warm, dry, and
well fed, mortality can be reduced.

Subclinical infections by rotavirus are common

(Crouch and Woode, 1978; Banatvala et al, 1978), and the

 

 

 

severity of
infections a
part, age-de
1002 mortali
lOZ mortalit
age. The c]
also are age
age do not <
(Sheridan e‘
also occur
cattle (Cro
some instan
gastroenter
instances c
1979).

Pigs l
disease cat
infection.
exPlain th
foufid youn
adults bee
e16€trolyt
not Younge
logs from
absorptior
include t1

intestine

26

severity of clinical signs in experimental rotavirus
infections are quite variable. This variation is, in
part, age-dependent. Woode et al (1976b) noted up to
100% mortality in piglets infected at 0-4 days of age,
10% mortality at 5-7 days, and no mortality at 28 days of
age. The clinical signs of rotavirus infection in mice
also are age—dependent. Mice greater than 10—14 days of
age do not develop clinical signs of rotavirus infection
(Sheridan et al, 1983). Subclinical rotavirus infections
also occur in adult swine (Benfield et a1, 1982) and
cattle (Crouch and Acres, 1984). Human rotaviruses, in
some instances, may affect adults causing severe
gastroenteritis (Echeverria et a1, 1983), but in other
instances only mild infections occur (Wenmann et al,
1979).

Pigs have a remarkable age—resistance to clinical
disease caused by transmissible gastroenteritis
infection. A number of explanations have been offered to
explain this age-resistance. Cornelius et a1 (1968)
found young pigs more vulnerable to dehydration than
adults because of changes in body water content and renal
electrolyte regulation with age. The colon of older, but
not younger, pigs can compensate for the increased fluid
loss from the small intestine by increased fluid
absorption (Argenzio et a1, 1984). Other explanations
include the greater involvement of the upper small

intestine and the slower proliferation of the intestinal

 

 

 

crypts resul
aborptive ep
One or more
age-resistar
age-depender
difference :
enterocytes
Riepenhoff—'
receptors 0'
than in mic
does not co
susceptibil
age are ref
infected, 2
Similar to
diarrhea (1
mice to re:
ability of
intestinal
Rotav
and Colost
1981; Bohl
nursing pi
of the lac
with trans

U
ilaltErmanr

 

27

crypts resulting in slower replacement of damaged
aborptive epithelial cells in younger pigs (Moon, 1971).
One or more of these explanations may apply to the
age-resistance shown by rotavirus infection. The
age-dependence of rotavirus infection may also reflect a
difference in the number of virus—specific receptors on
enterocytes of different maturity (Wolf et al, 1981).
Riepenhoff-Talty et al (1982a) found fewer rotavirus
receptors on enterocytes in the intestine of adult mice
than in mice less than 11 days of age. However, this
does not completely explain the difference in age
susceptibility. Although mice greater than 14 days of
age are refractory to clinical disease, they are
infected, and the extent and distribution of antigen is
similar to that seen in 7-day—old mice with rotaviral
diarrhea (Eydelloth et al, 1984). Thus, the ability of
mice to resist disease is not simply related to the
ability of the virus to infect certain portions of the
intestinal tract.

Rotavirus antibody is present in the serum, milk,

 

and colostrum of most, if not all, adult sows (Corthier,
1981; Bohl et a1, 1984). Thus the clinical signs in
nursing piglets will be influenced by the immune status
of the lactating sow, a situation similar to that found
with transmissible gastroenteritis (Hooper and
Haltermann, 1966). As expected, removal of nursing

piglets from the influence of "lactogenic" immunity, such

 

 

as occurs a

(Woode and

Serum
rotaviral d
(Mouwen et
al, l977).
children hc
found isotc
acidosis, e
the specif:
also was re
study of 2'
caused by 1
acidosis a
(Tallet et
Of these c
hypernatre
Serum then
increased
These labc
vomiting 6
illness it

Some
Piglets We
as the Cam

eff
‘veCt Of

 

28

as occurs at weaning, increases the severity of diarrhea
(Woode and Bohl, 1981).

Clinical Pathology

 

Serum biochemical alterations associated with
rotaviral diarrhea have not been studied in great detail
(Mouwen et al, 1972; Tallett et al, 1977; Rodriguez et
a1, 1977). Rodriguez et a1 (1977), in a study of 72
children hospitalized with rotavirus gastroenteritis,
found isotonic dehydration, compensated metabolic
acidosis, and elevations in blood urea nitrogen (BUN) and
the specific gravity of urine. Hypertonic dehydration
also was reported, although much less commonly. In a
study of 27 children hospitalized with acute diarrhea
caused by rotavirus, most had mild compensated metabolic
acidosis associated with decreased plasma bicarbonate
(Tallet et al, 1977). Serum sodium concentration in most
of these children was normal, but in 6 instances mild
hypernatremia was detected. Additional alterations in
serum chemistry included mildly decreased potassium,
increased chloride, and elevated, normal, or reduced BUN.
These laboratory findings reflect the high frequency of
vomiting and dehydration associated with rotaviral
illness in people.

Some of the biochemical aspects of "white scours" in
piglets were investigated before rotavirus was recognized
as the cause of this condition (Mouwen et al, 1972). The

effect of "white scours” on serum lipids indicated

 

 

 

increased n6
phospholipic
alterations
The pa'
but more se‘
serum bioch
studied mor
Cornelius e
2-day-old p
that became
and total I
fluids ass<
et al, l98.
protein se
(Cornelius
those repo
serum bioc
Ectabolic
hypoglycen
(Cornelius
a1 (1984)
that may 1
mOrtality
0r Older
from 6 to
rOtaviral

\7

29

increased neutral fat and decreased total cholesterol and
phospholipid. These investigators noted that serum lipid
alterations resembled those found in sprue in people.

The pathophysiology of TGE of swine is similar to,
but more severe than that of rotaviral enteritis. The
serum biochemical changes associated with TGE have been
studied more extensively (Reber and Whitehair, 1955;
Cornelius et a1, 1968; Drolet et al, 1984). When
2-day-old piglets were challenged with TGE virus, those
that became moribund had increased packed-cell—volumes
and total proteins which were attributed to the loss of
fluids associated with vomiting and dehydration (Drolet
et a1, 1984). Elevations in packed—cell—volume and total
protein secondary to TGE have been reported by others
(Cornelius et al, 1968), but are in disagreement with
those reported by Reber and Whitehair (1955). Additional
serum biochemical changes associated with TGE were
metabolic acidosis, increased BUN, elevated chloride,
hypoglycemia, and unchanged sodium and potassium
(Cornelius et al, 1968; Drolet et al, 1984). Drolet et
al (1984) considered hypoglycemia a major feature of TGE
that may be responsible for the high rate of neonatal
mortality with that disease. Piglets 5 to 6 days of age
or older are refractory to hypoglycemia even if fasted
from 6 to 7 days (Sampson et al, 1942). Because
rotaviral diarrhea is less common than TGE in piglets

younger than 7 days of age, hypoglycemia may not be as

 

 

 

 

important it

Rotavi'
the Villous
membrane-b0
sodium-pota
al, l984).
accompaniec
decreased e
l984).

The 3.
activities
the strain
example, D
lO-day-olc
had decree
“PPEI jEjl
infected 1
sodium-p0.
ileum Onl-

Loss
to nutrie
with Porc
Stool Osn
m0STU/lite
Of the o:

l

 

30

important in rotaviral infections.
Pathophysiology

Rotavirus infection in pigs causes desquamation of
the Villous epithelial cells resulting in loss of
membrane-bound enzymes such as lactose, sucrose, and
sodium-potassium ATPase (Davidson et al, 1977; Graham et
al, 1984). Loss of microvillus-associated enzymes is
accompanied by reduced transport of 3-0—methyglucose and
decreased absorption of sodium and water (Graham et al,
1984).

The segments of intestine in which altered enzyme
activities are detected may, in part, be influenced by
the strain of rotavirus used to infect the piglets. For
example, Davidson et a1 (1977) found that 8- to
lO-day-old piglets infected with a human rotavirus strain
had decreased sodium-potassium ATPase activity in the
upper jejunum. In contrast, when Graham et a1 (1984)
infected piglets with a porcine rotavirus (OSU strain),
sodium-potassium ATPase activities were depressed in the
ileum only.

Loss of membrane—associated digestive enzymes leads
to nutrient malabsorption. Miniature piglets infected
with porcine rotavirus (OSU strain) had an increase in
stool osmolarity from 248 i 20 mOsm/liter to 348 i 20
mOsm/liter at 75 hours after inoculation. The majority
of the osmotic gap in the feces could be accounted for by

lactose in the stools. Sodium concentration in the stool

 

 

 

 

increased, b1
1984). Male!
uptake of D-
rotavirus-in
al, l977;‘Wc

These 5
events in tl
rotavirus d
mucosal sur
reduced, an

malabsorpti

At ne
rotavirus
stomachs o
Partially
tine betwe
and Under(
refleCtS .
Small int
Cecum and
in lactea
rOtaVlrus
inoculat:
atrophy .

JerUUm,

31

increased, but potassium stayed the same (Graham et al,
1984). Malabsorption also has been measured by a reduced
uptake of D-xylose in the small intestine of
rotavirus-infected children and calves (Mavromichaelis et
al, 1977; Woode et al, 1978).

These studies indicate the following sequence of
events in the pathophysiology of rotaviral diarrhea: (a)
rotavirus destroys columnar Villous epithelial cells, (b)
mucosal surface area and important digestive enzymes are
reduced, and (c) osmotic diarrhea occurs due to nutrient
malabsorption.

Lesions and Pathogenesis

 

Gross Lesions

At necropsy, lesions in piglets infected with
rotavirus are confined to the intestinal tract. The
stomachs of experimentally—infected piglets are usually
partially filled with milk regardless of differences in
time between the last feeding and necropsy (Torres-Medina
and Underdahl, 1980). The color of the fecal material
reflects the diet and may be white, yellow, or grey. The
small intestinal wall is reduced in thickness, and the
cecum and colon may be distended with fluid. Visible fat
in lacteals in gnotobiotic piglets infected with
rotavirus is absent between 16 and 72 hours after
inoculation (Theil et al, 1978). Evidence of Villous
atrophy in the small intestine, especially in the

jejunum, can be seen by subgross examination (Pearson and

 

 

ltNulty, 197
Histopatholc
The hi:
are similar
al, l97lb; '
et al, 1983
shortening
epithelial
and exposur
quickly re'
with no mi
biochemica
1978).
Diarr
Small inte
within 1-
Villous a
after the
(BOhl et
hours aft
some gas,
Crouch a7
to 168 h
The
at whict
and Thej

early sw

32

McNulty, 1977).

Histopathologic Lesions

 

The histopathologic lesions of rotavirus infection
are similar in many mammalian and avian species (Mebus et
al, 1971b; Theil et al, 1978; Conner et a1, 1980; Johnson
et a1, 1983). The lesions are characterized by
shortening of small intestinal villi, desquamation of
epithelial cells lining the distal portion of the villus,
and exposure of the lamina propria. The latter is
quickly relined by squamous to cuboidal epithelial cells
with no microvillous border (Woode et al, 1976b) and with
biochemical characteristics of crypt cells (Middelton,
1978).

Diarrhea in piglets precedes the development of
small intestinal lesions. If small intestine is obtained
within 1-3 hours after the onset of diarrhea, little
Villous atrophy is evident, but if greater than 12 hours
after the onset of diarrhea, Villous atrophy is distinct
(Bohl et a1, 1978). Villous atrophy is most severe 24-72
hours after infection; villi may be essentially absent in
some cases (Pearson and McNulty, 1977; Theil et al, 1978;
Crouch and Woode, 1978). Villous atrophy persists for 48
to 168 hours after infection (Pearson and McNulty, 1977).

There is disagreement about the stage of infection
at which fusion of villi occurs. McAdaragh et a1 (1980)
and Theil et a1 (1978) reported fusion of villi in the

early stages of infection whereas Pearson and McNulty

 

 

(1977) and T
fusion of vi
The re}
lesions in 1
variable.
atrophy at
jejunum, an
villous atr
small intes
killed 36 c
severe vili
investigat
and distal
duodenum (
Torres-Med
In one of
Piglets, t
duodenal r
the pigs
lesions 0
intestine
gnotobiot
middle sn
in the up
Diffeteh(
methods ‘

33

(1977) and Torres—Medina and Underdahl (1980) noticed
fusion of villi only after several days.

The reported distribution of small intestinal
lesions in piglets infected with porcine rotavirus is
variable. Pearson and McNulty (1977) found villous
atrophy at all levels of the intestine (duodenum,
jejunum, and ileum). McAdaragh et a1 (1980) also noticed
villous atrophy and fusion of villi at all levels of the
small intestine. In fact, the duodenum of 2 piglets
killed 36 or 72 hours after infection, respectively, had
severe villous atrophy and fusion of villi. Other
investigators have found lesions primarily in the middle
and distal small intestine and "less strikingly" in the
duodenum (Woode et al, 1976; Crouch and Woode, 1978;
Torres-Medina and Underdahl, 1980; Graham et al, 1984).
In one of the earliest reports of "white scours" in
piglets, the distribution of lesions varied. The
duodenal mucosa was most severely altered in one—third of
the pigs (Mouwen et a1, 1971). One-half of the pigs had
lesions of greatest severity in the distal small
intestine. When a bovine rotavirus strain was given to
gnotobiotic piglets, lesions were most severe in the
middle small intestine, but milder lesions were detected
in the upper small intestine (Hall et a1, 1976).
Differences in age of the piglets, virus strains, and
methods of sampling the intestine might explain these

differences in lesion distribution.

 

 

Glandul
epithelial <
increased p1
gnotobiotic
and is maxi
et al, l980
porcine rot
and mid-sme
al, l984).
and ileum (
strain of ‘
(McAdaragh
(1983) not
gnotobioti
preceded t

The s
is influer
Et a1 (19.
Virus (gr
and mild
ePithelia
Villous 5
et a1 (1‘.
aSSOCiaU
(ch 1) a-
Specific

34

Glandular crypts respond to the loss of villous
epithelial cells caused by rotavirus infections by
increased proliferation. Crypt hyperplasia in
gnotobiotic piglets occurs by 48—60 hours after infection
and is maximal by 96 hours (Theil et al, 1978; McAdaragh
et a1, 1980). Piglets infected with the OSU strain of
porcine rotavirus had crypt hyperplasia in the jejunum
and mid-small intestine but not in the ileum (Graham et
al, 1984). However, crypt hyperplasia in the duodenum
and ileum of gnotobiotic piglets infected with a field
strain of porcine rotavirus has been reported by others
(McAdaragh et al, 1980). Interestingly, Johnson et a1
(1983) noted that crypt hyperplasia in the duodenum of
gnotobiotic pups infected with canine rotavirus was not
preceded by villous atrophy.

The severity and distribution of intestinal lesions
is influenced by the type of infecting rotavirus. Theil
et a1 (1985) studied the pathogeneis of a rotavirus-like
virus (group B) in gnotobiotic piglets and found variable
and mild histologic changes. Sloughing of villous
epithelial cells occurred only at the tips of villi and
villous atrophy was evident only occasionally. McNulty
et al (1983) compared the clinical and virologic findings
associated with infection by a group A avian rotavirus
(ch 1) and a non—group A avian rotavirus (ch 132) in
specific—pathogen—free chickens. There was a distinct

difference between the two rotavirus isolates in viral

 

antigen dis
had a predi
rotavirus s
the mid-sma
difference:
of rotavirr
Eosin'
observed h
mice infec
1967), rat
(Vonderfec
al, l98la)
only with
inclusion:
correlate
structure
The -
unCOmplic
gastric m
and McNul
Viral ant
1977; The
deSCribe(
epitheli;
rotaViru.
CEcum an

mlCrOsCO

35

antigen distribution. Rotavirus strain ch 1 (group A)
had a predilection for the duodenal mucosa, whereas
rotavirus strain ch 132 (non—group A) replicated best in
the mid-small intestine. This was the first report of
differences in intestinal tropism for different isolates
of rotavirus.

Eosinophilic cytoplasmic inclusions have been
observed histologically in the intestinal epithelium of
mice infected with murine rotavirus (Adams and Kraft,
1967), rats infected with an atypical rotavirus
(Vonderfecht et al, 1984), and in cell culture (Carpio et
al, 1981a). Inclusions, in most instances, were evident
only with the use of special stains. The nature of the
inclusions is unclear. Adams and Kraft (1967) could not
correlate inclusions seen by light microscopy with
structures seen by electron microscopy.

The pathogenesis of rotavirus infections tends to be
uncomplicated by lesions in other organs. Lesions in the
gastric mucosa have been reported (Woode 1975b; Pearson
and McNulty, 1977) but have not been associated with
viral antigen (McNulty et al, 1976a; Davidson et al,
1977; Theil et a1, 1978). Rodriguez et a1 (1980)
described cecal and colonic vacuolation of superficial
epithelial cells in mice infected with a murine
rotavirus. Murine rotavirus has been detected in the
cecum and colon of mice by immunofluorescent and electron

microscopic techniques (Adams and Kraft, 1967; Narita et

 

 

 

al, 1982).
Scanning Ele
Scannir
offers the a
specimens, :
visualized l
advantages,
studied by
Torres-Hedi
Underdahl (
the intesti
given a rot
case of p01
Visualized
lower smal
occluded t
36 hours a
lensth of
Cells expc
epithelial
epithelia]
dearly n01
There is <
fuSion 0c.
quiOn as
Edged V11

Underdahl

36

al, 1982).

Scanning Electron Microscopy

 

Scanning electron microscopy (SEM) of intestine
offers the advantages of tridimensional view, larger
specimens, and observation of details not readily
visualized by histologic examination. Despite these
advantages, porcine rotavirus infections have been
studied by SEM infrequently (McAdaragh et a1, 1980;
Torres-Medina and Underdahl, 1980). Torres-Medina and
Underdahl (1980) described the sequential SEM changes in
the intestinal mucosa of 6—day—old gnotobiotic piglets
given a rotavirus strain (B—3l7) isolated from a field
case of porcine rotaviral diarrhea. Lesions were first
visualized at 12 hours after infection in the middle and
lower small intestine. Enterocytes were swollen and
occluded transverse furrows and goblet cell openings. By
36 hours after inoculation, villi were one—third the
length of control villi and detachment of epithelial
cells exposed lamina propria. Microvilli on degenerate
epithelial cells were sparse and short. Regeneration of
epithelial cells was evident by 156 hours and villi were
nearly normal in dimension by 16 days after infection.
There is disagreement about the time at which villous
fusion occurs. McAdaragh et al (1980) observed villous
fusion as early as 12 hours post—inoculation (PI), and
fused villi were numerous by 72 hours. Torres—Medina and

Underdahl (1980) did not observe fused villi until 4.8

 

 

days PI.
Transmissic
Rotavi
electron mi
piglets by
Virions at
within ves:
The earlie;
between l8
swelling,
cytoplasmi
distended
endoplasmi
Cells with
be seen.
synthesize
Visible it
Narita St
The j
Observed }
after rot.
Piglets i
in the mi
(PEarson
Et al, 19

37

days PI.

Transmission Electron Microscopy

 

Rotavirus particles can be seen by transmission
electron microscopy (TEM) in the jejunum of gnotobiotic
piglets by 12 hours after infection (Narita et al, 1982).
Virions at this time are on and between microvilli and
within vesicles in the apical cytoplasm and terminal web.
The earliest cytopathologic alterations, detectable
between 18-24 hours after infection, are cytoplasmic
swelling, reduction in the number and size of microvilli,
cytoplasmic lipid droplets, rough endoplasmic reticulum
distended with vacuoles, and virus particles in the rough
endoplasmic reticulum. By 48 hours, immature cuboidal
cells with an uneven, incomplete microvillous border can
be seen. No virus particles are evident in newly
synthesized cells. Virus particles may or may not be
visible in goblet cells (Pearson and McNulty, 1979;
Narita et al, 1982).

Immunofluorescence and Location of Viral Antigens

 

The location and number of antigen—positive cells
observed by immunofluorecence (IF) is a function of time
after rotavirus infection. Maximal fluorescence in
piglets is observed in the enterocytes near villous tips
in the middle small intestine 24 hours after infection
(Pearson and McNulty, 1977; Crouch and Woode, 1978; Bohl
et al, 1978). McAdaragh et a1 (1980) first observed

fluorescence in the duodenum and upper jejunum of

 

 

 

 

gnotobiotic
fluorescenc
ileum by 36
with findir
1977; Bohl
and Underda
infected v:
intestine,
ileum. Du
not observ
rotaviruse
for 96 hou
et al, 197
infected v
than 24 h<
Viraf
than smal
seen in t'
Piglets (
and calve
nodes of
a1, l97lt
intesting
SPECifiC
Were Pre:
PIESehCe

prOPI‘ia '

38

gnotobiotic piglets 12 hours after infection. The
fluorescence progressively descended through the upper
ileum by 36 hours. These findings are in disagreement
with findings by other investigators (Chasey and Lucas,
1977; Bohl et a1, 1978; Theil et al, 1978; Torres—Medina
and Underdahl, 1980). The latter investigators found
infected villous epithelial cells throughout the small
intestine, but most consistently in the jejunum and
ileum. Duodenal fluorescence was sparse and variable or
not observed. Enterocytes infected by group A
rotaviruses desquamate so IF is only detected in piglets
for 96 hours (McAdaragh et al, 1980) to 168 hours (Theil
et a1, 1978) after infection. Gnotobiotic piglets
infected with a group B rotavirus had few IF cells later
than 24 hours after infection (Theil et al, 1985).

Viral antigen has been detected by IF in sites other
than small intestine epithelium. Immunofluorescence was
seen in the lamina propria of the small intestine of
piglets (Theil et al, 1978), dogs (Johnson et al, 1983),
and calves (Mebus et al, 1971b), and the mesenteric lymph
nodes of piglets and calves (Theil et a1, 1978; Mebus et
al, 1971b). Fluorescence observed in the small
intestinal lamina propria of calves was not regarded as
specific by Mebus et al (1971b) because many eosinophils
were present. But Stair et a1 (1983) substantiated the
presence of rotavirus in macrophages in the lamina

propria by TEM. A few infected enterocytes have been

 

 

 

 

detected in
(Narita et ;
(Banfield e
l983), and
1985).
Compar
negatively
time. Elev
content fr
signs cont
are nunero
greatest r
from pigs
1976; Bob?
Particles
5‘8 days

TOrres-Me

Resi
neutrali;
tract (P.
1978; Of
secreted
infectic

paSSiVel

39

detected in the cecum of rotavirus—infected piglets
(Narita et a1, 1982), the colon and rectum of mice
(Banfield et a1, 1968), colon of chickens (McNulty et al,
1983), and cecum and colon of calves (Reynolds et a1,
1985).
Viral Shedding

Compared to IF, rotavirus can be detected in
negatively stained fecal material for a longer period of
time. Electron microscopic examination of intestinal
content from piglets just beginning to show clinical
signs contain few rotaviral particles. Viral particles
are numerous by 24 hours after infection, and the
greatest number of particles are evident in specimens
from pigs 48 to 72 hours after infection (Lecce et al,
1976; Bohl et al, 1978). The number of rotavirus
particles is reduced or they cannot be detected between
5-8 days after infection (Tzipori and Williams, 1978;

Torres-Medina and Underdahl, 1980).

Law
Humoral Immunity

Resistance to rotavirus diarrhea is mediated by
neutralizing antibodies present within the intestinal
tract (Pearson and McNulty, 1977; Snodgrass and Wells,
1978; Offit and Clark, 1985). These antibodies may be
secreted into the intestinal lumen following intestinal
infection (Corthier and Vannier, 1983) or be obtained

passively via colostrum and milk (Bridger and Brown,

 

 

 

1981; Saif
antibody dc
infection
l985).
Passi
suckling p
adequate a
rotavirus
lgG, lgA,
Bachmann,
predomina1
persisting
class whi
class in
The
Pigs less
there are
infrequer
fEEd (Bot
antibody
weeks af'
on SElec-
an inCre
at this
Rot
inteStin

EECQS b>

40

1981; Saif et a1, 1984). The presence of serum rotavirus
antibody does not correlate with resistance to rotavirus
infection (Snodgrass and Wells, 1978; Offit and Clark,
1985).

Passive immunity against enteric pathogens occurs in
suckling pigs as a result of frequent ingestion of an
adequate amount of specific antibodies. Antibodies to
rotavirus in swine occur in the immunoglobulin classes
IgG, IgA, and IgM in mammary secretions (Hess and
Bachmann, 1981). The IgG class of immunoglobulin
predominates in colostrum, but the highest and longest
persisting rotavirus antibody is associated with the IgA
class which constitutes the predominant immunoglobulin
class in the milk of pigs (Vairman et a1, 1970).

The antirotaviral antibody in sows' milk protects
pigs less than 7-days-old from rotavirus infection unless
there are inadequate antibody titers in the milk,
infrequent suckling, or dilution of antibodies by creep
feed (Bohl et al, 1978). Protective IgA antirotaviral
antibody is found in sows' milk until weaning at 6 to 8
weeks after parturition (panel report on the Colloquium
on Selected Diarrheal Diseases of the Young, 1978). Thus
an increased incidence of rotavirus diarrhea is observed
at this time.

Rotavirus infection induces local production of
intestinal antibody. Immune complexes appeared in the

feces by 4 days after experimental inoculation of newborn

 

 

 

 

piglets with
were demonstr
and Vannier,
feces of hum:
clinical sig1
(Stals et al
evidence of
\dth rotavir
ammnestic r
production c
lived (42-7(
same rotavi:
infected (B
The ex
hOSt reSpop
understood,
some aSpect

infection

1983),

In
imflmnitv a
2 days aft
days after
9

SlmultaneC
aneStigat

41

piglets with rotavirus. Free IgA and IgM coproantibodies
were demonstrated by 7 days after inoculation (Corthier
and Vannier, 1983). Maximal excretion of IgA in the
feces of human infants occurred 7 days after the onset of
clinical signs and corresponded with clinical improvement
(Stals et al, 1984). In human infants, there also is
evidence of an immunologic memory because reinfection
with rotavirus is associated with an intestinal IgA
anamnestic response (Yamaguchi et al, 1985). The
production of intestinal antibody in calves may be short
lived (42-70 days), thereby allowing reinfection with the
same rotavirus strain with which they were originally
infected (Bachmann et al, 1983).

Cell-Mediated Immunity

 

The exact role of the different components of the
host response to rotavirus infection is not clearly
understood. Attention has been given only recently to
some aspects of the cellular immune response to rotavirus
infection (Riepenhoff—Talty et al, 1982b; Little et a1,
1983). In Balb—c mice, virus—specific cell mediated
immunity appeared in the splenic lymphocytes as early as
2 days after rotavirus infection and peaked around 10
days after infection (Riepenhoff—Talty et al, 1982).
Because cell—mediated immunity (CMI) increased
simultaneously with cessation of viral replication, these
investigators concluded that CMI may be important in

recovery from the disease.

 

 

 

Littl
lymphocyte
Infant mud
and T lymp
receiving
animals re
that did 1
important

Rota
by Eiden
experienc
which was
immunocon
seronegat
similar e
both the
resolved
antibody
dEEense
fUnction
antibody

Cel
transfei
IeUkOCYT
i985),
PaSsive

prOtth

42

Little et al (1983) studied the role of T and B
lymphocytes in recovery from rotaviral infections.
Infant nude mice were transfused with equal numbers of B
and T lymphocytes from syngenic adult mice. Animals
receiving spleen cells from immunologically mature

animals resolved their infections more quickly than those

 

that did not, which suggested that T and B cells were
important in the resolution of the rotavirus infection.
Rotavirus infection in athymic nude mice was studied

by Eiden et a1 (1986). Neonatal T-cell-deficient mice

 

experienced a self—limited gastrointestinal infection
which was identical to that observed in age-matched
immunocompetent mice. Also, adult T—cell-deficient

seronegative mice and age—matched normal mice showed a

 

similar extent of resistance to rotavirus infection. In
both the neonatal and adult mice, the infection was
resolved without the generation of antirotavirus
antibody. These investigators suggested that the host
defense against murine rotavirus requires neither
functional T—lymphocytes nor specific antirotaviral
antibody.

Cell—mediated immunity to rotavirus can be passively
transferred to neonatal calves by giving colostral
leukocytes from immunized heifers (Duhamel and Osburn,
1985). Although significant cell-mediated immunity is
passively acquired by the calves, it does not confer

protection when calves are orally challenged with live

 

bovine rotav
The big
gastroenter:
underscores
Commercial
rotavirus s
disappointi
1986).

Two a]
rotavirus
stinulatio
newborn ca
modified 1
afforded p
hours afte
Furthermo
neonatal
(Thurber
trials fa
RadOStitg

It 1
antibodi
newborn
Zaane et
rotaVirr

antibgdj

43

bovine rotavirus.
Immunization

The high morbidity and mortality caused by rotavirus
gastroenteritis in many mammalian and avian species
underscores the need for an efficacious and safe vaccine.
Commercial vaccines containing bovine or porcine
rotavirus strains are available, but results have been
disappointing (Hoblet et a1, 1984; Van Zaame et a1,
1986).

Two approaches have been used to try to prevent
rotavirus infections in calves. The first involves
stimulation of active immunity in the intestinal tract of
newborn calves by oral inoculation with attenuated
modified live rotavirus vaccines. These vaccines
afforded protection to gnotobiotic calves as early as 48
hours after inoculation (Mebus et a1, 1973).

Furthermore, use of this vaccine was reported to decrease
neonatal calf morbidity and mortality in field trials
(Thurber et al, 1977). However, other double-blind
trials failed to substantiate these findings (Acres and
Radostits, 1976).

It has been suggested that colostral rotavirus
antibodies may interfere with active immunization of the
newborn calf. This has been documented recently (Van
Zaane et a1, 1986). Oral vaccination, with modified live
rotavirus, of calves fed colostrum with rotavirus

antibodies did not induce a protective intestinal immune

 

response.
immunity i1
antibodies
delay or o:
A see
rotavirus
with subse
calf via c
cows with
increased
and milk,
lZ supple]
experimen
increase
multiple
81, 1983)
initially
cows to 1
bovine r<
(1984) h.
will res
With whi
instance
A u
rbtavirt
to nursj

.1 u
«lcense,

44

response. Oral vaccination did induce protective
immunity in calves which were not fed colostral rotavirus
antibodies, for practical purposes, it is not sensible to
delay or omit colostrum feeding.

A second approach used to protect calves from
rotavirus infection is immunization of the pregnant dam
with subsequent passive transfer of antibodies to the
calf via colostrum and milk. Parenteral immunization of
cows with an adjuvated modified live rotavirus vaccine
increased virus—neutralizing antibody titers in colostrum
and milk, which were protective when fed to calves as a
1% supplement (Saif et al, 1983). In the same
experiment, a commercially available vaccine did not
increase virus-neutralizing antibody titers. Because
multiple serotypes of bovine rotavirus exist (Murakami et
al, 1983) and do not show cross protection, it was
initially believed that for parenteral immunization of
cows to be effective, a vaccine must contain all of the
bovine rotavirus serotypes. However, Snodgrass et a1
(1984) have shown that after monovalent vaccination, cows
will respond heterotypically to all rotavirus serotypes
with which they were exposed. Therefore, in some
instances, single serotype vaccination may be sufficient.

A modified-live TGE virus and serotype 1 porcine
rotavirus vaccine to be administered to pregnant swine or
to nursing piglets prior to weaning is federally

licensed. There are no published controlled studies on

 

 

the efficac:
have shownii
of diarrhea
gains betwe
l984). The
calf vaccin
this has be
does not re
does it prc
Rotavf
causes of 2
years of a;
need for a:
has been r
develOpmen
Strain as
under acti
a1, 1979).
administre
induces re
moderate 1
rOtavirus.
Anot
adVantage
genetic r
CO-infeet

“O“‘CUIti

45

the efficacy of this vaccine, but preliminary studies
have shown no differences in the incidence or frequency
of diarrhea, rotavirus shedding, and post-weaning weight
gains between vaccinates and controls (Hoblet et al,
1984). The rotavirus strain in a commercially-available
calf vaccine has been used to vaccinate swine. However,
this has been shown to be ineffective because the virus
does not replicate in newborn colostrum-free piglets nor
does it protect them from disease (Lecce and King, 1979).

Rotaviruses are now considered one of the major
causes of gastroenteritis in children younger than two
years of age (Flewett and Woode, 1978). Thus, an urgent
need for an effective and safe vaccine for use in people
has been recognized. A number of approaches to vaccine
development are being studied. Use of a bovine rotavirus
strain as a potential vaccine candidate for people is
under active investigation (Zissis et a1, 1983, Wyatt et
al, 1979). Vesikari et a1 (1983) have shown that oral
administration of a bovine rotavirus (NCDV strain)
induces resistance in infants and young children against
moderate to severe diarrheal illness caused by human
rotaviruses.

Another approach to vaccine development takes
advantage of the propensity of rotaviruses to undergo
genetic reassortment (Greenberg et al, 1981).
Co-infection of cell cultures with animal rotaviruses and

non—cultivatable human rotaviruses has yielded

 

reassortar
the major
and the al
rotavirus

type repr

Ente
other ent
pigs. Al
assist ir
insuffici
definitii
the demOI
diarrhei
the pres

Man
been des
sensitiv
often he
StOpa, ]
The labc
are GHZj
micresc
Mm

En
are sin

utiliZc

46

reassortants which contain the gene segment coding for
the major neutralization protein from the human rotavirus
and the ability to replicate in yitgg from the animal
rotavirus (Midthun et a1, 1985). Reassortants of this
type represent potential live vaccine strains.

Diagnosis

Enteric disease caused by rotavirus and several
other enteropathogens is common in nursing and weanling
pigs. Although history, clinical, and necropsy findings
assist in obtaining an accurate diagnosis, they alone are
insufficient to distinguish among these diseases. A
definitive diagnosis of rotavirus infection depends on
the demonstration of rotaviral antigen in specimens from
diarrheic animals and correlation of these findings with
the presence of histopathologic lesions.

Many useful techniques for rotavirus detection have
been described, and the details of the relative
sensitivities, specificities, and advantages of each test
often have been provided (Yolken et a1, l978a; Yolken and
Stopa, 1979; Benfield et a1, 1984; Reynolds et al, 1984).
The laboratory tests that have been most widely compared
are enzyme—linked immunosorbent assays (ELISA), electron
microscopy (EM), and immunofluorescence (IF).

Enzyme—linked Immunosorbent Assays

 

Enzyme—linked immunosorbent assays are systems that
are similar in design to radioimmunoassays, but which

utilize an enzyme rather than radioactive isotope as the

 

immunoglol
enzyme, wd
antigen-a1
substrate
color, ca:
colorimet
determine
that can
a modific
was used
modificat
detectior
ELISA or
The
rotaviru;
were mor.
ReYnolds
several
of the E
microscc
l984).
Coo
for the
1984; Re
Compare.
detecri

ResUlts

47

immunoglobulin marker (Engvall and Perlmann, 1972). This
enzyme, when bound to the "solid phase" in a series of
antigen-antibody reactions, interacts with added
substrate to produce a product. The product, usually a
color, can be detected visually or with the aid of a
colorimeter. The sensitivity of these assays is
determined by the lowest concentration of visible color
that can be detected. Yolken and Stopa (1979) described
a modification of a rotavirus ELISA in which a substrate
was used that yielded a fluorescent product. This
modification improved the sensitivity of human rotavirus
detection by greater than 100 times compared to standard
ELISA or radioimmunoassay techniques.

The sensitivity of ELISA has been compared to other
rotavirus detection methods. In most instances, ELISA
were more sensitive than EM (Rubenstein and Mille, 1982;
Reynolds et al, 1984). For example, comparison of
several commercially—available ELISA indicated that one
of the ELISA was more sensitive (98%) than immunoelectron
microscopy (93%), IF (86%), and EM (842) (Mounet et a1,
1984).

Commercially—available ELISA correlate well with EM
for the detection of bovine rotavirus (Benfield et al,
1984; Reynolds et al, 1984). Reynolds et a1 (1984)
compared EM to enzyme—linked immunosorbent assay for the
detection of rotavirus and coronavirus in bovine feces.

Results showed excellent correlation with the detection

 

 

 

 

 

of rotavir
calves as
Benfield t
immunosorl
than IF 0:
rotavirus
enzyme-lit
detecting
enzyme-li
is not se
rotavirus
other lab
An a
viral ant
for detec
(1977) he
feces is
DisadVant
et al, 19
Viral ant
feces of
I984),
M
Bec.
animals f
remains

(I978b)

 

48

of rotavirus by EM in feces from experimentally infected
calves as well as from diagnostic laboratory specimens.
Benfield et a1 (1984) found a commercial enzyme-linked
immunosorbent assay as sensitive as EM but more sensitive
than IF or viral isolation for detecting bovine
rotavirus. However, EM was more sensitive than IF,
enzyme-linked immunosorbent assay, or viral isolation for
detecting porcine rotavirus. The commercially—available
enzyme-linked immunosorbent assay used by Benfield et a1
is not sensitive for the detection of porcine
rotaviruses, so it should be used in combination with
other laboratory procedures.

An advantage of ELISA when compared to EM is that
viral antigen need not be assembled into viral particles
for detection. This is useful because Mathan et a1
(1977) have shown that much of the rotavirus antigen in
feces is not assembled into viral particles.
Disadvantages of some ELISA are non—specificity (Chrystie
et al, 1983) and in some instances, inability to detect
viral antigen-antibody complexes which commonly occur in
feces of naturally infected animals (Crouch and Acres,
1984).

Electron Microscopy

 

Because many rotaviruses associated with diarrhea in
animals have been difficult to adapt to cell culture, EM
remains the primary method for their detection. Flewett

(l978b) reported that, with skill and perseverance, EM

 

 

will detect
should not
rapidly die
improved, 1
particles
This techn
The sensit
use of sol
technique
antirotavf
microscopj
direct EM
et al, 19
All
that can
and comp]
group spr
the grou‘
group A
of non~g
the COmn
inn
against
rOtavirr
these I,

49

will detect 105 viral particles/ml, 106 particles/ml
should not be missed, and 108 particles/ml will be
rapidly diagnosed. These sensitivities can be greatly
improved, however, by the aggregation of rotavirus
particles with antisera prior to EM (Saif et a1, 1977).
This technique is termed immunoelectron microscopy (IEM).
The sensitivity of EM also can be greatly enhanced by the
use of solid—phase immunoelectron microscopy. This
technique utilizes protein coated grids treated with
antirotavirus antibody. Solid phase immunoelectron
microscopy has been reported 30 times more sensitive than
direct EM and 10 times more sensitive than ELISA (Yolken
et al, 1979).

Immunofluorescence

 

All group A rotaviruses possess a common antigen
that can be detected by IF, IEM, ELISA, gel diffusion,
and complement fixation (Woode et al, 1976a). Thus,
group specific antigen and antisera prepared from any of
the group A rotaviruses can be used for the diagnosis of
group A rotavirus infections in any species. Detection
of non-group A rotaviruses requires antisera specific for
the common antigen of each group.

Immunofluorescence techniques which utilize antisera
against the common antigen of group A porcine
rotaviruses have been described (Theil et al, 1978). In
these reports, it has been demonstrated that rotaviral

antigen in intestinal smears or frozen sections can

 

 

usually 0T
Thus, in i
must be s<
infection
disadvant.
interpret
fluoresce
decomposi
Other Dia
Othe
describec
Woode an<
gnotobior
presence
negative
the most

rotaviru

At
Preventj
Vaccines
Van ZaaI
haVG as
of thes

Th
and mm

 

50

usually only be detected for 72-96 hours after infection.
Thus, in field cases of rotaviral diarrhea, specimens
must be selected from piglets in the early stages of
infection if IF tests are to be successful. Additional
disadvantages of IF procedures are subjective
interpretation, interference by non-specific background
fluorescence, and interference by post mortem
decomposition.

Other Diagnostic Techniques

 

Other techniques for rotavirus detection have been
described (Middleton et al, 1974; Middleton et al,l976;
Woode and Bohl, 1981). One of these, the inoculation of
gnotobiotic piglets, has been utilized to demonstrate the
presence of rotavirus when other techniques have proved
negative. Inoculation of the gnotobiotic piglet may be
the most sensitive means of demonstrating the presence of
rotavirus (Woode and Bohl, 1981).

Treatment and Control

 

At present, there are no practical methods for
preventing rotavirus infection; commercially available
vaccines have not proved efficacious (Hoblet et al, 1984;
Van Zaane et al, 1986). Management factors, therefore,
have assumed an important role in moderating the severity
of these infections.

Thorough cleaning and disinfection of the farrowing
and nursery units and the use of "all—in-all-out"

management systems will reduce the amount of virus in the

 

 

 

 

 

piglets'
to which
delay the
1980). R
environme
cannot be
(Snodgras
Unde
ambient t
associate
and Woode
piglets,
piglets 1
caused b
Res
neutrali
tract (S
Most sow
antibodj
immunity
and Bacl
Way is :
rotavir.
antibod-
and Boh
at birt

SeVerit

 

51

piglets' environment. Reduction of the quantity of virus
to which piglets are exposed may reduce the severity and
delay the onset of rotaviral diarrhea (Lecce and King,
1980). Rotaviruses are extremely resistant to
environmental conditions and disinfectants, so they
cannot be completely eliminated from the farm environment
(Snodgrass et a1, 1977; Sattar et a1, 1983).

Under natural and experimental conditions, a drop in
ambient temperature has been shown to increase mortality
associated with rotaviral infection in piglets (Crouch
and Woode, 1978). Meeting ambient temperature needs of
piglets, especially preventing chilling of neonates and
piglets with diarrhea, assists in controlling losses
caused by rotavirus.

Resistance to rotaviral diarrhea is mediated by
neutralizing antibodies present within the intestinal
tract (Snodgrass and Wells, 1978; Offit and Clark, 1985).
Most sows are positive for group A rotavirus serum
antibodies and will transfer variable amounts of passive
immunity to their piglets via colostrum and milk (Hess
and Bachmann, 1981). Passive protection obtained in this
way is important because a low infectious dose of
rotavirus in the presence of protective milk—secreted
antibody may result in immunity without disease (Woode
and Bohl, 1981). If piglets receive rotavirus antibodies
at birth, the onset of infection will be delayed, and the

severity of diarrhea and mortality caused by rotavirus

 

 

 

will be r
infection
2-weeks-o

In g
controlle
contamine
receive a

warm, drj

52

will be reduced. This is important because rotavirus
infections are most severe in piglets less than
2-weeks-old (Lecce et a1, 1976).

In general, rotaviral diarrhea of pigs can best be
controlled by avoiding environments that are heavily
contaminated with rotavirus and by ensuring that piglets
receive an adequate amount of colostrum and are kept

warm, dry, and well fed.

 

 

Six
hysterot
as descr
Control
inoculat
pigs wer
25-30°C.
York, NY
on the f
increase
Miniats
bEfore n

Inn
swabs fr
residual
each pie
Cultured
tergito]
aerobic
contamiI

examiner

 

 

 

 

MATERIALS AND METHODS

 

Gnotobiotic Pigs

Sixty-three pigs were obtained from 7 sows by closed
hysterotomy and maintained in stainless steel isolators
as described previously (Miniats and Joel, 1978).

Control pigs were kept in isolators separate from
inoculated pigs. Isolators containing these gnotobiotic
pigs were kept in a room with an ambient temperature of
25—30°C. The pigs were fed 50 m1 of SPF-lac (Borden, New
York, NY) three times (morning, afternoon, and evening)
on the first day. The volume of milk replacer was
increased by 13.5 ml/feeding daily as recommended by
Miniats and Joel (1978). Pigs were fasted 3 to 6 hours
before necropsy.

Immediately prior to inoculation, fecal and oral
swabs from 1 pig in each isolator and a composite swab of
residual milk in feed pans were collected. Ileum from
each pig was collected at necropsy. Swabs and ilea were
cultured for bacteria by use of sheep blood,
tergitol—seven, and brilliant-green agars incubated in
aerobic and anaerobic atmospheres. Evidence of bacterial
contamination also was obtained by histopathologic

examination of intestine.

53

'Pe

 

 

 

 

Micr¢
before in!
cultured
with the
enteric o
inoculati
Bacteria
intestine
with rote
necropsi.
Viabilit

by aerob

Por
Universj
(Ohio Ag
(OARDC)
TOtavir‘
(Bohl e
morphol
Strain
al, 197
intestj
Percin,
State .
DaVId

SD).

54

Microorganisms were not detected in any isolators
before inoculation of the pigs. Bacillus sp. was
cultured from the ileum of l pig 3 days after inoculation
with the SDSU strain of porcine rotavirus. A few
enteric organisms were cultured from 1 pig 7 days after
inoculation with the OSU strain of porcine rotavirus.
Bacteria were seen by histologic examination in the large
intestine of 2 pigs necropsied 1 day after inoculation
with rotavirus (l pig--OSU; l pig--SDSU) and 2 pigs
necropsied 3 days after inoculation with the OSU strain.
Viability of the observed bacteria was not demonstrated
by aerobic and anaerobic culture.

Kine-a

Porcine rotavirus designated as the Ohio State
University (OSU) isolate was obtained from Dr. Linda Saif
(Ohio Agricultural Research and Development Center
(OARDC), Wooster, OH). The OSU strain of porcine
rotavirus has serotype 1 and subgroup I specificities
(Bohl et a1, 1984; Hoshino et a1, 1984). Additional
morphologic and antigenic properties of this rotavirus
strain have been described (Saif et a1, 1977; Theil et
a1, 1978). The other porcine rotavirus was from an
intestinal homogenate of a pig with diarrhea. This
porcine rotavirus was designated as the South Dakota
State University (SDSU) isolate and was obtained from Dr.
David Benfield (South Dakota State University, Brookings,

SD). The SDSU strain of porcine rotavirus has not been

 

 

 

 

 

isolated
strain n
(Gottfri
(persona
Oh), so
rotaviru
times in
inoculat
gnotobic
contents
respectf
inocular
collectv
differei
tract a'
Prepare
New Yor
twice t
increas
intesti
10,000
Pipette
~70°c.
by fiii
(Millii
(APD) .

filtra

55

isolated in cell culture. Antisera against the SDSU
strain neutralized a serotype 2 porcine rotavirus
(Gottfried strain) at a titer of greater than 1:1024
(personal communication: Dr. Linda Saif, OARDC, Wooster,
OH), so this strain was considered a serotype 2 porcine
rotavirus. Each strain of virus was serially passaged 4
times in gnotobiotic pigs; diarrhea was observed in
inoculated pigs at each passage. At the fourth
gnotobiotic pig passage, the intestinal tract and
contents from 2 pigs given the OSU or SDSU isolates,
respectively, were aseptically collected 48 hours after
inoculation. To avoid cross-contamination, intestine was
collected and processed on different days and in
different locations. A 50% suspension of intestinal
tract and contents in Hanks' balanced salt solution was
prepared by homogenization in a blender (Waring Blender,
New York, NY). The homogenate was frozen and thawed
twice to rupture infected epithelial cells, thereby
increasing the quantity of released virus. The
intestinal homogenate was centrifuged 20 minutes at
10,000 x g. Aliquots (10 m1) of supernatant were
pipetted into sterile plastic tubes and refrigerated at
-70°C. As needed, supernatant was thawed and sterilized
by filtration through a series of Millipore filters
(Millipore Corp., Bedford, MA) with average pore diameter
(APD) of 0.65 pm, 0.45 mm, and 0.22 pm. Sterility of the

filtrate was confirmed by bacterial culture as described

 

 

 

 

above. 1
examined
containe
Feces) r

were obs

The
adapted
availabi
virus p;
and OSU
gnotobi

Tw
Pigs, w
virus p
10'6, 1
were p1
of pig:
Viruse
Piss <
totavi
with 1
Pools.
Signs
micro,
Pigle

in th

56

above. Negatively stained specimens of inocula were
examined by EM. Filtrates of the SDSU and OSU isolates
contained +4 (see Electron Microscopic Examination of
Feces) rotavirus particles. No contaminating organisms
were observed.

Virus Infectivity Titrations

 

The SDSU strain of porcine rotavirus has not been
adapted to cell culture, so an in yitgg method was not
available to determine accurately the number of infective
virus particles in the viral pools. Therefore, the SDSU
and OSU virus pools were titrated for infectivity in
gnotobiotic pigs.

Two litters, one containing 11 and the other 12

pigs, were used for the titration of the OSU and SDSU
5

9

virus pools, respectively. Various dilutions (10‘

10'6, 10‘7,

and 10_8) of the SDSU and OSU virus pools
were prepared in Hanks' balanced salt solution. Groups
of pigs inoculated with different dilutions and different
viruses were housed in separate isolators. Groups of 3

5 dilution of the osu

pigs (except the group given the 10'
rotavirus, which contained 2 pigs) were inoculated orally
with 1 m1 of either the diluted SDSU or OSU rotavirus
pools. Pigs were examined three times daily for clinical
signs of diarrhea. Feces were collected for electron
microscopic examination 48 hours after inoculation.

Piglets that became diarrheic and had rotavirus particles

in their feces at 48 hours after inoculation were

 

considers
caused d:
pig-infe
for the

10-5.8’

For
groups t
age, 28
contain:
rotaviru
rotavir'
pig was
appropr
Pigs) w
solutic
in eacl
Pigs we
Four p:
SDSU,

inocul

Signs
intern

SUSPQI

57

considered infected. The dilution of the virus pool that
caused diarrhea in 502 of the piglets was defined as one

pig-infectious—dose fifty (PIDSO). The calculated PID50

-7.2

for the SDSU and OSU rotavirus pools were 10 and

-5.8

10 , respectively.

Experimental Design

 

Forty gnotobiotic pigs were assigned to treatment
groups by use of a random number table. At 3 days of
age, 28 pigs were each inoculated orally with 1 ml
containing 105 PID50 of either the OSU strain of porcine 1
rotavirus (14 pigs) or the SDSU strain of porcine
rotavirus (14 pigs). Immediately after inoculation, each
pig was fed SPF-lac to which an additional 1 m1 of the
appropriate viral inoculum had been added. Controls (12
pigs) were inoculated with filtered Hanks' balanced salt
solution in the same manner as the principals. Five pigs
in each of the virus-inoculated groups and four control
pigs were killed at 24 and 168 hours after inoculation.
Four pigs in each of the three experimental groups (OSU,
SDSU, and control) were killed at 72 hours after
inoculation.

Clinical Signs and Body Weight

 

Control and infected pigs were observed for clinical
signs 3 times daily at the time of feeding. Body weight
was measured prior to inoculation and at 24 hour
intervals thereafter. Body weights were measured by

suspending pigs in a sling attached to a spring scale

 

 

 

(Ohaus, F

were fed

Bloc
vacutaine
for the e
and for
ethylene
Packed-c
microhen
profiles
Medicine
reagent:

Industr

Pi
bYani
barbite
the 8m:
micros.
Scanni
Was re
last P
appro;
The re
1/3,

flexu

58

(Ohaus, Florham Park, NJ) with accuracy to i20 gms. Pigs
were fed after body weights were recorded.

Clinical Pathology

 

Blood collected at necropsy was dispensed into
vacutainer tubes (Becton—Dikinson, Rutherford, NJ) which
for the collection of serum contained no anti-coagulant
and for the collection of whole blood contained
ethylenediamine—tetracetic acid (EDTA).
Packed—cell-volume of unclotted blood was measured by the
microhematocrit method (Benjamin, 1978). Serum chemistry
profiles were measured by the Laboratory of Clinical
Medicine (Sioux Falls, SD). Technicon SRA 2000 and
reagents for the "basic panel” from Technicon (Technicon
Industrial Systems, Tarrytown, NY) were used.

Necropsy

Pigs were bled by cardiac puncture and then killed
by an intracardiac injection of a solution containing
barbiturates. Segments (3 cm in length) from 6 sites in
the small intestine were removed immediately for light
microscopy (LM), fluorescent antibody testing (FAT), and
scanning electron microscopy (SEM). The first segment
was removed at the duodenal—jejunal flexure (DU) and the
last portion was removed from the lower ileum (LI)
approximately 5 cm proximal to the ileocecal junction.
The remaining segments were removed at approximately 1/6,
1/3, 1/2, and 2/3 the distance from the duodenal—jejunal

flexure to the ileocecal junction and were designated as

the uPPer
intestine
respecti‘
LM and S]
A small
intestin
Karnovsk
and refr
was inci
attachme
straigh‘
formali
rectum,
In addi
heart,
neutral
335%!
G
of por
serum
Were h
State
Storee
gamma
modif
(Plum

Uhles

59

the upper jejunum (UJ), mid-portion of the small
intestine (MI), lower jejunum (LJ), and upper ileum (U1),
respectively. The lumen of each section of intestine for
LM and SEM was rinsed with 10% neutral buffered formalin.
A small portion (0.5 cm2) of each segment of small
intestine, cecum, and colon was removed and immersed in
Karnovsky's fixative at pH 7.4 (Karovsky, 1965) for SEM
and refrigerated at 4°C. The remaining small intestine
was incised longitudinally along the mesenteric
attachments, pinned to rigid sections of cardboard for
straight flat fixation, and immersed in neutral buffered
formalin. Sections of unfixed stomach, cecum, colon,
rectum, and mesenteric lymph node were collected for FAT.
In addition to these tissues, lung, liver, kidney, brain,
heart, thymus, pancreas, and spleen were immersed in
neutral buffered formalin for LM.

Preparation of Fluorescein-Conjugated Gamma Globulin

Gnotobiotic pig hyperimmune serum to the SDSU strain

of porcine rotavirus (serotype 1) and rabbit hyperimmune
serum to the OSU strain of porcine rotavirus (serotype 2)
were kindly supplied by Dr. David Benfield (South Dakota
State University, Brookings, SD). Hyperimmune serum was
stored at -70°C until used. Fluorescein-conjugated
gamma-globulins were prepared from hyperimmune serum by a
modification of procedures developed by Dr. I. C. Pan \
(Plum Island Research Laboratories, Plum Island, NY).

Unless noted, all procedures were done at room

 

 

 

 

temperat'
flask.
addition
solutior
magnetic
been ade
Precipi1
x g for
decante
origina
before.
the lae
resuspe
Tris-hf
dissol‘
minute
SUpern
(Spect
dialyz
M Tris
total
deter;
Louis
isoth
was a

a fit

mixtt

6O

temperature. Serum was dispensed into an Erlenmyer
flask. The gamma-globulin was precipitated by the
addition of an equal volume of saturated ammonium sulfate
solution dropwise while stirring with a mechanical
magnetic stirrer. After all the ammonium sulfate had

been added, the mixture was stirred for 20 minutes.

 

Precipitated gamma-globulin was then centrifuged at 1,900

x g for 30 minutes at 4°C. After the supernatant was
decanted, the pellet was resuspended in one-half the
original volume of ammonium sulfate and centrifuged as
before. This step was repeated (usually 3 times) until
the last pellet was completely white. The pellet was
resuspended in the smallest amount possible of 0.01 M
Tris-hydrochloric acid buffer (HCl) (pH 9.0). The
dissolved pellet was centrifuged at 1,100 x g for 30
minutes at 4°C. To remove residual ammonium sulfate, the
supernatant was dispensed into Spectrapor membrane tubing
(Spectan Medical Industries, Inc., Los Angeles, CA) and
dialyzed overnight at 4°C against a large volume of 0.01
M Tris-HCl in a beaker with a magnetic stirrer. The
total protein of the concentrated gamma-globulin was
determined by the biuret method (Sigma Diagnostics, St.
Louis, MO). Sufficient isomer I of fluorescein
isothiocyanate (FITC) (Sigma Chemical Co., St. Louis, MO)
was added to the gamma globulin in a 50 ml beaker to give
a fluorochrome-to-protein ratio by weight of 1:100. The

mixture was slowly (to prevent denaturation of the

 

 

 

globulin)
refrigeré
the mixtt
cm in he:
(Pharmac
equilibr
(pH 7.3)
Bee
was adse
Louis, 1
(pH 7.3
mg of l
and all
Tissue
minute:
time a1
um APD
plasti
diluti
Stainj

dilut:

Place
Napex
tissr

(IUte

61

globulin) stirred overnight by a mechanical stirrer in a
refrigerator at 4°C. Unconjugated FITC was removed from
the mixture by gel filtration through a glass column (60
cm in height x 5 cm in diameter) packed with Sephadex—25
(Pharmacia Fine Chemicals, Inc., Piscataway, NJ)
equilibrated with 0.1 M phosphate buffered saline (PBS)
(pH 7.3).

Because of non-specific fluorescence, the conjugate
was adsorbed with liver powder (Sigma Chemical Co., St.
Louis, MO). The conjugate was adjusted with 0.1 M PBS
(pH 7.3) to a concentration of 3.3 mg of protein/ml. Ten
mg of liver powder per mg of conjugate protein were mixed
and allowed to stand 1 hour with occasional stirring.
Tissue powder was removed by centrifugation for 30
minutes at 20,000 x g. This process was repeated one
time and the conjugate was then filtered through a 0.45
pm APD Millipore filter. Small aliquots of conjugate in
plastic vials were stored at —20°C until used. Various
dilutions of conjugate in 0.1 M PBS were tested in
staining trials to determine the appropriate conjugate
dilution for FAT.

Examination by Immunofluorescence

 

Specimens for fluorescent antibody testing were
placed in embedding medium (Miles Laboratories, Inc.,
Naperville, IL), and quick frozen at —70°C. Frozen
tissues were cut 8—10 pm thick with a freezing microtome

(International Equipment Company, Needham Heights, MA)

 

 

 

and then
After fi:
tissue s
anti-rot
section
anti-rot
were ap1
for 30 I
minutes
minute
mountin
and cov
examine
immunoi
illumiI
German:
Site w
coarse
to con
rotavi

aS the

Secti
eOSin
Hunt,

QVale

62

and then two sections were placed on one glass slide.
After fixation in acetone for 10 minutes, one of these
tissue sections was stained with fluorescein-conjugated
anti—rotavirus (SDSU strain) globulin and the other
section was stained with fluorescein—conjugated
anti—rotavirus (OSU strain) globulin. After the stains
were applied, the slides were placed in a humid chamber
for 30 minutes at 37°C. Then slides were washed for 5
minutes in each of 2 rinses of 0.1 M PBS and then for 1
minute in distilled water. Slides were air-dried and
mounting medium pH 9 (Difco Laboratories, Detroit, MI)
and coverslips were applied. Stained preparations were
examined by immunofluorescence microscopy using a Zeiss
immunofluorescent microscope with epifluorescent
illumination and a Xenon light source (Carl Zeiss, West
Germany). At least 4 sections of small intestine per
site were evaluated. Specimens with intense fine to
coarsely granular cytoplasmic fluorescence when compared
to control tissues were considered positive (+) for
rotavirus antigen. Tissues from uninfected pigs served
as the controls.
Light Microscopy

Formalin—fixed tissues were embedded in paraffin,
sectioned 6 pm thick, and stained with hematoxylin and
eosin using standard histologic techniques (Thompson and
Hunt, 1966). Histopathologic changes in tissues were

evaluated with the use of a light microscope. In

 

 

 

addition
ocular m
height (
crypts v
intestir
mucosal
made to
Additio
subject
scale w
and a (
cuboide
conside
by the
segmen
(-) wh
which

villor

Piece
addit
other
Princ
each
M phc

Chang

63

addition, small intestine was examined using a calibrated
ocular micrometer to determine crypt depth and villous
height (Moon et a1, 1975). Five, well—oriented villi and
crypts were measured from each segment of small
intestine. Because submucosal lymphoid nodules affect
mucosal architecture (Mouwen et al, 1971), no attempt was
made to measure villi or crypts in these areas.
Additional changes in intestinal segments were
subjectively graded on a negative (-) to plus three (+3)
scale where (-) represented no change from the control
and a (+3) represented the greatest change. Squamous,
cuboidal, and low columnar epithelial cells were
considered immature. Cytoplasmic vacuolation was scored
by the number of vacuolated epithelial cells in each
segment of intestine. Vacuolation ranged from a negative
(—) which represented no vacuolation to a plus four (+4)
which equaled vacuolation of approximately 100% of the
villous epithelial cells.

Scanning Electron Microscopy

 

Tissues for SEM were refrigerated at 4°C until
processed for SEM (usually within 24—48 hours). All
additional processing was done at room temperature unless
otherwise specified. Duodenum, UJ, and LI from 3
principal pigs and 1 control pig per treatment group at
each time period were rinsed in each of 3 changes of 0.2
M phosphate buffer and washed for 8 hours in each of 3

changes of double—distilled water. Tissue was post—fixed

 

 

 

 

in 1% 03
wash, an
additior
Tie
(Polyscf
were d0]
and I-Iil
thiocar
treatme
tissue
over 3C
Ti
(Maser
acidif'
H(31/50
for 15
change
dehydr
ethane
Vacuur
Garbo]
Silve-

Co.

9

exami
(lute
acce]

magnj

64

in 1% osmium tetroxide for 30 minutes. The above rinse,
wash, and post-fixation steps were repeated one
additional time.

Tissue staining with thiocarbohydrazide
(Polysciences, Inc., Warrington, PA) and osmium tetroxide

were done using procedures described previously (Malick

 

and Wilson, 1975) with the following modifications: the
thiocarbohydrazide was a saturated solution; each
treatment with osmium tetroxide was for 30 minutes; and
tissue was washed in 3 changes of double—distilled water
over 30 minutes.

Tissue dehydration was done by reported methods
(Maser and Trimble, 1977) except that only 2 ml of
acidified dimethoxypropane (DMP) (1 drop of concentrated
HC1/50.0 m1 of DMP) was applied to each tissue section
for 15 minutes. The DMP was removed and each of 3
changes of 100% ethanol was applied for 15 minutes. The
dehydrated ethanol treated specimens were placed in 100%
ethanol in a critical point drying apparatus (Denton
Vacuum, Inc., Cherry Hill, NJ) and were dried in liquid
carbon dioxide. Dried specimens were attached with
silver conducting paint (Acme Chemicals and Insulation
Co., New Haven, CT) to aluminum SEM stubs and were
examined using a Super IIIA scanning electron microscope
(International Scientific Instruments, Korea) at an
accelerating voltage of 15kv and at various

magnifications and working distances. Images were

 

 

recorded

Cambridg

Fee
to inoce
isolator
Sterile
fecal s.
swabs w
control
stored
~70°C u

Ea
Suspenc
by cent
pellet
at 40,'
discar
distil

A
Ritchi
of dis
(PTA)
(BSA)
depre.
aPpli

Tnsti

65

recorded on Polaroid Land Film Type 55 (Polaroid Corp.,
Cambridge, MA).

Electron Microscopic Examination of Feces

 

Fecal specimens were collected from each pig prior
to inoculation and from the remaining pigs in each

isolator 24, 72, and 168 hours after inoculation.

 

Sterile plastic tubes (12 x 75 mm) were used to collect
fecal samples, or if pigs were not scouring, sterile
swabs were used. Cecal contents were collected from each
control and infected pig at necropsy. Specimens were
stored in a freezer (Revco, Inc., Deerfield, M1), at
—70°C until processed.

Each specimen was thawed at room temperature,
suspended in Hanks' balanced salt solution, and clarified
by centrifugation at 3,000 x g for 15 minutes. The
pellet was discarded and the supernatant was centrifuged
at 40,000 x g for 30 minutes. The supernatant was
discarded and the pellet was suspended in 0.5 ml of
distilled water.

A negative-staining procedure as described by
Ritchie and Fernelius (1968) was used. Briefly, 20 drops
of distilled water, 2 drops of 42 phosphotungstic acid
(PTA) pH 6.2-6.8, 1 drop of 0.1% bovine serum albumin
(BSA), and 1 drop of the fecal suspension were mixed in a
depression dish with a Pasteur pipette. The mixture was
applied with an all—glass nebulizer (Ted Pella, Inc.,

Tustin, CA) to nitrocellulose coated 200 mesh grids.

 

 

Specimer
HU—IZA t
Japan) i
the PTA
optimiz«

Th

criteri

Bo
analyze
varianc
by usir
from ce
level e
Dr. Wi

South

66

Specimens were examined immediately with an Hitachi
HU—12A transmission electron microscope (Hitachi, Tokyo,
Japan) at an accelerating voltage of 75kv. If needed,
the PTA, BSA, or sample concentrations were adjusted to
optimize spreading and contrast.

The density of viral particles was estimated by
criteria described by Benfield et a1 (1984).

Statistical Analysis

 

Body weights, villous heights, and crypt depths were
analyzed statistically by using the one-way analysis of
variance. Differences between group means were analyzed
by using the Waller-Duncan K-ratio T—test. Differences
from control values were considered significant at the
level of p<0.05. (The statistical analysis was done by
Dr. William Tucker, Agricultural Experiment Station,

South Dakota State University, Brookings, SD).

 

 

 

 

All
with the
observed
strain.
deveIOp .
with the
lengths
observed
strain,
not have

The
the SD31
Pigs wi'
undiges
hours p
Persist

inocula

RESULTS

Clinical Signs

A11 pigs in the control group and pigs inoculated
with the OSU or SDSU strains survived. Diarrhea was
observed in 10/14 pigs 19-36 hours P1 with the OSU
strain. Four pigs inoculated with the OSU strain did not
develop diarrhea until 44—48 hours PI. Pigs inoculated
with the SDSU strain had incubation periods of varying
lengths before the onset of diarrhea. Diarrhea was
observed in 11/14 pigs 24 to 54 hours P1 with the SDSU
strain. Three pigs inoculated with the SDSU strain did
not have diarrhea when killed at 24 hours PI.

The appearance of feces from pigs inoculated with
the SDSU or OSU strains were similar. Initially,
pigs with diarrhea had watery yellow feces. Curds of
undigested milk in the feces were first observed 48—96
hours PI. Diarrhea with curds of undigested milk
persisted until pigs were killed 168 hours PI. Pigs
inoculated with the OSU strain in most instances became
reluctant to drink milk shortly before the onset of
diarrhea, and milk remained in the feeding bowls at the
time of the next feeding. The pigs infected with the OSU

strain remained anorectic until 48-96 hours PI. Vomiting
67

g

 

 

was obsei
the OSU :
strain d'
Con

the expe
fter ea

characte

Pig
rotavir1
comparee
pigs in
pigs ha
The rot
gains 1
inocule
by the
by 144
a weig
the sa
pig in
152 wt
inocu]
of 27:
body \

stati

68

was observed in 5 of 14 (36%) of the pigs infected with
the OSU strain, whereas pigs inoculated with the SDSU
strain did not vomit.

Control pigs remained active and alert throughout
the experiment and rapidly consumed their entire ration
after each feeding. Control pigs had pasty golden feces
characteristic of milk-fed gnotobiotic pigs.

Body Weight

Pigs inoculated with the OSU or SDSU strains of
rotavirus had lower body weights and reduced weight gains
compared to the controls (Table 1). At 168 hours PI,
pigs infected with the OSU or SDSU strains and control
pigs had weight gains of 16, 22, and 46%, respectively.
The rotavirus-infected pigs had highly variable weight
gains irrespective of the strain of rotavirus used as the
inoculum. These differences in weight gain are reflected
by the high standard deviations (Table 1). For example,
by 144 hours PI, a pig inoculated with the OSU strain had
a weight loss of 25% whereas another pig inoculated with
the same virus had a weight gain of 21%. Similarly, a
pig inoculated with the SDSU strain had a weight loss of
15% whereas another pig inoculated with the same pool of
inoculum and kept in the same isolator had a weight gain
of 27%. Because of this extreme pig to pig variation,
body weights of virus infected pigs were not

statistically different (p<0.05) from controls.

 

 

Table 1
pigs in
rotavir‘

 

Treatme
rou
Control

OSU

SDSU

 

a

groups

69

Table 1. Mean body weights (g i SD) for gnotobiotic
pigs infected with the OSU or SDSU strains of porcine
rotavirus

 

Hours after inoculation
Treatment
group 0 24 48 72 96 120 144 168

Control 1254a 1295 1456 1520 1620 1652 1741 1832
i142 i142 i158 i143 i127 i153 i160 1138

OSU 1320 1323 1386 1363 1362 1386 1432 1526
i222 i255 i177 i232 i248 i265 i292 i311

SDSU 1257 1316 1309 1346 1378 1412 1464 1534
i224 i227 i220 i228 $88 $99 i112 i126

 

 

a Body weights of control, OSU, and SDSU treatment

groups were not significantly different (p<0.05).

 

 

 

He
pigs he
deliver
useful
with tl
chemis1
proteir
inocul.
total‘
24, 72
reduce
Howeve
l68 ho
contrc
pigs i
time.
before
serum
Chlor:

cont 1:.

Varied
as the
judged

PrOceE

Clinical Pathology
Hematocrit values varied among pigs because some
pigs had excessive umbilical cord bleeding after
delivery. Therefore, hematocrit measurements were not
useful as an index of hemoconcentration. Pigs inoculated
with the OSU or SDSU strains had similar changes in serum
chemistry values (Tables 2, 3, and 4). The total serum
protein and BUN values were greater in rotavirus
inoculated pigs than in controls. The elevations in
total protein and BUN were greatest at 168 hours PI. At
24, 72, and 168 hours PI, serum triglycerides were
reduced in virus infected pigs when compared to controls.
However, results of serum triglyceride measurements at
168 hours PI were not meaningful because sera from 3
controls, 3 pigs inoculated with the OSU strain, and 2
pigs inoculated with the SDSU strain were lipemic at this
time. Lipemia, which indicates insufficient fasting
before the collection of serum, may have interfered with
serum chemistry determinations. Sodium, potassium,
chloride, cholesterol, glucose, and creatinine values for
control and rotavirus infected pigs were similar.
Gross Lesions
The external appearance of virus—inoculated pigs
varied and was independent of the strain of rotavirus used
as the inoculum. Some pigs were thin and dehydrated (as
judged by decreased skin pliability and prominence of bony

processes) and had rough hair coats whereas others

 

 

Table 2
gnotobie
or SDSU

__—--——

ire-e
Hematoc
Total p
Glucose
BUN

Creatir
Sodium.
Potass
Chlori
Cholee

Triglj

‘—

l

71

Table 2. Serum chemistry and hematocrit values for
gnotobiotic pigs 24 hours after inoculation with the OSU
or SDSU strains of porcine rotavirus

 

Treatment group

 

 

Parameter Units Control OSU SQSU
Hematocrit Z 29.0 i 1.2 29.8 i 0.2 26.4 i 0.5
Total protein gm/dl 2.5 i 0.3 2.6 i 0.1 2.6 i 0.2
Glucose mg/dl 97 i 14 110 i 5 83 i 0
BUN mg/dl 7.8 i 0.3 9.4 i 4.9 18.6 i 14.
Creatinine mg/dl 1.0 i 0.1 1.0 i 0.1 1.1 i 0.1
Sodium meq/l 149 i 0.6 148 i 3.4 151 i 4.3
Potassium meq/l 5.4 i 0.2 5.8 i 0.6 5.6 i 0.7
Chloride meq/l 108 i 2.3 108 i 1.1 108 i 4.0
Cholesterol mg/dl 149 i 33 136 i 14 180 i 8
Triglyceride mg/dl 90 i 2 34 i 17 51 i 18

 

Data are expressed as the mean i SD.

Table 3.
otobio
or SDSU

#

mains
Hematocr
Total p1
Glucose
BUN
Creatin
Sodium
Potassi
Chlorid
Cholest

Triglyc

N

De

Table 3.

72

Serum chemistry and hematocrit values for

gnotobiotic pigs 72 hours after inoculation with the OSU
or SDSU strains of porcine rotavirus

 

 

Treatment group
OS

 

Parameter Uplpp Control SQSU
Hematocrit % 27.0 i 2.3 28.3 i 0.4 25.3 i 2.0
Total protein gm/dl 2.8 i 0.3 3.0 i 0.5 2.9 i 0.2
Glucose mg/dl 116 i 26 103 i 4 110 i 17
BUN mg/dl 7.5 i 0.6 27.0 i 18.5 26.3 i 28.
Creatinine mg/dl 1.1 i 0.1 0.9 i 0.2 1.1 i 0.3
Sodium meq/l 154 i 9.2 153 i 10.7 154 i 2.7
Potassium meq/l 6.1 i 0.8 6.5 i 0.7 5.7 i 0.1
Chloride meq/l 118 i 10.7 116 i 8.4 116 i 5.8
Cholesterol mg/dl 157 i 15 137 i 21 113 i 12
Triglyceride mg/dl 89 i 17 46 i 34 48 i 19

Data are expressed as the mean i SD.

 

 

Table 4.
gnotobic
or SDSU

#

rams!
Hematocr
Total p1
Glucose
BUN
Creatin:
Sodium
Potassi
Chlorid
Cholest

Triglyc

\—

Da

73

Table 4. Serum chemistry and hematocrit values for
gnotobiotic pigs 168 hours after inoculation with the OSU
or SDSU strains of porcine rotavirus

 

Treatment group

 

 

Parameter Upipg Control OSU SQSU
Hematocrit % 32.0 i 4.6 28.4 i 2.9 27.0 i 0.0
Total protein gm/dl 2.7 i 0.3 3.3 i 0.4 3.3 i 0.0
Glucose mg/dl 151 i 8 119 i 8 105 i 24
BUN mg/dl 5.0 i 0.0 37.2 i 30.4 31.6 i 22.8
Creatinine mg/dl 1.2 i 0.1 1.0 i 0.1 1.0 i 0.0
Sodium meq/l 148 i 2.3 154 i 12.1 150 i 9.6
Potassium meq/l 5.8 i 0.0 5.2 i 0.2 5.1 i 0.3
Chloride meq/l 111 i 4.0 119 i 12.8 114 i 10.1
Cholesterol mg/dl 143 i 25 157 i 26 128 i 51
Triglyceride mg/dl 203 i 13 120 i 92 62 i 25

 

Data are expressed as the mean i SD.

 

appearee
fecal 31
In1
infectee
The stor
amounts
with wa'
of incor
hours P
pale, f
PI. Th
strain
normal
Tb
Chyle i
Small 5
Of the
Organs
Gt
LYmpha
Contai

intest

L

\—

V

from p

rOtavi

PI (Tc

C

74

appeared healthy except for the presence of diarrhea and
fecal staining on the perineum.

Internally, gross changes were similar in pigs
infected with either the OSU or SDSU strain of rotavirus.
The stomachs of virus-infected pigs contained various

amounts of clotted milk. Ceca and colons were distended

 

with watery yellow contents which contained small clots

of incompletely digested milk in pigs killed at 72 and 168

hours PI. The small intestine in pigs with diarrhea was

pale, flaccid, and thin-walled at 24, 72, and 168 hours v
PI. The intestines of two pigs inoculated with the SDSU

strain which did not have diarrhea at 24 hours PI were

normal grossly.

The majority of virus-infected pigs did not have
chyle in mesenteric lymphatics 24 hours after inoculation.
Small amounts of chyle were seen in the proximal one—third
of the small intestine at 72 and 168 hours PI. Other
organs of infected pigs were normal grossly.

Gross lesions were not seen in control pigs.
Lymphatics in the small intestine of control pigs
contained various amounts of chyle in the proximal small
intestine 24, 72, and 168 hours PI.

Light and Scanning Electron Microscopic Changes

 

Villi in each of the small intestinal sites sampled
from pigs inoculated with the OSU or SDSU strains of
rotavirus were shorter than control villi at 24 hours

PI (Table 5). The DU was less severely affected by

 

 

 

 

Table
gnotol
porcir

 

H01
aft
inocul

 

2!

16

for t
lnocu

Value

beth

 

75

Table 5. Villous length (pm) in the small intestines of

gnotobiotic pigs infected with the OSU or SDSU strains of
porcine rotavirus

 

 

 

 

Hours
after Treatment Intestinal site
inoculation group DU UJ LJ MI UI L1
24 Control 855 906 813 825 781 815
i4 ea i12 i6 i48 e18
osu 645 342a 501 541 594 546
i5 i44 e12 i27 ill e11
snsu 675 526a 544 537 513 422
i31 e40 e30 i58 i0 i24 ,
72 Control 827 943 833 839 897 952
i7 $7 $28 e9 i63 e12
osu 614a’b 227a 124a“b 82a 194a 249a
i82 e37 e19 e7 e11 i4
sosu 3543’b 311a l96a’b 155a 247a 366a
113 e23 i2 i0 i3 i7
168 Control 845 810 823 939 1016 1002
$19 i27 i27 i9 144 i64
osu 874 419a 364a 321a"b 376a 686a
i42 ilO i9 e21 e10 e5
snsu 439‘?"b 322a 289a 245‘?"b 293a 640a
i1 i26 i4 i9 i7 e22

 

Data are expressed as the mean i SE (N = 20 except
for the OSU and SDSU groups at 24 and 168 hours after
inoculation in which N 25).

Values significantly different from control
values, p<0.05.

Significant difference (p<0. 05) in villous length
between the OSU and SDSU inoculated pigs.

 

 

 

villous
villous
differe
groups
24 hou1
the OSI
Villous
with tl
(Figure
infecte
controi
the UJ
SDSU 8'
mean v
SDSU s
168 ho-
strain
in the
hours

in the
were 6
of pie
Villi

or SDE
PI, b1
DU of

Villi

 

76

villous atrophy than the other sites. The reduction in
villous length in the UJ (Figure 1) was significantly
different (p<0.05) when each of the rotavirus inoculated
groups of pigs were compared to the control (Table 5). At
24 hours PI, mean villous lengths for pigs inoculated with
the OSU or SDSU strains were not significantly different.
Villous atrophy in the small intestines of pigs infected
with the OSU or SDSU strains was maximal at 72 hours PI
(Figure 2). At this time, villous lengths in virus
infected pigs were significantly (p<0.05) shorter than
controls (Table 5). Villous atrophy was most severe in
the UJ, LJ, MI, and UI of pigs infected with the OSU or
SDSU strains of rotavirus at 72 and 168 hours PI. The
mean villous lengths in the DU of pigs infected with the
SDSU strain were significantly shorter (p<0.05) at 72 and
168 hours PI when compared to pigs infected with the OSU
strain or control pigs (Figures 3, 4, 5, 6, and 7). Villi
in the duodenum of pigs infected with the OSU strain 168
hours PI were the same length as the control villi. Villi
in the LJ of pigs infected with the OSU strain 72 hours P1
were significantly shorter (p<0.05) than villi in the LJ
of pigs infected with the SDSU strain and control pigs.
Villi in the small intestine of pigs infected with the OSU
or SDSU strains were longer at 168 hours than at 72 hours
PI, but were still significantly shorter, except for the

DU of pigs infected with the OSU strain, than control

villi at each site.

 

 

Pigur
Pig 2
Porci
(arro

tips

77

 

Figure 1. Scanning electron micrograph of the UJ of a
pig 24 hours after inoculation with the OSU strain of
porcine rotavirus. Notice the rounded absorptive cells
(arrowhead) and the exposed lamina propria (arrow) on the

tips of atrophic villi. Bar = 100 um.

 

 

 

 

 

Figur
pig ?
Porci

denue

 

78

 

Figure 2. Section of small intestine from the MI of a
pig 72 hours after inoculation with the OSU strain of

porcine rotavirus. Villi are short, fused, and focally
denuded (arrow). Hematoxylin and eosin stain; bar = 10

um.

 

 

 

 

‘ Km
’ _. 16. (2m

Figu
Pig

pore

stai

79

 

testine from the DU of a

In

Section of small

3.

igure

F

th the SDSU strain of

ion w1

lat

anCH

72 hours after ’

pig

lined

Moderately shortened villi are

irus.

tav

porcine ro

d eosin

in an

o

Hematoxyl

by columnar absorptive cells.

; bar = 100 um.

stain

 

.- 10.5
1.11. e
val. can... . .2 M
I I -l U I
i’lto‘J’. 'u’
,_!

 

Figur

pig 1

porci

and l

Hemat

80

 

Figure 4. Section of small intestine from the DU of a

pig 168 hours after inoculation with the SDSU strain of
porcine rotavirus. Villi are short and fused (arrows)

and lined by low to tall columnar absorptive cells.

Hematoxylin and eosin stain; bar — 100 um.

 

 

 

 

Pig 1
Porci
fused

100 u

81

 

Figure 5. Scanning electron micrograph of the DU of a
pig 168 hours after inoculation with the SDSU strain of
porcine rotavirus. The tips of severely shortened and
fused villi have areas of exposed lamina propria. Bar =

100 um.

 

 

Figu:
pig 2
POrc:
Ville
lOO~

82

 

ine from the DU of a

f small intest

ion 0

Sect

Figure 6

lation with the OSU strain of

pig 168 hours after inocu

tologically normal

Notice the his

tavirus.

porcine ro

in and eosin stain; bar =

Hematoxyl

villous structure.

100 um.

 

 

 

 

 

Figure 7
big 168 ‘
Porcine

Control-

Seen (ar

83

 

Figure 7. Scanning electron micrograph of the DU of a
pig 168 hours after inoculation with the OSU strain of
porcine rotavirus. Villous structure is similar to

control villi. Epithelial cell extrusion zones can be

seen (arrows). Bar = 100 um.

 

 

 

 

Smal
principal
hours PI,
infected
crypts in
deeper in
hours PI
at all si
of pigs i
significa
crypt dep

Fusi
inoculate
(Tables 7
intestine
numerous
were Obse
with the
with the
The smal]
Villous f

Cont
Cells in
11). The
in the dj
aged. C3

rOtaviruE

 

84

Small intestinal crypt depths in control and
principal pigs were not significantly different at 24
hours PI, except for the crypt depths in the UI of pigs
infected with the SDSU strain (Table 6). At 72 hours PI,
crypts in pigs infected with the OSU or SDSU strains were
deeper in the UJ-LI than those of controls and by 168
hours PI this difference in crypt depths was significant
at all sites. Crypts at each site in the small intestine
of pigs infected with the SDSU strain 168 hours PI were
significantly deeper, except in the L1, when compared to
crypt depths in pigs infected with the OSU strain.

Fusion of villi in the small intestine of rotavirus
inoculated pigs was observed 24, 72, and 168 hours PI
(Tables 7, 8, and 9). All of the segments of small
intestine were affected, but fused villi were most
numerous in the UJ-UI (Figures 8, 9, and 10). Fused villi
were observed more frequently in the DU of pigs inoculated
with the SDSU strain (9 of 9 pigs) than in pigs inoculated
with the OSU strain (2 of 9 pigs) at 72 and 168 hours P1.
The small intestine of control pigs had a few areas of
Villous fusion 24 and 72 hours PI.

Control pigs had vacuolated absorptive epithelial
cells in the UJ—LI and less commonly in the DU (Figure
11). The amount of epithelial cell vacuolation increased
in the distal small intestine and decreased as the pigs
aged. Cytoplasmic vacuolation in the small intestine of

rotavirus infected pigs, particularly at 168 hours PI, was

 

 

 

Table 6.
gnotobioti
porcine rc

_________
Hours
after

inoculatie

24

Dat
the OSU
in Which

V
P<0.0S.

bELWEQHK

Table 6.

85

Crypt depths (pm) in the small intestines of

gnotobiotic pigs infected with the OSU or SDSU strains of
porcine rotavirus

 

 

 

Hours Treat-
after ment Intestinal site
inoculation group DU UJ LJ MI UI L1
24 Control 97 98 9o 77 91 85
$6.8 $0.8 $2.6 $11.6 $0.5 $0.9
osu 94 89 92 75 74 8o
$3.4 $1.4 $6.2 $0.6 $10.0 $0.5
snsu 85 82 71 70 66a 70
$1.6 $1.1 $1.4 $0.5 $2.4 $1.8
72 Control 120 105 93 93 89 101
$1.5 $1 7 $0.0 $0.1 $3.9 $0.6
030 122 122 135 143a 1288 121
$4.4 $5 1 $5.6 $1.5 $2.1 $3.0
snsu 125 130 121 118 124a 109
$0.0 $2 1 $0.6 $9.4 $3.7 $6.0
168 Control 113 107 101 102 89 98
$4.7 $6.9 $4.2 $1.4 $2.1 $8.3
osu 1518’b 165"?"b 149"?"b 153a"b 139a;b 136a
$2.9 $2.0 $4.3 $9.8 $4.3 $3.7
sosu ISIa’b 1898 b 1935"b 179a"b 1605"b 14oa
$4.3 $2.4 $6.9 $0.8 $3.1 $1.1

 

Data are expressed as the means i SE (N =
the OSU and SDSU groups 24 and 168 hours after inoculation

in which N =

25).

20 except for

a Values significantly different from control values

p<0.05.

b Significant difference (p<0.05) in crypt depth
between the OSU and SDSU inoculated pigs.

 

 

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Figure
from th
SDSU st

Stain;

89

 

Figure 8. Villous fusion in a section of small intestine
from the LJ of a pig 72 hours after inoculation with the
SDSU strain of porcine rotavirus. Hematoxylin and eosin

stain; bar = 10 um.

 

 

 

Figure

intesti
inocula
Loss of
of dema
exPosed

(arrow)

9O

‘9‘» ‘

u.
~

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5...
3‘ "j

w '

v¢_
.
‘I

I .

t

b
(4’ 1‘
r'
1' L
r F' v

 

Figure 9. Scanning electron micrograph of small
intestine from the LJ of a pig 168 hours after
inoculation with the SDSU strain of porcine rotavirus.
Loss of absorptive cells creates a sharp line

of demarcation (arrowheads) bordering an area of
xposed lamina propria. An area of villous fusion

(arrow) can be seen. Bar = 10 um.

 

 

Figure 1
168 houx
POrCine

100 um.

91

 

Figure 10. Villous fusion in a section of UJ from a pig
168 hours after inoculation with the OSU strain of

porcine rotavirus. Hematoxylin and eosin stain; bar —

100 um.

 

 

 

Figure 1
the LI (
media a]

bar = 1(

 

92

rm

 

an": :9

Q” vi

1..l.4¢o swan)
Mossy

o!&aau'm_

v

 

Us:

Most of the absorptive cells lining villi in

Figure 11.

the LI of a control pig 24 hours after inoculation with

Hematoxylin and eosin stain;

media are vacuolated (+4).

= 10 um.

bar

 

 

 

much les
7, 8, at
Ror
cells or
observed
pigs at
much pig
detachme
with exp
numerous
SDSU str
adjacent
disorgan:
microvili
microvilj
detachmex
With the
hours P1
With the
at 72 and
inoculate
cell deta
uncommon .
time. C01
or exposec
Immat

93

much less conspicuous (Figure 12) than in controls (Tables
7, 8, and 9).

Rounding and detachment of absorptive epithelial
cells on the tips and sometimes the sides of villi were
observed in the small intestine of rotavirus infected
pigs at 24 hours PI (Table 7). At this time, there was
much pig-to—pig variation in the amount of cell ’
detachment. Rounding and detachment of absorptive cells
with exposure of the underlying lamina propria were most
numerous in the UJ—UI of pigs infected with the OSU or
SDSU strains 72 hours PI (Table 8). Microvilli in areas
adjacent to rounded or detached absorptive cells were
disorganized or reduced in number. Areas devoid of
microvilli were often sharply demarcated from normal
microvilli on the same absorptive cell. Absorptive cell
detachment was observed in the duodenum of pigs infected
with the OSU strain 24 hours PI, but not at 72 or 168
hours PI (Tables 7, 8, and 9). In contrast, pigs infected
with the SDSU strain had cell detachment in the duodenum
at 72 and 168 hours PI, but not at 24 hours. Pigs
inoculated with the OSU strain did not have epithelial
cell detachment 168 hours PI, and cell detachment was
uncommon in pigs infected with the SDSU strain at this
time. Control pigs did not have areas of cell detachment
or exposed lamina propria (Figures l3, l4, and 15).

Immature villous epithelium lining the small

intestinal villi of rotavirus infected pigs was observed

 

 

Figure
cells
after

rotavj

94

  

-- x‘ i . I
' “$6. ‘1'! ‘5’?!
Figure 12. Immature cuboidal to low columnar absorptive
cells line an atrophic villus in the UJ of a pig 24 hours

after inoculation with the OSU strain of porcine

rotavirus. Hematoxylin and eosin stain; bar = 10 um.

 

 

 

Figure
POSt-iI

bar = j

 

95

 

Figure 13. Duodenum of a control pig killed at
post—inoculation hour 24. Hematoxylin and eosin stain;

bar = 100 um.

 

96

A

 

L

Figure 14. Scanning electron micrograph of the duodenum

of a control pig at post—inoculation hour 24. Long
slender villi have prominent transverse fissures

(arrows). Bar = 100 um.

 

97

 

Figure 15. Scanning electron micrograph of the lower
ileum of a control pig at post—inoculation hour 24.

Villi are broader in the lower ileum when compared to
villi in the duodenum (Figure 14). Enterocytes bulge
into the intestinal lumen giving a cobblestone appearance

to the mucosal surface. Bar = 100 um.

 

 

 

 

at 24 a1
majoritj
epithel:

As:
intestil
(Figure
transve1
of cont]
(Figure
changes
lung, 1:
tonsil,

infectec

of rote
in ente
small j
€ntero<
MI, LJ,
iDOCUlg
was det
72 hon}
Strain,

the DU

hOUIS,

98

at 24 and 72 hours PI (Figure 12). By 168 hours, the
majority of Villi were lined by low to tall columnar
epithelial cells (Tables 7, 8, and 9).

Aside from the occasional fused villi, small
intestine of control pigs was histologically normal
(Figure 13). Villi were long and slender with prominent
transverse fissures (Figure 14). Villi in the lower ileum
of controls were broader and had prominent enterocytes
(Figure 15) and numerous goblet cells. Histopathologic
changes were not observed in the cecum, colon, rectum,
lung, liver, heart, pancreas, spleen, brain, thymus,
tonsil, and mesenteric lymph node of control or rotavirus
infected pigs.

Immunofluorescence

Pigs inoculated with either the OSU or SDSU strains
of rotavirus had finely granular cytoplasmic fluorescence
in enterocytes lining the sides and tips of villi in the
small intestine (Tables 10 and 11). Immunofluorescent
enterocytes were most consistently detected in the UJ,
MI, LJ, UI, and LI of pigs 24 and 72 hours after
inoculation with either rotavirus strain. Viral antigen
was detected in the DU of pigs 24 hours (5 of 5 pigs) and
72 hours PI (2 of 4 pigs) after inoculation with the OSU
strain. Immunofluorescent cells were seldom observed in
the DU of pigs 24 hours (1 of 5 pigs) or 72 hours (0 of 4
pigs) after inoculation with the SDSU strain. By 168

hours, immunofluorescence was rarely observed in the

 

 

 

Table l(
gnotobic
porcine

 

 

Intestir
site

DU
UJ
LJ
MI
UI

LI

 

* .
specific
separate

99

Table 10. Immunofluorescence in the small intestine
gnotobiotic piglets infected with the SDSU strain of
porcine rotavirus

of

 

 

 

Intestinal Hours after inoculation

site 24 12

DU -, -, -, +, -* -, -, -, - -, —, -, -,

UJ -, +, +, +, — +, —, +, — —, —, -, —,

LJ -, +, +, +, - +, +, +, + -, —, -, —,

MI -, —, +, +, - +, +, +, + —, —, -, -,

UI +, —, -, +, - +, +, +, — -, -, -, -,

LI +, —, +, +, - +, —, -, - -, -, -, —,
* - = No specific fluorescence for rotavirus; + =

specific immunofluorescence for rotavirus; a comma
separates results obtained from each pig.

 

 

 

Table l
gnotobi
porcine

 

 

 

Intesti
site

DU
UJ
LJ
Ml
Ul
LI

 

*
specif
separa

100

Table 11. Immunofluorescence in the small intestine of
gnotobiotic piglets infected with the OSU strain of
porcine rotavirus

 

 

 

 

Intestinal Hours after inoculation

site __ 72

DU +, +, +, +, +* -, -, +, + -, -, -, -, -
UJ +, -, -, +, + +, +, +, + -, -, -, —, -
LJ +, +, -, +, + +, +, +, + -, -, -, —, —
MI +, —, +, +, + +, -, +, + -, —, -, -, —
UI +, -, +, +, + +, —, +, + -, -, —, -, —
L: +, -, +, +, + +, —, +, + -, —, +, —, —

* - = No specific fluorescence for rotavirus; + =

specific immunofluorescence for rotavirus; a comma
separates results from each pig.

 

 

 

 

small i
strain.
A
fluores
hours a
additic
contail
fluore:
inocul
immuno
of one
S
frozen
lymph
from c
immunc

antig«

101

small intestine of pigs infected with either virus
strain.

A few surface enterocytes with cytoplasmic
fluorescence were observed in the cecum from one pig 24
hours after inoculation with the OSU strain. Two
additional sections of cecum from this pig did not
contain immunofluorescent cells. Specific immuno-
fluorescence was not seen in the cecal mucosa from pigs
inoculated with the SDSU strain, but a few detached
immunofluorescent cells were observed in the cecal lumen
of one pig 24 hours after inoculation.

Specific immunofluorescence was not observed in
frozen sections of stomach, tonsil, spleen, mesenteric
lymph node, colon, or rectum. Frozen tissue sections
from control pigs did not have specific
immunofluorescence when stained for porcine rotaviral
antigen.

Electron Microscopic Examination of Feces

Feces collected from control and principal pigs
before inoculation did not contain viral particles. At
24 hours PI, most virus—inoculated pigs had +2 to +3
rotavirus particles in their feces (Table 12). Two pigs,
one inoculated with the OSU strain and another with the
SDSU, did not have clinical signs of diarrhea at 24 hours
PI, but rotavirus particles were identified in cecal
contents collected at necropsy. Cecum and colon in these

pigs were dilated with fluid yellow contents indicating

 

 

Table 12
examinat
infectec

 

M
Neg

+1

+2

+3

+4

Total n
of Spec

‘-

a
obtaine
b

C

Was est
Squarej

102

Table 12. Results of direct electron microscopic
examination of feces or cecal content from pigs
infected with the SDSU or OSU strain of porcine rotavirus

 

. a
Number of spec1mens

 

 

24 hrsb M 168 hrs
EM score we M 9_s_U M gs_U fig
Neg 6 6 0 l 2 1
+1 0 l l l 0 2
+2 3 4 l 3 3 2
+3 5 3 7 4 0 0 1
+4 9 9 2 2 .0. 9
Total no.
of specimens l4 l4 9 9 5 5

 

a Viral particles were not observed in feces
obtained before rotavirus inoculation.

b Hours after inoculation.

C The number of rotavirus particles per grid square.
was estimated and rated as +1 (0 to 5 virus particles/grid
square), +2 (6 to 20), +3 (21 to 100), or +4 (>100).

 

 

that d
allowe
24 hou
remain

V
virus
feces
strait
72 box
fluid
infect
to de<
seen :
PI. 1
hours
compl.
parti
colle
inocu

were

103

that diarrhea would have occurred had these pigs been
allowed to survive. Virus particles were not detected at
24 hours after inoculation in the feces from the
remainder of the non—diarrheic pigs.

Virus particles had been seen in the feces of all
virus infected pigs by 72 hours after inoculation. The
feces of one diarrheic pig inoculated with the SDSU
strain contained virus particles at 24 hours, but not at
72 hours PI. The feces from this pig were becoming less
fluid so it may have been recovering from the rotavirus
infection.

The number of rotavirus particles in the feces began
to decrease by 168 hours PI. Virus particles were not
seen in the feces of 3 pigs (2 OSU, l SDSU) at 168 hours
PI. Feces or cecal content collected at 24, 72, or 168
hours PI contained approximately equal numbers of
complete and incomplete virus particles. Rotaviral
particles were not observed in the fecal samples
collected from control pigs 24, 72, or 168 hours after
inoculation. In general, the virus excretion patterns

were similar for both rotavirus strains.

 

 

 

 

 

F
gnotob
strain
signs
caused
duoden
with t
freque
strain

V
strain
findin
McAdar
after
rEport
colost
McNult
Graham
report
Narita
r0tavi-

(1980)

DISCUSSION

Findings in this study indicate that in 3-day-old
gnotobiotic pigs serotype 1 (OSU) and serotype 2 (SDSU)
strains of porcine rotavirus cause different clinical
signs and histopathologic lesions. The SDSU strain
caused more severe villous atrophy and fusion at the
duodenal—jejunal flexure when compared to pigs inoculated
with the OSU strain. Pigs infected with the OSU strain
frequently vomited whereas pigs infected with the SDSU
strain did not vomit.

Vomiting by gnotobiotic pigs infected with the OSU
strain but not the SDSU strain is consistent with the
findings of other investigators (Theil et al, 1978;
McAdaragh et al, 1980; Graham et al, 1984). Vomiting
after infection by some strains of rotavirus has been
reported in conventional (Bohl et al, 1978),
colostrum—deprived (Lecce et al, 1976; Pearson and
McNulty, 1977), and gnotobiotic pigs (Theil et al, 1978;
Graham et al, 1984). In other instances, no vomiting was
reported (Tzipori et al, 1978; McAdaragh et a1, 1980;
Narita et a1, 1982). In fact, a strain of porcine
rotavirus (B—3l7) studied by Torres—Medina and Underdahl

(1980), has never been observed by these investigators to

104

 

 

 

cause v
in this
occurre
differe:
infecti«
caused l
occurre<
2 than 5
More rec
people,
vomiting
This is
al (l984
Signs ca
The OSU
Of human
all othe
Strain o
31, 1984
Used in .
VOm:
gaStroin1
chemorem
Present 5
severe ir
the SDSU

FuthErnor

 

 

105

cause vomiting in gnotobiotic pigs. Therefore, findings
in this and other studies suggest that differences in the
occurrence of vomiting are related to rotavirus strain
differences. Epidemiologic studies of human rotavirus

infections also indicate differences in clinical signs

 

caused by different strains of rotavirus. Vomiting
occurred more frequently following infection by serotype
2 than serotype 1 human rotavirus (Yolken et al, l978b).
More recently, Uhnoo and Svenson (1986), reported that in
people, subgroup II rotavirus strains caused diarrhea and
vomiting of greater severity when compared to subgroup I.
This is in contrast, however, to the findings of White et
a1 (1984), who could find no differences in clinical

signs caused by different subgroups of human rotavirus.

 

The OSU strain of porcine rotavirus and the DS—l strain
of human rotavirus are classified as subgroup I, whereas
all other human rotavirus strains and the Gottfried
strain of porcine rotavirus are subgroup II (Hoshino et
al, 1984). The subgroup specificity of the SDSU strain
used in the present study has not yet been identified.

Vomiting may be caused by irritation of the upper
gastrointestinal tract or by stimulation of
chemoreceptors in the medulla (Ganong, 1983). In the
present study, mucosal histopathologic lesions were most
severe in the upper small intestine of pigs infected with
the SDSU strain, yet none of these pigs vomited.

Futhermore, rotaviral antigen was not detected by IF in

 

the stoma
strain no
in other
1977; The
associate
mechanism
tract.

In t
the effec
course of
strains w
temperatu
were rand
Pig-tO-pi
infection
underwent
inoculati
Prevent a
Occur aft

Uude
infected
neCIOPSiE
SDSU virL
other in\
1980). I

InfectioT

l978)’

C(

106

the stomachs of pigs infected with either rotaviral
strain nor has it been detected in the stomachs of pigs
in other studies (McNulty et al, 1976a; Davidson et al,
1977; Theil et al, 1978). This suggests that vomiting
associated with rotavirus infection may be initiated by
mechanisms other than irritation of the gastrointestinal
tract.

In the present study, attempts were made to minimize
the effects of environmental and host factors on the
course of the rotaviral infections. The OSU and SDSU
strains were studied while keeping diet, environmental
temperature, and housing constant. Experimental animals
were randomly assigned to treatment groups to account for
pig-to-pig variation in susceptiblity to rotavirus
infection. Furthermore, the OSU strain of rotavirus
underwent four gnotobiotic pig passages prior to
inoculation of experimental animals. This was done to
prevent attenuation, i.e. loss of virulence which may
occur after passage in cell culture (Bohl et al, 1984).

Under these experimental conditions, all of the pigs
infected with the OSU or SDSU strains survived until
necropsied. This lack of mortality caused by the OSU and
SDSU virus strains is in agreement with the findings of
other investigators (Theil et al, 1978; McAdaragh et al,
1980). Mortality associated with porcine rotavirus
infection has been reported in conventional (Bohl et al,

1978), colostrum—deprived (Lecce et al, 1976; McNulty et

 

 

al, 19]
Narita
experit
host 01
differe
enviror
infect:
rotaviJ
Woode,
Newsome
rotavi:
colost
to l- (1;
used h
(persm
When 31
Virulm
factor

D
are ev
hetero
1986).
of rot
(bOVin
HOWeve
rOtavi

into a

 

 

107

a1, 1976), and gnotobiotic pigs (Tzipori et al, 1978;
Narita et al, 1982). However, in these studies
experimental conditions differed so it is possible that
host or environmental factors and not rotavirus strain
differences influenced the observed mortality. Age,
environmental temperature, and intercurrent bacterial
infection have been shown to influence the severity of
rotavirus infections (Woode et al, 1976b; Crouch and
Woode, 1978; Tzipori et al, 1980b; Torres-Medina, 1984;
Newsome and Coney, 1985). When a strain of porcine
rotavirus which caused high mortality in l-day—old
colostrum-deprived conventionally-derived pigs was given
to l-day—old gnotobiotic pigs kept under the conditions
used in this experiment, no mortality was observed
(personal data). Therefore, caution must be exercised
when suggesting that field strains of rotavirus differ in
virulence, particularly because environmental and host
factors may influence the severity of the disease.
Differences in virulence between rotavirus strains
are evident when rotaviruses are inoculated into a
heterologous host (Bridger and Brown, 1984; Offit et al,
1986). For instance, the SA—ll strain (primate origin)
of rotavirus was more virulent than the NCDV strain
(bovine origin) when inoculated orally into newborn mice.
However, evidence indicating that field strains of
rotavirus are more virulent than others when inoculated

into a homologous host such as in the present study is

 

 

 

 

 

 

scant
(Cast1
(Bridg
Carpic
indica
chara<
isolat
differ
ligate
The a1
of vi]
virule
it may
equate
villor
calves
EXperi
intrav
Chlora
shown
Chlora
have i

I
the 08
than t
Theil

and Va

108

scant (Woode, 1979) and often based on clinical reports
(Castrucci et a1, 1983) or personal communications
(Bridger, J. C. in Reynolds et al, 1985). A report by
Carpio et al (1981b) is frequently cited as evidence
indicating that field isolates with different virulence
characteristics exist. In their report, four field
isolates of bovine rotavirus consistently caused
different amounts of villous atrophy when inoculated into
ligated intestinal loops in colostrum-deprived calves.
The authors concluded that the difference in the severity
of villous atrophy corresponded to differences in
virulence among the rotavirus isolates tested. However,
it may be incorrect to assume that villous atrophy is
equated with virulence. Reynolds et a1 (1985) observed
villous atrophy in clinically normal (non-diarrheic)
calves infected with rotavirus. The calves used in the
experiment by Carpio et al (1981b) were maintained on
intravenous chloramphenicol throughout the experiment.
Chloramphenicol administered to healthy calves has been
shown to induce diarrhea (Huffman et al, 1981).
Chloramphenicol or the rotavirus strains, or both, may
have influenced the amount of villous atrophy observed.
The incubation periods after inoculating pigs with
the OSU or SDSU strains, were longer and more variable
than those previously reported (Crouch and Wood, 1978;
Theil et al, 1978; McAdaragh et al, 1980). The prolonged

and variable incubation periods were probably due to the

 

 

 

lower c
Previor
diarrhe
suspene
1978; l
rotavim
incubat
do not
et al,
Va
respon:
atrophj
OSU an.
infect
but 10:
infect
signif
strain
hours
becaus
differ
times.
R
When C
Variab
and in

Variat

 

109

lower dosage of rotavirus used in the present study.
Previous investigations on the pathogenesis of rotaviral
diarrhea caused by the OSU or SDSU strains used 102
suspensions of intestinal or fecal filtrate (Theil et al,
1978; McAdaragh et al, 1980). Decreasing dosages of
rotavirus have been shown to increase the length of the
incubation period before the onset of clinical signs but
do not affect the severity of the clinical disease (Woode
et al, 1976b).

Variation in incubation periods may have been
responsible for the differences in the amount of villous
atrophy observed in the UJ—UI of pigs infected with the
OSU and SDSU strains. Villi in the UJ-UI of pigs
infected with the OSU strain were shorter at 72 hours PI
but longer at 168 hours PI when compared to those of pigs
infected with the SDSU strain. Glandular crypts were
significantly deeper in pigs infected with the SDSU
strain than in pigs infected with the OSU strain 168
hours PI. Pigs infected with the OSU or SDSU strain,
because of different incubation periods, may have been at
different stages of the infection when studied at these
times.

Rotavirus—infected pigs had reduced body weights
when compared to controls. Weight gains were highly
variable even when pigs were kept in the same isolator
and inoculated with the same pool of virus. Pig—to—pig

variation in body weight gain also was reported by Woode

 

 

(1979)
R
but ma
instan
intest
rotavi
protei
(Offit
qualit
enzyme
of the
1976;
affect
recent
primat
2.0, l
inact:
1985)
intes'
entert

rotav:

infec
1972)
Susse
With

the 1

 

 

110

(1979).

Reasons for the pig—to-pig variation are not known,
but may be related to host physiological factors. For
instance, the availability of trypsin in the small
intestine might affect the expression of virulence by

rotaviruses. It is now clear that a trypsin-sensitive

 

protein plays a major role in gastrointestinal virulence
(Offit et al, 1986). It has been suggested that
qualitative or quantitative differences in intestinal
enzymes (trypsin) may be responsible for the resistance
of the human neonate to rotaviral diarrhea (Holmes et al,
1976; McClean and Holmes, 1981). Gastric pH might also
affect the clinical expression of rotavirus infection. A
recent report indicated that three bovine and several
primate-origin rotaviruses are rapidly inactivated at pH
2.0, but inactivated at a much slower rate at pH 3.0;
inactivation at pH 4.0 was minimal (Weiss and Clark,
1985). The effect of host factors such as gastric pH,
intestinal trypsin content, and the availability of
enterocyte receptors on the expression of virulence by
rotaviruses deserves further study.

Serum biochemical changes that occur in rotavirus
infected pigs seldom have been reported (Mouwen et al,
1972). In the present study, elevations in BUN which
suggested dehydration were detected in pigs inoculated
with the OSU or SDSU strains. However, as evidenced by

the large standard deviation of the mean, elevations in

 

 

BUN wer
pathoge
has bee
protei:
Serum 1
pigs wl
presen'
condit
to nor
trigly
of the
E
in the
regart
This ;
total
is is
more
in ch

(Tall

Straf
in t1
ShOr'
isol
give

freq

lll

BUN were detected inconsistently. Another enteric
pathogen of pigs, transmissible gastroenteritis virus,
has been reported to cause elevations in BUN and total
protein (Cornelius et a1, 1968; Drolet et al, 1984).
Serum triglycerides were decreased in rotavirus infected
pigs when compared to controls. Higher levels of fat are
present in the feces of piglets with "white scours", a
condition presumably caused by rotavirus, when compared
to normal pigs (Mouwen et al, 1972). Decreased serum
triglycerides may be related to the malabsorptive state
of the damaged small intestine.

Serum electrolyte values of rotavirus infected pigs
in the present study did not differ from the controls
regardless of the strain of rotavirus used as inoculum.
This finding, coupled with the elevations in BUN and
total protein suggests that, when dehydration occurs, it
is isotonic. Isotonic dehydration also has been reported
more frequently than hypertonic or hypotonic dehydration
in children hospitalized with rotavirus infection
(Tallette et al, 1977; Rodriguez et a1, 1977).

An important finding in this study was that the SDSU
strain of rotavirus produced more severe villous atrophy
in the upper small intestine. Villi were significantly
shorter in the DU of pigs inoculated with the SDSU
isolate at 72 and 168 hours PI when compared to pigs
given the OSU strain. Fusion of villi also was more

frequently seen in the duodenum of pigs infected with the

 

 

SDSU st
is the
mammali
histope
differe
P:
villou
instan
fusion
wherea
villou
McAdaI
used V
strait
B-3l7
and f
EXpla

the f

Pigs

hours
were

Stra:
SDSU

dete

the

inte

 

 

 

 

112

SDSU strain than pigs infected with the OSU strain. This
is the first time that different serotypes of the same
mammalian rotavirus have been shown to cause
histopathologic lesions of different severity in
different sites in the small intestine.

Previous investigators disagreed about the extent of
villous atrophy in the upper small intestine. For
instance, McAdaragh et a1 (1980) observed atrophy and
fusion of villi in the duodenum as early as 24 hours PI
whereas Torres-Medina and Underdahl (1980) found no
villous atrophy in the duodenum. In the experiment by
McAdaragh et a1 (1980), the SDSU strain of rotavirus was
used whereas Torres-Medina and Underdahl used a field
strain (serotype unknown) of porcine rotavirus designated
B-3l7. Differences in the severity of villous atrophy
and fusion in the upper small intestine might be
explained by rotavirus strain differences as suggested by
the findings in the present study.

Viral antigen was consistently detected in the DU of
pigs infected with the OSU but not SDSU strain 24 and 72
hours PI. This was unexpected since histologic lesions
were most severe in the DU of pigs infected with the SDSU
strain. Three of the pigs killed 24 hours PI with the
SDSU strain did not have diarrhea. Viral antigen was not
detected in the small intestine from 1 of these pigs and
the other two only had immunofluorescence in a few

intestinal segments. It is possible that duodenal

 

 

 

 

entero:
became
could'
M
differ
rotavi
predil
pathog
132 (r
intest
]
McNul'
some
clini
The G
isola
sugge
Mixec
rota\
Rodrf
infec
may .
eith
8988
Pore
atrc

Snal

 

113

enterocytes in the pigs infected with the SDSU strain
became infected between 24 and 72 hours PI, sloughed, and
could no longer be detected at 72 hours PI.

McNulty et al (1983) were the first to report
different intestinal tropisms for different strains of
rotavirus. Avian rotavirus strain ch 1 (group A) had a
predilection for the duodenal mucosa of specific
pathogen—free chickens, whereas avian rotavirus strain ch
132 (non—group A) replicated best in the mid-small
intestine.

Based on the differences in intestinal tropism,
McNulty et a1 (1983) suggested that mixed infections by
some strains of avian rotavirus may cause more severe
clinical disease than infection by either virus alone.
The Gottfried strain of porcine rotavirus was originally
isolated from a mixture of rotaviruses (Bohl et al, 1984)
suggesting that mixed infections occur naturally in pigs.
Mixed infections by different electropherotypes of human
rotavirus also have been reported (Spencer et a1, 1983;
Rodriguez et al, 1983). It is possible that mixed
infections in pigs by two or more strains of rotavirus
may cause more severe clinical disease than infection by
either strain alone. The findings in the present study
suggest that mixed infections by serotypes l and 2 of
porcine rotavirus might result in more severe villous
atrophy with the serotype 2 rotavirus producing the upper

small intestinal lesion. This hypothesis deserves to be

 

 

tested
H
findin
(Pears
Woode,
Underd
report
first
rotavi'
McAdar.
villou
contra
after :
porcine
In the
PI, so
al (19?
Gr
lesions
than tl
lamina
Pigs We
reporte
In
and cec

and has

 

 

 

114

tested.

Histopathologic and scanning electron microscopic
findings were similar to those reported previously
(Pearson and McNulty, 1977; Theil et al, 1978; Crouch and
Woode, 1978; McAdaragh et al, 1980; Torres-Medina and
Underdahl, 1980; Narita et al, 1982). However, previous
reports disagree about the time at which villous fusion
first appears after inoculating gnotobiotic pigs with
rotavirus. Theil et a1 (1978), using the OSU strain, and
McAdaragh et al (1980), using the SDSU strain, observed
villous fusion in gnotobiotic pigs by 24-48 hours PI. In
contrast, villous fusion was first observed 4.5 days
after inoculating pigs with a field strain (B-317) of
porcine rotavirus (Torres-Medina and Underdahl, 1980).

In the present study, villous fusion was seen at 24 hours
PI, so results are in agreement with findings by Theil et
a1 (1978) and McAdaragh et al (1980).

Gnotobiotic pigs in the present study did not have
lesions or specific immunofluorescence in organs other
than the small intestine. Immunofluorescence in the
lamina propria and mesenteric lymph node of gnotobiotic
pigs was not observed in the present study, but has been
reported by others (Theil et al, 1978).

Immunofluorescence was observed in the cecal mucosa
and cecal lumen of one pig infected with the OSU strain
and one pig infected with the SDSU strain at 24 hours PI

and has been observed in the cecum of gnotobiotic pigs by

 

 

other i
1982).
immunof
intesti
and MCI
1980).
Tl
the OSI
simila
al, 19
Torres
of rot
were s
of rot
could
A
(Torre
detect
a long
fluOn
infec
cells
(Thei
fluor
Peric

Strai

115

other investigators (McAdaragh et a1, 1980; Narita et al,
1982). In general, the absence of histologic lesions and
immunofluorescence in organs other than the small
intestine is consistent with previous reports (Pearson
and McNulty, 1977; Theil et al, 1978; McAdaragh et a1,
1980).

 

The virus-excretion patterns for pigs infected with
the OSU and SDSU strains in the present study were
similar and in agreement with previous reports (Lecce et
a1, 1976; Bohl et a1, 1978; Tzipori and Williams, 1978;
Torres-Medina and Underdahl, 1980). The greatest number
of rotavirus particles as detected by electron microscopy
were shed by gnotobiotic pigs at 72 hours PI. The number
of rotavirus particles was reduced, or virus particles
could not be detected 7 days after infection.

As has been shown by other investigations
(Torres—Medina and Underdahl, 1980), virus could be
detected by electron microscopic examination of feces for
a longer period of time when compared to immuno-
fluorescence of the small intestine. Enterocytes
infected by rotaviruses desquamate so immunofluorescent
cells can only be detected up to 96 to 168 hours PI
(Theil et a1, 1978; McAdaragh et a1, 1980). Immuno—
fluorescent cells may be detected for even shorter
periods of time after infection by some porcine rotavirus

strains (Theil et al, 1985).

 

 

 

Tl
seroty;
Dakota
determi
virus :
lesion
seroty}

F
inocul
homoge
the Oh
Dakota
rotavi
Mortal
monitc
inocul
and 16
the 3
killer
c011e<
immuH(

eXamit

 

SUMMARY

The purpose of the present study was to compare
serotypes 1 (Ohio State University strain) and 2 (South
Dakota State University strain) of porcine rotavirus to
determine if specific clinical signs, mortality rates,
virus shedding patterns, virus antigen distribution, or
lesion distribution and severity were related to the
serotype of rotavirus with which pigs were infected.

Forty hysterotomy derived gnotobiotic pigs were
inoculated orally at 3 days of age with 2 ml of
homogenate containing 105 pig-infectious dose50 of either
the Ohio State University (OSU) strain or the South
Dakota State University (SDSU) strain of porcine
rotavirus. Controls were inoculated with media only.
Mortality, clinical signs, and body weights were
monitored daily. Five pigs in each of the virus
inoculated groups and 4 control pigs were killed at 24
and 168 hours after inoculation. Four pigs in each of
the 3 experimental groups (OSU, SDSU, and control) were
killed 72 hours after inoculation. Specimens were
collected at necropsy for serum chemistry, histologic,

immunofluorescent, and scanning electron microscopic

examinations.

 

 

 

Al
strains
througf
SDSU st
hours t
observ«
OSU st
did no
strain
but tt
given
immun<
given
after
duode
and i
and f
flexx
comp;
Alth.
the
atro
3-da
Cont

of}

 

117

All of the pigs inoculated with the OSU or SDSU
strains survived. Control pigs remained healthy
throughout the study. Pigs inoculated with the OSU or
SDSU strains developed diarrhea 19-48 hours and 24—54
hours after inoculation, respectively. Vomiting was
observed in 5 of 14 (362) of the pigs infected with the
OSU strain whereas pigs inoculated with the SDSU strain
did not vomit. Pigs inoculated with the OSU or SDSU
strains had reduced weight gain compared to control pigs,
but there was no difference in weight gain between pigs
given the OSU or SDSU strains. Results of
immunofluorescent examinations were similar for pigs
given either rotavirus strain except that at 24 hours
after inoculation, viral antigen was detected at the
duodenal-jejunal flexure in 5/5 pigs given the OSU strain
and in 1/5 pigs given the SDSU strain. Villous atrophy
and fusion were more severe at the duodenal-jejunal
flexure of pigs inoculated with the SDSU strain when
compared to pigs inoculated with the OSU strain.

Although the OSU and SDSU strains did show differences in
the occurrence of vomiting and distribution of villous
atrophy, these strains were equally virulent for
3—day-old gnotobiotic pigs. Therefore, prevention and

control practices must be directed against both serotypes

of porcine rotavirus.

 

 

 

 

 

BIBLIOGRAPHY

 

 

 

Acree
modii
bacte
beef

Adams
the i
of e;
51:35

Argex
C.:
gastt

Bacht
Kinet
infEt
Neon;

l983

Banf:
obse:
mice
1968

Benf:
Shed:
farrt

Benf-

immu:
anti
and‘
1984

Benj
path.
71

Fish
T o

mneo
east

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and
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G. i
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str;

Crm
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Pig
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Davids
Hamilt
conven

Drolet
factor
piglet
virus.

Duhame
experf
calves
Confe:
Chica

Echev
Vibul
Rotav
J Cli

Elden
T—cel
exper
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E18,
SA-ll
1972,

EngL
iden
from
41:7

Engv
assa

Espe
poly
try;

Este
Rot;
43H

Est
enh
mec

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VITA

The author was born in St. Paul, Minnesota, on
October 1, 1953. He completed his high school education
at Columbia Heights, Minnesota. He received the Bachelor
of Science degree in 1976 and the degree of Doctor of
Veterinary Medicine in June, 1978, from the University of
Minnesota.

After one year of private veterinary practice, the
author accepted a position as resident/instructor at
Michigan State University. The residency and a Master of
Science degree in pathology were completed in June, 1982.
In July, 1982, the author began employment as an
assistant professor in the Department of Veterinary
Science at South Dakota State University.

The author was married to Barbara Bjorke Collins in

1974 and they have two sons, Brian and Daniel.

 

 

   

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