A SIMPLE AND RAPID METHOD FOR DETECTION
OF BLUETONGUE VIRUS IN CELL CULTURE USING RT-PCR
By
Afra'a Tajelsir Mohamed Elata
B.V.M, University of Khartoum, 2002
A thesis submitted to the University of Khartoum in fulfillment of the requirements for the Master degree in Veterinary Medicine
Supervisor:
Prof. Imadeldin Elamin Eltahir Aradaib
Department of Medicine, Pharmacology and Toxicology
Faculty of Veterinary Medicine
University of Khartoum
April, 2007
i
DEDICATION
TO THE SOUL OF MY FATHER …..
TO MY MOTHER …..
TO MY BROTHERS AND SISTERS …..
WITH ALL MY LOVE
ii
Acknowledgements
My gratitude and special thanks to Prof. Imadeldin
Elamin Eltahir Aradaib, Molecular Biology Laboratory,
Department of Medicine, Pharmacology and Toxicology,
Faculty of Veterinary Medicine, University of Khartoum, my
supervisor, for his encouragement, full assistance and
support during the completion of this work. It has been a
privilege and a unique opportunity to work under his
supervision. His suggestions and advices were of great value.
So, I am very indebted to him for his keen interest and
enthusiasm.
I would like to thank my colleagues at the Molecular
Biology Laboratory, namely; Dr. Khairalla Mohamed Saeed,
Ahmed Osman, Imadeldeen Osman, Samah Abdelrahman,
Amel Mahmoud and Tumadur Mohamed for their thorough
and unlimited support and comments.
Thanks are extended to my colleagues at the
Department of Medicine, Pharmacology and Toxicology for
their help and continuous assistance during my work in this
research.
iii
Last, but not least, I must express my gratitude to my
friends, especially Hanan, Rasha, Ranya, Reem, Siham and
Ahmed Omer for being close to me during the hard time of my
work, and for their great help and continuous encouragement.
I am very lucky to have kind friends such like them.
For all of those people, I send my appreciation and
respect.
iv
LIST OF CONTENTS
Page
Dedication……………………………………….. i
Acknowledgements……………………………… ii
List of contents…………………………………... iv
List of Figures…………………………………… vi
Abstract………………………………………….. vii
Arabic abstract………………………………….. ix
CHAPTER ONE: INTRODUCTION AND GENERAL
LITERATURE REVIEW
1.1 Introduction……………………………………... 1
1.2 The bluetongue virus……………………………. 3
1.2.1 Virus classification……………………………… 3
1.2.2 Virus structure…………………………………... 4
1.3 Vector and Transmission……………………….. 5
1.4 Epidemiology and Distribution………………… 8
1.5 Bluetongue disease in Sudan…………………… 10
1.6 Economic Importance…………………………... 11
1.7 Pathology of bluetongue disease………………... 12
1.7.1 Pathogenesis of BTV……………………………. 12
1.7.2 Immune response to bluetongue virus…………. 13
1.7.3 Clinical signs…………………………………….. 14
1.7.4 Post-mortem findings…………………………… 16
1.8 Diagnosis of BTV………………………………... 17
1.8.1 Virus Isolation…………………………………... 17
1.8.1.1 Embryonated Chicken Eggs (ECE)……………. 18
1.8.1.2 Sheep inoculation………………………………... 19
1.8.1.3 Cultured cells……………………………………. 19
v
1.8.2 Virus Identification……………………………... 20
1.8.2.1 Serological techniques…………………………... 20
1.8.2.1.1 Agar gel immunodiffusion (AGID)…………….. 21
1.8.2.1.2 Competitive enzyme-linked
Immuno sorbent assay (c-ELISA)………………
21
1.8.2.2 Virological techniques…………………………... 22
1.8.2.2.1 Virus neutralization test 22
1.8.2.3 Reverse transcriptase-polymerase chain
reaction (RT-PCR)………………………………
22
1.9 Prevention and control 23
CHAPTER TWO: MATERIALS AND METHODS
2.1 Bluetongue virus………………………………... 27
2.2 Virus propagation in tissue culture……………. 27
2.3 Viral nucleic acid extraction from infected cell
monolayers……………………….........................
28
2.4 Primers selection……………………………….. 29
2.5 Reverse transcriptase polymerase chain
reaction (RT-PCR)………………………………
30
2.6 Agarose gel electrophoresis…………………….. 31
CHAPTER THREE: RESULTS………………………………… 36 CHAPTER FOUR: DISCUSSION……………………………… 40 REFFERENCES…………………………………………………. 46
vi
LIST OF FIGURES
Figure 2.1: Thermal cycler TECHNE, TC-41……………………………… 33
Figure 2.2: Electrophoresis apparatus……………………………………… 34
Figure 2.3: Gel documentation apparatus………………………………….. 35
Figure 3.1: Detection of BTV serotypes in infected cell cultures…………. 37
Figure 3.2: Specificity of RT-PCR for BTV………………………………... 38
Figure 3.3: Sensitivity of RT-PCR for BTV………………………………... 39
vii
ABSTRACT
In this study, a reverse transcriptase polymerase chain reaction
(RT-PCR) protocol was evaluated for detection of bluetongue virus
(BTV) RNA in cell culture.
North American BTV serotypes 2, 10, 11, and 13, and Sudanese
BTV serotype 2 were studied. All these serotypes were propagated in cell
culture. RNAs were extracted from these serotypes and then, they were
detected by the described RT-PCR assay, using primers derived from
segment 6 of BTV-17 which codes for non-structural protein 1 (NS1). So,
this NS1 gene was targeted for PCR amplification.
The specific 614 bp PCR product was amplified from all BTV
serotypes used in the study and they were visualized on ethidium
bromide-stained agarose gel.
Amplification product was not detected when the RT-PCR assay
was applied to RNA from epizootic hemorrhagic disease virus (EHDV)
or Palyam virus.
Also this specific 614 bp PCR product was obtained from the
amounts of 1ng, 500pg, 250pg and 125pg RNA from Sudanese BTV
serotype 2.
viii
The results of this study indicated that the described RT-PCR
assay, using the mentioned primers, could be applied for detection of
BTV serogroups.
In conclusion, the described RT-PCR assay could be used as a
simple, rapid, sensitive and specific supportive diagnostic assay to the
current conventional procedures used for detection of BTV.
ix
ملخص األطروحة
في الكشف عن الحمض النووي م تفاعل البلمرة المتسلسل العكسيفي هذه الدراسة تم تقيي
. الريبي لفيروس اللسان األزرق من الزرع الخلوي
لمتوطنة في أمريكا ا13 و 11, 10, 2تمت دراسة األنماط المصلية لفيروس اللسان األزرق
ثارها في كل هذه األنماط المصلية تم إك. في السودان المتوطن2الشمالية و النمط المصلي
الكشف عنه بواسطة تمو من ثم, ص الحمض النووي الريبي منهاالزرع الخلوي و استخل
من جينوم النمط المصلي 6تفاعل البلمرة المتسلسل باستخدام بادئات مشتقة من القطعة رقم
. للفيروس1تين غير التركيبي رقم لفيروس اللسان األزرق و التي تشفر إلنتاج البرو17
زوج قاعدي تم إكثاره من جميع األنماط 614ناتج تفاعل البلمرة المتسلسل البالغ طوله
و تم إظهاره في هالم الجل , المصلية لفيروس اللسان األزرق المستخدمة في الدراسة
. المصبوغ ببروميد االيثيديوم
على ء تفاعل البلمرة المتسلسل العكسيتم إكثار الناتج عند إجرالم ي, في اختبارات الخصوصية
الحمض النووي الريبي المستخلص من فيروس المرض النزفي الوبائي أو من فيروس باليام
. مما يدل على انه خاص فقط بفيروس اللسان األزرق
بي هذا الناتج فائق الخصوصية تم إكثاره من كميات مختلفة من الحمض النووي الري
0.5, نانوجرام1.0: لفيروس اللسان األزرق وهي2المستخلص من النمط المصلي
ة تفاعل البلمرة المتسلسل مما يؤكد حساسي, نانوجرام0.125 نانوجرام و 0.25, نانوجرام
. في الكشف عن فيروس اللسان األزرقالعكسي
x
باستخدام البادئات ل العكسيلسفان نتائج هذه الدراسة توضح أن تفاعل البلمرة المتس, ختاماً
ذو خصوصية و حساسية , سريع, بسيطذكورة أعاله يمكن استخدامه كاختبار تشخيصيالم
.لدعم الطرق التقليدية المستخدمة في تشخيص فيروس اللسان األزرق
1
CHAPTER ONE
Introduction and General Literature Review
1.1. Introduction
Bluetongue (BT) is an infectious, non-contagious arthropod-borne
disease of ruminants, caused by bluetongue virus and transmitted between
vertebrate hosts via the bites of certain species of biting midges of the
genus Culicoides (Mellor, 1990). Bluetongue was first reported in the
year 1881 as a result of the introduction of European breeds of sheep into
southern Africa (Howell and Verwoerd, 1971).
The viral etiology of the disease was demonstrated in 1906 and its
strains have been identified in many tropical and temperate areas of the
world since that time. While the virus is classified antigenically and
taxonomically as bluetongue virus, each serotype is unique and may not
cause the disease (Callis, 1985).
Bluetongue virus (BTV) is a double stranded (ds) RNA orbivirus
of the family Reoviridae (Borden et al., 1971; Fenner et al., 1974; Gould
et al., 1992). BTV naturally infects domestic and wild ruminants,
camelids and some other herbivores such as elephants, but it is almost
exclusively a disease of sheep. In cattle and goats clinical disease is rare,
and, when present, is much milder than in sheep (Verwoerd and Erasmus,
2
1994). Bluetongue disease was thought to be limited to sheep until 1933
when the virus was isolated in South Africa from cattle with clinical signs
similar to those of foot and mouth disease (Bekker et al., 1934).
The virus has a world wide distribution and it exists in at least 25
serotypes (Davies et al., 1992). It occurs in the Americas, Africa, Asia
and Australia, and it can cause an acute, sub-acute, mild or inapparent
disease (Mellor and Wittmann, 2002).
Serotypes 1, 2, 4 and 16 are enzootic in the Sudan, while serotypes
2, 10, 11, 13 and 17 are enzootic in North America (Davies et al., 1992;
Mohammed and Taylor, 1987).
The mortality rate of bluetongue disease vary from 0% to 30%, but
can reach 75% (Mellor et al., 1983) in highly susceptible animals, but
that is dependent upon the serotype involved. The real significance of
bluetongue disease lies in the indirect losses sustained; these include
abortion in pregnant ewes and severe loss of condition during prolonged
convalescence (Tomori et al., 1991).
It has been estimated that BTV alone causes losses to international
livestock trade in excess of US$ 3 billion a year (Tabachnick et al.,
1996). Immunization of susceptible animals requires multivalent vaccines
because bluetongue vaccines are serotype specific.
Diagnostic tests currently used for the detection of BTV involve
the isolation and growth of virus isolates in eggs or sheep, followed by
3
passaging in tissue culture. The virus is characterized using serological
tests such as the agar gel immunodiffusion test or serum neutralization
test. These procedures are time-consuming and may fail to detect low
levels of virus. The use of enzyme-linked immuno-sorbent assay (ELISA)
for the detection of antibodies of BTV in infected animals is faster but
doesn’t confirm recent infection. These traditional methods may require
at least three to four weeks to provide a result. The polymerase chain
reaction technique may be used not only to detect the presence of BTV
but also to serogroup the virus and provide information on the serotype
within a few days (Zientara et al., 2004)
The objective of the present study was to develop a simple, rapid,
sensitive, specific and inexpensive method for detection of bluetongue
virus serogroup using a reverse transcriptase (RT) polymerase chain
reaction (RT-PCR).
1.2. The bluetongue virus
1.2.1. Virus classification
Family: Reoviridae
Contains twelve genera of multi-segmented dsRNA viruses.
Genus: Orbivirus
The members of this genus called ‘orbiviruses’. They are twenty
one species which characteristically have a ten segmented
4
dsRNA genome that is packaged within an icosahedral protein
capsid.
Species: Bluetongue virus
It is the prototype of twenty one different species of the genus
Orbivirus. It has a capsid composed of three distinct protein
layers: the subcore, the core-surface layer and the outer capsid
layer (Mertens et al., 2004).
1.2.2. Virus structure
Bluetongue virus is an icosahedral-shaped particle consisting of a
segmented double-stranded RNA genome, encapsidated in a double-
layered protein coat. Removal of the outer protein layer activates a viral-
associated RNA polymerase which transcribes the ten genome segments
into 10 mRNAs which are in turn translated into seven structural (VP1-
VP7) and three non-structural (NS1,NS2 and NS3) proteins (Huismans
and van Dijk, 1990). The virus particle is arranged as three concentric
capsid shells surrounding the viral dsRNA (Basak et al., 1997; Grimes et
al., 1995; Huismans and van Dijk, 1990). The outermost layer (the outer
capsid) is composed of two structural proteins, VP2 and VP5, which are
principally involved in virus attachment and penetration of the host cell
during the initiation of infection. These are the most variable of the viral
proteins and the specificity of their interactions with neutralizing
5
antibodies (particularly those of VP2) determines virus serotype (Eaton et
al, 1990; Huismans and van Dijk, 1990; Roy et al., 1990).VP2 is coded
for by genome segment 2, and VP5 is coded for by genome segment 5
(Verwoerd et al., 1970).
The two innermost protein shells that make up the transcriptionally
active virus core, are composed of VP3 and VP7, respectively. These are
more highly conserved proteins, showing serological cross reactions
within the BTV species (Grimes et al., 1995; Mertens, 1999; Mertens et
al., 1987; Verwoerd et al., 1972). VP3 is coded for by genome segment 3
(Huismans and van Dijk, 1990), and VP7 is coded for by genome
segment 7 (Huismans et al., 1987)
The non-structural proteins NS1, NS2 and NS3 are coded for by
genome segments 6, 8 and 10, respectively (Roy, 1992).
1.3. Vector and Transmission
Midges of the genus Culicoides act as biological vectors of
bluetongue virus. Of the approximately 1400 species of Culicoides world-
wide, less than 20 are considered actual or possible vectors (OIE, 1998;
Mellor, 1990).
The most well-studied vector species are C. varipennis and C.
insignis in the United States of America, C. fulvus, C. wadai, C. actoni
6
and C. brevitarsis in Australia, and C. imicola in Africa and the Middle
East (Erasmus, 1990).
The insect vectors of BTV breed in moist conditions in a variety of
habitats, particularly damp, muddy areas and in faecal and plant matter.
They have nocturnal feeding habits, preferring still, warm conditions,
pastures and open pens.
Females take a blood meal prior to egg laying, feed at roughly 4-
day intervals and live for about 2 to 3 weeks. The eggs hatch in 2 to 3
days, and, depending on the temperature, the larval stage lasts 12 to 16
days. Adults emerge 2 to 3 days after pupation and take a blood meal 1
day later and they also mate during this time (Roberts, 1990). The
activities of the midge are influenced by temperature and the optimum
lies between 13ºC and 35ºC (Sellers, 1981).
BTV has evolved a life cycle where alternate cycles of virus
replication in vertebrate and invertebrate hosts are essential for virus
persistence (Roberts, 1990).
The midges may be infected when biting viraemic vertebrates. The
chance of infection depends in part on the genotype of the midge, the
strain of virus, the level of viraemia, and environmental factors. The
incubation period (between feeding on infected blood and the appearance
of virus in the saliva of the midge) is 1-2 weeks (Mellor et al., 2000).
Viraemia must be of the order of 104 infectious units of virus per 1ml of
7
blood or greater for feeding midges to have much chance of infection.
The peak levels of viraemia, in virus infectious units per 1ml of blood,
were reported as 104.4 to 106.3 for cattle, 106.4 to 108 for sheep and 106
for goats. Viraemia peaks in the first two weeks after infection, before the
appearance of serum antibodies. Virus titres then drop rapidly and are
very low if infection persists for a month or more (OIE, 1998). The
duration of viraemia in the infected vertebrates is an important factor in
the transmission of BTV to biting midges (Mac Lachlan, 1994). The
duration of viraemia of most cattle is less than 4 weeks with less than 1%
exceeding 8 weeks (OIE, 1998). The maximum viraemia reported for
sheep is 54 days (Koumbati et al., 1999). Singer et al. (2001) analyzed a
large volume of existing data on the length of bluetongue viraemia of
cattle and concluded that this was equal to or less than 9 weeks in >99%
of adults.
There is no evidence of vertical transmission of the virus in the
invertebrate hosts. Any vertical transmission in vertebrates is considered
to be of no consequence to virus ecology because observations on the
placental transmission of virus in the vertebrate hosts are contradictory
(Roberts, 1990). There is a little evidence of direct or indirect contact
transmission in either host, other than rare instances of seminal
transmission in vertebrates (OIE, 1998). The virus can not be spread by
meat, milk or dairy products (Erasmus, 1990).
8
1.4. Epidemiology and Distribution
Bluetongue is a common, generally sub-clinical infection of
ruminants throughout the tropics and subtropics, within a number of
separate ecosystems. Seasonal incursions of the virus into more temperate
latitudes, sometimes accompanied by disease, may occur under favorable
climatic conditions at certain key locations (Gibbs and Greiner, 1994).
Bluetongue disease is the result of a complex interaction between the
animal, the virus and the environment. It is almost exclusively a disease
of sheep, with European breeds most susceptible. Most breeds of sheep,
especially in regions where the virus is endemic, are resistant to disease
though there is increasing information that native breeds in India and
China can be clinically affected. Outbreaks of disease typically occur
either when susceptible sheep are introduced to endemic areas, or when
infected midges carry the virus from endemic regions to adjacent areas
containing populations of immunologically-naive susceptible sheep
(Erasmus, 1990).
There is evidence that infected midges are carried on the wind for
long distances (Sellers, 1981). It has been postulated that the major
epidemics of bluetongue, in regions where disease occurs only
sporadically can often be traced to windborne carriage of infected
Culicoides from distant areas (Gibbs and Greiner, 1988).
9
Over the past 30 years, evidence of regular virus activity, but not
necessarily disease, has been found in most countries in the tropics and
subtropics with substantial populations of ruminants. The virus may be
found in a geographic band between latitudes 40º N and 35º S. The
presence of BTV within this band, whether year round or seasonal,
depends on the climatic zone type. Genetic studies (topotyping) indicate
that the virus existence in discrete, stable ecosystems, probably the result
of co-evolution of different strains of virus and vectors (OIE, 1998).
Numerous countries in the tropics and subtropics have bluetongue virus
unknowingly circulating subclinically in cattle and other ruminants.
A properly designed serological survey would reveal the presence
of the virus. The virus is endemic in areas of some countries, being more
or less continuously active. Depending on climatic factors affecting the
vector, in most years the virus will seasonally extend to adjacent areas
(Gibbs and Greiner, 1988).
Many strains of bluetongue virus appear incapable of causing
significant disease following natural or experimental infection of sheep
known to be susceptible to disease. Experimental reproduction of disease
can be inconsistent, except with the most virulent strains of virus. This
could be because exposure of sunlight can have a marked influence on the
severity of disease (Erasmus, 1990).
10
1.5. Bluetongue disease in Sudan
In Sudan, bluetongue disease was first reported in 1953 when
samples from the Blue Nile Province were confirmed by the Veterinary
Research Laboratory at Onderstepoort, South Africa, to contain
bluetongue virus (Anon, 1953).
Infection of sheep with BTV at Khartoum University Farm,
Shambat, was suspected on the basis of clinical symptoms but was not
confirmed by virus isolation (Pillai, 1961).
Bluetongue virus group specific antibodies have been detected
throughout the Sudan in the sera of many species of ruminants, which
suggested a wide distribution of the virus. Although the high levels of
bluetongue antibodies and antigens that have been found among cattle,
sheep, goats and camels in Sudan, no virus isolation was made in earlier
years (Eisa et al., 1979, 1983; Abu Elzein, 1983, 1985a, 1986; Abu
Elzein et al., 1987; Herniman et al., 1980).
An outbreak, from which BTV was isolated, involved indigenous
sheep in Western Sudan. The stress which these animals have been
exposed to while being driven over long distances enhanced the severity
of the disease (Eisa et al., 1980).
An outbreak of bluetongue disease in 3-6 months old indigenous
lambs was reported in Khartoum Province in 1982. The BTV was isolated
11
and the disease was experimentally induced (Abu Elzein and Tag Eldin,
1985).
Bluetongue virus was suggested as the cause of death of a Frezian
cross-herd calf at the Khartoum University Farm at Shambat, but the
virus was not isolated (Mohamed et al., 1980).
BTV serotype 5 was isolated from Culicoides species in Sudan
(Mellor et al., 1984).
1.6. Economic Importance
Bluetongue can be a costly infection for several reasons. The
clinical disease in sheep can be severe, resulting in wool break, weight
loss and death. In some countries where disease is endemic such as
Sudan, South Africa and USA, vaccination is a recurring cost. However,
the greater cost of bluetongue is to infected countries which export live
animals, germplasm and some animal products such as fetal calf serum.
Here the presence of bluetongue virus, even if wholly sub clinical, causes
loss of trade due to restriction on the source of animals, and the cost of
health testing. It has been estimated that in the late 1970s, the ban on US
cattle semen exports resulted in an annual loss of $24 million (Gibbs and
Greiner, 1988).
Bluetongue is included in the OIE list A diseases, largely because
of dramatic outbreaks of disease in Cyprus in 1943 and Portugal and
12
Spain in 1956. The Cyprus outbreak was due to a particularly virulent
strain of the virus causing between 60-70% losses in some flocks
(Gambles, 1949). Within the first 4 months, 46,000 sheep had died in
Portugal and 133,000 in Spain (Roberts, 1990). This listing of bluetongue
in the most serious list of animal diseases exacerbates the trade sensitivity
and associated costs to countries with the infection (Gibbs and Greiner,
1994).
1.7. Pathology of bluetongue disease
1.7.1. Pathogenesis of BTV
After introduction by the bite of an infected midge, bluetongue
virus first replicates in the local lymph node and subsequently induces a
primary viraemia which seeds other lymph nodes, spleen, lung and
vascular endothelium (Gibbs and Greiner, 1988). Circulating virus
associates with blood cells, mostly with erythrocytes and platelets, though
virus associated with mononuclear cells is critical for dissemination of
virus throughout the animal. Later in viraemia, the virus is exclusively
associated with erythrocytes (Mac Lachlan, 1994). Virus particles appear
to be sequestered in invaginations of the erythrocyte membrane, allowing
prolonged viraemia in the presence of neutralizing antibodies (OIE,
1998).
13
All of the pathology of bluetongue can be assigned to vascular
endothelial damage resulting in changes to capillary permeability and
fragility, with subsequent disseminated intravascular coagulation and
necrosis of tissues supplied by damaged capillaries. These changes result
in oedema, congestion, hemorrhage, inflammation and necrosis (Erasmus,
1990).
1.7.2. Immune response to bluetongue virus
The mechanism of immunity to BTV infection, and whether this
immunity is mediated by the humoral or the cellular components of the
immune system, is not fully understood.
Evidence of the role of cell-mediated immunity (CMI) in BTV
infection has been demonstrated in sheep (Stott et al., 1979). Sheep which
had been vaccinated with an inactivated BTV vaccine, were refractory to
challenge with homologous virus in the absence of neutralizing
antibodies. Bluetongue virus specific cytotoxic T-Lymphocytes (CTLs)
have been induced in sheep (Jeggo et al., 1985) and their important role
in clearing BTV in sheep was indicated (Ghalib et al., 1985). CTLs also
have been induced in mice infected with live BTV (Jeggo and Wardley,
1982).
The immune response to BTV infection in cattle is complex when
compared to that of sheep. Whereas the appearance of neutralizing
14
antibodies in the serum of infected sheep coincides with a subsequent
decline in circulating virus, this does not seem to be the case in cattle
(Luedke et al., 1977). Even in the presence of high titre of neutralizing
antibodies, the viraemia in cattle persists for months. The failure of cattle
to display clinical disease in the face of prolonged viraemia suggests an
impaired immune response (Osburn, 1985). Immunological tolerance and
viral persistence have been reported in congenitally infected calves
(Luedke, et al., 1977). Congenital infection has also been demonstrated in
bovine fetuses infected between 85 and 125 days of age. The virus was
not recovered from these calves at birth, but virus specific antibodies
were detected in precolostral serum samples (Mac Lachlan et al., 1985).
Clinical bluetongue disease in cattle is mediated by IgE antibodies, and
the role of CMI in bluetongue immunology in cattle is not fully
understood (Jochim, 1985).
1.7.3. Clinical signs
Fever is usual but not invariable. Other common clinical signs
include oedema (of lips, nose, face, submandibulum, eyelids and
sometimes ears), congestion (of mouth, nose, nasal cavity, conjunctiva,
skin and coronary bands), lameness and depression. The oedema of lips
and nose can give the sheep a ‘monkey-face’ appearance. There is
frequently a serous nasal discharge, later becoming mucopurulent. The
15
congestion of the nose and nasal cavity produces a ‘sore muzzle’ effect,
the term used to describe the disease seen in sheep in the USA before its
bluetongue virus etiology was realized. The mouth is sore and the sheep
may champ to produce a frothy oral discharge. Sheep are not strictly
anorexic, but eat less because of oral soreness and will hold food in their
mouths to soften it before chewing. Affected sheep occasionally have
swollen, congested, cyanotic tongues. Lameness, due to coronary band
congestion, may occur early in the disease, and lameness, as a result of
skeletal muscle damage, may occur later.
If fever occurs, sheep are first pyretic 4-10 days after infection. The
other clinical signs, soon followed with acute deaths, occurring during the
second week following infection. Many of theses deaths are the result of
pulmonary oedema and/or cardiac insufficiency. Further sheep may die
from chronic disease 3 to 5 weeks after infection with bacterial
complications, especially Pasteurellosis. The production loss due to
bluetongue may be the result of deaths, unthriftiness during prolonged
convalescence and possibly reproductive wastage (OIE, 1998).
Although the frequency of infection of cattle with BTV is generally
higher than in sheep, disease in cattle is rare. Clinical infection is actually
a hypersensitivity reaction including fever, stiffness or lameness and
increased salivation. The skin of the muzzle is often inflamed, and may
crack and peel. The lips and tongue may be swollen, with ulcers on the
16
oral mucosa. Similarly, the skin of the neck, flanks, perineum and teats
may be affected (Erasmus, 1990).
Hydranencephaly and congenital deformities may develop in
bovine and sheep fetuses of bluetongue virus-infected dams. The severity
of lesions is depending on the stage of gestation. Fetuses seem to be most
susceptible during the period of active brain development (Erasmus,
1990).
It is clear that cell culture-adapted virus more readily crosses the
placenta than unadapted virus, suggesting that the occasionally instances
of natural virus-induced teratogenesis may be due to strains of virus
derived from live virus vaccines (Mac Lachlan, 1994).
Bluetongue in dogs associated with use of a contaminated vaccine
was reported by Akita et al. (1994). Only pregnant bitches were affected.
1.7.4. Post-mortem findings
In animals dying acutely, the oral mucosa is hyperemic and
petechiae or ecchymoses may be present. Excoriations may be in areas
subject to mechanical abrasion; the edges of lips, dental pad, tongue and
cheeks opposite to molar teeth. There may be hyperemia in the fore-
stomach. The lungs may be hyperemic with severe alveolar and
interstitial oedema, froth in the bronchi, and excess fluids in the thoracic
cavity. The pericardial sac may have petechiae and excess fluids.
17
A variable sized hemorrhage in the tunica media near the base of
pulmonary artery is almost pathognomonic. Sub-epicardial and sub-
endothelial hemorrhages, particularly those involving the left ventricle
are common. Generalized damage to the cardiovascular system is
evidenced by widespread hyperemia, oedema and hemorrhage (Erasmus,
1990).
Animals that die later than 14 days after infection often show
dramatic degeneration and necrosis of the skeletal musculature. Muscles
lose pigmentation and the inter-muscular fasciae are infiltrated with clear
gelatinous fluids (Erasmus, 1990).
Microscopic examination of mucosal lesions shows mononuclear
cells infiltration, degeneration and necrosis of epithelial cells in which
large acidophilic intra-cytoplasmic masses accumulate. Affected muscles
have oedema, hemorrhage, hyaline degeneration and necrosis. Infiltration
by neutrophils, macrophages and lymphocytes is present in acute cases
(Verwoerd and Erasmus, 1994).
1.8. Diagnosis of BTV
1.8.1. Virus Isolation
A number of methods for the isolation of bluetongue virus have
been developed over the past fifty years in an attempt to increase the
efficiency with which virus in field materials can be amplified to
18
facilitate identification. Favored methods include replication in
embryonated chicken eggs (ECE), sheep, and a wide variety of cultured
cells (Clavigo et al., 2000; Gard et al., 1988; Gard et al., 1992; Gould et
al., 1989; Wechsler and McHolland, 1988).
1.8.1.1. Embryonated Chicken Eggs (ECE)
Mason, Coles and Alexander (1940) first reported the growth of BTV
in chicken embryos following inoculation into the yolk sac of ECE
(Mason et al., 1940). Over a quarter of a century later, Goldsmit and
Barzilai (Goldsmit and Barzilai, 1968) and Foster and Luedke (Foster et
al., 1972) showed that intravenous inoculation of ECE was 100-1000
times more sensitive than yolk sac administration. Since then, intravenous
inoculation of 10-13-day-old ECE has been widely used as the method of
choice in the isolation of BTV from clinical samples (Clavigo et al.,
2000). The preferred tissues for isolation include washed, unseparated
blood cells, spleen, lung and lymph nodes (Pearson et al., 1992).
Preparation of washed blood cells for inoculation into ECE is
straightforward, whereas tissues must be homogenized by grinding with
sand in a mortar and pestle or in tissue grinder. The number of ECE
inoculated per sample varies but is usually 10, the incubation temperature
33-34ºC and the inoculum dose 0.01ml. Although dead embryos are
usually the source of virus for identification, embryo deaths are neither an
19
indication of BTV replication nor are surviving embryos indicative of
virus absence (Eaton and White, 2004).
1.8.1.2. Sheep inoculation
Sheep have been variously described to be as efficient as ECE
(Foster et al., 1972; Goldsmit et al., 1975), less efficient than ECE
(Breckon et al., 1980) and more efficient than ECE (Luedke, 1969;
Parsonson et al., 1981). The latter suggested that the larger sample
volume that can be administered to sheep might account for the enhanced
efficiency of isolation compared with ECE. However, sheep inoculation
is often an impracticable option because of the requirement to maintain
the sheep for at least 30 days after inoculation to permit development of
the antibody response that provide evidence of virus infection (Eaton and
White, 2004).
1.8.1.3. Cultured cells
The first successful attempt to grow BTV in cultured cells was in
1956. BTV adapted for growth in eggs by serial passage in ECE was
shown to replicate in primary lamb kidneys (Haig et al., 1956). The first
successful isolation in tissue culture of wild-type non-egg adapted virus
from the blood of infected sheep was in 1959 (Fernandes, 1959). Shortly
20
thereafter, direct isolation of BTV in cultured cells was confirmed
(Livingston and Moore, 1962; Pini et al., 1966).
Among the large number of mammalian cell lines that have been
evaluated for their sensitivity to BTV, baby hamster kidney (BHK),
African green monkey (Vero) and calf pulmonary artery endothelium
(CPAE) are most frequently used (Pearson et al., 1992).
1.8.2. Virus Identification
Identification of BTV is an essential part of the laboratory
confirmation of BTV infection. This may be achieved in three different
ways:
a. Identification of antibodies by serological assay
b. Identification of the virus antigens by virological assay
c. Identification of the specific nucleic acids of BTV by reverse
transcriptase-polymerase chain reaction (RT-PCR) and sequence
analysis (Zientara et al., 2004).
1.8.2.1. Serological techniques
The outer capsid, structural viral proteins VP2 and VP5 of BTV are
the serotype determinants and are responsible for generation of serotype-
specific neutralizing antibodies (Roy et al., 1990).
21
Testing sera for the presence of BTV antibodies may be required for
serotype identification of field strains, for monitoring vaccination
campaigns, for serological surveillance, and to facilitate safe international
trade in live animals, animal products and germplasm (Hamblin, 2004).
Two prescribed tests were outlined by the OIE Manual (OIE, 2000)
for international trade, namely, the agar gel immunodiffusion (AGID)
(Pearson et al., 1979) and competitive enzyme-linked immunosorbent
assay (c-ELISA) (Jeggo et al., 1992).
1.8.2.1.1. Agar gel immunodiffusion (AGID)
The AGID test (Pearson et al., 1979) is well documented as a
serogroup-specific test for the detection of BTV antibodies. Although the
AGID test may still be used in some laboratories, the lack of sensitivity
(Gustafson et al., 1992; Pearson, et al., 1992) and documented cross-
reactions that can occur with other orbivirus serogroups (Pearson, et al.,
1992) makes the continued use of this assay questionable when more
rapid, sensitive and specific tests are readily available.
1.8.2.1.2. Competitive enzyme-linked immunosorbent assay
(c-ELISA)
The ELISA has been used for approximately 40 years (Voller et
al., 1979) and has provided a valuable means of studying numerous
22
antigens and their antibodies. ELISA is a serogroup-specific test,
identifying primarily the highly conserved BTV VP7 of all known
serotypes. Using the c-ELISA as a spot test will only provide a qualitative
measurement of positivity (Hamblin, 2004).
Competitive ELISA (cELISA) is probably the most widely used
and validated method (Jeggo et al., 1992).
1.8.2.2. Virological techniques
Several virus/ antibody-based methodologies for the identification
of BTV have been described and they fall into two categories, being
either serogroup-specific such as ELISA, or serotype-specific such as
virus neutralization test (Hamblin, 2004).
1.8.2.2.1. Virus neutralization test
It is a serotype-specific test which can be used to identify all
antigenically distinct serotypes of BTV. The sensitivity of this assay is
dependent on the titer of virus in the test sample (Hamblin, 2004).
1.8.2.3. Reverse transcriptase-polymerase chain reaction (RT-PCR)
The PCR is a method for in vitro amplification of DNA. It is a
series of multiple rounds of primer extension reactions in which
complementary strands of a defined region of a DNA molecule are
23
simultaneously synthesized by a thermo stable DNA polymerase
(Zientara, et al., 2004).
This primer-directed amplification of viral nucleic acid has
revolutionized BT diagnosis. Results to date indicate that PCR technique
may be used, not only to detect the presence of viral nucleic acid, but also
to ‘serogroup’ orbiviruses and provide information on the serotype and
possible geographic source of BTV isolates within a few days of receipt
of a clinical sample, such as infected sheep blood.
Oligonucleotide primers used to date have been derived from
RNA7 (VP7 gene), RNA6 (NS1 gene), RNA3 (VP3 gene) and RNA2
(VP2 gene).
The PCR assay involves three separate procedures. In the first,
BTV RNA is extracted. The second procedure is the denaturation of viral
ds-RNA and reverse transcription (RT) to generate DNA, which is
amplified by PCR. The final step of the process is the analysis of the PCR
product by electrophoresis (OIE, 2000; Dadhich, 2004).
1.9. Prevention and control
Bluetongue is a disease of sheep, but cattle are the principal
vertebrate reservoirs of the virus. Once established, it is impossible to
actively eradicate bluetongue virus. The virus will circulate, generally sub
clinically, in cattle and other ruminants, and in midges. In countries
24
marginally suitable for virus persistence, the virus may be maintained for
several years before dying out (Roberts, 1990).
In seasonally infected areas, the onset of cold weather will reduce
midge populations to ineffective levels and cause the virus to retreat to
regions of year-round activity.
The bluetongue virus cycle could be interrupted by the
immunization of vertebrate hosts, especially cattle, removal of vectors, or
prevention of vector attack. Understandably, the immunization of animals
that will not suffer from the disease is not acceptable to farmers. The
control of midges by the application of insecticides and larvicides to
insect resting and breeding sites, or systemically to cattle, has not been
fully investigated but is likely to have local success only. Protecting
sheep from exposure to midges is a more practical approach and can be
achieved by moving sheep from insect resting and breeding sites, stabling
animals overnight, or the use of insect repellents. Mixing cattle with
sheep will draw vectors with a host preference for cattle from sheep, but
may raise the virus infection level of the midge population. Prophylactic
immunization of sheep is the most practical and effective control
measure, especially when the threat is from an epidemic due to a single
serotype. Multiple serotypes of virus are usual in endemic situations
(Hawkes, 1996), requiring multivalent vaccines because bluetongue
vaccines are serotype specific.
25
The first method of immunization against bluetongue developed
around 1900 in South Africa with inoculation of immune serum and
infective blood. The attenuation of a strain of virus was achieved after
limited serial passages in sheep. This was referred to as Theiler's vaccine,
and over a period of 40 years more than 50 million doses were used. This
vaccine was inadequate because of the plurality of virus strains occurring
in nature (Howell and Verwoerd, 1971). The use of embryonated chicken
egg-attenuated BTV in the production of polyvalent vaccine for sheep
was found to be effective and the virus did not regain its virulence
(Alexander and Haig, 1951; McKercher et al., 1957).
However, multivalent vaccines have attendant problems resulting
from interference between virus strains, differences in immunogenicity
and growth rates between various strains, as well as differences in the
response of individual animals to the components of such vaccines
(Verwoerd and Erasmus, 1994).
Additionally, there is growing concern by some scientists about the
use of live attenuated bluetongue vaccines. Murray and Eaton (1996)
summarized these concerns into four areas. These areas are: the known
teratogenicity of attenuated virus for the developing fetus; the propensity
for vaccine virus to be excreted in the semen of bulls and rams; the
possibility that vaccine virus will infect vectors and establish in the
environment; and the generation of recombinant progeny virus with novel
26
genetic and biological properties after the reassortment of genes from
wild and vaccine virus in the vaccinated animal or the vector.
Alternatives to live attenuated vaccines are described by Murray
and Eaton (1996). Vaccines based on inactivated whole virus,
recombinant virus-like particles or recombinant core-like particles all
show promise, but require more research. If a commercial product of any
of these achieved, it will likely cost considerably more than a live
attenuated vaccine.
Live attenuated bluetongue vaccines have wide use in South
Africa, and more limited use in USA and a few other countries. The
vaccines are compromises between attenuation and immunogenicity and
may have residual pathogenicity for some vaccinated sheep. The
application of the vaccines has to be well managed. Colostral immunity in
young sheep can interfere with the development of active immunity to the
vaccine, and breeding ewes and rams should be vaccinated before mating.
27
CHAPTER TWO
Materials and Methods
2.1. Bluetongue virus
The North American Bluetongue virus prototypes serotypes 2, 10, 11
and 13 were obtained from Arthropod-Borne Animal Disease Research
Laboratory, Laramie, WY. The Sudanese isolates of BTV serotypes were
recovered from Khartoum University farm at Shambat (Mohammed and
Mellor; 1990).
2.2. Virus propagation in tissue culture
Vero cells were cultured in 25 ml tissue culture flasks containing
minimal essential medium (MEM). Fetal Bovine Serum (FBS) was used
at a concentration of 10% for growth and maintenance of cells, and they
were incubated at 37 ºC for 2-3 days. All viruses were propagated on
confluent monolayers of Vero cells. The infectious materials were
harvested When 80 % cytopathic effect (CPE) was observed (Usually 3-5
days after infection). The virus-infected cell culture was then kept at 4ºC
till used for the dsRNA extraction.
28
2.3. Viral nucleic acid extraction from infected cell monolayers
Viral nucleic acid was extracted by the QIAamp viral RNA
extraction kit following the mini spin protocol (QIAGEN GmbH, Hilden,
Germany). The carrier RNA was dissolved in 1ml AVL buffer and
transferred to the AVL bottle and mixed thoroughly. 560 microlitre (µl)
from this preparation were dispensed in each 1.5ml micro-centrifuge tube
and kept at 4ºC. All samples and reagents were equilibrated to room
temperature (about 25ºC). 140 µl from the infected cell culture were
added to the Buffer AVL/Carrier RNA in the micro-centrifuge tube, and
mixed by pulse-vortexing for 15 seconds. The mixture incubated at room
temperature (25ºC) for 10 minutes. The tube was briefly centrifuged to
remove drops from inside of the lid. 560 µl of ethanol (100%) were
added and mixed by pulse vortexing for 15 seconds followed by brief
centrifugation. 630 µl from the mixture were applied to the QIAamp spin
column (in a 2ml collection tube) and centrifuged at 8000 rpm for 1
minute. The column was placed into a clean 2ml collection tube, the
remaining 630 µl of the mixture were applied to it and the previous
centrifugation step was repeated. 500 µl of buffer AW1 were added to
the column, after placing it into a clean 2ml collection tube, and
centrifuged at 8000 rpm for 1 minute. 500 µl of wash buffer AW2 were
added to the column, after placing it into a clean 2ml collection tube, and
centrifuged at 14000 rpm for 3 minutes. The spin column was placed into
29
1.5ml micro-centrifuge tube and 60 µl of Buffer AVE were added. After
it had been left at room temperature for 1 minute, the micro-centrifuge
tube was centrifuged at 8000 rpm for 1 minute. Finally the micro-
centrifuge tubes containing RNA extracts were labeled and kept at -20ºC
till used in RT-PCR. The dsRNA concentration was determined by
spectrophotometer at 260 nm wave length .
2.4. Primers selection
Primers (P1 and P2) were selected from the published sequence of
genome segment 6, which codes for non structural protein 1 (NS1) of
BTV-17 (Hwang et al., 1993). P1 included bases 648-667 of the positive
sense strand of genome segment 6: 5´-GCC CTT ACA CTG GAT ACA
GA-3´. P2 was designed from the complementary strand of the above
sequence between bases 1242-1261: 5´-CCT CGC TCC AGT GTA ACA
AT-3´. PCR amplification using P1 and P2 would be expected to produce
a 614 base pair (bp) PCR product.
Primers were synthesized on a DNA synthesizer (Milligen
iosearch/Millipore, Burlington, MA) and purified using Oligo-Pak oligo-
nucleotide purification columns (Glen Research, Sterling, VA) as per
manufacturer’s instructions.
30
2.5. Reverse transcriptase polymerase chain reaction (RT-PCR)
For each PCR amplification, 1.0 µl methyl mercuric hydroxide of
80 mM concentration was used to denature a mixture of 5 µl of viral
RNA and 2 µl of primers (P1 and P2). The primers were used at a
concentration of 20µg/µl. The mixture was then incubated at 25ºC for 10
minutes. The mixture was neutralized by 10 µl of neutralizing solution
containing 1 µl of β-mercapto ethanol, 8 µl of dNTPs (2 µl of each dATP,
dTTP, dGTP and dCTP) and 1 µl of enzyme RNAse inhibitor.
A reverse transcription step was performed to synthesize a
complementary DNA (cDNA) from RNA templates using a reverse
transcriptase mixture composed of 2.7 µl of 10X PCR buffer, 5 µl MgCl2
of 1.5 mM concentration and 1.1 µl of the enzyme reverse transcriptase.
8.8 µl of the reverse transcriptase mixture was added to each PCR tube.
The PCR tubes were placed in the thermal cycler at 40ºC for 30 minutes.
For amplification, a mixture of 7.3 µl of 10X PCR buffer, 8 µl of
MgCl2 and 1 µl of Taq DNA polymerase at a concentration of 5 units/µl
was added to each PCR tube. The total volume of the PCR mixture was
brought to 100 µl using double distilled water.
The RT-PCR was performed in (TECHNE, TC-412, USA) thermal
cycler (figure 2.1) following a program of a 2 minute incubation at 95ºC,
followed by 40 cycles of (95ºC for 1 minute as denaturing temperature;
56 ºC for 30 seconds as annealing temperature and 72 º C for 45 seconds
31
for extension of the predicted amplified PCR product). A final
incubation at 60ºC for 5 minutes was performed to complete the
extension of uncompleted fragments of the PCR products.
2.6. Agarose gel electrophoresis
Following RT-PCR assay, the amplified products of cDNA
transcribed from viral RNA molecules by RT-PCR were analyzed in
agarose gel (SeaKem, agarose FMC Bioproducts, Rockland, ME). 1%
agarose gel was prepared by suspending 1 gram of agarose powder in
100ml of 1X Tris-boric acid-EDTA (TBE) buffer. The suspension was
placed in the microwave for 2 minutes until the agarose was completely
melted. Then 35ml of melted agarose was cooled and poured in the gel
tray loaded with a comb.
The agarose gel was submerged in the buffer basin of the
electrophoresis apparatus (figure 2.2) filled with 1X TBE buffer
containing 10 µl/500 ml ethidium bromide.
To prepare 10X TBE buffer, 108 gram of Tris- (hydromethyl)-
aminomethane, 55 gram of Boric acid and 7.4 gram of EDTA were mixed
and brought to 1 litre using double distilled water. The buffer was kept at
room temperature and used at 1x concentration for gel preparation and
electrophoresis.
32
Per each electrophoresis run, 5 µl of 100 bp ladder molecular
weight marker (MW marker) stained with an indicator dye was placed in
the first lane of the gel. 12 µl of each RT-PCR product were loaded in the
gel after being stained with 3 µl of an indicator dye (Bromophenol blue).
Constant electric current of 90 mV then switched on for about 40
minutes to allow the migration of the amplified PCR products. The
standard molecular weight marker (1KB) was incorporated in each
reaction for determination of the size of the amplified product by
comparing this size with the separate distinguishable bands of the MW
marker which were seen when the gel was stained with ethidium bromide.
The gel was then visualized under UV light using gel
documentation apparatus (figure 2.3).
33
Figure 2.1: Thermal cycler (TECHNE, TC-412, USA)
34
Figure 2.2: Electrophoresis apparatus
35
Figure 2.3: Gel documentation apparatus
36
CHAPTER THREE
Results
The BTV RT-PCR assay was a simple procedure that efficiently
detected all BTV serotypes used in this study.
The described RT-PCR assay, with primers derived from genome
segment 6 of BTV serotype 17, afforded sensitive and specific detection
of all BTV serotypes used in this study. The specific 614 bp PCR product
was visualized on ethidium bromide-stained agarose gel from 1ng RNA
of North American BTV serotypes 2, 10, 11, and 13, and from 1ng RNA
of Sudanese BTV serotype 2 (Figure 3.1).
37
Figure 3.1: Detection of BTV serotypes in infected cell cultures.
Visualization of the 614 bp PCR product on ethidium bromide-stained
agarose gel from 1ng RNA of North American BTV serotypes 2, 10, 11,
and 13; and Sudanese serotype 2. Lane MW: molecular weight marker;
Lane 2: North American BTV-2; Lane 2: Sudanese BTV-2; Lane 3:
North American BTV-10; Lane 4: North American BTV-11; Lane 5:
North American BTV-13. Lane 6: Non-infected Vero cell (negative
control).
MW 1 2 3 4 5 6
500 bp
38
Specificity of BTV RT-PCR
The specificity studies indicated that the specific 614 bp PCR
product was not detected from 1ng of RNA from epizootic hemorrhagic
disease virus (EHDV) serotype 1 and 1ng of RNA from Palyam virus
(Figure 3.2).
Figure 3.2: Specificity of RT-PCR for BTV.
614 bp Amplification product was not detected from 1ng RNA of EHDV-
1 and Palyam virus. Lane MW: molecular weight marker; Lane 1:
Sudanese BTV-2 (positive control); Lane 2: EHDV-1; Lane 3: Palyam
virus.
1000 bp
500 bp
MW 1 2 3
39
Sensitivity of BTV RT-PCR
The sensitivity studies indicated that the specific 614 bp PCR
product was obtained from the amounts of 1ng, 500 pg, 250 pg and 125
pg RNA from Sudanese BTV serotype 2 (Figure 3.3).
Figure 3.3: Sensitivity of RT-PCR for BTV.
The specific 614 bp PCR product was obtained from the amounts of 1.0
ng, 500 pg, 250 pg and 125 pg RNA from Sudanese BTV serotype 2.
Lane MW: molecular weight marker; Lanes 1-4: (Sudanese BTV-2) 1.0
ng, 500 pg, 250 pg and 125 pg, respectively.
MW 1 2 3 4
1000 bp
500 bp
40
CHAPTER FOUR
Discussion
Bluetongue virus is a serious veterinary problem (Shope et al.,
1960), and the economic importance of the disease is mainly attributed to
clinical disease in sheep (Holf and Trainer, 1974; Pini, 1976; Jessup,
1985). There is restriction on the international trade of livestock and
animals products unless the animals are certified BTV-free by
conventional virus isolation and serology (Osburn et al., 1994). Thus the
disease is of interest to dairy producers and wildlife managers.
Rapid detection of BTV is important in disease outbreaks as well
as for determining disease-free status of exporting animals. Many
diagnostic tests have been developed for detection of BTV including
antibody, antigen and nucleic acid detection techniques (Mecham and
Wilson, 1994). Antibody detection indicates that an animal has been
previously exposed to the virus but not necessarily an indicator of
viraemia. An indirect enzyme-linked immunosorbent assay (ELISA) and
a competitive (C) ELISA, using a group-specific monoclonal antibody
against bluetongue virus (BTV), are described for the detection of
antibodies to BTV in cattle and sheep sera (Afshar et al., 1987).
41
Antigen and nucleic acid detection assays are more indicative of
viraemia, but can detect residual non-infectious molecules from a recent
infection. None of the available antigen or nucleic acid detection assays
have been validated for all the 25 serotypes of BTV. Although antigen
detection assays are very sensitive, inexpensive and reliable, they take
long time to perform. In addition, reagents are more difficult to develop
than nucleic acid detection tests. Therefore, the development of a nucleic
acid amplification-based assay for all serotypes of BTV was necessary
(Aradaib et al., 1998; Dangler et al., 1990; Katz et al., 1993; Shad et al.,
1997; Wilson and Chase, 1993).
Conventional virus isolation and serology are time consuming and
laborious (Pearson et al., 1992). The traditional approaches that rely on
virus isolation followed by virus identification may require at least three
to four weeks to generate information on BTV serogroup and serotypes.
Also, conventional virus isolation may not provide data on the possible
geographic origin of the isolates (Zientara et al., 2004). So, the
development of molecular diagnostic techniques for detection of BTV
would be advantageous in a variety of circumstances including clinical
and sub clinical disease investigation, vaccination programs and
epidemiological studies (Pearson et al., 1992; Aradaib et al., 1994, 1995).
RT-PCR based detection assays have been described for detection
of bluetongue virus infection in susceptible ruminants (McColl and
42
Gould, 1991; Wade-Evans et al., 1991; Wilson and Chase, 1993). BTV
RT-PCR can provide rapid, sensitive and specific viral identification for
BTV infections (Zientara et al., 2004). The primary gene target for group-
specific amplification was genome segment 6, which codes for non-
structural protein 1 (NS1) as it is highly conserved among cognate genes
of BTV serogroup (Jensen and Wilson, 1995).
The BTV RT-PCR assay was a simple procedure that efficiently
detected all BTV isolates used in this study. The described RT-PCR assay
specifically detected BTV RNA in infected vero cell cultures. Selection
of the primers was based on the observation that NS1 gene of BTV is the
most conserved among cognate genes of BTV serogroup (Aradaib et al.,
1998).
The specific 614 bp PCR products, visualized on ethidium
bromide-stained agarose gel, were obtained from all BTV RNA samples
tested (1.0 ng each). The specificity studies indicated that the specific 614
bp PCR product was not amplified from 1.0 ng of RNA from epizootic
hemorrhagic disease virus (EHDV)-1 and RNA from Palyam virus under
the same stringency condition described in this study. This confirmed that
BTV NS1 genome is highly conserved among cognate genes of BTV
serogroup.
A nested PCR assay using primers derived from the non structural
protein 1 (NS1) gene of North American BTV serotype 11 was
43
developed to detect the United States BTV serotypes 2, 10, 11, 13 and 17
and the Sudanese BTV serotypes 1, 2, 4 and 16 and BTV serotype 4 from
South Africa and BTV serotype 2 from Senegal. The primary specific 790
bp PCR products and the nested 520 bp amplification products were not
detected from closely related Orbiviruses including, EHDV serotypes 1,
2, 4; Sudanese isolate of Palyam virus and total nucleic acid extracts from
uninfected Vero cells (Aradaib et al., 2005).
A duplex, one-step RT-PCR assay was developed and evaluated to
detect genome segment 7 from any of the BTV serotypes. Assay
sensitivity was evaluated using tissue culture derived virus, infected
vector insects and clinical samples (blood and other tissues). No cross-
reactions were detected with members of closely related Orbivirus
species (African horse sickness virus, Epizootic hemorrhagic disease
virus and Palyam virus) (Anthony et al., 2007).
Two new Real Time qPCRs were developed and validated to detect
and amplify BTV segments 1 and 5 from all of the BTV serotypes. These
two methods are complementary and could be used in parallel to confirm
the diagnosis of a possible incursion of BTV (Toussaint et al., 2006).
A new rapid single step RT-PCR with infra red (IR)-dye- labeled
primers was reported as a sensitive and specific assay for detecting BTV
RNA in Culicoides biting midges. All serotypes of BTV and none of the
44
eight serotypes of the closely related EHDV were detected (Kato and
Mayer, 2006).
The sensitivity studies indicated that the specific 614 bp PCR
product was obtained from the amounts of 1 ng, 500 pg, 250 pg and 125
pg RNA from Sudanese BTV serotype 2. Thus, in the present study, the
described BTV RT-PCR protocol indicated that the PCR assay was
capable of detecting the amount of 125 pg of BTV genomic dsRNA.
However, the interpretation of positive BTV PCR results must be
analyzed carefully, particularly in BTV-free areas before officially
reporting BTV cases. In the absence of epidemiological data, virus
isolation is strongly recommended to confirm molecular diagnosis
(Zientara et al., 2004).
The BTV RT-PCR assay provides supportive diagnostic technique
to the lengthy cumbersome conventional virus isolation procedures.
While the nested PCR assay required 7 hours for submission of the final
results (Aradaib et al., 2005), the BTV RT-PCR assay described in this
study, required 4 hours to obtain the final results.
The rapidity, sensitivity and specificity of the RT-PCR assay
would greatly facilitate detection of BTV infection among susceptible
ruminants.
45
In conclusion, the described BTV RT-PCR assay, using primers
derived from genome segment 6 of BTV-17, provides a simple, rapid,
specific and sensitive diagnostic method for detection of BTV.
In addition, the PCR assay could be used for detection of the virus
in areas of endemicity or incursion of the virus in BTV- free zones.
46
REFERENCES
Abu Elzein, M. E. M. (1983). Precipitating antibodies against bluetongue
and foot and mouth disease viruses in cattle between the two Nile
in Khartoum Province, Sudan. Rev. Sci. Tech. Off. Int. Epiz. 2 (4),
1059-1066.
Abu Elzein, M. E. M. (1985). Bluetongue in the Sudan. Rev. Sci. Tech.
Off. Int. Epiz. 4 (4), 795-801.
Abu Elzein. E. M. E. (1985a). Bluetongue in camels: A serological
survey of the one-humped camel (Camelus dromedaries). Rev.
Elev. Med. Vet. Pays. Trop. 38 (4), 438-442.
Abu Elzein, M. E. M. (1986). Recovery of bluetongue virus serogroup
from sera collected for a serological survey from apparently
healthy cattlefrom the Sudan. J. of Hyg. Camb. 96, 529-533.
Abu Elzein, E. M. E. and Tag Eldin, M. H. (1985). The first outbreak of
sheep bluetongue in Khartoum Province, Sudan. Rev. Sci. Tech.
Off. Int. Epiz. 4 (3), 509-515.
47
Abu Elzein. E. M. E., Fayza, A. O., Fagieri, I. (1987). Natural exposure
of exotic cattle to bluetongue virus in the Sudan as reflected by
sero conversion. Bull. Anim. Hlth. Prod. Afri. 35, 4.
Afshar. A., Thomas, F. C., Wright, P. F., Shapiro, J. L., Shettigara, P. T.,
Anderson, J. (1987). Comparison of competitive and indirect
enzyme-linked immunosorbent assays for detection of bluetongue
virus antibodies in serum and whole blood. J. clin. Microbiol. 25
(9), 1705-1710.
Alexander, R. A., Haig, D. A. (1951). The use of egg attenuated
bluetongue virus in the production of a polyvalent vaccine for
sheep. A propagation of the virus in sheep. Onderstepoort J. Vet.
Res. 25, 3-15.
Anon, (1953). Annual report, Department of Animal Production, Sudan
Government.
Anthony, S., Johnes, H., Darpel, K. E., Elliott, H., Maan, S., Samuel, A.,
Mellor, P.S., Mertens, P. P. (2007). A duplex RT-PCR assay for
detection of genome segment 7 (VP7 gene) from 24 BTV
serotypes. J Virol. Methods.
48
Aradaib, I. E., Akita, G. Y., Osburn, B. I. (1994). Detection of epizootic
hemorrhagic disease virus serotypes 1 and 2 in cell culture and
clinical samples using polymerase chain reaction. J. Vet. Diagn.
Invest. 6, 143-147.
Aradaib, I. E., Akita, G. Y., Pearson, J. E., Osburn, B. I. (1995).
Comparison of polymerase chain reaction and virus isolation for
detection epizootic hemorrhagic disease virus in clinical samples
from clinically infected deer. J. Vet. Diagn. Invest. 7, 196-200.
Aradaib, I. E., Schore, C. E., Cullor, J. S., Osburn, B. I. (1998). A nested
PCR for detection of North American isolates of bluetongue virus
based on NS1 genome sequence analysis of BTV17. Vet.
Microbiol. 59, 99-108.
Aradaib, I. E., Wilson, W.C., Schore, C. E., Mohammed, M. E., Yilma, T.
D., Cullor, J. S., Osburn, B. I. (1998). PCR detection of North
American and Central African isolates of epizootic hemorrhagic
disease virus (EHDV) based on genome segment 10 of EHDV
serotype 1. J. Clin. Microbiol. 36, 2604-2608.
Aradaib, I. E., Mohamed, M. E., Abdalla, T. M., Sarr, J., Abdalla, M. A.,
Yousof, M. A., Hassan, Y. A., Karrar, A. R. (2005). Serogrouping
49
of United States and some African serotypes of bluetongue virus
using RT-PCR. Vet. Microbiol. 111 (3-4), 145-150.
Basak, A. K., Grimes, J., Gouet, P., Roy, P., Stuart, D. (1997). Structures
of orbivirus VP7: implications for the role of this protein in the
viral life-cycle. Structure 5, 871-883.
Bekker, J. G., De Kook, G., Quinlan, J. B. (1934). The occurrence and
identification of bluetongue in cattle, the so-called psudo foot and
mouth disease in South Africa. Ondersteport. J. Vet. Sci. Anim. Ind.
2, 393-507.
Borden, E. C., Shope, R. E., Murphy, F. A. (1971). Physicochemical and
morphological relationships of some arthropod-borne viruses to
bluetongue virus-a new taxonomic group. Physicochemical and
serological studies. J. Gen. Virol. 3, 261-271.
Breckon, R. D., Luedke, A. J., Walton, T.E. (1980). Bluetongue virus in
bovine semen: viral isolation. Am. J. Vet. Res. 41, 439-442.
Callis, J. (1985). Bluetongue in the United States. In Bluetongue and
related orbiviruses (T. L. Barber, M. M. Jochim and B. I. Osburn,
eds). Proc. First international symposium, Monterey, California,
50
16-20 January 1984. A. R. Liss, New York, Progr. Clin. Biol. Res.
178, 37-42.
Clavijo, A., Heckert, R. A., Dulac, G. C., Afshar, A. (2000). Isolation and
identification of bluetongue virus. J. Virol. Methods 87, 13-23.
Dadhich, H. (2004). Bluetongue: an overview of recent trends in
diagnostics. Vet. Ital. 40(4), 564-566.
Dangler, C. A., de Mattos, C. A., de Mattos, C.C., Osburn, B.I. (1990).
Identifying bluetongue virus ribonucleic acid sequences by the
polymerase chain reaction. J. Virol. Methods 28, 281-292.
Davies, F. G., Mungai, J. N., Pini, A. (1992). A new bluetongue virus
serotype isolated in Kenya. Vet. Microbiol. 31, 25-32.
Eaton, B. T., Hyatt, A. D., Brookes, S. M. (1990). Replication of
bluetongue viruses. In Current Topics in Microbiology and
Immunology: bluetongue viruses. (P. Roy and B. M. Gorman, eds).
Springer- Verlag, New York, 89-118.
Eaton, B. T., White, J. R. (2004). Developing new orbivirus diagnostic
platforms. Vet. Ital. 40(4), 525-530.
51
Eisa, M., Karrar, A. E., Abdel Rahim, A. H. (1979). Incidence of
bluetongue virus precipitating antibodies in sera of some domestic
animals in the Sudan. J. Hyg. Camb. 83, 539-545.
Eisa, M., Osman, O. M., Karrar, A. E., Abdel Rahim, A. H. (1980). An
outbreak of bluetongue in sheep in the Sudan. Vet. Rec. 106 (23),
481-482.
Eisa, M., Mc Grane, J. J., Taylor, W.P., Ballouh, A. (1983). A survey of
precipitating antibodies of bluetongue virus in domestic animals of
the Sudan. Bulletin of Animal Health and Production in Africa. 31,
95-99.
Erasmus, B. J. (1990). Bluetongue virus. Virus infections of ruminants
227-237.
Fenner, F., Pereira, H. G., Porterfield, J. S. (1974). Family and generic
names for virus approved by the International Committee on
Taxonomy of Viruses. Intervirology 3, 193-194.
Fernandes, M. V. (1959). Isolation and propagation of bluetongue virus in
tissue culture. Am. J. Vet. Res. 20, 398-408.
52
Foster, N. M., Luedke, A. J., Metcalf, H. E. (1972). Bluetongue in sheep
and cattle: efficacy of embryonating chicken eggs in viral isolation.
Am. J. Vet. Res. 33, 77-81.
Gambles, R. M. (1949). Bluetongue of sheep in Cyprus. Journal of
Comparative Pathology 59, 176-190.
Gard, G. P., Eaton, B. T., Gould, A. R. (1992). Virus isolation technology
for Australian orbiviruses. In Bluetongue, African horse sickness
and related orbiviruses (T. E. Walton and B. I. Osburn, eds). Proc.
Second International Symposium, Paris, 17-21 June 1991.CRC
Press, Boca Raton, 694-700.
Gard, G. P., Weir, R. P., Walsh, S. J. (1988). Arboviruses recovered from
sentinel cattle using several virus isolation methods. Vet.
Microbiol. 18, 119-125.
Ghalib, H. W., Shore, O. E., Osburn, B. I. (1985). Immune responses of
sheep to bluetongue virus: in vitro included lymphocyte
blastogenesis. Vet. Immuno. Immunopath. 10, 177-188.
Gibbs, E. P. J., Greiner, E. C. (1988). Bluetongue and epizootic
hemorrhagic disease. The arboviruses: epidemiology and ecology.
Volume II, 39-70.
53
Gibbs, E. P. J., Greiner, E. C. (1994). The epidemiology of bluetongue.
Comparative Immunology, Microbiology and Infectious Diseases
17 (3/4), 207-220.
Goldsmit, L., Barzilai, E. (1968). An improved method for the isolation
and identification of bluetongue virus by intravenous inoculation of
embryonating chicken eggs. J. Comp. Pathol. 78, 477-487.
Goldsmit, L., Barzilai, E., Tadmor, A. (1975). The comparative
sensitivity of sheep and chicken embryos to bluetongue virus and
observations in experimentally infected sheep. Aust. Vet. J. 51,
190-196.
Gould, A. R., Hyatt, A. D., Eaton, B. T., White, J. R., Hooper, P. T.,
Blacksell, S. D., Le Blanc Smith, P. M. (1989). Current techniques
in rapid bluetongue virus diagnosis. Aust. Vet. J. 66, 450-454.
Gould, A. R., McColl, K. A., Prichchord, L. I. (1992). Phylogenetic
relationship between bluetongue virus and other orbiviruses. In:
(Walton, T.E., Osburb, B.I. eds.), Bluetongue African Horse
Sickness and Related Orbiviruses. CRC Press, Boca Raton, FL, pp.
452-460.
Grimes, J., Basak, A. K., Roy, P., Stuart, D. (1995). The crystal structure
of bluetongue virus VP7. Nature 373, 167-170.
54
Gustafson, G. A., Pearson, J. E., Moser, K. M. (1992). A comparison of
the bluetongue competitive ELISA to other serologic tests. In
Bluetongue, African horse sickness and related orbiviruses (T. E.
Walton and B. I. Osburn, eds). Proc. Second International
Symposium, Paris, 17-21 June 1991.CRC Press, Boca Raton, 570-
574.
Haig, D. A., Mc Kercher, D. G., Alexander, R. A. (1956). The
cytopathogenic action of bluetongue virus on tissue cultures and its
application to the detection of antibodies in the serum of sheep.
Onderstepoort J. Vet. Res. 27, 171-177.
Hamblin, C. (2004). Bluetongue virus antigen and antibody detection,
and the application of laboratory diagnostic techniques. Vet. Ital.
40(4), 538-545.
Hawkes, R. A. (1996). The global distribution of bluetongue. Bluetongue
disease in Southeast Asia and the Pacific: Proceedings of the
Southeast Asia and Pacific Regional Bluetongue Symposium,
Greenlake Hotel, Kunming, P. R. China, 22-24 August 1995., 6-14.
Herniman, K. A., J., Gumm, I. D., Owen, L., Taylor, W. P., Sellers, R. F.
(1980). Distribution of bluetongue virus and antibodies in some
countries of the Eastern Hemispher. Bull. Off. Epiz. 92, 581-586.
55
Holf, G. L., Trainer, D. O. (1974). Observation on bluetongue and
epizootic hemorrhagic disease viruses in white-tailed deer: (1)
distribution of virus in blood (2) cross-challenge. J. Wildl. Dis. 10,
25-31.
Howell, P. G., Verwoerd, D. W. (1971). Bluetongue virus. In Virology
monographs, Vol. 9 (S. Gard, C. Hallaver and K. F. Meyer, eds).
Springer Verlag, New York, 35-74.
Huismans, H., Dijk, A.A.van., Els, H. J. (1987). Uncoating of parental
bluetongue virus to core and subcore particles in infected L cells.
Virology 157, 180-188.
Huismans, H. and Dijk, A.A.van. (1990). Bluetongue virus structural
components. Current Topics in Microbiology and Immunology
162, 21-41.
Huismans, H. and Dijk, A.A.van. (1990). Bluetongue virus structural
components. In Current Topics in Microbiology and Immunology:
bluetongue viruses. (P. Roy and B. M. Gorman, eds). Springer-
Verlag, New York, 21-41.
Hwang, G. Y., Chiou, J. F., Yang, Y. Y., Li Li, J. K. (1993). High
sequence conservation among the United States bluetongue viruses
56
cognate M2 genes which encode the nonstructural NS1 tubule
protein. Virology 192, 321-327.
Jeggo, M. H., Wardley, R. C. (1982). The induction of murine cytotoxic
T. Lymphocytes by bluetongue virus. Arch. Virol. 71, 197-206.
Jeggo, M. H., Wardley, R. C., Brownlie, J. (1985). Importance of ovine T
cells in protection against BTV infection. In bluetongue and related
orbiviruses. pp. 477-487. Edited by T. L. Barber and M. M.
Jochim. New York, Alan R. Liss.
Jeggo, M., Wright, P., Anderson, J., Eaton, B., Afshar, A., Pearson, J.,
Kirkland, P., Ozawa, Y. (1992). Review of the IAEA meeting in
Vienna on Standardization of the competitive ELISA test and
reagents for the diagnosis of bluetongue. In Bluetongue, African
horse sickness and related orbiviruses (T. E. Walton and B. I.
Osburn, eds). Proc. Second International Symposium, Paris, 17-21
June 1991.CRC Press, Boca Raton, 547-560.
Jensen, M. J., Wilson, W. C. (1995). A model for the membrane topology
of the NS3 protein as predicted from the sequence of segment 10 of
epizootic hemorrhagic disease virus serotype 1. Arch. Virol. 140,
799-805.
57
Jessup, D. A. (1985). Epidemiology of two orbiviruses in California’s
native wild ruminants: preliminary report. In: (Barber, T., Jochim,
M., eds). Bluetongue and Related Orbiviruses, Progress in Clinical
and Biological Research, Alan R. Liss, New York, 178, 53-65.
Jochim, M. M. (1985). Bluetongue: A review of the immune response of
sheep and cattle to bluetongue virus. In bluetongue and related
orbiviruses. pp. 455-460. Edited by T. L. Barber and M. M.
Jochim. New York, Alan R. Liss.
Kato, C. Y., Mayer, R. T. (2006). An improved, high-throughput method
for detection of bluetongue virus RNA in Culicoides midges
utilizing infra red-dye- labeled primers for Reverse Transcriptase
PCR. J Virol. Methods.
Katz, J. B., Alstad, A.D., Gustafson, G.A., Moser, K.M. (1993). Sensitive
identification of bluetongue virus serogroup by a colorimetric dual
oligonucleotide sorbent assay of amplified viral nucleic acid. J.
Clin. Microbiol. 31, 3028-3030.
Koumbati, M., Mangana, O., Nomikou, K., Mellor, P. S., Papadopoulos,
O. (1999). Duration of bluetongue viraemia and serological
responses in experimentally infected European breeds of sheep and
goats. Veterinary Microbiology 64 (4), 277-285.
58
Livingston, C. W., Moore, M. S. (1962). Cytochemical changes of
bluetongue virus in tissue cultures. Am. J. Vet. Res. 23, 701-710.
Luedke, A. J. (1969). Bluetongue in sheep: viral assay and viraemia. Am.
J. Vet. Res. 30, 499-509.
Leudke, A. J., Jochim, M. M., Jones, R.H. (1977). Bluetongue in cattle:
repeated exposure of two immunologically tolerant calves to
bluetongue virus by vector bites. Am. J. Vet. Res. 38, 1701-1704.
Leudke, A. J., Jochim, M. M., Jones, R.H. (1977). Bluetongue in cattle:
effects of Culicoides variipennis transmitted bluetongue virus on
pregnant heifers and their calves. Am. J. Vet. Res. 38, 1687-1695.
Mac Lachlan, N. J., Osburn, B. I., Stott, J. L., Ghalib, H. W. (1985).
Orbivirus infection of the bovine fetus. In Bluetongue and related
orbiviruses. pp. 79-84. Edited by T. L. Barber and M. M. Jochim.
New York, Alan R. Liss.
Mac Lachlan, N. J. (1994). The pathogenesis and immunology of
bluetongue virus infection of ruminants. Comparative Immunology,
Microbiology and Infectious Diseases 17 (3/4), 197-206.
59
Mason, J. H., Coles, J. D. W. A., Alexander, R. A. (1940). Cultivation of
bluetongue virus in fertile eggs produced in vitamin-deficient diet.
Nature 145, 1022.
McColl, K. A., Gould, R. A. (1991). Detection and characterization of
bluetongue virus using the polymerase chain reaction. Virus Res.
21, 19-34.
Mc Kercher, D. G., Mcgowan, B., Cabasso, V. J., Roberts, G. I. (1957).
Studies on bluetongue 111. The development of a modified live
virus vaccine employing American strains of the bluetongue virus.
Am. J. Vet. Res. 18, 310-316.
Mecham, J. O., Wilson, W.C. (1994). Strategies for improved bluetongue
diagnostics. Proc. US Anim. Hlth Assoc. 98, 43-50.
Mellor, P. S., Boorman, J. P. T., Wilkinson, P. J., Martinez-Gomez, F.
(1983). Potential vectors of bluetongue and African horse sickness
viruses in Spain. Vet. Rec. 112, 229-230.
Mellor, P. S., Osborne, R., Jenning, D. M. (1984). Isolation of bluetongue
and related viruses from Culicoides spp. in the Sudan. J. Hyg.
Camb. 93, 621-628.
60
Mellor, P. S. (1990). The replication of bluetongue virus in Culicoides
vectors. Bluetongue viruses. Current Topics in Microbiology and
Immunology 162, 143-161.
Mellor, P. S., Boorman, J., Baylis, M. (2000). Culicoides biting midges:
their role as arbovirus vectors. Annual Review of Entomology 45,
307-340.
Mellor, P. S., Wittmann, E. J. (2002). Bluetongue virus in the
Mediterranean Basin 1998-2001. Vet. J. 164, 20-37.
Mertens, P. P. C., Burroughs, J. N., Anderson, J. (1987). Purification and
properties of virus particles, infectious sub-viral particles and cores
of bluetongue virus serotypes 1 and 4. Virology 157, 375-386.
Mertens, P. P. C. (1999). Orbivirus and Coltivirus. In Encyclopedia of
virology, 2nd Ed. (A. Granoff and R. G. Webster, eds). Academic
Press, London, 1043-1074.
Mertens, P. P. C., Diprose, J., Maan, S., Singh, K. P., Attoui, H., Samuel,
A. R. (2004). Bluetongue virus replication, molecular and
structural biology. Vet. Ital. 40(4), 426-437.
61
Mohamed, K. A., Eisa, M., Nayil, A. A., Ballouh, A., Abd Elaziz, M.
(1980). A fatal bluetongue infection in a calf: A case report. Sudan
J. Vet. Res. 2, 69-73.
Mohammed, M.E.H. and Taylor, W. P. (1987). Infection with bluetongue
and related orbiviruses in the Sudan detected by the study of
sentinel calf herds. Epidemiol. Infect. 99, 533-545.
Mohammed, M.E.H., Mellor, P. S. (1990). Further studies on bluetongue-
related orbiviruses in the Sudan. Epidemiol. Infect. 105, 619-632.
Murray, P. K., Eaton, B. T. (1996). Vaccines for bluetongue. Australian
Veterinary Journal 73 (6), 207-210.
Office International des Epizooties. (1998). Supporting document for the
OIE International Animal Health Code chapter 2. 1. 9 on
bluetongue. OIE AD hoc working group on bluetongue, September
1998.
Office International des Epizooties. (OIE) (2000). Bluetongue. In Manual
of standards for diagnostic tests and vaccines. OIE, Paris, 153-167.
Osburn, B. I. (1985). Role of the immune system in bluetongue host-viral
interactins. In Bluetongue and related orbiviruses. pp. 417-422.
62
Edited by T. L. Barber and M. M. Jochim. New York, Alan R.
Liss.
Osburn, B. I., Aradaib, I. E., Schore, C. E. (1994). Comparison of
bluetongue and epizootic hemorrhagic disease complex. Proc. of
the 13th International Symposium of the World Association of
Veterinary Microbiologists, Immunologists and Infectious Disease
Specialists, Peruguia and Montova, Italy, pp. 219-224.
Parsonson, I. M., Della-Porta, A. J., McPhee, D. A., Cybinski, D. H.,
Squire, K. R., Standfast, H. A., Uren, M. f. (1981). Isolation of
bluetongue virus serotype 20 from the semen of an experimentally
infected bull. Aust. Vet. J. 57, 252-253.
Pearson, J. E., Jochim, M. M. (1979). Protocol for the immunodiffusion
test for bluetongue. Proc. Am. Assoc. Vet. Lab. Diagn. 22, 463-471.
Pearson, J. E., Gustafson, G. A., Shafer, A. L., Alstad, A.D. (1992).
Diagnosis of bluetongue and epizootic hemorrhagic disease. In
Bluetongue, African horse sickness and related orbiviruses (T. E.
Walton and B. I. Osburn, eds). Proc. Second International
Symposium, Paris, 17-21 June 1991.CRC Press, Boca Raton, 533-
546.
63
Pillai, C. (1961). Suspected cases of bluetongue in sheep. Sudan J. Vet.
Sci. Anim. Husb. 8, 47-50.
Pini, A., Coackley, W., Ohder, H. (1966). The adverse effect of some calf
sera on the isolation and propagation of bluetongue virus in tissue
culture. Arch. Gesammte. Virusforsch. 18, 88-95.
Pini, A. (1976). Study on the pathogenesis of bluetongue: replication of
the virus in the organs of infected sheep. Onderstepoort J. Vet. Res.
43, 159-164.
Roberts, D. H. (1990). Bluetongue: a review. State Veterinary Journal 44
(124), 66-80.
Roy, P., Marshall, J. J. A., French, T. J. (1990). Structure of the
bluetongue virus genome and its encoded proteins. In Current
Topics in Microbiology and Immunology: bluetongue viruses. (P.
Roy and B. M. Gorman, eds). Springer- Verlag, New York, 43-87.
Roy, P., Marshall, J. J. A., French, T. J. (1990). Structure of the
bluetongue virus genome and its encoded proteins. In Bluetongue
viruses (P. Roy and B. M. Gorman, eds). Curr. Top. Microbiol.
Immunol. 162, 43-87.
64
Roy, P. (1992). Bluetongue virus proteins. J. Gen. Virol. 73, 3051-3064.
Sellers, R. F. (1981). Bluetongue and related diseases. In: (Gibbs, E. P. J.,
eds). Virus Diseases of Food Animals. London, UK: Academic
Press, 567-584.
Shad, G., Wilson, w. C., Mecham, J. O., Evermann, J.F. (1997).
Bluetongue virus detection: a safer reverse-transcriptase
polymerase chain reaction for prediction of viremia in sheep. J.
Vet. Diag. Invest. 9, 118-124.
Shope, R. E., MacNamara, K. G., Mangold, R. (1960). A virus-induced
epizootic hemorrhagic disease of the Virginia white-tailed deer
(Odocoileus virginianus). J. Exp. Med. 111, 155-170.
Singer, R. S., Mac Lachlan, N. J., Carpenter, T. E. (2001). Maximal
predicted duration of viraemia in bluetongue virus-infected cattle.
Journal of Veterinary Diagnostic Investigation 13, 43-49.
Stott, J. L., Osburn, B. I., Barber, T. L. (1979). The current status of
research on an experimental bluetongue virus vaccine. Proc. 83rd
Ann. Meet. US Anim. Hlth. Assoc. 28-31 Oct., 1-2 Nov., 1979, pp.
55-62.
65
Tabachnick, W. J., Robertson, M. A., Murphy, k. E. (1996). Culicoides
variipennis and bluetongue disease. Ann. NY Acad. Sci. 791, 219-
226.
Tomori, O., Baba, S., Adu, F., Adeniji, J. (1991). An overview and
perspective on orbivirus disease prevalence and occurrence of
vectors in Africa. In Bluetongue, African horse sickness and
related orbiviruses (T. E. Walton and B. I. Osburn, eds). Proc.
Second International Symposium, Paris, 17-21 June 1991.CRC
Press, Boca Raton, 23-33.
Toussaint, J. F., sailleau, C., Breard, E., Zientara, S., De Clercq, K.
(2006). Bluetongue virus detection by two real-time RT-qPCRs
targeting two different genomic segments. J Virol. Methods.
Verwoerd, D. W. and Erasmus, B. J. (1994). Bluetongue. In: (Coetzer, J.
A. W., Thomson, G. R., Tustin, R. C., eds). Infectious Diseases of
Livestock with special reference to Southern Africa. Cape Town,
South Africa: Oxford University press, 443-459.
Verwoerd, D. W., Els, H. J., De Villiers, E., Huismans, H. (1972).
Structure of the bluetongue virus capsid. J. Virol. 10, 783-794.
66
Verwoerd, D. W., Louw, H., Oellermann, R. A. (1970). Characterization
of bluetongue virus ribonucleic acid. J. Virol. 5, 1-7.
Voller, A., Bidwell, D. E., Bartlett, A. (1979). The enzyme linked
immunosorbent assay (ELISA). In A guide with abstracts of
microplate applications. Dynatech Europe, Guernsey, 128 pp.
Wade-Evans, A. M., Mertens, P. P. C., Bostock, J. C. (1991).
Development of polymerase chain reaction for the identification of
bluetongue virus in tissue samples. J. Virol. Methods 30, 15-24.
Wechsler, S. J. and McHolland, L. E. (1988). Susceptibilities of 14 cell
lines to bluetongue virus infection. J. Clin. Microbiol. 26, 2324-
2327.
Wilson, W. C., Chase, C. C. L. (1993). Nested and multiplex polymerase
chain reaction for the identification of bluetongue virus infection in
the biting midge Culicoides variipennis. J. Virol. Methods 45, 39-
47.
Zientara, S., Breard, E., Sailleau, C. (2004). Bluetongue diagnosis by
reverse transcriptase-polymerase chain reaction. Vet. Ital. 40(4),
531-537.