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

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Abu Elzein, M. E. M. (1985). Bluetongue in the Sudan. Rev. Sci. Tech.

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Abu Elzein. E. M. E. (1985a). Bluetongue in camels: A serological

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