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PREVALENCE AND ASSOCIATED RISK FACTORS OF
BLUETONGUE VIRUS IN PUNJAB AND BALOCHISTAN
TAYYEBAH SOHAIL
2009-VA-209
A THESIS SUBMITTED IN THE PARTIAL FULFILLMENT OF THE
REQUIREMENTS FOR THE DEGREE
OF
DOCTOR OF PHILOSOPHY
IN
MICROBIOLOGY
UNIVERSITY OF VETERINARY AND ANIMAL SCIENCES,
LAHORE
2019
To,
The Controller of Examinations,
University of Veterinary and Animal Sciences,
Lahore.
We, the Supervisory Committee, certify that the contents and form of the thesis, submitted by Miss
Tayyebah Sohail, Registration No. 2009-VA-209 have been found satisfactory and recommended that it
to be processed for the evaluation by the External Examiner(s) for award of the Degree.
Supervisor
Dr. Muhammad Zubair shabbir
Member
Prof. Dr. Masood Rabbani
Member
Muhammad Yasir Zahoor
i
DEDICATION
I would like to dedicate my thesis to my beloved parents for being the lantern of hope throughout
my life, my teachers and friends whose support gave me strength and confidence to complete my
studies.
ii
ACKNOWLEDGEMENTS
In the name of ALLAH, the Most Gracious and the Most Merciful. All praises to ALLAH for giving me
strength, opportunity and his blessing to complete my thesis. I am also thankful to the holy prophet
MUHAMMAD (PBUH), as a symbol of guidance and hope for me in life.
Foremost, I would like to express sincere gratitude to my supervisor Dr. Muhammad Zubair
Shabbir, Associate Professor, Department of Microbiology, University of Veterinary and Animal
Sciences, Lahore for his continuous support, patience, motivation, enthusiasm, and immense knowledge
throughout my study and research. His invaluable help with constructive comments and suggestions have
contributed to the success of this research.
Besides my supervisor, I would like to pay my special thanks to my supervisory committee:
Prof. Dr. Masood Rabbani, Pro Vice Chanceller/Dean Faculty of Veterinary Sciences, UVAS Lahore
and Dr. Muhammad Yasir Zahoor, Assistant Professor, Institute of Biochemistry and Biotechnology,
UVAS Lahore for their encouragement, insightful comments, and sincere advice.
I am thankful to the Chairman of Department of Microbiology, Prof. Dr. Tahir Yaqub.
Finally, my deepest gratitude to my beloved friends and class fellows for all their moral support,
encouragement and sincere advice during my research work.
Tayyebah Sohail
iii
CONTENTS
DEDICATION (i)
ACKNOWLEDGEMENTS (ii)
LIST OF TABLES (iv)
LIST OF FIGURES (v)
LIST OF ABBREVIATIONS (vi)
LIST OF ANNEXURES (vii)
ABSTRACT (viii)
SR. NO. CHAPTERS PAGE NO.
1 INTRODUCTION 1
2 REVIEW OF LITERATURE 3
3 MATERIALS AND METHODS 10
4 RESULTS 15
5 DISCUSSION 31
6 SUMMARY 35
7 LITERATURE CITED 36
iv
LIST OF TABLES
TABLE.NO. TITLE PAGE.NO.
2.1 Epidemiological pattern and distribution of different serotypes of
bluetongue virus worldwide
8
4.1. Herd-wise seroprevalence and serotype distribution of BTV among
different districts of Punjab.
16
4.2. Animal species-wise distribution and seroprevalence of sheep, goat, cattle and buffalo in Punjab province.
17
4.3 Univariate analysis of anti-VP7 antibodies in small ruminants and their
potential association to categorical variables.
19
4.4 Univariate analysis of anti-VP7 antibodies in large ruminants and their
potential association to categorical variables.
21
4.5 Multivariable mixed effects logistic regression model (p < 0.05) for BTV
serological status amongst ruminants originating from traditional
farms/villages in Punjab province (2013-15).
23
4.6 Summary of the number of sheep and goats sampled, the prevalence of anti-
VP7 antibodies and the BTV serotypes present in the Kech, Mastung, Killa
Saifullah and Quetta districts of Balochistan province.
26
4.7 Univariate analysis of anti-VP7 antibodies in goats and their potential
association to categorical variables.
27
4.8 Univariate analysis of anti-VP7 antibodies in sheep and their potential
association to categorical variables.
28
4.9 Multivariate logistic regression model determining the relationship between
categorical predictors and seropositivity in goats.
28
4.10 Multivariate logistic regression model determining the relationship between
categorical predictors and seropositivity in sheep.
29
v
LIST OF FIGURES
FIGURE NO. TITLE PAGE NO.
3.1 The cELISA plate showing positive and negative results of samples
along with positive control (PC) and negative control (NC). 12
4.1 Geographical distribution pattern of sites for blood sampling and
identified serotypes across selected districts of Punjab province,
Pakistan
25
4.2 Geographical distribution pattern of sites for blood sampling and
identified serotypes across selected districts of Balochistan province,
Pakistan
30
vi
LIST OF ABBREVIATIONS
BT: Bluetongue
BTV: Bluetongue virus
VP: Viral protein
NS: Non-structural protein
cELISA: Competitive enzyme-linked immune sorbent assay
PCR: Polymerase chain reaction
RT-PCR: Real-time polymerase chain reaction
OIE: Office International des Epizooties
DTRA: Defense threat reduction agency
HEC: Higher Education Commission
vii
LIST OF ANNEXURES
SR. NO. TITLE PAGE NO.
1 Blood sample collection Performa ix
2 Seroprevalence of Bluetongue Virus in small ruminants in Balochistan
province, Pakistan
x
3 Seroprevalence of Bluetongue Virus in small and large ruminants in Punjab
province, Pakistan xi
viii
ABSTRACT
Bluetongue (BT) is a vector-borne disease of immense economic importance for small and large
ruminants. Despite frequent disease reports from neighboring countries, a little is known about current
disease status and prevalent serotypes in Pakistan. We screened a total of 1,312 healthy animals (sheep=
326, goat= 476, cattle= 234, buffalo= 276) from Punjab and 876 from clinically healthy sheep (475) and
goats (401) from Balochistan for the detection of group-specific antibodies and serotype-specific genome
for BT virus through competitive ELISA and real-time PCR, respectively. An overall prevalence of
group-specific VP7 antibodies [28.81% (n= 378/1312, 95% CI=26.4 – 31.4)] was observed in Punjab.
The prevalence was higher in goats [40.75% (n=194/476, 95% CI=36.4 – 45.3)] followed by buffalo
[29.34% (n=81/276, 95% CI=24.3 – 34.9)], sheep [18.40% (n= 60/326, 95% CI=14.5 – 22.9)] and cattle
[17.94% (n= 42/234, 95% CI= 13.56 – 23.4)]. The odds of seropositivity were more in buffalo of Nili
breed (OR= 2.06, 95% CI= 1.19-3.58) as well as those found with a presence of vector (OR= 2.04, 95%
CI= 1.16-3.59). Buffalo and cattle with history of abortion [(OR= 3.95, 95% CI= 1.33-11.69) and (OR=
5.89, 95% CI= 1.80-19.27) respectively] were much likely to be infected with the disease. Serotype 8
was detected in all animal species while, serotypes 4 and 6 were detected in sheep, 2, 6 and 11 in goat,
and 2 and 16 in buffalo. The study concludes a much frequent exposure of different serotypes of
Bluetongue virus (BTV) in small and large ruminants and indicates its expansion to enzootic range
worldwide.In Balochistan, none of the study herds (n = 97) were seronegative for BTV, and at the
individual level, the overall prevalence of BTV seroconversion was 47.26% (n = 414/876, 95%
CI=43.92-50.63%). A higher percentage of goats (50.87%, 95% CI = 45.99-55.73%) were seropositive
for anti-VP7 immunoglobulins (IgG) than sheep (44.21%, 95% CI= 39.81-48.70%). Odds of
seroconversion for goats were associated with breed-type (χ2 = 16.84, p = 0.01), parity (χ2 = 23.66, p =
0.00) and presence of vector (χ2 = 2.63, p = 0.10), whereas for sheep, it was associated with breed-type
(χ2 = 13.80, p = 0.01) and parity (χ2 = 53.40, p = 0.00). The presence of vector was also observed to be a
risk factor in goat. Serotype 8 was the most prevalent (26.82%, 95% CI=14.75-43.21%) followed by an
equal prevalence of serotypes 2 and 9 (7.31%, 95% CI= 1.91-21.01%). To the best of our knowledge,
this is the first study conducted in Balochistan province and the results indicate that there is a necessity to
initiate intervention strategies to control BT disease burden not only in this region of Pakistan but also in
adjacent areas of the neighboring countries, Iran and Afghanistan.
1
CHAPTER 1
INTRODUCTION
Bluetongue (BT) is a vector borne viral disease of small and large ruminants including both
domestic and wild animals. The virus responsible of disease transmission is Bluetongue virus, which
belongs to genus Orbivirus within the family Reoviridae. Bluetongue virus (BTV) is icosahedral, 90 nm
in diameter and is non-enveloped with triple layered protein capsid. It has segmented dsRNA genome
consisting of ten segments each segment coding for different viral proteins. Seven structural (VP1–VP7)
and four non-structural (NS1–NS3 and NS3A) proteins are produced by viral genome. Numerous species
of Culicoides biting midges serve as a vector for transmission of virus from one susceptible host to
another (Legisa et al. 2013). Also some mechanical vectors play role in disease transmission such as
ticks and mosquitoes. These vectors do not help in viral replication in the carrier but just act as a vehicle
to transmit the virus from one host to the other (Bouwknegt et al. 2010; Hoffmann et al. 2012). To-date,
27 BTV serotypes are identified worldwide (Maan et al. 2012; Jenckel et al. 2015). Bluetongue is widely
spread to latitudes 35ºS and 40ºN, extending further to North-Western America, China and Kazakhstan
up to 50º (Lundervold 2011). It is believed that BTV is established worldwide in each continent but
Antarctica (Maclachalan 2011).
Office International des Epizooties (OIE) has declared Bluetongue, ‘notifiable disease’ due to
direct losses by mortality, infertility, rare abortions, reduced milk and weight gain and indirect losses by
international animals trade restriction, cost on countrywide disease surveillance, mass vaccination,
treatment and vector control strategies (Wilson and Mellor 2009; Coetzee et al. 2012; OIE 2014). An
annual loss of US$ 125 million has been estimated in US alone due to this disease (Nicoletti et al. 2014).
Clinical signs include erosions and oedema of oral mucosa, conjunctivitis, salivation, fever, apathy and
tiredness, lameness, and dysphagia, muscle necrosis, coronitis and limb stiffness.
Although BT equally infects all ruminants but the disease is mainly observed in goat, sheep,
cattle, llamas, camels, antelopes and deer. Comparatively severe clinical disease is observed in sheep and
white-tailed deer (Odocoileus virginianus) but often is subclinical in cattle (Tabachnick 1996; Bitew et
al. 2013; Arun et al. 2014). Except ruminants, BT is also reported in carnivores. In a report, it was
observed that BTV was found in dogs that were given BTV contaminated vaccine (Wilbur et al. 1994;
Evermann, 2008).
Pakistan shares borders with India to the east, Iran to the southwest, China to the northeast and
Afghanistan to the west. In China, Afghanistan and Iran, the prevalence of BTV has been investigated by
several researchers using indirect or competitive ELISAs (Panda et al. 2011; Joardar et al. 2013;
Mozaffari et al. 2014; Ma. et al. 2017). A prevalence of group-specific anti-VP7 antibodies in sheep
(68.6%), goats (71.8%) and cattle (48.6%) were reported from Afghanistan (Ali et al. 2014). The
seroconversion rates for anti-VP7 antibodies in sheep (58.82%), goats (66.95%) and cattle (70.0%) were
reported from India (Panda et al. 2011; Joardar et al. 2013). Seroprevalence of BTV in goats (67.7%)
and sheep (46%) was studied in Iran (Mozaffari et al. 2014; Yavari et al. 2018), whilst a prevalence of
20.3% and 13.3% in sheep and yaks was reported from the Tibetan Plateau of China, respectively (Ma et
al. 2017). In Jilin province, China a recent study showed 17% prevalence in Sika deer (Liu et al. 2018).
In this context, one can assess potential spread and persistency of BTV in Pakistan, particularly in
scenarios when cross-border movement of animals and animal products is not unusual, such as that in
areas adjacent to Afghanistan and Iran (Raziq et al. 2010; Zahur et al. 2011). The open-access and free-
movement of the disease vector (Culicoides spp., mosquitoes and ticks) presents another potential source
for transboundary transmission of BTV in the region (Saegerman et al. 2008; Lam et al. 2011; Busch et
al. 2014).
Bluetongue virus infection is considered as endemic in Pakistan (Sarwar 1962; Akhtar et al.
1995; Sreenivasulu et al. 2003). High seropositivity (48.4%) was documented in 34 sheep flocks of
Introduction
2
North West Frontier Province Pakistan (Akhtar et al. 1997). It was also found that seropositivity
was associated with abortion risk and distance travelled by flock during transhumant movements. A
recent study conducted in Khyber Pakhtunkhwa province showed a seroprevalence of 56.60% and
42.86% in sheep and goat respectively (Malik et al. 2018). In the same district, BTV serotypes 2, 6, 8
and 9 were detected (Malik et al. 2019).
However, currently in Pakistan, there is an absolute lack of surveillance, routine diagnosis or
vaccination for BTV. Given the evolutionary dynamics and global relevance of BTV as a trans-boundary
pathogen, data on the current disease status, epidemiology and prevalent serotypes are pre-requisites for
devising appropriate interventions for disease control strategies. To collect such data, we conducted a
surveillance study in areas of the Punjab and Balochistan province of Pakistan that were selected due to
their close proximity to the borders with Afghanistan, Iran and India. The outcomes of this study provide
novel details on the epidemiology of BTV in this previously unexplored geographical area that will
contribute to an enhanced understanding of this pathogen and the development of intervention strategies
to control it.
3
CHAPTER 2
REVIEW OF LITERATURE
Bluetongue is an infection of small and large ruminants caused by Bluetongue virus. Virus is
transmitted by the bite of insect vector among domestic and wild ruminant species. Disease is found in
temperate, tropical and subtropical regions worldwide (Pandrangi 2012). Bluetongue is included as
notifiable disease according to the classification mentioned by OIE due to its economic importance (OIE
2008). Notifiable diseases are basically the diseases that have the potential of fast spread and
communicability. These diseases are usually important regarding socioeconomic impact and public
health concerns (Alexander et al. 1996).
2.1. Etiology: Bluetongue virus belongs to genus Orbivirus and family Reoviridae. It is non-developed virus
with icosahedral symmetry. Virus diameter is 90nm and has triple layered capsid. Genome consists of
double-stranded RNA that is segmented to 10 segments while each segment coding for different protein.
These segments code for seven structural and four non-structural proteins (VP1–VP7 and NS1–NS3,
NS3A respectively). Two proteins VP2 and VP5 form the outermost layer of the virus, (Roy and Noad
2006). Protein responsible for determining the virus serotype is VP2 because it is responsible for binding
to the receptors, haemagglutination and stimulating host-specific immune response. The second major
protein, VP5 is responsible for interaction with host cell endosomal membrane that is responsible for
playing a slight role in provoking antibody response against the virus (Roy 2008). The VP7 protein forms
the middle or inner capsid called core (Nason et al. 2004). This protein is the main determinant of
serotype specificity and the target for most of the serological tests for identification of antibody against
BTV. The other proteins VP1, VP3, VP4 and VP6 form the innermost layer (subcore), that plays role in
genome replication and transcription (Anthony et al. 2007; Schwartz-Cornil et al. 2008). The non-
structural proteins control many other important functions such as virus replication, maturation and
export of new virus progeny from the infected cell (Ratinier et al. 2011).
2.2. Historical Background: Bluetongue disease has been endemic in Africa for more than a hundred years especially in wild
ruminants in sub-Saharan regions (Verwoerd and Erasmus 2004). In South Africa, the first report came
to light in 18th century from the province “Cape” in an imported Marino sheep from Europe. The disease
was first referred to as “fever” or “epizootic catarrh” but after that name was mistakenly changed to
“malarial catarrhal fever” due to its similarity with the disease caused by intracorpuscular parasite
(Hutcheon 1881; Hutcheon 1902). Later the distinct symptoms and cyanotic tongues in infected sheep
lead to changing the name to bluetongue in 1905 (Spreull 1905).
The first epidemic outside the African continent occurred in the European continent amongst
sheep of Cyprus in 1943 with mortality reaching up to 70% in some sheep flocks. It is estimated that
about 2500 sheep died in that outbreak (Gambles 1949; MacLachlan 2004). Serotype 10 of BTV entered
Portugal and South West Spain between 1956-1960 causing deaths in almost 180,000 sheep (Lopez and
Botija 1958; Manso-Ribeiro and Noronha 1958; Gorman 1990).
In 1948 the virus was observed in Texas, USA (Hourrigan and Klingsporn 1975). European
serotypes (1, 2, 4, 5, 8, 9, 11, 16) were also observed from Florida, Mississippi, and Lousiana (Mellor et
al. 2009; Wilson et al. 2009). Reports of similar outbreaks were observed from other continents as in
Israel the disease was observed in 1949 (Komarov and Goldsmit 1951). Afterwards, the disease was also
observed Asia, Middle East and Southern Europe (MacLachlan 2004). In 1980s it was also observed in
South America (Clavijo et al. 2002). Bluetongue virus of serotype 20 in Australia was first isolated in the
Northern region of the country from Culicoides in 1976 (Kirkland 2004). Bluetongue has successively
been detected from all continents as Africa, the Americas, the Indian subcontinent, the Middle East,
northern Australia, China, South-East Asia and Europe. Antarctica is the only continent exempt of BTV
infection (Maclachlan et al. 2009).
Review of Literature
4
Bluetongue in Greece was first reported in an island named Lesbos in 1979. In a period of just
three months sheep flocks suffered from morbidity (10-90%) and mortality (28%) in 17 communes with
29,300 sheep population and 68 flocks each harboring 5000 sheep (Vassalos 1980; Mastrogiannis et al.
1981; Nomikou et al. 2004), while goats and cattle did not show any clinical signs (Dragonas 1981). In
Turkey, BT was first reported in Aydin province in 1977 in west Anatolia (Yonguç et al. 1982). The
serotype 4 caused an outbreak covering western Provinces Antalya, Balikesir, Çanakkale, Manisa,
Kocaeli and Denizli on the Black Sea coast (Erturk et al. 2004). After 1998, bluetongue distribution was
changed dramatically spreading to North-western Europe and Scandinavia (Tollersrud 2009; Mellor et al.
2008). In late 1999 and early 2000 BT was first observed in Tunisia in north-eastern part (Hammami
2004). BTV-2 was reported in July-September 2000, from the eastern Bulgeria near Tunisian border
(Hammami 2004).
2.3. Bluetongue Virus Life Cycle: VP2 protein helps the virus for attachment to the cell surface. Virus internalization in the
endosome occurs with the help of clathrin-dependent endocytosis pathway (Forzan et al. 2007). From the
capsid layer the dissociation of VP2 protein takes place. The process of acidification inside endosome
stimulates the fusion of VP5 protein with the endosomal membrane (Forzan et al. 2004). This process
results in transport of active virus core protein into the cytoplasm after which replication takes place
inside cytoplasm. Transcription of all the ten segments to positive sense ssRNA takes place with help of
VP1 protein (Boyce et al. 2004). Capping of mRNAs is done with the help of transmethylase and
guanylyl transferase and enzymes produced from VP4 gene (Sutton et al. 2007). VP1 then constructs the
negative strand RNA using that template producing dsRNA (Boyce et al. 2004). The outer capsid protein
is constructed with VP2 and VP5 that surrounds the core particles of progeny viruses while entering to
the host cell cytoplasm. Transportation of virus particles occurs with the assistance of microtubules
(Bhattacharya et al. 2007). Finally, the cell membrane destabilizes and virions are released from the
infected cells. This process is mediated by the activity of NS3 protein (Han and Harty 2004).
2.4. Culicoides species:
Culicoides (C.) imicola is considered the major bluetongue disease vector especially in the parts
of Mediterranean areas that encountered the disease before 1998 such as Israel, the Greek islands of
Rhodes, Anatolian Turkey, Morocco, Portugal and southwestern Spain (Mellor and Wittmann 2002). The
vector was assumed to be very important in disease transmission in Mediterranean Basin. Despite the
movement of this vector to the north parts, BTV was also observed to the areas where either C. imicola
was absent or very rare (Sicily, Lazio and Tuscany in Italy) suggesting that some novel vector other than
C. imicola were responsible for the disease transmission (De Liberato et al. 2005; Purse et al. 2005). For
instance, C. obsoletus along with C. pulicaris and C. dewulfi were reported more frequently in non-
Imicola and BTV outbreak reported areas suggesting that may be these vectors were responsible for the
disease outbreaks (De Liberato et al. 2003; Torina et al. 2004; De Liberato et al. 2005). However, these
vectors were found to have lower susceptibility to the virus (Jennings and Mellor 1988). In Europe due to
increased global warming, these vectors gained much importance due to increase growth, increase in
feeding, egg production and oral susceptibility as described by (Mellor 2004).
2.5. Transmission by Biting Midges: Adult female haematophagous midges belonging to genus Culicoides (Diptera: Ceratopogonidae)
is mainly responsible for BTV transmission (Du Toit 1944). These midges feed blood to obtain protein to
produce eggs. Once virus is in the gut of insect vector, it infects the wall of posterior mid gut. After that
inside the body cavity, the progeny virus particles are released. From there, these particles move towards
the salivary glands. Second cycle of replication takes place in salivary glands. Once the virus is turned to
active virus it is freely available with the higher chances to get entry to the blood stream of susceptible
host in the process of blood feeding by the vector (Wilson and Mellor 2008). Depending upon the
Review of Literature
5
temperature and virus serotype the time required from insect gut to salivary gland varies from a few days
to several weeks (Mullens et al. 1995; Wittmann et al. 2002).
There are 1400 known species of midges from which only about 30 species are BTV vectors
(Meiswinkel et al. 2008). The susceptibility of the vector to the infection by normal blood feeding is low
but selective breeding or/and artificial feeding techniques can encourage probability of infection. Even
low virus particle number is sufficient to cause an infection with a probability of almost 100%
(O'Connell 2002).
2.6. Factors contributing towards bluetongue transmission: The vector activity is dependent on the competent midges species, temperature and climatic
conditions and certain other factors.
2.6.1. Overwintering:
The BT outbreaks usually follow a pattern with re-appearing after some months and the term
used for this phenomenon is called overwintering (Yonguç et al. 1982; Taylor and Mellor 1994; Wilson
et al. 2007). During this period the virus either stays hidden in the host or the vector. Exact mechanism is
unknown. The average time of incubation of the virus inside the host cattle and sheep is only a few
weeks (Koumbati et al. 1999; Bonneau et al. 2002; Gubbins et al. 2008). But occasionally in some
animals can last up to 100 days (Sellers and Taylor 1980; MacLachlan et al. 1991). After the viremia
stage is over, the virus can come to the skin for over a month. The virus can cause a persistent infection
in γδ T-cells. Skin fibroblast cells have counter ligands for WC-1 surface specific molecules on γδ T-cells.
The interaction of γδ T-cells and skin fibroblast-cells can cause increased virus production and hence
increased chances of virus transmission at biting sites on animal’s skin (Takamatsu et al. 2003).
Most of the Culicoides species are capable of surviving at a temperature of 10 ˚C and some may
survive for upto 3 months (Lysyk and Danyk 2007) but at this temperature the viral replication is
effectively reduced (Mullens et al. 1995). Likewise, BTV serotype 8 infection reappeared in the spring
2007 when the average incubation period in the vector after the cessation of previous transmission period
(Wilson et al. 2007). Vertical transmission of viruses in vectors was not observed (Mellor 1990; Mellor
2000). However, sometimes the virus transmission can occur via mechanical vectors as ticks and
mosquitoes (Brown et al. 1992; Bouwknegt et al. 2010).
2.6.2. Temperature:
For the purpose of breeding and feeding these vectors necessitate warmness and humidity
between 28°C and 30°C for maximum activity. A cold temperature and dry hot weather can markedly
reduce vector numbers. So, climate is a major risk factor for BTV transmission (Purse et al. 2005). So for
BTV transmission the climate change plays vital role regarding the distribution and availability of
suitable habitat for the vectors to live in. for example, C. obsoletus adults like shady and leafy habitats
i.e., forest. C. imicola mostly inhabit the vegetation more inclined towards daylight (Conte et al. 2007). Animal dung, irrigation, agriculture practice, river floodings, storms and rise of see level are used by C.
pulicaris, C. newsteadi, C. salinarius, C. halophilus and other species like C. circumscriptus, and C.
maritimus as these provide the breeding areas for these vectors. However, the role of these midges in
disease transmission is unknown (Wilson and Mellor 2008).
2.6.3. Moisture availability:
Precipitation is important in terms of its role in determining the area and tenacity of the semi-
aquatic sites used for breeding of insect larvae, moist sites for adults in autumn/summer and protection
from desiccation (Murray 1991; Mellor et al. 2000). For example, C. imicola breeds in wet soil
especially rich in organic materials but on the other hand, the pupae are prone to flooding of these sites
(Braverman et al. 1974; Braverman 1978).
2.7. Transmission by Other Means: Some other arthropods also showed the presence of BTV such as sheep ked (Melophagus ovinus)
Review of Literature
6
and in some tick species (Bouwknegt et al. 2010). However, these arthropods act as mechanical vectors
which play a minor role in disease establishment. BTV transmission can also occur from one animal to
another through placenta (Santman-Berends et al. 2010) and through semen (Kirkland et al. 2004). Live
attenuated vaccines of bluetongue virus or vaccines against other antigens contaminated with bluetongue
virus can transmit virus (Pandrangi 2012). Oral transmission can occur by direct ingestion of the
contaminated placental or foetal material (Menzies et al. 2008). Fetus can be infected from mother
through placenta. In most of the cases it causes abortion, still birth or viable offsprings with defected
nervous system (Osburn 1994; MacLachlan et al. 2000).
2.8. Intra-region transmission: Transmission of BTV from one area into another can occur in four ways, firstly, through the
movement of animal hosts or their products like semen and embryos, secondly, by the movement of
infected vector species. These vectors are carried by various living or inanimate means. The third way is
the active flight of infected vector. Fourth way is through the passive transport of infected vector by the
wind. This mode of transmission is responsible for long distance dissemination of virus (Saegerman et
al. 2008).
2.9. Mechanism of Pathogenesis: Infected midge transmits the bluetongue virus through the bite to the susceptible animal. From
the skin, virus is transported to the local lymph nodes via host dendritic cells (Hemati et al. 2009). Virus
then infects the endothelial linings of blood vessels, mononuclear phagocytic cells and dendritic cells
(Mahrt and Osburn 1986; Barratt-Boyes et al. 1992; Hemati et al. 2009). This results in necrosis, tissue
infarction and vascular thrombosis resulting in hemorrhage (Pini 1976; Mahrt and Osburn 1986;
Verwoerd and Erasmus 2004). Subsequently, the virus spreads to the other parts of the body via blood
circulation inducing primary viremia. From blood, virus is transmitted to secondary organs such as lungs,
spleen and lymph nodes (Sánchez-Cordón et al. 2010). In early viremia, virus associates itself with blood
components. On following stages, the virus particles attach and are sequestered in the invaginations of
erythrocyte membrane (MacLachlan 2004). The tissue damage results in the clinical manifestation of the
disease as malaise, serous or sometimes bloody discharge from nose, severe pulmonary and facial
oedema, ulcers in mouth and lameness (Verwoerd and Erasmus 2004; Maclachlan et al. 2009).
BTV infection causes cell apoptosis and necrosis (Mortola et al. 2004). p38MAP kinase is
activated that increases the vascular permeability. This initiates the production of TNFα, IFN-I, IL-1, IL-
6, IL-8, and cyclooxygenase-2 that cause the increased production of prostacyclin and thromboxane in
blood plasma resulting in excessive inflammatory response. This inflammatory response is responsible
for the development of clinical signs of the disease caused by damage to infected animal cells and tissues
(Drew et al. 2010).
The mortality rate in sheep is ranged between 2-30% (Verwoerd and Erasmus 2004) but
occasionally can be up to 70% (Gambles 1949). The recovery of the infected animals can be prolonged
and during that period animals can suffer reduced fertility, milk production and wool quality (Verwoerd
and Erasmus 2004; Maclachlan et al. 2009). These cause the direct economic loss due to mortality and
morbidity in susceptible and infected animals. Furthermore, the restrictions on animal trades
internationally and the germ plasm cells from the BT endemic countries cause significant indirect
economic losses (MacLachlan and Osburn 2006). The cost for surveillance, mass vaccination aming
susceptible animals, vector control strategies and treatment of infected animals also contributes to
economic loss (Wilson and Mellor 2008).
2.10. Host Range and Clinical Signs: Bluetongue virus infects ruminant species, including goat, sheep, cattle, deer and camelids. The
virus usually remains subclinical and asymptomatic in cattle and goats. Sheep and deer are more
susceptible to the disease with the development of clinical disease (Elbers et al. 2008). Clinical signs
developed in infected animals include fever, lameness, cattaral inflammation of lips and hooves,
Review of Literature
7
conjunctivitis, salivation and hyperaemia (Darpel et al. 2007). In severe cases, it can also cause abortion
in infected pregnant animals. Even after the disease recovery of the animal, it may exhibit different long-
lasting secondary effects. These include reduced milk production, less weight gain, wool breakage and
sometimes sterility in animal (Wilson and Mellor 2009).
The virus can cause an acute infection in an immunologically naïve population where BTV is not
usually prevalent. For instance, in Europe BTV infection of serotype 8 caused the clinical signs of
disease in both cattle and goat despite of the fact that the disease is usually sub-clinical in cattle. The
clinical signs observed in cattle included conjunctivitis, discharge from eyes, necrotic lesions and ulcers
and odema of lips and tongue and ocular discharge, oral mucosal congestion, ulcer formation and
necrosis on the lips and tongue (Dal Pozzo et al. 2009), while the signs observed in goats included facial
odema, odema of lips, discharge from nose and erythema of skin and udder (Dercksen et al. 2007).
Cattle usually plays the role of a virus amplifying host in BTV endemic areas (Barratt-Boyes et al.
1992).
2.11. Impact of Bluetongue on economy:
OIE classified BT as a ‘notifiable disease’ in 1960s. This decision lead to many restrictions on
animal trade and the germ plasm from BT endemic countries which had severe economic loss in BT
infected countries and overall international livestock industry which was more than the damage due to
the disease itself (Gibbs and Greiner 1994). Despite of these restrictions BT continues to emerge in BT
non-infected areas like observed in northern Europe (Scandinavia) and in south-eastern regions of US
and Australia (Purse et al. 2005; Maclachlan and Guthrie 2010).
2.12. Global Epidemiology and Distribution:
The global dissemination of BTV corresponds to the worldwide distribution of Culicoides vector
and changes in climatic. Climate change is a potential cause of this change especially in Europe. There
are twenty-seven BTV serotypes circulating worldwide (Maan et al. 2012; Jenckel et al. 2015) of which
twenty-two serotypes are endemic and co-circulate in southern Africa (Zulu and Venter 2014). Many
serotypes (BTV-1, 2, 4, 6, 8, 9, 11 and 16) entered Europe since 1998. BTV-10, 11, 13 and 17 are
present throughout the North America. Many serotypes as 1, 3, 5, 6, 9, 12, 14, 19, 22 and 24 have found
to exist in the southeastern parts of United States after 1998 (Maclachlan et al. 2013). The virus was
detected in 12 countries of Europe between 1998-2005 that caused death and culling of over I million
sheep (Purse et al. 2005).
In Australia, ten serotypes (1, 2, 3, 7, 9, 15, 16, 20, 21 and 23) of BTV were reported (Boyle et
al. 2012). BTV-6 was reported from Netherlands and Germany while, BTV-11 was identified from
Belgium in 2008 (Maclachlan and Guthrie 2010). In 2008, new serotype BTV-25 was isolated in
Switzerland. In Kuwait serotype 26 was identified in 2010 (Maan et al. 2012). Twenty-two different
serotypes of BTV were observed from all over India (Shafiq et al. 2013).
During 2003, a serological survey in cattle was performed in European Turkey and 23 provinces
in Anatolia that showed up to 90% of seroprevalence. BTV serotypes 4, 9 and 16 were observed
circulating in Anatolia, Turkey (see: Final Report EU project QLK2-CT-2000-00611). To device the
constrains on disease spread different strategies were applied such as disease surveillance, constraints on
animal whereabouts and live vaccination of animals with locally produced live attenuated vaccine
against serotype 4 (Mellor and Wittmann 2002).
A brief overview of epidemiological pattern and distribution of various serotypes in
chronological order is given in the table 2.1.
Review of Literature
8
Table 2.1: Epidemiological pattern and distribution of different serotypes of bluetongue virus
worldwide
Sr.
No.
Location Serotypes Year of
discovery
Reference
1 Portugal and South
West Spain
10 1956-1960 (Lopez and Botija 1958;
Manso-Ribeiro and Noronha
1958; Gorman 1990).
2 Australia 20 1976 (Kirkland 2004)
3 Europe 1, 2, 4, 6, 8, 9, 11, 16 1998 (Maclachlan et al. 2013)
4 Western and central
Europe
1, 6, 8, 11, 14, 25, 27 ---- (Jenckel et al. 2015;
Martinelle et al. 2016)
5 North America 10, 11, 13, 17 1998 (Maclachlan et al. 2013)
6 South-eastern United
States
1, 3, 5, 6, 9, 12, 14,
19, 22, 24
1998 (Maclachlan et al. 2013)
7 Australia 1, 2, 3, 7, 9, 15, 16,
20, 21, 23
---- (Boyle et al. 2012)
8 India 1–4, 6, 9, 10, 12, 16–
18, 21, 23
1967-
2014
(Rao et al. 2014)
9 Bulgaria 2 2000 (Hammami 2004)
10 Netherland and
Germany
6 2008 (Maclachlan and Guthrie
2010)
11 Belgium 11 2008 (Maclachlan and Guthrie
2010)
12 Switzerland 25 2008 (Maclachlan and Guthrie
2010)
13 Kuwait 26 2010 (Maan et al. 2012)
14 Anatolia, Turkey 4, 9, 16 ---- (Mellor and Wittmann 2002).
15 Western provinces,
Turkey
4 ---- (Erturk et al. 2004)
16 Corsica, France 27 2014 (Jenckel et al. 2014)
17 Morocco 1, 4, 8 2004-2015 (Cêtre-Sossah et al. 2011)
18 Pakistan 2, 6, 8, 9 2019 (Malik et al. 2019)
2.13. Molecular and Serological Diagnosis:
For antigen identification, both serological and molecular detection approach can be used. For
serological analysis, sandwich enzyme linked immunosorbent assay (ELISA) is used. Molecular assays
include serotype specific reverse transcriptase Polymerase chain reaction (RT-PCR), Serogroup-specific
RT-PCR, RNA polyacrylamide gel electrophoresis (RNA-PAGE), Real time quantitative PCR,
Restriction enzyme profile analysis (REPA), Sequencing and phylogenetic analysis, Molecular probes
for BTV identification (Bitew et al. 2013).
For antibody detection variety of serological tests are used. These are haemagglutination-
inhibition (HI), complement fixation test (CF), agar gel immunodiffusion (AGID), competitive enzyme-
linked immunosorbent assay (c-ELISA) and serotype-specific serum neutralization (SN) test (Khezri and
Azimi 2013). Competitive ELISA is higher in sensitivity and specificity than other serological tests that
exclude the chances of cross-reactivity at serogroup-level. It is rapid, reliable and ideally suitable test for
serotype confirmation (Yilmaz et al. 2012).
Review of Literature
9
2.14. Prevention and Control:
No specific therapy is available for the treatment of infected animals with BTV. The control
strategies include restriction on animal trade and movement, establishment of vector control by usage of
insecticides, slaughtering of infected animals and vaccination of vulnerable animals. Prophylaxis is most
effective in controlling bluetongue infection. Both the inactivated and live attenuated vaccines are
currently available and used (Bhanuprakash et al. 2009). Other vaccines are also being developed such
as recombinant vector vaccines and sub-unit vaccines etc. The development of new vaccines is necessary
because it rules out the risk of virus transmission and development of rapid onset of immune response
(Sperlova and Zendulkova 2011).
2.15. Statement of problem: Scarce information is available about seroprevalence and association of risk factors with
bluetongue virus in Pakistan. To establish an overall prevalence status of BTV in different livestock rich
areas of the country and later determine if the prevalence has any potential association with a range of
categorical variables or not was the main purpose of the study. Hence the study has been designed with
the following main objectives:
i. To determine the serum-based surveillance of antibodies against bluetongue virus in
different species of animals to fill up the paucity of data in wide geographical area of
Punjab and Balochistan, Pakistan
ii. To evaluate the association of potential risk factor with the absence or presence of
antibodies against bluetongue virus in different animals
iii. To perform molecular surveillance of BTV with the help of RT-PCR to establish the
understanding about genotype of BTV
10
CHAPTER 3
MATERIALS AND METHODS
3.1. Research funding statement:
The research activity was conducted under Higher Education Commission of Pakistan funded
project named “sero-epidemiology and molecular characterization of Bluetongue virus in Pakistan”
(grant no. 20-3307/NRPU/R&D/HEC/13/674) and DTRA (Defense threat reduction agency), USA
funded project named, “spatial ecology and epidemiology of soil borne human and animal bacterial
pathogens and its public health significance in Pakistan” (USA funded project (Project ID: HDTRA1-10-
1-0080) respectively.
3.2. Work station:
All the experimental studies and pratical laboratory work was done in molecular virology lab
(room# 104), Quality Operations Laboratories, UVAS, Lahore.
3.3. Experimental design:
Livestock is the major sub-sector that contributes 56% to country’s agriculture sector and 11% to
overall economy (Rehman et al. 2017). Punjab and Balochistan provinces were selected in the study. The
basis for selection of these provinces was to represent the seroprevalence status in economically most
important but representative areas that could also represent the two different animal breeding cultures.
Punjab is the most populous province contributing the major part of gross domestic product (GDP) of the
country and on the other hand Balochistan is the largest province in terms of area. Punjab represents the
in-house and manger feeding culture but on the other hand Balochistan represents totally nomadic
breeding. Keeping this in mind, a different sampling strategy was employed for both the provinces.
3.3.1. Sampling strategy for Punjab:
The used sera (n = 1312) and data were collected from select districts of Punjab province in
2013-2015 under an international collaborative research project related to epidemiology of soil-borne
zoonotic pathogens in Pakistan (Project ID: HDTRA1-10-1-0080). In multi-stage random sampling,
1312 animals (476 goats, 326 sheep, 234 cattle and 276 buffaloes) were selected from small select
villages where animal numbers in livestock barns ranged from 8-41. At first stage nine districts of the
province were selected through purposive sampling approach. The select districts named Faisalabad,
Chakwal, Attock, Gujranwala, Lahore, Dera Ghazi Khan, Sahiwal, Sargodha and Sheikhupura the main
rearing areas for livestock in Punjab, represent traditional agro-livestock mixed production system in the
country, with higher number of human and animal diseases incidence annually (Directorate of Animal
and Human Health, Punjab). In second stage, a sample size of villages (357) was estimated out of total
villages (4883) assuming prevalence (50%), confidence interval (95%), intra-class coefficient (0.4), and
margin of error (5%). According to probability proportional to size sampling, ten percent villages were
selected for valid sampling. Third stage involved convenient blood sampling of animals according to
prior history of soil-borne pathogen detection, farmer’s consent, presence of adult animals at livestock
barn, available resources and logistics. Given the frequent contact among animals while sharing water-
canal, grazing fields and nature of disease transmission being vector-borne, a village was considered as a
single large herd. In case, owner was not available or unwilling to cooperate, the next nearest neighbor
was approached for sampling. For better response from resident farms, the sampling was carried out in
close liaison with Livestock and Dairy Development Department of Punjab province and private
veterinary practitioners, whichever available for a particular region. A summary of number of samples
and herds from each district is given in Table 4.1, while different sampling locations are presented in
Figure 4.1.
3.3.2. Sampling strategy for Balochistan:
We used a multi-stage sampling technique for collection of blood from February to May in 2016.
Materials and Methods
11
Four districts; Kech, Killa Saifullah, Mastung and Quetta were selected for sampling. Quetta, Killa
Saifullah and Mastung are adjacent to Afghanistan whilst Kech shares a boundary with Iran and, to some
extent, coastal region via district Gawadar. According to a livestock census completed in 2015, the total
number of herds in these districts was 4768 (Livestock and Dairy Development Department,
Balochistan). Based on this, assuming 50% disease prevalence, 95% CI and 10% precision, the number
of herds required for sampling was estimated to be 97 using a proportional allocation method of stratified
sampling [ni= n×(Ni/N)] as described in Figure 4.2 (Pandey & Verma, 2008). A brief summary of the
number of herds and animals selected for blood collection from each district is given in Table 4.6.
Collection of blood involved convenient sampling of 10% of animals in each herd depending upon
secure accessibility and consent of the herd-owner. Collectively, 475 sheep and 401 goat samples were
collected (total n= 876).
3.4. Ethical approval: Prior approval of sampling and processing was obtained from the “Ethical Review Committee for
the Use of Laboratory Animals (ERCULA)” vide letter no. DR/69, dated: 17-1-2016 of UVAS, Lahore.
3.5. Sample collection: Approximately 5ml of blood samples from different species including goat, sheep, cattle and
buffalo were collected aseptically from the jugular vein directly and blood was added into the gel clot
activator tubes (VACUETTE®, VWR, Radnor, Pennsylvania, USA) placed on ice followed by
transportation at 4 ºC. A questionnaire was also filled at the sample collection site including the
information such as age, sex, breed, history of abortion, BTV vaccination, herd size, presence of midges
and use of acaricides etc. This information was used for further risk factor-based analysis in the study. A
copy of questionnaire is also attached at the end of this section.
3.6. Serum analysis through cELISA: Serum was separated from the blood samples contained in Gel/clot activator tube by
centrifugation for 10-15 minutes at 1300-1800xg. These serum samples were collected into separate
tubes and until further use was kept at -80 ºC. These samples were thawed and analyzed through
Competitive ELISA against BTV VP7 antigen for the detection of anti-VP7 antibodies. ID.VET kit
(Grabels, FRANCE) was used for this purpose. This kit (ID SCREEN® BLUETONGUE
COMPETITION) was suitable for sheep, goat, cattle, buffalo serum or plasma. The kit components were,
microplate coated with recombinant VP7 antigen (12 strips of 8 micro wells each), Anti-VP7-HRP-
conjugate (10X concentrated), Positive and Negative control, Wash concentrate (20X), Dilution buffer 2,
Substrate solution (TMB), Stop solution (H2SO4, 0.5 M). As per manufacturer’s instructions the protocol
was followed.
Anti-VP7-HRP-conjugate solution (1X) was prepared by diluting the Anti-VP7-HRP-conjugate
(10X) to 1/10 in Dilution Buffer 2. Washing solution (1X) was prepared by diluting the Wash
Concentrate (20X) to 1/20 in Distilled water. Dilution Buffer 2 (50 µl) was added in each well of
microtiter plate coated with recombinant VP7 protein of virus. Positive Control (50 µl) was added in well
A1 and 50 µl of negative Control was added in well B1. Serum samples to be tested were added 50 µl in
remaining wells. Plate was incubated at 21°C (±5 ºC) for 45 minutes ± 4 min. Then 100 µl of Anti-VP7-
conjugate (1X) was added in each well of plate. Plate was incubated for 30 minutes ±3 min at 21°C (±5
ºC). To wash each well three times 300 µl of washing solution (1X). Then in each well substrate solution
(100 µl) was added. Plate was incubated in dark at 21°C (±5 °C) for 15 minutes ±2 min. To stop the
reaction, 100 µl stop solution was added in each well. Optical density (OD) values were observed and
recorded at 450 nm by iMark™ Microplate Absorbance Reader (BIO-RAD) at serology lab in University
Diagnostic Laboratory of UVAS. For each sample competition percentage (S/N %) was calculated by
(S/N % = ODsample /ODNC × 100). Samples presenting S/N % greater than or equal to 40% were
considered negative, less than 40% were considered positive.
Materials and Methods
12
Figure 3.1: The cELISA plate showing positive and negative results of samples along with positive
control (PC) and negative control (NC)
3.7. Risk factor analysis of BTV:
The data obtained by cELISA and the information obtained through the questionnaire was
compiled in Microsoft Excel file and a univariate analysis was carried out in IBM SPSS statistics for
Windows version 21.0. (IBM SPSS Inc, Armonk, NY). In our study we used Chi-square test and
multivariable logistic regression model to analyze the association between different risk factors and
seroprevalence status of bluetongue disease. Risk factors or predictor variables are independent variables
that can predict the outcome variables. In our study the outcome variable was seropositive status in
animals. The predictor variables and outcome variables were both categorical. When both the predictor
variables and outcome are categorical then the suitable statistical tool is Chi-square test and/or regression
model. The logistic regression relates the natural logarithm of the odds ratio to linear combination of the
predictor variables (Rizzo et al. 2014). The odds ratio is a statistic that quantifies the strength of
association between two events. The methodology was followed as done previously for similar studies
for estimation of risk factors and the association with the disease seropositivity such as Shabbir et al.
2016. Association between seropositivity and categorical predictor variables was analyzed by Pearson’s
chi-square (χ2) test as mentioned by (Shabbir et al. 2016). Fisher’s Exact test was used as an alternate
method when any assumption of the chi-square test was violated. All variables with p-value ≤ 0.2 (two-
sided) in a univariate analysis were included in a mixed-effects logistic regression model with villages as
random effect. The mixed model was used to account for clustering effect as BTV is a vector borne
disease and frequent contact amongst animals within village is not unexpected. The regression model
included serological status of animals (positive versus negative) as dependent variable. The independent
fixed effects variables in the model were age, sex, breed, parity, presence of vector, history of abortion,
and herd size depending on outcome of univariate analysis. For each explanatory variable, category with
lowest prevalence was used as a reference. Any variable showing an association of ≤ 0.05 in regression
analysis carried out in R-software version 3.4.4 (lme4 package) was considered as statistically significant
for the disease. The sampling sites and the study outcomes in terms of seropositive or seronegative herd
and the distribution of BTV serotypes were mapped with Q GIS version 2.18.14.
3.8. Pooled samples from Balochistan:
A total of 41 herds were screened for BTV serotyping. The selected herds were comprised of 11
from Killa Saifullah, 8 from Mastung, 9 from Quetta and 13 from Kech district. From each herd, a pool
of 5-8 sera was processed for RNA extraction using QIAamp® DSP Virus Spin Kit (QIAGEN, Hilden,
Germany). The samples were pooled irrespective of their species (sheep or goat) as the two species are
reared in close proximity within the study sites and are equally susceptible to the virus.
PC
NC
Materials and Methods
13
3.9. Pooled samples from Punjab: A pool of 6-8 sera of each animal species from seropositive and seronegative herds was
processed in equal numbers for genome extraction and real-time PCR based genotyping. The pooled
samples (n = 30) included sheep (n = 9), goat (n = 11), cattle (n = 4) and buffalo (n = 6).
3.10. Genome extraction and quantification:
Total RNA was extracted from the serum samples using QIAamp viral RNA extraction kit
(QIAGEN, Hilden, Germany). An amount of 300 µl of serum sample was taken in vial to which 5.60 µl
RNA carrier was added and vortexed for 2-3 min. Then 560 µl a viral lysis buffer (AVL) was added and
incubated for 10 minutes at room temperature. Thereafter, chilled ethanol was added followed by
centrifugation of the all contents at 8000 rpm for 1 minute. After centrifugation, the column was washed
with wash buffer 1 (AW1) and wash buffer 2 (AW2) respectively. Finally, elusion buffer was added and
tube was centrifuged to collect the fluid having extracted RNA. Quantification was done through Nano-
drop sample retention system (Thermo Scientific™). An amount of 0.1 µl of extracted fluid was placed
on pedestal and concentration of genome was measured spectrophotometrically and value was given on
monitor screen connected with Nano-drop.
3.11. Real-time based genotyping:
The RNA samples were subjected to genome-based genotyping with the help of LSI VetMAXTM
European BTV Typing Real Time PCR kit (Thermo-Fisher Scientific-LSI, Lissieu, France) that is
capable of detection of eight serotypes individually (1, 2, 4, 6, 8, 9, 11 and 16) simultaneously from one
sample aliquot in eight different reaction tubes. The same well was used for the detection of both the
virus RNA as well as IPC (internal positive control). The detection of IPC in the sample indicates both
the efficiency of the extraction as well as the absence of the inhibitors in the sample. Eight external
positive controls (EPC BTVEUG(X) corresponding to eight different serotypes consisted of already
extracted genome of respective serotype were used to be denatured and then amplified in RT-PCR in the
same way as the samples.
3.12. RNA denaturation and amplification:
A total of 40µl of each RNA sample was used for one analysis requiring 5µl for eight serotypes.
5µl of RNA sample was added to each well in eight-well PCR strip. The strip was covered with the
caps and briefly spun the samples to settle them in mini centrifuge (MyFuge; Benchmark Scientific,
USA). The sample strip was then put into the Applied Biosystems™ MiniAmp™ Thermal Cycler
(Applied Biosystems, Foster City, California, USA) and the samples were heated for 3 minutes at 94 ºC
for denaturation of genome. After denaturation, the samples were placed on the ice block until further
use. Thawed all the eight tubes of Mix-BTVEUG(X) on ice block, where X corresponds to different
serotypes. Tubes were gently homogenized with the help of Digital mini vortex mixer
(VWR international, Germany) for about 10 seconds then were briefly centrifuged to settle down the
contents at bottom. Added 20µl of the Mix-BTVEUG(X) to each well of the eight-well PCR strip
containing denatured RNA samples. Positive amplification control, negative extraction control (NCS)
and negative amplification controls (NC) were also run along with the samples. For positive
amplification control, 5µl of denatured EPC BTVEUG(X) were used instead of samples. NCS
consisted of the contents used in the RNA extraction but the sample was replaced with RNA-free water
that undergoes the same treatment for extraction followed by RT-PCR as the sample. NC consists of
the amplification mix but the sample is replaced with RNA-free water. The strip wells were covered
with the lids and were placed in CFX96TM Real-time PCR Detection System (BioRad, California, USA)
and set up as follows; step 1: 45 ºC for 10 minutes (1x), step 2: 95 ºC for 10 minutes (1x), step 3: 95 ºC
for 15 seconds followed by 60 ºC for 45 seconds (40x). The fluorescence detection was measured at 60
ºC and 45 seconds part in step 3. The detectors for BTV(X) detections were set for FAMTM reporter
(non-fluorescent quencher) while for IPC, were set for VIC® reporter (non-fluorescent quencher) in
thermocycler with ROXTM as passive reference. The kit has been reported to be highly sensitive and
Materials and Methods
14
specific with high efficiency (95-100%). Determined by a series of dilution (usually 10-fold), the
efficiency of an assay corresponds to constant increase in target amplicon per cycle during exponential
phase. An ideal assay should have an efficiency value ranging from 90-100%, where one can have
confidence on primer and probe specificity, reaction condition and pipetting error etc. The assay was
performed, and Ct-values were recorded for interpretation as per manufacturer’s instruction keeping the threshold Ct-value to 40.
15
CHAPTER 4
RESULTS
4.1. Seroprevalence of BTV in Punjab:
A total of 1,312 samples were processed for detection of group-specific (VP7 protein) antibodies
against BTV from nine districts of Punjab province. Out of 117 total herds sampled, 71 were found
positive for group-specific antibodies against BTV giving an overall herd prevalence of 60.68% (95%
CI= 51.20-69.45) in the selected geographical area of the province. A maximum herd-wise
seroprevalence was found in DG Khan, Sahiwal, Chakwal, Attock i.e, 100% while least seroprevalence
was detected in district Sheikhupura i.e, 30.0% (Table. 4.1). An overall prevalence of 28.81% (95% CI=
26.39-31.36) was observed in both small and large ruminants combined. The level of seroconversion was
more in goats [40.76% (n= 194/476, 95% CI= 36.33-45.34)] followed by buffalo [29.34% (n= 81/276,
95% CI= 24.12-35.16)], sheep [18.40% (n= 60/326, 95% CI= 14.43-23.13)], and cattle [17.94% (n=
42/234, 95% CI= 13.38-23.60)]. The seroprevalence varied with respect to species and districts. A
district-wise summary of seroprevalence in each animal species is given in Table. 4.2.
Results
16
Table. 4.1. Herd-wise seroprevalence and serotype distribution of BTV among different districts of Punjab
No. Districts Herds Samples Seroprevalence Serotype distribution
Positive % prevalence 95% CI Sheep Goat Cattle Buffalo
1 Faisalabad 25 252 15 60.00 38.89 - 78.19 6 11 8 8
2 Sahiwal 9 65 9 100.0 62.88 - 100.0 8 - - -
3 Chakwal 4 20 4 100.0 39.58-100.0 8 2, 8, 16
4 Sargodha 8 56 3 37.50 10.24 - 74.11 - - - -
5 Attock 4 29 4 100.0 39.58-100.0 4 8
6 Gujranwala 15 157 11 73.33 44.83 - 91.08 8
7 Sheikhupura 30 524 9 30.00 15.41 - 49.56 - - - -
8 Lahore 20 199 14 70.00 45.67 - 87.16 2, 6, 8 8
9 Dera Ghazi
Khan 2 10 2 100.0 19.79 - 100.0 - - - -
Total 117 1312 71 60.68 51.20 - 69.45 -- -- -- --
Results
17
Table. 4.2. Animal species-wise distribution and seroprevalence of sheep, goat, cattle and buffalo in
Punjab province
District Type of
animal Animals tested
Animals
tested positive
Animal level
Prevalence % 95% CI
Attock*
Cattle 3 3 100.0 31.00 - 100.0
Sheep 5 5 100.0 46.29 - 100.0
Goat 21 21 100.0 80.76 - 100.0
Chakwal
Cattle 5 5 100.0 46.29 - 100.0
Buffalo 2 2 100.0 19.79 - 100.0
Sheep 4 2 50.00 09.19 - 90.81
Goat 9 7 77.78 40.19 - 96.05
Dera Ghazi
Khan*
Sheep 1 0 00.00 00.00
Goat 9 8 88.89 50.67 - 99.42
Faisalabad
Cattle 54 10 18.52 09.70 - 31.87
Buffalo 59 28 47.46 34.48 - 60.77
Sheep 64 21 32.81 21.90 - 45.80
Goat 75 41 54.67 42.81 - 66.05
Gujaranwala
Cattle 18 1 5.56 00.29 - 29.38
Buffalo 56 19 33.93 22.16 - 47.91
Sheep 34 7 20.59 09.34 - 38.41
Goat 49 16 32.65 20.36 - 47.65
Lahore
Cattle 48 6 12.50 05.19 - 25.94
Buffalo 59 12 20.34 11.39 - 33.20
Sheep 17 5 29.41 11.38 - 55.95
Goat 75 22 29.33 19.67 - 41.13
Sahiwal* Sheep 32 8 25.00 12.13 - 43.75
Goat 33 32 96.97 82.49 - 99.84
Sargodha Cattle 12 7 58.33 28.60 - 83.50
Buffalo 3 0 00.00 00.00
Results
18
Sheep 6 1 16.67 00.88 - 63.52
Goat 35 10 28.57 15.24 - 46.52
Sheikhupura
Cattle 94 10 10.64 05.50 - 19.12
Buffalo 97 20 20.62 13.34 - 30.28
Sheep 163 11 06.75 03.59 - 12.06
Goat 170 37 21.76 15.96 - 28.87
* The districts from which no sample was available for one or two species.
4.2. Risk factors analysis in Punjab: Outcome of association between seropositivity and risk factors were determined by univariate
analysis for small (Table 4.3) and large ruminants (Table 4.4), whereas the outcome of risk factors as
definite disease determinants is given in Table 4.5. Following univariate analysis for small ruminants,
though an initial association was observed for presence of vector in sheep, and herd size and age in goat
(Table 4.3); the odds of seroconversion were found insignificant when further analyzed in multivariate
logistic regression (Table 4.5). For cattle, the odds of seroconversion were five times more in animals
with a history of abortion (OR= 5.89, 95% CI= 1.80-19.27, p=0.003). Age, breed and presence of vector,
that initially showed an association with virus exposure (Table 4.4), were found to be insignificant as
disease determinants (Table 4.5). On the other hand, for buffaloes, the odds of seroconversion were more
for Nili breed (OR= 2.06, 95% CI= 1.19-3.58, p = 0.010), animals with history of abortion (OR= 3.95,
95% CI= 1.33-11.69, p = 0.013) and those that had an exposure to vectors (OR= 2.04, 95% CI=1.16-
3.59, p = 0.013) (Table 4.5). A village to village clustering effect was estimated by ICC value through
mixed-effects logistic regression model (Table 4.5). Such a congregation effect within a cluster is used to
group study farms/herds with similar production system and thereby, excluding any discriminatory effect
of a particular geographical area or farm conditions comparable to any other rearing and production
system in a country. A zero intra-class correlation coefficient (ICC) was observed for buffalo that
showed a lack of clustering effect between villages or an absolute lack of correlation between the
responses within the cluster. On the contrary, a highest clustering effect or ICC value was observed for
sheep (0.547), followed by goat (0.452) and cattle (0.248) and, hence a higher reliability of responses
within the cluster could be expected.
Results
19
Table. 4.3. Univariate analysis of anti-VP7 antibodies in small ruminants and their potential
association to categorical variables
Animal Variable Levels ELISA
positive
Total
tested
Seropositivity
(%) χ2 df p-value
Sheep
Age Young 33 195 16.92
0.710 1 0.400 Adult 27 131 20.61
Sex Female 45 260 17.31
1.030 1 0.310 Male 15 66 22.73
Breed
Buchi 19 76 25.00
2.963 2 0.227 Kajli 15 97 15.46
Lohi 26 153 16.99
Parity Nulliparous 47 259 18.15
0.056 1 0.813 Multiparous 13 67 19.40
Presence
of vector
No 18 77 23.38 1.659 1 0.198
Yes 42 249 16.87
History of
abortion
No 56 309 18.12 0.314 1 0.575
Yes 4 17 23.53
Herd/flock
size
<20 46 256 17.97 0.151 1 0.698
>20 14 70 20.00
Goats
Age Young 99 278 35.60
7.326 1 0.007 Adult 95 198 48.00
Sex Female 160 397 40.30
0.204 1 0.651 Male 34 79 43.00
Breeds
Beetal 68 141 48.20
5.497 4 0.240 Kajli 90 233 38.60
Nachi 25 71 35.20
Teddy 11 31 36.70
Parity Nulliparous 149 378 39.40
1.362 1 0.243 Multiparous 45 98 45.90
Results
20
Presence
of vector
No 136 340 40.00 0.282 1 0.595
Yes 58 136 42.60
History of
abortion
No 188 463 40.60 0.161 1 0.688
Yes 6 13 46.20
Herd/flock
size
<20 108 291 37.10 4.115 1 0.043
>20 86 185 46.50
* Presence of vector in sheep while, herd size and age in goat were found to be significant in goat in univariate analysis.
Results
21
Table. 4.4. Univariate analysis of anti-VP7 antibodies in large ruminants and their potential
association to categorical variables
Animal Variable Levels ELISA
positive
Total
tested
Seropositivity
(%) χ2 df p-value
Buffalo
Age Young 22 68 32.35
0.393 1 0.531 Adult 59 208 28.37
Sex Female 67 214 31.31
1.766 1 0.184 Male 14 62 22.58
Breed Nili 38 97 39.18
6.966 1 0.008 Nili-Ravi 43 179 24.02
Parity Nulliparous 34 128 26.56
0.893 1 0.345 Multiparous 47 148 31.76
Presence
of vector
No 26 123 21.14 7.212 1 0.007
Yes 55 153 35.95
History of
abortion
No 71 260 27.31 9.003 1 0.003
Yes 10 16 62.50
Herd/flock
size
<20 63 207 30.43 0.472 1 0.492
>20 18 69 26.09
Cattle
Age Young 18 131 13.74
3.579 1 0.059 Adult 24 103 23.30
Sex Female 32 184 17.39
0.182 1 0.670 Male 10 50 20.00
Breed
Cholistani 16 65 24.62
6.077 4 0.193
Dajal 6 31 19.35
Rajan 6 33 18.18
Red Sindhi 7 31 22.58
Sahiwal 7 74 09.46
Parity Nulliparous 30 171 17.54
0.071 1 0.790 Multiparous 12 63 19.05
Results
22
Presence
of vector
No 25 116 21.55 2.028 1 0.154
Yes 17 118 14.41
History of
abortion
No 33 215 15.35 12.15 1 0.000
Yes 9 61 14.75
*Breed, history of abortion, presence of vector and gender in buffalo while breed, history of abortion, presence of vector and age were found to be
significant in univariate analysis.
Results
23
Table. 4.5. Multivariable mixed effects logistic regression model (p < 0.05) for BTV serological
status amongst ruminants originating from traditional farms/villages in Punjab province (2013-15)
Animal Variable OR 95% CI Std. Error p-value
Buffalo
Fixed Parts
(Intercept) -- 00.16 00.08 - 00.33 00.06 <0.001
Breed (Nili) -- 02.06 01.19 - 03.58 00.58 0.010
History of
abortion(Yes) -- 03.95 01.33 -11.69 02.19 0.013
Presence of vector
(Yes) -- 02.04 01.16 - 03.59 00.59 0.013
Sex (Female) -- 01.24 00.62 - 02.48 00.44 0.537
Random Parts
Variance 00.00 -- -- -- --
ICC 00.00 -- -- -- --
Cattle
Fixed Parts
(Intercept) -- 00.06 00.02 - 0.18 00.03 <0.001
Breed (Cholistani) -- 03.43 00.89 - 13.25 02.37 0.073
Breed (Dajal) -- 02.27 00.45 - 11.45 01.88 0.319
Breed (Rajan) -- 02.20 00.45 - 10.92 01.80 0.333
Breed (Red Sindhi) -- 03.92 00.62 - 24.95 03.70 0.148
History of abortion
(Yes) -- 05.89 01.80 - 19.27 03.56 0.003
Presence of vector
(No) -- 01.29 00.46 - 03.65 00.68 0.629
Age group (Adult) -- 00.90 00.29 - 02.81 00.52 0.858
Random Parts
Variance 1.086 -- -- -- --
ICC 0.248 -- -- -- --
Sheep Fixed Parts
(Intercept) -- 00.12 00.04 - 00.38 00.07 <0.001
Results
24
Presence of
vector(Yes) -- 01.26 00.42 - 03.75 00.70 0.681
Random Parts
Variance 3.977
ICC 0.547
Goat
Fixed Parts
(Intercept) -- 00.54 00.30 - 00.98 00.16 0.043
Flock size (>20) -- 01.11 00.51 - 02.40 00.44 0.790
Age group(Adult) -- 01.12 00.54 - 02.30 00.41 0.759
Random Parts
Variance 2.712 -- -- -- --
ICC 0.452 -- -- -- --
4.3. Genotyping of BTV in Punjab:
Of the total samples processed for BTV genome detection, only 43.33% were found positive. The
prevalence of BTV serotype specific genome was detected in 6 of 9 districts only (Table 4.1). Serotypes
2, 4, 6, 8 and 11 in small ruminants, while 2, 8 and 16 were found in large ruminants. Interestingly,
serotype 8 was found exclusively in all positive districts and animal species indicating it to be the main
serotype of BTV prevailing in the province. In addition, serotype 4 was found exclusively in sheep,
serotype 11 was only found in goat while serotype 16 was found in cattle alone. The serotype distribution
in different districts of Punjab along with the sampling sites is presented in Figure 4.1.
Results
25
Figure. 4.1: Geographical distribution pattern of sites for blood sampling and identified serotypes
across selected districts of Punjab province, Pakistan
4.4. Seroprevalence of BTV in Balochistan:
Results
26
We processed a total of 876 sera of small ruminants originating from 4 selected districts of
Balochistan province (Quetta, Killa Saifullah, Mastung and Kech). An overall seroprevalance of 47.26%
(n = 414/876, 95% CI = 43.92-50.63%) was observed. Interestingly, none of the study herds (n=97) were
found to be seronegative. The prevalence of anti-VP7 antibodies was highest in Quetta district (49.6%,
95% CI = 43.89-55.46%) followed by Killa Saifullah (49.3%, 95% CI = 37.70-61.03%), Mastung
(47.7%, 95% CI = 40.72-54.89%), and Kech (44.0%, 95% CI = 38.33-49.82%) as shown in Table 4.6.
The frequency of seroconversion was higher in goats 50.87 % (210/475, 95% CI = 45.99-55.73%) than
sheep 44.21% (204/401, 95% CI = 39.81-48.70%) but statistically no difference (OR= 0.765) was
observed between both prevalence values. The prevalence of anti-VP7 antibodies in sheep was highest in
Mastung district (63%, 95% CI = 52.71-72.27%), followed by Killa Saifullah (49.33%, 95% CI = 37.70-
61.03%), Quetta (38.0 %, 95% CI = 30.31-46.31%) and Kech (35.33 %, 95% CI = 27.83-43.60%). In
contrast, Quetta district was found to have the highest seroconversion rate for goats (61.33 %, 95% CI =
53.01-69.06%) followed by Kech (52.66 %, 95% CI = 44.39-60.82%) and Mastung 32.67 % (95% CI
=23.86-43.82%), respectively.
Table 4.6. Summary of the number of sheep and goats sampled, the prevalence of anti-VP7
antibodies and the BTV serotypes present in the Kech, Mastung, Killa Saifullah and Quetta
districts of Balochistan province
No. District Total
herds
Herds
sampled
No. of Sera
samples
Prevalence
Sheep Goat Sheep Goat Anti-VP7
antibodies
Serotype
1 Kech 1352 13 17 150 150 44.00% 2
2 Mastung 823 12 9 100 101 47.76% 8
3 Killa
Saifullah
1137 13 0* 75 0 49.33% 8
4 Quetta 1456 21 12 150 150 49.60% 8, 9
Total 4768 59 38 475 401 47.26% - *= Due to limitation of accessibility and cooperation by respective herd-owner, no goat sample from Killa Saifullah could be obtained.
4.5. Risk factors analysis in Balochistan:
Chi-square based univariate analysis for seroconversion of goats revealed significant associations
(p<0.20) with breed (χ2 = 16.84, p= 0.01), parity (χ2 = 23.66, p = 0.00), presence of vector (χ2 = 2.63, p =
0.10) and herd size (χ2 = 8.67, p = 0.02). However, other risk factors such as age, gender and history of
abortion did not show any association with seroconversion (Table 4.7). For sheep, only breed (χ2= 13.80,
p= 0.01), parity (χ2= 53.40, p = 0.00) and history of abortion (χ2= 5.30, p= 0.02) showed significant
association for seroconversion (Table 4.8). Because use of acaricide was not practiced by any of the
herd-owners involved in the study, it was excluded from the risk factor analysis. Variables that showed
significant association in univariate analysis (p < 0.20) were further analyzed in a multivariate logistic
regression model analyses conducted separately for sheep (Table 4.9) and goats (Table 4.10). For goats,
the odds for seroconversion were greater in herds with the presence of vector (OR = 1.73, 95% CI =
1.06-2.85), for animals that were multi-parous [no parity (OR = 2.32, 95 % CI = 1.21-4.46), 2nd parity
(OR= 6.74, 95% CI = 2.73-16.65) and 3rd parity (OR = 5.16, 95% CI =1.46-18.25)], and for animals of
specific breeds [Jatan (OR = 3.92, 95% CI = 1.46-10.54) and Kajli (OR = 2.24, 95% CI = 1.01-5.00)].
Similarly, for sheep, the odds for seroconversion were found to be associated with parity and animal
breed. The seropositivity was found to be 6 times more in multi-parous animals (OR= 6.94, 95% CI =
3.10-15.53), while it was found to be 2 times more in animals of the Mengali breed (OR = 2.69, 95% CI
= 1.20- 6.03).
Results
27
Table 4.7. Univariate analysis of anti-VP7 antibodies in goats and their potential association to
categorical variables
Risk factors Categories Animals
tested
Seropositive
animals 95% CI χ2 p-value
Age <2 years
≥2 years
124
80
49.8%
52.6%
43.44-56.16%
44.40-60.72%
0.30 0.58
Sex Female
Male
124
80
51.2%
50.3%
44.77-57.67%
42.32-58.29%
0.03 0.85
Breed Barbari
Beetal
Jatan
Kajli
Kamori
Khursani
Lehri
75
29
30
26
10
25
9
48.4%
44.6%
75.0%
55.3%
58.8%
39.1%
69.2%
40.34-56.52%
32.47-57.41%
58.48-86.75%
40.24-69.54%
33.45-80.57%
27.36-52.08%
38.88-89.64%
16.84 0.01
Parity 0
1
2
3
140
18
35
11
50.5%
29.5%
74.5%
68.8%
44.51-56.56%
18.87-42.74%
59.36-85.58%
41.48-87.87%
23.66 0.00
Presence of
vector
No
Yes
85
119
46.4%
54.6%
39.11-53.94%
47.73-61.29%
2.63 0.10
History of
abortion
No
Yes
201
3
50.9%
50.0%
45.85-55.91%
13.95-86.05%
0.00 0.96
Herd size <100
≥100
99
105
44.0%
59.7%
37.45-50.75%
51.99-66.90%
8.67 0.02
Results
28
Table 4.8. Univariate analysis of anti-VP7 antibodies in sheep and their potential association to
categorical variables
Risk factors Categories Animals tested Seropositive
animals 95% CI χ2 p-value
Age ˂2 years
≥2 years
138
72
45.5%
41.9%
76.91-88.76%
34.47-49.64%
0.60 0.43
Sex Female
Male
117
93
45.9%
42.3%
39.68-52.21%
35.71-49.10%
0.62 0.43
Breed
Balochi
Bibrik
Harnai
Mengali
Rakhshani
Shinwari
58
4
43
53
14
38
37.9%
66.7%
50.6%
55.2%
29.8%
43.2%
30.30-46.14%
24.11-94.00%
39.60-61.52%
44.74-65.26%
17.79-45.08%
32.80-54.16%
13.80 0.01
Parity
0
1
2
3
151
10
39
10
44.8%
14.5%
83.0%
45.5%
39.44-50.30%
7.54-25.50%
68.65-91.86%
25.07-67.32%
53.40 0.00
Presence of
vector
No
Yes
107
103
44.4%
44.0%
38.06-50.92%
37.60-50.64%
0.00 0.93
History of
abortion
No
Yes
202
8
43.4%
80.0%
38.90-48.09%
44.22-96.46%
5.30 0.02
Herd size <100
≥100
122
88
44.0%
44.4%
38.14-50.11%
37.45-51.65%
0.00 0.93
Table 4.9. Multivariate logistic regression model determining the relationship between categorical
predictors and seropositivity in goats
Risk Factors B S.E Wald df Sig. Exp (B) 95% CI for Exp (B)
Lower Upper
Breed (Khursani)*
Breed (Barbari)
Breed (Beetal)
Breed (Jatan)
Breed (Kajli)
Breed (Kamori)
Breed (Lehri)
.519
-.094
1.367
.811
.854
.811
.320
.415
.504
.408
.592
.671
14.667
2.632
.051
7.359
3.942
2.082
1.460
6
1
1
1
1
1
1
.023
.105
.822
.007
.047
.149
.227
1.680
.911
3.925
2.249
2.350
2.249
.898
.404
1.461
1.010
.736
.604
3.143
2.055
10.542
5.006
7.497
8.379
Parity (1)*
Parity (0)
Parity (2)
Parity (3)
.844
1.909
1.642
.332
.461
.644
18.931
6.447
17.161
6.501
3
1
1
1
.000
.011
.000
.011
2.326
6.749
5.166
1.212
2.735
1.462
4.462
16.655
18.251
Presence of vector (Yes) .554 .253 4.796 1 .029 1.739 1.060 2.855
Herd (≥ 100)
Constant
.924
-2.008
.239
.461
14.976
18.991
1
1
.000
.000
2.518
.134 1.578 4.021 B= Unstandardized regression coefficient, S. E= Standard error, df= Degree of freedom, Sig.= p-value, EXP(B)= Odd ratio. *Khusrani breed, 1st parity, absence of vector and <100 Herd size was taken as reference.
Results
29
Table 4.10. Multivariate logistic regression model determining the relationship between categorical
predictors and seropositivity in sheep
Risk Factors B S.E Wald df Sig. Exp (B) 95% CI for Exp (B)
Lower Upper
Breed (Rakhshani) *
Breed (Balochi)
Breed (Bibrik)
Breed (Harnai)
Breed (Mengali)
Breed (Shinwari)
.311
1.737
.669
.991
.475
.394
.963
.426
.412
.418
10.512
.626
3.251
2.457
5.797
1.294
5
1
1
1
1
1
.062
.429
.071
.117
.016
.255
1.365
5.679
1.951
2.694
1.608
.631
.860
.846
1.202
.709
2.952
37.514
4.501
6.035
3.647
Parity (0) *
Parity (1)
Parity (2)
Parity (3)
-1.384
1.938
.043
.368
.411
.458
40.635
14.115
22.237
.009
3
1
1
1
.000
.000
.000
.926
.251
6.943
1.044
.122
3.103
.425
.516
15.536
2.562
Constant -.786 .357 4.839 1 .028 .456 B= Unstandardized regression coefficient, S.E= Standard error, df= Degree of freedom, Sig.= p-value, EXP(B)= Odd ratio.*Rakhshani breed, zero parity was taken as reference.
4.6. Genotyping of BTV in Balochistan: Pooled samples of sheep and goat from 41 herds were subjected to RNA extraction and qPCR-
based genotyping. Overall 14 of 41 (34.14%) herds had BTV genomes corresponding to serotypes 2, 8
and 9. Serotype 8 was the most frequent (26.82%, 95% CI = 14.75-43.21%) followed by an equal
occurrence of serotype 2 and 9 (7.31%, 95% CI= 1.91-21.01%). Although definitive statements would
require completion of large scale genome-based identification studies involving larger numbers of herds
and samples, we observed a distinct pattern of BTV distribution in the studied districts. Serotype 8 was
detected in 45.45% of herds in Killa Saifullah (5/11), 37.5% of herds in Mastung (3/8) and 33.3% of
herds in Quetta (3/9). In contrast serotype 9 was only detected in herds from Quetta (33.3% - 3/9) as a
co-infection with serotype 8 and serotype 2 was detected exclusively in herds from Kech (23.07% - 3/13)
as shown in Figure 4.2.
Results
30
Figure. 4.2: Geographical distribution pattern of sites for blood sampling and identified serotypes
across selected districts of Balochistan province, Pakistan
31
CHAPTER 5
DISCUSSION
This is the first large-scale study on prevalence status of BTV group-specific antibodies and its
circulating serotypes in Pakistan. Different assays such as agar gel immunodiffusion (AGID),
haemagglutination-inhibition (HI), complement fixation test (CFT), and serotype-specific serum
neutralization (SN) test have been employed for serological evidence of BTV previously (Boulanger et
al. 1967; Van der Walt 1980; Herniman et al. 1983; Koumbati et al. 1999). However, each of these
assays have certain limitations including cross-reactivity among closely related serotypes, laborious and
time-consuming protocols, sub-optimal sensitivity and specificity and subjective interpretation which
could lead to inter-laboratory and inter-researcher variability (OIE, 2014; Khezri and Azimi 2013). In
contrast, cELISA provides rapid, accurate and precise assessment of exposure to BTV (Yilmaz et al.
2012). The serological marker (VP7) is considered sero-group specific for all BTV serotypes and,
therefore, is being used extensively for routine disease surveillance and diagnostics (van Regenmortel et
al. 2000; de Diego et al. 2011) has previously been validated by others (Vandenbussche et al. 2008).
Since it involves a competition between test serum and monoclonal antibodies, potential sero-group level
cross-reactivity by other arboviruses is excluded (Afshar 1994; Yilmaz et al. 2012). Similarly, a
commercially available RT-PCR protocol has previously been validated with a high efficiency and
absolute lack of cross-reactivity to accurately identify each of eight different serotypes in animal sera
(Thermo-Fisher Scientific-LSI, Lissieu, France).
Although varying, we observed seroconversion in all studied animal species. From Punjab, A
greater proportion of small ruminants, 31.67% (254/802, 95% CI= 28.48-35.03) were seropositive than
large ruminants, 24.12% (123/510, 95% CI= 20.52-28.12). The rate of seroconversion was higher in
goats, 40.76% (194/476, 95% CI= 36.33-45.34) followed by buffalo, 29.34% (81/276, 95% CI= 24.12-
35.16), sheep, 18.40% (60/326. 95% CI= 14.43-23.13) and cattle, 17.94% (42/234, 95% CI= 13.38-
23.60). While in Balochistan, we found a higher seroconversion rate in goats 50.87 % (204/401, 95% CI
= 45.99-55.73%) than sheep 44.21% (210/475, 95% CI = 39.81-48.70%).
Previously a seropositivity of 48.4% in sheep flocks has been documented from KPK province
(Akhtar et al. 1997). Similar observations for BTV have also been reported from the neighboring
countries. For instance, a prevalence of 85.3% in goat and 74.4% in sheep has been reported from Iran
(Oryan et al. 2014). Another study from the same country evidenced a seroprevalence of 74.2% and
72.9% in goat and sheep, respectively (Mohammadi et al. 2012). Joardar et al. (2013) reported a higher
rate of seropositivity in sheep (58.82 %) than goat (31.79 %) from North-Eastern state of India. A study
originating from West-Bengal (India) demonstrated a higher occurrence of seropositivity in goat
(66.95%), followed by sheep (57.66%) and cattle (52%) (Panda et al. 2011). Another study from India
(northern Kerala) revealed a prevalence of 16%, 7.5% and 6.9% in sheep, goat and cattle, respectively
(Arun et al. 2014). Gaire et al. (2016) reported a seropositivity of 29.3% in dairy cattle in Nepal (Gaire et
al. 2016). Taken together, one can speculate that all the animals are equally susceptible to BTV. There
may be differences in clinical form of disease where severity could be more often observed in one specie
than others. For example, sheep is considered more vulnerable to infection and exhibit more severe form
of infection even at same level of viremia in goat (Koumbati et al. 1999). Similarly, seroprevalence is
also reported in other animal species like camel in Iran (100%) (Mahdevi 2006), Saudi Arabia (25.7%)
(Yousef et al. 2012) and Gujrat, India (19.3%) (Chandel et al. 2003). Previously it was known that dogs
can get infected with BTV by ingestion of infected meat but a study provided evidence of dogs being
naturally infected (21%) with the virus (Oura and El Harrak 2011). Wild animals in Spain were also
found to be seropositive such as Red deer (21.9%), Fallow deer (35.4%), Roe deer (5.1%), Mouflon
(13.2%) and Aoudad (25%) (Ruiz-Fons et al. 2008).
Discussion
32
In fact, the incidence rate and severity of clinical disease depend upon various factors that
include, but are not limited to levels of herd immunity, virulence of circulating BTV strains and the
inherent genetic resistance of different animals (Caporale et al. 2014). Occurrence of BTV is considered
to be subclinical in cattle (Arun et al. 2014), however it could serve as a source of infection through virus
transmission to and from vectors to susceptible animals (Schwartz-Cornil et al. 2008). The outcomes
revealed a higher ICC value for sheep followed by goat, cattle and buffalo. Considering cluster sampling
and subsequent analysis, therefore, one can speculate a higher consistency for disease occurrence in
small ruminants than large ruminants. Hence, to better elucidate clustering effect, a smaller sampling size
for small ruminants than large ruminants may be considered enough to predict a reliable disease status in
the region.
In Punjab, The Nili breed of buffalo was found at a greater risk of BTV infection than any other
study breed of small and large ruminants. While Balochistan, the frequency of seroconversion was
observed to vary between different breeds of animals; the frequency of BTV serconversion was highest
in Jatan and Kajli breeds of goat and the Mengali breed of sheep. A little or no impact of breed
susceptibility to infection has previously been reported for small ruminants (Caporale et al. 2014).
Similarly, in the Mediterranean area (Sardinian and Italian mixed breed) and Northern Europe (Dorset
poll breed) animal breed was not found to be a major factor in susceptibility to BTV seroconversion
(Koumbati et al. 1999). In another study both indigenous and exotic cattle breeds were found to be
equally susceptible in a previous study (Adam et al. 2014). Although a significant difference in
seroconversion was observed for type of animal species (Tibetan sheep vs yak), rate of difference in
seroconversion within breeds of sheep and yak was found negligible (Ma et al. 2017).
History of abortion was observed to be associated with an increased seropositivity in cattle and
buffalo in Punjab. In previous studies it was found that, some serotypes like serotype 8 can also cause
abortion in cattle as observed in outbreak in Netherlands in 2007, that caused abortions and brain defects
in calves (Wouda et al. 2008). The same serotype was also observed in cattle in our study. However, we
are uncertain that the main cause of infection was previous BTV infection as it rarely causes abortion in
cattle. Secondly, we did not isolate the virus in aborted fetuses. This area is needed to be explored further
in future studies. A similar relationship between abortion and seropositvity has previously been
documented (Akhtar et al. 1997) and (Yavri et al. 2018) in sheep, whereas in Balochistan, abortion and
seropositivity status was insignificant similar to Gaire et al. (2016) and Sabaghan et al. (2014), who
documented a lack of any association between abortion and BTV seropositivity (Sabaghan et al. 2014;
Gaire et al. 2016).
The presence of vector was also found to be a significant factor for BTV infection in buffalo.
Similarly, we also found this susceptibility in goats of Balochistan. It is quite obvious for being a vector-
borne disease of small and large ruminants where a remarkable association with presence of disease
vector in animal vicinity has been reported (Bouwknegt et al. 2010).
Though we did not find an association between animal age with disease seroconversion in the
animals of both provinces, adult animals are reported to be more seropositive than the younger ones
(Radostits et al. 2007; Adam et al. 2014; Khair et al. 2014) simply due to fact that chances of exposure
increase with age. A potential reason for almost equal susceptibility in all age groups may be that all
animals were living in close vicinity corresponding to a recent observation made by Ma et al. (2017),
who also reported lack of association between seroconversion and season, age, gender or region.
Numerous factors such as warm temperature, humid environment, transhumant animal
movement, introduction of animal in a herd, rainfall and insect spray are reported to have a positive and
negative association for animal seropositivity (Akhtar et al. 1997; Becker et al. 2003; Adam et al. 2014;
Khair et al. 2014; Gao et al. 2017). Since collection of information pertaining to these factors require
periodic farm visits or access to farmers, environmental variables, nationwide disease surveillance
records, implementation of disease control strategy in terms of use of acaricide, we were not able to
correlate study outcome with these factors and, therefore owing to these limitations, study outcomes and
its subjective interpretation should be considered accordingly.
Discussion
33
Because determining the prevalent serotypes in a particular geographical setting provides an
important insight towards disease epidemiology, its subsequent diagnostics and control strategies, we
evidenced real-time PCR based prevalence of BTV serotype for the first time in Pakistan. More than one
serotype was detected from different animal species where serotype 8 was the most dominant in all of the
studied animals across the positive districts. Serotypes 6, 8 and 11 were found in Faisalabad, 2, 8, 16 in
Chakwal, 2, 8 and 16 in Lahore, while serotype 4 and 8 were found in Attock. Only one serotype, 8 was
found in both Sahiwal and Gujranwala while in three districts Sargodha, Sheikhupura and Dera Ghazi
Khan, no serotype was observed. Three different serotypes (2, 8 and 9) were observed in Balochistan in
which serotype 8 being the most frequent and seen in multiple districts, serotype 9 seen only in Quetta
district and only in a subset of the samples that were positive for serotype 8, and serotype 2 only seen in
Kech district where neither serotype 8 or 9 were evident.
The occurrence of more than one serotype has been documented worldwide. Notably, in many
instances of such ‘mixed’ infections, the clinical severity of particular strains shows marked variability.
For instance, serotypes 1, 6, 8, 11, 14, 25, and 27 were detected in Western and Central Europe (Jenckel
et al. 2015; Martinelle et al. 2016) and of these, serotype 8 was found to be the most devastating,
causing the death of 5-10% of the national flock in Belgium alone (Saegerman et al. 2011), while, a
negligible clinical impact was reported from France (Sailleau et al. 2017). Similary, whilst serotypes 1, 4
and 8 were identified in Morocco between 2004 and 2015, severe clinical disease was ascribed to
serotypes 1 and 4 rather than 8 (Cêtre-Sossah et al. 2011). A similar pattern of disease severity for
serotype 1 and 4 was observed in Central and Western Europe in 2006 (Saegerman et al. 2011).
Serotypes 1, 2, 3, 4, 6 and 10 are considered highly pathogenic that have potential to cause severe
epidemics (Dungu et al. 2004). Genetic shift and drift are responsible for the genetic diversity in
different serotype and also within the same serotype in different regions (OIE, 2007). An outbreak in
Italy was reported to be caused by serotype 2 and 9 that caused 11% mortality (Savini et al. 2005). A
total of 23 serotypes have been reported from India (Sairaju et al. 2013). Despite non-virulent nature of
serotype 21 worldwide, clinical cases were observed in goats (Susmitha et al. 2012; Rao et al. 2016).
Taken together, an apparent variation in clinical nature of disease induced either by same and/or different
serotypes within same geographical area could be ascribed to segmented BTV genome where
reassortment among different serotypes could yield emergence of novel serotypes with varying clinical
outcome (Shafiq et al. 2013; Caporale et al. 2014). Although we observed an increased prevalence of
disease, clinical cases of BT are not much frequently reported in study areas unlike other diseases with
similar clinical symptoms such as peste des petits ruminants virus. There could be many reasons that
includes a relatively less-virulent nature of prevailing serotypes such as observed in Australia where
BTV serotypes are either non-virulent or less virulent in nature leading to absence of clinical disease in
the country (Kirkland et al. 2004), lack of routine diagnostics and disease surveillance, lack of
differential diagnosis with other closely related diseases like vesicular stomatitis, goat pox, sheep pox,
foot and mouth disease and peste des petits.
There were several study limitations. For seroprevalence study in Punjab, we used archived sera
collected previously in another project. Therefore, sample number was not uniform and/or representative
of animal population across select districts and a very little information was available to correlate with
respect to BTV infection. Also we were not able to discriminate titers suggestive of either current or past
infection because of lack of proper animal tracking system in the country where resampling from the
same animal is not possible in another time. Sensitivity and specificity of cELISA has previously been
evaluated for cattle and sheep sera only (Vandenbussche et al. 2008); nonetheless, we also used buffalo
and goat sera. Test validity has neither been assessed for buffalo and goat sera previously anywhere nor
available validity results are pertaining to Pakistan in particular; above reference study was conducted in
Belgium. This is important because disease outbreaks have been reported from Europe and Belgium with
BTV serotype 8 whereas, despite abundant prevalence in all animal species studied, there has been no
such evidence of clinical disease in Pakistan so far. Though needs further investigation in subject matter,
together it indicates potential variability in disease susceptibility to indigenous breed and thus, variation
in prevalence rate may be expected despite using previously published sensitivity and specificity of
Discussion
34
cELISA assay. Considering real-time PCR, we were able to process only a limited number of pooled sera
within available budget using a commercially available kit. Because the kit offers detection of 8 different
serotypes in a sample when same genome is processed separately in 8 different reaction tubes, pooling of
sera originating from a herd may not affect presence of more than one prevalent serotype in that
particular sample.
Conclusion:
Bluetongue is prevalent in both Punjab and Balochistan. Seroprevalence and serotype distribution
is different in different districts irrespective of animal species. The disease was also found to be
associated with certain risk factors that can be potential determinants of disease seropositivity. In Punjab,
buffalo of Nili breed, history of abortion, presence of vector in the rearing area were potential risk factors
for seropositive status. In cattle, history of abortion was the potential risk factor. Serotypes detected in
Punjab were serotype 2, 4, 6, 8, 11 and 16. In Balochistan the potential risk factors in goats were breed
(Jatan and Kajli), parity and presence of vector. In sheep, mengali breed and parity were found to be a
risk factor. Serotype 2, 8 and 9 were the serotypes detected in the country.
Suggestion:
Bluetongue virus is considered subclinical in Pakistan. In the present study different serotypes
were detected in both the provinces. It is quite possible that these serotypes are sub-clinical in the
country despite of the fact that some of these serotypes such as serotype 4 and show severe clinical
disease in animals especially in goat. On contrary, the higher seroprevalence status of BTV in all the
species suggests that there is possible misdiagnosis or under-diagnosis of this disease due to its clinical
signs similarity with other viral diseases. So in future, this aspect should be considered in field diagnosis.
Secondly there is complete absence of surveillance, vaccination, vector control and treatment strategies
at government level. Higher seroprevalence status and prevailing serotypes in the country suggest that
there is need to implement these strategies at mass level. Thirdly, as we detected virus serotypes from
serum, there is need to detect and isolate active virus from field so that we can use it as control for
further studies making detection of other than these 8 European serotypes possible.
35
CHAPTER 6
SUMMARY
Bluetongue is an infectious disease of ruminants. The causative agent is Bluetongue virus that
has double stranded RNA genome. Culicoides biting midges act as a vector for virus transmission. There
are 27 bluetongue virus serotypes present worldwide. Bluetongue disease mainly affects sheep, goat,
cattle, and camelids species. The current status of bluetongue virus existence is unknown in Pakistan.
Competitive ELISA is proven to be a rapid and more specific test for the identification of antibodies to
bluetongue virus. Molecular surveillance further confirms the results of serology and provides
opportunity for further analysis. The hypothesis of the study was; “Different serotypes of bluetongue
virus are prevalent in Pakistan and their prevalence is associated with certain risk factors”.
Blood samples were collected from selected districts of Punjab and Balochistan. In Punjab, 1312
samples of sheep (326), goat (476), cattle (234) and buffalo (276) were collected from Faisalabad,
Chakwal, Attock, Gujranwala, Lahore, Dera Ghazi Khan, Sahiwal, Sargodha and Sheikhupura. While, in
Balochistan, total 876 blood samples were collected from sheep (n = 475) and goats (n= 401) from Kech,
Killa Saifullah, Mastung and Quetta. Serum separated from blood collected in Gel/Clot activator tubes
was tested by Competitive ELISA by kit method (ID SCREEN® BLUETONGUE COMPETITION).
Both the seropositive and seronegative herds were selected from each district from Punjab and
Balochistan. Herd-wise sample distribution from each herd is given in the table 4.1 and 4.6 from Punjab
and Balochistan respectively. This selection was performed irrespective of the animal species assuming
the fact that each animal species was equally likely to be infected. Later, these samples were subjected to
RNA extraction using RNA extraction kit (QIAamp viral RNA extraction kit). Those RNA samples were
then used for real-time based genomic surveillance for serotype detection using LSI VetMAXTM
European BTV Typing Real-Time PCR kit.
The association between seropositive/seronegative status and categorical variables (risk factors) was
determined with the help of Pearson’s Chi-square (χ2) test. At first the p-value was set to ≤ 0.2 in
univariate analysis to include more of the possible risk factors in the study. After that the multivariate
logistic regression model with a backward LR method of selection was used to estimate the predictive
values of individual risk factors keeping the p-value at 0.05.
In Punjab, the overall prevalence was observed to be 28.81% in small and large ruminants. The
highest prevalence found in the province was in goats (40.75%) followed by buffalo (29.34%), sheep
(18.40%) and cattle (17.94%). The risk factor analysis showed that buffalo of Nili breed and found with
presence of vector were at higher risk of being seropositive for BTV. Both cattle and buffalo with history
of abortion were found to be much likely carrying the disease. Real-time based serotyping showed the
detection of serotypes 4, 6 and 8 in sheep, 2, 6, 8 and 11 in goats, 2, 8 and 16 in buffalo and only 8 in
cattle. As for as Balochistan is concerned, the overall seroprevalence of BTV was found to be 47.26% in
sheep and goats. Seroprevalence was more in goats (50.87%) than sheep (44.21%). Parity and breed
(Jatan and Kajli breed in goats, Mengali breed in sheep) were found to be risk factors for disease
occurrence. The presence of vector was also observed to be a risk factor in goats. Serotype 8 was again
the most prevalent serotype in Balochistan followed by an equal prevalence of serotypes 2 and 9.
This study contributed to appraise the overall status of BTV prevalence in Pakistan in association
to potential risk factors. Molecular surveillance helped to assess the country’s status regarding prevailing
serotypes and also to estimate its contribution in circulation of those prevailing serotypes in comparison
to nearby countries.
36
CHAPTER 7
LITERATURE CITED
Adam IA, Abdalla MA, Mohamed ME, Aradaib IE. 2014. Prevalence of bluetongue virus infection and
associated risk factors among cattle in North Kordufan State, Western Sudan. BMC Vet. Res. 10:
94.
Afshar A. 1994. Bluetongue: laboratory diagnosis. Comp. Immunol. Microbiol. Infect. Dis. 17: 221-242.
Akhtar S, Djallem N, Shad G, Thieme O. 1997. Bluetongue virus seropositivity in sheep flocks in North
West Frontier Province, Pakistan. Prev. Vet. Med. 29: 293-298.
Akhtar S, Howe R, Jadoon J, Naqvi M. 1995. Prevalence of five serotypes of bluetongue virus in a
Rambouillet sheep flock in Pakistan. Vet. Record. 136: 495-495.
Alexander G, Alexander M, St George T 1996. Bluetongue-its impact on international trade in meat and
livestock. In Bluetongue Disease in Southeast Asia, Volume 66. (Australian Centre for
International Agricultural Research), pp. 254-258.
Ali AM, Dennison T, Schauwers SA, Ziay G, Qanee AH. 2014. Bluetongue virus sero survey in sheep,
goats and cattle flocks of afghanistan. BTV survey report.
https://www.researchgate.net/profile/Ali_Safar_Maken_Ali/publication/269396828_Authors_Dr
_Maken_Ali/links/548935670cf2ef344790acbc/Authors-Dr-Maken-Ali.
Anon. 2000. Disease outbreaks reported to the OIE in September–October 2000. Bull. Off. Int. Epizoot.,
112, 552.
Anthony S, Jones H, Darpel K, Elliott H, Maan S, Samuel A, Mellor P, Mertens P. 2007. A duplex RT-
PCR assay for detection of genome segment 7 (VP7 gene) from 24 BTV serotypes. J. Virol.
Methods. 141: 188-197.
Arun S, John K, Ravishankar C, Mini M, Ravindran R, Prejit N. 2014. Seroprevalence of bluetongue
among domestic ruminants in Northern Kerala, India. Tropic. Biomed. 31: 26-30.
Barratt-Boyes S, Rossitto P, Stott J, MacLachlan N. 1992. Flow cytometric analysis of in vitro
bluetongue virus infection of bovine blood mononuclear cells. J. Gen. Virol. 73: 1953-1960.
Becker N, Petrić D, Boase C, Lane J, Zgomba M, Dahl C, Kaiser A. 2003. Mosquitoes and their control.
New York, Boston, Dordrecht, London, Moscow, Plenum Publishers. 2: 498.
Bhanuprakash V, Indrani B, Hosamani M, Balamurugan V, Singh R. 2009. Bluetongue vaccines: the
past, present and future. Expert Rev. Vaccines. 8: 191-204.
Bhattacharya B, Noad RJ, Roy P. 2007. Interaction between Bluetongue virus outer capsid protein VP2
and vimentin is necessary for virus egress. Virol. J. 4: 1.
Bitew M, Sukdeb N, Ravishankar C. 2013. Bluetongue: Virus proteins and recent diagnostic approaches.
Afr. J. Microbiol. Res. 7: 5771-5780.
Bonneau K, DeMaula C, Mullens B, MacLachlan N. 2002. Duration of viraemia infectious to Culicoides
sonorensis in bluetongue virus-infected cattle and sheep. Vet. Microbiol. 88: 115-125.
Boulanger P, Ruckerbauer G, Bannister G, Gray D, Girard A. 1967. III. Comparison of Two
Complement-Fixation Methods. Can. J. Comp. Med. Vet. Sci. 31: 166.
Bouwknegt C, van Rijn PA, Schipper JJ, Hölzel D, Boonstra J, Nijhof AM, van Rooij EM, Jongejan F.
2010. Potential role of ticks as vectors of bluetongue virus. Exp. App. Acarol. 52: 183-192.
Boyce M, Wehrfritz J, Noad R, Roy P. 2004. Purified recombinant bluetongue virus VP1 exhibits RNA
replicase activity. J. Virol. 78: 3994-4002.
Caporale M, Di Gialleonorado L, Janowicz A, Wilkie G, Shaw A, Savini G, Van Rijn PA, Mertens P, Di
Ventura M, Palmarini M. 2014. Virus and host factors affecting the clinical outcome of
bluetongue virus infection. J. Virol. 88: 10399-10411.
Literature Cited
37
Cêtre-Sossah C, Madani H, Sailleau C, Nomikou K, Sadaoui H, Zientara S, Maan S, Maan N, Mertens P,
Albina E. 2011. Molecular epidemiology of bluetongue virus serotype 1 isolated in 2006 from
Algeria. Res. Vet Sci. 91: 486-497.
Chandel BS, Chauhan HC, Kher HN. 2003. Comparison of the standard AGID test and competitive
ELISA for detecting bluetongue virus antibodies in camels in Gujarat, India. Trop. Anim. Health
Prod, 35(2): 99-104.
Clavijo A, Sepulveda L, Riva J, Pessoa-Silva M, Tailor-Ruthes A, Lopez J. 2002. Isolation of bluetongue
virus serotype 12 from an outbreak of the disease in South America. Vet. Rec. 151: 301-301.
Coetzee P, Van Vuuren M, Stokstad M, Myrmel M, Venter EH. 2012. Bluetongue virus genetic and
phenotypic diversity: towards identifying the molecular determinants that influence virulence and
transmission potential. Vet. Microbiol. 161: 1-12.
Conte A, Goffredo M, Ippoliti C, Meiswinkel R. 2007. Influence of biotic and abiotic factors on the
distribution and abundance of Culicoides imicola and the Obsoletus Complex in Italy. Vet.
Parasitol. 150: 333-344.
Dal Pozzo F, Saegerman C, Thiry E. 2009. Bovine infection with bluetongue virus with special emphasis
on European serotype 8. Vet. J. 182: 142-151.
Darpel K, Batten C, Veronesi E, Shaw A, Anthony S, Bachanek-Bankowska K, Kgosana L, Bin-Tarif A,
Carpenter S, Müller-Doblies U. 2007. Clinical signs and pathology shown by British sheep and
cattle infected with bluetongue virus serotype 8 derived from the 2006 outbreak in northern
Europen. Vet. Rec. 161: 253-261.
de Diego ACP, Athmaram TN, Stewart M, Rodríguez-Sánchez B, Sánchez-Vizcaíno JM, Noad R, Roy P.
2011. Characterization of protection afforded by a bivalent virus-like particle vaccine against
bluetongue virus serotypes 1 and 4 in sheep. PloS One. 6: e26666.
De Liberato C, Purse B, Goffredo M, Scholl F, Scaramozzino P. 2003. Geographical and seasonal
distribution of the bluetongue virus vector, Culicoides imicola, in central Italy. Med. Vet.
Entomol. 17: 388-394.
De Liberato C, Scavia G, Lorenzetti R, Scaramozzino P, Amaddeo D, Cardeti G, Scicluna M, Ferrari G,
Autorino G. 2005. Identification of Culicoides obsoletus (Diptera: Ceratopogonidae) as a vector
of bluetongue virus in central Italy. Vet. Rec. 156: 301.
Dercksen D, Groot NN, Paauwe R, Backx A, Vellema P. 2007. First outbreak of bluetongue in goats in
the Netherlands. Tijdschr. Diergeneesk. 132: 786-790.
Diprose J, Burroughs J, Sutton G, Goldsmith A, Gouet P, Malby R, Overton I, Zientara S, Mertens P,
Stuart D. 2001. Translocation portals for the substrates and products of a viral transcription
complex: the bluetongue virus core. EMBO J. 20: 7229-7239.
Dragonas PN. 1981. Evolution of bluetongue in Greece. OIE Monthly Circ, 9(417), 10-11.
Drew CP, Heller MC, Mayo C, Watson JL, MacLachlan NJ. 2010. Bluetongue virus infection activates
bovine monocyte-derived macrophages and pulmonary artery endothelial cells. Vet. Immunol
Immunopathol. 136: 292-296.
Dragonas PN. 1981. Evolution of bluetongue in Greece. OIE Monthly Circ, 9(417), 10-11.
Drew CP, Heller MC, Mayo C, Watson JL, MacLachlan NJ. 2010. Bluetongue virus infection activates
bovine monocyte-derived macrophages and pulmonary artery endothelial cells. Vet. Immunol
Immunopathol. 136: 292-296.
Du Toit R. 1944. The transmission of bluetongue and horse sickness by Culicoides. Onderstepoort J. Vet.
Sci. Anim. Ind. 19: 7-16.
Dungu B, Gerdes T, Smit T. 2004. The use of vaccination in the control of bluetongue in southern
Africa. Vet. Ital. 40(4): 616-22.
Elbers AR, Backx A, Mintiens K, Gerbier G, Staubach C, Hendrickx G, van der Spek A. 2008. Field
observations during the Bluetongue serotype 8 epidemic in 2006: II. Morbidity and mortality
rate, case fatality and clinical recovery in sheep and cattle in the Netherlands. Prev. Vet. Med. 87:
31-40.
Literature Cited
38
Erturk A, Tatar N, Kabakli O, Incoglu S, Cizmeci S, Barut F. 2004. The current situation of bluetongue
in Turkey. Vet Ital. 40: 137-140.
Evermann JF. 2008. Accidental introduction of viruses into companion animals by commercial
vaccines. Vet Clin North Am Small Anim Pract. 38(4): 919-929.
Forzan M, Marsh M, Roy P. 2007. Bluetongue virus entry into cells. J. Virol. 81: 4819-4827.
Forzan M, Wirblich C, Roy P. 2004. A capsid protein of nonenveloped Bluetongue virus exhibits
membrane fusion activity. Proceedings of the National Academy of Sciences. 101: 2100-2105.
Gaire T, Karki S, Dhakal I, Khanal D, Bowen R. 2016. Serosurveillance and factors associated with the
presence of antibodies against bluetongue virus in dairy cattle in two eco-zones of Nepal. Rev.
Sci. Tech. (International Office of Epizootics). 35: 779.
Gambles R. 1949. Bluetongue of sheep in Cyprus. J. Comp. Pathol. Therap. 59: 176-190.
Gao X, Wang H, Qin H, Xiao J. 2017. Influence of climate variations on the epidemiology of bluetongue
in sheep in Mainland China. Small Rumin. Res. 146: 23-27.
Gibbs EPJ, Greiner EC. 1994. The epidemiology of bluetongue. Comp. Immunol. Microbiol. Infect. Dis.
17: 207-220.
Gorman BM.1990. The bluetongue viruses. In Bluetongue viruses. (Springer), pp. 1-19.
Gubbins S, Carpenter S, Baylis M, Wood JL, Mellor PS. 2008. Assessing the risk of bluetongue to UK
livestock: uncertainty and sensitivity analyses of a temperature-dependent model for the basic
reproduction number. J. Royal Soc. Interface. 5: 363-371.
Hammami S. 2004. North Africa: a regional overview of bluetongue virus, vectors, surveillance and
unique features. Vet. Ital. 40: 43-46.
Han Z, Harty RN. 2004. The NS3 protein of bluetongue virus exhibits viroporin-like properties. J. Biol.
Chem. 279: 43092-43097.
Hemati B, Contreras V, Urien C, Bonneau M, Takamatsu H-H, Mertens PP, Bréard E, Sailleau C,
Zientara S, Schwartz-Cornil I. 2009. Bluetongue virus targets conventional dendritic cells in skin
lymph. J. Virol. 83: 8789-8799.
Herniman K, Boorman J, Taylor W. 1983. Bluetongue virus in a Nigerian dairy cattle herd: 1.
Serological studies and correlation of virus activity to vector population. Epidemiol. Infect. 90:
177-193.
Hoffmann B, Scheuch M, Höper D, Jungblut R, Holsteg M, Schirrmeier H, Eschbaumer M, Goller KV,
Wernike K, Fischer M. 2012. Novel orthobunyavirus in cattle, Europe, 2011. Emerg. Infect. Dis.
18: 469.
Hourrigan J, Klingsporn A. 1975. Epizootiology of bluetongue: the situation in the United States of
America. Aust. Vet. J. 51: 203-208.
Hutcheon D. 1881. Fever of epizootic catarrh. Rep Coll Vet Surg. 1880: 12-15.
Hutcheon D. 1902. Malarial catarrhal fever of sheep. Vet. Rec. 14: 629-633.
Jenckel M, Bréard E, Schulz C, Sailleau C, Viarouge C, Hoffmann B, Höper D, Beer M, Zientara S.
2015. Complete coding genome sequence of putative novel bluetongue virus serotype 27.
Genome Announc. 3: e00016-00015.
Jennings D, Mellor P. 1988. The vector potential of British Culicoides species for bluetongue virus. Vet.
Microbiol. 17: 1-10.
Joardar SN, Barkataki B, Halder A, Lodh C, Sarma D. 2013. Seroprevalence of bluetongue in north
eastern Indian state-Assam. Vet World. 6: 196-199.
Khair HO, Adam IA, Bushara SB, Eltom KH, Musa NO, Aradaib IE. 2014. Prevalence of bluetongue
virus antibodies and associated risk factors among cattle in East Darfur State, Western Sudan.
Irish Vet. J. 67: 4.
Khezri M, Azimi SM. 2013. Epidemiological investigation of bluetongue virus antibodies in sheep in
Iran. Vet. World. 6: 122-125.
Kirkland P. 2004. Bluetongue viruses, vectors and surveillance in Australia–the current situation and
unique features. Vet Ital. 40: 47-50.
Literature Cited
39
Kirkland P, Melville L, Hunt N, Williams C, Davis R. 2004. Excretion of bluetongue virus in cattle
semen: a feature of laboratory-adapted virus. Vet Ital. 40: 497-501.
Komarov A, Goldsmit L. 1951. A disease similar to bluetongue in cattle and sheep in Israel. Refuah vet.
8: 100.
Koumbati M, Mangana O, Nomikou K, Mellor P, Papadopoulos O. 1999. Duration of bluetongue
viraemia and serological responses in experimentally infected European breeds of sheep and
goats. Vet. Microbiol. 64: 277-285.
Lam SK, Burke D, Capeding MR, Chong CK, Coudeville L, Farrar J, Gubler D, Hadinegoro SR, Hanna
J, Lang J. 2011. Preparing for introduction of a dengue vaccine: recommendations from the 1st
Dengue v2V Asia-Pacific Meeting. Vaccine. 29: 9417-9422.
Legisa D, Gonzalez F, De Stefano G, Pereda A, Santos MD. 2013. Phylogenetic analysis of bluetongue
virus serotype 4 field isolates from Argentina. J. Gen. Virol. 94: 652-662.
Liu F. Li J M. Zeng F L. Zong Y. Leng X. Shi K. Diao NC. Li D. Li BY. Zhao Q. Du R. (2018).
Prevalence and risk factors of brucellosis, Chlamydiosis, and bluetongue among sika deer in Jilin
Province in China. Vector Borne Zoonotic Dis. 18(4): 226-230.
Lopez AC, Botija CS. 1958. Epizootie de fièvre catarrhale ovine en Espagne (blue tongue). Bull. Off. Int.
Epiz. 50: 65-93.
Lundervold M, Milner-Gulland E, O’callaghan C, Hamblin C. 2003. First evidence of bluetongue virus
in Kazakhstan. Vet. Microbiol. 92: 281-287.
Lysyk TJ, Danyk T. 2007. Effect of temperature on life history parameters of adult Culicoides sonorensis
(Diptera: Ceratopogonidae) in relation to geographic origin and vectorial capacity for bluetongue
virus. J. Med. Entomol. 44: 741-751.
Ma J-G, Zhang X-X, Zheng W-B, Xu Y-T, Zhu X-Q, Hu G-X, Zhou D-H. 2017. Seroprevalence and
Risk Factors of Bluetongue Virus Infection in Tibetan Sheep and Yaks in Tibetan Plateau, China.
BioMed Res. Int. 2017. (https://doi.org/10.1155/2017/5139703).
Maan NS, Maan S, Belaganahalli MN, Ostlund EN, Johnson DJ, Nomikou K, Mertens PP. 2012.
Identification and differentiation of the twenty six bluetongue virus serotypes by RT–PCR
amplification of the serotype-specific genome segment 2. PloS One. 7: e32601.
MacLachlan N. 2004. Bluetongue: pathogenesis and duration of viraemia. Vet Ital. 40: 462-467.
MacLachlan N, Conley A, Kennedy P. 2000. Bluetongue and equine viral arteritis viruses as models of
virus-induced fetal injury and abortion. Animal reproduction science. 60: 643-651.
Maclachlan N, Drew C, Darpel K, Worwa G. 2009. The pathology and pathogenesis of bluetongue. J.
Comp. Pathol. 141: 1-16.
Maclachlan NJ. 2011. Bluetongue: history, global epidemiology, and pathogenesis. Prev. Vet. Med. 102:
107-111.
MacLachlan NJ, Barratt-Boyes B, Brewer AW, Stott JL. 1991. Bluetongue virus infection of cattle. In:
Walton, TE, Osburn, BI (eds) Bluetongue, African horse sickness and related orbiviruses. CRC,
FL, pp 725–736.
Maclachlan NJ, Guthrie AJ. 2010. Re-emergence of bluetongue, African horse sickness, and other
orbivirus diseases. Vet. Res. 41: 35.
MacLachlan NJ, Osburn BI. 2006. Impact of bluetongue virus infection on the international movement
and trade of ruminants. J. Am. Vet. Med. Assoc. 228: 1346-1349.
Maclachlan NJ, Wilson WC, Crossley BM, Mayo CE, Jasperson DC, Breitmeyer RE, Whiteford AM.
2013. Novel serotype of bluetongue virus, western North America. Emerg. Infec. Dis. 19: 665.
Mahdavi S, Khedmati KAMBIZ, Pishraft Sabet L. 2006. Serologic evidence of bluetongue infection in
one-humped camels (Camelus dromedarius) in Kerman province, Iran. Iran. J. Vet. Res. 7(3): 85-
87.
Mahrt C, Osburn B. 1986. Experimental bluetongue virus infection of sheep; effect of vaccination:
pathologic, immunofluorescent, and ultrastructural studies. Am. J. Vet. Res. 47: 1198-1203.
Literature Cited
40
Malik AI, Ijaz M, Yaqub T, Shabir MZ, Avais M, Ghaffar A, Ali A, Farooqi SH, Mehmood K. 2019.
The first report on serotyping of bluetongue virus in small ruminants of Khyber Pakhtunkhwa
province, Pakistan. Tropical animal health and production. 51(4): 977-82.
Malik AI, Ijaz M, Yaqub T, Avais M, Shabbir MZ, Aslam HB, Aqib AI, Farooqi SH, Sohail T, Ghaffar
A, Ali A. 2018. Sero-epidemiology of bluetongue virus (BTV) infection in sheep and goats of
Khyber Pakhtunkhwa province of Pakistan. Acta trop. 182: 207-11.
Manso-Ribeiro J, Noronha F. 1958. Fievre catarrhale du mouton au Portugal (Blue Tongue). Bull. Off.
Int. Epizoot. 50: 46-64.
Martinelle L, Pozzo F, Sarradin P, Campe W, Leeuw I, Clercq K, Thys C, Thiry E, Saegerman C. 2016.
Experimental bluetongue virus superinfection in calves previously immunized with bluetongue
virus serotype 8. Vet. Res. 47: 73.
Mastrogiannis M, Axiotis I, Stoforos E. 1981. Study of the first outbreak of bluetonge disease in sheep in
Greece. Deltio tis Ellinikis Ktiniatrikis Etaireias (Greece).32 (2): 138-144.
Meiswinkel R, Baldet T, De Deken R, Takken W, Delécolle J-C, Mellor PS. 2008. The 2006 outbreak of
bluetongue in northern Europe—the entomological perspective. Prev. Vet. Med. 87: 55-63.
Mellor P, Baylis M, Mertens PPC. 2009. Introduction. In P. S. Mellor, M. Baylis, P. P. C. Mertens
(Eds.), Bluetongue (1 ed, pp. 1-6). London: Academic Press.
Mellor P 2004. Environmental influences on arbovirus infections and vectors. In symposia-society for
general microbiology. (Cambridge; Cambridge University Press; 1999), pp. 181-198.
Mellor P. 2000. Replication of arboviruses in insect vectors. J. Comp. Path. 123: 231-247.
Mellor P.1990. The replication of bluetongue virus in Culicoides vectors. In Bluetongue viruses.
(Springer), pp. 143-161.
Mellor P, Boorman J, Baylis M. 2000. Culicoides biting midges: their role as arbovirus vectors. Annual
review of entomology. 45: 307-340.
Mellor P, Wittmann E. 2002. Bluetongue virus in the Mediterranean Basin 1998–2001. Vet. J. 164: 20-
37.
Mellor PS, Carpenter S, Harrup L, Baylis M, Mertens PP. 2008. Bluetongue in Europe and the
Mediterranean Basin: history of occurrence prior to 2006. Prev. Vet. Med. 87: 4-20.
Menzies F, McCullough S, McKeown I, Forster J, Jess S, Batten C, Murchie A, Gloster J, Fallows J,
Pelgrim W. 2008. Evidence for transplacental and contact transmission of bluetongue virus in
cattle. Vet. Rec. 163: 203-209.
Mohammadi A, Tanzifi P, Nemati Y. 2012. Seroepidemiology of bluetongue disease and risk factors in
small ruminants of Shiraz suburb, Fars province, Iran. Trop Biomed. 29: 632-637.
Mortola E, Noad R, Roy P. 2004. Bluetongue virus outer capsid proteins are sufficient to trigger
apoptosis in mammalian cells. J. Virol. 78: 2875-2883.
Mozaffari AA, Khalili M, Sabahi S. 2014. High seroprevalence of bluetongue virus antibodies in goats in
southeast Iran. Asian Pac. J. Trop. Biomed. 4: S275-S278.
Mullens BA, Tabachnick WJ, Holbrook FR, Thompson LH. 1995. Effects of temperature on virogenesis
of bluetongue virus serotype 11 in Culicoides variipennis sonorensis. Med. Vet. Entomol. 9: 71-
76.
Murray M. 1991. The seasonal abundance of female biting-midges, Culicoides-brevitarsis Kieffer
(Diptera, Ceratopogonidae), in coastal south-eastern Australia. Aust. J. Zool. 39: 333-342.
Nason EL, Rothagel R, Mukherjee SK, Kar AK, Forzan M, Prasad BV, Roy P. 2004. Interactions
between the inner and outer capsids of bluetongue virus. J. Virol. 78: 8059-8067.
Nicoletti M, Murugan K, Del Serrone P. 2014. Current mosquito-borne disease emergencies in Italy and
climate changes. The neem opportunity. Trends Vector Res. Parasitol. 1: 2.
Nomikou K, Mangana-Vougiouka O, Panagiotatos D. 2004. Overview of bluetongue in Greece. Vet. Ital.
40: 108-115.
O'Connell L 2002. Entomological aspects of the transmission of arboviral diseases by Culidoides biting
midges. (Doctoral dissertation, University of Bristol).
Literature Cited
41
OIE. 2007. Saegerman C, Hubaux M, Urbain B, Lengelé L, Berkvens D. Regulatory aspects concerning
temporary authorisation of animal vaccination in case of an emergency situation: example of
bluetongue in Europe. Revue Scientifique et Technique de l’Office International des Epizooties.
2007; 26:395–414 [cited 2019 April 24]. Available
from http://www.oie.int/eng/publicat/rt/2602/PDF%2026-2/09-saegerman395-414.
OIE A. 2008. Manual of diagnostic tests and vaccines for terrestrial animals. Office International des
Epizooties, Paris, France. 1092-1106.
OIE. 2014. Mannual of diagnostic tests and vaccines for terrestrial animals. Chapter. 2.01.03.
Bluetongue. http://www.oie.int/manual-of-diagnostic-tests-and-vaccines-for-terrestrial-animals.
(Accessed 28 February 2018).
Oryan A, Amrabadi O, Mohagheghzadeh M. 2014. Seroprevalence of bluetongue in sheep and goats in
southern Iran with an overview of four decades of its epidemiological status in Iran. Comp. Clin.
Path. 23: 1515-1523.
Osburn BI. 1994. The impact of bluetongue virus on reproduction. Comp. Immunol. Microbiol. Infect.
Dis. 17: 189-196.
Oura CAL, El Harrak, M. 2011. Midge-transmitted bluetongue in domestic dogs. Epidemiol.
Infect. 139(9): 1396-1400.
Panda M, Mondal A, Joardar S. 2011. Seroprevalence of bluetongue virus in sheep, goat, and cattle in
West Bengal, India. Anim. Sci. Reporter. 5: 105-110.
Pandrangi A. 2012. Etiology, pathogenesis and future prospects for developing improved vaccines
against bluetongue virus: A Review. Afric. J. Environ. Sci. Technol. 7: 68-80.
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.
Purse BV, Mellor PS, Rogers DJ, Samuel AR, Mertens PP, Baylis M. 2005. Climate change and the
recent emergence of bluetongue in Europe. Nat. Rev. Microbiol. 3: 171-181.
Radostits O, Gay C, Hinchcliff K, Constable P 2007. Veterinary medicine, a text book of the diseases of
cattle, sheep, pig, goat, and horse, 10th edn. Saunders. (Elsevier, Spain).
Rao P, Hegde N, Reddy Y, Krishnajyothi Y, Reddy Y, Susmitha B, Gollapalli S, Putty K, Reddy G.
2016. Epidemiology of bluetongue in India. Transbound. Emerg. Dis. 63(2), e151-e164.
Ratinier M, Caporale M, Golder M, Franzoni G, Allan K, Nunes SF, Armezzani A, Bayoumy A, Rixon
F, Shaw A. 2011. Identification and characterization of a novel non-structural protein of
bluetongue virus. PLoS Pathog. 7: e1002477.
Raziq A, Younas M, Rehman Z. 2010. Continuing education article prospects of livestock production in
Balochistan. Vet J. 30: 181-186.
Rehman A, Jingdong L, Chandio AA, Hussain I. 2017. Livestock production and population census in
Pakistan: Determining their relationship with agricultural GDP using econometric
analysis. Inform Process Agric. 4(2): 168-177.
Rizzo JA, Chen J, Fang H, Ziganshin BA, Elefteriades JA. (2014). Statistical challenges in identifying
risk factors for aortic disease. Aorta, 2(2): 45-55.
Roy P. 2008. Functional mapping of bluetongue virus proteins and their interactions with host proteins
during virus replication. Cell Biochem. Biophys. 50: 143-157.
Roy P, Noad R.2006. Bluetongue virus assembly and morphogenesis. In Reoviruses: Entry, Assembly
and Morphogenesis. (Springer), pp. 87-116.
Ruiz-Fons F, Reyes-García ÁR, Alcaide V, Gortázar C. 2008. Spatial and temporal evolution of
bluetongue virus in wild ruminants, Spain. Emerg. Infec. Dis. 14(6): 951.
Sabaghan M, Borujeni MP, Shapouri MRSA, Rasooli A, Norouzi M, Samimi S, Mansouri S 2014.
Seroprevalence of Bluetongue in sheep in Kohgiluyeh and Boyer-Ahmad province, Iran. In
Veterinary Research Forum, Volume 5. (Faculty of Veterinary Medicine, Urmia University,
Urmia, Iran), p. 325.
Literature Cited
42
Shabbir MZ, Akram S, ul Hassan Z, Hanif K, Rabbani M, Muhammad J, Chaudhary MH, Abbas T,
Ghori MT, Rashid H, Jamil T. 2016. Evidence of Coxiella burnetii in Punjab province, Pakistan.
Acta trop. 163:61-9.
Saegerman C, Berkvens D, Mellor PS. 2008. Bluetongue epidemiology in the European Union. Emerg.
Infect. Dis. 14(4): 539.
Saegerman C, Bolkaerts B, Baricalla C, Raes M, Wiggers L, de Leeuw I, Vandenbussche F, Zimmer J-Y,
Haubruge E, Cassart D. 2011. The impact of naturally-occurring, trans-placental bluetongue virus
serotype-8 infection on reproductive performance in sheep. Vet. J. 187: 72-80.
Sailleau C, Bréard E, Viarouge C, Vitour D, Romey A, Garnier A, Fablet A, Lowenski S, Gorna K,
Caignard G. 2017. Re‐Emergence of bluetongue Virus serotype 8 in France, 2015. Transbound.
Emerg. Dis. 64: 998-1000.
Sairaju V, Susmitha B, Rao PP, Hegde NR, Meena K, Reddy YN. 2013. Type-specific seroprevalence of
bluetongue in Andhra Pradesh, India, during 2005–2009. Indian J. Virol. 24: 394-397.
Sánchez-Cordón P, Rodríguez-Sánchez B, Risalde M, Molina V, Pedrera M, Sánchez-Vizcaíno J,
Gómez-Villamandos J. 2010. Immunohistochemical detection of bluetongue virus in fixed tissue.
J. Comp. Pathol. 143: 20-28.
Santman-Berends IM, van Wuijckhuise L, Vellema P, Van Rijn P. 2010. Vertical transmission of
bluetongue virus serotype 8 virus in Dutch dairy herds in 2007. Vet. Microbiol. 141: 31-35.
Sarwar M. 1962. A note on bluetongue in sheep in West Pakistan. Pak J Anim Sci. 1: 1-2.
Savini G, Goffredo M, Monaco F, Di Gennaro A, Cafiero MA, Baldi L, De Santis P, Meiswinkel R,
Caporale V. 2005. Bluetongue virus isolations from midges belonging to the Obsoletus complex
(Culicoides, Diptera: Ceratopogonidae) in Italy. Vet. Rec. 157(5):133-9.
Schwartz-Cornil I, Mertens PP, Contreras V, Hemati B, Pascale F, Bréard E, Mellor PS, MacLachlan NJ,
Zientara S. 2008. Bluetongue virus: virology, pathogenesis and immunity. Vet. Res. 39: 1.
Sellers R, Taylor W. 1980. Epidemiology of bluetongue and the import and export of livestock, semen
and embryos [sheep, virus, domestic ruminants, Culicoides]. Bulletin de l'Office International des
Epizooties. 92:587–592.
Shafiq M, Minakshi P, Bhateja A, Ranjan K, Prasad G. 2013. Evidence of genetic reassortment between
Indian isolate of bluetongue virus serotype 21 (BTV-21) and bluetongue virus serotype 16 (BTV-
16). Virus Res. 173: 336-343.
Sperlova A, Zendulkova D. 2011. Bluetongue: a review. Vet Med-Czech. 56: 430-452.
Spreull J. 1905. Malarial catarrhal fever (bluetongue) of sheep in South Africa. J. Comp. Pathol. Ther.
18: 321-337.
Sreenivasulu D, Subba RM, Reddy Y, Gard G. 2003. Overview of bluetongue disease, viruses, vectors,
surveillance and unique features: the Indian sub-continent and adjacent regions. Vet. Ital. 40: 73-
77.
Susmitha B, Sudheer D, Rao PP, Uma M, Prasad G, Minakshi P, Hegde NR, Reddy YN. 2012. Evidence
of bluetongue virus serotype 21 (BTV-21) divergence. Virus Genes. 44: 466-469.
Sutton G, Grimes JM, Stuart DI, Roy P. 2007. Bluetongue virus VP4 is an RNA-capping assembly line.
Nat. Struct. Mol. Biol. 14: 449.
Tabachnick WJ. 1996. Culicoides variipennis and bluetongue-virus epidemiology in the United States.
Annu. Rev. Entomol. 41: 23-43.
Takamatsu H, Mellor P, Mertens P, Kirkham P, Burroughs J, Parkhouse R. 2003. A possible
overwintering mechanism for bluetongue virus in the absence of the insect vectorFN1. J. Gen.
Virol. 84: 227-235.
Taylor W, Mellor P. 1994. Distribution of bluetongue virus in Turkey, 1978–81. Epidemiol. Infec. 112:
623-633.
Tollersrud T 2009. Bluetongue-Europe (06): Norway, First Cases Detected. Archive No. 20090221.0729.
(http://www.promedmail.org [accesed 02.04.2009]).
Torina A, Caracappa S, Mellor P, Baylis M, Purse B. 2004. Spatial distribution of bluetongue virus and
its Culicoides vectors in Sicily. Med. Vet. Entomol. 18: 81-89.
Literature Cited
43
Van der Walt N. 1980. A haemagglutination and haemagglutination inhibition test for bluetongue virus.
Onderstepoort J. Vet. Res. 47: 113-117.
van Regenmortel MH, Fauquet C, Bishop D, Carstens E, Estes M, Lemon S, Maniloff J, Mayo M,
McGeoch D, Pringle C. 2000. Virus taxonomy: classification and nomenclature of viruses.
Seventh report of the International Committee on Taxonomy of Viruses, (Academic Press).
Vandenbussche F, Vanbinst T, Verheyden B, Van Dessel W, Demeestere L, Houdart P, Bertels G, Praet
N, Berkvens D, Mintiens K. 2008. Evaluation of antibody-ELISA and real-time RT-PCR for the
diagnosis and profiling of bluetongue virus serotype 8 during the epidemic in Belgium in 2006.
Vet. Microbiol. 129: 15-27.
Vassalos M. 1980. Cas de fièvre catarrhale du mouton dans l'île de Lesbos (Grèce), (Office International
des Epizooties). 92(7/8): 547-555.
Verwoerd D, Erasmus B. 2004. Bluetongue. Infectious diseases of livestock. 2: 1201-1220.
Wilbur LA, Evermann JF, Levings RL, Stoll IR, Starling DE, Spillers CA, Gustafson GA, McKeirnan
AJ. 1994. Abortion and death in pregnant bitches associated with a canine vaccine contaminated
with bluetongue virus. J Am Vet Med Assoc. 204(11): 1762.
Wilson WC, Mecham JO, Schmidtmann E, Sanchez CJ, Herrero M, Lager I. 2009. Current status of
bluetongue virus in the Americas. In P. S. Mellor, M. Baylis, P. P. C. Mertens (Eds.), Bluetongue
(1 ed., pp. 197-221). London: Academic Press.
Wilson A, Carpenter S, Gloster J, Mellor P. 2007. Re-emergence of bluetongue in northern Europe in
2007. Vet. Record. 161: 487.
Wilson A, Mellor P. 2008. Bluetongue in Europe: vectors, epidemiology and climate change. Parasitol.
Res. 103: 69-77.
Wilson AJ, Mellor PS. 2009. Bluetongue in Europe: past, present and future. Philos. Trans. R. Soc.
Lond. B. Biol. Sci. 364: 2669-2681.
Wittmann E, Mellor P, Baylis M. 2002. Effect of temperature on the transmission of orbiviruses by the
biting midge, Culicoides sonorensis. Med. Vet. Entomol. 16: 147-156.
Wouda W, Roumen MP, Peperkamp NH, Vos JH, Van Garderen E, Muskens J. 2008. Hydranencephaly
in calves following the bluetongue serotype 8 epidemic in the Netherlands. Vet. Record. 62(13):
422-3.
Yavari M, Gharekhani J, Mohammad ZA. 2018. Bluetongue virus seropositivity and some risk factors
affecting bluetongue virus infection in sheep flocks. Comp. Clin. Path. 1-6.
Yilmaz V, Yildirim Y, Otlu S. 2012. The seroprevalance of Bluetongue virus infectıon in cattle in the
Kars Dıstrıct of Turkey. Isr. J. Vet. Med. 67: 4.
Yonguç A, Taylor W, Csontos L, Worrall E. 1982. Bluetongue in western Turkey. Vet. Record. 111:
144-146.
Yousef MR, Al-Eesa AA, Al-Blowi MH. 2012. High seroprevalence of bluetongue virus antibodies in
Sheep, Goats, Cattle and Camel in different districts of Saudi Arabia. Vet. World. 5(7).
Zahur A, Ullah A, Hussain M, Irshad H, Hameed A, Jahangir M, Farooq M. 2011. Sero-epidemiology of
peste des petits ruminants (PPR) in Pakistan. Prev. Vet. Med. 102: 87-92.
Zulu GB, Venter EH. 2014. Evaluation of cross-protection of bluetongue virus serotype 4 with other
serotypes in sheep. J. S. Afr. Vet. Assoc. 85: 01-02.
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