Full Length Research Paper
of competitive enzyme-linked immunosorbent assay and one
step RT-PCR tests for
Bluetongue virus in south west of Iran
Shahin Nejat2, Negar Souod3, Monochehr
Momeni4 and Sohrab Safari4
Microbiology, Faculty of Veterinary Medicine, Islamic Azad
University, Shahrekord Branch, Shahrekord-Iran.
of Clinical Sciences, Faculty of Veterinary Medicine,
Islamic Azad University, Shahrekord Branch, Shahrekord-Iran.
3Master of Science of
Microbiology, Member of young researchers club, Islamic Azad
University, Jahrom Branch, Jahrom-Iran.
Center of Biotechnology,
Islamic Azad University- Shahrekord Branch- Shahrekord-
*Corresponding author. E-mail:
Competitive enzyme-linked immunosorbent assay;
reverse transcriptas- polymerase chain reaction; BTV,
Accepted 9 June, 2011
Bluetongue is a noncontagious, arthropod-borne viral disease of
both domestic and wild ruminants. Bluetongue virus (BTV) is the
species of the genus Orbivirus within the family Reoviridae. BTV
is endemic in some areas with cattle and wild ruminants serving
as reservoirs for the virus. Clinical symptoms are often seen in
sheep. There are several methods for
detection of Bluetongue virus,
among them the molecular technique like RT-PCR is considered as
the most sensitive and reliable one. The aim of this study was
to comprise competitive enzyme-linked immunosorbent assay
(C-ELISA) with one step RT-PCR test for
detection of BTV in sheep.
A total of 770 blood samples
obtained from sheep (265 serum positive samples and 505 serum
negative samples in C-ELISA). According to
data, out of
265 serum positive samples in ELISA test, 234 were positive in
RT-PCR assay whereas all serum negative samples were negative in
RT-PCR experiment. According to the results, the PCR assay
more sensitive and reliable than ELISA technique for
Bluetongue virus, C-ELISA, RT- PCR, Sheep, Iran.
Bluetongue virus (BTV)
belongs to the genus Orbivirus of the family Reoviridae. The
virus particle is nonenveloped and icosahedral in shape and
consists of a double-layer-ed protein coat. Nowadays, at least
24 serotypes of this virus have been identified (Davies et al.,
1992; Lee et al., 2010; Roy, 1992; Schwartz-Cornil et al.,
2008). BTV has been found where suitable vectors are present (Tabachnick,
1996). Bluetongue disease is on the multispecies list of
notifiable diseases by the office of international epizootics (OIE)
because of its substantial economic impact and potential for
rapid spread. Infection with BTV can cause serious hemorrhagic
disease with high mortality rates in sheep and deer (Howerth et
al., 1988; Osburn, 1994; Rodriguez-Sanchez et al., 2010). BTV is
a small icosahedral virus with a ten-segmented, double stranded
RNA (dsRNA) genome. Each of the ten segments codes for at least
one of ten distinct viral pro-teins, seven of which are
constructed components of the virus particle and three of which
are nonstructural. The inner capsid of BTV is composed of five
poly-peptides: three minor proteins (VP1, VP4 and VP6) and two
major proteins (VP3 and VP7) (Roy, 1992). The outer capsid is
composed of two major viral proteins, VP2 and VP5, which
determine the antigenic variability of the BTV. The core
proteins and nonstructural proteins NS1, NS2, NS3/3A
(encoded by seg. 5, 8 and 10, respectively) are thought to be
relatively conserved and are antigenically cross-reactive
between different strains and serotypes of BTV. However,
cross-hybridization and sequencing studies have shown that these
genome segments can vary in manner that reflects the
geographical origin of the virus strains (Pritchard et al.,
2004). Efficiency of PCR in diagnosis of BTV in several parts of
the world, verified this technique as a rapid, reliable and
sensitive diagnosis methods. The highly specific and sensitive
nature of RT-PCR based assay makes it ideal for rapid detection
of BTV genome segments in clinical samples without requirement
to virus isolation (Anthony et al., 2007). BTV infection
previously has been determined in several parts of Middle East
in Iran (Shoorijeh et al., 2010). The history of BTV detection
in this part of the world is more than 50 years. Serological
evidence indicates that the virus has been present in Iran, at
least since 1974 (Afshar and Keyvanfar, 1974). The purpose of
this study was to use a valid RT-PCR assay to detect any BTV
isolates in South West of Iran, from sheep blood specimen.
Materials And Methods
Blood and serum samples were collected randomly from male and
female sheep of 3 different geographic regions in Iran between
May 2009 and October 2010.
Sera collected from 770
sheep of 120 flocks in 10 different regions of Chaharmahal va
Bakhtiari, Khozestan and Isfahan provinces (166 samples from
male sheep and 604 samples from female sheep) were screened for
group specific antibodies to BTV, using a commercial competitive
enzyme linked immunosorbent assay (C-ELISA; ID Screen1, ID VET
Co., Montpellier, France) and an ELISA micro plate reader (model
Stat Fax 2100, Awareness Technology Inc., USA). The sera were
collected from 284 aborted and 320 non-aborted sheep during 2009
to 2010. The clinical signs of BTV disease were not described in
any of these animals.
Extraction of viral RNA
The dsRNA extractions were
carried out using the RNeasy® mini
kit (Qiagen) from whole blood samples according to the
Two pairs of primers: (TF-1: 5'-GTTAAAAATCTATAGAG-3'; TR-1:
5'-GTAAGTGTAATCTAAGAG-3') and (TF-2:5'- GTTAAAAATC
TATAGAGATG-3'; TR-2:5'- GTAAGTGTAATCTAAGAGA-3') which amplify
full length of BTV serogroup S7 gene (1156 bp), were used. For
nested PCR, internal primers (IF-1: 5'-ACAACTGATGCTGCG AATGA-3';
IR-1: 5'-AACCCACACCCGTGCTAAGTGG-3' was applied (Bréard
et al., 2003; Wade-Evans et al., 1990). The
primer set amplified internal part of S7 segment in length of
769 bp. All oligonucleotide primers were synthesized
commercially (Cinnagen Co., Iran).
RNA extracted from reference strain BTV1 (RSAvvvv/01, which was
received from Institution of Animal Health, Pirbright, UK) was
used as positive control and sterile distilled water was used as
negative control in RT-PCR assay.
One step RT-PCR
The one step RT-PCR kit (QIAGEN®
OneStep RT-PCR Kit) was used for
detection of S7 BTV gene in blood samples. The master mix was
made as follows: 10 μl of 5X Qiagen RT-PCR buffer, 2 μl dNTPs
mixture (0.2 mM each), 0.5 μl (20 pmol) of each of four primers
(TF-1, TR-1, TF-2, TR-2), 2 μl Qiagen enzyme mix and 28 μl of
RNase free water. Then 6 μl of denatured RNA
added to master mix. In RT-PCR,
the RNA was initially reverse-transcribed at 45°C for 30 min;
then, followed by a step at 95°C for 15 min to simultaneous
activation of DNA polymerase and inactivation of reverse
transcriptase. Forty amplification cycles were performed at 95°C
for 1 min, 45°C for 1 min and 72°C for 2 min. The PCR cycles was
terminated by final extension step at 72°C for 10 min (Anthony
et al., 2004).
PCR products of first amplification (RT-PCR)
were used as template
in nested PCR. The mixture of master mix contained, 100 μM dNTPs,
1.5 mM MgCl2, 1 µM of each primer (IF-1 and IR-2), 1
U of Taq DNA polymerase and 35 μl of RNase free water, at the
end, 5 μl of template was added to the reaction in a final
volume of 50 μl. The Mastercycler gradient PCR machine (Eppendorf,
Mastercycler® 5330, Eppendorf-Netheler-Hinz GmbH,
Hamburg, Germany) was set to amplify the nested fragment as
follow: first step was at 95°C for 1 min, then 30 cycles were
performed at 95°C for 1 min, 59°C for 1 min and 72°C for 1 min.
The reaction was stopped by extension at 72°C for 10 min
(Anthony et al., 2004).
Analysis of PCR products
final PCR products were run on a 1%
agarose gel containing ethidium bromide in 1X TBE buffer along
with 1 Kb DNA ladder (Fermentas). The gels were analysed using
gel documentation system (Uvidoc imaging system).
Results and Discussion
Out of 770 samples, 265 specimens had
antiviral anti-bodies with C-ELISA test.
All the samples were tested with PCR
assay too. The results are shown in
Table 1. According to these results, the
769 bp segment relates to bluetongue
positive samples in Nested-PCR assay.
In this study, PCR
technique was employed as a quick,
sensitive and specific method for
diagnosis of Bluetongue virus from the
cases suspected to the disease. This
research is the first report on BTV
identification based on molecular method
in South West of Iran. It is worth
mentioning that the situation of
diagnosis of this virus in neighboring
countries and the Middle-East
Turkey and Occupied Palestine) is not
better than our country. In such
countries as Saudi Arabia, Syria, Yemen,
Oman and Pakistan,
presence of the virus has been
documented only relying on serological
tests (Akhtar et al., 1997; Lundervold
et al., 2003; Shoorijeh et al., 2010).
According to recent
studies, there is an evidence of
occurrence of BT disease in tropical and
subtropical countries (such as Iran). In
such areas generally, the disease
appears subclinically and does not
attract attention. In such
presence of the virus often confirms via
serological evidences. It should be
mentioned that in such foci, in spite of
unrevealed disease and manifestations,
sometimes sudden inciden-ce of acute
forms of the disease sustain a loss (Basak
et al., 1997; Nikolakaki et al., 2005).
identification of BTV in suspected
cattle and sheep based on clinical
manifestations was performed. However,
there are some limitations and problems.
it should be considered that clinical
expression of BTV regarding strain and
virus intensity, cattle race and
environmental condition varies from
peracute to subclinical. Secondly, symptoms of
disease in sheep can be mistaken with
those of other viral much diseases and
even some of the non-viral diseases.
For the first time,
Afshar and Kayvanfar (1974) reported the
existence of antibody against this virus
via AGID test in Iran. In recent years,
based on serological methods, infection
with this virus has been proved in
various pro-vinces of Iran (based on
veterinary organization reports). It
should be mentioned that some suspected
cases have been reported by
veterinarians but unfortunately because
of the lack of reliable and quick
laboratory techniques, we could not
accept such cases.
During the BTV
epidemics in Europe in 2008,
Williamson et al. (2008)
considered clinical signs for diag-nosis
of the disease. The results showed low
specificity of this method. These
researchers believe that some-times
clinical signs of BTV in sheep are
mistaken with those of such diseases as
FMD, PPR, contagious ecthyma and
haemonchosis (Elbers et al., 2008; Tan
et al., 2001).
many methods have been employed in order
to detect BTV. The most important of
them are isolation of virus via
embryonic egg or cell culture,
serological and molecular methods.
laboratory diagnostic methods, molecular
techniques enjoy the highest level of
sensitivity and specificity for
diagnosis of arboviruses including BTV.
This method is so sensitive that even 6
molecules of the virus genome in blood
can be traced (Anthony et al., 2004).
Biteau- Coroller et
al. (2006) and Afshar, (1994) in their
studies introduced the PCR method as a
'golden test' for BTV diagnosis compared
with other procedures. In sheep, being
infected by vector anopheles, PCR
readings from 5 days to 38 up to 54 days
the onset of infection becomes positive.
This period in infected cattle is up to
100 days following onset of infection (Afshar,
1994; Biteau-Coroller et al., 2006;
Koumbati et al., 1999)
In a study
conducted in 2005 by Bréard et al.
(2005) from the 46 feverish sheep blood
and suspected to BTV,
only one cast of virus isolation via
embryonated chicken egg (ECE) injection
was possible, while a great number of
them were PCR-positive (Bréard et al.,
2003). Also, through a study conducted
by MacLachlan et al. (1994), the
sensitivity of various diagnostic
methods like injection to ECE and PCR
and feeding colicoides mosquitoes were
compared. Their study revealed that it
is possible to diagnose the virus in
cattle blood, with infecting mosquito
method for about 2 weeks, injecting to
ECE about 2 to 8 weeks and in PCR
procedure between 16 to 20 weeks
following the onset of infection.
Steinrigl et al. (2010) from Australia
injected an in-activated vaccine of the
BTV 8 serotype to the understudy sheep
and showed that RT-PCR is the best
detection of trace amount of BTV in
Yin et al. (2010) used RT-PCR method for
detection of NS1 gene. They succeeded in
tracing all 24 serotypes of BTV in
China. Vandenbussche et al. (2010)
employed multiplex RT-PCR as a routine
method for BTV identification in their
study. They also used this method in
order to find other genes simultaneously
in BTV and showed that this method
enjoys extraordinary sensitivity for
identification of some different genes
So far, various methods of RT-PCR have
been em-ployed for BTV diagnosis.
Usually, the genes used to identify BTV
serogroups are retained in all
serotypes. These genes produce such
core-proteins as (VP7), (VP1) L1, (VP3)
L3 and (NSI) M6 and/or nonstructural
proteins such as NS10 (NS3/NS3A) (Bréard
et al., 2004; Dangler et al., 1990;
Hwang et al., 1992; MacLachlan et al.,
1994; Orrù et al., 2006; Parsonson and
Among these genes, S7 segment is the
best candidate for
identification of this virus because it
is a stable protein in all BTV serotypes
and topotypes and it’s highly sensitive
in PCR (Anthony et al., 2007; Billinis
et al., 2001; Zientara et al., 2004).
Therefore in this study, we used S7
segment as a marker for
identification of infected samples.
Moreover, we do not have enough
information about the prevalent strain
of BTV in Iran, so we used RT-PCR-duplex:
two supplementary primer pairs at 3' and
5', for performing the reaction
simulta-neously. In addition, recently,
it has been identified that noncoding
ends of the S7 segment are similar in
the 7, 10 and 19 serotypes and also vary
a little in other serotypes (Anthony et
al., 2007, 2004). Therefore, by using
two primer pairs, all serotypes of BTV,
apart from their serotypes would be
we applied one step RT-PCR too. The
advantage of this procedure is
decreasing the test time in comparison
with two-step RT-PCR method and to
decrease the number of false positive
cases by omitting additional
manipulations at cDNA generation and
reducing number of pipette used.
study, we gathered suspected sheep blood.
we applied RT-PCR then we used nested
PCR assay to verify the product of the
primary PCR and to in-crease sensitivity
of virus identification in clinical
samples. As the matter of fact, this
method is a simple, fast and sensitive
assay for confirming of PCR products.
The sensitivity of nested PCR is at the
rate of 10 copies of the target gene in
the clinical samples (equal to
sensitivity of real time PCR) since in
this method, we use four specific
primers (Biteau-Coroller et al., 2006).
According to many studies, the second
PCR (nested PCR) in comparison with the
first PCR is 10 to 100 times more
sensitive. So, when the amount of RNA is
less than 100 fg, the nested PCR method
is so valuable. It was shown that this
method is able to identify even 0.1 fg
of the BTV genome (= 5 molecules of the
BTV genome) in the cell culture (Aradaib
et al., 2005, 2003, 1998).
Eaton and White (2004) also introduced
nested PCR of the S7 gene as a sensitive
and appropriate method for
identification of BTV in suspicious
cases. This point on employing RT- PCR
in BTV diagnosis is that most of the
researchers have designed and optimized
this method based on laboratory
specimens obtained from cell cul-tures
inoculated with virus and/or
experimental samples supplied through
injecting the acute virus to sheep.
However, they have not evaluated
efficiency of their procedure in
clinical cases (Aradaib et al., 2003).
In this study, the results are in
agreement with other investigators;
so the first PCR products were unclear
or weak. This
probably due to the low level of virus,
quality of samples and/or low quantity
or quality of the extracted RNA. Anyway,
by using nested PCR assay, we obtained
specific and clear bands and the earlier
mentioned problem were solved.
Meanwhile, some negative specimens in
the first PCR assay were positivein
this procedure, so that it is obvious
that the first bands were weak.
As it was mentioned earlier, one of the
exclusive applications of PCR, which has
made it superior to other diagnostic
techniques, is the possibility of
molecular epidemiologic surveys, study
of genetic modifications and
determination of origin of virus
transfer to other foci (Gould
and Pritchard, 1990;
2005; Pritchard et al., 1995;
Another important method of BTV
identification is the competitive ELISA.
In this method, anti-VP7 antibodies are
traced. Nowadays, this method due to
high specificity, rapid action and ease
of application is considered as a
standard method such that in all the
international reference laboratories in
addition to molecular methods, this
procedure is also used for evaluating
In fact, the competitive ELISA test is
considered as a powerful tool in
seroepidemilogic studies of BTV (Batten
et al., 2008).
The only weal point of C-ELISA is its
limited sensitivity which is due to
delayed antibody formation. This time,
it is about 7 to 28 days post-infection.
During this period, the test sensitivity
is low and the test will show false
negative responses. To remove this
defect, employment of PCR is
recommended. Because of this reason,
many of the investigators recommend that
in order to study the status of disease
in endemic foci and/or in foci where the
status of disease is not clear, both
procedures could be employed
et al., 2006; Singer et al., 1998).
Those investigators, who have used
molecular and serologic method in
identifying BTV, unanimously believe
that PCR and ELISA results complement
each other and that one of them being
positive necessarily does not influence
the result of the other.
Bréard et al. (2005) and
Singer et al. (1998)
in their study fields found that PCR and
ELISA results have no direct relations
to each other.
In this study, a number of samples were
positive both in PCR and ELISA tests and
this is in agreement with other
investigator’s reports. To explain this
matter, we can say the positivity of the
cattle’s serum does not have any
incompatibility with the present of
virus in its blood. This can be due to
the reinfection of serum-positive cattle
with heterogenous serotype of the virus
et al., 2001; Bréard et al., 2005;
Singer et al., 1998)
On the other hand, it should be noted
that in C-ELISA, anti-VP7 antibodies are
traced. As it was mentioned earlier,
this type of antibody does not have any
role in neutralizing the BTV in
vertebrate hosts. Thus, in spite of
formation of such antibodies, virus can
continue its activity.
In this study, a number of samples were
positive only in ELISA or in PCR assays.
The reason for inconsistency of PCR
results with those of ELISA goes back to
the difference between the viremia times
(incubation period) and sero-conversion.
Viremia in sheep starts between 4 and 8
onset of infection and continues till
the next 54 days. With seroconversion,
however, on average starts 14 days post
induced infection but con-tinues for
long periods (Biteau-Coroller
et al., 2006; Maan et al., 2004).
In fact, PCR is very valuable in new
infected cattle when no serum response
has been produced.
The reason for increased serum-positive
cases to PCR is that following sero-conversion
unlimitedly (occasionally for years)
blood antibody level remains high.
Clavijo et al. (2000) and Dadhich
believes that one of the reasons for
long-term serologic response in BTV
infection as the continuous stimulation
of the cattle's immunity system is due
attachment of virus to the blood cell
membranes and remaining with them up to
the end of the cell's life.
Definitively, such cattle are also PCR-positive
and as soon as infected red blood cells
are removed from their blood, they will
become negative from this point of view.
From the serological point of view
however, they will remain positive thus,
accordingly. Therefore, one of the
suitable tools for BTV diagnosis in foci
where the disease is found in quiescent
(latent) state is to use the serum ELISA
(Biteau-Coroller et al.,
In order to know the BTV status in South
West of Iran, first it is necessary to
isolate circulating viruses in foci.
we have to analyze at least 2 to 3
segments of their genome. This is
important since genotyping of a gene
only reveals part of genomic
characteristics of the virus. Regarding
independence of various segments of BTV
in transferring to subsequent
generations, it is necessary to study a
greater number of them. Accor-dingly,
which increasing precision of topotyping,
the possibility of study of probable
rearrangements between the viruses
prevalent in the country will be
competitive enzyme linked immunosorbent
(C-ELISA) with RT-PCR for
The authors would like to thank
Dr. E. Tajbakhsh, Dr. S. Mollamohseni
and Dr. F. Zeiai
at the Biotechnology Research Center of
the Islamic Azad University of
Shahrekord for their sincere technical
and clinical support.
Afshar A (1994).
Bluetongue: laboratory diagnosis.
Comp. Immunol. Microbiol. . Infect. Dis.
Kayvanfar H (1974). Occurrence of precipitating antibodies to
bluetongue virus in sera of farm animals in Iran. Vet. Rec. 94:
(1997). Bluetongue virus seropositivity in sheep flocks in North
West Frontier Province, Pakistan.
Prevent. Vet. Med.
Anthony S, Maan
S, Samuel AR, Mellor PS, Mertens PP (2004).
Differential diagnosis of bluetongue virus using a reverse
transcriptase-polymerase chain reaction for genome segment 7.
(1998). A nested PCR for detection of North American isolates of
bluetongue virus based on NS1 genome sequence analysis of BTV-17.
(2003). A multiplex PCR for simultaneous detection and
differentiation of North American serotypes of bluetongue and
epizootic hemorrhagic disease viruses.
Comp. Immunol. Microbiol. Infect. Dis.
(1997). Structures of orbivirus VP7: implications for the role of
this protein in the viral life cycle.
(2004). Bluetongue: an overview of recent trends in diagnostics.
Davies FG, Mungai
JN, Pini A (1992). A new bluetongue virus serotype isolated in
(2004). Developing new orbivirus diagnostic platforms.
van der Spek AN,
van Rijn PA
(2008). Performance of clinical signs to detect bluetongue virus
serotype 8 outbreaks in cattle and sheep during the 2006-epidemic in
(1990). Relationships amongst bluetongue viruses revealed by
comparisons of capsid and outer coat protein nucleotide sequences.
Greene CE, Prestwood AK (1988). Experimentally induced bluetongue
virus infection in white-tailed deer: coagulation, clinical
pathologic, and gross pathological changes. Am. J. Vet. Res. 49:
(1992). Comparative sequence analyses of the cognate structural
protein VP6 genes of five US bluetongue viruses.
(1999). Duration of bluetongue viraemia and serological responses in
experimentally infected European breeds of sheep and goats.
(2010). Subclinical bluetongue virus infection in domestic ruminants
(2004). Development of reverse transcriptase-polymerase chain
reaction-based assays and sequencing for typing European strains of
bluetongue virus and differential diagnosis of field and vaccine
(1994). Detection of bluetongue virus in the blood of inoculated
calves: comparison of virus isolation, PCR assay, and in vitro
feeding of Culicoides variipennis.
(2005). Persistence of bluetongue virus serotype 2 (BTV-2) in the
southeast United States.
(2005). Molecular analysis of the NS3/NS3A gene of Bluetongue virus
isolates from the 1979 and 1998-2001 epizootics in Greece and their
segregation into two distinct groups.
De Santis P
(2006). Rapid detection and quantitation of Bluetongue virus (BTV)
using a Molecular Beacon fluorescent probe assay. J. Virol. Methods,
Osburn B I
(1994). Bluetongue virus, The Veterinary Clinics of North America.
Food Anim. Prac. 10: 547-560.
(1995). Retrospective diagnosis of bluetongue virus in stored frozen
and fixed tissue samples using PCR.
(1995). Complete nucleotide sequence of RNA segment 3 of bluetongue
virus serotype 2 (Ona-A), Phylogenetic analyses reveal the probable
origin and relationship with other orbiviruses.
Sendow I, Lunt R, Hassan SH, Kattenbelt J, Gould AR, Daniels PW,
Eaton BT (2004). Genetic diversity of bluetongue viruses in south
east Asia. Virus Res. 101: 193-201.
(1988). Genetic relationships of bluetongue virus serotypes isolated
from different parts of the world.
Roy P (1992). Bluetongue virus
proteins. J. Gen. Virol. 73: 3051-3064.
I, Mertens PPC, Contreras V, Hemati B, Pascale F, Breard E, Mellor
PS, MacLachlan NJ, Zientara S (2008). Bluetongue virus: virology,
pathogenesis and immunity. Vet. Res. 39: 46-61.
Ramin AG, Maclachlan NJ, Osburn BI, Tamadon A, Behzadi MA, Mahdavi
M, Araskhani A, Samani D, Rezajou N, Amin Pour A (2010). High
seroprevalence of bluetongue virus infection in sheep flocks in West
Comp. Immunol. Microbiol. Infect. Dis.
(2010). Bluetongue virus RNA detection by RT-qPCR in blood samples
of sheep vaccinated with a commercially available inactivated BTV-8
vaccine. Vaccine, 28: 5573-5581.
(1996). Culicoides variipennis and Bluetongue-virus epidemiology in
the United States. Ann. Rev. Entomol. 41: 23-43.
(2001). RGD tripeptide of bluetongue virus VP7 protein is
responsible for core attachment to Culicoides cells.
De Clercq K
(2010). Simultaneous detection of bluetongue virus RNA, internal
control GAPDH mRNA, and external control synthetic RNA by multiplex
Methods Mol. Biol.
Mertens PPC, Bostock CJ (1990). Development of the polymerase chain
reaction for the detection of bluetongue virus in tissue samples. J.
Virol. Methods, 30: 15-24.
Williamson S, Woodger N, Darpel
K (2008). Differential diagnosis of bluetongue in cattle and sheep.
(2010). Detection and quantitation of bluetongue virus serotypes by
a TaqMan probe-based real-time RT-PCR and differentiation from
epizootic hemorrhagic disease virus. J. Virol. Methods, 168:
(2004). Bluetongue diagnosis by reverse transcriptase-polymerase