Download Bluetongue virus diagnosis of clinical cases by a duplex reverse

Journal of Virological Methods 98 (2001) 77 – 89
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Bluetongue virus diagnosis of clinical cases by a duplex
reverse transcription-PCR: a comparison with conventional
methods
Charalambos Billinis a, Maria Koumbati a, Vassiliki Spyrou a,
Kyriaki Nomikou b, Olga Mangana b, Christos A. Panagiotidis c,
Orestis Papadopoulos a,*
a
Laboratory of Microbiology and Infectious Diseases, Faculty of Veterinary Medicine, Aristotle Uni6ersity,
GR-54006 Thessaloniki, Greece
b
Institute of Infectious and Parasitic Diseases, GR-15341 Agia Paraske6i, Attiki, Greece
c
Department of Pharmacy, Aristotle Uni6ersity, GR-54006 Thessaloniki, Greece
Received 25 January 2001; received in revised form 21 June 2001; accepted 22 June 2001
Abstract
A duplex reverse transcription polymerase chain reaction (RT-PCR) assay for the detection of bluetongue virus
(BTV) in clinical samples was developed. This assay, which detects the highly conserved S10 region of BTV, was
assessed for sensitivity and application as a rapid and dependable diagnostic tool by comparison with standard assays
of virus detection, such as virus isolation in embryonated chicken eggs and cell culture. Simultaneous detection of
BTV and host b-actin RNAs minimizes the possibility of false negative results. The sensitivity of the assay was found
to be equal to five cell culture infectious dose (CCID50) units and its specificity was confirmed as no RT-PCR product
was detected with RNAs from two closely related orbiviruses, i.e. epizootic haemorrhagic disease virus (serotypes 1,
2 and 318) and African horse sickness virus, serotype 9, or RNAs from uninfected BHK-21 cells and blood samples
from uninfected sheep or goats. In this study, 36 blood samples from naturally infected mixed flocks of sheep and
goats were examined. Seventeen animals were identified as BTV-positive by RT-PCR, whereas only 13 were found
positive by virus isolation in embryonated chicken eggs and nine by cell culture assays. These results indicate that the
duplex RT-PCR could be a useful technique for monitoring BTV infection in the field. © 2001 Elsevier Science B.V.
All rights reserved.
Keywords: Bluetongue virus; Reverse transcription duplex polymerase chain reaction; Bluetongue diagnosis; Clinical cases
1. Introduction
* Corresponding author. Tel.: + 30-31-999951; fax: +3031-999959.
E-mail address: [email protected] (O. Papadopoulos).
Bluetongue is an arthropod-transmitted disease
of wild and domestic ruminants. It is a virusborne disease that is caused by the bluetongue
0166-0934/01/$ - see front matter © 2001 Elsevier Science B.V. All rights reserved.
PII: S 0 1 6 6 - 0 9 3 4 ( 0 1 ) 0 0 3 6 0 - 3
78
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
virus (BTV), which is the prototype species of the
genus Orbi6irus in the family Reo6iridae
(MacLachlan, 1994). There are at least 24
serotypes of BTV worldwide (Davies et al., 1992),
three of which (4, 9 and 16) have been isolated in
Greece (K. Nomikou et al., unpublished data).
BTV infection occurs throughout temperate and
tropical regions of the world, and infection is
dependent on the presence of competent vector
midges of Culicoides spp. (Gibbs and Greiner,
1994; Mellor, 1990). These insects become infected persistently with BTV and are infectious to
ruminants on which they feed after an extrinsic
incubation period of 10– 14 days (Tabachnick,
1996).
Bluetongue is an International Office of Epizootics List A disease (Alexander et al., 1994;
OIE, 2000) described as an economically devastating affliction of sheep (Alexander et al., 1994).
Indeed, the vast economic effects of BTV infection in many parts of the world, due to the high
morbidity and mortality rates in the affected animals have been established (Abu Elzein et al.,
1992; Eisa et al., 1980; Hourrigan and
Klingsporn, 1975; Quist et al., 1997; Barnard et
al., 1998; Erasmus, 1975; Gee, 1975; Geering,
1975; Kvasnicka, 1985; Metcalf et al., 1980; Mulhern et al., 1985; Mulhern, 1985; Murphy et al.,
1985; Tabachnick, 1996). Early detection of infected animals could reduce drastically the consequences of the disease by reducing the virus pool
and by containing the dissemination of the disease
through export of potentially infectious animals.
The latter can be a very significant problem as
infected animals, especially cattle, present viremia
for long periods (up to 100 days) with no clinical
signs (Erasmus, 1990; Roy, 1996).
BTV routine diagnosis is based primarily on
serological methods that detect virus-specific antibodies in serum (Pearson et al., 1992). A number
of other procedures are also used currently to
detect BTV from blood or tissues of infected
animals. These include direct inoculation of cultured mammalian or insect cells, or intravenous
inoculation into 10– 12 day embryonated chicken
eggs, followed by one passage in insect cell culture
and up to three passages in mammalian cell cultures (Foster and Leudke, 1968; Gard and
Kirkland, 1993; Wechsler and McHolland, 1988).
In particular, the inoculation of embryonated
chicken eggs and passaging through cell culture is
the generally accepted method for testing of animals for export and other regulatory purposes.
This is, however, a laborious and time-consuming
protocol that may take up to 5 weeks for completion. Consequently, alternative methods of virus
detection have been sought. These include antigen
capture enzyme linked immunosorbent assay
(ELISA), dot immunobinding assay (DIA), immunoelectron microscopy and polymerase chain
reaction (PCR) (Hawkes et al., 2000; Katz et al.,
1993; McColl and Gould, 1991; Mecham, 1993;
Mecham et al., 1990; Mecham and Nunamaker,
1994; Nunamaker et al., 1997a,b; Shad et al.,
1997). The use of antigen capture ELISA for the
detection of BTV in the blood of infected ruminants has either been unsuccessful (Mecham,
1993), has detected antigen only in animals with
high viremias (Stanislawek et al., 1996), or was
not consistent enough to allow for the reliable
diagnosis of BTV (Hawkes et al., 2000). A major
problem in the diagnosis of BTV infection by
immunological methods is also the cross-reactivity
with proteins from other orbiviruses (Lunt et al.,
1988), although this may be circumvented by the
use of c-ELISA (Afshar et al., 1989, 1987).
To avoid these problems, PCR-based assays
were developed and evaluated for the detection of
BTV serotypes based on nucleotide sequences of
different genome segments (Akita et al., 1992,
1993; Aradaib et al., 1998; Dangler et al., 1990;
McColl and Gould, 1991; Parsonson and McColl,
1995; Shad et al., 1997; Wade-Evans et al.,
1990; Wilson, 1999; Wilson and Chase, 1993).
Although these assays were found to detect BTV
in BTV-infected cell cultures and in infected experimentally ruminants, their usefulness and dependability under field conditions have not been
demonstrated.
The purpose of this study was to develop a
sensitive and dependable PCR assay for BTV
detection in clinical (field) specimens (blood samples) from sheep and goats infected naturally and
evaluate its sensitivity in comparison with conventional methods such as embryonated chicken eggs
inoculation and cell culture assays.
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
2. Materials and methods
2.1. Cells and 6iruses
Purified preparations of bluetongue virus
(BTV) serotypes 1, 2, 3, 4, 9 and 16 were obtained
from the Onderstepoort Veterinary Institute,
South Africa. Greek isolates of the BTV serotypes
4, 9 and 16, which represent different topotypes
than those obtained from South Africa, were also
used. The epizootic haemorrhagic disease virus
serotypes 1, 2 and 318, and the African horse
sickness virus serotype 9, were obtained from the
Institute of Animal Health, Pirbright Laboratory,
UK. When necessary, the viruses were propagated
on confluent BHK-21 cell monolayers, as described previously (Koumbati et al., 1999). In
brief, BHK-21 cell monolayers were grown in
BHK-21 Glasgow medium (Gibco BRL) supplemented with 10% fetal calf serum (FCS). The cells
were infected with 1 ml of treated blood cells and
the culture was incubated at 37 °C until the cytopathic effects became apparent. At that time, the
infected cell culture fluid was frozen and kept at
− 80 °C.
2.2. Clinical samples
Blood samples were collected from 24 sheep
and 12 goats, from naturally infected mixed
flocks, in EDTA-containing tubes (VACUTAINER™), during the bluetongue, serotype 9,
epizootic on the island of Rhodes, Greece in 1998.
The majority of the sheep in those flocks had
shown bluetongue clinical signs during the month
prior to sampling, whereas the goats appeared to
be healthy.
2.3. Virus isolation in embryonated chicken eggs,
cell culture assays and 6irus identification
Virus isolation and identification were processed as described previously (Koumbati et al.,
1999). Briefly, 0.1 ml of washed, packed and
ultrasonic-treated blood cells was inoculated intravenously into each of six 12-day-old embryonated chicken eggs. BTV-positive samples caused
embryo haemorrhage that usually resulted in em-
79
bryo death, in most cases. Dead embryos were
collected between 2 and 7 days post-inoculation
for virus identification.
BHK-21 cell monolayers were inoculated with
1.0 ml of treated sample and virus adsorption was
allowed to take place for 1 h and 30 min at 37 °C.
The monolayers were then washed twice with
phosphate-buffered saline (PBS) and BHK-21
Glasgow medium (Gibco BRL, Grand Island,
NY) was added. Monolayers were examined daily
for BTV cytopathic effects for a maximum of 10
days. If no obvious cytopathic effects were evident
for 10 days, the cells were removed and passaged
into a new monolayer. A sample was characterized as negative when it had shown no cytopathic
effects after three blind passages.
For virus identification, Vero cells were cultivated in 8-chambered slides (Lab-tek) and inoculated with either 0.2 ml of infected cell culture
medium or 0.2 ml of dispersed heart supernatant
from infected chick embryos. Positive and negative controls were included. Forty-eight hours
post-inoculation, the cells were fixed in situ with
acetone at 4 °C for 15 min and at − 70 °C for
another 30 min (Jochim et al., 1974). Virus identification was carried out by indirect immunofluorescence using the Pirbright monoclonal
antibody 3-17-A3 (Anderson, 1984). The presence
of intracytoplasmic fluorescent inclusions, which
are characteristic of bluetongue virus, was observed in the positive cultures.
2.4. RNA extraction procedures
The RNA extractions were carried using the
single-step method described by Chomczynski and
Sacchi (Chomczynski and Sacchi, 1987) with the
TRIzol™ LS reagent (Gibco BRL) available commercially. The precautions recommended (Sambrook et al., 1989) to avoid contamination with
ribonucleases were observed.
2.4.1. RNA extraction from blood samples and
chicken embryos
About 0.25 ml of total blood sample (diluted
1:1 with water), or 100 mg of heart tissue from
chicken embryos was homogenized in 0.75 ml
TRIzol™ LS reagent (Gibco BRL). Following a 5
80
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
min incubation, 0.2 ml of chloroform was added
to each sample and the solution was shaken by
hand and then incubated at room temperature for
approximately 10 min. The samples were centrifuged at 15,000× g for 15 min at 4 °C and the
RNA-containing aqueous phase was transferred
to a fresh tube. Following the addition of 0.5 ml
ice-cold isopropanol and a 30 min incubation at
− 20 °C, the RNAs were precipitated by centrifugation at 15,000×g for 15 min at 4 °C. The
resulting RNA pellets were washed with 75% ethanol before air-drying, and each RNA sample was
dissolved in 50 ml diethyl pyrocarbonate treated
H2O by incubating 10 min at 55 °C.
2.4.2. RNA extraction from infected cell cultures
Frozen, infected cell culture fluid, prepared as
described above, was thawed and centrifuged at
6000×g for 5 min. The virus was pelleted from
10 ml of the resulting supernatant by centrifugation at 25,000× g for 16 h at 4 °C. Subsequently,
the virus pellet was dissolved in 0.75 ml TRIzol™
LS reagent (Gibco BRL). The mixture of virus
pellet and the monophasic solution of phenol and
guanidine isothiocyanate was incubated for 5 min
at room temperature to permit the complete dissociation of nucleoprotein complexes. The RNAs
were then isolated as described above.
2.5. Oligonucleotide primers
The oligonucleotide primers CB1 and CB2
(Table 1) were designed based on published sequence (Pierce et al., 1998) of the BTV10 segment
10, (EMBL accession no. AF044381). All oligonucleotides were synthesized commercially (MWG,
Germany).
The
control
primers
BA1
(GAGAAGCTGTGCTACGTCCGC) and BA2
(CCAGACACGCACTGTGTTGGC), whose sequences are based on the conserved bovine b-actin
gene, were included in our duplex RT-PCR assay
since they provide a positive control for sample
quality, the integrity of the extracted RNA and
the efficiency of the reactions. BA1 and BA2 have
been shown to work well on RNA from all domestic species such as goat, pig, cattle and sheep
(Reid et al., 1998).
2.6. Re6erse transcription (RT) and polymerase
chain reaction (PCR) amplification
Reverse-transcription (RT) reactions were performed by mixing 7.6 ml of RNA extraction
product with 12.4 ml of an RT premix to obtain a
final concentration of 1× first strand buffer
(Gibco BRL), 0.5 mM dNTP mix (Gibco BRL),
10 mM DTT (Gibco BRL), 100 U of Moloney
murine leukaemia virus (MMTV) reverse transcriptase (Gibco BRL), and 3.5 pmol/ml of random hexamers (Pharmacia Biotech) per reaction.
Following preparation of the RT reaction mixtures on ice, reverse transcription reactions were
carried out at 37 °C for 30 min, they were terminated by heating for 5 min at 95 °C.
PCR amplifications were carried out observing
the guidelines described by Kwok and Higuchi
(Kwok and Higuchi, 1989). Each PCR reaction
mixture contained 2 ml of RT product, 1× PCR
buffer (Gibco BRL), 1 mM MgCl2 (Gibco BRL),
0.2 mM of a dNTP mix (Gibco BRL), 0.15 U of
Taq Polymerase (Gibco BRL), and 30 pmol of
each primer. The reaction volume was adjusted to
20 ml with DEPC treated H2O. The PCR mixtures
were prepared on ice and the reactions were initiated by heating for 3 min at 94 °C, followed by
30 cycles of: 1 min at 94 °C, 2 min at 52 °C and
1 min at 72 °C . The mixtures were then brought
to 72 °C for 5 min and then held at 4 °C. The
PCR conditions remained the same for both simple and duplex PCR reactions, with the CB1/CB2
and BA1/BA2 primer pairs. Following the amplification, 10 ml of each RT-PCR product was
analyzed by electrophoresis on a 2.5% agarose gel
and stained with ethidium bromide (0.5 mg/ml). A
100 bp DNA ladder (Gibco, BRL) was analyzed
on the same gel to serve as a size marker. The
expected sizes of the RT-PCR products were 792
bp for the BTV S10 amplimer and 275 bp for the
b-actin amplimer.
2.7. Specificity and sensiti6ity of the RT-PCR
procedure
The specificity of the assay was determined by
applying the above described RT-PCR protocol
to RNAs prepared from stocks of either the BTV
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
serotypes 1, 2, 3, 4, 9 and 16, or the closely related
orbiviruses epizootic haemorrhagic disease virus,
serotypes 1, 2 and 318, and the African horse
sickness virus serotype 9, or to RNAs extracted
either from cultures of uninfected BHK-21 cells or
blood samples from uninfected sheep or goats.
The sensitivity of the assay was evaluated by
81
applying it to virus detection using logarithmic
dilutions of a titrated BTV, serotype 4, suspension
in sterile 0.04 M phosphate buffer, pH 7.2. Each
virus dilution was successively subjected to the
RNA isolation, reverse transcription and PCR
amplification steps, as outlined previously, and
the products were analyzed on a 2.5% agarose gel.
Table 1
Homology of the BTV primers used to S10 region sequences from field and laboratory strains of bluetongue virus
BTV strain/accession
numbera
CB1 primer (bp 21–41)
TGCTATCCGGGCTGATCCAAAb
CB2 primer (bp 813–796)
TAGCGCCGCGTACCCTCCb
VAC10/AF044376
10B80Z/AF044379
10B81X/AF044382
10B81U/AF044381
10B9OZ/AF044384
10O80Z/AF044380
10O90H/AF044385
VAC11/AF044377
11B80Z/AF044702
11B81P/AF044383
11C81Z/AF044703
11O79X/AF044386
11O81X/AF044704
13B80Z/AF044702
13B81K/AF044712
13B89Z/AF044710
13O79Z/AF044713
VAC17/AF044378
17B80Z/AF044705
17B81Y/AF044707
17B90Z/AF044708
17O79Y/AF044706
17O90Y/AF044709
BTV4/AF135226
BTV9
BTV16/AF135229
Austral BTV/D00253
BTV1/AF135223
BTV2/AF135230
BTV3/AF135225
BTV12/AF135227
BTV15/AF135228
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
(+)
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
(+)c
(+)
(+)
+
(+)
(+)
(+)
?c
?
a
The primer sequences were compared with all available BTV sequences. The accession number is given next to the virus
designation.
b
Numbers indicate the location of primer sequences within BTV S10 region, and are based on the BTV10 sequence (10B81U)
(Pierce et al., 1998).
c
Plus in parenthesis (+) means that the RT-PCR was found to work well with the isolates tested, despite absence of match
between primer sequences and the published sequence of the particular serotype, and (?) that the primer sequence is not present in
the published BTV sequence and no experimental verification was done due to unavailability of the particular serotype.
82
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
Fig. 1. Specificity of the RT-PCR method with the CB1 and
CB2 primers. (A) Total RNAs were isolated either from
uninfected BHK-21 cells (lanes 2 and 3) or from purified virus
preparations of bluetongue virus serotypes 4 (BTV4, lane 4), 9
(BTV9, lane 5) and 16 (BTV16, lane 6), or epizootic haemorrhagic disease virus serotype 1(EHDV1, lane 7), 2 (EHDV2,
lane 8) and 318 (EHDV318, lane 9), and African horse sickness virus serotype 9 (AHSV9, lane 10). All BTV viruses used
in this panel were isolated in Greece. (B) Total RNAs were
isolated either from control uninfected goat blood (Blood, lane
2), or from purified virus preparations of BTV serotypes 1
(BTV1, lane 3), 2 (BTV2, lane 4), 3 (BTV3, lane 5), 4 (BTV4,
lane 6), 9 (BTV9, lane 7) and 16 (BTV16, lane 8). All BTV
viruses analyzed in this panel were obtained from the Onderstepoort Veterinary Institute, South Africa, and they represent
different topotypes than those analyzed in panel A. The RTPCR reactions were carried out, with the CB1/CB2 primer
pair, as described in Section 2 and the reaction products were
analyzed on a 2.5% agarose gel. The size markers used in both
panels (M, lane 1) were a 100-bp DNA ladder (Gibco-BRL).
Fig. 1. (Continued)
3. Results
3.1. Specific bluetongue 6irus RNA detection
using re6erse transcription PCR with the CB1
and CB2 primers
The CB1 and CB2 primers were designed based
on the sequence of the highly conserved S10 region, coding for the nonstructural proteins NS3
and NS3A, of the BTV10 strain sequence
(10B81U, Accession no. AF044381) (Pierce et al.,
1998). The CB1 primer hybridizes to the 5% end of
the NS3/NS3A gene (bases 21– 41) (Pierce et al.,
1998), and CB2 hybridizes close to the edge of the
noncoding 3% end (bases 813–796) of the NS3/
NS3A gene. Care was taken, in primer design, to
avoid primer internal stability problems, dimer
formation, or homologies with RNAs from other
orbiviruses, other regions of the BTV RNA, or
cellular RNAs. As shown in Table 1, the sequences of both the CB1 and the CB2 primers
were found to exist in the majority of the published BTV S10 region (NS3 and NS3A gene)
sequences. The apparent absence of CB2 sequences in the published sequences of some BTV
serotypes might be due to incomplete sequencing
at the far 3% end of the S10 RNA region. This
notion is based on the finding that the CB1/CB2
primers were found to amplify a fragment of the
appropriate size (792 bp) from at least five BTV
serotypes (1, 2, 3, 4 and 16) whose published S10
region sequences lack homology with the CB2
primer (Fig. 1).
As stated above, the RT-PCR method described here was found to detect BTV RNA isolated from purified preparations of the serotypes
1, 2, 3, 4, 9 and 16 (Fig. 1). BTV RNAs for
serotypes 4, 9 and 16 isolated from different geographical regions (topotypes), i.e. Greece (Fig.
1A) and South Africa (Fig. 1B) were detected
equally well by our method. In addition, two
Greek isolates from different geographical regions
(topotypes) of bluetongue serotype 4 were also
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
83
detected (data not shown). In contrast, no PCR
product was detected when total RNA, isolated
either from control uninfected BHK-21 cells (Fig.
1A) or blood samples from uninfected sheep or
goats (Fig. 1B), were used. The specificity of the
method was further demonstrated by the absence
of amplification products when RNAs from three
serotypes (1, 2 and 318) of the closely related
epizootic haemorrhagic disease virus or the
African horse sickness virus, serotype 9 were used
as templates (Fig. 1A). The above data argue
against the possibility that false positive results
will be obtained with the RT-PCR method, due to
the presence of either cellular RNAs or RNAs of
related viruses.
3.2. Sensiti6ity of the RT-PCR procedure
The sensitivity of the assay was evaluated by
using serial dilutions of a titrated BTV4 suspension. The virus had been titrated on BHK-21 cells
and the titer is expressed as cell culture infectious
dose (CCID50) units. It was found that our RTPCR assay could detect amounts of BTV as low
as 5 CCID50 (Fig. 2).
Fig. 2. Sensitivity of the RT-PCR method. The sensitivity of
the assay was evaluated by performing RT-PCR reactions on
RNAs prepared from serial dilutions of a BTV4 suspension
that had been titrated in BHK-21 cells. The titration units are
expressed as CCID50, and the virus load equivalent used in
each RT-PCR reaction is 109 (lane 2), 107 (lane 3), 105 (lane
4), 103 (lane 5), 100 (lane 6), 10 (lane 7), 5 (lane 8), 2.5 (lane
9) and 0 (lane 10). The reaction products were analyzed on a
2.5% agarose gel and the size markers used (M, lane 1) were a
100-bp DNA ladder (Gibco-BRL).
3.3. Application of the RT-PCR assay in
bluetongue 6irus RNA detection to field specimens
To determine the actual value of the above-described RT-PCR assay we applied it to BTV
RNA detection using blood samples collected
from 24 sheep and 12 goats, from naturally infected mixed flocks, during a bluetongue epizootic
on the island of Rhodes, Greece in 1998. The
results of this assay were compared with the results of two other standard tests used in routine
BTV diagnosis, namely cytopathic effects on
BHK-21 cells in culture and inoculation into 12day embryonated chicken eggs.
Virus detection in the field diagnostic samples
using inoculation of embryonated chicken eggs
indicated that 13 out of the total 36 samples were
positive for the presence of the virus (Table 2).
Embryonic death was noted, in the positive samples, between days 2 and 7 post inoculation and
the number of dead embryos varied from one to
five out of six eggs that had been inoculated with
each field sample. All positive blood samples were
derived from sheep, whereas all goat samples were
found negative. Further analysis, by indirect immunofluorescence, indicated that all the positive
samples were indeed positive for BTV antigens
(data not shown). Therefore, 13 out of the 36
samples were confirmed as positive for the presence of the bluetongue virus using embryonated
chicken eggs inoculation (Table 2).
BTV detection by measuring virus cytopathic
effects on cultured BHK-21 cell monolayers was
less efficient, since only nine positive samples were
detected (Table 2). These nine positive samples,
were also found positive by indirect immunofluorescence with an anti-BTV monoclonal antibody, and were a subset of the 13 samples found
positive by the embryonated chicken eggs assay
(Table 2).
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
84
PCR (Table 2). Inclusion of the b-actin and BTV
primers in the same RT-PCR reactions provides
an additional control against possible false negative results. Indeed, the presence of the 275 bp
b-actin RNA amplification product, using the
conserved BA1 and BA2 b-actin primers in the
In contrast, the duplex RT-PCR assay was far
more sensitive since it detected 17 positive samples (Fig. 3; Table 2). It is noted that the 13
samples found positive by the embryonated
chicken eggs inoculation and cell culture assays
were among those identified as positive by RT-
Table 2
Comparison of the RT-PCR with two conventional BTV diagnosis assays
Number
S1
S2
S3
S4
S5
S6
S7
S8
S9
S10
S11
S12
S13
S14
S15
S16
S17
S18
S19
S20
S21
S22
S23
S24
G1
G2
G3
G4
G5
G6
G7
G8
G9
G10
G11
G12
RT-PCR
+
+
+
+
+
+
+
+
+
+
+
−
−
+
−
−
+
+
+
+
+
−
−
−
−
−
−
−
−
−
−
−
−
−
ECEs
+
+
+
−
−
+
+
+
+
+
+
−
−
+
−
−
−
−
+
+
−
+
−
−
−
−
−
−
−
−
−
−
−
−
−
−
Cell culture
+
+
+
−
−
+
+
+
+
+
−
−
+
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
RT-PCRa
ECEs
Cell culture
+
+
+
ND
ND
+
+
+
+
+
+
ND
ND
+
ND
ND
ND
ND
+
+
ND
+
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
+
+
+
+
+
+
+
+
+
+
+
−
−
+
−
−
+
+
+
+
−
+
−
−
−
−
−
−
−
−
−
−
−
−
−
−
The conventional BTV diagnosis assays were (i) inoculation of 12-day embryonated chicken eggs (ECEs) and (ii) inoculation of
BHK-21 cell cultures (cell culture). Thirty six clinical samples (blood) were tested. (+)= BTV-positive sample, (−)=negative
sample, (ND) = not done.
a
RT-PCR was performed on RNAs isolated either from positive ECEs (dead embryos) or from cell cultures inoculated with
clinical samples.
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
85
Fig. 3. Application of the duplex RT-PCR method in field diagnosis. Total RNAs were isolated either from BHK-21 cells infected
with BTV9, as control, or from 24 blood samples from sheep (S1 – S24) and 12 from goats (G1 – G12). RT-PCR reactions were
performed and the products (792 bp for the BTV product and 275 bp for that of b-actin, indicated by arrows) were analyzed on
2.5% agarose gels (A and B) and a 100-bp DNA ladder was used as size markers (M).
same reactions (Fig. 3), practically nullifies the
possibility that the lack of amplification in the
negative samples is due to low quality RNA or
other technical problems.
The increased specificity of the RT-PCR
method became more obvious when it was applied
on RNAs obtained from BHK-21 cell cultures
that had been inoculated with the clinical samples.
Positive RT-PCR results, for the presence of
BTV, were obtained not only for the nine samples
showing cytopathic effects, but for an additional
eight samples (Table 2). All of the above samples
had been identified as positive by RT-PCR on
RNAs isolated from the blood samples (Table 2).
RT-PCR assays on RNAs from dead chicken
embryos further verified the presence of BTV in
these samples (Table 2).
4. Discussion
Monitoring and control of BTV infection in
cattle, sheep and goats remains a top priority in
BTV-endemic and epidemic countries interested in
exporting livestock free of this disease or in restricting the introduction of new serotypes into
already existing endemic populations (Morley,
1993; Roberts et al., 1993). While serological assays are sensitive and easy to use they only
provide evidence indicating earlier animal expo-
86
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
sure to BTV but not necessarily an ongoing infection. Earlier work on cattle and sheep has provided evidence that seropositivity does not
correlate with circulating BTV RNA or the presence of infectious virus in the blood of the
seropositive animals (Singer et al., 1998).
A number of procedures have been developed
to detect the presence of BTV antigens or nucleic
acids. PCR is a powerful tool in the field of
diagnostic medicine and it has been used successfully in identifying several infectious diseases of
veterinary importance (Belak and Ballagi-Pordany, 1993). By using PCR-based techniques, researchers may circumvent problems such as
serologic cross-reactivity among related orbiviruses (Ristow et al., 1988). Several RT-PCR
assays have been described for the detection of
BTV (Akita et al., 1992, 1993; Aradaib et al.,
1998; Dangler et al., 1990; McColl and Gould,
1991; Parsonson and McColl, 1995; Shad et al.,
1997; Wade-Evans et al., 1990; Wilson, 1999;
Wilson and Chase, 1993). However, despite the
fact that these assays have been tested using virusinfected cultures or experimentally infected animals, their usefulness in clinical diagnosis remains
mostly unproven due to a lack of comparative
evaluation with standard methods using field samples. The above is underscored by a recent report
(Tiwari et al., 2000) which shows that an RT-PCR
method that works well with purified BTV RNA
fails to detect the virus RNA in clinical samples,
unless a second nested-PCR step is included.
In an earlier report of a BTV PCR assay (McColl and Gould, 1991), virus nucleic acid was only
detected in the leukocyte fraction of blood from
infected animals. The aim of the present study,
however, was to develop a simpler RT-PCR assay, that could detect the virus in total unfractionated blood. It was also imperative that the assay
should detect all known BTV serotypes but no
other related viruses (such as epizootic haemorrhagic disease virus), a problem that has plagued
many serological assays for this virus.
To achieve this primers were designed that
could amplify segment 10 of the BTV10 genome,
a region that codes for the non structural proteins
NS3 and NS3A. Selection of the particular
genome segment, as a target for a diagnostic
RT-PCR assay was based on the observation that
it is one of the most highly conserved segments of
BTV genome (Hwang et al., 1992; Pierce et al.,
1998). Additional care was taken, in primer design, to avoid homologies between primer sequences and sequences of known virus or
mammalian genes. As a result we obtained a set
of primers (CB1 and CB2) that could amplify
specifically the BTV genome without any obvious
background when either uninfected BHK-21 cells
(Fig. 1A) or blood samples from uninfected sheep
or goats (Fig. 1B) were used. Additionally, no
cross-reactivity was observed with two other
closely related orbiviruses, such as the epizootic
haemorrhagic disease virus and the African horse
sickness virus (Fig. 1A). These data minimize the
possibility that false positive results will be obtained due to amplification of either cellular or
related orbivirus RNAs.
Furthermore, the assay proved to be sensitive,
as it could detect quantities of BTV as low as 5
CCID50 (Fig. 2). There is always a possibility that
a PCR-based assay will yield false negative results
due to low quality of some clinical samples, technical problems or experimental error. Therefore, it
is crucial to include an internal control for the
identification of such false negatives. The presence
of the b-actin primers (BA1 and BA2) in our
duplex RT-PCR represents such an internal control, since the absence of a b-actin amplification
product would indicate that the particular sample
is a false negative. The conditions were optimized
so that the double BTV/b-actin PCR reactions
could take place in the same tubes without lose of
specificity or sensitivity (data not shown).
The results presented in Fig. 3, where the
method was applied in clinical diagnosis using 36
whole blood samples, argue that this method does
achieve the goals set at the beginning of the work.
Furthermore, the diagnostic sensitivity of the RTPCR assay is greater than the virus inoculation in
embryonated chicken eggs or assay of cytopathic
effects on cultured cells (Table 2). This is not
surprising since it has been reported previously
that the presence of BTV RNA in the blood of
affected animals, as indicated by positive RTPCR results, does not always correlate with infectivity (Katz et al., 1994; Tabachnick et al., 1996).
C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89
These data indicate that the RT-PCR method is
specific and more sensitive than the accepted BTV
detection methods that were used comparatively
in this study. Moreover, it is far simpler and much
faster than these other methods.
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