2. Materials and Methods
2.1 Ethics statement
Blood samples were collected from animals in public slaughterhouses
during the mandatory ante-mortem clinical examination. All procedures
performed in this study followed common good clinical practices and
received institutional approval from the Ethical Animal Care and Use
Committee of the University of Naples Federico II (PG/2017/ 0099607).
All farmers were previously informed and in agreement with the purpose
and methods used.
2.2 Liquid biopsy samples and DNA extraction
Blood samples from 103 apparently healthy 1- to 3-year-old sheep were
collected from the jugular vein in vacutainers containing
ethylenediaminetetraacetic acid (EDTA). A total of 40 samples were
obtained from sheep living in Sardinia (Sar) (20) and Campania (Cam)
(20), 48 samples from Calabria (Cal) (24) and Basilicata (Bas) (24), and
15 samples from Apulia (Apu). All sheep, excluding those from Cam, were
from flocks that lived and shared the bracken fern-infested lands that
they grazed on with pasture-residing cattle. Sheep from Cam were from
flocks living in closed pens without any contact with other animals.
Total DNA was extracted using a DNeasy Blood & Tissue Kit (Qiagen,
Wilmington, DE, USA), according to the manufacturer’s instructions.
2.3. RT-qPCR
RT-qPCR was performed in a final volume of 20 μL containing 10 μL of
TaqMan Universal Master Mix (Applied Biosystems, Foster City, CA, USA),
900 nM of each of the forward and reverse primers (Bio-Rad Laboratories,
Hercules, CA, USA), 250 nM of the probe (Bio-Rad Laboratories), and 100
ng of the DNA sample. The primers and probes for the detection of four
BPV genotypes (BPV-1, -2, -13, and -14) were used as reported elsewhere
(De Falco et al., 2020). The reaction was performed on the CFX96
Real-Time System of the C1000 TouchTM Thermal Cycler
(Bio-Rad Laboratories). The thermal cycling conditions were as follows:
50 °C for 2 min, 95 °C for 10 min, and 40 cycles of 95 °C for 15 s and
58 °C for 60 s (acquiring FAM and VIC dyes). Each sample was analyzed in
duplicate, and negative controls were included in all runs. Data
acquisition and data analyses were performed using CFX
MaestroTM (Bio-Rad Laboratories) software. The same
samples used as positive controls for ddPCR were also tested using
RT-qPCR.
For ddPCR, Bio-Rad’s QX100 ddPCR System was used according to the
manufacturer’s instructions. The reaction was performed in a final
volume of 20 μL containing 10 μL of ddPCR Supermix for Probes (no dUTP
2×; Bio-Rad), 0.9 μM primer, and 0.25 μM probe together with 5 μL sample
DNA (100 ng). A black hole quencher was used in combination with FAM and
VIC fluorescent dye reporters (Bio-Rad Laboratories). The reaction
mixture was placed into the sample well of a DG8 cartridge (Bio-Rad
Laboratories). A volume of 70 μL of droplet generation oil was loaded
into the oil well, and droplets were formed in the droplet generator
(Bio-Rad Laboratories). After processing, the droplets were transferred
to a 96-well PCR plate (Eppendorf, Hamburg, Germany). PCR amplification
was carried out on a T100 Thermal Cycler (Bio-Rad Laboratories) with the
following thermal profile: hold at 95 °C for 10 min, 40 cycles of 94 °C
for 30 s and 58 °C for 1 min, 1 cycle at 98 °C for 10 min, and ending at
4 °C. After amplification, the plate was loaded onto a droplet reader
(Bio-Rad Laboratories) and the droplets from each well of the plate were
read automatically. QuantaSoft software was used to count the
PCR-positive and PCR-negative droplets to provide absolute
quantification of the target DNA. Therefore, the ddPCR results could be
directly converted into copies/µL in the initial samples simply by
multiplying them by the total volume of the reaction mixture (20 µL) and
then dividing that number by the volume of DNA sample added to the
reaction mixture (5 µL) at the beginning of the assay. Samples with very
few positive droplets were re-analyzed to ensure that these low copy
number samples were not due to cross-contamination.
2.5 Statistical analysis
Differences in the proportions of detected cases were tested using the
chi-square test by Campbell and Richardson (Richardson, 2011).
Furthermore, regarding the significance relative to the number of copies
of BPV DNA detected in sheep in the different regions, the t-test was
used after adjusting for the Bonferroni multiple comparison correction
of means. P-values ≤ .05 were considered to be statistically
significant. All analyses were performed using R statistical software
(The R Foundation, Vienna, Austria).
3. Results
Overall, our results showed that BPV DNA was found in 68 out of 103
blood samples (66%) from healthy sheep using ddPCR. The same liquid
biopsies were also investigated using RT-qPCR, which revealed BPV DNA in
approximately 9% of blood samples (Fig 1). In 42 of the positive
samples (61.8%), a single BPV infection was observed (Fig. 2), 26 of
which were caused by BPV-2 (61.9%) and 7 by BPV-13 (16.7%). BPV-14 was
responsible for 7 single infections (16.7%), and BPV-1 for two single
infections (4.7%) (Fig. 3). Multiple BPV infections were seen in 26
(38.2%) positive samples. BPV coinfections caused by two genotypes were
seen in 22 positive cases (84.6%), with dual BPV-2/BPV-13 infection
being the most prevalent. BPV coinfections by triple and quadruple
genotypes were detected in 11.5% (3/26) and 3.8% (1/26) of blood
samples, respectively (Fig. 4). In sheep flocks that lived and shared
lands with cattle, BPV DNA was detected in approximately 53% of blood
samples collected in Apu (8/15), 75% of samples acquired in both Bas
and Cal (18/24), and 100% of blood samples harvested from Sar (20/20).
In sheep flocks from Cam that lived in isolated and closed pens without
any contact with cattle, BPV DNA was detected in 20% of blood samples
examined (4/20). The percentage differences in BPV infections in all
sheep flocks with cattle contact were statistically significant compared
to the percentage observed in sheep flocks without any contact with
cattle, as the Campbell-Ricardson’s chi-square test resulted in a
p-value < 0.05. Furthermore, in all geographical areas except
for Apu, BPV-2 was the most prevalent genotype. BPV-13 and BPV-14, as
well as BPV-1 were also observed. Furthermore, Apu BPV-14 showed very
high numbers of copies/µL (mean value of 895.2); it was the most
prevalent BPV genotype at a detection level of 40% in the examined
samples (6/15) and a statistically significant Campbell-Ricardson’s
chi-square test p-value < .05.
The overall quantification results showed that viral copy numbers/µL
ranged from 76 to 568 for BPV-1 (mean value = 183.6 ), 65 to 3768 for
BPV-2 (mean value = 397.1), 66 to 2112 for BPV-13 (246.5), and 76
to 1768 for BPV-14 (mean value = 447.5). When sheep flocks from Sar
alone were considered, BPV-2 showed the highest copy numbers/µL with a
mean of 1162. Using t-tests, the differences between the copy numbers of
BPV-2 in Sar compared to the other means found in Cal, Cam, Bas, and Apu
were statistically significant with p-values < .05. Indeed,
after adjusting for the Bonferroni multiple comparison correction, their
p-values were .003 (Sar-Cal), .04 (Sar-Cam), .002 (Sar-Bas), and .01
(Sar-Apu), respectively. Table 1 summarizes the quantitative data using
QuantaSoft software, demonstrating the numbers of copies/µL in blood
samples from sheep flocks located in the five regions in South Italy.