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\begin{document}

\title{Evidence of a novel cross-species transmission by ovine papillomaviruses}



\author[1]{Francesca  De Falco}%
\author[2]{Anna  Cutarelli}%
\author[1]{Bianca Cuccaro}%
\author[3]{Cornel Catoi}%
\author[2]{Esterina De Carlo}%
\author[1]{Sante Roperto}%
\affil[1]{Universita degli Studi di Napoli Federico II Dipartimento di Medicina Veterinaria e Produzioni Animali}%
\affil[2]{Istituto Zooprofilattico Sperimentale del Mezzogiorno}%
\affil[3]{Universitatea de Stiinte Agricole si Medicina Veterinara Cluj-Napoca Facultatea de Agricultura}%


\vspace{-1em}



  
  \date{May 26, 2024}


\begingroup
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\selectlanguage{english}
\begin{abstract}
Ovine papillomavirus (OaPV) comprises four genotypes; OaPV1, OaPV2, and
OaPV4 are fibropapillomaviruses within the genus
\emph{Delta}-papillomavirus ( \emph{Delta-}PV \emph{)}, whereas OaPV3 is
an epitheliotropic virus that belongs to the genus
\emph{Dyokappa}-papillomavirus ( \emph{Dyokappa-}PV \emph{)}. To date,
all of them have been known to infect sheep only. OaPV1, OaPV2, and
OaPV4 have been associated with ovine cutaneous and mucosal
fibropapillomas, while OaPV3 is a key factor in the squamous cell
carcinoma (SCC) pathway of the sheep skin. Peripheral blood mononuclear
cell (PBMC) samples obtained from 128 cattle at public slaughterhouses
were investigated using droplet digital polymerase chain reaction
(ddPCR). ddPCR is a new-generation PCR technique that enables accurate
and absolute quantification of target molecules with high sensitivity
and specificity. All OaPVs were detected by identification and
quantification of nucleic acids using specific fluorescent probes. Of
128 PBMC samples, 100 (\textasciitilde{}78\%) showed OaPV infections.
Further, 42, 35, and 23 PBMC samples showed single, double, and triple
OaPV infections, respectively. OaPV1 was responsible for 22 single
infections, OaPV2 caused 16 single infections, and OaPV3 and OaPV4
caused two single infections each. OaPV1 and OaPV2 were the most
frequent ovine viruses in dual and triple infections. In many PBMC
samples, both ovine \emph{Delta-}PV and \emph{Dyokappa}-PV were found to
be transcriptionally active, as shown by the detection and
quantification of E5 oncogene transcripts for OaPV1, L1 transcripts for
OaPV2, E6 and E7 transcripts for OaPV3, and E6 for OaPV4. OaPVs were
found in the blood samples from cattle that shared grasslands rich in
bracken ferns known to contain immunosuppressant substances.
Furthermore, OaPVs were also found in cattle from intensive livestock
farming without any contact with sheep. Because OaPV DNA was detected in
both grass hay and corn silage, it is conceivable that these feed may be
the viral sources.%
\end{abstract}%



\sloppy


Evidence of a novel cross-species transmission by ovine papillomaviruses

Short title: OaPV transmission in cattle

Francesca De Falco\textsuperscript{1}, Anna
Cutarelli\textsuperscript{2}, Bianca Cuccaro\textsuperscript{1}, Cornel
Catoi\textsuperscript{\textbf{3}} , Esterina De
Carlo\textsuperscript{2}, Sante Roperto\textsuperscript{1*}

\textbf{1} Dipartimento di Medicina Veterinaria e delle Produzioni
Animali, Universit\selectlanguage{ngerman}à degli Studi di Napoli Federico II, Napoli,
Italia;\textbf{2} Istituto Zooprofilattico Sperimentale del Mezzogiorno,
Portici (NA), Italia: \textbf{3} Pathology Department, Faculty of
Veterinary Medicine, University of Agricultural Sciences and Veterinary
Medicine, Cluj-Napoca, Romania

Corresponding author: Dr. Sante Roperto

email:\emph{sante.roperto@unina.it}

\textbf{Summary}

Ovine papillomavirus (OaPV) comprises four genotypes; OaPV1, OaPV2, and
OaPV4 are fibropapillomaviruses within the genus\emph{Delta}
-papillomavirus (\emph{Delta-} PV\emph{)} , whereas OaPV3 is an
epitheliotropic virus that belongs to the genus\emph{Dyokappa}
-papillomavirus (\emph{Dyokappa-} PV\emph{)} . To date, all of them have
been known to infect sheep only. OaPV1, OaPV2, and OaPV4 have been
associated with ovine cutaneous and mucosal fibropapillomas, while OaPV3
is a key factor in the squamous cell carcinoma (SCC) pathway of the
sheep skin. Peripheral blood mononuclear cell (PBMC) samples obtained
from 128 cattle at public slaughterhouses were investigated using
droplet digital polymerase chain reaction (ddPCR). ddPCR is a
new-generation PCR technique that enables accurate and absolute
quantification of target molecules with high sensitivity and
specificity. All OaPVs were detected by identification and
quantification of nucleic acids using specific fluorescent probes. Of
128 PBMC samples, 100 (\textasciitilde{}78\%) showed OaPV infections.
Further, 42, 35, and 23 PBMC samples showed single, double, and triple
OaPV infections, respectively. OaPV1 was responsible for 22 single
infections, OaPV2 caused 16 single infections, and OaPV3 and OaPV4
caused two single infections each. OaPV1 and OaPV2 were the most
frequent ovine viruses in dual and triple infections. In many PBMC
samples, both ovine \emph{Delta-} PV and \emph{Dyokappa} -PV were found
to be transcriptionally active, as shown by the detection and
quantification of E5 oncogene transcripts for OaPV1, L1 transcripts for
OaPV2, E6 and E7 transcripts for OaPV3, and E6 for OaPV4. OaPVs were
found in the blood samples from cattle that shared grasslands rich in
bracken ferns known to contain immunosuppressant substances.
Furthermore, OaPVs were also found in cattle from intensive livestock
farming without any contact with sheep. Because OaPV DNA was detected in
both grass hay and corn silage, it is conceivable that these feed may be
the viral sources.

\textbf{Keywords:} Cross-species transmission; Ovine papillomaviruses;
cattle; blood; grass hay; corn silage.

\textbf{Introduction}

Papillomaviruses (PVs) are small, non-enveloped, double-stranded DNA
viruses infecting mucosal and cutaneous epithelia of mammals, reptiles,
birds, and fish (IARC, 2007; Willemsen et al., 2020). As part of the
commensal flora, these viruses can be found in the healthy skin and
mucosa in a latent state; reactivation occurs following the loss of
immunity, resulting in a persistent infection which causes oncogenic
risk, with occurrence of tumors at several body sites (Sichero et al.,
2019; Strickley et al., 2019).

Ovine papillomavirus (OaPV) infections occur in sheep and are caused by
four oncogenic genotypes. OaPV1, OaPV2, and OaPV4 belong to the genus
\emph{Delta} -PV, while OaPV3 belongs to the genus \emph{Dyokappa-} PV
(http://pave.niaid.nih.gov/). Ovine \emph{Delta-} PV is characterized by
marked tropism for both mesenchymal and epithelial cells (Tore et al.,
2017), whereas OaPV3 exclusively infects epithelial cells (Alberti et
al., 2010). OaPVs have sporadically been associated with ruminal
fibropapillomas, papillomas, papillomatosis, and fibropapillomas of the
skin (Gibbs et al., 1970; Vanselow et al., 1982; Norval et al., 1985;
Trenfield et al., 1990; Tilbrook et al., 1992; Hayward et al., 1993;
Uzal et al., 2000). Although it has been suggested that OaPVs may be
responsible for the progression of cutaneous papillomas to squamous cell
carcinomas (SCCs) in sheep (Vanselow et al., 1982), a novel OaPV, namely
OaPV3, was only recently identified in a high number of SCCs in sheep,
suggesting that OaPV3 could represent a key infectious agent in the
onset of SCC in ovine species (Alberti et al., 2010; Vitiello et al.,
2015). OaPV3 and OaPV4 are well-characterized molecularly, as they are
the only OaPVs identified in tumor samples from sheep. Indeed, it has
been shown that the E6 and E7 oncogenes of OaPV3 and OaPV4 can
immortalize primary sheep keratinocytes and regulate the levels of
proliferative proteins such as cyclin A and cyclin-dependent kinases
(CDKs). However, it has been suggested that only OaPV3 E7 can strongly
promote the cleavage and degradation of ovine retinoblastoma protein
(pRb) (Tore et al., 2019). Calpain-mediated cleavage of pRb may result
in the dysregulation of E2F transcription factors, which play crucial
roles in the cell cycle, cell proliferation, and viral replication
(Darnell et al., 2007; Scarth et al., 2021). OaPV1 and OaPV2 are not
well characterized molecularly so far; however, it has been postulated
that they could be associated with tumors in sheep. DNA sequences
related to OaPV2 E5 have been found in the mass of the buccal cavity of
a pig, suggesting that similar to bovine \emph{Delta} -PVs, ovine
fibropapillomaviruses may also be responsible for cross-species
transmission (Munday et al., 2020). Unlike OaPV3 that induces cell
transformation by the E6 and E7 oncoproteins (Tore et al., 2019), ovine
\emph{Delta} -PVs may exert their main oncogenic activity through the
oncoprotein encoded by the E5 gene, as verified in most artiodactyl
fibropapillomaviruses (Munger and Howley, 2002). OaPV1, OaPV2, and OaPV4
are fibropapillomaviruses and belong to the \emph{Delta} -PV clade,
which is known to encode the most highly conserved E5 oncoproteins (Van
Doorslaer, 2013); this is likely because of the integration of the E5
ORF in the \emph{Delta-} PV genus occurring between 65 and 23 million
years ago (Garcia-Vallvé et al., 2005).

Recently, the first systematic research on the molecular epidemiology of
OaPV infection was conducted in sheep and revealed a divergent
geographical prevalence of OaPV genotypes. Furthermore, this survey
showed a high prevalence of OaPV infection since OaPV DNA was found in
up to 76.4\% of the peripheral blood of apparently healthy sheep (De
Falco et al., 2021b).

This study aimed to provide evidence of a novel cross-species
transmission and infection by OaPVs which were detected, quantified, and
found to be expressed in the peripheral blood mononuclear cells (PBMCs)
of cattle.

\textbf{Materials and Methods}

\textbf{Ethics statement}

We did not perform any animal experiments in this study. All samples
were collected postmortem from slaughterhouses, and hence, ethics
approval was not required.

\textbf{DNA Extraction from blood and matrix samples}

Blood samples of 128 healthy cattle aged 2\selectlanguage{english}-14 years were collected from
regions of southern Italy (Basilicata, Calabria, Campania). All animals
were pasture-free, except for 30 cattle from intensive livestock
farming. Blood samples harvested in ethylenediaminetetraacetic acid
(EDTA)-acid-containing vacutainers were obtained from healthy cattle at
public slaughterhouses. Total DNA was extracted using a DNeasy Blood \&
Tissue Kit (Qiagen, Wilmington, DE, USA) according to the manufacturer's
instructions. Furthermore, DNA was extracted from samples of grass hay
and maize silage (used as feed for cattle) obtained from intensive
livestock farming. The DNeasy plant Mini Kit (Qiagen, Wilmington, DE,
USA) was used for the extraction of hay and maize DNA, according to the
manufacturer's instructions.

\textbf{Positive Controls}

The positive controls for \emph{Delta} and \emph{Dyokappa} OaPVs were
obtained as previously reported (De Falco et al., 2021b).

\textbf{ddPCR}

Table 1 lists the primers and probes used for ddPCR. Primers and probes
were obtained as a mixture containing a primer-to-probe ratio of 3.6
(final concentration of 900 nM of each primer and 250 nM of probe). For
ddPCR, a Bio-Rad QX100 ddPCR system was used according to the
manufacturer's instructions. The reaction was performed in a final
volume of 22 \selectlanguage{greek}μ\selectlanguage{english}L and contained 11 \selectlanguage{greek}μ\selectlanguage{english}L of ddPCR Supermix for Probes (2X;
Bio-Rad Laboratories, Hercules, CA, USA), 1 \selectlanguage{greek}μ\selectlanguage{english}L of OaPVs primer and probe
mixture, 7 \selectlanguage{greek}μ\selectlanguage{english}L of DNA samples (corresponding to 100 ng), and 3 \selectlanguage{greek}μ\selectlanguage{english}L of
DNAase-free water. The plate containing the reactions was subsequently
transferred to an automated droplet generator (AutoDG; Bio-Rad
Laboratories, Hercules, CA, USA). AutoDG added 70 \selectlanguage{greek}μ\selectlanguage{english}L of droplet
generation oil to each well, and each sample was partitioned into
approximately 20,000 stable nanodroplets. The droplet generator
transferred each row of 8 droplet emulsion (40 \selectlanguage{greek}μ\selectlanguage{english}L) champions into a new
96 well PCR plate, which was subsequently coated with a pierceable film
heat-sealed using a PX1 PCR Plate Sealer (Bio-Rad Laboratories,
Hercules, CA, USA). PCR amplification was performed using a T100 Thermal
Cycler (Bio-Rad Laboratories, Hercules, CA, USA) with the following
thermal profile: hold at 95\selectlanguage{ngerman}°C for 10 min, 40 cycles of 94°C for 30 s,
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, Hercules, CA, USA), and the droplets from each well of the
plate were read automatically. The data were analyzed using the
QuantaSoft analysis tool (Bio-Rad Laboratories, Hercules, CA, USA).
Poisson statistics were used to calculate the absolute concentration of
the OaPV DNA in each sample. A manual threshold line was used to
discriminate between positive (blue) and negative (gray) droplets. There
were also differences in the fluorescence amplitude range of the
background (negative) droplets among the OaPV samples: 1,000--2,500 for
OaPV1 E5, 3,500--7,000 for OaPV2 L1, 500--2,700 for OaPV3 E6,
2,000--7,000 for OaPV3 E7, and 1,000--4,000 for OaPV4 E6. Therefore, the
ddPCR results could be directly converted into copies/\selectlanguage{greek}μ\selectlanguage{english}L in the initial
samples simply by multiplying them by the total volume of the reaction
mixture (22 \selectlanguage{greek}μ\selectlanguage{english}L) and then dividing that number by the volume of the DNA
sample added to the reaction mixture (7 \selectlanguage{greek}μ\selectlanguage{english}L) at the beginning of the
assay. Each sample was analyzed in duplicate. According to previous
studies on PV detection and quantification using ddPCR (De Falco et al.,
2021a; 2021b; Jeannot et al., 2016; Jeannot et al., 2021), blood samples
were considered OaPV-positive in the presence of at least three positive
droplets at the same amplitude as positive controls. A sample was
considered OaPV-negative when fewer than three droplets or no droplets
containing OaPV amplicons were observed.

\textbf{Limit of Detection (LoD) Determination}

The four OaPV genes were detected using ddPCR standard curves of the
positive controls used in the serial dilutions. A calibration curve of
the positive sample dilutions (log10) was plotted against the number of
PCR cycles. The linear range was determined by diluting the positive
controls from 10\textsuperscript{5}~to 10\textsuperscript{-1}~copies/\selectlanguage{greek}μ\selectlanguage{english}L,
detecting each dilution thrice, taking the average value, and
correlating the result with the theoretical value. The lower detection
limit obtained by ddPCR, with values \textless{}1 copies/\selectlanguage{greek}μ\selectlanguage{english}L, indicated
high sensitivity.

\textbf{qPCR}

The primers and probes used for the real-time qPCR assay are listed in
Table 1. The qPCR reaction mixture (total volume of 20 \selectlanguage{greek}µ\selectlanguage{english}L) was prepared
by adding 7 \selectlanguage{greek}µ\selectlanguage{english}L of template (100 ng genomic DNA), 10 \selectlanguage{greek}µ\selectlanguage{english}L of 2X
SsoAdvanced\textsuperscript{TM} Universal Probes Supermix (Bio-Rad
Laboratories, Hercules, CA, USA), and 1 \selectlanguage{greek}µ\selectlanguage{english}L of target probe (FAM)/primer
mix. The DNA quality and concentration were evaluated using a NanoDrop
spectrophotometer (Thermo Scientific, MA, USA). Four separate PCR were
performed using the CFX96 Real-Time System of the C1000
Touch\textsuperscript{TM} Thermal Cycler (Bio-Rad Laboratories,
Hercules, CA, USA). The thermal cycling conditions were as follows: 95\selectlanguage{ngerman}°C
for 5 min, and 40 cycles of 95°C for 15 s and 58°C for 30 s. Each sample
was analyzed in duplicate, and positive as well as negative controls
were included in all runs. Data acquisition and analysis were performed
using CFX Maestro\textsuperscript{TM} (BioRad Laboratories, Hercules,
CA, USA) software.

\textbf{PCR}

PCR was performed with DNA isolated from the blood, grass hay, and maize
silage samples using EconoTaq PLUS~ (Lucigen, WI, USA) and the OaPV
primers (see Table 1) according to the manufacturer's instructions.
Conditions for PCR were as follows: 94°C for 2 min; 35 cycles at 94°C
for 30 s, 58°C for 30 s, and 72°C for 20s; final extension at 72°C for 5
min. All the amplicons were sequenced.

\textbf{Statistical Analysis}

McNemar's test for two related binomial proportions (conditional) was
used to evaluate the agreement between two tests performed on the same
animals. To evaluate the actual differences in the prevalence of the
four types of PVs in the same animals, the Cochran-Armitage test was
performed. ~\emph{P} ~\textless{}0.05 was considered to be statistically
significant.

\textbf{RNA Extraction and Reverse Transcription-Polymerase Chain
Reaction}

Total RNA was extracted from 34 blood samples using an RNeasy Plus Mini
Kit (Qiagen, Hilden, Germany), according to the manufacturer's
instructions. Genomic DNA was removed from the RNA samples using
RNase-free DNase I (Fermentas Life Sciences, Thermo Fisher Scientific,
MA, USA). One microgram of the Total RNA was used to generate a single
strand of cDNA using the QuantiTect Reverse Transcription Kit (Qiagen
TM, Germany), according to the manufacturer's instructions. PCR was
performed on samples with and without reverse transcriptase (RT) added
to the reaction mix, using the primers reported in Table 1. The
conditions used for PCR were: 94 °C for 5 min, followed by 35 cycles at
95 °C for 30 s, 58 °C for 30 s, and 72 °C for 30 s. All the amplicons
were sequenced.

\textbf{One-Step reverse transcription (RT)-ddPCR}

Total RNA was extracted from 34 healthy cows (as negative controls) as
previously reported. 100 ng of total RNA was used for One-Step RT-ddPCR
Advanced Kit for Probes (Bio-Rad Laboratories, Hercules, CA, USA)
according to the manufacturer's instructions. The reaction was performed
in a final volume of 22 \selectlanguage{greek}μ\selectlanguage{english}L, containing 11 \selectlanguage{greek}μ\selectlanguage{english}L of ddPCR Supermix 2x for
Probes, 1 \selectlanguage{greek}μ\selectlanguage{english}L of primer and probe mix for OaPVs (Table 1), 2 \selectlanguage{greek}μ\selectlanguage{english}L reverse
transcriptase, and 1 \selectlanguage{greek}μ\selectlanguage{english}L DTT. The plate was transferred to an automated
droplet generator (AutoDG, Bio-Rad Laboratories, Hercules, CA, USA) as
described above. PCR amplification was carried out on a T100 Thermal
Cycler (Bio-Rad Laboratories Hercules, CA, USA) with the following
thermal profile: 50\selectlanguage{ngerman}°C for 60min, 95°C for 10 min, 40 cycles of 94°C for
30 s, 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, Hercules, CA, USA), and the droplets from each well of the
plate were read automatically. Therefore, the ddPCR results could be
directly converted into copies/\selectlanguage{greek}µ\selectlanguage{english}L in the initial samples simply by
multiplying them by the total volume of the reaction mixture (22 \selectlanguage{greek}µ\selectlanguage{english}L) and
then dividing the number by the volume of the RNA sample added to the
reaction mixture (5 \selectlanguage{greek}µ\selectlanguage{english}L) at the beginning of the assay. Each sample was
analyzed in duplicate.

\textbf{Sequence Analysis}

PCR products from DNA and cDNA were purified using a Qiaquick PCR
purification Kit (Qiagen TM, ME, DE) and bidirectionally sequenced using
a BigDye Terminator v1.1 Cycle Sequencing Kit (Applied Biosystems, CA,
USA) following the manufacturer's recommendations. Sequences were
removed with a DyeEx\_2.0 spin kit (Qiagen TM, DE) and run on a
SeqStudio Genetic Analyzer (Thermo Fischer Scientific, CA, USA).
Electropherograms were analyzed using Sequencing analysis v5.2 and
sequence scanner v1.0 software (Thermo Fischer Scientific, CA, USA). The
obtained sequences were compared to other sequences in GenBank using
BLAST.

\textbf{Results}

OaPV DNA was detected in 100 of 128 PBMC samples (\textasciitilde{}78\%)
examined through ddPCR. Detection and quantification of OaPVs were
performed by ddPCR. In particular, OaPV1 DNA quantification ranged from
0.22 to 34.4 copy number/\selectlanguage{greek}μ\selectlanguage{english}L; OaPV2 DNA from 0.23 to 14.92 copy
number/\selectlanguage{greek}μ\selectlanguage{english}L; OaPV3 DNA showed a range from 0.25 to 6.22 copies per \selectlanguage{greek}μ\selectlanguage{english}L, and
finally OaPV4 DNA from 0.25 to 12.03 copies per \selectlanguage{greek}μ\selectlanguage{english}L. These detailed
results are listed in Figure 1 SuppInfo.

Among 100 OaPV positive samples, single infections were detected in 42
(42\%) using ddPCR, whereas only 18 (18\%) were detected using qPCR.
Differences between the two molecular methods in detecting OaPV DNA were
statistically significant, as shown by McNemar's test (p
\textless{}0.05).

OaPV1 and OaPV2 infections were the most representative being found in
22 (52.4\%) and 16 (38\%) single infections by ddPCR. On the other hand,
qPCR revealed OaPV1 and OaPV2 infections in 9 (50\%) and 8 (44.4\%)
samples, respectively. Hence, OaPV1 and OaPV2 infections were the most
prevalent single infections as per results obtained from both the
methods. OaPV3 and OaPV4 DNAs were found in two single infections
(4.8\%) by ddPCR, while qPCR revealed only a single OaPV3 infection
(5.6\%). No OaPV4 infection was detected by qPCR. Figure 1 summarizes
the results obtained by ddPCR.

Differences in OaPV DNA genotype detection were statistically
significant, as shown by the Cochran-Armitage test (p \textless{}0.05).

Table 2 summarizes the results of the coinfection studies. Coinfections
were observed in 58 out of 100 (58\%) positive samples by ddPCR. In
particular, 35 (35\%) were dual infections, and 23 (23\%) were triple
infections. Dual coinfections by OaPV1/2 were the most frequent being
observed in 23/35 (\textasciitilde{}66\%); coinfections by OaPV1/3,
OaPV2/3, OaPV2/4, and OaPV3/4 were more rarely detected. Coinfections
with OaPV1/2/3 were the most frequent triple infections, as observed in
18/23 samples (\textasciitilde{}78\%). OaPV1/2/4 infections have also
been reported. qPCR failed to detect most of the multiple coinfections
and identified only seven dual infections. Furthermore, qPCR revealed
only one genotype in many dual and triple coinfections confirmed by
ddPCR. Triple coinfections were not observed by qPCR. However, ddPCR
revealed that OaPV1 was the most prevalent genotype in multiple
coinfections being detected in 52 of them. OaPV2, OaPV3, and OaPV4 were
detected in 51, 29, and 7 samples, respectively. In the same multiple
coinfections, qPCR revealed presence of the OaPV1, OaPV2, OaPV3, and
OaPV4 genotypes in 13, 7, 2, and 1 samples, respectively.

PCR analysis, using DNA isolated from PBMC samples, detected amplicons
using primers specific to all OaPV genotypes. Sequencing revealed the
presence of DNA fragments with 100\% identity with OaPV1 E5, OaPV2 L1,
OaPV3 E7, and OaPV4 E6 DNAs reported in GenBank (accession number:
U83594.1., U83595.1., NC\_038516.1, and KX954121.1, respectively)
(Figure 2).

One-step reverse transcription (RT)-ddPCR was performed on 34 randomized
positive samples. We detected and quantified the transcripts of OaPV1 E5
and OaPV2 L1 as well as transcripts of OaPV3 E6, E7, and OaPV4 E6, which
showed that all OaPV genotypes can be transcriptionally active in
healthy cattle. The Bio-Rad system quantified mRNA in copies per \selectlanguage{greek}μ\selectlanguage{english}L.
Samples were considered positive if they had at least three or more
positive droplets at the same amplitude as the positive control (Figure
3). Details of this investigation are reported in Figure 2 SuppInfo.

Furthermore, we performed RT-PCR analysis of RNA from the PBMC samples.
We detected amplicons, the sequencing of which showed 100\% identity
with OaPV1 E5, OaPV2 L1, OaPV3 E6 and E7, and OaPV E6 mRNAs reported in
GenBank, thus validating the one step RT-ddPCR results (Figure 4).

OaPV coinfections were most prevalent in cattle that shared grasslands
with sheep. In cattle from intensive dairy farms without any apparent
contact with sheep, double coinfections with OaPV1 and OaPV2 were also
observed (detected in 11 out of 30 examined PBMC samples). OaPV1 E5 and
OaPV2 L1 DNAs were found in feed composed of grass hay and corn silage,
which are known to be fed. ddPCR detected and quantified high copy
numbers of OaPV1 DNA as it was found in grass hay samples (from 4.2 to
7.7 copies per \selectlanguage{greek}µ\selectlanguage{english}L) as well as in feed composed of maize silage (from
3.43 to 5.7/\selectlanguage{greek}µ\selectlanguage{english}L copy number). OaPV2 DNA was also detected in grass hay
(up to 14.4 copies per \selectlanguage{greek}µ\selectlanguage{english}L) and corn silage samples (up to 10.9 copies
per \selectlanguage{greek}µ\selectlanguage{english}L) (Figure 3A SuppInfo). PCR analysis performed on all these matrix
samples revealed amplicons, and the sequencing of obtained DNA fragments
showed 100\% identity with OaPV1, and OaPV2 DNA deposited in GenBank
(Figure 3B SuppInfo).

\textbf{Discussion}

Oncogenic OaPVs infect only ovine species. Accordingly, all benign and
malignant OaPV-related tumors have been described exclusively in sheep.
OaPVs have never been associated with any pathology in other domestic
animals. This study shows, for the first time, that cross-species
transmission by both \emph{Delta} and epitheliotropic
\emph{Dyokappa}OaPVs occurs between sheep (\emph{Ovis aries} ) and
cattle (\emph{Bos taurus} ). All OaPV genotypes were detected through
both DNA detection and transcriptionally active forms in PBMC samples
from healthy cattle. Transcripts of E oncogenes as well as the L1 gene
were peculiar molecular findings of this study, which suggested that the
biological properties of OaPVs might be characterized by inducing both
abortive and productive infections in cattle.

The scant information on the epidemiology of OaPV infections is the
reason for the poor understanding of the biological significance of
these viruses in cattle. PV transmission by blood in sheep and other
species appears to be a key event in their pathology and epidemiology
(Cutarelli et al., 2021b). It has been suggested that the blood infected
with PV yields infections at permissive sites with detectable viral DNA,
RNA transcripts, and viral proteins (Cladel et al., 2019; Syrj\selectlanguage{ngerman}änen and
Syrjänen, 2021). Similar to humans, it is conceivable that ecological
factors can influence the virulence of several PVs from different
genera, and the concomitant ecological changes in different hosts linked
to the human domestication of farm animals, including sheep, may have
increased their susceptibility to OaPV cross-species transmission and/or
simply increased the frequency of physical contact to grant OaPVs
improved access to a potential new host.

PVs have a long history of co-divergence with their hosts, and hence,
these viruses are relatively more host-specific than other viruses
(Geoghegan et al., 2017). Indeed, to date, Bovine \emph{Delta} -PVs have
been the only PVs responsible for documented cases of natural
cross-species transmission leading to carcinogenetic events via
oncoproteins encoded by E genes (IARC, 2017; Roperto et al., 2013). Most
OaPV-harboring cattle in this study have in common highlands rich in
bracken ferns with sheep that live in the same geographical zones. Close
physical proximity and/or sharing of grazing lands may be a prerequisite
for PV types to cross host-species barriers, as suggested by the
detection of various BPVs in other hoofed domestic animals (de Villiers
et al., 2004; Cutarelli et al., 2021; Roperto et al., 2021). It is
conceivable that animal husbandry practices and/or mammalian sympatry
may contribute toward the cross-species transmission of OaPVs. This is
corroborated by the fact that a high prevalence of OaPVs was found in
cattle sharing large enclosures with sheep, which facilitates direct and
indirect contacts. Therefore, our study strengthens the assumption that
cross-species transmission may occur among related hosts inhabiting the
same geographic areas (Parrish et al., 2008). Successful cross-species
transmission has been suggested to occur among phylogenetically related
hosts, likely because they share fewer divergent cell receptors (Murthy
et al., 2013). Indeed, it is believed that the closer the phylogenetic
relationship between hosts, the more likely it is that a pathogen will
be able to jump between them with appropriate exposure.
Immunosuppressants of bracken coupled with more frequent exposure
between sympatric hosts may help OaPVs jump host species, resulting in
host switching.

OaPVs have also been detected in cattle from intensive dairy farms
without any apparent contact with sheep. The cows were fed grass hay and
corn silage prepared with grass and maize grown using irrigation water
from the Volturno River. PVs have recently been detected and quantified
in surface water of rivers (Iaconelli et al., 2015). It has been shown
that PVs can be detected in vegetables and irrigation water and
long-term consumption of HPV-polluted water can be associated with cell
transformation (Ghaffar et al., 2018; Itarte et al., 2021). Several PV
genotypes of different genera are known to be responsible for oral
infection; however, the information on associated risk factors is still
limited (Wong et al., 2018). It is possible that polluted irrigation
water may have played an overlooked role in the OaPV epidemiology. The
molecular findings of this study appear to strengthen our suggestions
since OaPV DNA has been detected and sequenced in hay as well as corn
silage samples. However, further studies on virus isolation from these
feed should be conducted to better understand the actual risk of virus
transmission from feed. It is well known that PVs can survive without
significant loss of infectivity during desiccation, as well as at low pH
and high temperatures, chemical features that characterize these feed
(Roden et al., 1997; Nielsen et al., 2021). Furthermore, a recent
scientific report by the European Food Safety Authority (EFSA) showed
that some viruses, including African swine fever virus (ASFV), can be
transmitted through feed based on hay and maize (Nielsen et al., 2021).

As the number of cross-species transmissions continues to rise and viral
diseases pose a continual threat to animal populations, understanding
the ecological diversity of OaPV prevalence and genotype distribution
among new host species in different geographical regions remain
essential. The need to understand how PV transmit within a given
species, as well as to new host species, has become increasingly
important as the cross-species transmission of viruses from one host
species to another is responsible for the majority of emerging
infections that can profoundly affect animal health (Geoghegan et al.,
2017).

Finally, both circulating OaPV DNA and OaPV RNA have been reported in
the blood samples of cattle found healthy by ante- and post-mortem
anatomo-clinical observations, confirming that blood represents an
important primary route of PV infection and that OaPVs can disseminate
to any organ via the bloodstream. We detected and sequenced OaPVs in the
urinary bladder of healthy cattle (personal observations), which
supports the hypothesis that OaPVs may contribute to the composition of
the normal bladder microbiota of cattle. BPV and HPV DNA have also been
reported in the blood of healthy cattle (De Falco et al., 2021a) and
asymptomatic blood donors (Vergara et al., 2019). It has been suggested
that there is an actual likelihood that HPVs could reach epithelial
target sites in the blood, which might explain how and why HPVs are
associated with tumors of several organs (Cladel et al., 2019; Syrjänen
and Syrjänen, 2021 Vergara et al., 2019; Conceição Gomes Nascimento et
al., 2021). Accordingly, PV transmission by blood represents a
conceptually novel idea, and if accepted more generally, this would
change the current thinking about the modes of PV spread, including HPV
within the host (Cladel et al., 2019; Roperto et al., 2013; Syrjänen and
Syrjänen, 2021). In this context, a naturally occurring OaPV infection
may be an additional animal model that sheds light on the PV biology,
including issues related to viral tropism in the One Health approach.

\textbf{Data Availability} All relevant data are within the manuscript
and its~\emph{Supporting information}~files.

\textbf{Funding:} This research was partially supported by Istituto
Zooprofilattico Sperimentale del Mezzogiorno. The funders of the work
did not influence study design, data collection and analysis, decision
to publish, or preparation of the manuscript.

\textbf{Conflict of interest statement:} The authors have declared that
no competing interests exist.

\textbf{Author Contributions:} Francesca De Falco and Anna Cutarelli:
Methodology; Data curation; Formal Analysis, Investigation: Bianca
Cuccaro: Formal analysis; Investigation; Cornel Catoi and Esterina De
Carlo: Validation; Visualization; Writing -- original draft; Sante
Roperto: Conceptualization, Supervision, Validation, Visualization, Data
curation; Funding Acquisition, Writing -- original draft; Writing --
review and editing

\textbf{Acknowledgments}

The authors wish to thank Dr. G. Salvatore of the Regione Basilicata,
Dr. S. Morace of the University of Catanzaro `Magna Graecia', Drs. F. Di
Domenico, E. Grieco, G. Milone, F. Di Santi from Azienda Sanitaria
Locale (ASL) of Salerno, Dr. R.N. La Rizza from ASL of Vibo Valentia, Dr
Marcellino Riccitelli from Istituto Zooprofilattico Sperimentale del
Mezzogiorno, and Dr. A. Russillo from Azienda Sanitaria of Potenza for
their technical support.

\textbf{References}

\begin{enumerate}
\tightlist
\item
  Alberti, A., Pirino, S., Pintore, F., Addis, M.F., Chessa, B.,
  Cacciotto, C., Cubeddu, T., Anfossi, A., Benenati, G., Coradduzza, E.,
  Lecis, R., Antuofermo, E., Carcangiu, L., Pittau, M. (2010). Ovis
  aries papillomavirus 3: a prototype of a novel genus in the family
  Papillomaviridae associated with ovine squamous cell carcinoma.
  Virology, 407, 352-359. doi: 10.1016/j.virol.2010.08.034.
\item
  Cladel, N.M., Jiang, P., Li, J.J., Peng, X., Cooper, T.K., Majerciak,
  V., Balogh, K.K., Meyer, T.J., Brendle, S.A., Budgeon, L.R., Shearer,
  D.A., Munden, T., Cam, M., Vallur, R., Christensen, N.D., Zheng, Z.M.,
  Hu, J. (2019). Papillomavirus can be transmitted through the blood and
  produce infections in blood recipients: Evidence from two animal
  models. Emerging Microbes \& Infections, 8, 1108-1121. doi:
  10.1080/22221751.2019.1637072.
\item
  Conceição Gomes Nascimento, K., Gonçalves Lima, E., Mota Nunes, Z.,
  Rêgo Barros Júnior, M., de Aragão Batista, M.V., Lucena-Araujo,
  A.R.,~da Costa Silva Neto, J., Simas Chagas, B., Diniz Gurgel, A.P.A.,
  de Freitas, C.A.C. (2021). Detection of human papillomavirus DNA in
  paired peripheral blood and cervix samples in patients with cervical
  lesions and healthy individuals. Journal of Clinical Medicine, 10,
  5209, doi:10.3390/jcm10215209.
\item
  Cutarelli, A., De Falco, F., Uleri, V., Buonavoglia, C., Roperto, S.
  (2021). The diagnostic value of the droplet digital PCR for the
  detection of bovine deltapapillomavirus in goats by liquid biopsy.
  Transboundary and Emerging Diseases, 68, 3624-3630. doi:
  10.1111/tbed.13971.
\item
  Darnell, G.A., Schroder, W.A., Antalis, T.M., Lambley, E., Major, L.,
  Gardner, J., Birrell, G., Cid-Arregui, A., Suhrbier, A. (2007). Human
  papillomavirus E7 requires the protease calpain to degrade the
  retinoblastoma protein. Journal of Biological Chemistry, 282,
  37492-37500. doi: 10.1074/jbc.M706860200.
\item
  De Falco, F., Corrado, F., Cutarelli, A., Leonardi, L., Roperto, S.
  (2021a). Digital droplet PCR for detection and quantification of
  circulating bovine Deltapapillomavirus. Transboundary and Emerging
  Diseases, 68, 1345-1352. doi: 10.1111/tbed.13795.
\item
  De Falco, F., Cutarelli, A., D'Alessio, N., Cerino, P., Catoi, C.,
  Roperto, S. (2021b). Molecular epidemiology of ovine papillomavirus
  infections among sheep in southern Italy. Frontiers in Veterinary
  Science, 8, 790392, doi:10.3389/fvets.2021.790392.
\item
  de Villiers, E.M., Fauquet, C., Broker, T.R., Bernard, H.U., zur
  Hausen, A. (2004). Classification of papillomaviruses. Virology,
  324,17-27. doi: 10.1016/j.virol.2004.03.033.
\item
  Garcia-Vallvé, S., Alonso, A., Bravo, I.C. (2005). Papillomaviruses:
  different genes have different histories. Trends in Microbiology, 13,
  514-521. doi: 10.1016/j.tim.2005.09.003.
\item
  Geoghegan, J.L., Duchêne, S., Holmes, E.C. (2017). Comparative
  analysis estimates the relative frequencies of co-divergence and
  cross-species transmission within viral families. PLoS Pathogens, 13,
  e106215. doi:10.1371/journal.ppat.1006215.
\item
  Ghaffar, M., Li, J., Zhang, L., Khodahemmati, S., Wang, M., Wang, Y.,
  Zhao, L., Jia, R., Chen, S., Zeng, Y. (2018). Water carcinogenicity
  and prevalence of HPV infection in esophageal cancer patients in
  Huaihe River Basin, China. Gastroenterology Research and Practice,
  2018:2028986, doi:10.1155/2018/2028986.
\item
  Gibbs, E.P., Smale, C.J., Lawman, M.J. (1975). Warts in sheep.
  Identification of a papillomavirus and transmission of infection to
  sheep. Journal of Comparative Pathology, 85, 327-334. doi:
  10.1016/0021-9975(75)90075-4.
\item
  Hayward, M.L., Baird, P.J., Meischke, H.R. (1993). Filiform viral
  squamous papillomas on sheep. The Veterinary Record, 132, 86-88. doi:
  10.1136/vr.132.4.86.
\item
  Iaconelli, M., Petricca, S., Della Libera, S., Di Bonito, P., La Rosa,
  G. (2015). First detection of human papillomaviruses and human
  polyomaviruses in River waters in Italy. Food and Environmental
  Virology, 7, 309-315. doi: 10.1007/s12560-015-9203-7.
\item
  Itarte, M., Martínez-Puchol, S., Forés, E., Hundesa, A., Timoneda, N.,
  Bofill-Mas, S., Girones, R., Rusiñol, M. (2021). NGS techniques reveal
  a high diversity of RNA viral pathogens and papillomaviruses in fresh
  produce and irrigation water. Foods, 10, 1820. doi:
  10.3390/foods10081820.
\item
  Jeannot, E., Becette, V., Campitelli, M., Calméjane, M.A.,
  Lappartient, E., Ruff, E., Saada, S., Holmes, A., Bellet, D.,
  Sastre-Garau, X. (2016). Circulating human papillomavirus DNA detected
  using droplet digital PCR in the serum of patients diagnosed with
  early-stage human papillomavirus-associated invasive carcinoma.
  Journal of Pathology and Clinical Research, 2, 201--209. doi:
  10.1002/cjp2.47.
\item
  Jeannot, E., Latouche, A., Bonneau, C., Calméjane, M.A., Beaufort, C.,
  Ruigrok-Ritstier, A., Bataillon, G., Chérif, L.L., Dupain, C., Lecerf,
  C., Popovic, M., de la Rochefordière, A., Lecuru, F., Fourchotte, V.,
  Jordanova, E.S., von der Leyen, H., Tran-Perennou, C., Legrier, M.E.,
  Dureau, S., Raizonville, L., Roufai, D.B., Le Tourneau, C., Bièche,
  I., Rouzier, R., Berns, E.M.J.J., Kamal, M., Scholl, S. (2021).
  Circulating HPV DNA as a marker for early detection of relapse in
  patients with cervical cancer. Clinical Cancer Research, 27,
  5869-5877. doi: 10.1158/1078-0432.CCR-21-0625.
\item
  Munday, J.S., Fairley, R., Lowery, I. (2020). Detection of Ovis aries
  papillomavirus type 2 DNA sequences in a sarcoid-like mass in the
  mouth of a pig. Veterinary Microbiology, 248, 108801. doi:
  10.1016/j.vetmic.2020.108801.
\item
  Munger, K., \& Howley, R.M. (2002). Human papillomavirus
  immortalization and transformation functions. Virus Research, 89,
  213-228. doi: 10.1016/s0168-1702(02)00190-9.
\item
  Murthy, S., Couacy-Hymann, E., Metzger, S., Nowak, K., De Nys, H.,
  Boesch, C., Wittig, R., Jarvis, R.A., Leendertz, F.H., Ehlers, B.
  (2013). Absence of frequent herpesvirus transmission in a nonhuman
  primate predator-prey system in the wild. Journal of Virology, 87,
  10651--10659. doi: 10.1128/JVI.01104-13.
\item
  Nielsen, S.S., Alvarez, J., Bicout, D.J., Calistri, P., Canali, E.,
  Drewe, J.A., Garin-Bastuji, B., Rojas, J.L.G., Schmidt, G.C., Herskin,
  M., Chueca, M.A,M., Michel, V., Padalino, B., Pasquali, P., Sihvonen,
  L.H., Spoolder, H., Stabl, K., Velarde, A., Viltrop, A., Winckler, C.,
  Boklund, A., Botner, A., Gervelmeyer, A., Mosbach-Schulz, O., Roberts,
  H.C. (2021). Ability of different matrices to transmit African swine
  fever virus. EFSA Journal, 19, e06558, doi: 10.2903/j.efsa.2021.6558.
\item
  Norval, M., Michie, J.R., Apps, M.V. (1985). Rumen papillomas in
  sheep. Veterinary Microbiology, 10, 219-229. doi:
  10.1016/0378-1135(85)90048-3.
\item
  Parrish, C.R., Holmes, E.C., Morens, D.M., Park, E.C., Burke, D.S.,
  Calisher, C.H., Laughlin, C.A., Saif, L.J., Daszak, P. (2008).
  Cross-species virus transmission and the emergence of new epidemic
  diseases. Microbiology and Molecular Biology Reviews, 72, 457-- 470.
  doi: 10.1128/MMBR.00004-08.
\item
  PaVE (2022, July 27). The papillomavirus episteme. Retrieved
  from:\emph{http://pave.niaid.nih.gov/.}
\item
  Roden, R.B., Lowy, D.R., Schiller, J.T. (1997). Papillomavirus is
  resistant to desiccation. The Journal of Infectious Diseases, 176,
  1076-1079. doi: 10.1086/516515.
\item
  Roperto, S., Cutarelli, A., Corrado, F., De Falco, F., Buonavoglia, C.
  (2021). Detection and quantification of bovine papillomavirus DNA by
  digital droplet PCR in sheep blood. Scientific Reports, 11, 10292,
  doi: 10.1038/s41598-021-89782-4.
\item
  Roperto, S., Russo, V., Ozkul, A., Sepici-Dincel, A., Maiolino, P.,
  Borzacchiello, G., Marcus, I., Esposito, I., Riccardi, M.G., Roperto,
  F. (2013). Bovine papillomavirus type 2 infects the urinary bladder of
  water buffalo (Bubalus bubalis) and plays a crucial role in bubaline
  urothelial carcinogenesis. ~Journal of General Virology, 94, 403--408.
  doi: 10.1099/vir.0.047662-0.
\item
  Scarth, J.A., Patterson, M.R., Morgan, E.L., Macdonald, A. (2021). The
  human papillomavirus oncoproteins: a review of the host pathways
  targeted on the road to transformation. Journal of General Virology,
  102, 001540. doi: 10.1099/jgv.0.001540.
\item
  Sichero, L., Rollison, D.A., Amorrortu, R.P., Tommasino, M. (2019).
  Beta human papillomavirus and associated diseases. Acta Cytologica,
  63, 100-108. doi: 10.1159/000492659.
\item
  Strickley, J.D., Messerschmidt, J.L., Awad, M.E., Li, T., Hasegawa,
  T., Ha, D.T., Nabeta, H.W., Bevins, P.A., Ngo, K,H., Asgari, M.M.,
  Nazaryan, R.M., Neel, V.A., Bennett Jenson, A., Joh, J., Demehri, S.
  (2019). Immunity to commensal papillomaviruses protects against skin
  cancers. Nature, 575, 519-522. doi: 10.1038/s41586-019-1719-9.
\item
  Syrjänen, S., \& Syrjänen, K. (2021). HPV-associated benign squamous
  cell papillomas in the upper aero-digestive tract and their malignant
  potential. Viruses, 13, 1624, doi: 10.3390/v13081624.
\item
  Tilbrook, P.A., Sterrett, G., Kulski, J.K. (1992). Detection of
  papillomaviral-like DNA sequences in premalignant and malignant
  perineal lesions of sheep. Veterinary Microbiology, 31, 327-341. doi:
  10.1016/0378-1135(92)90125-d.
\item
  Tore, G., Cacciotto, C., Anfossi, A.G., Dore, G.M., Antuofermo, E.,
  Scagliarini, A., Burrai, G.P., Pau, S., Zedda, M.T., Masala, G.,
  Pittau, M., Alberti, A. (2017). Host cell tropism, genome
  characterization, and evolutionary features of OaPV4, a novel
  Deltapapillomavirus identified in sheep fibropapilloma. Veterinary
  Microbiology, 204, 151-158. doi: 10.1016/j.vetmic.2017.04.024.
\item
  Tore, G., Dore, G.M., Cacciotto, C., Accardi, R., Anfossi, A.G.,
  Bogliolo, L., Pittau, M., Pirino, S., Cubeddu, T., Tommasino, M.,
  Alberti, A. (2019). Transforming properties of ovine papillomaviruses
  E6 and E7 oncogenes. Veterinary Microbiology, 230, 14-22. doi:
  10.1016/j.vetmic.2019.01.010.
\item
  Trenfield, K., Spradbrow, P.B., Vanselow, B.A. (1990). Detection of
  papillomavirus DNA in precancerous lesions of the ears of sheep.
  Veterinary Microbiology, 25\textbf{,} 103-116. doi:
  10.1016/0378-1135(90)90070-c.
\item
  Uzal, F.A., Latorraca, A., Ghoddusi, M., Horn, M., Adamson, M., Kelly,
  W.R., Schenkel, R. (2000). An apparent outbreak of cutaneous
  papillomatosis in merino sheep in Patagonia, Argentina. Veterinary
  Research Communications, 24, 197-202. doi: 10.1023/a:1006460432270.
\item
  Van Doorslaer, K. (2013). Evolution of the
  \emph{Papillomaviridae.}Virology, 445\textbf{,} 11-20. doi:
  10.1016/j.virol.2013.05.012.
\item
  Vanselow, B.A., Spradbrow, P.B., Jackson, A.R. (1982).
  Papillomaviruses, papillomas and squamous cell carcinomas in sheep.
  The Veterinary Record, 110, 561-562. doi: 10.1136/vr.110.24.561.
\item
  Vergara, N., Balanda, M., Vidal, D., Roldán, F., Martin, H.S.,
  Ramírez, E. (2019). Detection and quantification of human
  papillomavirus DNA in peripheral blood mononuclear cells from blood
  donors. Journal of Medical Virology, 91, 2009-2015. doi:
  10.1002/jmv.25551.
\item
  Willemsen, A., van den Boom, A., Dietz, J., Dagalp, S.B., Dogan, F.,
  Bravo, I.G., Ehrhardt, A., Ehrke-Schulz, E. (2020). Genomic and
  phylogenetic characterization of ChPV2, a novel goat PV closely
  related to the \emph{Xi-PV1} species infecting bovines. Virology
  Journal, 17, 167, doi:10.1186/s12985-020-01440-9.
\item
  Vitiello, V., Burrai, G.P., Anfossi, A.M., Alberti, A., Antuofermo,
  E., Rocca, S., Cubeddu, T., Pirino, S. (2017). Ovis aries
  papillomavirus 3 in ovine cutaneous squamous cell carcinoma.
  Veterinary Pathology, 54, 775-782. doi: 10.1177/0300985817705171.
\item
  Wong, M.C.S., Viantis, A.C., Liang, M., Wong, P.Y., Ho, W.C.S., Boon,
  S.S., Sze, R.K.H., Leung, C., Chan, P.K.S., Chen, Z. (2018).
  Prevalence and epidemiologic profile of oral infection with Alpha,
  Beta, and Gamma papillomaviruses in an Asian Chinese population.
  Journal of Infectious Diseases, 218, 388-397. doi:
  10.1093/infdis/jiy160.
\item
  World Health Organization -- International Agency for Research on
  Cancer Monographs on the Evaluation of Carcinogenic Risks to Humans,
  vol 90 -- Human Papillomaviruses, Lyon, France, 2007.
\end{enumerate}

\textbf{Figure legends}

\textbf{Figure 1.} Detection rates of OaPV single infections.
Differences were significant after Cochran-Armitage test (p
\textless{}0.05).

\textbf{Figure 2.} (A) Electrophoresis of PCR products for evaluating
OaPV1 E5, OaPV2 L1, OaPV3 E7, and OaPV4 E6 DNA in bovine PBMC samples.
MW: DNA molecular weight marker (100 bp). C: PCR negative control. (B),
(C), (D), and (E):100\% identity between the sequences of the amplicons
OaPV1 Seq, OaPV2 Seq, OaPV3 Seq, and OaPV4 Seq, and the sequences
reported in GenBank, respectively (accession number: U83594.1; U83595.1;
NC\_038516.1; and KX954121.1).

\textbf{Figure 3.} One Step ddPCR. The rain plots for the OaPV1 E5,
OaPV2 L1, OaPV3 E6 and E7, and OaPV4 E6 are shown. Representative
samples have positive droplets at the same amplitude as the positive
control for each OaPVs.

\textbf{Figure 4.} (A) Electrophoresis of RT-PCR products to evaluate
OaPV1 E5, OaPV2 L1, OaPV3 E6 and E7, and OaPV4 E6 mRNA expression in
PBMC samples. MW: DNA molecular weight marker (100bp); C+: PCR positive
control. PCR was performed on PBMC samples with (RT+) and without (RT-)
the addition of reverse transcriptase to the reaction mixture, using the
same amount of RNA. (B) The amplicon sequences showed 100\% identity
with the respective sequences reported in S1 Fig. Furthermore, 100\%
identity between the OaPV3 E6 cDNA amplicon and the OaPV3 E6 sequence
reported in GenBank (accession number: NC\_038516.1) is shown.

\textbf{Figure 1 SuppInfo.} Detection and quantification of OaPV DNA
using ddPCR. The Bio Rad system quantified DNA copies per \selectlanguage{greek}μ\selectlanguage{english}L. Bov =
bovine; N = negative

\textbf{Figure 2 SuppInfo.} Detection and quantification of mRNA using
one-step reverse transcription (RT)-ddPCR. The Bio Rad system quantified
mRNA copies per \selectlanguage{greek}μ\selectlanguage{english}L. Bov = bovine; N = negative

\textbf{Figure 3 SuppInfo. (A)} . Representative positive samples and a
positive control for the OaPV1 E5 and OaPV2 L1 assessed by ddPCR in
grass hay and maize silage samples. Positive samples have droplets at
the same amplitude as the positive control. Amplitude of OaPV1 E5 ranged
from 1.500 to 5.500; amplitude of OaPV2 L1 ranged from 1.500 to 11.500.
(B). PCR products of the above samples. MW: DNA molecular weight marker
(100bp). C+: PCR positive control; C-: PCR negative control. The
amplicon sequences showed 100\% identity with the respective sequences
reported in GenBank (see Figure 2).

Table 1 shows the primers and probes used for the detection and
quantification of OaPVs.

Table 2- Genotype coinfections evaluated by ddPCR with number and
related percentages of their combination are shown.\selectlanguage{english}
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\textbf{Hosted file}

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\textbf{Hosted file}

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