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Respiratory disease in ball pythons ( Python regius ) experimentally infected with ball python nidovirus


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Circumstantial evidence has linked a new group of nidoviruses with respiratory disease in pythons, lizards, and cattle. We conducted experimental infections in ball pythons (Python regius) to test the hypothesis that ball python nidovirus (BPNV) infection results in respiratory disease. Three ball pythons were inoculated orally and intratracheally with cell culture isolated BPNV and two were sham inoculated. Antemortem choanal, oroesophageal, and cloacal swabs and postmortem tissues of infected snakes were positive for viral RNA, protein, and infectious virus by qRT-PCR, immunohistochemistry, western blot and virus isolation. Clinical signs included oral mucosal reddening, abundant mucus secretions, open-mouthed breathing, and anorexia. Histologic lesions included chronic-active mucinous rhinitis, stomatitis, tracheitis, esophagitis and proliferative interstitial pneumonia. Control snakes remained negative and free of clinical signs throughout the experiment. Our findings establish a causal relationship between nidovirus infection and respiratory disease in ball pythons and shed light on disease progression and transmission.
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Respiratory disease in ball pythons (Python regius) experimentally infected
with ball python nidovirus
Laura L. Hoon-Hanks
, Marylee L. Layton
, Robert J. Ossibo
, John S.L. Parker
Edward J. Dubovi
, Mark D. Stenglein
Department of Microbiology, Immunology, and Pathology, College of Veterinary Medicine and Biomedical Sciences, Colorado State University, Fort Collins, CO, USA
Department of Population Medicine and Diagnostic Sciences, College of Veterinary Medicine, Cornell University, Ithaca, NY, USA
Baker Institute for Animal Health, College of Veterinary Medicine, Cornell University, Ithaca, NY, USA
Ball python
Experimental infection
Respiratory disease
Koch's postulates
Circumstantial evidence has linked a new group of nidoviruses with respiratory disease in pythons, lizards, and
cattle. We conducted experimental infections in ball pythons (Python regius) to test the hypothesis that ball
python nidovirus (BPNV) infection results in respiratory disease. Three ball pythons were inoculated orally and
intratracheally with cell culture isolated BPNV and two were sham inoculated. Antemortem choanal, or-
oesophageal, and cloacal swabs and postmortem tissues of infected snakes were positive for viral RNA, protein,
and infectious virus by qRT-PCR, immunohistochemistry, western blot and virus isolation. Clinical signs in-
cluded oral mucosal reddening, abundant mucus secretions, open-mouthed breathing, and anorexia. Histologic
lesions included chronic-active mucinous rhinitis, stomatitis, tracheitis, esophagitis and proliferative interstitial
pneumonia. Control snakes remained negative and free of clinical signs throughout the experiment. Our ndings
establish a causal relationship between nidovirus infection and respiratory disease in ball pythons and shed light
on disease progression and transmission.
1. Importance
Over the past several years, nidovirus infection has been circum-
stantially linked to fatal respiratory disease in multiple python species,
but a causal relationship has not been denitively established. Through
experimental infections, our study fullls Koch's postulates and con-
rms ball python nidovirus as a primary respiratory pathogen in this
species. Our ndings will provide veterinarians valuable information
for the diagnosis and management of this disease and lay the ground-
work for continued scientic investigation of this sometimes fatal dis-
ease. Python nidoviruses are members of a growing group of viruses
that have been associated with severe respiratory disease, including
bovine nidovirus and shingleback lizard nidovirus. The establishment of
BPNV as a primary pathogen in pythons is an important step in un-
derstanding the pathogenic potential of this emerging group of viruses.
2. Introduction
The nidoviruses (order Nidovirales) are a large and diverse group of
viruses that includes notable human and veterinary pathogens (De
Groot et al., 2012; Graham et al., 2013; Lauber et al., 2012; Masters and
Perlman, 2013; Snijder et al., 2013; Snijder and Kikkert, 2013). The
discovery of a group of related nidoviruses in snakes, lizards, cattle, and
nematodes has recently expanded the order (Bodewes et al., 2014;
Dervas et al., 2017; Marschang and Kolesnik, 2017; ODea et al., 2016;
Shi et al., 2016; Stenglein et al., 2014; Tokarz et al., 2015; Uccellini
et al., 2014). These novel nidoviruses cluster most closely with viruses
in the subfamily Torovirinae within the Coronaviridae family of the Ni-
dovirales order, and form a distinct clade from viruses in the Banivirus
and Torovirus genuses, which infect ray-nned sh and mammals, re-
spectively. Based on phylogenetic analysis, it has been proposed that
the reptile nidoviruses be classied within a distinct genus named
Barnivirus, and that Torovirinae be classied as its own family due to the
growing evidence of the paraphyly of Coronaviridae, though these
viruses have not yet been formally classied (Adams et al., 2017; Batts
et al., 2012; Gonzalez et al., 2003; Nga et al., 2011; Stenglein et al.,
2014). Toroviruses share similar tissue tropisms of the gastrointestinal
(GI) and respiratory epithelium, ultrastructural features, and genome
Received 29 September 2017; Received in revised form 1 December 2017; Accepted 11 December 2017
Correspondence to: Colorado State University, 200 W Lake St. 2025 Campus Delivery, Fort Collins, CO 80523, USA.
Co-corresponding author.
Current address: Department of Comparative, Diagnostic, and Population Medicine, College of Veterinary Medicine, University of Florida, Gainesville, FL, USA.
E-mail addresses: (L.L. Hoon-Hanks), (M.D. Stenglein).
Virology 517 (2018) 77–87
Available online 10 January 2018
0042-6822/ © 2017 The Authors. Published by Elsevier Inc. This is an open access article under the CC BY license (
organization (Batts et al., 2012; Pradesh et al., 2014; Schutze et al.,
2006), and represent a group of emerging pathogens of unknown, and
possibly underestimated, signicance in veterinary and human medi-
The snake-associated nidoviruses were rst discovered in ball py-
thons (Python regius) and Indian rock pythons (P. molurus) with severe
respiratory disease that had tested negative for known snake respiratory
pathogens (Bodewes et al., 2014; Stenglein et al., 2014; Uccellini et al.,
2014). Postmortem ndings in sick pythons included stomatitis, sinu-
sitis, pharyngitis, tracheitis, esophagitis, and proliferative pneumonia
with signicant mucus secretion in aected tissues; secondary bacterial
infections were also noted within the respiratory tract or systemically in
some snakes. In 2017, similar ndings were detected in green tree
pythons (Morelia [M.]viridis) infected by a related nidovirus (Morelia
viridis nidovirus) (Dervas et al., 2017). Additionally, nidoviruses have
been detected in antemortem oral swabs (or rarely blood) from a Bur-
mese python (P. bivittatus), ball pythons, Indian rock pythons, green
tree pythons, a carpet python (M. spilota), and boa constrictors (Boa
constrictor) with or without documented respiratory signs (Marschang
and Kolesnik, 2017). Related nidoviruses associated with respiratory
disease in wild shingleback lizards and cattle have also been recently
described (ODea et al., 2016; Tokarz et al., 2015).
Reports of nidovirus in multiple python species are highly sugges-
tive of, but do not denitively establish, a causal relationship between
viral infection and respiratory disease. This study sought to fulll
Koch's postulates through experimental infections of ball pythons with
ball python nidovirus (BPNV). The goal was to conclusively establish a
causative relationship between infection and respiratory disease as well
as further characterize the clinical course of disease, describe useful
diagnostic techniques, and to investigate possible routes of transmis-
3. Materials and methods
3.1. Generation of a diamond python cell line
A non-immortalized cell line was generated from heart tissue col-
lected from a diamond python (Morelia spilota). Multiple ~ 2 mm cubes
of myocardium were collected from a diamond python directly fol-
lowing humane barbiturate overdose euthanasia for chronic vertebral
disease. Tissues were collected within 2 h of euthanasia and placed in
1.5 ml, ice-cold, sterile phosphate buered saline (PBS) in 2 ml mi-
crocentrifuge tubes for transport to the laboratory. Tissue samples were
individually transferred to a 6-well cell culture plate (Corning), washed
three times with ice cold PBS, and manually minced with a sterile
scalpel blade in 1.5 ml PBS with 0.25% trypsin (Gibco) and 1 mM
ethylenediaminetetraacetic acid (EDTA). Samples were incubated at
37 °C with gentle agitation every 20 min (m) for a total of 60 m.
Following incubation, 0.5 ml of the digested product was added per
well of a 12-well cell culture plate (Corning) along with 2 ml of com-
plete cell growth medium [Minimum Essential Medium with Earle's
Balanced Salts, L-Glutamine, and Nonessential Amino Acids (MEM/
EBSS; Hyclone); 10% irradiated fetal bovine serum (FBS; HyClone); 100
U penicillin; 100 μg streptomycin; 0.25 μg amphotericin B (Cellgro);
and 50 μg gentamicin (Cellgro)] and placed at 30 °C in a humidied 5%
atmosphere. Wells were monitored regularly for evidence of cell
adherence and replication. Partial (~ 50%) medium changes were
performed weekly. When cell monolayers reached ~ 70% conuence,
monolayers were washed twice with room temperature sterile PBS, 1 ml
enzyme free cell dissociation buer (Gibco) was added to each well,
and the samples were incubated for 5 m at 30 °C. Cell monolayers were
disrupted by gently pipetting samples up and down, and the cell/dis-
sociation buer mixture was transferred to a 60 mm tissue culture dish
(Corning) with 7 ml of complete cell growth medium and returned to a
30 °C, humidied, 5% CO
atmosphere. The cells were monitored reg-
ularly for evidence of cellular replication with weekly, partial (~ 50%)
medium changes. At ~ 70% conuence, monolayers were passed using
0.25% trypsin rst into T25, and then into T75 tissue culture asks
(Corning). At 100% conuence, T75 asks were trypsinized, washed in
complete cell growth medium, and resuspended in 1 ml of complete cell
growth medium with 20% irradiated FBS and 10% DMSO for storage in
liquid nitrogen in 1.2 ml cryovials (Corning).
3.2. Isolation of BPNV
Oral swabs were collected from a ball python with upper respiratory
disease that was part of a colony with a documented history of BPNV
infections (Uccellini et al., 2014). Swabs were placed in 1.5 ml of viral
transport medium (MEM/EBSS, 0.5% bovine serum albumin, 200 U
penicillin, 200 μg streptomycin, 0.25 μg fungizone, and 10 μg cipro-
oxacin; Gibco) prior to inoculation on diamond python heart (DPHt)
cells. Briey, 1 ml of the swab extracts were added to DPHt cells in T25
culture asks. After a 3 h incubation at 30 °C, monolayers were rinsed
and cell growth medium added (MEM/EBSS, 10% irradiated FBS, 200 U
penicillin, 200 μg streptomycin, 0.25 μg fungizone, and 10 μg cipro-
oxacin; Gibco). Cultures were maintained at 30 °C and monitored
daily for cytopathic eects. At 7 days post inoculation cells were frozen
at 70 °C, thawed, and were re-inoculated onto new DPHt mono-
layers. The study challenge virus (deemed BPNV-148) was a passage 2
3.3. Plaque assay
DPHt cells were incubated in complete cell medium [MEM/EBSS
(HyClone), 10% irradiated FBS (HyClone), 10% Nu-Serum1 (Corning),
and 2x penicillin-streptomycin solution (HyClone)] in a 6-well
CELLSTAR cell culture plate (Greiner Bio-one) at 30 °C in 5% CO
90% conuence was attained. BPNV-148 stock was diluted in serum-
free MEM/EBSS to generate 5 dilutions of 1 × 10
through 1 × 10
For cell inoculation, all medium was removed and 900 μl of each di-
lution was placed on the cells, with serum-free MEM/EBSS added to the
last well as a negative control. Cells were incubated at 30 °C in 5% CO
for 1 h, after which the infected medium was removed and an agarose
overlay was placed [complete cell medium with 0.8% UltraPure LMP
Agarose (Invitrogen)]. Assays were incubated at 30 °C in 5% CO
for 6
days, at which time 1 ml of 4% paraformaldehyde (EM grade; Electron
Microscopy Sciences) in DPBS (Corning) was added to each well and
incubated for an additional hour. The agarose overlay was removed,
cells were rinsed with DPBS, and an additional 1 ml of paraformalde-
hyde mixture was added. Cells were placed at 4 °C overnight. The for-
maldehyde was removed, cells were rinsed with sterile water, and 100
μl of crystal violet (0.5% crystal violet in 25% methanol and 75% sterile
water) was added and incubated for 10 min at room temperature.
Crystal violet was rinsed owith sterile water, assays were dried, and
plaques were counted. Plaque assays were also performed using sam-
ples collected during experimental infection studies; the same protocol
was utilized.
3.4. Experimental infection
Five captive-bred ball pythons (BP A-E; 4 males and one un-
determined sex) were acquired, each approximately 6 weeks old and
varying in size from 77 to 106 g. All pythons were housed and treated
according to the IACUC protocol (156063A) and Colorado State
University Laboratory Animal Resources standards. Infected snakes
were housed in a cubicle with separate HEPA-ltered air supply from
control snakes and all snakes were housed in separate cages without
direct contact. Uninfected snakes were always handled prior to infected
snakes to prevent fomite transmission. Physical exams were performed
and all snakes were deemed clinically healthy at the time of acquisition.
Pre-infection choanal (CHS), oroesophageal (OES), and cloacal swabs
(CLS) were collected and tested by qRT-PCR (see below) for BPNV. One
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
week after arrival, three snakes were inoculated with BPNV infected
DPHt cell culture supernatant discussed above (BP-A, B, and C) and two
were sham inoculated (BP-D and E). Inoculation was performed both
orally (200 μl) and intratracheally (100 μl) for each snake with 1.1 ×
10^5 PFU in 300 μl for the infected snakes and a similar volume of
uninfected cell culture medium for the control snakes. Snakes were
monitored daily, weights were taken weekly, and CHS, OES, and CLS
were collected weekly from all snakes using PurFlock Ultra sterile
ocked 6plastic-handle swabs (Puritan Diagnostics). Swabs were
placed in 2 ml Bacto brain-heart infusion medium (Becton, Dickinson
and Company), incubated at room temperature (RT) for approximately
30 min, vortexed, and then stored at 80 °C. BP-C was euthanized at 5
weeks post infection (PI) as a demonstration of early infection. BP-A
was euthanized at 10 weeks PI and BP-B at 12 weeks PI based on
clinical signs and established euthanasia criteria. BP-D and E were eu-
thanized at 12 weeks to end the study. Final CHS, OES, and CLS and
culture swabs of the oral cavity (BBL CultureSwab plus Amies gel
without charcoal; Becton, Dickinson and Company) were collected at
the time of euthanasia. Sections of the glottis, nasal and oral cavity,
cranial, middle, and caudal trachea and esophagus, lungs, heart, liver,
kidneys, gallbladder, spleen, pancreas, stomach, small intestine, colon,
feces, blood, urates, gonads, head and vertebrae with brain and spinal
cord were saved fresh and/or placed in 10% neutral buered formalin.
3.5. RNA extraction
RNA from swabs and fresh-frozen tissues (lung, cranial trachea/
esophagus, liver, kidney, heart, stomach, small intestine, colon, feces,
urates) was extracted using a combination of TRIzol (tissue; Ambion
Life Technologies) or TRIzol LS (swabs in BHI; Ambion Life
Technologies) with RNA clean and concentrator columns (CC-5; Zymo
Research). Approximately 100 mg of tissue was added to 1 ml of TRIzol
and 250 μl of BHI swab medium was added to 750 μl of TRIzol LS and
incubated at room temperature (RT) for 5 min. Tissue samples were
macerated using a single sterile metal BB shaken in a TissueLyzer
(Qiagen) at 30 Hz for 3 min. Then, 200 μl of chloroform (Sigma-
Aldrich) was added, shaken for 15 s by hand, and incubated at RT for
2 min. Samples were spun at 12,000 RPM for 10 min at RT. The aqu-
eous phase was removed (approximately 450 μl) and was added to a
mixture of 450 μl of RNA binding buer (CC-5; Zymo Research) and
450 μl of 100% ethanol (EtOH). This was added to an RNA clean and
concentrator column (CC-5; Zymo Research). The interphase and or-
ganic phase were discarded. The RNA column was washed with 400 μl
RNA wash buer and then incubated with 6 U DNase enzyme (NEB), 1x
DNase buer (NEB), and RNA wash buer for 15 min. The column was
spun to remove DNase mixture and then washed with 400 μl RNA prep
buer. An additional wash with 800 μl RNA wash buer was per-
formed, the column was dried with a 1 min high-speed spin, and then
RNA samples were eluted in 30 μl of RNase-free water.
3.6. Viral RNA detection
RNA extracted from swabs and fresh-frozen tissues was reverse
transcribed into cDNA as follows. Five microliters of RNA were added
to 200 pmol of a random pentadecamer oligonucleotide (MDS-911;
Table 1) and incubated for 5 min at 37 °C; a water template control was
also used. Reverse transcription reaction mixture containing 1x Su-
perScript III FS reaction buer (Invitrogen), 5 mM dithiothreitol (In-
vitrogen), 1 mM each deoxynucleoside triphosphates (dNTPs), and 100
U SuperScript III reverse transcriptase enzyme (Invitrogen) was added
to the RNA-oligomer mix (12 μl total reaction volume) and incubated
for 30 min at 42 °C, then 30 min at 50 °C, then 15 min at 70 °C.
Quantitative reverse transcription polymerase chain reaction (qRT-
PCR) was performed using 1x HOT FIREPol DNA Polymerase (Solis
BioDyne), 3 μM of each degenerate nidovirus primer (MDS-918 and
MDS-919; Table 1), and 5 μl of diluted (1:10) cDNA in a 30 μl reaction.
Reaction mixtures were placed in a TempPlate semi-skirted 96-well
PCR plate and were run in a Roche LightCycler 480 II with the fol-
lowing cycle parameters: 95 °C for 15 min; 95 °C for 10 s, 60 °C for 12 s,
and 72 °C for 12 s with 40 cycles; and a melting curve. All samples were
run in duplicate, Ct values were averaged and standard deviations were
calculated. The PCR reaction eciency for each primer-pair was mea-
sured using a dilution series of positive samples (BP-B terminal OES for
nidovirus primers and BP-B trachea/esophagus for GAPDH primers);
the dilution series samples were run in duplicate. Relative viral RNA for
all CHS, OES, and CLS samples was determined by comparison of each
sample Ct to the sample with the highest Ct (lowest viral RNA) at the
rst collection time point following inoculation (BP-B OES at week 1
PI). Relative viral RNA from tissues was determined by normalization to
snake GAPDH within each sample (same qRT-PCR conditions with
MDS-921and MDS-923 primers; Table 1).
3.7. Antibody development
The predicted amino acid (aa) sequence for the ball python nido-
virus 1 nucleocapsid protein (152 aa protein; GenBank: AIJ50569.1)
and nidovirus nucleocapsid protein sequences isolated from green tree
pythons (unpublished data) were used by our lab to identify a relatively
conserved peptide sequence with predicted high immunogenicity and
epitope exposure. The peptide (aa 136152 of the N protein of a green
tree python nidoviral isolate: Cys-RAFIPLKHEGAETEEEV) was sub-
mitted to Pacic Immunology (Ramona, CA) for synthesis and poly-
clonal anti-nidoviral nucleocapsid antisera (NdvNcAb) was developed
in two rabbits.
3.8. Histopathology/immunohistochemistry
Formalin-xed tissue was paran-embedded and 5 µm sections
were stained by hematoxylin and eosin (H&E), Gram, periodic acid-
Schi(PAS), and Ziehl-Neelsen acid fast for light microscopy (per-
formed by Colorado State University Veterinary Diagnostic Laboratory;
CSUVDL). Immunohistochemistry was also performed by CSUVDL using
the Bond Polymer Redene Red Detection kit (Leica) and a 10 min
incubation with Epitope Retrieval Solution 1 (Leica). NdvNcAb
(0.32 μg/ml) was used as the primary antibody and the slides were
counter stained with hematoxylin. Lung tissue from a green tree python
that was nidovirus positive (PCR and virus isolation) and that died of
respiratory disease was used as a positive control (data not shown).
Table 1
Primers. List of primers used during qRT-PCR for detection of BPNV or GAPDH and used for sequencing library generation. Forward (F); Reverse (R); N/A (not applicable).
Primer name Target Sequence (5-3) Direction (F/R) Reference
MDS-143 Sequencing library adaptors CAAGCAGAAGACGGCATACG F (Runckel et al., 2011)
MDS-921 Python GAPDH gene AATATCTGCCCCATCAGCTG R (Stenglein et al., 2017)
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
3.9. Virus isolation and immunouorescence
Oroesophageal swabs collected at the time of euthanasia from all
infected and uninfected snakes were ltered (Merck Millipore
UltraFree-MC 0.22 µm centrifugal lters) and 40 μl was inoculated onto
DPHt cells at 80% conuence in 35 mm diameter glass-bottom plates
(MatTek corporation). Cells were maintained in 2 ml of complete cell
medium and incubated at 30 °C with 5% CO
; medium was refreshed
every other day. Cells infected with BP-A OES were formalin-xed as
previously described at 1, 12, 24, 48, 96, 144, and 192 h PI; all other
OES-infected cells (BP-B, C, D, and E) were formalin-xed at 4 days PI.
Approximately 50 mg of lung or feces from infected and uninfected
pythons was homogenized in 500 μl of DPBS, claried, and then ltered
(0.22 µm). Infection of cell culture was as previously described. Lung-
infected cells were formalin-xed at 10 days PI and fecal-infected cells
at 3 days PI.
Fixed cells were washed 3 times with 1 ml of PBS. Cells were per-
meabilized in 0.1% Triton X-100 (reagent grade; Amresco) in PBS for
5 min. Washes were repeated and then cells were incubated in blocking
buer (1% bovine serum albumin (Fisher Scientic) in PBS) for 1 h. A
1:2000 dilution of NdvNcAb rabbit serum (primary antibody) was
added to the blocking buer and incubated for an additional hour.
Wash steps were repeated and then new blocking buer with 5 μg/ml of
secondary antibody (Alexa Fluor 488 goat anti-rabbit IgG antibodies;
A11008 Life Technologies) was added and incubated for 1 h. Wash
steps were repeated and then cells were stained with Hoechst 33342
(1 μg/ml nal concentration; Life Technologies) to stain DNA. Cells
were imaged on an Olympus IX81 motorized inverted system confocal
microscope with FluoView 4.2 software. Images were processed in
Adobe Photoshop CC (2017) and both infected and uninfected were
processed equally.
3.10. Western blot
DPHt cells inoculated with OES from all infected and uninfected
snakes, as previously described, as well as a sham inoculated control
(BHI only) were harvested at 4 days PI. Cells were lysed using equal
volumes of sample and SDS-based tissue lysis buer (40 mM TrisCl pH
7.6, 120 mM NaCl, 0.5% Triton X-100, 0.3% SDS, Roche complete
protease inhibitor cocktail tablet), mixed for 30 min at 4 °C, and clar-
ied by centrifugation at 4 °C for 10 min at 10,000 rpm. Twelve mi-
croliters of sample or 4 μl of ladder with 8 μl of PBS (precision plus
protein western C; BioRad) were combined with 1x NuPage LDS sample
buer (Life Technologies) and separated using a 412% polyacrylamide
gel (Invitrogen). Protein was transferred to a nitrocellulose membrane
using a Trans-Blot turbo (low molecular weight protein transfer; Bio-
Rad). A 1 h incubation of the membrane in blocking buer [1x PBS,
0.05% Tween20, 1% Carnation nonfat dry milk, and 1:1000 Kathon
CG/ICP preservative (Dow Chemical)] was followed by a 1 h incubation
with 1.6 μg/ml NdvNcAb in blocking buer. The membrane was wa-
shed (1x PBS and 0.05% Tween20) 3 times for 5 min each followed by a
1 h incubation with a 1:50,000 dilution of goat anti-rabbit IgG antibody
conjugated to horseradish peroxidase (HrP; Pierce 31460 Invitrogen)
and 1:4000 dilution of streptactin-HrP (ladder) in blocking buer. A
second wash was performed and the blot was developed using a 5 min
incubation with clarity western ECL substrate (BioRad). Imaging was
via chemiluminescence for 60 s (BioRad Gel Doc).
3.11. Metagenomic sequencing
Shotgun libraries were generated from total RNA extracted from BP-
A, B, C, D, and E lung and cranial trachea/esophagus and BPNV-148
inoculum. Library preparation was as follows: Ten microliters of un-
diluted cDNA (see polymerase chain reaction for cDNA preparation)
was treated with 1 U RNase H (NEB) diluted in 5 μl 1x SuperScript III FS
reaction buer and 160 pmol MDS-911 to degrade RNA templates.
Samples were incubated at 37 °C for 20 min followed by 94 °C for
2 min. Then, single-stranded cDNA was converted to double-stranded
DNA by adding 2.5 U Klenow DNA polymerase (3to 5exo- NEB) in
5μl 1x SuperScript III FS reaction buer and 2 mM each dNTPs and
incubated at 37 °C for 15 min. DNA was puried using SPRI beads at a
1.4:1 bead/DNA volume ratio. DNA was eluted in 20 μl molecular grade
water (Sigma-Aldrich). The dsDNA concentration from each sample was
measured uorometrically and 10 ng was used as a template in 6.5 μlof
1x Tagment DNA buer and 0.5 μl Nextera Tagment DNA enzyme
(Illumina). The mixture was incubated at 55 °C for 10 min and then
placed directly on ice. Tagmented DNA was cleaned with SPRI beads
and used as a template (5.8 μl) in the addition of full-length adaptors
with unique bar-code combinations by PCR. The 25 μl PCR reaction
contained 1x Kapa real-time library amplication master mix (Kapa
Biosystems), 0.33 μM (each) MDS-143 and MDS-445 primers (Table 1),
and 0.020 μM each of adaptor 1 and 2 bar-coded primers (Stenglein
et al., 2015). Thermocycling conditions in consecutive order were 72 °C
for 3 min, 98 °C for 30 s, and 8 cycles of 98 °C for 10 s, 63 °C for 30 s,
and 72 °C for 3 min. Relative concentrations of libraries were measured
in qRT-PCR reactions containing 1x qRT-PCR master mix [10 mM Tris-
HCl pH 8.6, 50 mM KCl, 1.5 mM MgCl
, 0.2 mM of each dNTP, 5%
glycerol, 0.08% NP-40, 0.05% Tween-20, 1x Sybr green (Life Tech-
nologies) and 0.5 U Taq polymerase] and 0.5 μM MDS-143 and MDS-
445 primers. Equivalent amounts of DNA from each sample were
pooled and then cleaned using SPRI beads. The pooled libraries were
run on a 2% agarose gel and size selected (400500 nucleotides) by gel
extraction with a gel DNA recovery kit (Zymo Research) according to
the manufacturer's protocol. Size-selected pooled libraries were am-
plied once more in a PCR mixture containing 1x Kapa real-time library
amplication mix, 500 pmol of MDS-143 and -445 each, and 5 μlof
library template in a 50 μl total reaction volume. This PCR also included
single reactions of 4 separate uorometric standards (Kapa). Thermo-
cycler conditions were 98 °C for 45 s and 14 cycles of 98 °C for 10 s,
63 °C for 30 s, and 72 °C for 2 min, which was the cycle at which the
sample curve passed standard 1. DNA was puried using SPRI beads as
previously described. Library quantication was performed with the
Illumina library quantication kit (Kapa Biosystems) according to the
manufacturer's protocol. Paired-end 2 × 150 sequencing was per-
formed on an Illumina NextSeq. 500 instrument with a NextSeq. 500/
550 Mid Output Kit v2 (300 cycles).
3.12. Sequence analysis
Sequences were trimmed using Cutadapt (version 1.9.1) in order to
trim adaptor sequences and low-quality bases, and remove trimmed
sequences that were shorter than 80 nucleotides (nt) long (Martin,
2011). Quality base was set to 33 (default) and quality cutowas set to
30 for the 5and 3ends. The rst base of each sequence was also
trimmed. The CD-HIT-DUP sequence clustering tool was then used to
collapse reads with 99% global pairwise identity, leaving unique reads
(Li and Godzik, 2006). Python-derived sequences were then ltered
using the Bowtie2 alignment tool (version 2.2.5) (Langmead and
Salzberg, 2012). First, a bowtie index was generated from the host
genomic sequence [Python bivittatus (Burmese python) genome as-
sembly (NC_021479.1)] and then sequences aligning with a local mode
alignment score greater than 60 were removed. SPAdes genome as-
sembler (version 3.5.0) was used to generate contiguous sequences
(contigs) (Bankevich et al., 2012). Then, to taxonomically categorize
sequences, the NCBI nt database was queried with all contigs greater
than 150 nt using the BLASTn alignment tool (version 2.2.30+)
(Altschul et al., 1990; Camacho et al., 2009). Any hit with an expect
value less than 10
was assigned taxonomically according to the se-
quence with the highest alignment score (Altschul et al., 1990;BLAST+
Command Line Applications User Manual, n.d.). Additionally, to at-
tempt to categorize contigs that were too divergent to produce a high
scoring nt-nt alignment, the NCBI nr database was queried using
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
Diamond (version with an expect value of 0.001 (Buchnk
et al., 2015). The same process was performed using all the reads that
did not form contiguous sequences from SPAdes genomic assembly,
except GSNAP alignment tool (version 2017-05-08) was used instead of
BLASTn (Wu and Nacu, 2010). Lastly, a bowtie index was generated
from ball python nidovirus 1 (NC_024709.1) and sequences aligning
with a local mode alignment score greater than 60 were evaluated in
Geneious (version 9.0.5) for percent identity. The inoculum sequence
was deposited in Genbank (accession MG752895) and raw sequence
data was deposited in the NCBI Short Read Archive database (accession
3.13. Bacteriology
Oral swabs collected in agar medium (see Experimental infection)
were submitted to the Colorado State University Veterinary Diagnostic
Lab for aerobic bacterial culture.
4. Results
4.1. Antemortem clinical ndings
Choanal, cloacal, and oroesophageal swabs tested negative for
BPNV by qRT-PCR in all snakes prior to inoculation. Clinical signs in
infected snakes (BP-A, B, and C) began at 4 weeks PI and progressed
over time. Initial clinical signs were moderate reddening of the choanal
and oral mucosa and excessive oral mucus secretion. With progression,
oral reddening and mucus secretions became more severe resulting in
excessive swallowing and ventral oral swelling. This was accompanied
by small mucosal hemorrhages (petechiations), increased respiratory
eort and rate, open-mouthed breathing, and anorexia by weeks 1012
(Fig. 1).
4.2. Postmortem gross and histologic ndings
BP-C displayed mild clinical signs (mucinous exudate in the oral
cavity) at 5 weeks PI and was euthanized to assess lesions of early in-
fection. Grossly, the oral mucosa was diusely and mildly reddened
with moderate mucinous secretions in the oral cavity and cranial eso-
phagus. Histologically, there was moderate chronic-active mucinous
rhinitis, stomatitis, glossitis, tracheitis, and cranial esophagitis with
variable epithelial proliferation. Inammatory inltrates were mixed
with moderate numbers of lymphocytes, plasma cells, heterophils and
macrophages. There was mild faveolar pneumocyte hyperplasia re-
gionally with the accumulation of luminal proteinaceous material. The
caudal esophagus was normal.
BP-A was euthanized at 10 weeks PI and BP-B was euthanized at 12
weeks PI, both due to anorexia, intermittent increased respiratory eort
and open-mouthed breathing, following IACUC protocol euthanasia
criteria. Grossly, oral cavities of each snake were similar to BP-C but
with signicantly more mucinous exudate. BP-A also had a focal ul-
ceration of the glottis, the caudal esophagus adjacent to the lungs was
markedly dilated with air and mucus, and the cranial 1/3 of the lungs
were wet and red (Fig. 2). BP-B lungs were slightly reddened and wet in
the cranial portion but the caudal esophagus was grossly normal. His-
tologically, both snakes had similar but more severe lesions in the upper
respiratory tract (URT) and cranial esophagus as compared to BP-C
(Fig. 3A and B). Additionally, there were regions of erosion and ul-
ceration in areas of inammation as well as individual epithelial cell
necrosis and regions of marked epithelial proliferation. The caudal
esophagus of BP-A also had similar inammatory inltrates to that in
the cranial esophagus but these were signicantly milder. Lumina of the
URT, cranial esophagus, central lumen of the lung, and faveoli con-
tained mucus, necrotic debris, heterophils, hemorrhage, and occasional
colonies of short Gram-negative bacterial rods. Both snakes (BP-A
greater than BP-B) had a mild interstitial pneumonia of the cranial lung
eld with pneumocyte proliferation (Fig. 3C). Lesions were character-
ized by multifocal hyperplasia of respiratory epithelial cells lining the
central lumen, hypertrophy and hyperplasia of faveolar pneumocytes
(predominately in the luminal 1/3 of the faveoli), and expansion of the
interstitium by edema and similar inammatory cells to that in the
Sham inoculated snakes (BP-D and E) did not show clinical signs nor
have histologic lesions of the respiratory tract or esophagus (Fig. 3D-F).
Both the infected and control snakes had moderate to severe lympho-
plasmacytic and heterophilic, non-ulcerative colitis of unknown origin
and mild lymphohistiocytic to granulomatous embolic hepatitis, which
are considered unrelated to the clinical and histologic signs found only
in the infected snakes. Gram, PAS, and Ziehl-Neelsen acid fast stains did
not elucidate an infectious agent associated with these lesions. Re-
maining tissues were histologically normal (heart, kidneys, spleen,
stomach, small intestine, pancreas, gall bladder, adrenal glands, go-
nads, brain, spinal cord, vertebral bone, bone marrow, skin, and ske-
letal muscle).
4.3. Western blot
DPHt cells inoculated with oroesophageal swabs from both infected
and uninfected snakes were analyzed by western blot using anti-nu-
cleocapsid protein polyclonal antisera to determine antibody speci-
city. This polyclonal antibody was designed in our laboratory and de-
veloped in rabbits, and this is the rst demonstration of its specicity.
The predicted length and molecular mass of the BPNV nucleocapsid
protein is 152 aa and 16.7 kDa (GenBank: AIJ50569.1) and a protein of
approximately this size was detected in BP-A, B, and C (infected) but
not BP-D or E (control; Fig. 4).
Fig. 1. Antemortem ndings. Clinical signs in in-
fected snakes (BP-A, B, and C) began at 4 weeks PI
and included moderate reddening of choanal and
oral mucosa and abundant oral mucus secretion (A).
This progressed to excessive swallowing, ventral oral
swelling, mucosal petechiations, increased re-
spiratory eort and rate, open-mouthed breathing
(B), and anorexia by weeks 1012. Control snakes
were clinically normal throughout the experiment.
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
4.4. Immunohistochemistry
Immunohistochemical staining of viral antigen (nucleocapsid
protein) was present in all infected snakes within the epithelial surface
of the oral mucosa, nasal mucosa, trachea, and esophagus (Fig. 5A-C).
Immunopositive staining was detected in the cytoplasm of presumed
Fig. 2. Postmortem ndings. Early lesions (BP-C)
included diuse reddening of the oral mucosa with
moderate mucinous secretions in the oral cavity and
cranial esophagus. Later lesions (BP-A and BP-B)
included increased severity of early lesions with
focal ulceration of the glottis in BP-A (A; arrowhead).
The caudal esophagus adjacent to the lungs was
markedly dilated with air and mucus (B; star), and
the cranial 1/3 of the lungs were wet and red in BP-A
(B; arrow). Control snakes were normal.
Fig. 3. Histopathology. Infected snakes had severe
chronic-active mucinous rhinitis and stomatitis (A),
tracheitis and esophagitis with epithelial prolifera-
tion (B), and interstitial proliferative pneumonia (C).
Control snakes (D-F) were histologically normal.
Arrows indicate inammation; stars indicate epithe-
lial proliferation. Hematoxylin and eosin. Boxes are
represented in higher magnication in the insets (A,
C, D, F). Scale bars: Inset scale 200 µm. (A) and (D)
1000 µm. (B) 200 µm. (E) 100 µm. (C) and (F)
500 µm. Esophagus (Es). Trachea (Tr).
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
epithelial cells, predominately in regions of inammation. Intact and
degenerate cells containing viral antigen, or free viral antigen admixed
with mucus was frequently present in the lumen of the URT and GI
tract, or rarely faveoli (BP-A) (Fig. 5D). Within the caudal esophagus,
stomach, small intestine, and colon viral antigen was restricted to the
luminal contents and not detected in epithelial cells. However, regions
of mucosal-associated lymphoid tissue of the caudal esophagus, small
intestine, and colon contained low to moderate numbers of im-
munopositive cells suspected to be associated with M-cell-like uptake
and sampling of the luminal contents (Fig. 6). Viral antigen was not
detected in any other tissues in infected snakes (heart, kidneys, spleen,
pancreas, gall bladder, adrenal glands, gonads, brain, spinal cord,
vertebral bone, bone marrow, skin, and skeletal muscle). No viral an-
tigen was detected in the control snakes.
4.5. Viral RNA detection
Viral RNA was detectable in oroesophageal, choanal, and cloacal
swabs by qRT-PCR beginning at 1 week PI in all infected snakes, with
an increase noted at 4 weeks PI in choanal and oroesophageal swab
samples, coinciding with the onset of clinical signs. Levels of viral RNA
increased steadily over the course of the experiment and reached levels
exceeding 1000x more than the initial sampling time point (Fig. 7).
Viral RNA was detected in multiple postmortem tissues from infected
snakes, including trachea and esophagus, liver, kidney, heart, stomach,
and feces, the lung of BP-A, and the small intestine and colon of BP-A
and BP-C (Fig. 8). Respiratory and GI tract tissues and feces contained
Fig. 4. Western blot of BPNV nucleoprotein. Viral nucleocapsid protein (approximately
16.7 kDa) was detected in DPHt cells inoculated with OES from BP-A, B, and C (infected)
but not BP-D or E (control).
Fig. 5. Immunohistochemistry of the respiratory tract and cranial esophagus. Viral antigen (red staining) was prominent in the epithelial layer of the oral mucosa, trachea, and cranial
esophagus of infected snakes (A-C). There were only rare positive cells within the pulmonary epithelium (D top left inset) and faveolar lumen (D top right inset) of one infected snake (BP-
A), as compared with the positive IHC control from a nidovirus-positive green tree python that died of respiratory disease (E, lung). Intact and degenerate cells that contained viral
antigen were also found in the lumen of these tissues (arrowheads) admixed with cell-free viral antigen in mucus (arrow) and hemorrhage (star). No viral antigen was detected in the
control snakes (F-I). The IHC negative control (J) was the same lung tissue as that used for the positive control, but lacking primary antibody application. Fav, faveolar lumen; IHC,
immunohistochemistry. Primary antibody: polyclonal rabbit NdvNcAb. Counter stain: hematoxylin. Scale bars: A-C and F-H = 50 µm; D-E and I-J lower = 500 µm, upper (inset) =
20 µm.
Fig. 6. Immunohistochemistry of the gastrointestinal tract. Representative images of the
stomach (A) and small intestine (B) of infected snakes. Viral antigen (red staining) was
detected in the lumen of the GI tract admixed with intact and degenerate cells and mucus
(large black arrow). Epithelial cells (arrowheads) were immunonegative throughout the
caudal esophagus and GI tract. However, viral antigen was detected in cells along the
mucosal surface (small black arrows) overlying mucosal-associated lymphoid tissue
(MALT; asterisk) of the esophagus, small intestine, and colon. Immunopositive cells ex-
tended into the center of MALT (white arrows). Primary antibody: polyclonal rabbit
NdvNcAb. Counter stain: hematoxylin. Scale bars = 100 µm.
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
the highest viral RNA per host mRNA (GAPDH), with remaining organs
having detectable but lower viral RNA levels. Viral RNA was not de-
tected in any swabs or tissues from control snakes.
4.6. Virus isolation
The presence of infectious virus in collected swabs, tissues, and
excreta was evaluated by virus isolation in cell culture. Inoculation of
DPHt cells with terminal oroesophageal swabs from infected snakes
(BP-A, B, and C) resulted in viral replication, as determined by im-
munouorescence using antibodies targeting the BPNV nucleocapsid
protein, and cytopathic eects (Fig. 9). Speckled immunouorescent
staining was restricted to the cytoplasm of infected cells. Cytopathic
eects included cell death and syncytial cell formation (Fig. 9 2DPI HM
panel). Viral infection of cells was detected as early as 12 h PI with
greater than 50% of cells infected by 2 days PI and signicant cell death
detected by day 4 PI. Plaque assay of the terminal oroesophageal swab
from BP-A revealed a viral titer of 1.67 × 10
PFU/ml. Infectious virus
was also isolated from feces of infected snakes, as determined by im-
munouorescence of cell culture, but viral titers were not measured.
Inoculation of DPHt cells with oral swabs and feces from control snakes
(BP-D and E) did not produce detectable virus by immunouorescence
nor result in cytopathic eects. Virus isolation attempts with post-
mortem fresh lung samples from all snakes (infected and control) were
4.7. Metagenomic sequencing
Metagenomic sequencing was used to validate the purity of the in-
oculum, to rule out other possible etiologic agents, and to evaluate
genomic sequence of virus reisolated from infected snakes. The average
number of read pairs per sample was 4.4 × 10
. On average, 93%, 11%,
and 3.6% of sequences remained following adaptor and quality l-
tering, collapsing to unique reads, and ltration of python derived se-
quences, respectively. Remaining sequences were compared against
nucleotide and protein databases for taxonomic assessment. Sequences
aligning to ball python nidovirus 1 (NCBI taxonomy ID: 1986118) and
python nidovirus (NCBI taxonomy ID: 1526652) were detected in the
lung and trachea/esophagus of BP-A, B, and C (infected snakes) and
BPNV-148 inoculum. Nidovirus sequences were not detected in BP-D or
E (control snakes). In addition to python nidovirus, BP-A lung had se-
quences aligning to Pseudomonas species. In all samples, Python molurus
and curtus endogenous retrovirus-like sequences were detected. Other
sequences aligning to organisms in the queried databases were pre-
dominately non-specic alignments due to low complexity or tax-
onomically ambiguous sequences that span multiple taxa. Other than
Pseudomonas reads found in BP-A lung, no other sequences specically
aligning to known primary pathogenic or opportunistic infectious
agents were identied. We analyzed the sequences generated through
bowtie alignment of the BPNV inoculum and the viruses re-isolated
from infected snakes. The sequence of the nidovirus in the inoculum
was 96.2% identical to BPNV (NC_024709.1), and we obtained cov-
erage across the complete genome. We did not obtain complete genome
coverage for the recovered viruses, but the bowtie-mapped reads that
did align to the BPNV sequence were 99.7100% identical to the in-
oculum virus.
4.8. Bacteriology
Aerobic culture was performed on oral swabs from BP-A, B, D, and
E. BP-A yielded heavy growth of Pseudomonas aeruginosa and
Stenotrophomonas maltophilia and BP-B yielded moderate growth of
Bordatella and Pseudomonas species. In the control snakes, BP-D yielded
light growth of Acinetobacter baumannii,Brevundimonas species, Delftia
acidovorans, and Pseudomonas species; BP-E yielded light growth of
Pseudomonas aeruginosa.
5. Discussion
Novel nidoviruses were recently found in multiple python species
with respiratory disease, but a causal relationship between infection
and disease has not been established (Bodewes et al., 2014; Dervas
et al., 2017; Marschang and Kolesnik, 2017; Stenglein et al., 2014;
Uccellini et al., 2014). Through the use of experimental infections, our
study is the rst to fulll Koch's postulates and demonstrate a causal
relationship between python nidovirus infection and respiratory and
esophageal disease in ball pythons. Our ndings demonstrate that
BPNV infection caused marked mucinous inammation of the URT and
cranial esophagus with progression towards proliferative interstitial
pneumonia. These ndings are consistent with previous reports of ni-
dovirus infection in multiple python species (Bodewes et al., 2014;
Dervas et al., 2017; Stenglein et al., 2014; Uccellini et al., 2014). In our
study, the clinical hallmark of this disease was excessive mucous pro-
duction in the oral cavity, which was accompanied histologically by
marked inammation and epithelial proliferation. Additionally, cranial
esophagitis was a prominent lesion, similar to that in previous reports
(Uccellini et al., 2014). Mucous production and esophagitis are not
characteristic ndings with other respiratory viruses in snakes and,
therefore, may be useful clinical and histologic features for dierential
Fig. 7. Relative viral RNA in antemortem swabs. Total RNA was extracted from choanal
(CHS), oroesophageal (OES), and cloacal (CLS) swabs and was analyzed by qRT-PCR
using primers targeting BPNV RNA. Relative viral RNA was determined by comparison of
each sample Ct to the sample with the highest Ct (lowest viral RNA) at the rst collection
time point following inoculation (BP-B OES week 1 PI). All samples were run in duplicate
and error bars indicate standard deviations. Viral RNA was detectable beginning at 1
week PI in all swabs of the infected snakes, with an increase noted at 4 weeks PI, cor-
relating with the initiation of clinical signs. Viral RNA continued to increase over the
course of the experiment, consistent with amplication in the host. Early clinical signs
included reddening of the oral mucosa and abundant oral mucus secretions. Late clinical
signs included increased respiratory distress, open-mouthed breathing, and anorexia.
Control snakes were negative throughout the experiment.
Fig. 8. Relative viral RNA in postmortem tissues. Total RNA was extracted from fresh
tissues and analyzed by qRT-PCR. Relative viral RNA was determined by normalization of
BPNV Ct to GAPDH (a cellular mRNA) Ct within each sample. All samples were run in
duplicate and standard deviation is represented by error bars. Viral RNA was detected in
nearly all tissues of infected snakes (BP-A, B, and C). Control snakes were negative in all
tissues. T/E (trachea/esophagus); SI (small intestine).
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
diagnosis of nidovirus infection in pythons. However, the overlap in
clinical and histologic lesions with other viral agents (e.g. respiratory
distress, anorexia, and proliferative pneumonia) could warrant the de-
velopment of a multiplex PCR test to screen for multiple reptile re-
spiratory viruses.
Viral antigen was detected in the mucosa of the oral and nasal
cavity, trachea, and esophagus of all infected snakes, indicating a
tropism for epithelial cells, especially ciliated cells of the respiratory
tract and upper esophagus. Viral antigen was rarely detected in pul-
monary epithelial cells, which may be due to the short time course of
the experimental infections as discussed below. Viral antigen was not
detected in the epithelium of the caudal esophagus or GI tract, but was
present in the GI lumen by IHC and viral RNA was detected in these
samples by qRT-PCR. This suggests that GI epithelium is not a site of
viral replication, but the GI tract is a conduit for virus that is swallowed
and passed in the feces. Additionally, viral antigen was detected in
mucosal-associated lymphoid tissue of the caudal esophagus, small in-
testine, and colon. Sampling of the luminal contents by M-cell-like
uptake could be a method for systemic spread, as indicated by viral
RNA detection in non-respiratory/GI tissues.
Until recently, primary viral causes of pneumonia in pythons in-
cluded paramyxovirus (ferlaviruses) and reovirus (Ariel, 2011;
Jacobson, 2007a; Marschang, 2011). Other causes of proliferative
pneumonia include chlamydophilosis, mycoplasmosis, chronic bacterial
or parasitic infection, or toxin exposure (Bodetti et al., 2002; Jacobson,
2007b; Penner et al., 1997). Histopathology, bacterial culture, and next
generation sequencing did not yield evidence of other primary in-
fectious agents in the infected snakes or inoculum, consistent with
BPNV as the principal cause of respiratory disease. The bacteria de-
tected by oral aerobic culture and sequencing of the lung have been
found as oral ora in clinically healthy snakes, (Blaylock, 2001;
Goldstein et al., 1981; Plenz et al., 2015) but the presence of moderate
to heavy growth in the infected snakes is suspected to be secondary to
viral infection and disruption of physical or immune barriers. Sec-
ondary bacterial infections are a common sequelae to viral disease in all
types of animals and humans and have been documented in previous
cases of python nidoviral infection (Bodewes et al., 2014; Uccellini
et al., 2014). The role of secondary infections and the progression of
disease in pythons has yet to be determined and warrants further in-
In previous reports, the degree of pneumonia associated with ni-
dovirus infection was more advanced at the time of death when
compared to our study (Bodewes et al., 2014; Dervas et al., 2017;
Stenglein et al., 2014; Uccellini et al., 2014). This is highlighted by our
immunohistochemistry control (Fig. 5E) collected from a green tree
python that had been nidovirus positive for over 6 months and even-
tually died of respiratory disease. Had we allowed for a longer time
course of infection in this study, it is likely that the infected snakes
would have progressed to more severe disease. Another possibility
unexplored by this study, though suggested by other studies within our
lab (unpublished data), is that some snakes clear nidovirus infection
and recover.
Pythons have an overcapacity for oxygen consumption, and there-
fore rarely show clinical signs of respiratory disease until the oxygen
exchange capacity is severely limited (Starck et al., 2015). A recent
report in green tree pythons demonstrated infection of respiratory and
faveolar epithelial cells to be associated with apoptosis, proliferation,
and high numbers of serous/mucous granules within the cytoplasm
(Dervas et al., 2017). Respiratory epithelial proliferation and mucus
production were postulated to result in mechanical and physiologic
inhibition of gas exchange within the lungs, resulting in death. Our
study ended at 12 weeks PI based on euthanasia criteria, however the
degree of respiratory eort was greater than would be expected for the
mild pneumonia in the infected snakes. It is thought that the excessive
production of mucus in the oral cavity and upper airway contributed to
respiratory diculty through obstruction of the airway and glottis.
Therefore, the obstructive mechanical eects of mucus could also play a
role in the upper respiratory tract by aecting overall air intake, in
addition to the eects in the lung on direct oxygen exchange (Dervas
et al., 2017).
Although the disease we observed closely resembles that seen in
naturally infected snakes, there are several aspects of our study that
may not have recapitulated natural infection. First, disease may be
dose-dependent, and the 1.1 × 10
PFU administered may exceed a
typical natural infectious dose. Second, the natural route(s) of trans-
mission are unknown. Nidovirus RNA is detected in oral swabs
(Marschang and Kolesnik, 2017) and respiratory/pulmonary epithelial
cells of infected snakes (Dervas et al., 2017; Uccellini et al., 2014),
indicating the oral cavity and respiratory tract as regions of viral re-
plication and possible routes of exposure. We inoculated snakes in the
oral cavity and upper trachea to mimic URT exposure and subsequently
detected infectious virus in these regions indicating that virus is re-
leased in respiratory and oral secretions. Based on this nding, trans-
mission may involve fomites or aerosolization. The presence of
Fig. 9. Immunouorescence of DPHt cells inoculated with infectious swabs and tissues. DPHt cells were inoculated with ltered terminal oroesophageal swabs (OES) and feces from
infected snakes (BP-A OES represented in the bottom panel) resulting in viral infection, amplication, and cytopathic eects (syncytial formation and cell death). Inoculation with lung
homogenate from infected snakes yielded negative results. Swabs and tissues from control snakes also yielded negative results. The top panel represents uninfected DPHt cells. HrPI (hours
post infection); DPI (days post infection); HM (high magnication). Magnication is 100x in all images, except HM (high magnication) which is 1000x. Fluorescence is Hoescht 33342
(blue) for nuclear staining and Alexa 488 (green) for the BPNV nucleocapsid protein detection using NdvNcAb. All images include overlay of both uorescence lters.
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
infectious virus in the feces indicates that fecal-oral transmission is also
Other factors, including husbandry, age, sex, and immune status
could modulate disease progression. Care of the snakes in this study
followed best practices for ball python husbandry. Snakes were housed
separately with appropriate enclosures and climate control and handled
minimally to limit stress. Although all snakes in this study were juvenile
males (one was of undetermined sex), reports have documented nido-
virus-associated respiratory disease in snakes of various sexes and ages
(Dervas et al., 2017; Stenglein et al., 2014; Uccellini et al., 2014).
The python nidoviruses are part of an expanding clade of respiratory
disease-associated toroviruses. A related virus has also been identied
in wild shingleback lizards with respiratory disease (ODea et al., 2016).
Clinical signs in lizards were similar to those observed in pythons, in-
cluding excessive mucous in the oral cavity and anorexia. Additionally,
a related nidovirus in cattle has been associated with severe tracheitis
and pneumonia (Tokarz et al., 2015). The similar clinical ndings
suggest that viruses in this clade may share tissue tropism and patho-
genic mechanisms. Related virus sequences have also been detected in
snake-associated nematodes, though no information beyond sequence is
available (Shi et al., 2016). That this clade includes viruses with rep-
tilian, mammalian, and invertebrate hosts highlights its diversity, and
emphasizes the need for further investigation into this expanding col-
lection of potential pathogens.
The authors would like to thank the following groups and in-
dividuals: The histology laboratory at Colorado State University, Fort
Collins Veterinary Diagnostic Laboratory for performing the histolo-
gical and immunohistochemical procedures; Justin Lee and the CSU
Next Generation Sequencing facility; the CSU Laboratory Animal
Resources faculty and staand Dr. Matt Johnston for help in snake
handling and care; and Alora Lavoy and Lauren Kuechler for their
contributions to this study.
Funding sources
Funding to develop and characterize the Diamond Python Heart
cells used in this project was provided by the David R. Atkinson Center
Academic Venture Fund at Cornell University (to J.S.L.P). Funding for
experimental infections and subsequent disease characterization was
provided by Colorado State University.
Conict of interest
There is no conict of interest.
Adams, M.J., Lefkowitz, E.J., King, A.M.Q., Harrach, B., Harrison, R.L., Knowles, N.J.,
Kropinski, A.M., Krupovic, M., Kuhn, J.H., Mushegian, A.R., Nibert, M.,
Sabanadzovic, S., Sanfaçon, H., Siddell, S.G., Simmonds, P., Varsani, A., Zerbini,
F.M., Gorbalenya, A.E., Davison, A.J., 2017. Changes to taxonomy and the interna-
tional code of virus classication and Nomenclature ratied by the international
committee on taxonomy of viruses (2017). Arch. Virol. 162, 25052538. http://dx.
Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment
search tool. J. Mol. Biol. 215, 403410.
Ariel, E., 2011. Viruses in reptiles. Vet. Res. 42, 100.
Bankevich, A., Nurk, S., Antipov, D., Gurevich, A.A., Dvorkin, M., Kulikov, A.S., Lesin,
V.M., Nikolenko, S.I., Pham, S., Prjibelski, A.D., 2012. SPAdes: a new genome as-
sembly algorithm and its applications to single-cell sequencing. J. Comput. Biol. 19,
Batts, W.N., Goodwin, A.E., Winton, J.R., 2012. Genetic analysis of a novel nidovirus from
fathead minnows. J. Gen. Virol. 93, 12471252.
BLAST+Command Line Applications User Manual, n.d. [WWW Document]. URL http://
Blaylock, R.S., 2001. Normal oral bacterial ora from some southern African snakes.
Onderstepoort J. Vet. Res. 68, 175182.
Bodetti, T.J., Jacobson, E., Wan, C., Hafner, L., Pospischil, A., Rose, K., Timms, P., 2002.
Molecular evidence to support the expansion of the hostrange of Chlamydophila
pneumoniae to include reptiles as well as humans, horses, koalas and amphibians.
Syst. Appl. Microbiol. 25, 146152.
Bodewes, R., Lempp, C., Schurch, A.C., Habierski, A., Hahn, K., Lamers, M., von
Dornberg, K., Wohlsein, P., Drexler, J.F., Haagmans, B.L., Smits, S.L., Baumgartner,
W., Osterhaus, A.D.M.E., 2014. Novel divergent nidovirus in a python with pneu-
monia. J. Gen. Virol. 95, 24802485.
Buchnk, B., Xie, C., Huson, D.H., 2015. Fast and sensitive protein alignment using
DIAMOND. Nat. Methods 12, 5960.
Camacho, C., Coulouris, G., Avagyan, V., Ma, N., Papadopoulos, J., Bealer, K., Madden,
T.L., 2009. BLAST+: architecture and applications. BMC Bioinforma. 10, 421. http://
De Groot, R., Cowley, J., Enjuanes, L., Faaberg, K., Perlman, S., Rottier, P., Snijder, E.,
Ziebuhr, J., Gorbalenya, A., 2012. Order - nidovirales. In: KIng, A.M., Adams, M.J.,
Carstens, E.B., Lefkowitz, E.J. (Eds.), Virus Taxonomy: Classication and
Nomenclature of Viruses: Ninth Report of the International Committee on Taxonomy
of Viruses. Elsevier, San Diego, pp. 784794.
Dervas, E., Hepojoki, J., Laimbacher, A., Romero-Palomo, F., Jelinek, C., Keller, S.,
Smura, T., Hepojoki, S., Kipar, A., Hetzel, U., 2017. Nidovirus-associated Proliferative
Pneumonia in the green tree Python (Morelia viridis). J. Virol.
Goldstein, E.J., Agyare, E.O., Vagvolgyi, A.E., Halpern, M., 1981. Aerobic bacterial oral
ora of garter snakes: development of normal ora and pathogenic potential for
snakes and humans. J. Clin. Microbiol. 13, 954956.
Gonzalez, J.M., Gomez-Puertas, P., Cavanagh, D., Gorbalenya, A.E., Enjuanes, L., 2003. A
comparative sequence analysis to revise the current taxonomy of the family
Coronaviridae. Arch. Virol. 148, 22072235.
Graham, R.L., Donaldson, E.F., Baric, R.S., 2013. A decade after SARS: strategies for
controlling emerging coronaviruses. Nat. Rev. Micro 11, 836848.
Jacobson, E.R., 2007a. Infectious Diseases and Pathology of Reptiles. CRC Press (Chapter
9. Viruses and Viral Diseases of Reptiles).
Jacobson, E.R., 2007b. Infectious Diseases and Pathology of Reptiles. CRC Press (Chapter
5. Host response to infectious agents and identication of pathogens in tissue sec-
Langmead, B., Salzberg, S.L., 2012. Fast gapped-read alignment with Bowtie 2. Nat.
Methods 9, 357359.
Lauber, C., Ziebuhr, J., Junglen, S., Drosten, C., Zirkel, F., Nga, P.T., Morita, K., Snijder,
E.J., Gorbalenya, A.E., 2012. Mesoniviridae: a proposed new family in the order
Nidovirales formed by a single species of mosquito-borne viruses. Arch. Virol. 157,
Li, W., Godzik, A., 2006. Cd-hit: a fast program for clustering and comparing large sets of
protein or nucleotide sequences. Bioinformatics 22, 16581659.
Marschang, R.E., 2011. Viruses infecting reptiles. Viruses 3, 20872126. http://dx.doi.
Marschang, R.E., Kolesnik, E., 2017. Detection of nidoviruses in live pythons and boas.
Tierarztl. Prax. Ausg. K. Klient. Heimtiere 45, 2226.
Martin, M., 2011. Cutadapt removes adapter sequences from high-throughput sequencing
reads. EMBnet. J. 17.
Masters, P.S., Perlman, S., 2013. Coronaviridae. In: Knipe, D.M., Howley, P.M. (Eds.),
Fields Virology. Lippincott Williams & Wilkins, pp. 825858.
Nga, P.T., Parquet, M., del, C., Lauber, C., Parida, M., Nabeshima, T., Yu, F., Thuy, N.T.,
Inoue, S., Ito, T., Okamoto, K., Ichinose, A., Snijder, E.J., Morita, K., Gorbalenya, A.E.,
2011. Discovery of the rst insect nidovirus, a missing evolutionary link in the
emergence of the largest RNA virus genomes. PLoS Pathog. 7, e1002215. http://dx.
ODea, M.A., Jackson, B., Jackson, C., Xavier, P., Warren, K., 2016. Discovery and partial
genomic characterisation of a novel nidovirus associated with respiratory disease in
wild Shingleback Lizards (Tiliqua rugosa). PLoS One 11, e0165209. http://dx.doi.
Penner, J.D., Jacobson, E.R., Brown, D.R., Adams, H.P., Besch-Williford, C.L., 1997. A
novel Mycoplasma sp. associated with proliferative tracheitis and pneumonia in a
Burmese python (Python molurus bivittatus). J. Comp. Pathol. 117, 283288.
Plenz, B., Schmidt, V., Grosse-Herrenthey, A., Kruger, M., Pees, M., 2015.
Characterisation of the aerobic bacterial ora of boid snakes: application of. Vet. Rec.
176, 285.
Pradesh, U., Chikitsa, P.D.D.U.P., Vishwa, V., Dhcirnci, C., Izcitncigcir, B., 2014.
Toroviruses aecting animals and humans: a review. Asian J. Anim. Vet. Adv. 9,
Runckel, C., Flenniken, M.L., Engel, J.C., Ruby, J.G., Ganem, D., Andino, R., DeRisi, J.L.,
2011. Temporal analysis of the honey bee microbiome reveals four novel viruses and
seasonal prevalence of known viruses, Nosema, and Crithidia. PLoS One 6, e20656.
Schutze, H., Ulferts, R., Schelle, B., Bayer, S., Granzow, H., Homann, B., Mettenleiter,
T.C., Ziebuhr, J., 2006. Characterization of White bream virus reveals a novel genetic
cluster of nidoviruses. J. Virol. 80, 1159811609.
Shi, M., Lin, X.-D., Tian, J.-H., Chen, L.-J., Chen, X., Li, C.-X., Qin, X.-C., Li, J., Cao, J.-P.,
Eden, J.-S., Buchmann, J., Wang, W., Xu, J., Holmes, E.C., Zhang, Y.-Z., 2016.
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
Redening the invertebrate RNA virosphere. Nature.
Snijder, E.J., Kikkert, M., 2013a. Arteriviruses. In: Knipe, D.M., Howley, P.M. (Eds.),
Fields Virology. Lippincott Williams & Wilkins, pp. 859879.
Snijder, E.J., Kikkert, M., Fang, Y., 2013b. Arterivirus molecular biology and pathogen-
esis. J. Gen. Virol. 94, 21412163.
Starck, J.M., Weimer, I., Aupperle, H., Muller, K., Marschang, R.E., Kiefer, I., Pees, M.,
2015. Morphological pulmonary diusion capacity for oxygen of Burmese Pythons
(Python molurus): a comparison of animals in healthy condition and with dierent
pulmonary infections. J. Comp. Pathol. 153, 333351.
Stenglein, M.D., Jacobson, E.R., Wozniak, E.J., Wellehan, J.F.X., Kincaid, A., Gordon, M.,
Porter, B.F., Baumgartner, W., Stahl, S., Kelley, K., Towner, J.S., DeRisi, J.L., 2014.
Ball python nidovirus: a candidate etiologic agent for severe respiratory disease in
Python regius. mBio 5, e014841414.
Stenglein, M.D., Jacobson, E.R., Chang, L.-W., Sanders, C., Hawkins, M.G., Guzman, D.S.-
M., Drazenovich, T., Dunker, F., Kamaka, E.K., Fisher, D., Reavill, D.R., Meola, L.F.,
Levens, G., DeRisi, J.L., 2015. Widespread recombination, reassortment, and
transmission of unbalanced compound viral genotypes in natural arenavirus infec-
tions. PLoS Pathog. 11, e1004900.
Stenglein, M.D., Sanchez-Migallon Guzman, D., Garcia, V.E., Layton, M.L., Hoon-Hanks,
L.L., Boback, S.M., Keel, M.K., Drazenovich, T., Hawkins, M.G., DeRisi, J.L., 2017.
Dierential disease susceptibilities in experimentally REptarenavirus-infected Boa
Constrictors and Ball Pythons. J. Virol. 91.
Tokarz, R., Samero, S., Hesse, R.A., Hause, B.M., Desai, A., Jain, K., Lipkin, W.I., 2015.
Discovery of a novel nidovirus in cattle with respiratory disease. J. Gen. Virol. 96,
Uccellini, L., Ossibo, R.J., de Matos, R.E.C., Morrisey, J.K., Petrosov, A., Navarrete-
Macias, I., Jain, K., Hicks, A.L., Buckles, E.L., Tokarz, R., McAloose, D., Lipkin, W.I.,
2014. Identication of a novel nidovirus in an outbreak of fatal respiratory disease in
ball pythons (Python regius). Virol. J. 11, 144.
Wu, T.D., Nacu, S., 2010. Fast and SNP-tolerant detection of complex variants and spli-
cing in short reads. Bioinformatics 26, 873881.
L.L. Hoon-Hanks et al. Virology 517 (2018) 77–87
... In reptiles, clinical signs associated with infection of the respiratory tract appear to be the most common feature of nidovirus infection (15,16,18,21,41,55). Initial clinical signs in captive pythons include increased amounts of clear or mucoid material in the nose and mouth and oral inflammation (stomatitis). ...
... Clinical signs were observed 4 weeks after exposure and included mucosal hyperaemia and profuse mucus secretion, followed by a progression to the appearance of petechial mucosal haemorrhages, open mouth breathing, and anorexia by 10-12 weeks post exposure. The presence of infectious virus was confirmed using virus isolation from oroesophageal swabs taken on the day of euthanasia; 5, 10, and 12 weeks post exposure (55). ...
... The tropism for respiratory epithelium has also been confirmed using in situ hybridisation (ISH) (15,18). Experimental infection of P. regius resulted in histological findings consistent with a chronicactive mucinous rhinitis, stomatitis, tracheitis, oesophagitis, and proliferative interstitial pneumonia (55). The proliferative interstitial pneumonia has been a consistent finding in clinical cases of nidovirus infection in snakes and has more recently been called "nidovirus associated proliferative disease-NPD" (16,57). ...
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Since their discovery in 2014, reptile nidoviruses (also known as serpentoviruses) have emerged as significant pathogens worldwide. They are known for causing severe and often fatal respiratory disease in various captive snake species, especially pythons. Related viruses have been detected in other reptiles with and without respiratory disease, including captive and wild populations of lizards, and wild populations of freshwater turtles. There are many opportunities to better understand the viral diversity, species susceptibility, and clinical presentation in different species in this relatively new field of research. In captive snake collections, reptile nidoviruses can spread quickly and be associated with high morbidity and mortality, yet the potential disease risk to wild reptile populations remains largely unknown, despite reptile species declining on a global scale. Experimental studies or investigations of disease outbreaks in wild reptile populations are scarce, leaving the available literature limited mostly to exploring findings of naturally infected animals in captivity. Further studies into the pathogenesis of different reptile nidoviruses in a variety of reptile species is required to explore the complexity of disease and routes of transmission. This review focuses on the biology of these viruses, hosts and geographic distribution, clinical signs and pathology, laboratory diagnosis and management of reptile nidovirus infections to better understand nidovirus infections in reptiles.
... These viruses have been shown to be an important cause of upper-and lower-respiratory disease in pythons. [4][5][6][7] Similar viruses have also been found in boas 7,8 and colubrids. 7 Serpentoviruses have also been described in skinks (Tiliqua spp) with upper-respiratory signs in Australia 9 and in association with a severe disease outbreak in Bellinger River snapping turtles (Myuchelys georgisi) with swollen eyelids and conjunctivitis in Australia. ...
... Virus is shed in oroesophageal, choanal, and cloacal swabs and has also been isolated from feces of infected snakes. 6 Later in infection, highest viral loads may be found in the lungs of infected snakes. 4 Infection has been shown to persist for extended periods of time. ...
... Transmission of serpentoviruses in pythons has been shown to be possible via the oral cavity. 6 Because the viruses have a tropism for epithelial cells of the respiratory tract and upper esophagus, it is considered likely that they are transmitted via oral secretions and possibly fomites and aerosols. Fecal-oral transmission may also occur because virus is also shed via the cloaca. ...
Methods for the detection of pathogens associated with respiratory disease in reptiles, including viruses, bacteria, fungi, and parasites, are constantly evolving as is the understanding of the specific roles played by various pathogens in disease processes. Some are known to be primary pathogens with high prevalence in captive reptiles, for example, serpentoviruses in pythons or mycoplasma in tortoises. Others are very commonly found in reptiles with respiratory disease but are most often considered secondary, for example, gram-negative bacteria. Detection methods as well as specific pathogens associated with upper- and lower-respiratory disease are discussed.
... Total RNA was extracted from fresh-frozen lung, liver, and kidney pools (VC1 and VC2) and oral mucosa, trachea, and lung pool (VC3) using a combination of TRIzol (Ambion Life Technologies, Carlsbad, CA, USA) with RNA clean and concentrator columns (CC-5; Zymo Research, Irvine, CA, USA) as previously described [15]. RNA libraries were generated using the Kapa RNA HyperPrep Kit (Kapa Biosystems, Wilmington, MA, USA) according to the manufacturer's instructions with an input concentration of approximately 50-100 ng of RNA and 6-10 rounds of amplification. ...
... Data analysis was performed as previously described [15]. Briefly, adaptor sequences, low-quality bases, and short sequences (<80 bp) were removed (Cutadapt version 1.18). ...
... Four cell lines were used for virus isolation attempts with fresh-frozen tissue homogenates from VC3 (no tissue remained from VC1-2 for virus isolation attempts): JK cells (boa constrictor kidney) [31], DPHt cells (diamond python heart) [15], IgH2 cells (iguana heart; ATCC, CCCL-108), and VH2 (viper heart; ATCC, CCL-140). Pooled tissues (oral mucosa, lung, trachea) from VC3 were homogenized in brain heart infusion (Becton Dickinson) by manual disruption with a plastic sterile pestle in a 1.5 mL Eppendorf tube. ...
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Serpentoviruses are an emerging group of nidoviruses known to cause respiratory disease in snakes and have been associated with disease in other non-avian reptile species (lizards and turtles). This study describes multiple episodes of respiratory disease-associated mortalities in a collection of juvenile veiled chameleons (Chamaeleo calyptratus). Histopathologic lesions included rhinitis and interstitial pneumonia with epithelial proliferation and abundant mucus. Metagenomic sequencing detected coinfection with two novel serpentoviruses and a novel orthoreovirus. Veiled chameleon serpentoviruses are most closely related to serpentoviruses identified in snakes, lizards, and turtles (approximately 40–50% nucleotide and amino acid identity of ORF1b). Veiled chameleon orthoreovirus is most closely related to reptilian orthoreoviruses identified in snakes (approximately 80–90% nucleotide and amino acid identity of the RNA-dependent RNA polymerase). A high prevalence of serpentovirus infection (>80%) was found in clinically healthy subadult and adult veiled chameleons, suggesting the potential for chronic subclinical carriers. Juvenile veiled chameleons typically exhibited a more rapid progression compared to subadults and adults, indicating a possible age association with morbidity and mortality. This is the first description of a serpentovirus infection in any chameleon species. A causal relationship between serpentovirus infection and respiratory disease in chameleons is suspected. The significance of orthoreovirus coinfection remains unknown.
... Recently, experimental infection of ball pythons established the causal relationship between serpentovirus infection and inflammation with excess mucus production of the upper respiratory and the gastrointestinal tract as the main pathological process early, i.e., within the first 12 weeks, after infection (7). Infected pythons showed severe respiratory distress with only minimal pneumonia, most likely because the mucus overproduction resulted in obstruction of the upper airways (7). ...
... Recently, experimental infection of ball pythons established the causal relationship between serpentovirus infection and inflammation with excess mucus production of the upper respiratory and the gastrointestinal tract as the main pathological process early, i.e., within the first 12 weeks, after infection (7). Infected pythons showed severe respiratory distress with only minimal pneumonia, most likely because the mucus overproduction resulted in obstruction of the upper airways (7). These findings correlate with those we made when studying naturally infected cases; some animals were submitted for euthanasia due to acute respiratory distress but only exhibited inflammatory processes in the nasal and oral cavity, without histologic evidence of tracheitis or pneumonia. ...
... This is particularly relevant in association with hyperplasia of the pulmonary epithelium, which has an impact on the blood gas exchange (16). In some snakes, we additionally observed an esophagitis; this is interpreted as a consequence of overspill and swallowing of virus-laden mucus, a theory also supported by previous studies (6,7,16). The esophageal epithelium contains ciliated cells in various snake species (13), which represent the primary site of viral replication in human coronaviruses (17). ...
During the last years, python nidoviruses (now reclassified as serpentoviruses) have become a primary cause of fatal disease in pythons. Serpentoviruses represent a threat to captive snake collections, as they spread rapidly and can be associated with high morbidity and mortality. Our study indicates that, different from previous evidence, the viruses do not only affect the respiratory tract, but can spread in the entire body with blood monocytes, have a broad spectrum of target cells, and can induce a variety of lesions. Nidovirales is an order of animal and human viruses that comprises important zoonotic pathogens such as Middle East respiratory syndrome coronavirus (MERS-CoV), severe acute respiratory syndrome coronavirus (SARS-CoV), and SARS-CoV-2. Serpentoviruses belong to the same order as the above-mentioned human viruses and show similar characteristics (rapid spread, respiratory and gastrointestinal tropism, etc.). The present study confirms the relevance of natural animal diseases to better understand the complexity of viruses of the order Nidovirales .
... Nidoviruses are linked to respiratory disease in pythons [2,11], and the pathogenicity of nidoviruses in pythons has been recently confirmed by an in vivo study [3]. In addition to pythons, nidoviruses have been reported to cause proliferative interstitial pneumonia in a variety of animal species, including cattle, lizards, and turtles, suggesting that the nidovirus is potential emerging pathogens warranting further investigation [4,8,12]. ...
... Based on the gross and microscopic findings, the primary differential diagnosis was serpentovirus infection. However, other etiologies including toxic gases, ophidian paramyxovirus (OPMV), and bacterial infection such as mycoplasma could not be ruled out [3]. ...
... Furthermore, the amino acid sequences of full-length ORF-1a and 1b were used for phylogenetic analysis [16]. The current serpentovirus was grouped with the serpentovirus identified in ball pythons (KJ541759 and MG752895) and a woma python (Aspidites ramsayi) (MN161571) in the US by the phylogenetic tree based on the amino acid sequence of ORF 1a and 1b (Fig. 3) [3,4,11]. Previous studies conducted in Europe and the US have demonstrated that serpentovirus can be detected by PCR using oral swabs or blood samples from several snake species, including pythons (Pythonidae family), Boids (Boidae family), and Colubrids (Colubridae family), with or without documented respiratory diseases [4,7]. ...
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Reptile-associated nidoviruses (serpentoviruses) have been reported to cause proliferative interstitial pneumonia in pythons and other reptile species. A captive, younger than 2 years old, intact female ball python (Python regius) showed increased oral mucus, wheezing, and audible breathing with weight loss. Gross and microscopic examination revealed large amounts of mucus in the esophagus and proliferative interstitial pneumonia. Serpentovirus genes were detected from the lung tissues by polymerase chain reaction. The current serpentoviruses was phylogenetically grouped with the serpentovirus previously identified in the US. No case of serpentovirus infection has been reported in Asia. The present report provides information of complete genome sequence and global distribution of serpentovirus.
... These viruses are mostly found in pythons, but genetically related viruses have been found in a wide variety of other species (Hoon-Hanks et al., 2019). These nidoviruses have been shown to cause respiratory disease in infected pythons (Hoon-Hanks et al., 2018). ...
... Nidoviruses have been increasingly detected in pet snakes in Europe and the United States (Stenglein et al., 2014;Dervas et al., 2017;Hoon-Hanks et al., 2019;Blahak et al., 2020). These viruses have been proven to cause severe respiratory disease in infected snakes (Hoon-Hanks et al., 2018). Interestingly, in the present study, 10 of the 13 nidovirus-positive snakes had no clinical signs of disease at the time of testing, and respiratory signs were only reported in 1 snake. ...
... The bacteria involved are most often Gram-negative rods, and often include species that can also be found in healthy snakes (Jacobson, 2007;Schmidt et al., 2013). There are also a number of viruses that have been described in association with respiratory disease in snakes, including paramyxoviruses (Hyndman et al., 2013) and nidoviruses (Stenglein et al., 2014;Hoon-Hanks et al., 2018). Mycoplasma (Penner et al., 1997) and chlamydia (Hilf et al., 1990) have also been suggested as causes of respiratory disease in snakes. ...
Serpentoviruses (order Nidovirales) are an important cause of respiratory disease in snakes. Although transmission studies have shown that serpentoviruses can cause respiratory disease in pythons, the possible role of additional potential pathogens is not yet understood. Very little information is available on the role of mycoplasma and chlamydia infections in disease in pythons. Diagnostic samples from 271 pythons of different genera submitted to a laboratory for detection of serpentoviruses were also screened for mycoplasma and chlamydia infections by PCR. Most of the samples were oral swabs. Almost 30% of the samples were positive for serpentoviruses, and mycoplasmas were detected in more than 60% of the pythons. The occurrence of these two pathogens correlated significantly (P < 0.001). Additionally, about 3% of the snakes tested positive for Chlamydia. This study found a high prevalence of mycoplasmas in the tested pythons and a correlation between infections with these bacteria and serpentoviruses in python samples submitted for diagnostic testing. Because the role mycoplasmas play in respiratory diseases of snakes is still largely unknown, further investigations are necessary to evaluate the role of mixed infections in disease.
Rift Valley fever virus (RVFV) is a mosquito‐borne pathogen with significant human and veterinary health consequences that periodically emerges in epizootics. RVFV causes fetal loss and death in ruminants and in humans can lead to liver and renal disease, delayed‐onset encephalitis, retinitis, and in some cases severe hemorrhagic fever. A live attenuated vaccine candidate (DDVax), was developed by the deletion of the virulence factors NSs and NSm from a clinical isolate, ZH501, and has proven safe and immunogenic in rodents, pregnant sheep and non‐human primates. Deletion of NSm also severely restricted mosquito midgut infection and inhibited vector‐borne transmission. To demonstrate environmental safety, this study investigated the replication, dissemination and transmission efficiency of DDVax in mosquitoes following oral exposure compared to RVFV strains MP‐12 and ZH501. Infection and dissemination profiles were also measured in mosquitoes 7 days after they fed on goats inoculated with DDvax or MP‐12. We hypothesized that DDVax would infect mosquitoes at significantly lower rates than other RVFV strains and, due to lack of NSm, be transmission incompetent. Exposure of Ae. aegypti and Cx. tarsalis to 8 log10 plaque forming units (PFU)/mL DDVax by artificial bloodmeal resulted in significantly reduced DDVax infection rates in mosquito bodies compared to controls. Plaque assays indicated negligible transmission of infectious DDVax in Cx. tarsalis saliva (1/140 sampled) and none in Ae aegypti saliva (0/120). Serum from goats inoculated with DDVax or MP‐12 did not harbor detectable infectious virus by plaque assay at 1, 2, or 3 days‐post‐inoculation. Infectious virus was, however, recovered from Aedes and Culex bodies that fed on goats vaccinated with MP‐12 (13.8% and 4.6%, respectively), but strikingly, DDvax positive mosquito bodies were greatly reduced (4%, and 0%, respectively). Furthermore, DDVax did not disseminate to legs/wings in any of the goat‐fed mosquitoes. Collectively, these results are consistent with a beneficial environmental safety profile. This article is protected by copyright. All rights reserved
Although reptiles have often been overlooked in research, information on viruses of reptiles has been growing steadily in recent decades as has our understanding of the importance of these animals in the ecosystem. As ectotherms, their immune systems are dependent on temperature, among other factors, and interactions between infection and disease are complex and dependent on host, pathogen, and environmental factors. This chapter provides an overview of the viruses described in reptiles so far, as well as insight into some of the diseases caused by viruses in this group of animals. It also discusses the reptile immune system and the host reaction to infection. Influences of the environment on development of disease are in many cases not well understood, and this chapter includes a discussion of some important progress in this field. Studies of the effects of viruses on wild, pet, and farmed reptiles are limited, but indicate that viral disease can strongly affect individual populations in the wild, and that human action and the animal trade likely play a role in disease epidemiology.
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Inclusion body disease (IBD) is an infectious disease originally described in captive snakes. It has traditionally been diagnosed by the presence of large eosinophilic cytoplasmic inclusions and is associated with neurological, gastrointestinal, and lymphoproliferative disorders. Previously, we identified and established a culture system for a novel lineage of arenaviruses isolated from boa constrictors diagnosed with IBD. Although ample circumstantial evidence suggested that these viruses, now known as reptarenaviruses, cause IBD, there has been no formal demonstration of disease causality since their discovery. We therefore conducted a long-term challenge experiment to test the hypothesis that reptarenaviruses cause IBD. We infected boa constrictors and ball pythons by cardiac injection of purified virus. We monitored the progression of viral growth in tissues, blood, and environmental samples. Infection produced dramatically different disease outcomes in snakes of the two species. Ball pythons infected with Golden Gate virus (GoGV) and with another reptarenavirus displayed severe neurological signs within 2 months, and viral replication was detected only in central nervous system tissues. In contrast, GoGV-infected boa constrictors remained free of clinical signs for 2 years, despite high viral loads and the accumulation of large intracellular inclusions in multiple tissues, including the brain. Inflammation was associated with infection in ball pythons but not in boa constrictors. Thus, reptarenavirus infection produces inclusions and inclusion body disease, although inclusions per se are neither necessarily associated with nor required for disease. Although the natural distribution of reptarenaviruses has yet to be described, the different outcomes of infection may reflect differences in geographical origin. IMPORTANCE New DNA sequencing technologies have made it easier than ever to identify the sequences of microorganisms in diseased tissues, i.e., to identify organisms that appear to cause disease, but to be certain that a candidate pathogen actually causes disease, it is necessary to provide additional evidence of causality. We have done this to demonstrate that reptarenaviruses cause inclusion body disease (IBD), a serious transmissible disease of snakes. We infected boa constrictors and ball pythons with purified reptarenavirus. Ball pythons fell ill within 2 months of infection and displayed signs of neurological disease typical of IBD. In contrast, boa constrictors remained healthy over 2 years, despite high levels of virus throughout their bodies. This difference matches previous reports that pythons are more susceptible to IBD than boas and could reflect the possibility that boas are natural hosts of these viruses in the wild.
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This article lists the changes to virus taxonomy approved and ratified by the International Committee on Taxonomy of Viruses (ICTV) in March 2017.
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Current knowledge of RNA virus biodiversity is both biased and fragmentary, reflecting a focus on culturable or disease-causing agents. Here we profile the transcriptomes of over 220 invertebrate species sampled across nine animal phyla and report the discovery of 1,445 RNA viruses, including some that are sufficiently divergent to comprise new families. The identified viruses fill major gaps in the RNA virus phylogeny and reveal an evolutionary history that is characterized by both host switching and co-divergence. The invertebrate virome also reveals remarkable genomic flexibility that includes frequent recombination, lateral gene transfer among viruses and hosts, gene gain and loss, and complex genomic rearrangements. Together, these data present a view of the RNA virosphere that is more phylogenetically and genomically diverse than that depicted in current classification schemes and provide a more solid foundation for studies in virus ecology and evolution.
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A respiratory disease syndrome has been observed in large numbers of wild shingleback lizards (Tiliqua rugosa) admitted to wildlife care facilities in the Perth metropolitan region of Western Australia. Mortality rates are reportedly high without supportive treatment and care. Here we used next generation sequencing techniques to screen affected and unaffected individuals admitted to Kanyana Wildlife Rehabilitation Centre in Perth between April and December 2015, with the resultant discovery of a novel nidovirus significantly associated with cases of respiratory disease according to a case definition based on clinical signs. Interestingly this virus was also found in 12% of apparently healthy individuals, which may reflect testing during the incubation period or a carrier status, or it may be that this agent is not causative in the disease process. This is the first report of a nidovirus in lizards globally. In addition to detection of this virus, characterisation of a 23,832 nt segment of the viral genome revealed the presence of characteristic nidoviral genomic elements providing phylogenetic support for the inclusion of this virus in a novel genus alongside Ball Python nidovirus, within the Torovirinae sub-family of the Coronaviridae. This study highlights the importance of next generation sequencing technologies to detect and describe emerging infectious diseases in wildlife species, as well as the importance of rehabilitation centres to enhance early detection mechanisms through passive and targeted health surveillance. Further development of diagnostic tools from these findings will aid in detection and control of this agent across Australia, and potentially in wild lizard populations globally.
In 2014 we observed a noticeable increase in sudden deaths of green tree pythons (Morelia 28 viridis). Pathological examination revealed accumulation of mucoid material within airways and lung, associated with enlargement of the entire lung. We performed full necropsy and histological examination on affected green tree pythons from different breeders to characterise the pathogenesis of this “mucinous” pneumonia. By histology we could show a marked hyperplasia of the airway epithelium and of faveolar type II pneumocytes. Since routine microbiological tests failed to identify a causative agent, we studied lung samples of a few diseased snakes by next-generation sequencing (NGS). From the NGS data we could assemble a piece of RNA genome <85% identical to nidoviruses previously identified in ball pythons and Indian pythons. We then employed RT-PCR to demonstrate the presence of the novel nidovirus in all diseased snakes. To attempt virus isolation, we established primary cell cultures of Morelia viridis liver and brain, which we inoculated with lung homogenates of infected individuals. Ultrastructural examination of concentrated cell culture supernatants showed the presence of nidovirus particles, and subsequent NGS analysis yielded the full genome of the novel virus, Morelia viridis nidovirus (MVNV). We then generated an antibody against MVNV nucleoprotein, which we used alongside RNA in situ hybridisation to demonstrate viral antigen and RNA in the affected lungs. This suggests that in natural infection MVNV damages the respiratory tract epithelium which then results in epithelial hyperplasia, most likely as an exaggerated regenerative attempt in association with increased epithelial turnover.
OBJECTIVES Nidoviruses have recently been described as a putative cause of severe respiratory disease in pythons in the USA and Europe. The objective of this study was to establish the use of a conventional PCR for the detection of nidoviruses in samples from live animals and to extend the list of susceptible species. MATERIALS AND METHODS A PCR targeting a portion of ORF1a of python nidoviruses was used to detect nidoviruses in diagnostic samples from live boas and pythons. A total of 95 pythons, 84 boas and 22 snakes of unknown species were included in the study. Samples tested included oral swabs and whole blood. RESULTS Nidoviruses were detected in 27.4% of the pythons and 2.4% of the boas tested. They were most commonly detected in ball pythons (Python [P.] regius) and Indian rock pythons (P. molurus), but were also detected for the first time in other python species, including Morelia spp. and Boa constrictor. Oral swabs were most commonly tested positive. CONCLUSION The PCR described here can be used for the detection of nidoviruses in oral swabs from live snakes. These viruses appear to be relatively common among snakes in captivity in Europe and screening for these viruses should be considered in the clinical work-up. CLINICAL RELEVANCE Nidoviruses are believed to be an important cause of respiratory disease in pythons, but can also infect boas. Detection of these viruses in live animals is now possible and can be of interest both in diseased animals as well as in quarantine situations.