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Shelter-housed cats show no evidence of faecal shedding of canine parvovirus DNA
The Veterinary Journal 239 (2018) 54–58
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Original Article
Shelter-housed cats show no evidence of faecal shedding of canine parvovirus DNA P. Byrnea , J.A. Beattya,b , J. Šlapetaa , S.W. Corleyc , R.E. Lyonsc , L. McMichaelc, M.T. Kyaw-Tannerc, P.T. Dungc , N. Decarod , J. Meersc, V.R. Barrsa,b,* a
University of Sydney, Sydney School of Veterinary Science, Faculty of Science, NSW 2006, Australia University of Sydney, Marie Bashir Institute, NSW 2006 Australia University of Queensland, School of Veterinary Science, Gatton, QLD 4343, Australia d University of Bari, Department of Veterinary Medicine, Valenzano (Bari), Italy b c
A R T I C L E I N F O
A B S T R A C T
Keywords: Canine Feline Panleukopenia Parvovirus Shelter
Canine parvovirus (CPV) and feline panleukopenia virus (FPV) are deoxyriboncucleic acid (DNA) viruses in the taxon Carnivore protoparvovirus 1. Exposure of cats to either CPV or FPV results in productive infection and faecal shedding of virus. Asymptomatic shedding of CPVs by one-third of shelter-housed cats in a UK study suggests that cats may be an important reservoir for parvoviral disease in dogs. The aim of this cross-sectional study was to determine the prevalence of faecal shedding of CPVs in asymptomatic shelter-housed cats in Australia. Faecal samples (n = 218) were collected from cats housed in three shelters receiving both cats and dogs, in Queensland and NSW. Molecular testing for Carnivore protoparvovirus 1 DNA was performed by polymerase chain reaction (PCR) amplification followed by DNA sequencing of the VP2 region to differentiate CPV from FPV. Carnivore protoparvovirus 1 DNA was detected in only four (1.8%, 95% confidence interval 0.49–4.53%) faecal samples from a single shelter. Sequencing identified all four positive samples as FPV. Faecal shedding of CPV by shelter-cats was not detected in this study. While the potential for cross-species transmission of CPV between cats and dogs is high, this study found no evidence of a role for cats in maintaining CPV in cat and dog populations through faecal shedding in the regions tested. © 2018 Elsevier Ltd. All rights reserved.
Introduction Canine parvovirus (CPV) and feline panleukopenia virus (FPV) are small, (5 Kb) single-stranded, icosahedral DNA viruses in the taxon Carnivore protoparvovirus 1. CPV and FPV share greater than 98% nucleotide identity and both viruses can infect and cause disease in felids (Hoelzer and Parrish, 2010). Viral host range and pathogenicity are determined by binding of the viral VP2 capsid region to cellular transferrin receptors (TfR) (Allison et al., 2013). Frequent cross-species transmission of parvoviruses between wildlife carnivore species suggests that CPV and FPV have evolved independently from common sylvatic ancestors (Allison et al., 2013; Allison et al., 2014). The clinical syndrome of feline panleukopenia, characterised by enteritis and panleukopenia, has been recognised in cats for over a
* Corresponding author. E-mail address: [email protected] (V.R. Barrs). https://doi.org/10.1016/j.tvjl.2018.08.005 1090-0233/© 2018 Elsevier Ltd. All rights reserved.
hundred years (Hoelzer and Parrish, 2010), although it was not until 1965 that FPV was first isolated (Johnson, 1965). When CPV emerged as a new virus in dogs in the mid-to-late 1970s, the virus, termed CPV-2, was unable to infect cats (Parrish, 1999). However, by 1980 a new variant, CPV-2a, had emerged, replacing CPV-2 that had acquired the feline host range through four mutations in VP2 (L87M, I101T, A300G, D305Y) that likely enable capsid binding to the feline TfR. Two additional CPV variants VP2 N426D (CPV-2b) and VP2 D426E (CPV-2c) have since emerged that co-circulate with CPV-2a in varying proportions in different geographic regions (Hoelzer and Parrish, 2010; Decaro and Buonavoglia, 2012). CPV variants have been isolated from the blood (Ikeda et al., 2000) and faeces of cats worldwide, in both natural (Mochizuki et al., 1993; Mochizuki et al., 1996; Truyen et al., 1996; Gamoh et al., 2003a; Battilani et al., 2006; Battilani et al., 2011) and experimental (Nakamura et al., 2001) infections. While FPV is the most common aetiological agent of feline panleukopenia, CPVs can cause clinical signs in cats that are indistinguishable from those caused by FPV (Stuetzer and Hartmann, 2014). DNA sequencing of
P. Byrne et al. / The Veterinary Journal 239 (2018) 54–58
parvovirus isolates from cats presenting with signs of feline panleukopenia has revealed most to be FPV, with CPVs causing approximately 5% of feline panleukopenia cases (Truyen et al., 1996). Clinical disease caused by CPV variants have been reported in Italy, Germany, USA, Japan and in Portugal (Mochizuki et al., 1996; Truyen et al., 1996; Gamoh et al., 2003b; Decaro et al., 2010; Miranda et al., 2014). In the UK, faecal shedding of CPV was detected in more than a third of faecal samples collected from healthy cats from two mixed canine and feline shelters (Clegg et al., 2012). The ability of CPV variants to infect cats has raised concerns about the role of cats as reservoirs of infection for dogs, and has important implications for biosecurity especially in mixed animal shelters housing both cats and dogs (Clegg et al., 2012). The aim of this study was to determine the prevalence of faecal shedding of CPV in asymptomatic shelter-housed cats to understand whether faecal shedding of CPV by cats poses a significant infection risk in mixed cat-dog shelters in the regions tested. Materials and methods Samples Shelters 1 and 2 Faecal samples (n = 118) were collected from healthy cats from two mixed catdog rescue shelters in Brisbane, Queensland within 24 h of admission to the shelter. Cats were individually housed except for litters where up to three litter-mates shared an enclosure. On entry to the shelter all cats were administered an attenuated trivalent vaccination containing feline herpesvirus-1, feline calicivirus and feline panleukopenia virus (Companion F3, MSD Animal Health Australia). Sixty-one faecal samples were collected from Shelter 1 on four occasions between 19th October 2015 and 1st February 2016 and 57 faecal samples were collected from Shelter 2 on three occasions between 4th December 2015 and 20th January 2016. The holding capacity of shelter 1 was 180 dogs and 150 cats, and for shelter 2 was 110 dogs and 120 cats. At both shelters the majority of cats were housed one to a cage, but there were a few group housing rooms. Veterinary shelter personnel could move freely between canine and feline areas of the shelter. Shelter 3 Faecal samples (n = 100) were collected from a mixed cat-dog shelter in Sydney, NSW on three occasions at any time point after admission to the shelter (March 2016–February 2017). An outbreak of feline panleukopenia occurred at this shelter in the two months prior to the third sample collection. On admission to the shelter dogs were routinely administered two inactivated canine parvovirus vaccines, 7 days apart (Parvac, Zoetis Australia, Silverwater Australia). Cats were not vaccinated. Healthy cats were housed individually or in groups of up to six cats in enclosures (n = 14). One faecal sample was collected from individually caged cats while all faecal samples were collected from multicat enclosures. In some multicat enclosures the number of faecal samples collected exceeded the number of cats because individual cats had defecated more than once since the litter trays had been cleaned. The shelter also housed approximately 100 dogs. All shelter personnel moved freely between canine and feline areas of the shelter. At all shelters sampled, enclosures were cleaned daily and litter trays were emptied and cleaned at least once daily. At each visit, fresh faecal samples were collected from the litter tray or enclosure floor prior to cleaning. Data including age, sex, breed and date of admission of the cats was obtained at each visit. Faecal samples were stored at 20 C or 80 C until processing.
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Molecular detection of CPV and FPV from faeces Shelters 1 and 2 DNA was extracted from faecal samples using the QIAmp DNA Stool Mini Kit (Qiagen) to remove PCR inhibitors as described previously, and stored at 20 C until PCR testing (Meers et al., 2007). A duplicate set of DNA extractions was performed using boiling and homogenization, as described previously and stored at 4 C for PCR testing within 7 days of extraction (Clegg et al., 2012). The presence of parvovirus in faecal samples was detected in duplicate DNA samples extracted by both methods using a conventional PCR to amplify the VP2 gene of Carnivore protoparvovirus 1, using primer pair JS1F and JS2R as previously described (Table 1) (Meers et al., 2007). For the kit-extracted DNA samples, each 25 mL reaction contained 0.1 mL of 5U/mL KAPA Taq DNA polymerase (KAPA Biosystems Inc., Wilmington, MA), 2.5 mL KAPA Buffer A (containing 15 mM MgCl2), 0.5 mL of 10 mM dNTP mix, 0.3 mL DMSO (final concentration of 1.5%), 1.25 mL betaine (final concentration 0.25 mM), 1 mL of 0.2 mmol of each primer (10 mM each) and 2 mL DNA template. For the DNA extracted by homogenization and boiling, each 25 mL reaction contained 2.5 mL of 10 reaction buffer (containing 15 mM MgCl2), 0.5 mL of 25 mM MgCl2, 4 mL of 1.25 mM dNTPs, 10 pmol of each primer, 0.5 mL of 5U/mL HotStar Taq DNA polymerase (Qiagen Australia) and 5 mL DNA template. Cycling conditions were initial denaturation at 94 C for 5 min, followed by 40 cycles of denaturation at 94 C for 30 s, annealing at 52 C for 30 s and polymerization at 72 C for 2 min. A negative control (no DNA) and a positive control were included in each run. Positive controls were faecal DNAs from a dog with parvovirus enteritis infected with CPV-2b and/or from a cat with feline panleukopenia infected with FPV diagnosed using the methods above. Shelter 3 DNA was extracted from faeces by homogenization and boiling as described previously, (Clegg et al., 2012). The resulting supernatants were diluted in molecular-grade water (1:10, 1:20, and 1:50) and stored at 4 C for PCR testing within 7 days of extraction. To test for PCR inhibition, DNA extracts (1:10 and 1:20 dilutions) were spiked with 1 mL of feline genomic DNA (138 ng/mL) extracted from the lymph node of a healthy cat. A conventional PCR using primer set GAPDH-F and GAPDH-R to amplify the feline housekeeping GAPDH gene was performed (Table 1) (McLuckie et al., 2016). Each 50 mL reaction contained 5 mL of 10 X PCR buffer (containing 10 mMol dNTP and 75 mM MgCl2), 1 mL (10 pmol) of each primer, 0.2 mL (0.2 U) of Taq QIAGEN DNA Polymerase (QIAGEN Germantown, MD), 1 mL of feline genomic DNA template and 5 mL faecal DNA. Cycling conditions were initial denaturation at 94 C for 3 min, followed by 35 cycles of denaturation at 94 C for 45 s, annealing at 55 C for 30 s and polymerization at 72 C for 30 s, with final extension 72 C for 7 min. If PCR inhibition was detected additional dilutions of the sample were tested to identify the dilution at which PCR inhibition was abolished. The presence of parvovirus in each faecal sample was detected using a conventional PCR to amplify the VP2 gene using primer pair 555-F and 555-R (Decaro et al., 2008) (Table 1). PCR screening of all faecal samples was performed using both 1:10 and 1:20 dilutions, as well as 1:50 dilutions for samples in which PCR inhibition was detected. Each 50 mL reaction contained 10 mL PCR buffer (5 mM dNTPs, 15 mM MgCl2), 0.5 mL (0.5U) MyTaq Hot Start polymerase (Bioline, Meriden Life Science, Alexandria, Australia), 1 mL (10 pmol) forward and reverse primer, 7 mL DNA and 30.5 mL water. Cycling conditions were initial denaturation at 94 C for 1 min, followed by 35 cycles of denaturation at 94 C for 30 s, annealing at 55 C for 30 s and polymerization at 72 C for 30 s, with final extension 72 C for 1 min. PCR was performed using sample dilutions that had no evidence of PCR inhibition. A blank DNA control (PBS used during the extraction process) and a positive control were included in each run. The positive control was faecal DNA from a dog with parvovirus enteritis extracted and sequenced using the methods above, and identified as CPV-2a. The sensitivity of this conventional PCR was determined using DNA extracted from a feline faecal sample confirmed to contain FPV by sequencing. The FPV copy
Table 1 Primers used in this study for PCR amplification of feline DNA (GAPDH) and of canine and feline parvovirus DNA, targeting the VP2 region (VP2). Primer name
Target
Sequence
Size (nta )
Reference
JS1F JS2F GAPDH-F GAPDH-R 555-F 555-R 2655F 3511R 3381F 4116R
VP2
50 -AGCTACAGGATCTGGGAACG-30 50 -CCACCCACACCATAACAACA-30 50 -AAGGCTGAGAACGGGAAAC-30 50 -CATTTGATGTTGGCGGGATC-30 50 -CAGGAAGATATCCAGAAGGAA-30 50 -GGTGCTAGTTGATATGTAATAAACA-30 50 -CCAGATCATCCATCAACATCA-30 50 -TGAACATCATCTGGATCTGTACC-30 50 -CCATGGAAACCAACCATACC-30 50 -AGTTAATTCCTGTTTTACCTCCAA-30
1975
Meers et al. (2007)
80
McLuckie et al. (2016)
583
Decaro et al. (2008)
837
Decaro et al. (2008
717
Decaro et al., 2008
a
Nucleotide.
GAPDH VP2 VP2 VP2
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P. Byrne et al. / The Veterinary Journal 239 (2018) 54–58
number per gram of faeces was determined for this sample by quantitative PCR (qPCR), using a previously validated multiplex assay (Meggiolaro et al., 2017), and was 3 108 FPV copies per gram of faeces. An end-point dilution of the FPV positive faecal DNA was performed using serial ten-fold dilutions (from 101 to 109) in the conventional VP2 PCR (primer pair 555F–555R). The second highest dilution at which a positive result was detected on gel electrophoresis was considered to be the cPCR sensitivity threshold. Sequence and phylogenetic analysis PCR products were detected by electrophoresis (1% agarose gel) and visualization under UV light. To differentiate CPV from FPV, the remainder of the VP2 gene was amplified in positive samples using primers 3381F–4116R, and 2679F–3511R (Decaro et al., 2008) (Table 1). Sanger sequencing was performed at a commercial laboratory (Macrogen Seoul, Korea). DNA sequence chromatographs were visually inspected and assembled in CLC Workbench. Viral subtype (FPV, CPV 300G (CPV-2a), CPV 426D (CPV-2b) or CPV 426E (CPV-2c)) was determined from key amino acid substitutions (Decaro and Buonavoglia, 2012). Unique strains identified in this study were deposited on GenBank (submission #2092491).
Results A total of 118 faecal samples were collected from shelters 1 and 2 in Brisbane. Four of these samples were collected from cages that held 2 or 3 litter-mates. Of 66 samples for which age was recorded, the median age of the cats was 12 months (range 4 weeks to 7 years), with 52% of cats being 12 months of age or younger. Of 87 cats for which sex was recorded, 42 were male and 45 were female. From shelter 3 in Sydney 100 faecal samples were obtained from 79 cats. The median age of the cats was 4 months (range 5 weeks to 5 years) and the majority of cats for which age was recorded (66/79, 84%), were 12 months of age or younger. Of 77 cats for which sex was recorded, there were 29 males and 48 females. Of 100 faecal DNA samples from Sydney in which PCR inhibition was tested, six had PCR inhibition at a 1:10 dilution, which was abolished at a 1:20 dilution in five samples and at a 1:50 dilution in one sample. Quantification of the number of viral copies present in the known FPV positive control sample, using the qPCR assay was determined to be 299, 290 copies/g of faeces. The end-point dilution study of the conventional FPV/CPV VP2 PCR assay revealed a 1 105 limit of detection, which was repeatable (Fig. 1). Thus, the limit of sensitivity of detection of this conventional PCR assay was 300 viral copies/gram (Fig. 1) of faeces or 15 viral copies/1 mL template. Carnivore protoparvovirus 1 DNA was detected in 4/218 (1.8%, 95% confidence interval 0.49–4.53%) samples from all three shelters by conventional PCR. The four positive samples were collected from four separate enclosures in shelter 3 on the third sampling occasion, after an outbreak of feline panleukopenia had
Fig. 1. A dilution assay was performed using a feline panleukopenia virus-positive faecal DNA sample containing 299, 290 viral copies/g faeces, as determined by a validated qPCR assay to calculate the sensitivity of the conventional PCR assay used to detect Carnivore protoparvovirus1 in shelter 3.
occurred within the previous two months at this shelter. Sequencing of the VP2-gene showed that VP2 key residues typical of FPV were present in all obtained sequences, so that the four parvovirus strains were identified as FPV. The FPV strains were identical to the outbreak strain detected in sick cats in a separate investigation (Barrs et al., 2017). In contrast, CPV shedding was not detected in any samples from the three shelters in this study. Discussion Cats are susceptible to subclinical or clinical infection by CPV variants including CPV 300G (CPV-2a), CPV 426D (CPV-2b) and CPV 426E (CPV-2c) (Nakamura et al., 2001; Gamoh et al., 2003a). This has been demonstrated in experimental infections, in which CPV-2 was unable to replicate in feline tissues but CPV 300G (CPV-2a) and CPV 426D (CPV-2b) underwent efficient replication (Truyen et al., 1994; Nakamura et al., 2001), as well as in naturally occurring infections in which CPV-2a, CPV-2b and CPV-2c but not CPV-2 isolates were detected in cats with clinical signs similar to those caused by FPV infection (Mochizuki et al., 1996; Truyen et al., 1996; Gamoh et al., 2003b; Decaro et al., 2010; Miranda et al., 2014). The absence of faecal CPV shedding in this cohort of healthy Australian shelter-housed cats contrasts the UK study where 26% of shelterhoused cats from one shelter, sampled at a single time point, shed CPV in their faeces (Clegg et al., 2012). In that study, longitudinal sampling was performed in a second shelter, demonstrating that 34 of 74 cats (46%) shed CPV in faeces on at least one occasion over an 8-week period, while neither 122 canine faecal samples or 160 environmental samples from that shelter were positive (Clegg et al., 2012). Our results align more closely with those of a US investigation, where quantitative PCR was used to detect faecal shedding of parvoviruses in healthy shelter cats (Leutenegger et al., 2015). In that study although 58% of cats were found to be shedding Carnivore protoparvovirus 1, only 21% of these (12% overall) had high parvoviral loads consistent with active infection and <1% of cats overall were shedding high viral loads of CPV. The reasons why active CPV infections are highly prevalent in cats entering shelters in the UK while none were detected in Australian shelter-housed cats are not immediately obvious. The difference in the observed prevalence of CPV shedding between Australia and UK cannot be accounted for by methodological differences since similar DNA extraction and PCR methods were used and we also tested a larger number of cats (Clegg et al., 2012). DNA extracted from faeces frequently contains PCR inhibitors, especially at low dilutions (Schrader et al., 2012), which can be a cause of false negative PCR results. PCR inhibitors in faeces include complex polysaccharides, bile salts, lipids, urea and haemoglobin (Schrader et al., 2012). Additional steps were taken in this study to exclude PCR inhibition as a cause of negative results including the use of a commercial DNA extraction method or an extraction control. It is possible that low-level faecal CPV shedding was not detected using the cPCR assays in this study, although cPCR was also used in the UK study (Clegg et al., 2012). To explore this possibility, we determined the limit of detection of one cPCR assay, which was 45 viral copies/3 mL template. In comparison, a qPCR designed to detect FPV and CPV was reported to have a 95% limit of detection of 20 copies/3 mL DNA template (Freisl et al., 2017). In another investigation using the same qPCR protocol, faecal samples from cats and dogs with suspected parvovirus infection were tested using IFA, conventional PCR, and qPCR. A marked difference in sensitivity between the three assays was detected, with 12%, 32% and 58% of samples testing positive for FPV or CPV by IFA, cPCR and qPCR, respectively (Streck et al., 2013). Quantitative results of CPV viral loads shed in faeces by healthy cats as determined by qPCR have not as yet been published. CPV loads
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shed in faeces in dogs after administration of a modified live CPV vaccine were 3.87 102–8.39 105 viral copies/gram of faeces, which were 4–5 fold lower than CPV loads shed in faeces after infection with field strains of CPV (Decaro et al., 2005; Freisl et al., 2017). If CPV faecal loads in cats are similar to dogs, then it is unlikely that cats undergoing active CPV infections were not detected in this study since the limit of detection was 3 102 viral copies/gram. However, to further investigate CPV shedding in asymptomatic cats, use of an optimized qPCR would be prudent. As expected, vaccine strains of FPV were not detected in the faeces of any cats from shelters 1 and 2, since the incubation period for FPV is usually at least two days (Csiza et al., 1971a,b). While the cats tested here were not shedding CPV, our results do not exclude that these cats might have been latently infected with CPV. CPV DNA has been detected by PCR from the bone marrow aspirates of healthy unowned Australian cats (Haynes and Holloway, 2012). Carnivore protoparvovirus 1 becomes latent in peripheral blood mononuclear cells (PBMC) after viral shedding ceases, leaving behind a convenient molecular “footprint” of a previous active infection (Miyazawa et al., 1999; Allison et al., 2013; Allison et al., 2014). There were several limitations to our study. Investigations of CPV-shedding in Brisbane (Shelters 1 and 2) and Sydney (Shelter 3) were initiated independently and without knowledge of each other, hence the methodological differences in primers used for Carnivore protoparvovirus 1 amplification, DNA extraction methods and how PCR inhibition was addressed. Clegg et al. (2012), used primer EF (2748–2765) and JS2R (Table 1) for VP2 amplification of parvoviruses from shelter cat faeces as well as slightly different PCR conditions to what were used in our studies (Clegg et al., 2011). It is possible that the sensitivity of the conventional PCR used by Clegg et al. (2011, 2012) was higher than the PCRs we used. Canine parvovirus is a common cause of enteritis in dogs worldwide. In dogs with acute, severe diarrhoea presenting to charity-sponsored primary care veterinary clinics in the UK, CPV was detected in 58% of 355 faecal samples (Godsall et al., 2010). In Australia it is estimated that 20,000 new cases of canine parvovirus enteritis are presented to veterinarians annually, using data collected in a national epidemiological survey (Kelman et al., 2017). Clusters of CPV-associated disease in Australia occur more commonly in areas with lower socioeconomic indices (Brady et al., 2012). While it is possible that we did not detect CPV shedding in shelter cats because they had not been exposed to CPV, Shelter 3 is located in a CPV “hot spot” suggesting that environmental exposure of free roaming cats presented to the shelter to CPV is not unlikely. Other potential contributors to the disparity between shedding of CPV by shelter cats in UK compared with Australia include differences in population density, territory size and climactic factors could lead to behavioural differences between cat populations, such as frequency of interaction with dogs and other cats resulting in different frequencies of exposure to CPV. Frequent cross-species transmission events between domestic and wildlife carnivores have been implicated in the evolution of CPVs in regions of the world with endemic carnivore species diversity (Allison et al., 2014). The role of wildlife carnivore contact in transmission of CPV to cats has not been investigated. Canine parvoviruses (CPVs) can infect multiple different families of carnivores in both Carnivora suborders, Feliformia and Caniformia (Allison et al., 2013). The diversity of carnivores in Australia and New Zealand is low, being limited to a few species in the Felidae, Canidae and Mustelidae families such as the domestic cat, dog and ferret. In Australia there is only one endemic terrestrial carnivore, the dingo, while in New Zealand there are none. The extent of carnivore species diversity in the UK is higher than Australia and New Zealand but lower than that in Europe and North America. Molecular surveillance of CPVs in Australia and New Zealand, has
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shown a ‘lag’ in CPV evolution, with CPV-2a being reported as the dominant subtype in Australia until 2011–2013, and the only subtype in NZ until at least 2009, likely reflecting both geographical isolation and the absence of endemic species which facilitate viral evolution (Ohneiser et al., 2015; Clark et al., 2018). Although we found no evidence of CPV shedding in shelter cats in Australia, cats can be infected by and shed CPV and can develop clinical disease indistinguishable from that caused by FPV. Considering that CPV-induced clinical disease has been reported in cats (Decaro et al., 2010; Miranda et al., 2014), stringent disinfection and biosecurity protocols that prevent both intra- and cross-species infectious diseases are essential for animal welfare in shelters. Conclusions Overall, the prevalence of faecal shedding of Carnivore protoparvovirus 1 in shelter-housed cats from three shelters in Australia was low. CPV shedding by shelter cats was not detected. Conflict of interest statement None of the authors of this paper has a financial or personal relationship with other people or organisations that could inappropriately influence or bias the content of the paper. Acknowledgements This study was supported by the June Rose Bullock bequest at the University of Sydney, the NSW Cat Protection Society, Boehringer Ingelheim, and MSD Animal Health. None of the organizations providing financial support were involved in the design or execution of the study. The authors thank staff at all of the shelters for their time and efforts in assisting with sample collection. References Allison, A.B., Kohler, D.J., Fox, K.A., Brown, J.D., Gerhold, R.W., Shearn-Bochsler, V.I., Dubovi, E.J., Parrish, C.R., Holmes, E.C., 2013. Frequent cross-species transmission of parvoviruses among diverse carnivore hosts. Journal of Virology 87, 2342–2347. Allison, A.B., Kohler, D.J., Ortega, A., Hoover, E.A., Grove, D.M., Holmes, E.C., Parrish, C.R., 2014. Host-specific parvovirus evolution in nature is recapitulated by in vitro adaptation to different carnivore species. PLoS Pathogens 10, e1004475. Barrs, V.R., Brailey, J., Allison, A.B., Kelman, M., Meers, J., Beatty, J.A., Holmes, E.C., 2017. Re-emergence of feline panleukopenia in Australia. 27th ECVIM-CA Congress, St Julian’s, Malta September 14–16, ISCAID_0_4. Battilani, M., Balboni, A., Ustulin, M., Giunti, M., Scagliarini, A., Prosperi, S., 2011. Genetic complexity and multiple infections with more Parvovirus species in naturally infected cats. Veterinary Research 42, 43. Battilani, M., Scagliarini, A., Ciulli, S., Morganti, L., Prosperi, S., 2006. High genetic diversity of the VP2 gene of a canine parvovirus strain detected in a domestic cat. Virology 352, 22–26. Brady, S., Norris, J.M., Kelman, M., Ward, M.P., 2012. Canine parvovirus in Australia: the role of socio-economic factors in disease clusters. Veterinary Journal 193, 522–528. Clark, N.J., Seddon, J.M., Kyaw-Tanner, M., Al-Alawneh, J., Harper, G., McDonagh, P., Meers, J., 2018. Emergence of canine parvovirus subtype 2b (CPV-2b) infections in Australian dogs. Infection, Genetics and Evolution: Journal of Molecular Epidemiology and Evolutionary Genetics in Infectious Diseases 58, 50–55. Clegg, S.R., Coyne, K.P., Dawson, S., Spibey, N., Gaskell, R.M., Radford, A.D., 2012. Canine parvovirus in asymptomatic feline carriers. Veterinary Microbiology 157, 78–85. Clegg, S.R., Coyne, K.P., Parker, J., Dawson, S., Godsall, S.A., Pinchbeck, G., Cripps, P.J., Gaskell, R.M., Radford, A.D., 2011. Molecular epidemiology and phylogeny reveal complex spatial dynamics in areas where canine parvovirus is endemic. Journal of Virology 85, 7892–7899. Csiza, C.K., De Lahunta, A., Scott, F.W., Gillespie, J.H., 1971a. Pathogenesis of feline panleukopenia virus in susceptible newborn kittens II. Pathology and immunofluorescence. Infection and Immunity 3, 838–846. Csiza, C.K., Scott, F.W., De Lahunta, A., Gillespie, J.H., 1971b. Pathogenesis of feline panleukopenia virus in susceptible newborn kittens I. Clinical signs hematology, serology, and virology. Infection and Immunity 3, 833–837.
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Original Article
Shelter-housed cats show no evidence of faecal shedding of canine parvovirus DNA P. Byrnea , J.A. Beattya,b , J. Šlapetaa , S.W. Corleyc , R.E. Lyonsc , L. McMichaelc, M.T. Kyaw-Tannerc, P.T. Dungc , N. Decarod , J. Meersc, V.R. Barrsa,b,* a
University of Sydney, Sydney School of Veterinary Science, Faculty of Science, NSW 2006, Australia University of Sydney, Marie Bashir Institute, NSW 2006 Australia University of Queensland, School of Veterinary Science, Gatton, QLD 4343, Australia d University of Bari, Department of Veterinary Medicine, Valenzano (Bari), Italy b c
A R T I C L E I N F O
A B S T R A C T
Keywords: Canine Feline Panleukopenia Parvovirus Shelter
Canine parvovirus (CPV) and feline panleukopenia virus (FPV) are deoxyriboncucleic acid (DNA) viruses in the taxon Carnivore protoparvovirus 1. Exposure of cats to either CPV or FPV results in productive infection and faecal shedding of virus. Asymptomatic shedding of CPVs by one-third of shelter-housed cats in a UK study suggests that cats may be an important reservoir for parvoviral disease in dogs. The aim of this cross-sectional study was to determine the prevalence of faecal shedding of CPVs in asymptomatic shelter-housed cats in Australia. Faecal samples (n = 218) were collected from cats housed in three shelters receiving both cats and dogs, in Queensland and NSW. Molecular testing for Carnivore protoparvovirus 1 DNA was performed by polymerase chain reaction (PCR) amplification followed by DNA sequencing of the VP2 region to differentiate CPV from FPV. Carnivore protoparvovirus 1 DNA was detected in only four (1.8%, 95% confidence interval 0.49–4.53%) faecal samples from a single shelter. Sequencing identified all four positive samples as FPV. Faecal shedding of CPV by shelter-cats was not detected in this study. While the potential for cross-species transmission of CPV between cats and dogs is high, this study found no evidence of a role for cats in maintaining CPV in cat and dog populations through faecal shedding in the regions tested. © 2018 Elsevier Ltd. All rights reserved.
Introduction Canine parvovirus (CPV) and feline panleukopenia virus (FPV) are small, (5 Kb) single-stranded, icosahedral DNA viruses in the taxon Carnivore protoparvovirus 1. CPV and FPV share greater than 98% nucleotide identity and both viruses can infect and cause disease in felids (Hoelzer and Parrish, 2010). Viral host range and pathogenicity are determined by binding of the viral VP2 capsid region to cellular transferrin receptors (TfR) (Allison et al., 2013). Frequent cross-species transmission of parvoviruses between wildlife carnivore species suggests that CPV and FPV have evolved independently from common sylvatic ancestors (Allison et al., 2013; Allison et al., 2014). The clinical syndrome of feline panleukopenia, characterised by enteritis and panleukopenia, has been recognised in cats for over a
* Corresponding author. E-mail address: [email protected] (V.R. Barrs). https://doi.org/10.1016/j.tvjl.2018.08.005 1090-0233/© 2018 Elsevier Ltd. All rights reserved.
hundred years (Hoelzer and Parrish, 2010), although it was not until 1965 that FPV was first isolated (Johnson, 1965). When CPV emerged as a new virus in dogs in the mid-to-late 1970s, the virus, termed CPV-2, was unable to infect cats (Parrish, 1999). However, by 1980 a new variant, CPV-2a, had emerged, replacing CPV-2 that had acquired the feline host range through four mutations in VP2 (L87M, I101T, A300G, D305Y) that likely enable capsid binding to the feline TfR. Two additional CPV variants VP2 N426D (CPV-2b) and VP2 D426E (CPV-2c) have since emerged that co-circulate with CPV-2a in varying proportions in different geographic regions (Hoelzer and Parrish, 2010; Decaro and Buonavoglia, 2012). CPV variants have been isolated from the blood (Ikeda et al., 2000) and faeces of cats worldwide, in both natural (Mochizuki et al., 1993; Mochizuki et al., 1996; Truyen et al., 1996; Gamoh et al., 2003a; Battilani et al., 2006; Battilani et al., 2011) and experimental (Nakamura et al., 2001) infections. While FPV is the most common aetiological agent of feline panleukopenia, CPVs can cause clinical signs in cats that are indistinguishable from those caused by FPV (Stuetzer and Hartmann, 2014). DNA sequencing of
P. Byrne et al. / The Veterinary Journal 239 (2018) 54–58
parvovirus isolates from cats presenting with signs of feline panleukopenia has revealed most to be FPV, with CPVs causing approximately 5% of feline panleukopenia cases (Truyen et al., 1996). Clinical disease caused by CPV variants have been reported in Italy, Germany, USA, Japan and in Portugal (Mochizuki et al., 1996; Truyen et al., 1996; Gamoh et al., 2003b; Decaro et al., 2010; Miranda et al., 2014). In the UK, faecal shedding of CPV was detected in more than a third of faecal samples collected from healthy cats from two mixed canine and feline shelters (Clegg et al., 2012). The ability of CPV variants to infect cats has raised concerns about the role of cats as reservoirs of infection for dogs, and has important implications for biosecurity especially in mixed animal shelters housing both cats and dogs (Clegg et al., 2012). The aim of this study was to determine the prevalence of faecal shedding of CPV in asymptomatic shelter-housed cats to understand whether faecal shedding of CPV by cats poses a significant infection risk in mixed cat-dog shelters in the regions tested. Materials and methods Samples Shelters 1 and 2 Faecal samples (n = 118) were collected from healthy cats from two mixed catdog rescue shelters in Brisbane, Queensland within 24 h of admission to the shelter. Cats were individually housed except for litters where up to three litter-mates shared an enclosure. On entry to the shelter all cats were administered an attenuated trivalent vaccination containing feline herpesvirus-1, feline calicivirus and feline panleukopenia virus (Companion F3, MSD Animal Health Australia). Sixty-one faecal samples were collected from Shelter 1 on four occasions between 19th October 2015 and 1st February 2016 and 57 faecal samples were collected from Shelter 2 on three occasions between 4th December 2015 and 20th January 2016. The holding capacity of shelter 1 was 180 dogs and 150 cats, and for shelter 2 was 110 dogs and 120 cats. At both shelters the majority of cats were housed one to a cage, but there were a few group housing rooms. Veterinary shelter personnel could move freely between canine and feline areas of the shelter. Shelter 3 Faecal samples (n = 100) were collected from a mixed cat-dog shelter in Sydney, NSW on three occasions at any time point after admission to the shelter (March 2016–February 2017). An outbreak of feline panleukopenia occurred at this shelter in the two months prior to the third sample collection. On admission to the shelter dogs were routinely administered two inactivated canine parvovirus vaccines, 7 days apart (Parvac, Zoetis Australia, Silverwater Australia). Cats were not vaccinated. Healthy cats were housed individually or in groups of up to six cats in enclosures (n = 14). One faecal sample was collected from individually caged cats while all faecal samples were collected from multicat enclosures. In some multicat enclosures the number of faecal samples collected exceeded the number of cats because individual cats had defecated more than once since the litter trays had been cleaned. The shelter also housed approximately 100 dogs. All shelter personnel moved freely between canine and feline areas of the shelter. At all shelters sampled, enclosures were cleaned daily and litter trays were emptied and cleaned at least once daily. At each visit, fresh faecal samples were collected from the litter tray or enclosure floor prior to cleaning. Data including age, sex, breed and date of admission of the cats was obtained at each visit. Faecal samples were stored at 20 C or 80 C until processing.
55
Molecular detection of CPV and FPV from faeces Shelters 1 and 2 DNA was extracted from faecal samples using the QIAmp DNA Stool Mini Kit (Qiagen) to remove PCR inhibitors as described previously, and stored at 20 C until PCR testing (Meers et al., 2007). A duplicate set of DNA extractions was performed using boiling and homogenization, as described previously and stored at 4 C for PCR testing within 7 days of extraction (Clegg et al., 2012). The presence of parvovirus in faecal samples was detected in duplicate DNA samples extracted by both methods using a conventional PCR to amplify the VP2 gene of Carnivore protoparvovirus 1, using primer pair JS1F and JS2R as previously described (Table 1) (Meers et al., 2007). For the kit-extracted DNA samples, each 25 mL reaction contained 0.1 mL of 5U/mL KAPA Taq DNA polymerase (KAPA Biosystems Inc., Wilmington, MA), 2.5 mL KAPA Buffer A (containing 15 mM MgCl2), 0.5 mL of 10 mM dNTP mix, 0.3 mL DMSO (final concentration of 1.5%), 1.25 mL betaine (final concentration 0.25 mM), 1 mL of 0.2 mmol of each primer (10 mM each) and 2 mL DNA template. For the DNA extracted by homogenization and boiling, each 25 mL reaction contained 2.5 mL of 10 reaction buffer (containing 15 mM MgCl2), 0.5 mL of 25 mM MgCl2, 4 mL of 1.25 mM dNTPs, 10 pmol of each primer, 0.5 mL of 5U/mL HotStar Taq DNA polymerase (Qiagen Australia) and 5 mL DNA template. Cycling conditions were initial denaturation at 94 C for 5 min, followed by 40 cycles of denaturation at 94 C for 30 s, annealing at 52 C for 30 s and polymerization at 72 C for 2 min. A negative control (no DNA) and a positive control were included in each run. Positive controls were faecal DNAs from a dog with parvovirus enteritis infected with CPV-2b and/or from a cat with feline panleukopenia infected with FPV diagnosed using the methods above. Shelter 3 DNA was extracted from faeces by homogenization and boiling as described previously, (Clegg et al., 2012). The resulting supernatants were diluted in molecular-grade water (1:10, 1:20, and 1:50) and stored at 4 C for PCR testing within 7 days of extraction. To test for PCR inhibition, DNA extracts (1:10 and 1:20 dilutions) were spiked with 1 mL of feline genomic DNA (138 ng/mL) extracted from the lymph node of a healthy cat. A conventional PCR using primer set GAPDH-F and GAPDH-R to amplify the feline housekeeping GAPDH gene was performed (Table 1) (McLuckie et al., 2016). Each 50 mL reaction contained 5 mL of 10 X PCR buffer (containing 10 mMol dNTP and 75 mM MgCl2), 1 mL (10 pmol) of each primer, 0.2 mL (0.2 U) of Taq QIAGEN DNA Polymerase (QIAGEN Germantown, MD), 1 mL of feline genomic DNA template and 5 mL faecal DNA. Cycling conditions were initial denaturation at 94 C for 3 min, followed by 35 cycles of denaturation at 94 C for 45 s, annealing at 55 C for 30 s and polymerization at 72 C for 30 s, with final extension 72 C for 7 min. If PCR inhibition was detected additional dilutions of the sample were tested to identify the dilution at which PCR inhibition was abolished. The presence of parvovirus in each faecal sample was detected using a conventional PCR to amplify the VP2 gene using primer pair 555-F and 555-R (Decaro et al., 2008) (Table 1). PCR screening of all faecal samples was performed using both 1:10 and 1:20 dilutions, as well as 1:50 dilutions for samples in which PCR inhibition was detected. Each 50 mL reaction contained 10 mL PCR buffer (5 mM dNTPs, 15 mM MgCl2), 0.5 mL (0.5U) MyTaq Hot Start polymerase (Bioline, Meriden Life Science, Alexandria, Australia), 1 mL (10 pmol) forward and reverse primer, 7 mL DNA and 30.5 mL water. Cycling conditions were initial denaturation at 94 C for 1 min, followed by 35 cycles of denaturation at 94 C for 30 s, annealing at 55 C for 30 s and polymerization at 72 C for 30 s, with final extension 72 C for 1 min. PCR was performed using sample dilutions that had no evidence of PCR inhibition. A blank DNA control (PBS used during the extraction process) and a positive control were included in each run. The positive control was faecal DNA from a dog with parvovirus enteritis extracted and sequenced using the methods above, and identified as CPV-2a. The sensitivity of this conventional PCR was determined using DNA extracted from a feline faecal sample confirmed to contain FPV by sequencing. The FPV copy
Table 1 Primers used in this study for PCR amplification of feline DNA (GAPDH) and of canine and feline parvovirus DNA, targeting the VP2 region (VP2). Primer name
Target
Sequence
Size (nta )
Reference
JS1F JS2F GAPDH-F GAPDH-R 555-F 555-R 2655F 3511R 3381F 4116R
VP2
50 -AGCTACAGGATCTGGGAACG-30 50 -CCACCCACACCATAACAACA-30 50 -AAGGCTGAGAACGGGAAAC-30 50 -CATTTGATGTTGGCGGGATC-30 50 -CAGGAAGATATCCAGAAGGAA-30 50 -GGTGCTAGTTGATATGTAATAAACA-30 50 -CCAGATCATCCATCAACATCA-30 50 -TGAACATCATCTGGATCTGTACC-30 50 -CCATGGAAACCAACCATACC-30 50 -AGTTAATTCCTGTTTTACCTCCAA-30
1975
Meers et al. (2007)
80
McLuckie et al. (2016)
583
Decaro et al. (2008)
837
Decaro et al. (2008
717
Decaro et al., 2008
a
Nucleotide.
GAPDH VP2 VP2 VP2
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number per gram of faeces was determined for this sample by quantitative PCR (qPCR), using a previously validated multiplex assay (Meggiolaro et al., 2017), and was 3 108 FPV copies per gram of faeces. An end-point dilution of the FPV positive faecal DNA was performed using serial ten-fold dilutions (from 101 to 109) in the conventional VP2 PCR (primer pair 555F–555R). The second highest dilution at which a positive result was detected on gel electrophoresis was considered to be the cPCR sensitivity threshold. Sequence and phylogenetic analysis PCR products were detected by electrophoresis (1% agarose gel) and visualization under UV light. To differentiate CPV from FPV, the remainder of the VP2 gene was amplified in positive samples using primers 3381F–4116R, and 2679F–3511R (Decaro et al., 2008) (Table 1). Sanger sequencing was performed at a commercial laboratory (Macrogen Seoul, Korea). DNA sequence chromatographs were visually inspected and assembled in CLC Workbench. Viral subtype (FPV, CPV 300G (CPV-2a), CPV 426D (CPV-2b) or CPV 426E (CPV-2c)) was determined from key amino acid substitutions (Decaro and Buonavoglia, 2012). Unique strains identified in this study were deposited on GenBank (submission #2092491).
Results A total of 118 faecal samples were collected from shelters 1 and 2 in Brisbane. Four of these samples were collected from cages that held 2 or 3 litter-mates. Of 66 samples for which age was recorded, the median age of the cats was 12 months (range 4 weeks to 7 years), with 52% of cats being 12 months of age or younger. Of 87 cats for which sex was recorded, 42 were male and 45 were female. From shelter 3 in Sydney 100 faecal samples were obtained from 79 cats. The median age of the cats was 4 months (range 5 weeks to 5 years) and the majority of cats for which age was recorded (66/79, 84%), were 12 months of age or younger. Of 77 cats for which sex was recorded, there were 29 males and 48 females. Of 100 faecal DNA samples from Sydney in which PCR inhibition was tested, six had PCR inhibition at a 1:10 dilution, which was abolished at a 1:20 dilution in five samples and at a 1:50 dilution in one sample. Quantification of the number of viral copies present in the known FPV positive control sample, using the qPCR assay was determined to be 299, 290 copies/g of faeces. The end-point dilution study of the conventional FPV/CPV VP2 PCR assay revealed a 1 105 limit of detection, which was repeatable (Fig. 1). Thus, the limit of sensitivity of detection of this conventional PCR assay was 300 viral copies/gram (Fig. 1) of faeces or 15 viral copies/1 mL template. Carnivore protoparvovirus 1 DNA was detected in 4/218 (1.8%, 95% confidence interval 0.49–4.53%) samples from all three shelters by conventional PCR. The four positive samples were collected from four separate enclosures in shelter 3 on the third sampling occasion, after an outbreak of feline panleukopenia had
Fig. 1. A dilution assay was performed using a feline panleukopenia virus-positive faecal DNA sample containing 299, 290 viral copies/g faeces, as determined by a validated qPCR assay to calculate the sensitivity of the conventional PCR assay used to detect Carnivore protoparvovirus1 in shelter 3.
occurred within the previous two months at this shelter. Sequencing of the VP2-gene showed that VP2 key residues typical of FPV were present in all obtained sequences, so that the four parvovirus strains were identified as FPV. The FPV strains were identical to the outbreak strain detected in sick cats in a separate investigation (Barrs et al., 2017). In contrast, CPV shedding was not detected in any samples from the three shelters in this study. Discussion Cats are susceptible to subclinical or clinical infection by CPV variants including CPV 300G (CPV-2a), CPV 426D (CPV-2b) and CPV 426E (CPV-2c) (Nakamura et al., 2001; Gamoh et al., 2003a). This has been demonstrated in experimental infections, in which CPV-2 was unable to replicate in feline tissues but CPV 300G (CPV-2a) and CPV 426D (CPV-2b) underwent efficient replication (Truyen et al., 1994; Nakamura et al., 2001), as well as in naturally occurring infections in which CPV-2a, CPV-2b and CPV-2c but not CPV-2 isolates were detected in cats with clinical signs similar to those caused by FPV infection (Mochizuki et al., 1996; Truyen et al., 1996; Gamoh et al., 2003b; Decaro et al., 2010; Miranda et al., 2014). The absence of faecal CPV shedding in this cohort of healthy Australian shelter-housed cats contrasts the UK study where 26% of shelterhoused cats from one shelter, sampled at a single time point, shed CPV in their faeces (Clegg et al., 2012). In that study, longitudinal sampling was performed in a second shelter, demonstrating that 34 of 74 cats (46%) shed CPV in faeces on at least one occasion over an 8-week period, while neither 122 canine faecal samples or 160 environmental samples from that shelter were positive (Clegg et al., 2012). Our results align more closely with those of a US investigation, where quantitative PCR was used to detect faecal shedding of parvoviruses in healthy shelter cats (Leutenegger et al., 2015). In that study although 58% of cats were found to be shedding Carnivore protoparvovirus 1, only 21% of these (12% overall) had high parvoviral loads consistent with active infection and <1% of cats overall were shedding high viral loads of CPV. The reasons why active CPV infections are highly prevalent in cats entering shelters in the UK while none were detected in Australian shelter-housed cats are not immediately obvious. The difference in the observed prevalence of CPV shedding between Australia and UK cannot be accounted for by methodological differences since similar DNA extraction and PCR methods were used and we also tested a larger number of cats (Clegg et al., 2012). DNA extracted from faeces frequently contains PCR inhibitors, especially at low dilutions (Schrader et al., 2012), which can be a cause of false negative PCR results. PCR inhibitors in faeces include complex polysaccharides, bile salts, lipids, urea and haemoglobin (Schrader et al., 2012). Additional steps were taken in this study to exclude PCR inhibition as a cause of negative results including the use of a commercial DNA extraction method or an extraction control. It is possible that low-level faecal CPV shedding was not detected using the cPCR assays in this study, although cPCR was also used in the UK study (Clegg et al., 2012). To explore this possibility, we determined the limit of detection of one cPCR assay, which was 45 viral copies/3 mL template. In comparison, a qPCR designed to detect FPV and CPV was reported to have a 95% limit of detection of 20 copies/3 mL DNA template (Freisl et al., 2017). In another investigation using the same qPCR protocol, faecal samples from cats and dogs with suspected parvovirus infection were tested using IFA, conventional PCR, and qPCR. A marked difference in sensitivity between the three assays was detected, with 12%, 32% and 58% of samples testing positive for FPV or CPV by IFA, cPCR and qPCR, respectively (Streck et al., 2013). Quantitative results of CPV viral loads shed in faeces by healthy cats as determined by qPCR have not as yet been published. CPV loads
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shed in faeces in dogs after administration of a modified live CPV vaccine were 3.87 102–8.39 105 viral copies/gram of faeces, which were 4–5 fold lower than CPV loads shed in faeces after infection with field strains of CPV (Decaro et al., 2005; Freisl et al., 2017). If CPV faecal loads in cats are similar to dogs, then it is unlikely that cats undergoing active CPV infections were not detected in this study since the limit of detection was 3 102 viral copies/gram. However, to further investigate CPV shedding in asymptomatic cats, use of an optimized qPCR would be prudent. As expected, vaccine strains of FPV were not detected in the faeces of any cats from shelters 1 and 2, since the incubation period for FPV is usually at least two days (Csiza et al., 1971a,b). While the cats tested here were not shedding CPV, our results do not exclude that these cats might have been latently infected with CPV. CPV DNA has been detected by PCR from the bone marrow aspirates of healthy unowned Australian cats (Haynes and Holloway, 2012). Carnivore protoparvovirus 1 becomes latent in peripheral blood mononuclear cells (PBMC) after viral shedding ceases, leaving behind a convenient molecular “footprint” of a previous active infection (Miyazawa et al., 1999; Allison et al., 2013; Allison et al., 2014). There were several limitations to our study. Investigations of CPV-shedding in Brisbane (Shelters 1 and 2) and Sydney (Shelter 3) were initiated independently and without knowledge of each other, hence the methodological differences in primers used for Carnivore protoparvovirus 1 amplification, DNA extraction methods and how PCR inhibition was addressed. Clegg et al. (2012), used primer EF (2748–2765) and JS2R (Table 1) for VP2 amplification of parvoviruses from shelter cat faeces as well as slightly different PCR conditions to what were used in our studies (Clegg et al., 2011). It is possible that the sensitivity of the conventional PCR used by Clegg et al. (2011, 2012) was higher than the PCRs we used. Canine parvovirus is a common cause of enteritis in dogs worldwide. In dogs with acute, severe diarrhoea presenting to charity-sponsored primary care veterinary clinics in the UK, CPV was detected in 58% of 355 faecal samples (Godsall et al., 2010). In Australia it is estimated that 20,000 new cases of canine parvovirus enteritis are presented to veterinarians annually, using data collected in a national epidemiological survey (Kelman et al., 2017). Clusters of CPV-associated disease in Australia occur more commonly in areas with lower socioeconomic indices (Brady et al., 2012). While it is possible that we did not detect CPV shedding in shelter cats because they had not been exposed to CPV, Shelter 3 is located in a CPV “hot spot” suggesting that environmental exposure of free roaming cats presented to the shelter to CPV is not unlikely. Other potential contributors to the disparity between shedding of CPV by shelter cats in UK compared with Australia include differences in population density, territory size and climactic factors could lead to behavioural differences between cat populations, such as frequency of interaction with dogs and other cats resulting in different frequencies of exposure to CPV. Frequent cross-species transmission events between domestic and wildlife carnivores have been implicated in the evolution of CPVs in regions of the world with endemic carnivore species diversity (Allison et al., 2014). The role of wildlife carnivore contact in transmission of CPV to cats has not been investigated. Canine parvoviruses (CPVs) can infect multiple different families of carnivores in both Carnivora suborders, Feliformia and Caniformia (Allison et al., 2013). The diversity of carnivores in Australia and New Zealand is low, being limited to a few species in the Felidae, Canidae and Mustelidae families such as the domestic cat, dog and ferret. In Australia there is only one endemic terrestrial carnivore, the dingo, while in New Zealand there are none. The extent of carnivore species diversity in the UK is higher than Australia and New Zealand but lower than that in Europe and North America. Molecular surveillance of CPVs in Australia and New Zealand, has
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shown a ‘lag’ in CPV evolution, with CPV-2a being reported as the dominant subtype in Australia until 2011–2013, and the only subtype in NZ until at least 2009, likely reflecting both geographical isolation and the absence of endemic species which facilitate viral evolution (Ohneiser et al., 2015; Clark et al., 2018). Although we found no evidence of CPV shedding in shelter cats in Australia, cats can be infected by and shed CPV and can develop clinical disease indistinguishable from that caused by FPV. Considering that CPV-induced clinical disease has been reported in cats (Decaro et al., 2010; Miranda et al., 2014), stringent disinfection and biosecurity protocols that prevent both intra- and cross-species infectious diseases are essential for animal welfare in shelters. Conclusions Overall, the prevalence of faecal shedding of Carnivore protoparvovirus 1 in shelter-housed cats from three shelters in Australia was low. CPV shedding by shelter cats was not detected. Conflict of interest statement None of the authors of this paper has a financial or personal relationship with other people or organisations that could inappropriately influence or bias the content of the paper. Acknowledgements This study was supported by the June Rose Bullock bequest at the University of Sydney, the NSW Cat Protection Society, Boehringer Ingelheim, and MSD Animal Health. None of the organizations providing financial support were involved in the design or execution of the study. The authors thank staff at all of the shelters for their time and efforts in assisting with sample collection. References Allison, A.B., Kohler, D.J., Fox, K.A., Brown, J.D., Gerhold, R.W., Shearn-Bochsler, V.I., Dubovi, E.J., Parrish, C.R., Holmes, E.C., 2013. Frequent cross-species transmission of parvoviruses among diverse carnivore hosts. Journal of Virology 87, 2342–2347. Allison, A.B., Kohler, D.J., Ortega, A., Hoover, E.A., Grove, D.M., Holmes, E.C., Parrish, C.R., 2014. Host-specific parvovirus evolution in nature is recapitulated by in vitro adaptation to different carnivore species. PLoS Pathogens 10, e1004475. Barrs, V.R., Brailey, J., Allison, A.B., Kelman, M., Meers, J., Beatty, J.A., Holmes, E.C., 2017. Re-emergence of feline panleukopenia in Australia. 27th ECVIM-CA Congress, St Julian’s, Malta September 14–16, ISCAID_0_4. Battilani, M., Balboni, A., Ustulin, M., Giunti, M., Scagliarini, A., Prosperi, S., 2011. Genetic complexity and multiple infections with more Parvovirus species in naturally infected cats. Veterinary Research 42, 43. Battilani, M., Scagliarini, A., Ciulli, S., Morganti, L., Prosperi, S., 2006. High genetic diversity of the VP2 gene of a canine parvovirus strain detected in a domestic cat. Virology 352, 22–26. Brady, S., Norris, J.M., Kelman, M., Ward, M.P., 2012. Canine parvovirus in Australia: the role of socio-economic factors in disease clusters. Veterinary Journal 193, 522–528. Clark, N.J., Seddon, J.M., Kyaw-Tanner, M., Al-Alawneh, J., Harper, G., McDonagh, P., Meers, J., 2018. Emergence of canine parvovirus subtype 2b (CPV-2b) infections in Australian dogs. Infection, Genetics and Evolution: Journal of Molecular Epidemiology and Evolutionary Genetics in Infectious Diseases 58, 50–55. Clegg, S.R., Coyne, K.P., Dawson, S., Spibey, N., Gaskell, R.M., Radford, A.D., 2012. Canine parvovirus in asymptomatic feline carriers. Veterinary Microbiology 157, 78–85. Clegg, S.R., Coyne, K.P., Parker, J., Dawson, S., Godsall, S.A., Pinchbeck, G., Cripps, P.J., Gaskell, R.M., Radford, A.D., 2011. Molecular epidemiology and phylogeny reveal complex spatial dynamics in areas where canine parvovirus is endemic. Journal of Virology 85, 7892–7899. Csiza, C.K., De Lahunta, A., Scott, F.W., Gillespie, J.H., 1971a. Pathogenesis of feline panleukopenia virus in susceptible newborn kittens II. Pathology and immunofluorescence. Infection and Immunity 3, 838–846. Csiza, C.K., Scott, F.W., De Lahunta, A., Gillespie, J.H., 1971b. Pathogenesis of feline panleukopenia virus in susceptible newborn kittens I. Clinical signs hematology, serology, and virology. Infection and Immunity 3, 833–837.
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