Persistence and chronic urinary shedding of the aphthovirus equine rhinitis A virus

Comparative Immunology, Microbiology and Infectious Diseases 36 (2013) 95–103

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Comparative Immunology, Microbiology and Infectious Diseases journal homepage: www.elsevier.com/locate/cimid

Persistence and chronic urinary shedding of the aphthovirus equine rhinitis A virus Stacey E. Lynch a , James R. Gilkerson a , Sally J. Symes a , Jin-an Huang b , Carol A. Hartley a,∗ a b

Equine Infectious Diseases Laboratory, Department of Veterinary Science, The University of Melbourne, Australia Veterinary Medicine Research and Development, Pfizer Animal Health, Australia

a r t i c l e

i n f o

Article history: Received 2 August 2012 Received in revised form 10 October 2012 Accepted 12 October 2012 Keywords: Equine rhinitis A virus Aphthovirus Picornavirus Persistence

a b s t r a c t Equine rhinitis A virus (ERAV) is a member of the Aphthovirus genus, and has many physical and structural similarities to the prototype Aphthovirus foot-and-mouth disease virus (FMDV). The pathogenesis of FMDV has been extensively studied, however, the similarities in the pathogenesis of ERAV and FMDV disease has not been well documented. This study describes and compares the pathogenesis of ERAV both in the natural host and a small animal model alternative (CBA mice). Distinct parallels in the pathogenesis of the acute infection of these two viruses are described where infection in the upper respiratory tract precedes shedding of high levels of virus from the nasopharynx and a transient viraemic phase before dissemination to distal sites. The finding that ERAV is maintained at high levels in the urine of infected horses for at least 37 days post infection, however, is a feature unique to ERAV amongst all of the picornaviruses. © 2012 Elsevier Ltd. All rights reserved.

1. Introduction Equine rhinitis A virus (ERAV) is a respiratory pathogen of horses classified in the family Picornaviridae, genus Aphthovirus, alongside foot-and-mouth disease virus (FMDV) [1], a systemic pathogen of cloven hoofed animals. ERAV shares many pathogenic features with the prototype aphthovirus FMDV such as a respiratory infection, establishment of viraemia and persistent infection [1]. Furthermore, the key FMDV virulence factors, Lpro and 3Cpro share significant functional similarity to the homologue ERAV proteins [2]. Further investigation into the pathogenesis of ERAV may also highlight additional parallels between these two aphthoviruses, which has previously been described ERAV as a surrogate for FMDV, without the requirement for high-level biocontainment [3–5].

∗ Corresponding author at: Equine Infectious Diseases Laboratory, Department of Veterinary Science, The University of Melbourne, VIC 3010, Australia. Tel.: +61 3 8344 7375; fax: +61 3 8344 7374. E-mail address: [email protected] (C.A. Hartley). 0147-9571/$ – see front matter © 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.cimid.2012.10.003

Sporadic outbreaks of acute febrile respiratory disease following ERAV infection have been reported [6–10], although, given the high seroprevalence of ERAV within the population, sub-clinical infections of ERAV are likely to be common [8,11,12]. It is difficult, however, to establish an estimate of the prevalence of disease caused by ERAV, due to difficulties associated with isolation and propagation of this non-cytopathic virus [7]. Furthermore, recent ERAV challenge studies using highly cell culture adapted virus inoculums have resulted in sub-clinical infections [13], in contrast to the acute febrile respiratory disease observed in the early challenge studies [14]. There has been limited investigation of the host, viral and other factors that may explain the disparate outcomes of experimental infection. Early experimental challenge studies that documented viraemia and urinary shedding in guinea pigs and rabbits [15] suggested the progression of ERAV infection in a small animal model may be similar to that observed in the horse. This study describes an investigation into the pathogenesis of ERAV both in the natural host and a small animal model alternative (CBA mice). Using these two models the

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pathogenesis of ERAV is compared to that of the prototype member, FMDV. 2. Methods 2.1. Cells Viruses were propagated in primary equine foetal kidney (EFK) cells and Vero cells. Cell monolayers were grown in a base media of Dulbecco’s minimal essential medium (DMEM, Sigma) containing 5 mM sodium bicarbonate, 10 mM HEPES and 50 ␮g/ml ampicillin. Uninfected cells were cultured in growth media containing 4 (Vero) or 10% (v/v) (EFK) foetal bovine serum (FBS). Virus was propagated in maintenance media containing 0.5% (v/v) FBS. Virus titrations and neutralisation assays were performed in assay media supplemented with 2% (v/v) FBS, increased sodium bicarbonate (30 mM) and cultured in an atmosphere containing 5% CO2 . 2.2. Challenge viruses As there is no reproducible, clinical model for ERAV associated febrile respiratory disease, two different Australian ERAV isolates were examined as challenge viruses to investigate ERAV pathogenesis and clinical impact of infection. ERAV.393/76 [1,6], the laboratory prototype, and ERAV.967/90 [7,16], the only other Australian ERAV isolate. Both isolated in Australia from horses with acute febrile respiratory disease. The endpoint titre of ERAV inoculums was determined using Vero cells [6] and all contained between 8.9 and 9.6 log10 genome equivalents/ml as determined by RT-qPCR. The discordance between the infectivity titre and genome copies in the inoculums administered to horse 1 is likely to be due to growth of this virus in EFK cells and titration on Vero cells. Equine foetal kidney cells were not available at the time of the study to further propagate the virus. 2.3. Experimental infection of horses Two 8-month-old horses (Percheron Standardbred cross) were used for the experiment. Horses were seronegative to ERAV by virus neutralisation assay as well as western blotting analysis of serum on purified ERAV. Clarified supernatant from virus infected cell culture was administered via a catheter into the nasopharynx guided by an endoscope. Horses were infected with one of two non-plaque purified ERAV isolates. Horse 1 received 12 ml of ERAV.393/76 (passage 15 in primary equine foetal kidney cells (EFK)) at 3.8 log10 TCID50 /ml and Horse 2 received 25 ml of ERAV.967/90 (passage 8, 7 passages in EFK cells and 1 passage in Vero cells) at 8.0 log10 TCID50 /ml. Blood for serum, plasma and peripheral blood mononuclear cell isolation (PBMCs) was collected from the jugular vein of horses using a Vacutainer and an 18 gauge needle. Nasopharyngeal swabs were collected by inserting a gauze cloth swab held by a wire loop approximately 70 cm in length (protected by a long, soft rubber tube) into one of the horse’s nostrils. Oral swabs were collected by placing a gauze cloth on the inside cheek of the horse for

1 min to absorb saliva fluids. Swabs were placed into 3 ml of virus transport media (DMEM containing 3% (v/v) FBS and 100 ␮g/ml ampicillin), transported on ice and stored at −70 ◦ C prior to assay. Urine was collected following the intravenous administration of the diuretic Frusemide (Ilium). Urine was stored at 4 ◦ C for up to a week before testing, then at −70 ◦ C. Blood, nasopharyngeal and oral swabs were collected on alternate days for two weeks following infection, as well as on days 21 and 37 post infection (p.i.). Urine was collected on days 4, 15, 21 and 37 p.i. Horses were observed daily for clinical signs associated with acute febrile respiratory disease (i.e. rectal temperature, nasal discharge, coughing, lethargy). 2.4. Post-racing urine samples Two hundred and fifteen urine samples were obtained from Racing Analytical Services Ltd. (Flemington, Victoria) from samples collected from Thoroughbred horses for routine post-race testing. Urine samples, stored at 4 ◦ C for less than one month prior to testing, were filtered through 0.22 ␮M pore sized filters and stored at −70 ◦ C. Samples were either diluted in virus titrated media for virus isolation, or RNA was extracted using a Viral RNA extraction kit (QIAGEN), according to manufacturer’s instructions. 2.5. Experimental infection of CBA mice Thirty 6–8-week-old male CBA mice were obtained from the Walter and Eliza Hall Medical Research Institute. Animal husbandry was in accordance with the standard operating procedures of the facility and the trial was carried out with the approval of the appropriate animal ethics committee. This model was developed using the well-characterised laboratory prototype ERAV isolate, ERAV.393/76 (passage 22, 18 passages in EFK cells and 4 passages in Vero cells) at 8.0 log10 TCID50 /ml or 50 ␮L of uninfected Vero cell lysate. Mice (n = 30) were anaesthetised in an induction chamber containing 5% (v/v) isofluorane (Veterinary Companies of Australia Pty Ltd.) and intranasally inoculated with either 50 ␮L of clarified ERAV.393/76 (infected, n = 25) or 50 ␮l of clarified uninfected Vero cell lysate (uninfected, n = 5). At 1, 3, 5, 7 and 28 days after infection, five ERAV infected mice and one uninfected control mouse were euthanased by CO2 asphyxiation. Organs and urine were collected at necropsy. Urine, if present at the time of necropsy, was aspirated from the bladder using a 29½ gauge ultra fine needle (Becton Dickinson). Serum was processed from blood collected from the thoracic cavity upon dissection. Samples for total RNA isolation were placed in 1.5 mL RNAlater (Ambion). For immunohistochemistry, organs were placed into a CryoMOLD (Tissue-Tek) containing optimal cutting temperature gel (Tissue-Tek) and placed over a vessel containing liquid nitrogen until frozen. 2.6. Virological methods Virus was isolated from clinical samples after incubation with a semi confluent Vero cell monolayer at 37 ◦ C

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for 7 days, after which time any cytopathic effects (CPE) characteristic of ERAV were recorded. For post-racing urine samples, 200 ␮l of filtered urine (diluted 1 in 2 in maintenance media) was added to a semi-confluent Vero cell monolayer in a 25 cm2 flask containing 5 ml of maintenance media. For CBA mice urine and serum samples, 10–20 ␮l of clarified (1500 × g for 1 min) sample was added to one well of a 96-well tray containing a semi-confluent Vero cell monolayer and 200 ␮l of maintenance media. Immunoperoxidase staining for the detection of noncytopathic virus was performed as described [17] using rabbit ERAV.393/76 antiserum [13]. The virus neutralisation and virus titration assays were performed as previously described [6]. Nasopharyngeal swab samples collected from experimentally infected horses, where treated with 10% (v/v) chloroform, to remove potentially contaminating envelope viruses, prior to titration. 2.7. Immunohistochemistry for ERAV antigen Air dried 5 ␮m frozen tissue sections were fixed with acetone and treated with a peroxidase block reagent (Dako) and a serum free protein block solution (Dako) according to the manufacturer’s instructions. Viral proteins were detected by probing tissue sections with a rabbit ERAV.393/76 antiserum [13] diluted 1 in 1000 in PBS containing 1% FBS (v/v) for 45 min in a humidified container. Following incubation, sections were washed once for 10 min in PBS and probed with a swine anti-rabbit IgG HRP conjugate (Dako), diluted 1 in 100 with PBS containing 1% (v/v) FBS for 45 min. Unbound antibodies were removed with one wash in PBS for 10 min. Bound antibodies were detected by the addition of the chromogenic substrate, 3,3 -diaminobenzidine (DAB, Sigma) for 10 min. Sections were washed for 10 min with PBS, counter stained with Mayer’s haematoxylin for 40 s and washed in water for 10 min. Sections were dehydrated through a series of ethanol washes (70%, v/v ethanol 5 min; 100%, v/v ethanol 8 min) then incubated with 100% xylene for 16 min. Slides were mounted in p-xylene-bis-pyridinium bromide (DPX, Sigma). 2.8. RNA extraction Viral RNA was extracted from clinical samples in high-throughput system using a QIA-Xtractor (QIAGEN) according to the Vx Viral DNA/RNA extraction protocol (QIAGEN). For small-scale preparations of viral RNA, the QIAamp Viral RNA Mini Kit (QIAGEN) was used according to the manufacturer’s directions. Total RNA was extracted from CBA mouse tissue with the QIA-Xtractor (QIAGEN) using a modified Vx Viral DNA/RNA extraction protocol. Twenty milligrams of tissue was homogenised with a three-way tap (Discofix C, Braun) in 500 ␮l of tissue digest buffer (DXT, QIAGEN) containing 5% (v/v) lysis reagent (VXL, QIAGEN). The homogenised samples were heated at 55 ◦ C for 30 min and centrifuged (1500 × g for 1 min). RNA was extracted from 300 ␮l of supernatant according to manufacturer’s instructions.

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2.9. Reverse transcription quantitative PCR Primers and TaqMan probes were designed to the 5 untranslated region (UTR) of ERAV using Beacon Designer software, version 7.0. (Premier Biosoft International) and manufactured by GeneWorks and Biosearch Technologies, respectively. Complementary DNA was generated from 5 ␮l of RNA or 1 ␮g of total RNA using the reverse transcriptase SuperScript III (Invitrogen), the gene specific primer ERAV R427 (5 -CTTGGTGACACCATCCAGAGC-3 ), which anneals to position 406–427 of ERAV.393/76 (GeneBank Accession no. L43052) and 4 mM Oligo (dT)15 sequence (GeneWorks) (total RNA only), according to the manufacturer’s instructions. Quantitative PCR was performed using primers ERAV F303 (5 -TCGTCACTTGGCTGTTCTATCG-3 ), which anneals to position 303–324 of ERAV.393/76 and ERAV R427. The ERAV R427 and ERAV F303 primer pair generates a 124 base pair product. For a SYBR Green I based assay the Quantitect SYBR qPCR master mix (QIAGEN), supplemented with additional MgCl2 (to a final concentration of 4.5 mM), was used according to the manufacturer’s instructions. For the probe-based assay, the Brilliant qPCR Multiplex master mix (Stratagene) and probe ERAV P401 (FAM-5 -TGGCACCGGGAAAATCCAGCACGC-3 BHQ1), which anneals to position 401–424 of ERAV.393/76, were used according to the manufacturer’s instructions. For both assays, an annealing temperature of 60 ◦ C was used. Quantitative PCR assays were performed using an MxPro Mx3000P Real Time PCR machine (Stratagene). The Mx3000P software constructed a standard curve from samples of known copy number and assigned copy numbers to unknown samples. Feline calicivirus was included as an exogenous internal positive control (IPC) to detect inhibitors in the extraction and quantitation of ERAV in clinical samples from experimentally infected horses. There was no significant difference in the mean Ct values for ERAV tested in the monoplex or multiplex RT-qPCR assay (data not shown), indicating that the sensitivity of the ERAV assay is retained when processed in a multiplex assay with the IPC. Furthermore, in these experiments the IPC was consistently detected at a Ct value of 24.4 (standard deviation 0.1, Cv% 0.41, n = 54) over a wide range of ERAV input starting concentrations (0.01–1584 TCID50 per reaction). To detect tissue specific inhibitors in the assay to quantify ERAV genome copies in tissue samples from experimentally infected mice, an endogenous internal control (cyclophilin A), as described in was included [18]. Cyclophilin A was consistently detected by RT-qPCR at Ct values ranging between 20 and 24 in total RNA extracted mice tissue, demonstrating consistent amplification of RNA targets from diverse clinical samples. 2.10. Preparation of synthetic RNA standard The ERAV qPCR amplicon (spanning a 124 base pair region of the 5 UTR) was cloned into pGEM-T downstream of a T7 promoter and a synthetic RNA transcript was generated using MAXIscript (Ambion), according to the manufacturer’s instructions. The original DNA template

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was digested twice with DNase I (Invitrogen) according to the manufacturer’s instructions. The mean concentration of synthetic transcripts was determined spectrophotometrically from triplicate readings. Standards were generated from 10-fold serial dilutions (8–2 log10 RNA copies/RTqPCR) in pyrogen free water containing 0.122 ␮g/␮l carrier RNA. Standards were reverse transcribed and amplified in parallel with clinical samples. Controls included RT negative control, where a parallel RT reaction was performed, without the addition of the RT enzyme, to confirm the complete digestion of DNA template.

2.11. Sequencing and phylogenetic analysis New ERAV isolates were further characterised based on capsid viral protein 1 (VP1) nucleotide sequence [16]. Viral protein 1 was amplified from viral cDNA using primers R2A and VP3/1 F as previously described [16] and sequenced using the BigDye Terminator v3.1 reagent according to the manufacturer’s instructions. Precipitated sequencing products were analysed on an ABI313x1 capillary genetic analyser (The University of Melbourne, Department of Pathology). Sequencing results were analysed using VectorNTI (Invitrogen).

3. Results 3.1. RT-qPCR sensitivity and specificity A RT-qPCR assay was designed to target the nonstructural internal ribosomal entry site (IRES) of ERAV. The assay was linear over 7 orders of magnitude extending 108 –102 copies of in vitro transcribed RNA/RT-qPCR reaction and could detect 10–0.8 TCID50 , of virus per reaction, which is 1000 times more virus than previously published assays [19]. The ERAV RT-qPCR assay was highly reproducible with a low inter-assay variation (0.40–2.5 cv% median 0.52) and low intra-assay variation (0.5–1.8 cv% median 0.77). DNA sequencing was use to confirm the specificity of the ERAV amplicon, which was 100% identical to the expected amplicon sequence. There was no amplification of non-specific products from nucleic acid prepared from uninfected clinical samples or equine rhinitis B viral RNA (ERBV.1436/71). Amplification was detected with comparable Ct values and inter- and intra-assay variation when using either TaqMan or SYBR Green I based chemistries in the assay (data not shown). No significant differences in Ct values and amplification efficiencies where found when ERAV controls were extracted and amplified from culture media, equine nasopharyngeal and oral swabs, plasma or urine (data not shown).

3.2. Experimental infection of horses with ERAV Nasopharyngeal and oral swabs, plasma and urine collected from the two experimentally infected horses up to 37 days p.i. were tested for the presence of viral RNA using an ERAV specific RT-qPCR assay (Fig. 1). Despite the detection of ERAV from respiratory secretions and in plasma, clinical

signs of acute febrile respiratory disease were not observed in the 14 days following infection. 3.2.1. Chronic urinary shedding of ERAV ERAV was excreted at high levels in the urine of both horses. ERAV RNA was first detected in urine samples collect at day 14 p.i. (horse 1) and on day 4 p.i. (horse 2) (Fig. 1 D and H). Viral RNA was consistently detected in all subsequent urine samples collected until day 37 p.i. The re-isolation of ERAV from the urine correlated well with detection of ERAV RNA by RT-qPCR. Four log10 TCID50/ ml was re-isolated from the horses at first detection and 2 log10 TCID50 /ml at the completion of the trial on day 37 p.i. These results show the highest level (3.97/4.63 log10 TCID50 /ml and 7.9/7.28 log10 RNA copies/ml, for horse 1 and 2, respectively) of virus detected was in the urine, compared with other anatomical sites, of experimentally infected horses. This implies the bulk of ERAV is shed through this route; considering a normal daily excretion of urine is 15–30 ml per kg bodyweight per day [20], and represents a significant source of excreted virus (Fig. 1). 3.2.2. Neutralising antibody response A rapid virus neutralising antibody response was detected at day 6 (horse 2) and day 8 (horse 1) after ERAV infection (Fig. 2A). The appearance of virus neutralising antibodies corresponded to the cessation of viraemia as determined either by RT-qPCR and/or virus isolation (Fig. 1) and infectious ERAV was not re-isolated from the nasopharynx of infected horses. 3.3. Detection and analysis of ERAV in post-race horse urine The high level and long duration of ERAV shedding in urine when compared with samples from other anatomical sites (Fig. 1) suggests that urine would be a more sensitive sample than nasal swabs for ERAV detection. Therefore urine samples collected from Thoroughbred racehorses post-racing were tested for ERAV to investigate the chronic urinary shedding of ERAV and assess ERAV shedding in clinically normal horses. ERAV was detected by RT-qPCR in 50 (23.3%) of the 215 urine samples tested. ERAV was most prevalent in urine samples collected from horses aged between 2 and 4 years (29.4%, 42 of 143 urine samples collected in this age group), and was detected in samples from horses of both sexes. The cycle threshold value ranged between 16 and 36 for the ERAV positive samples, with 80% (n = 40) of positive samples detected at Ct values less than 30 (data not shown). Sixteen RT-qPCR positive urine samples were inoculated onto 3 different cell lines; EFK, Vero and RK13 cells. ERAV was isolated from 5 of these urine samples following the observation of CPE characteristic of ERAV infection and an increase in viral load over time by RTqPCR (data not shown). Rabbit kidney (RK-13) cells and Vero cells were most sensitive to infection, supporting the growth of 4 out of the 5 isolates. The fifth isolate was isolated only on EFK cells. As non-cytopathic ERAVs have been cultured from clinical samples [7], all cultures inoculated with RT-qPCR positive urine samples were stained

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8

Nasopharyngeal

7 6 5

E) log10 RNA copies/ml

log10 RNA copies/ml

A)

4 0

10

20

30

8

Nasopharyngeal

7 6 5 4

40

0

day post infection

8

Oral

7 6 5

F)

log10 RNA copies/ml

log10 RNA copies/ml

B)

4 0

10

20

30

Plasma

7 6 5

Oral

7 6 5

0

G)

4 0

10

20

30

8

Urine

7 6 5 4 0

10

20

30

day post infection

40

40

7 6 5 4

40

0

H)

log10 RNA copies/ml

log10 RNA copies/ml

8

10 20 30 day post infection

Plasma

day post infection

D)

40

4

40

log10 RNA copies/ml

log10 RNA copies/ml

8

10 20 30 day post infection

8

day post infection

C)

99

10 20 30 day post infection

8

40

Urine

7 6 5 4 0

10 20 30 day post infection

40

Fig. 1. ERAV shedding in horse 1 (A–D) infected with ERAV.393/76, or horse 2 (E–H) infected with ERAV.967/90. Viral load is represented as RNA copies/ml by RT-qPCR, and is measured in samples from nasopharyngeal swabs (A and E); oral swabs (B and F); plasma (C and G) and urine (D and H). The limit of detection for each assay is displayed with a line.

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(A) 10

6

10 5

VN titre

10 4

10 3

10 2

10 1

10 0 0

5

10

15

20

25

30

35

Day post infection

(B) 10

4

Fig. 3. Unrooted phylogenetic tree of VP1 nucleotide sequences of ERAV isolates (nucleotide position 1650–2321) inferred using the maximum likelihood method. The reproducibility of the analysis was assessed by 100 bootstrapping replicates. Bootstrapping values are indicated on branches. ERAV 3117/09 sequences are the post race horse urine isolates. Comparison ERAV sequences were from Varrasso et al. [16].

VN titre

10 3

10 2

3.4. Quantification of ERAV in experimentally infected CBA mice 10 1

0

5

10

15

20

25

Day post infection Fig. 2. Virus neutralising (VN) serum antibody response to ERAV infection. VN titres in serum samples collected from (A) horse 1, infected with ERAV.393/76 (closed circle) and horse 2, infected with ERAV.967/90 (open circle) for 37 days following infection. (B) VN titres of serum from CBA mice infected with ERAV.393/76 and euthanased up to day 28 p.i. Antibodies titres from individual mice (up to five per time point) are indicated. The limit of detection for the assay is shown with a dashed line. Virus neutralising antibody titres were determined as the reciprocal of the lowest dilution to neutralise 100 TCID50 of the homologous ERAV challenge isolate.

using ERAV.393/76 polyclonal rabbit antiserum and bound antibodies detected using a horseradish peroxidase conjugated secondary antibody. The cytopathic effect correlated well with immunoperoxidase staining and there was no immunoperoxidase staining cells in cultures without CPE (data not shown). The phylogenetic relationship between the VP1 nucleotide sequence of the 5 ERAV racehorse urine isolates (ERAV.3117/09) and 9 genetically distinct archival ERAV isolates [16] were examined (Fig. 3). The phylogenetic relationship inferred using the maximum likelihood method divides the VP1 sequences into three clusters with ERAV.3117/09 sequences and ERAV.967/90 forming a discrete cluster. There was considerable nucleotide identity (94–98%) between the 5 VP1 nucleotide sequences obtained from post-race horse urine samples and an 89–90% identity with ERAV.967/90. There was a lower percentage nucleotide identity (79%) between isolates from the horse urine (ERAV.3117/09) and the laboratory prototype ERAV.393/76.

Although ERAV is reported to have a broad host range by experimental infection [15], the replication, dissemination and persistence of ERAV infection in species other than the natural host has not been investigated. The dynamics of ERAV replication in CBA mice was further investigated. There was no evidence of weight loss or other clinical signs of disease in infected mice when compared to the uninfected controls during this experiment. The presence of infectious ERAV in urine and serum samples was determined by virus isolation and the viral load in organ samples collected on days 1, 3, 5, 7 and 28 was determined by RTqPCR (Table 1). 3.4.1. Viraemia and the neutralising antibody response A transient viraemia was detected in intranasally inoculated mice following infection. Infectious ERAV was detected in all serum samples collected from infected mice on day 1 p.i. On day 3 p.i., infectious virus was detected in the serum sample from one individual mouse, however, not detected in any mouse serum on subsequent days (Table 1). A rapid virus neutralising antibody response was detected at day 3 after ERAV infection in experimentally infected mice (Fig. 2B). The appearance of virus neutralising antibodies corresponded to the cessation of viraemia as determined either by RT-qPCR and/or virus isolation and the level of ERAV RNA detected by RT-qPCR decreased in the lungs of infected mice (Table 1). 3.4.2. Replication sites and viral load in other tissues Quantifying ERAV load in the organs of infected mice showed the highest levels and most consistent detection in the lungs of the infected mice. The viral load peaked on days 1 and 3 p.i. (Table 1) and reduced approximately

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Table 1 Detection of ERAV by virus isolation and RT-qPCR in organs and samples collected from CBA mice for 28 days post intranasal infection. Daya

IDb

Virus isolation Urine

RT-qPCR (log10 viral RNA copies/100 ng total RNA) Urine f

d

Lung

Heart

Kidney

Bladder

Liver

Spleen

1

1 2 3 4 5

− − NS − −

+ NSe + + NS

− 4.09 − NS −

5.91 5.97 4.66 5.37 6.01

2.36 2.68 2.10 2.55 2.02

− − − − −

− − − NS NS

− − − − −

− − − − −

3

6 7 8 9 10

− + − − NS

− + − − NS

− 4.57 5.62 − NS

5.68 5.64 5.47 5.84 6.17

2.02 2.30 2.71 2.16 −

2.45 2.11 − − −

− − − NS NS

2.26 2.08 − − 2.02

− 2.54 − − −

5

11 12 13 14 15

− − + − −

NS − − NS −

4.62 − NS 4.92 NS

5.42 5.37 4.76 5.30 4.81

− − − − −

2.67 − − −

2.69 − 3.29 NS NS

− 2.67 − − −

− − − − −

7

16 17 18 19 20

− − NS − NS

− NS NS − NS

NS − NS NS NS

3.47 4.10 4.00 4.37 4.22

− − − − −

− − − − −

2.73 − 2.91 NS NS

− − − − −

− − − − 2.06

28

21 22 23 24 25

− − NS − −

NS − − − NS

NS − − − −

− − 2.2 2.3 −

− − − − −

− − − − −

− − − − −

− − − − −

− − − − −

a b c d e f

c

Serum

Days post infection. Animal identification number. ERAV not detected. Virus isolated from 10 ␮l of plasma or 10–20 ␮l of urine. Sample not available for analysis. Viral load in 100 ␮l of urine.

10-fold at every 48 h time point until day 7 p.i., with ERAV RNA detected in only two of the five mice on 28 p.i. Detection of virus in all other organs was less consistent, and at levels 100–1000 fold lower than in the lungs (Table 1). Detection of immunohistochemistry ERAV antigen positive cells was consistent with this finding, since positive cells indicative of actively replicating virus, were found only in the lungs and in no other tissue (Fig. 4). Infectious virus or ERAV RNA was detected in a proportion of the urine samples collected from infected mice until day 5 p.i. (Table 1). Therefore despite the absence of immunohistochemistry ERAV antigen positive cells in the bladder and kidney, the urine contained viral loads comparable to the lungs (Table 1). It is not known whether virus is concentrated to these high levels in the urine from viraemia, or whether a specific site of ERAV replication exists in the genitourinary tract that was not captured in our sections for immunohistochemistry. 4. Discussion ERAV has been classified as an aphthovirus alongside FMDV since 1996, and although several physical and structural features are conserved among these viruses [1–5], it is largely unknown whether these also translate into consistencies in pathogenesis. Furthermore the significance of ERAV as a pathogen and a cause of poor

performance in horses may be underestimated in part, due to a lack of understanding of the dynamics of viral shedding following infection and sensitive techniques to detect virus. This study describes the development of two ERAV infection models; the first involving the natural host, while the second, an alternate small animal model to study ERAV pathogenesis. This study demonstrated the experimental infection of horses with two different Australian ERAV isolates, and confirms and extends previous studies examining the pathogenesis of this picornavirus. Like FMDV, local replication of ERAV in the respiratory tract preceded the dissemination of the virus via a transient viraemia that ceased with the induction of virus neutralising antibodies. Despite the detection of virus in the plasma and shedding in the nasopharynx, clinical signs of acute febrile respiratory disease were not observed following experimental ERAV infection, suggesting additional factors may contribute to the clinical disease reported to be associated with ERAV. Consistent with the infection of ERAV in horses, primary replication and viraemia progressed sub clinically in ERAV infected CBA mice. The establishment of a persistent infection by FMDV and ERAV is a noted feature of these aphthoviruses, although the type of persistent infections established are distinct. In contrast to the high levels of FMDV shed during acute infection (as reviewed by [21]), the persistent infection

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Fig. 4. Immunohistochemistry staining of ERAV antigen in frozen lung sections prepared from (A) uninfected and (B and C) ERAV.393/76 infected CBA mice. Sections were probed with rabbit ERAV.393/76 antiserum, with bound antibodies detected by HRP conjugated swine anti-rabbit IgG and the chromogenic substrate DAB. Sections were counter stained with haematoxylin. Viral antigen positive cells are indicated with brown precipitate in C and D (black arrows). Magnification 200×.

established by FMDV is characterised by the isolation of low levels of infectious virus (10–100 TCID50/ ml) from the oesophageal–pharyngeal fluid 28 days p.i. By contrast in this study, persistent infection established by ERAV resulted in highest levels of infectious virus and viral RNA in the urine of experimentally infected horses, and this was

consistent with the very high levels of ERAV detected in naturally infected horses. Furthermore, the establishment of a persistent infection appears common, since high loads of ERAV (Ct = 16–38) were detected in 23% of the 215 urine samples collected from race horses and this finding is consistent with other studies [22]. High levels of viral RNA are also generated during the persistent infection established by certain strains of Theiler’s murine encephalomyelitis virus (TMEV) of genus Enterovirus, family Picornaviridae [23], however in contrast to ERAV, comparable levels of infectious TMEV are not recovered. Clearly the mechanism of ERAV persistence is unique among picornaviruses since virus levels in urine are comparable during the acute and convalescent phases. Given the high levels of virus detected in the urine of infected horses and mice, it seems most likely that a secondary replication site is established in the genitourinary system, although this site was not detected in this study. The alternate explanation that virus is concentrated in the urine from filtration of virus in plasma seems less likely, given virus particles are much larger than the macromolecules filtered by the kidney. Although ERAV was detected by RT-qPCR in both the mouse bladder and kidneys, active replication was not confirmed at either site by IHC. Replication within an immunologically privileged niche, such as the intact microvascular endothelium and continuous basement membrane of the kidney, would explain the chronic urinary shedding of ERAV in the presence of a robust neutralising antibody response, however, the inability to culture ERAV from the kidneys of seropositive horses shedding virus in the urine may reflect secondary replication in the bladder, rather than the kidneys [12]. ERAV was not detected by RT-qPCR or virus isolation in the urine of the experimentally infected horses on days 145 and 179 p.i. in this study (data not shown). Furthermore, ERAV was detected less frequently in the urine of older animals, while epidemiological studies indicate ERAV infects younger horses [24]. Taken together, this suggests the persistent infection established by ERAV is not life long. Viral specific cytopathology within an immunoprivileged site, such as the kidney, may eventually disrupted the anatomic barrier preventing efficient immune surveillance such that the infection may ultimately be cleared [25]. The clinical implications of the persistent urinary shedding, may be difficult to detect, as overt renal dysfunction is not apparent until the function of 75% of the nephrons are affected [20] and further studies are required to understand the clinical impact of ERAV persistence with high urinary shedding in details. This study expands and consolidates previous studies examining the pathogenesis of ERAV in horses, and establishes the use of mice as a small animal model alternative. The dynamics of viral shedding in ERAV infected horses has shown a transient viraemia preceding primary replication in the nasopharnyx and the induction of an early robust neutralising antibody response. In contrast to FMDV, ERAV remains antigenically stable [16] and maintained consistent levels of replication from the acute phase to persistence. The mechanisms surrounding chronic urinary shedding and viral

S.E. Lynch et al. / Comparative Immunology, Microbiology and Infectious Diseases 36 (2013) 95–103

persistence in immunoprivileged sites requires further investigation.

[11]

Acknowledgements [12]

Project funding was from the Australian Research Council and Pfizer Animal Health. S.E.L was a recipient of an Australian Postgraduate Award (Industry) scholarship with Pfizer Animal Health as the industry partner. We thank Drs Gary Muscatello (The University of Sydney), Laura Fenell (The University of Melbourne), John Moody (CSL Ltd.) and Bob Geyer (The University of Melbourne) for assistance with animal experiments, and Nino Ficorilli and Cynthia Brown, Department of Veterinary Science, The University of Melbourne, for excellent technical assistance.

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[16]

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