Clinical review of two fatal equine cases of infection with theinsectivorous bat strain of Australian bat lyssavirus
EJ Annanda* and PA Reidb
Case series The first two confirmed cases of Australian batlyssavirus (ABLV) infection in horses are presented. Both casesoccurred in the same week in May 2013 in paddock mates in south-east Queensland. Australia has been one of only a few countriesconsidered free from rabies-like viruses in domestic animal species.ABLV infection had previously only been confirmed in bats andhumans. All three confirmed human cases were fatal, the latest inFebruary 2013. An additional human case of possible abortiveinfection in 1996 has also been reported. Both equine casesreported here resulted in euthanasia. The risks of infection acrossother mammalian species are still to be determined.
Conclusions These two equine cases highlight that ABLV shouldbe considered as a differential diagnosis in animals with similarclinical presentations in Australia. There is a need for greater aware-ness regarding the zoonotic risk, use of personal protective equip-ment, pre- and post-exposure prophylactic measures andlaboratory diagnostic options. The authors recommend ABLVtesting for all Australian cases of progressive equine neurologicaldisease.
Keywords Australian bat lyssavirus; equine encephalitis; equinerabies; zoonoses
Abbreviations ABLV, Australian bat lyssavirus; ABLVp, Australianbat lyssavirus pteropus strain; ABLVs, Australian bat lyssavirusyellow-bellied sheathtail bat strain; CNS, central nervous system;EHV-1,4, equine herpesvirus strain 1, strain 4; FAT, fluorescent anti-body test; HeV, Hendra virus; PEP, post-exposure prophylaxis; PPE,personal protective equipment; RABV, rabies virus; RT-PCR, reversetranscriptase PCR.Aust Vet J 2014;92:324–332 doi: 10.1111/avj.12227
Australian bat lyssavirus (ABLV) is a recently emergedmember of the family Rhabdoviridae, genus Lyssavirus.ABLV causes a fatal neurological disease that is indistin-
guishable from clinical rabies. It was first identified in 1996, as a resultof bat research related to Hendra virus (HeV),1 and is among anumber of recently emerged pathogens in Australia which have pre-sented with an encephalitis syndrome.2 Outside Australia, rabies virus(RABV) and other bat lyssaviruses have caused infections in numer-ous mammalian species.3
Lyssaviruses are enveloped, bullet-shaped viruses with a single-stranded, negative sense RNA genome. There are 12 species (geno-
types) of lyssavirus; three phylotypes are currently classified andanother three bat lyssaviruses await official classification.4 Allmembers of the genus are capable of causing fatal acute encephalitisindistinguishable from clinical rabies in humans and other mammals.4
Sequence comparisons of the ABLV N protein and other lyssavirus Nproteins have shown that ABLV is most closely related to European batlyssavirus-1 (EBLV-1) and RABV.1,5 RABV bat-variant genotype 1 andEBLV-1 have been reported to cause spillover infections in terrestrialspecies.6–9
Globally, bats are the primary reservoir of lyssaviruses10 and all Aus-tralian bat species are considered susceptible.11 Bats (orderChiroptera) occur in two suborders: the Megachiroptera (flying foxes),of which Australia has 1 family that includes 5 genera and 13 species,and the Microchiroptera (small, mostly insectivorous bats), of whichAustralia has 6 families containing 20 genera and 65 species.12
ABLV has been isolated from all species of Australian flying fox and inthe yellow-bellied sheathtail bat (Saccolaimus flaviventris). In addition,serological evidence of exposure to the virus has been reported in 7genera, representing 5 of the 6 families of AustralianMicrochiroptera.13 The distribution of species known to have beeninfected with ABLV includes all of Australia except southern WesternAustralia and parts of South Australia. (Figure 1)4,11,12,14
Two genetically stable variants of ABLV are recognised in Australia:one that circulates in four species of flying foxes of the genus Pteropus(ABLVp) and the other identified in the yellow-bellied sheathtail bat,(ABLVs).15,16 Recently, it has been shown that numerous cell linesderived from species including rodents, rabbits, humans, monkeys andhorses were permissive to viral entry mediated by the G glycoproteinsof both ABLVp and ABLVs17 and that the unknown receptor is broadlyconserved among mammalian species.4
In Australia, ABLV is considered endemic in flying fox and insectivo-rous bat populations, and is a notifiable disease.
We present the first two confirmed cases of ABLV infection in aspecies other than bats and humans, including the clinical and labo-ratory findings. Both cases occurred in horses in May 2013 on aproperty in south-east Queensland and both were confirmed to beABLVs infections. These cases raise important concerns for Australianveterinary practitioners.
Case series
Case 1An 18-month-old sport horse filly presented with subtle signs ofhindlimb ataxia. Over the preceding 3 weeks, she had been observed to
*Corresponding author.aRandwick Equine Centre, Sydney, New South Wales, Australia; [email protected], Queensland, Australia
have intermittent periods of reduced interaction with paddock matesand other subtle behavioural changes. The filly had been kept in apaddock with two other horses of the same age and breed, and all hadbeen bred and continuously reared on the property.
On examination, the ataxia worsened when the filly was blindfoldedand there was instability when backing up, but no significantproprioceptive deficit when performing tight circles. She also showedmild hyperaesthesia, mild pyrexia (38.7°C) and moderate protrusionof the nictitating membrane. All other clinical parameters werenormal, and cranial nerve and ophthalmic examinations revealed noabnormalities.
Haematology and biochemistry parameters were within normallimits. Treatments with an oral paste containing balanced electrolytesand vitamins (B1, B2, B6, B12 and E), as well as dexamethasone(0.07 mg/kg IV) and flunixin meglumine (1.1 mg/kg IV), were admin-istered. The filly was isolated in a stable for care and observation andpersonal protective equipment (PPE) protocols were started. The twoasymptomatic paddock mates were also isolated into individual smallpaddocks.
The initial differential diagnoses included cervical vertebral stenoticmyelopathy, equine herpesvirus-1 myeloencephalopathy, Ixodesholocyclus paralysis, ergot alkaloid toxicity, lead toxicity, brown snakeenvenomation, botulism, Murray Valley encephalitis, West Nile virusKunjin strain infection, HeV infection, hepatotoxic encephalopathy,tetanus or traumatic neuropathy. The filly and both paddock mates
had current tetanus vaccination, but had not received vaccinationagainst equine herpesvirus strain 1 (EHV-1) or HeV.
At 24 h after initial presentation, the filly was obtunded and pro-foundly ataxic, with a wide stance of all four limbs and low headcarriage. Repeat clinical assessment revealed hyperaesthesia, abnormalprotrusion of nictitating membranes, mild dehydration, but normalrectal temperature and heart rate. Nasal, oral and rectal swabs andblood samples were taken for HeV PCR exclusion testing.
Flunixin meglumine (1.1 mg/kg IV) and Hartmann’s solution (5 L IV)were administered, as well as penicillin (25,000 IU/kg IM) and tetanusantitoxin (20,000 IU SC), as treatment for possible tetanus.
That evening the filly was recumbent and making intermittent pan-icked attempts to stand. Oral bloody mucoid discharge was evident,possibly from increased salivation and decreased swallowing, com-bined with mild buccal mucosal bleeding from superficial trauma.Rectal temperature remained normal. The filly was sedated withdetomidine (0.01 mg/kg IV) and butorphanol (0.015 mg/kg IV).
At 54 h after initial presentation, the filly was in lateral recumbencydemonstrating uncontrolled paddling of all four limbs, dorsoventralmovement of the head and neck and rapid eye movement (all typicalof a generalised seizure). Diazepam (0.2 mg/kg IV) was ineffective incontrolling the seizures. The filly was euthanased.
Case 2On the morning following the initial presentation of horse 1, whilestill in isolation, one of the filly’s paddock mates, an 18-month-old
Figure 1. Distribution of bat species known tobe infected with Australian bat lyssavirus(ABLV). Map adapted from Van Dyke et al,14
AUSVETPLAN,11 Richards and Hall.12 Pteropidbats are highly mobile and there are occa-sional incursions into new areas (e.g. therehave been occasional sightings of Pteropusalecto as far south as Melbourne, Victoria andof P. poliocephalus as far south as Tasmania).From a public health perspective, all Austral-ian bats are considered capable of beinginfected with ABLV.
sport horse gelding, was reported as briefly displaying subtle hindlimbataxia and altered demeanour. He had no prior history of abnormalbehaviour. A clinical examination at the walk, trot and canter did notreveal any abnormalities. Haematology and biochemistry parameterswere within normal limits.
The gelding was observed closely for the next 2 days and showed nofurther clinical, behavioural or gait abnormalities. On the evening ofthe third day, the gelding demonstrated mild hindlimb ataxia, similarto horse 1, and was stabled in isolation. The PPE protocol wasreinstated.
At 12 h after clinical onset, the gelding was severely obtunded anddemonstrated photophobia, head pressing, cervical ventroflexion,severe ataxia and a wide-based stance. Clinical examination revealedmydriasis, pyrexia (39.0°C), mild tachycardia (45 beats/min), reducedabdominal borborygmi, normal hydration and respiration. He wasadministered flunixin meglumine (1.1 mg/kg IV) as well as vitamin C(30 mg/kg), vitamin E (13 IU/kg) and dimethyl sulfoxide (0.7 g/kg)diluted in 1 L of Hartmann’s solution administered by IV infusion.Samples were taken for HeV exclusion.
At 36 h after clinical onset, the gelding was in sternal recumbency andunable to stand, but otherwise had normal clinical parameters, dem-onstrated an improved demeanour and was drinking but anorexic.Another series of HeV exclusion samples was collected and sent withthe first series for same-day testing. Dexamethasone (0.07 mg/kg IV)as well as further doses of vitamin C (30 mg/kg IM) and flunixinmeglumine (1.1 mg/kg IV) were administered.
That afternoon the gelding was standing, having stood earlier in theday without assistance. Profound hypermetria of all four limbs wasevident, as well as head pressing, tachycardia (60 beats/min), absentborborygmi and reduced faecal output. The gelding was drinking butdysphagic, with a normal rectal temperature, and a wide-based stance(Figure 2).
At 54 h after clinical onset,5.5 days after the transient episode of altereddemeanour, clinical signs were observed that were consistent with aseizure, similar to those seen in horse 1. These included lateral recum-bency with paddling movements,head nodding and nystagmus,as wellas increased sweating and oral discharge. The gelding was euthanased.
Video footage of both these cases has been published elsewhere4 and isfreely available for download from the publisher’s website.
Postmortem examinationNecropsy of the first horse was not performed. In horse 2, the caudalcranium was removed and the meninges, brain, brain stem and cervi-cal spinal cord were examined. Marked injection of the meningealvasculature was evident.
The cerebrospinal fluid (CSF) had normal gross appearance and swabswere taken for bacterial and viral testing. Samples of brain, brain stemand cervical spinal cord tissue were fixed in formalin and sent forlaboratory investigation, together with the remaining brain, brainstem and cervical spinal cord tissue, which were transported fresh andchilled.
Laboratory findingsHistopathology, microbiology, PCR and fluorescent antibody test(FAT) assays were all initially performed at the Department of Agri-culture, Fisheries and Forestry Biosecurity Queensland Veterinary Ser-vices Laboratory.18
Histopathology. Histopathological examination of the cerebralcortex, cerebellum, mid-brain and cervical spinal cord samples fromhorse 2 demonstrated moderate subacute, diffuse, non-suppurativemeningo-encephalomyelitis. Some neuronal degeneration was evidentin the cerebrum and cerebellum. Eosinophilic intracytoplasmic viralinclusions (Negri bodies) within neurones were seen occasionally in allsections and were most prominent in cerebellar Purkinje neurones(Figure 3). Predominantly lymphocytic perivascular cuffing (paren-
Figure 2. Veterinarian and handler in personal protective equipmentwith horse 2 at 12 h after clinical onset demonstrating a wide-basedstance because of ataxia, head pressing and mydriasis.
Figure 3. Histological image of cerebellum, showing Purkinje cells witheosinophilic intracytoplasmic inclusion bodies (arrows). H&E, scale bar =50 µm. Courtesy of Dr John Bingham, CSIRO Australian Animal HealthLaboratory.
chyma and meninges) with some recent petechiation was demon-strated in all sections (Figure 4). Both white and grey matter wereaffected. Other changes included oedema, haemorrhage and vacuolarchange within the white matter.
Bacteriology. Aerobic culture of CSF from horse 2 produced amixed growth signifying sample contamination.
Horse 2 virology. Real-time PCR performed on nucleic acidsextracted from brain tissue did not detect EHV-1 or -4 or HeV.
Lyssavirus antigens were detected on impression smears of sectionedfresh brain using the FAT assay. ABLV RNA was detected by reversetranscriptase PCR (RT-PCR) in brain and from an oral swab.
Sequencing of virus isolated from the brain in a mouse neuroblastomacell line revealed a 97.7% identity with ABLVs.18
Samples were forwarded to the CSIRO Australian Animal HealthLaboratory, where lyssavirus antigen was again detected by FAT per-formed on brain tissue. Real-time RT-PCR tests on brain tissuedetected ABLVs but not ABLVp. Real-time RT-PCR tests performedon RNA extracted from brain tissue did not detect Murray Valleyencephalitis virus or West Nile group viruses.
Queensland Health Forensic and Scientific Services Laboratorydetected lyssavirus RNA from brain tissue by conventional RT-PCRand real-time RT-PCR. Lyssavirus antigen was detected, for a thirdtime, by FAT on brain tissue (Figure 5).
Horse 1 virology. Following the results from horse 2, viral swabssubmitted to Biosecurity Queensland for HeV exclusion for horse 1were tested by real-time RT-PCR for ABLV. Only the oral swab testedpositive for ABLVs. This result was confirmed at Queensland HealthForensic and Scientific Services Laboratory by conventional and real-time RT-PCR.
Activities subsequent to confirmed ABLV diagnosisQuarantine restrictions were placed on the property by BiosecurityQueensland and risk assessment of animals was undertaken acting onestablished AUSVETPLAN11 and Biosecurity Queensland protocols.19
The three dogs and four horses in closest contact with the affectedhorses were each administered two doses of Nobivac® inactivatedrabies vaccine (MSD Animal Health, Bendigo East, VIC, Australia) 7days apart. Serological antibody testing by rapid fluorescent focusinhibition on blood sampled 21 days after the second vaccinationconfirmed that all vaccinated animals had seroconverted (titres>2 IU). No animals had demonstrated positive antibody titres prior tovaccination.All animals were monitored in quarantine for a further 21days.19
Queensland Health identified nine in-contact people and they electedto receive post-exposure prophylaxis (PEP).20 The attending veteri-narian, who had prior rabies vaccination prophylaxis, received twodoses of killed strain rabies vaccine. The other eight people, who hadnot been previously vaccinated, received the recommended PEP.
All in-contact people and animals remained well 12 monthspost-exposure.
Potential routes of exposureThe vector was not identified. Insectivorous microbats identified asGould’s wattled bat (Chalinolobus gouldii) were observed roosting in afeed shed on the property. This genus is one of those reported to havedemonstrated serological evidence of ABLV exposure.13 Soundrecordings made over 3 days by a bat epidemiology team identified 10species of microbat but not the yellow-bellied sheathtail bat. Therewere trees suitable for harbouring microbats in the infected horses’paddock.
Figure 4. Histological image of brain stem, showing mononuclear cellperivascular cuffing around small blood vessels (arrows) and glial reac-tion. H&E, scale bar = 100 µm. Courtesy of Dr John Bingham, CSIRO Aus-tralian Animal Health Laboratory.
Figure 5. Image of a fluorescent antibody test performed on brain tissuedemonstrating presence of lyssavirus antigen. (Image courtesy ofQueensland Health Forensic and Scientific Services Laboratory andDepartment of Agriculture, Fisheries and Forestry Biosecurity Queens-land Veterinary Services Laboratory.)
The two equine cases reported here demonstrate that ABLV can infectdomestic animal species, which has previously been acknowledged butnot confirmed.
The prevalence of ABLV in Australian bats is similar to that of otherlyssaviruses in bats worldwide.7,21 ABLV prevalence in wild healthybats is estimated at <1%,13,15 but appears to vary between bat species.15
It can be far higher in injured, orphaned or sick bats and has beenfound in 30% of bats showing central nervous system (CNS)signs.13,15,20 Seroprevalence of lyssavirus antibody in the yellow-belliedsheathtail bat was significantly higher (up to 62.5%) than in otherspecies, suggesting that this bat plays an important role in the ecologyof ABLV.13
In developed countries, lyssavirus is mainly confined to sylvatic cyclesof small carnivores and bats, and domestic animals represent a minor-ity of total lyssavirus infections in most regions that have undergoneeffective surveillance.7,21
Rabies and lyssavirus infections are regarded as low-incidence buthigh-consequence diseases because of the high fatality rate, approach-ing 100%. All three of the recorded human fatal cases of ABLV inAustralia resulted from contact with infected bats.22–25 High-riskhuman interactions with bats are relatively common and most fre-quently occur with sick or injured bats.2 Domestic animals have beenshown to be involved in 12% of human–bat interactions, usuallywhere a domestic pet had attacked or retrieved a bat.2
In the 4 years after the discovery of ABLV, 189 notifications of poten-tial human exposure requiring PEP were recorded in Queensland.2
Notifications were lower during the following period until 2013, when494 were reported, which represented a three-fold increase over2012.26
These statistics demonstrate the potential for spillover of infection.
Lyssavirus transmissionTransmission of lyssavirus to humans from domestic species is typi-cally by a bite of an infected animal,27 The bite of an infected batinoculates lyssavirus-laden saliva into the epidermis, dermis, subcuta-neous and possibly muscle tissue. Bites by insectivorous bats areminute, likely to go unnoticed and often not reported.27 Somelyssavirus variants may be more infectious than others when they aresuperficially inoculated into epidermis, where they may replicate morereadily in non-neuronal cells and at lower temperatures.28
Virus transmission by contamination of scratches or skin abrasionswith saliva of an infected animal is also possible; isolation of RABVfrom equine salivary glands has further demonstrated this potential.29
In 2012 in the USA, 47 of the 519 domestic animal cases of RABV werein horses and mules, which represented 5% of equine cases submittedfor testing.7 Young animals may be at greater risk because of theirinquisitive nature and immature immune response.30 In 19th centuryEurope, RABV was reported to have spread from dogs to horses andcoachmen,31,32 and rare cases of rabies in people contracted frominfected equids have been reported.29 One human death resulted froma rabid donkey bite in Ethiopia33 and two cases of equine-transmittedrabies were reported in Brazil.34
ABLV was present in the saliva of both horses in our study, as shownby PCR of oral swabs, demonstrating the potential for ABLV infec-tions in horses to pose a zoonotic risk comparable to RABV. Obtaininga prompt and accurate laboratory diagnosis of ABLV in animal casesis fundamental in identifying exposure.29
A veterinarian in Brazil died after contracting the disease throughtreatment of infected ruminants.35 Transmission by handling infectedcarcases and aerosol transmission in a laboratory environment havealso been reported.36
In the ABLV cases reported here, multiple bat species were identifiedin the infected horses’ paddocks and insectivorous bats were foundroosting in a feed shed. Transmission by eating feed contaminatedwith urine and/or faeces or an infected fresh carcass is consideredmore unlikely than by direct contact with bats, because of the insta-bility of lyssaviruses in the environment. A plausible scenario is thatthe inquisitive young horses approached an infected bat and werebitten on the face, as has been suggested in equine RABV cases.31 It isalso theoretically possible that the infections resulted from a bite of anunidentified infected carnivore.
Incubation periodIncubation periods of lyssavirus infections are extremely variable,ranging from days to years, and are thought to be affected by the virusstrain, host species, inoculum dose and proximity of the inoculationsite to the CNS.37 The average incubation in 21 experimental cases ofequine RABV, where inoculation had occurred in the masseter muscle,was 12.3 days.38 Retention of the virus in the myocytes and replicationin non-neural tissue at the site of inoculation may be a mechanism forthe prolonged and varied incubation.39 Virus migration and replica-tion may be associated with periods of stress.30 In most cases of bat-related infection the exposure is not witnessed, making assessment ofincubation periods difficult. Preliminary analysis in mice has shownthat ABLVs may have a shorter incubation period than ABLVp.40
In the two cases of ABLV reported here, the timing of infection wasunknown. The close timing of clinical onset between cases suggeststhat incubation periods were likely to have been similar and that asingle bat may have infected both horses.
Clinical signsIt is important for veterinarians to be aware of the variable clinicalsigns likely in horses infected with ABLV, given the limitations, delaysand difficulties in diagnosis. The clinical signs and disease courses ofABLV in these two horses were consistent with those seen in equinecases of RABV outside Australia.30-32,37,38,41
Clinical signs of rabies. No clinical signs are pathognomonic andovert neurological signs may not appear until late in the diseasecourse.31 Early signs can be vague, subtle, inconsistent and non-specific, ranging from lameness or colic to sudden death and includechanges in behaviour, mild pyrexia, hyperesthesia, ataxia, anorexia,frequent whinnying, apparent thirst but inability to swallow andparalysis or paresis.30,31,37
Neurological signs of lyssavirus infection in all animals may be clas-sified into spinal (paralytic), brainstem (dumb) and cerebral (furious),
depending on the neuro-anatomical location of the viral infection.37
Infected horses may demonstrate one, two or all three categories ofclinical signs, reflecting progression of infection through the CNS.
In horses with RABV, spinal signs typically include progressiveascending paralysis, ataxia, shifting lameness, hyperaesthesia,areflexia, analgesia, atonia and self-mutilation of an extremity.Brainstem signs typically include depression, anorexia, head tilt, cir-cling, ataxia, dementia, excess salivation, facial and pharyngeal paraly-sis, blindness, flaccid tail and anus, urinary incontinence and self-mutilation. Cerebral signs typically include aggressive behaviour,photophobia, hyperaesthesia, straining, paddling and cycling move-ments, muscular tremors and convulsions. Severe, overt, repeated anduncharacteristic aggression has been reported in a few cases.30,32,37,38
Progressive spinal to brainstem clinical signs have been describedmost frequently,30 with cerebral signs appearing mostly later in thedisease course. Infected horses may remain bright enough to eat anddrink even when recumbent;30 death usually occurs 3–7 days afteronset of clinical signs.31
Rapidly progressing neurological signs or those reflecting diffuse dys-function are strong warnings for RABV.37 One clinical descriptionstrongly suggestive of RABV is a recumbent horse that is ataxic, hasparalysis and anaesthesia of the limbs, loses tail and anal sphinctertone and demonstrates pyrexia and hyperesthesia.30
Clinical pathologyNo specific abnormalities in haematological or serum biochemistryvalues have been associated with equine RABV.30,37 Results of CSFsampling show either no changes41 or non-specific abnormalities con-sistent with viral encephalomyelitis.30,37
In the two cases of equine ABLV described here, no specific abnor-malities in haematological or serum biochemical values were observedand CSF analysis other than for bacteriological examination in horse2 was not performed.
Necropsy findingsIn cases of equine RABV, there may be no grossly appreciable post-mortem changes, or only inconsistent and non-specific changes,including mild/diffuse cerebral oedema, meningeal congestion andfocal areas of haemorrhage,30,38,41 consistent with the findings for horse2, in which there was marked injection of the meningeal vasculature.
One experimental case of rabies demonstrated red and malacicmultifocal areas within the grey matter of the spinal cord and othersshowed discolouration of the trigeminal ganglion with a greyishcast (likely to have been the nerve that underwent viral axoplasmicmigration).38
Laboratory diagnosisMethods for diagnosing ABLV infection are similar to those used forRABV and other lyssaviruses.11
Hitstopathological analysis. Histopathological changes consistentwith lyssavirus infection are seen most commonly in thehippocampus, brainstem, cerebellum and grey matter of the spinalcord and comprise a mild, non-suppurative encephalomyelitis,perivascular cuffing by mononuclear cells, gliosis, glial nodules and
neuronal degeneration.37 Negri bodies are considered pathognomonicfor lyssavirus infection37 and may be more likely to be seen in horsesthat survive >4 days after the onset of clinical signs.30 In experimentalcases involving inoculation of RABV into the masseter muscle,lymphocytic ganglioneuritis was seen in the trigeminal ganglion.38
The similar histopathological changes seen in horse 2 provided strongindication for performing subsequent laboratory tests.
Diagnostic tests. Testing of the brain, where possible, is importantbefore ruling out the possibility of ABLV.11,15 FAT performed on CNStissue is the initial test of choice because it is the most rapid, and PCRtesting has shown that FAT for ABLV on fresh brain touch impres-sions is highly sensitive.15,29 It involves the application of fluorescein-labelled antibodies specific to the viral nucleocapsid proteins tosmears of brain, brainstem and spinal cord tissues. Typically, thebrainstem and cervical medullar regions present the highest levels ofviral antigen.29 Current FAT reagents react to all lyssaviruses anddifferentiation of ABLV requires characterisation of the viral genomeby molecular genetic techniques, such as real-time RT-PCR andsequencing.11
Antemortem testing has proven effective in both human and animalcases of RABV.35,42 Serum and CSF may be tested for rabies viralneutralising antibodies by rapid fluorescent focus inhibition. Allsamples except serum can be processed for detection of rabies viralRNA by real-time RT-PCR.43 Saliva also may be tested by virus isola-tion. Skin biopsy specimens are examined using immunofluorescencefor rabies antigen in the cutaneous nerves at the base of hairfollicles.44,45
One study that included 12 horses found a high correlation ofimmunofluorescent positivity between skin biopsied cranium-derivednerve fibres and postmortem brain tissue samples from clinical andexperimental animal rabies cases.42
A study from India showed antemortem rabies diagnosis in 84.6% ofcases using RT-PCR, with diagnosis in all cases when combined withrabies viral neutralising antibodies detection in CSF.43
In the presented horses, ABLV was diagnosed through demonstrationof viral antigen in saliva by real-time RT-PCR performed on oralswabs. Minimally invasive collections for HeV exclusion include asaliva swab and whole blood (serum). Similar samples could beassayed for ABLV exclusion and flavivirus. A small optimised skin andnerve-ending biopsy may also be useful.
Diagnostic challengesRisk to personnel. Equine CNS sample collection is difficult toperform and places the veterinarian at heightened risk of viral expo-sure.35 Appropriate pre-exposure prophylaxis through completeimmunisation is essential before performing necropsy and CNSsample collection.35 In Australia, necropsy should not be performedon horses that have demonstrated neurological signs until HeV hasbeen excluded. Recommended options for tissue sampling and labo-ratory testing are listed in AUSVETPLAN.11 Before shipping speci-mens, submitters should contact the receiving laboratory to discussarrangements for sampling, transport and sample reception.11,19
Underdiagnosis of lyssavirusIn human medicine, recent publications have highlighted concernsregarding the possibility of under-diagnosis of lyssaviruses.25,28,46-48
Reasons for failure of diagnosis in both human and veterinary medi-cine include that transmission of the virus may go unnoticed, post-mortem testing is not always performed and clinical signs are oftenvaried and non-specific.27,28,32,46
A retrospective study of human patients with unexplained encepha-litis in the Northern Territory found that very few samples underwentor were available for lyssavirus testing.47 Another study found up to70% of adult encephalitis hospitalisations in Australia have no causeidentified,48 even though encephalitis syndrome may include novelpathogens with outbreak potential. As individual treatment and aneffective public health response rely on identifying a specific patho-gen, the authors suggest that improved encephalitis surveillance andthe use of a standardised testing algorithm, including ABLV, may bebeneficial. A recently published serological survey of human bat han-dlers in 1996 detected a rising antibody titre to ABLV in a 40-year-oldman, who then underwent PEP vaccination and has remained well todate.49
It is possible that ABLV may not have been considered in the pastwhen equine patients showed clinical signs consistent with infectionand were euthanased or died as a result of their illness. There areanecdotal accounts of cases in which clinical signs were very similar tothose seen in the two cases presented here, but in which no diagnosiswas reached and ABLV laboratory screening was not performed.
Treatment of horsesThere is no effective treatment for animals clinically affected bylyssaviruses. Prolonging the disease course may be consideredcontraindicated because it significantly increases the risk of transmis-sion to other animals, clinicians and handlers.37
Treatment of equine cases of RABV has been symptomatic and sup-portive, usually targeted at undifferentiated neurological disease andwhich, at best, may prolong the disease course to allow completion ofdiagnostics.30,37 Treatment similar to that instituted for both of thepresented horses has been reported for the treatment of equine RABV,including antibiotics, B vitamins, dimethyl sulfoxide in lactatedRingers solution, flunixin meglumine, oral electrolytes, sodiumselenite/vitamin E and dexamethasone.41
VaccinationPre-exposure vaccination is the most successful form of RABV pre-vention in humans and animals.28 No rabies deaths are known to haveoccurred in humans who received a pre-exposure vaccine course fol-lowed by a booster dose after exposure.28
A murine study showed that both of the human and animal rabiesvaccines used in Australia were effective against ABLVp when usedpre-exposure and in PEP, in conjunction with anti-rabies immuno-globulin (an ABLVs isolate was not available for challenge).40 An invitro and murine study concluded that although a rabies humandiploid cell vaccine effectively neutralised ABLVs measured by in vitroantibody assay, only half of the mice survived intracranial challengeafter two doses of vaccine. Following foot pad challenge, protectionwas significant but only 80%.50 Given also that ABLVs may be associ-ated with shorter incubation periods and so could provide a greaterchallenge for successful PEP, further research is warranted.40
Australia currently meets the World Organisation for Animal Health(OIE) requirement for rabies freedom. With the exception of meetingimport and export requirements, routine pre-exposure vaccination ofanimals has not been permitted in Australia to date (Barrett J, CrookA, Queensland Dept of Agriculture Fisheries and Forestry, pers.comm.)
Post-exposure prophylaxisPEP in humans consists of appropriate and early washing of wounds,or flushing of exposed mucous membranes, a course of vaccinationsand possibly wound infiltration with rabies immunoglobulin.20 Batbites or scratch wounds on animals should be washed for 5 min withsoap, lavaged and a virucidal agent such as povidone iodine applied.There are reports of the use of infiltrated rabies immunoglobulin inanimals overseas.7 Rabies immunoglobulin of human origin is limitedand restricted to human use. That of equine origin is considerablycheaper but availability is limited51 and approval for its importationand use in Australia could be problematic.
Post-exposure vaccination of domestic animals with rabies vaccineduring the preclinical period may prevent the development of clinicaldisease by preventing local viral replication and neuronal transmis-sion. In the USA between 2000 and 2009, 1014 animals received PEPagainst RABV, without any failures recorded.52
In Australia, Queensland guidelines specify a protocol for the approvalfor PEP through vaccination of domestic animals.19 Several animalshave been vaccinated following exposure to bats, with no confirmedsubsequent infections to date. The protocol involves a primary dose ofrabies vaccine (day 0) administered together with the collection of ablood sample, a second RABV vaccine administered 5–7 days later anda second blood sample 28–35 days following the first. The pairedsamples are tested for seroconversion. The animal should be moni-tored for clinical signs for a minimum of 60 days, after which restric-tions may be lifted.19 Permission for use of vaccine must be soughtthrough the Chief Veterinary Officer and conditions for monitoringthe animals are regulated.19 Strict compliance with quarantine andmonitoring of exposed, or potentially exposed, vaccinated animals isrequired.19
Conclusions
ABLV presents a significant zoonotic risk and, as with otherlyssaviruses worldwide, under-diagnosis is likely. The spillover tohorses reported here and in vitro ABLV tropism analyses17 indicatethat animals other than bats could pose potential human healththreats. Further neurological disease surveillance would be beneficialto increase our understanding of the disease and its zoonotic risk.
PEP in humans is effective but relies on prompt administration fol-lowing exposure. Where exposure is through domestic animals, anearly, accurate diagnosis is critical. In these situations, veterinary prac-titioners can facilitate diagnosis through clinical evaluation, collectionof samples, viral exclusion testing and performing or facilitating CNSnecropsy. Sample collection at necropsy should only be considered inAustralian horses after adequate immunisation, HeV exclusion andthe use of PPE.
The authors recommend that veterinarians strongly consider receiv-ing pre-exposure prophylactic vaccination. The subtleness and lack ofspecificity of initial signs means that appropriate caution in avoidingexposure to the virus through PPE is almost certain to be oftendelayed. Consequently, protection through avoiding contact in‘suspect’ cases may be ineffective and, for this reason, vaccination ofveterinarians is even more important.
It is hoped that the reporting of these cases will increase Australianveterinary and public health awareness of the potential for ABLVinfection in terrestrial animals and encourage consideration of ABLVas a differential diagnosis in progressive neurological cases.
Acknowledgments
The authors acknowledge the assistance of Biosecurity Queensland,Department of Agriculture, Fisheries and Forestry (BiosecurityScience Laboratory, Queensland Centre for Emerging InfectiousDisease and field operation), Coopers Plains, Queensland; QueenslandGovernment Department of Science, Information Technology, Inno-vation and the Arts; the Australasian Bat Society, which providedtechnical expertise on microbat surveys; Pathology Queensland Sci-entific Services (QHFSS) Coopers Plains, Queensland and CSIROAustralian Animal Health Laboratory, Geelong, Victoria.
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(Accepted for publication 14 June 2014)
OBITUARY
Allan McTackett
1920–2014
A llan McTackett was born on 5 July1920 in Maitland, NSW where hisfather managed a milk run/dairy farm
at nearby Bolwarra. He attended MaitlandBoys’ High School and graduated from theUniversity of Sydney in 1943 and became aregistered veterinarian.
Allan started his veterinary career conduct-ing research into the values of supplemen-tary feeding of dairy cattle in a time whenlimited feed was available. In 1944, Allanreturned to Maitland where he set up thefirst graduate veterinary practice in the area.
Allan married his wife Shirley in 1948. Theyboth grew up in Maitland and knew of eachother but only started dating when theyquite literally collided. Shirley was riding herbike and was knocked off it by Allan as he swung open his car door.
After spending 4 years in Maitland, Allan and Shirley moved tonorth-east Victoria, near Wangaratta, where Allan worked withthree cooperative butter factories as a veterinarian to their suppli-ers. In 1953, Allan accepted a research officer position with theQueensland Department of Agriculture and Stock. He investigatedherd fertility in dairy cattle, which incidentally became the subjectof his thesis. After 10 years of investigation into ‘Infertility in dairyherds of south east Queensland’, Allan was granted a Mastersdegree by the University of Queensland.
In 1963, Allan and Shirley moved back to Maitland where they grewtheir mixed practice. Allan once described Shirley’s role as, ‘whatnow-a-days you would call a practice manager. She also filled theshoes of a veterinary nurse on many occasions. Without Shirley Iwouldn’t have been anything,’ he said. While in Maitland, Allan alsospent some time serving as the official veterinarian to the Maitland,Cessnock and Newcastle Trotting Clubs and to the GreyhoundRacing Club and voluntary service to local pony clubs.
Allan became the veterinarian at the African Lion Safari nearRaymond Terrace and, on more than one occasion, he brought lionand tiger cubs home for treatment. The laundry also became akoala sanctuary where sick and injured koalas could recuperate.
He once recalled that, unlike most mixedpractice vets, who go to treat a cow and getasked to look at a rooster’s claw while you’rehere, ‘I was at the lion park treating a lion andthat all too familiar while you’re here cameout of one of the staff’s mouth and all of asudden I’m 18 feet up a ladder peering in agiraffe’s mouth waiting for Shirley to arrivewith some stuff to treat worms!’ he said.
After spending 15 years in Maitland, Allanand Shirley relocated to Nelson Bay wherethey set up another small veterinary clinic.They treated their patients like they werefamily and when setting up a veterinarypractice it would always be on the sameproperty as their home so that they couldcare for them throughout the night. It meant
that the McTackett home often resembled a menagerie. They ranthe practice until they retired in 2001. They remained in PortStephens for a short while before returning to their hometown ofMaitland, where it all began.
Allan was granted a honourary life veterinary surgeon registrationby the New South Wales Veterinary Board and a honourary lifemembership of the AVA and also of the Newcastle Branch. He alsoserved a time as the President of the Hunter River Division of theAVA. He had been a rotary member since 1968, being named a PaulHarris Fellow. He was a cricket enthusiast with a 1500 volume libraryfull of cricket memorabilia and literature.
Allan thoroughly enjoyed his career as a veterinarian but oftenremarked that his greatest satisfaction came from the years hespent in research, as he viewed his work in this area as a way tocontribute to the advancement of veterinary science.
Allan died on 3 January 2014, just two days shy of his 94th birthday.He is survived by his loving wife Shirley, three children, two grand-children and one great-grandchild.
Warren McTackett
doi: 10.1111/avj.12235
The University of Sydney veterinaryscience students, 1942. Allan is picturedin the far right of the photo (middlerow).
