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1 Rabies and Wildlife Zoonoses Group, WHO Collaborating Centre for the Characterisation of Rabies and Rabies-Related Viruses, Veterinary Laboratories Agency – Weybridge, Woodham Lane, Addlestone, Surrey KT15 3NB, UK
2 IDT Biologika GmbH, Am Pharmapark, D-06861 Dessau-Roßlau, Germany
3 Institute for Epidemiology, WHO Collaborating Centre for Rabies Surveillance and Research, Friedrich-Loeffler-Institute, Federal Research Institute for Animal Health, Seestrasse 55, D-16868 Wusterhausen, Germany
4 Zoologisches Institut, Eberhard Karls University, Auf der Morgenstelle 28, 72076 Tuebingen, Germany
5 Centre for Disease Control and Prevention, 1600 Clifton Road NE, Atlanta GA30333, USA
Correspondence
N. Johnson
n.johnson2{at}vla.defra.gsi.gov.uk
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). Inoculation of brain homogenate from the bat into mice induced progressive fatal encephalitis, and Negri bodies were detected in brain smears. At about the same time, rabies virus was also identified in insectivorous bats in North America (Scatterday, 1954). Sporadic reports of rabies-like virus followed in Europe, usually associated with the Serotine bat (Eptesicus serotinus) and mainly from Germany (Seidler et al., 1987). It was only through the development of monoclonal antibody (mAb) typing (Wiktor & Koprowski, 1978) that the lyssaviruses isolated from European bats could be differentiated from classical rabies virus; they were found to be antigenically similar to Duvenhage virus, a lyssavirus isolated exclusively from Africa (Tignor et al., 1977). Both Duvenhage and European lyssaviruses were classified as lyssavirus serotype 4. However, virus-typing using extended panels of mAbs identified a number of European exceptions. The first was a virus isolated from a human case of rabies in a bat ecologist who died in Finland (Lumio et al., 1986), the second were a number of viruses isolated from pond bats (Myotis dasycneme) in The Netherlands (Van der Poel et al., 2005). These viruses were tentatively classified as serotype 5 (King et al., 1990). Subsequent genomic sequencing confirmed this distinction and led to a revision of lyssavirus classification with two distinct viruses, European bat lyssavirus (EBLV)-1 and EBLV-2, being recognized (Bourhy et al., 1992). The first UK case of EBLV-2 in a Daubenton's bat (Myotis daubentonii) was recorded in 1996 (Whitby et al., 2000), with a second case isolated from the same species in 2002 (Johnson et al., 2003). This coincided with a human case in a bat worker in Angus, Scotland (Fooks et al., 2003a). Extensive surveillance in a number of countries in Europe has established that the reservoir species of EBLV-1 is the Serotine bat (Van der Poel et al., 2005; Müller et al., 2007), whilst the reservoir species of EBLV-2 are bats of the Myotis genus, exclusively M. dascycneme and M. daubentonii. For reasons that are not understood, EBLV-1 is prevalent throughout mainland Europe but not in the UK, whereas EBLV-2 appears to be restricted to the UK, The Netherlands and the Swiss/German border (Freuling et al., 2007).
EBLV-2 has been isolated from six Daubenton's bats in the UK between 1996 and 2007 (Harris et al., 2007) and serosurveys of bat populations throughout England and Scotland suggest that the virus is endemic in the species but prevalence is low (Brookes et al., 2005a). The Daubenton's bat belongs to the family Vespertilionidae and is found throughout Europe as far south as Greece, with a northern limit of central Finland. Colonies of Daubenton's bats have been found throughout the UK, where it is the third most abundant bat species after the common pipistrelle (Pipistrellus pipistrellus) and the brown long-eared bat (Plecotus auritis). All bats within the UK are protected by both national and European legislation, although the Daubenton's bat is not considered to be under threat and numbers may be increasing. After two well-documented cases of spill-over to humans, in both cases causing rabies, it is clear that the virus has the capacity for transmission to humans. In both cases, the individuals were unvaccinated and had recent contact with Daubenton's bats. This implies that bite transmission, in a similar manner to North American bats (Messenger et al., 2002), is the principal means of dissemination of this virus.
Cross-neutralization and cross-protection studies have demonstrated that current anti-rabies vaccines provide protection against EBLV-2 (Brookes et al., 2005b) and all bat handlers in the UK are required to receive vaccination. A limited number of pathogenesis studies have been conducted on the EBLVs using a range of animal models (Vos et al., 2004; Johnson et al., 2006a; Brookes et al., 2007). These have suggested that EBLV-2 is less virulent in animal models including mouse, sheep and fox, when compared directly with EBLV-1, and both are less virulent than rabies virus (RABV). However, it is not clear if this is true for the virus within its reservoir host. The case-reports of EBLV-2-infected bats indicate that unprovoked biting is commonly associated with rabies in bats (Table 1). This contrasts strongly with studies in Spain where apparently healthy bats have been shown to excrete EBLV-1 in saliva (Echevarria et al., 2001; Vazquez et al., 2006), suggesting the possibility that an asymptomatic carrier state could exist in lyssavirus-infected bats. From a public health perspective it is important to confirm or exclude this possibility.
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) and that this species is likely to be the reservoir host. The principal aim of this study was to investigate the susceptibility of the Daubenton's bat to inoculation with EBLV-2 and to study the spread of virus within its reservoir host. |
METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Thirty-three bats were assigned into five groups and housed in purpose-built metal-framed cages (75 cmx75 cmx75 cm) with fine-mazed wire gauzed side sheets. The remaining bat was not considered to be sufficiently healthy to enter the study and was retained by a bat rehabilitator. Two cages were held within one scantainer (Scanbur), which provided a closed, filtered-air system with constant temperature and humidity. Three scantainers were held within a secure containment laboratory under negative air pressure and controlled access. Animals were inspected daily to monitor their condition and development of disease.
Virus.
The EBLV-2 isolate used in this study was obtained from a Daubenton's bat found in Lancashire (Johnson et al., 2003) and designated RV1332. Virus was passaged three times by intracerebral (i.c.) inoculation in female 5-week-old Swiss OF-1 mice (Charles River Laboratories, France) until a titre of 4.9 log10(MLD50) per ml was reached. A 20 % mouse brain homogenate was prepared and clarified by centrifugation at 3200 g for 10 min, then immediately frozen at –80 °C until required.
Inoculation and monitoring.
Four routes of inoculation of 20 µl of EBLV-2 infected mouse brain homogenate were used. Intranasal (i.n.) and i.c. inoculations were performed under light ether anaesthesia. For i.n. inoculation, the inocula was placed on the nasal passage and inhaled. For intramuscular (i.m.) inoculation, either EBLV-2 or negative control mouse brain homogenate was injected into the pectoral muscle. Subdermal (s.d.) inoculation was performed by injecting EBLV-2 homogenate under the skin in the scruff of the neck. This gave a final inoculation dose of 3.2 log10(MLD50) that consistently induced disease in mice through both i.c. and s.d. routes. Lower doses of virus have produced variable outcomes in the murine model. All animals were monitored closely for one hour after inoculation. Subsequently, all animals were observed twice daily for 120 days. Saliva swabs were taken 1 week prior to inoculation and at weekly intervals, or daily if individual bats showed signs of disease. A cotton swab was held within the mouth until moistened and placed in 0.4 ml minimal essential medium with gentamicin (50 mg ml–1) and amphotericin B (2.5 mg ml–1). All swabs were frozen immediately and stored at –20 °C until processing. Body weight was measured at weekly intervals throughout the study.
Blood samples were taken either from the vena brachialis during the study period, or by cardiac puncture on the development of severe disease (paralysis) or at the completion of the experiment (terminal bleed after euthanasia). Serum samples were prepared and stored at –20 °C until testing. Bat carcasses were frozen immediately and stored at –20 °C until necropsy. One animal from each group was fixed in formalin immediately after death for pathological examination.
Virus detection.
The fluorescent antibody test (FAT) was performed following standard protocols (Dean & Abelseth, 1973). Virus isolation from saliva swabs (200 µl) was performed using the rapid tissue culture inoculation test (RTCIT) as described previously (Webster & Casey, 1996). At least three consecutive passages were conducted to prove a negative result by RTCIT.
Detection of EBLV-2 RNA.
RNA was extracted from organ samples using Trizol (Invitrogen) following the manufacturer's protocol. RNA was extracted from saliva swabs (200 µl) using an RNeasy column (Qiagen) following the manufacturer's protocol. For detection of virus genome, two RT-PCR methods were utilized. The first was the nested RT-PCR described by Heaton et al. (1997). Amplicons were separated on 1 % agarose gels and visualized by ethidium bromide (1 µg µl–1) staining and UV illumination. The second assay was a probe-based real-time (q)RT-PCR assay described by Wakeley et al. (2005). This was modified using primers Jw12 (5'-ATGTAACACCYCTACAATG-3') and N165-146 (5'-GCAGGGTAYTTRTACTACTCATA-3') where indicated, to incorporate SyBr green using the SyBr green JumpStart Taq ReadyMix assay (Sigma) to enable quantification of genome copies as described by Johnson et al. (2006a). β-actin primers (Wakeley et al., 2005) and 18S RNA primers (Ambion) were used to detect host mRNA within samples.
Rapid fluorescent focus inhibition test (RFFIT).
Virus neutralizing antibody (VNA) against EBLV-2 was measured with a modified RFFIT (Cox & Schneider, 1976) by replacing challenge virus standard (RABV) with EBLV-2. Samples were tested in duplicate with twofold serum dilutions starting at 1 : 10. A heterologous WHO international standard immunoglobulin (2nd human rabies immunoglobulin preparation, National Institute for Standards and Control, Potters Bar, UK), adjusted to 1.5 IU ml–1, served as a positive control. The VNA titre was expressed as the reciprocal of the serum dilution showing a 50 % reduction in concentration of the EBLV-2 test virus in vitro. Sera from bats were assessed prior to inoculation and at the end of the study.
Immunohistochemistry.
Following fixation in 10 % buffered formalin, brain, spinal cord, salivary glands, liver, kidney, lung, spleen and intestinal samples were blocked and routinely processed to paraffin wax. Immunohistochemical detection of lyssavirus nucleocapsid was performed as described by Hicks et al. (2008). Briefly, serial 4 µm sections were hydrated and successively incubated with hydrogen peroxide block (BHD), normal goat serum block, primary antibody anti-rabies nucleocapsid mAb HAM (obtained from the Swiss Rabies Centre, Bern, Switzerland), biotinylated goat–anti-mouse secondary antibody, avidin–biotin-peroxidase conjugate (Vector Laboratories) and 3,3-diaminobenzidine (Sigma). Sections were then counterstained with Meyer's haematoxylin (Surgipath) and mounted.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). Of the animals challenged by peripheral routes, only one animal (Y2070) developed rabies, at 33 days following inoculation by the subdermal route. At day 30, it developed disease that progressed rapidly to severe paralysis by day 33 and it was euthanized. Weight of this bat had decreased from 10 g at inoculation to 5.4 g at death. Antigen detection in the brain confirmed that these six animals (Y2495, Y2505, Y2516, Y2556, Y2566 and Y2070) had died of EBLV-2 infection (Table 2). One animal from each group was investigated for lyssavirus antigen using immunohistochemistry. Of five animals investigated this way, only the bat showing clinical signs of disease from the i.c. group (Y2556) showed evidence of lyssavirus nucleoprotein immunolabelling within the brain and spinal cord. Specific immunolabelling was detected in the perikarion and neuronal processes of neurons scattered through most regions of the brain (Fig. 2a, b) and spinal cord segments (Fig. 2c). Immunolabelled neurons were also observed outside the central nervous system in dorsal root ganglia neurons (Fig. 2d) and enteric plexus neurons. No viral nucleoprotein was detected in other tissues, including salivary glands, using this technique.
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Two animals died during the 123 day experimental period, although these could not be attributed to EBLV-2 infection and were considered non-specific. The remaining animals survived to the end of the experiment. Both nested RT-PCR and a qRT-PCR assay corroborated the antigen detection assays, with both detecting EBLV-2 genes in all five bats from the i.c. group and the single bat from the s.d. group (Table 3).
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). Host (18S) RNA presence was confirmed in all but two samples. In contrast, EBLV-2 RNA was detected in the salivary gland, tongue, heart, lung and kidney samples of the single bat that succumbed following s.d. inoculation (Table 4 and Fig. 3a). Furthermore, EBLV-2 RNA was detected in saliva samples taken from this bat between days 28 and 32 (Fig. 3a), preceding the development of clinical signs of rabies by 2 days. The genomic sequence generated from these amplicons demonstrated that all sequences showed 100 % identity with EBLV-2 strain RV1332 (data not shown). This finding was confirmed by virus isolation from saliva swabs taken from this s.d.-inoculated bat on three consecutive days prior to its death. However, this required three cell passages of each sample before virus was detected in the cells, verifying the low levels of viable virus in saliva. Further analysis of organ samples from this bat using qRT-PCR detected EBLV-2 RNA in all samples tested, but at very low levels (Fig. 3b), with the exception of the brain where very high levels of virus were observed. Relatively higher levels of genome copies (100–1000 copies per ml RNA) were found in the salivary gland, tongue, heart, lung and kidney, in agreement with nested RT-PCR detection of viral RNA.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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), although distinctly divergent at the sequence level (Bourhy et al., 1992). Whereas RABV causes over 50 000 human deaths a year, is endemic within a range of animal reservoirs and is present throughout the world, EBLV-2 has only been detected in two humans, and is restricted to two bat species in a geographically limited area within Europe. Despite these differences, the virus clearly has the ability to kill animals, including its reservoir species, with disease manifestations that are virtually identical to classical rabies. EBLV-2 also displays a strict neurotropism (Johnson et al., 2006b) with sections of brain and spinal cord showing infected neurons (Fig. 2). The small numbers of natural cases of EBLV-2 also limit our understanding of the pathogenesis of this infection and it is this fact that has prompted us to investigate the virulence of this virus under experimental conditions. Surprisingly, experimental inoculation with a relatively large virus dose did not induce disease in the majority of animals (Table 2). Both the i.m. and i.n. inoculation routes used were ineffective at infecting Daubenton's bats. Only one of seven bats challenged by the s.d. route developed disease. Intensive investigation of this bat's saliva demonstrated that EBLV-2 genome could be detected in this bat up to 2 days before the development of disease. Previous studies that have infected bats with lyssaviruses have demonstrated virus excretion in saliva (Kuzmin et al., 2008) but this study is the first demonstration of EBLV-2, either by detection of viral RNA or virus isolation in bat saliva. However, the levels of viral RNA detected were at the lower level of reliable quantification, suggesting that the amount of virus shed is low, possibly below 100 genome copies per µg extracted RNA. We have only had the opportunity to assess viral load in the saliva of one bat. It is possible that some bats may have a greater virus concentration in the saliva. EBLV-2 genome was also observed in bladder and kidney samples, although at low levels (Fig. 3). Further studies to investigate possible virus secretion in bat urine may identify an alternative means of virus egress and transmission within bats.
The data obtained in this study are limited, with only one of seven bats inoculated by the s.d route developing disease and all bats challenged by the i.m. route surviving to the end of the experiment. This implies that the reservoir species is surprisingly resilient to infection. In most lyssavirus challenge models, the i.m. route is effective at inducing disease (Moreno & Baer, 1980) so the relative differences in efficacy of inoculation may reflect differences in innervation at particular sites in the Daubenton's bat. The inability of EBLV-2 to infect bats by i.n. inoculation repeats a previous observation made for EBLV-1 in Eptesicus fuscus bats (Franka et al., 2008). In roosts of European bats, in contrast with those observed in bat species in other parts of the world, the number of individuals of species such as the Daubenton's bat is relatively small (<100 individuals). This would not favour aerosol spread and thus EBLV-2 could have evolved away from transmission by this route. When directly compared under experimental conditions, RABV proved more effective than EBLV-2 at infecting mice by the airborne route (Johnson et al., 2006c), although a mouse is not the natural host for lyssaviruses, so these results should be interpreted with caution.
One possible explanation for sustained transmissibility could be that EBLV-2, like the silver-haired bat rabies variant (SHBRV) in North America, has become adapted to replication at the relatively lower body temperatures that are found in the dermis. SHBRV was shown to replicate faster at 34 °C than a RABV variant isolated from a coyote (Morimoto et al., 1996). Superficial bites from Daubenton's bats were believed to be responsible for both human cases of EBLV-2 (Fooks et al., 2003b). Another possible factor could be an extended incubation period. All of the Daubenton's bats in our study developed rabies within 32 days, with the animals inoculated by the i.c. route developing disease between 12 and 14 days. In one case of a naturally infected Daubenton's bat, an incubation period of 8 weeks was reported (Johnson et al., 2003). Although a 90 day observation period was planned, this was extended to 123 days to lengthen the opportunity to detect disease after a longer incubation period; however, none occurred and there was no evidence of salivary excretion of virus without development of disease as has been reported for RABV (Aguilar-Setien et al., 2005) and EBLV-1 (Echevarria et al., 2001). However, much longer incubation periods have been observed for RABV infections (Smith et al., 1991).
In the single case of rabies following peripheral inoculation of EBLV-2 registered in our study, viral RNA was detected in saliva 2 days before the development of disease. Similar observations have been made for RABV infection in both insectivorous (Bell et al., 1969) and vampire (Moreno & Baer, 1980) bats and Eurasian bat lyssaviruses in E. fuscus (Hughes et al., 2006). This indicates that EBLV-2 shows similar pathogenic characteristics under experimental conditions to other lyssaviruses. Within this experiment, we did not find evidence for an asymptomatic carrier state with virus excretion. However, virus excretion from an apparently healthy bat was observed by detection of viral RNA; the bat then developed clinical signs of rabies over the next 2 days. This reinforces the need for those that have contact with bats to have rabies pre-exposure vaccination and to wear protective gloves to prevent being exposed to a bite.
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ACKNOWLEDGEMENTS |
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Received 2 May 2008;
accepted 16 July 2008.
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