Monitoring of West Nile Virus Infections in Germany

Authors

  • U. Ziegler,

    1.  Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Institute of Novel and Emerging Infectious Diseases, Greifswald-Insel Riems, Germany
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  • D. Seidowski,

    1.  Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Institute of Novel and Emerging Infectious Diseases, Greifswald-Insel Riems, Germany
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  • J. Angenvoort,

    1.  Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Institute of Novel and Emerging Infectious Diseases, Greifswald-Insel Riems, Germany
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  • M. Eiden,

    1.  Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Institute of Novel and Emerging Infectious Diseases, Greifswald-Insel Riems, Germany
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  • K. Müller,

    1.  Department of Veterinary Medicine, Small Animal Clinic, Freie Universität Berlin, Berlin, Germany
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  • N. Nowotny,

    1.  Zoonoses and Emerging Infections Group, Clinical Virology, Department of Pathobiology, University of Veterinary Medicine, Vienna, Austria
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  • M. H. Groschup

    1.  Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Institute of Novel and Emerging Infectious Diseases, Greifswald-Insel Riems, Germany
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M. H. Groschup. Institute of Novel and Emerging Infectious Diseases at the Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Südufer 10, 17493 Greifswald-Insel Riems, Germany. Tel.: +49 038351 7-0; Fax: +49 038351 7-1219; E-mail: martin.groschup@fli.bund.de

Summary

West Nile virus (WNV) is a flavivirus that is maintained in an enzootic cycle between ornithophilic mosquitoes, mainly of the Culex genus, and certain wild bird species. Other bird species like ravens, jays and raptors are highly susceptible to the infection and may develop deadly encephalitis, while further species of birds are only going through subclinical infection. The objective of this study was to continue in years 2009–2011 the serological and molecular surveillance in wild birds in Germany (see Vector Borne Zoonotic Dis. 10, 639) and to expand these investigations for the first time also to sera from domestic poultry and horses collected between 2005 and 2009. All three cohorts function as indicators for the endemic circulation of WNV. The presence of WNV-specific antibodies was detected in all samples by virus neutralization test (VNT), indirect immunofluorescence test (IFT) and/or enzyme-linked immunosorbent assay (ELISA). The presence of WNV genomes was monitored in relevant sera using two qRT-PCRs that amplify lineage 1 and 2 strains. A total of 364 migratory and resident wild bird serum samples (with emphasis on Passeriformes and Falconiformes) as well as 1119 serum samples from domestic poultry and 1282 sera from horses were analysed. With the exception of one hooded crow, antibody carriers were exclusively found in migratory birds, but not in resident birds/domestic poultry or in local horses. Crows are facultative, short-distance winter migrants in Germany. WNV-specific nucleic acids could not be demonstrated in any of the samples. According to these data, there is no convincing evidence for indigenous WNV infections in equines and in wild/domestic birds in Germany. However, since a few years, WNV infections are endemic in other European countries such as Austria, Hungary, Greece and Italy, a state-of-the-art surveillance system for the detection of incursions of WNV into Germany deems mandatory.

Impacts

  •  The German monitoring study identified WNV antibodies in migratory birds, but not in resident birds, domestic poultry or local horse populations.
  •  The WNV antibody–positive species are birds that migrate to tropical Africa or to southern Europe; WNV-specific RNA could not be found in any of the samples.
  •  There is currently no indication for the presence of active WNV infections in resident and migratory birds and in local horses in Germany.

Introduction

West Nile virus (WNV) is considered the most widespread flavivirus in the world, endemic in Africa, Asia, Europe, Australia and on the American continent (Komar, 2003; Trevejo and Eidson, 2008; Calstri et al., 2010; Weissenböck et al., 2010; Papa et al., 2011). WNV infections have been described in a wide variety of vertebrates, and the virus was found in more than 150 species of wild and domestic birds (van der Meulen et al., 2005). Following infection, many bird species produce levels of viraemia that are sufficient for transmitting the virus to mosquitoes (Komar et al., 2003).

Recently, it was suggested that WNV could be classified into seven distinct lineages (Mackenzie and Williams, 2009), of which the first two lineages constitute the major groups. WNV lineage 1 strains are distributed worldwide and have a high potential of being neuroinvasive. WNV lineage 2 strains were originally found exclusively in sub-Saharan Africa and Madagascar, but emerged in 2004 in Hungary (Bakonyi et al., 2006) and dispersed since then in central and south-eastern Europe. WNV lineage 2 strains were initially considered to be less virulent, compared to lineage 1 strains. This assumption seems to have changed, however, as highly virulent and neuroinvasive lineage 2 WNV strains were detected recently both in South Africa (Venter and Swanepoel, 2010) and in Greece (Papa et al., 2011).

In Europe, a variety of WNV strains of diverse virulence, belonging either to lineage 1 or to lineage 2, have been isolated to date. Until 2004, only lineage 1 strains were circulating, primarily in Mediterranean countries (Calstri et al., 2010). Lineage 2 WNV infections were first observed in 2004 in Hungary in birds of prey (Bakonyi et al., 2006; Erdélyi et al., 2007). This virus strain established itself in the region, dispersed in the following years locally and showed explosive geographic spread in 2008 throughout Hungary and into eastern Austria (Wodak et al., 2011). This virus strain dispersed further and caused a severe outbreak in human beings in 2010 in northern Greece; the increased virulence and neuroinvasiveness of the Greek virus may be contributed to a certain amino acid change (Papa et al., 2011) of the otherwise Hungarian/Austrian virus strain.

The long-distance spread of this lineage 2 WNV strongly points to the ability of the virus to be transmitted outside the traditional routes of viral circulation and to adaptation to local vectors.

Until lately, only few data were available on WNV epidemiology in Germany. In two previous studies, antibodies to WNV were found in several migratory and resident bird species (Linke et al., 2007; Seidowski et al., 2010), but WNV-specific RNA has so far never been detected in any of these studies (Schirrmeier et al., 2004; Hlinak et al., 2006; Linke et al., 2007; Seidowski et al., 2010; Ziegler et al., 2010), indicating no evidence for indigenous WNV infections in the past, even though potential vector mosquitoes are present in Germany (Timmermann and Becker, 2010). In August 2010, Usutu virus (USUV), a closely related virus to WNV, was isolated from a pool of Culex pipiens pipiens mosquitoes, trapped in south-west Germany, yet there was no reported increase in the mortality of wild and captive bird species in 2009 and 2010 (Jöst et al., 2011). However, starting in June 2011, dead blackbirds infected with USUV were frequently found in south-west Germany (Becker et al. 2012).

In the study presented here, we report WNV-monitoring data of free-ranging poultry and of horses in Germany, which were obtained most recently. Moreover, we describe a strategy to discriminate serological cross-reactions of WNV with other members of the Flavivirus group.

Methods

A total of 364 serum samples from migratory and resident birds belonging to forty different bird species collected between 2009 and 2011 were investigated. Bird species belonged to eleven bird orders (see Tables 1 and 2) with emphasis on Passeriformes and Falconiformes. Moreover, 1119 samples from free-ranging poultry (taken between 2005 and 2008) were tested. This sample panel included serum material from 633 geese, 391 ducks and 95 chickens. A total of 1282 sera from German horses (collected 2007–2009) were also tested. The geographic localization of the sampling sites in different federal states is listed in Tables 3 and 4. Most wild birds were captured in the larger Berlin/Brandenburg suburban area and submitted for a variety of reasons to the Small Animal Clinic, Veterinary department, Berlin University. The region of Berlin and Brandenburg is part of the Central European lowland area and characterized by a mixture of forests, agriculture areas and large diversity of wetlands, rivers and lakes, but coast and mountains are absent with moderate climate.

Table 1.   Total number of wild bird orders sampled in Germany from 2009 to 2011
Order (-formes)Year 2009Year 2010Year 2011Total
Anseriformes3027259
Passeriformes2837166
Falconiformes77511129
Strigiformes410115
Ciconiformes11415
Gruiformes57113
Columbiformes281442
Apodiformes123
Charadriiformes59115
Piciformes156
Cuculiformes11
Total1911667364
Table 2.   WNV and USUV neutralization assay results for wild bird samples (detailed by species). Positive samples are highlighted in bold, neutralisation titres in brackets
Bird’s orderBird’s nameScientific nameSamples testedWNV pos. (ND50)USUV pos. (ND50)
  1. aNot done because insufficient serum volume.

AnseriformesMute swan Cygnus olor 49 1 (10)0 (<10)
Whooper Swan Cygnus cygnus 100
Mallard Anas platyrhyn. 8 1 (40)0 (<10)
Canada Goose Branta canadensis 100
PasseriformesHooded Crow Corvus cornix 46 1 (30)0 (<10)
Rook Corvus frugilegus 100
Common Raven Corvus corax 500
Eurasian Blackbird Turdus merula 300
Common Magpie Pica pica 500
Eurasian Jay Garrulus glandarius 500
Common Starling Sturnus vulgaris 100
FalconiformesCommon Buzzard Buteo buteo 37 1 (20)0 (10)
Northern Goshawk Accipiter gentilis 1100
European Kestrel Falco tinnunculus 18 1 (40)0 (<10)
White-tailed Sea-eagle Haliaeetus albicilla 3700
Eurasian Sparrowhawk Accipiter nisus 1800
Osprey Pandion haliaetus 2 1 (320)0 (a)
Western Marsh-harrier Circus aeruginosus 2 1 (40)0 (15)
Lesser Spotted Eagle Aquila pomarina 100
Europ. Honey Buzzard Pernis apivorus 3 1 (60)0 (<10)
Strigi formesTawny Owl Strix aluco 600
Long-eared Owl Asio otus 600
Eurasian Eagle-owl Bubo bubo 200
Barn Owl Tyto alba 100
CiconiformesWhite Stork Ciconia ciconia 4 2 (10+80)0 (<10)
Black Stork Ciconia nigra 100
Grey Heron Ardea cinerea 900
Great Egret Casmerodius albus 100
Grui formesCommon Coot Fulica atra 11 6 (10–40)0 (<10–15)
Common Crane Grus grus 200
ColumbiformesCommon Wood Pigeon Columba palumbus 3300
Rock Pigeon Columba livia 900
ApodiformesCommon Swift Apus apus 300
CharadriiformesBlack-headed Gull Larus ridibundus 3 1 (10)0 (<10)
Europ. Herring Gull Larus argentatus 500
Eurasian Woodcock Scolopax rusticola 700
PiciformesGreen Woodpecker Picus viridis 100
Great Spotted Woodpecker Dendrocopos major 400
Black Woodpecker Dryocopus martius 100
CuculiformesCommon Cuckoo Cuculus canorus 100
Total    364 17 0
Table 3.   Origin of blood samples (serum, plasma) from free-ranging poultry collected in years 2005 to 2008
Federal state of GermanyYear 2005Year 2006Year 2007Year 2008Total
Brandenburg5521940200514
Mecklenburg-Western Pomerania100100200
Hesse2020
Saxony-Anhalt4040
Thuringia185185
North Rhine-Westphalia50110160
Total552194354101119
Table 4.   Origin of horse samples from Germany from 2007 to 2009
Federal state of GermanyYear 2007Year 2008Year 2009Total
Lower Saxony633194
Berlin12811180319
Mecklenburg-Western Pomerania21820
Hesse9164100255
Saxony708351204
Thuringia244569
North Rhine-Westphalia8080160
Baden-Württemberg8080
Bavaria8181
Total3155274401282

All serum samples were tested for antibodies to WNV strain NY99 by virus neutralization test (VNT) as described previously (Seidowski et al., 2010). To reveal cross-reactions with other members of the Japanese encephalitis virus serogroup, all wild bird samples were assayed in parallel for USUV-neutralizing antibodies (using the same experimental conditions but USUV strain Vienna 2001). Where applicable, sera were probed also for neutralizing antibodies to tick-borne encephalitis virus (TBEV) using strain Neudoerfl (kindly provided by F. Hufert, Institute for Virology, Göttingen).

Free-ranging poultry samples were also probed by indirect immunofluorescence test (IFT) as described previously (Seidowski et al., 2010; Ziegler et al., 2010).

The majority of the 1200 horse samples was additionally investigated by a commercially available competition ELISA, which is suitable for measurement of WNV IgG antibodies in serum of horses, following the manufacturer’s instructions (ID Screen© West Nile Competition, IDVet, Montpellier, France). Positive IgG samples were also tested by IgM-ELISA (West Nile Virus IgM Antibody Test from IDEXX).

Blood samples drawn from wild birds in 2009 and 2010 (357 samples) were also screened by quantitative real-time RT-PCR (RT-qPCR) to detect WNV NS1 and/or E protein–specific nucleotide sequences (Eiden et al., 2010) as well as USUV (Jöst et al., 2011) sequences. For these assays, RNA was extracted from whole blood or cruor by using a QIAamp Viral RNA kit or an RNeasy Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions.

Results

Serum samples from seventeen of 364 wild birds were positive for WNV-neutralizing antibodies (see Table 2). This includes five of 129 tested birds of prey, while only one of 66 Passeriformes carried WNV-specific antibodies. Six of the eleven common coots (order Gruiformes) were also positive.

VNT titres ranged from 10 to 320 (Table 2). To reveal cross-reactions with other Flaviviruses, wild bird sera were also tested in an USUV VNT. All 347 WNV antibody–negative wild birds were also negative for USUV-neutralizing antibodies. Of the seventeen WNV antibody–positive wild birds, nine were clearly negative for USUV antibodies and another seven had WNV titres that were on average 2-fold higher than the corresponding USUV titres, indicating WNV infections as primary causes. Because of insufficient serum volume, one WNV-positive Osprey sample was excluded from this comparative study.

No WNV-specific or USUV-specific RNA was detected by RT-qPCR in the wild bird blood samples collected in 2009 and 2010 (357 samples). Sixteen of them had WNV-neutralizing antibodies in the VNT. All 1119 poultry samples originating from animals in free-ranging areas were negative for WNV-specific antibodies by VNT and IFT.

Similar results were found in the horse population. With the exception of four horses, all of the 1282 tested sera were negative for WNV-specific antibodies using VNT. WNV-specific antibody titres in the four horses ranged from 15 to 320 (see Fig. 1). Further investigations revealed that three of the four horses had been vaccinated with a killed WNV vaccine for horses [Duvaxyn© (Pfizer)] in the past while one animal was imported from Hungary.

Figure 1.

 Discrimination of WNV and TBEV antibodies in equine sera (No 1–8), which were breaking ranks in a WNV ELISA (results depicted in Fig. b). Sera were further investigated by WNV strain NY99 (a) and TBEV strain Neudoerfl (c) virus neutralization tests. Results illustrate that horses No 1–4 had elicited WNV-specific antibodies, whereas horses No 5–8 carried TBEV-neutralizing antibodies.

By using a commercial ELISA for horse sera, these results could be confirmed in general. However, another four sera were reacting in the ELISA, albeit being negative in the WNV-specific VNT (Fig. 1). These four reactive samples could be linked to cross-reacting antibodies within the Flavivirus group, because they showed neutralizing antibodies to TBEV strain Neudoerfl in the VNT. TBEV titres in these sera ranged between 20 and 480. None of the eight IgG ELISA–positive horses were positive for IgM (West Nile Virus IgM Antibody Test from IDEXX).

Discussion

In endemic areas, wild birds are the main reservoirs for WNV and other related Flaviviruses. Migratory birds are considered to act as carry-over vectors and may be one different mode of introductions of virus into new regions and countries (Brault, 2009.)

There is no evidence for the circulation of WNV in Germany. Several WNV surveillance studies were conducted in the past (Schirrmeier et al., 2004; Hlinak et al., 2006; Ziegler et al., 2010) without demonstrating WNV-specific RNA in wild birds. WNV antibodies were shown only in migratory bird species (Linke et al., 2007; Seidowski et al., 2010), which had overwintering grounds in Africa and/or southern Europe. These birds had generally very low antibody titres, which resulted most likely from infections in their overwintering quarters.

No WNV-infected mosquitoes were detected in Germany to date. Timmermann and Becker (2010) tested 11 073 host-seeking adult female mosquitoes (of 13 species) collected in the Upper Rhine Valley in 2007 and 2008 for WNV. However, USUV was detected in Culex pipiens pipiens mosquitoes in south-west Germany most recently (Jöst et al., 2011). Anymore since June 2011, dead blackbirds infected with USUV were frequently found around the cities of Mannheim and Heidelberg in south-west Germany (ProMED-mail; http://www.promedmail.org/Archive Number: 20110916.2827). New data suggest that after the initial isolation of USUV from German mosquitoes collected in 2010, the virus emerged in 2011 and caused mass mortality of wild and captive birds in south-west Germany (J. Schmidt-Chanasit, personal communication, Bernhard Nocht Institute for Tropical Medicine, Hamburg).

No evidence for an epidemic circulation of WNV in Germany has been obtained to date. The antibody levels and distributions found in animals in the here-presented study compared well to those observed in previous studies. Only 17 of 364 samples (=4.6%) of wild birds tested positive for WNV-specific antibodies. In six of these seventeen WNV antibody–positive samples, USUV cross-reactive antibodies were found, that is, USUV titres were significantly lower than the corresponding WNV neutralization titres. No USUV sequences were detected in the German wild bird samples.

These results are in accordance with the results from other European countries where WNV antibodies were found but no WNV antigens or nucleic acid sequences. In Poland, WNV-specific antibodies were detected in wild birds and USUV antibodies in a black-headed gull (Larus ridibundus) (Hubálek et al., 2008). In the Netherlands, low WNV antibody titres were demonstrated in a stochastic model for migratory birds (Havelaar et al., 2010), but WNV-specific RNA was not detected.

In WNV epidemic areas, a high number of wild migratory and resident birds tested positive for WNV-specific antibodies and for WNV-specific RNA. During the WNV surveillance programme in Italy in 2009 (Monaco et al., 2011), resident birds, such as European magpies (Pica pica), Eurasian jays (Garrulus glandarius) and carrion crows (Corvus corone), carried WNV-specific RNA and such RNA was also detected in several mosquito species (e.g. Culex pipiens and Ochlerotatus caspius). In Italy, overwintering of WNV in the mosquitoes or – less likely – the virus persistence in the hosts may apply.

Wodak et al. (2011) described WNV lineage 2 outbreaks in birds of prey in the eastern part of Austria in 2008 and 2009 and identified virus-neutralizing antibodies in 38.7 per cent of the investigated sera of various wild birds. Moreover, a variety of wild birds tested positive for WNV-specific antibodies in Spain (Figuerola et al., 2008), and lineage 1 WNV was isolated from two dead golden eagles (Aquila chrysaetos) (Jiménez-Clavero et al., 2007; Höfle et al., 2008).

Free-ranging poultry is a good and useful sentinel system for WNV activity in many countries. This can be found in different published studies (Komar, 2001; Rizzoli et al., 2007; Chevalier et al., 2011). However, not all species are equally suitable because of differences in their WNV susceptibility.

In the past, equines were used as indicators to monitor the circulation of WNV in the environment (Leblond et al., 2007; Jiménez-Clavero et al., 2010; Chevalier et al., 2011). However, such sentinels work only in areas without vaccination, because infection antibodies cannot be discriminated from vaccination-derived antibodies.

In the here-studied sample panel from Germany, all sera tested negative for WNV-specific antibodies with the exception of four samples (one imported and three vaccinated horses). False-positive ELISA results, however, can also result from cross-reacting antibodies raised to other Flaviviruses. A clear discrimination is only possible by VNT. For example, we observed four false-positive ELISA results, caused by previous TBEV infections (i.e. WNV VNT results were negative, while TBEV VNT was clearly positive).

The number of TBEV infections in humans has increased more than 3-fold in Europe since 1983 (Süss, 2008). TBEV infections in horses are usually asymptomatic, but rare exceptions occur. Tick-borne encephalitis was described in 1981 in a horse in Switzerland (Waldvogel et al., 1981) for the first time. The animal showed central nervous symptoms. TBEV was demonstrated in the brain, and correlating antibodies were detected in the serum. Horses from TBEV endemic regions in former Yugoslavia tested also positive for TBEV antibodies without having shown clinical symptoms before (Waldvogel et al., 1981). In Germany, TBEV-related diseases are reported sporadically in horses, and only low levels of TBEV-specific antibodies are found (C. Klaus, personal communication, National reference laboratory for tick-borne diseases, Jena). Consequently, TBEV should be considered as differential diagnosis in horses, although this infection seems to occur in horses at a low frequency only.

Taken together, testing sentinel or wild birds in risk areas (mosquito-rich wetlands) as well as conducting monitoring programmes for horses is a useful approach to monitor the human WNV infection risk in these areas. Because of trade and climate change effects, the risk of WNV introduction to Germany is increasing steadily. Also, WNV is already endemic in neighbouring Austria and might disperse to Germany at any time. Therefore, a state-of-the-art monitoring system for WNV deems mandatory. Based on already obtained data, the presence of a larger WNV epidemic can be excluded for Germany, but not the presence of silent endemic cycles in restricted areas.

Acknowledgements

The authors would like to thank Ines Nedow and Birke Kalb for their assistance with this study. We also thank the equine clinics at the departments of Veterinary Medicine in Berlin, Gießen, Hannover, Leipzig and München for the horse samples and the veterinary authorities in Brandenburg, Hesse, Mecklenburg-Western Pomerania, North Rhine-Westphalia, Saxony-Anhalt and Thuringia for the samples of the domestic poultry. This project was supported by the German Federal Ministry of Education and Research (network project ‘Emerging Arthropod-Borne Viral Infections in Germany: Pathogenesis, Diagnostics and Surveillance’).

Conflicts of interest

The authors have declared no potential conflicts.

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