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ABSTRACT |
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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INTRODUCTION |
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Millions of birds migrate every year between Europe and Africa, wintering in or passing through WNV-endemic areas. Bird migration seems to play a major role in the dissemination of WNV and USUV.6,7 In Europe, antibodies to WNV have been detected in several bird species.8,9 In the United Kingdom, investigations on the prevalence and incidence of WNV infections were carried out in 2001–2002 and showed that migrating and resident birds had antibodies to WNV or viral genomes.10 In France, virus isolation was successful in various samples from mammals such as horses.3 There are no data on endogenous WNV and USUV infections in Germany, whereas USUV infections have been diagnosed in neighboring Austria.2 The aim of the present study was to investigate migrating and resident birds as a potential reservoir of WNV and USUV by using serologic and molecular methods, and to assess the possibility that migrating species import these viruses from their wintering areas in Africa.
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MATERIALS AND METHODS |
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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), were collected in the context of bird-ringing activities or other research interests. The birds were captured by traps and mist nets, bled, and released or downed in pest-control programs. Sampling of white stork nestlings and raptor nestlings was performed by bleeding nestlings at nest sites. Morphologic examinations were performed to determine sex, age, and species. Rings engraved with a serial number from German ringing centers were used for individual marking. Several samples were collected from injured wild birds during veterinary care. National laws and regulations regarding protection, conservation, and animal welfare were respected.
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Diagnostic assays. Two methods were used for serologic diagnosis: an indirect immunofluorescence assay (IFA) and a neutralization test (NT). The first screening was done using a WNV IFA kit (EUROIMMUN, Lübeck, Germany) following the manufacturer’s instructions. The samples were diluted 1: 10 with sampling buffer and 25 µL of the diluted samples were applied to a biochip slide and incubated for 30 minutes at room temperature. After the slide was washed for 10 minutes, a non-labeled rabbit anti-dove IgG hyperimmune serum diluted 1:300 was added to the biochips and incubated for 30 minutes. The rabbit anti-dove IgG reacted with a broad spectrum of antibodies derived from different bird species (Grund C, unpublished data). After a second washing step, 25 µL of a mixture of a goat anti-bird fluorescein isothiocyanate (FITC)–labeled antibody (Bethyl Inc., Montgomery, AL) diluted 1:50 in phosphate-buffered saline (PBS) plus Tween 20 and an anti-rabbit FITC-labeled antibody (Dianova, Hamburg, Germany) for detection of the rabbit anti-dove IgG hyper immune serum diluted 1:300 were applied to the bio-chips. After a second incubation for 30 minutes and a washing step, the slide was embedded with a drop of glycerol/PBS according to the manufacturer’s instructions and analyzed under a fluorescence microscope (Axioskop MC80, filters 495 nm and 528 nm; Zeiss, Oberkochen, Germany). Reactive samples were titrated in dilution steps of 1:10, 1:100, and 1: 1000 according to the manufacturer’s instructions. Reference goose serum with a known titer of antibodies to WNV was used as a control sample.
The method of choice for flavivirus diagnostics and differentiation is the virus NT. A WNV-specific NT was performed in a 96-well plate format (Nunc, Wiesbaden, Germany). Vero cells were grown in Eagle’s minimal essential medium (MEM) (PAA Laboratories, Coelbe, Germany) supplemented with 10% fetal bovine serum (PAA Laboratories) in 96-well microtiter plates. Samples were diluted 1:10 in PBS. The stock solution of WNV strain Israel was diluted in Eagle’s MEM with 1% ciprofloxacin (PAA Laboratories) to a constant concentration of 500 50% tissue culture infective doses (TCID50)/mL. A total of 25 µL of virus suspension were mixed with 25 µL of serum or plasma dilution and incubated for 1 hour at 37°C. After incubation, the inoculum of serum and virus was added to the cells and incubated for 3 days at 37° C in an atmosphere of 5% CO2. The suspension of plasma and virus was then removed from the cells and 100 µL of fresh medium were added to each well and incubated for 3 days at 37° C in an atmosphere of 5% CO2. After incubation, the cells were fixed and stained with naphthalene black and analyzed under a light microscope. Neutralizing goose sera were included in all experiments. Each test sample was investigated in duplicate with a single well of serum-cell control without virus. Samples with neutralizing antibodies at a dilution of 1:10 were titrated (two-fold serial dilutions from 1:10 to 1:2,560) to determine end point titers for WNV. Antibody titers were determined as the highest dilution of serum or plasma at which 50% of the wells did not show a cytopathic effect. Samples with a WNV antibody titer 
10 were considered positive because serum dilutions < 1:10 were often toxic for cells.
Some samples were further investigated in a USUV-specific plaque reduction neutralization test (PRNT). The test procedure was performed according to published studies13,14 with slight modifications. The test was performed in 48-well microtiter plates (Nunc) with Vero cells cultivated in Eagle’s MEM (PAA Laboratories) with 10% fetal bovine serum (PAA Laboratories) and 1% ciprofloxacin (PAA Laboratories). The serum or plasma samples were diluted two-fold from 1:10 to 1:640 in PBS. Aliquots (50 µL) of USUV strain Vienna 2001 from Austria containing 150 TCID50/mL were added to 50 µL of the sample and incubated for 1 hour at 37°C. The virus-sample suspension was applied to the cells and incubated for 1 hour at 37°C. After incubation, the mixture of plasma and virus was removed from the cell layer and fresh medium was added. An overlay of carboxymethyl cellulose medium (BDH Ltd., Poole, United Kingdom) was added to the cells and the cells were incubated at 37°C in an atmosphere of 5% CO2 for 3 days. The plates were fixed and stained with naphthalene black. Plaques were counted and the 50% PRNT (PRNT50) titer was calculated according to Reed and Münch.15
Viral RNA was isolated from 552 samples from white storks; 100 µL of plasma was centrifuged for 1.5 hours at 14,000 rpm. The pellet was used for virus extraction and the supernatant was used for serologic testing. RNA extraction was performed using the Qiagen RNeasy Mini Kit (Qiagen, Hilden, Germany). RNA was eluted in 60 µL of RNase-free water (Fluka Chemikalien GmbH, Buchs, Switzerland) supplemented with RNA (100 ng/µL) (Roche, Mannheim, Germany). RNA was stored at –70°C before further use.
cDNA was synthesized by reverse transcription of 11.6 µL of extracted RNA in a 20-µL reaction volume. For transcription, 1 µL of the specific reverse primer (10 µM; Table 2) and 1 µL of reverse transcriptase (200 U/µL) (Invitrogen, Karlsruhe, Germany) were used. The cDNA was stored at –20°C before further use.
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). As an internal control standard of the RNA extraction, of cDNA synthesis, and PCR, plasma was spiked with a constant volume of heat-inactivated supernatant of Squirrel monkey retrovirus (SMRV), family Retroviridae. A specific RT-PCR using the SMRV envelope protein sequence was established (Table 2). |
RESULTS |
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Antibodies against WNV were detected using IFA in nine species (Table 3). Washed samples were analyzed by a WNV NT, neutralizing antibodies to WNV were detected in five species (Table 3). Fifty-nine blood samples from birds had antibodies to WNV by IFA, 27 of which were also positive in the NT. Using the NT, 24 additional sera were identified that were not reactive by IFA.
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). On the basis of the IFA results, the percentage of samples with antibodies to WNV was low. Thirty-three (5.8%) of 569 Ciconia ciconia samples were reactive by WNV IFA, and only 13 (2.3%) samples were positive by WNV NT. Three sera from Cygnus olor showed neutralizing activity, but these sera showed negative results in the WNV IFA. Sixteen (11.4%) of 140 sera of Pandion haliaetus were reactive in the WNV IFA. In neutralization assays, the number of positive sera was higher (n = 28, 20%). Only one serum each of samples from Haliaeetus albicilla (n = 36), Milvus migrans (n = 1), and Accipiter gentilis (n = 19) had neutralizing antibodies to WNV.
To gain insight into the potential cross reactivity of bird-infecting flaviviruses, 25 samples reactive against WNV in the IFA but negative in the NT were investigated in the USUV NT. Three samples were positive and had PRNT50 titers of 17, 21, and 37, respectively. The antibody titers against WNV varied in different serologic test systems and among the individuals of the species. Tables 4 and 5 show that antibody titers determined using IFA and NT were generally lower in nestlings than in adults (titers of adult white storks were 3-fold to 100-fold higher than in nestlings). In ospreys, only one nestling bird of 106 samples tested was reactive in WNV IFA but not in WNV NT; all other reactive samples were from adult birds.
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Regarding the importation of WNV from Africa to Germany, bird species with WNV-specific antibodies were correlated with their migration status. Five species, most of which were migratory birds, had antibodies against WNV. While C. ciconia, M. migrans, and P. haliaetus migrate to tropical Africa, birds of the species Cygnus olor show a more complex migration behavior. Accipiter gentilis is a resident bird species, whereas birds of the species Cygnus olor are partial migrants.
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DISCUSSION |
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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). We identified 5 of 87 species with neutralizing antibodies, 3 of which can be correlated with wetland habitats. In most of the bird species, the percentage with neutralizing antibodies was low, with the exception of ospreys. We detected specific antibodies in adults and in nestlings. The antibody titer of the nestlings most likely reflects the maternal antibody status because it is well known that maternal antibodies to WNV are transmitted from the mother during egg production.17,18 We investigated nestlings of ospreys and white storks at 2–9 weeks of age, but only one osprey had antibodies to WNV by the IFA. In contrast, we detected neutralizing antibodies in adult birds living in the same region. However, we found a higher percentage of WNV-specific antibodies in nestlings of white storks. These results either reflect differences in the transmission of WNV-specific antibodies via egg yolk or differences in the half-life of maternal antibodies in the offspring. There is limited information on the persistence of maternal antibodies against WNV in birds, which complicates interpretation of serologic results.18 Viral genomes were not detected in nestling nor adult white storks by PCR, which implies that the antibody response observed in the birds seems to be an indirect marker of the serologic status of the parent birds.
The serologic data obtained by IFA and NT were not congruent in all cases. It is well known that there is high cross-reactivity of closely related flaviviruses in antigen detection systems such as the IFA and an enzyme-linked immunosorbent assay.19 The method of choice for detecting antibodies against flaviviruses is the NT, but cross-reacting antibodies have also been observed in this assay. High titers of neutralizing antibodies may represent cross-reacting antibodies from flaviviruses, especially of the Japanese encephalitis serocomplex. Differences in antibody titers against different flavivirus isolates can give further information on the virus responsible for the reduction of the immune response. The NT procedure used is very stringent and used a 100% knockout of virus infectivity for antibody detection. Under these highly demanding conditions, it could be assumed that antibodies to WNV were acquired through exposure to WNV.
Three samples of C. ciconia, Phoenicurus phoenicurus, and P. haliaetus showed positive results in the WNV IFA and had low-level neutralizing antibody titers to USUV. Findings on the emergence of USUV in Austria, which was probably introduced by migrating birds, showed that in the summer of 2001 several bird species were infected with USUV, and the epidemic, especially in blackbirds, was stable for more than one summer in a central European region.2
Little is known about WNV infections in wild birds in central Europe. There are limited data on the WNV seroprevalence in passerine birds in Poland, indicating that it is at a low level.8 A study of WNV infections in Austria did not detect WNV in dead wild birds, and the investigators concluded that WNV has no measurable impact in Austria.20 However, in the United Kingdom serologic evidence for WNV and USUV was reported in wild resident and migrating birds, as well as in sentinel chickens.21 Detection of the WNV genome was demonstrated in crows.10 In our study, none of 41 Corvidae samples showed evidence of an antibody response to WNV.
Infections with WNV in migratory birds in Europe can occur either through exposure in Europe, on migratory routes, or at wintering sites in Africa. The sporadic outbreaks of WNV infections in areas in southern Europe such as Romania, Tuscany in Italy, and Camargue in France do not support the assumption that WNV is endemic in these regions but may indicate that WNV is occasionally imported to these areas. In tracking the migratory route of birds from Europe to Africa, it is noteworthy that there are two major destinations, one to western regions of Africa and the other to eastern parts of Africa. On the route via southeastern Europe it was observed that WNV epidemics occur in countries such as Romania and Israel.22–24 Many bird species rest in Israel before or after the flight across the desert during autumn and spring migration.6 Sporadic introduction of WNV by migrating birds to central Europe may therefore be possible.
However, there are limited data on the viremic phase in WNV-infected birds. In experimental infections of different bird species, viremia in birds was high for approximately four days to enable infection of mosquitoes.25 It is doubtful whether this period is long enough for direct import of the virus from disease-endemic areas in Africa to central Europe. However, infections with USUV in Austria support the assumption that flavivirus infections might be imported and maintained in temperate regions.2
It remains unclear why there is a low level percentage of antibodies to WNV in European migrating birds without clear evidence of WNV-diseased birds. In contrast to birds in the United States showing clinical symptoms, European birds have long been exposed to WNV on their migration routes to and from Africa. Over centuries, this exposure might have induced a natural resistance to WNV infections in European birds, whereas in America WNV was introduced into a highly susceptible bird population that had never been exposed to this virus.
There is evidence of only a few human WNV infections imported from the United States into Germany. Because of climate warming, it must be assumed that further WNV infections limited in time and region might appear, similar to the current situation in southeastern Europe.
Received January 8, 2007. Accepted for publication May 3, 2007.
Acknowledgments: We thank I. Nehlmeier and A. Teichmann for excellent technical support; U. Erikli for careful copyediting; G. Wengler (Justus-Liebig-Universität Gießen, Germany) for providing WNV isolate B956; H. Zeller (Institut Pasteur, Paris, France) for Kunjin virus; H. Bin (Sheba Medical Center, Tel Hashomer, Israel) for WNV isolate Israel; T. R. Kreil (Baxter GmbH, Vienna, Austria) for providing WNV isolate New York; M. Pfeffer (Sanitätsakademie der Bundeswehr, Munich, Germany) for providing Usutu virus, strain Vienna; E. Firenzi (Berencsi György National Center for Epidemiology, Budapest, Hungary), C. Banet-Noach (Kimron Veterinary Institute, Beit Dagan, Israel), and M. A. Drebot (Viral Zoonoses, National Microbiology Laboratory, Winnipeg, Manitoba, Canada) for providing WNV-positive bird samples, especially sera from geese; H. Will (Heinrich-Pette-Institute, Hamburg, Germany) for providing blood samples; C. Grund (Faculty of Veterinary Medicine, Ludwig-Maximilians-University Munich, Germany) for supplying hyperimmune serum; K. Sonnenberg (EUROIMMUN AG, Lübeck, Germany) for supplying IFA kits for serologic analysis; R. Altenkamp for capturing adult goshawks; H. Eggers, P. Gottschalk, U. Hilfers, M. Hug, M. M. and M. Kaatz, D. Kasper, S. Martens, B. Metzger, R. Neumann, U. Querner, J. von Rönn, F. Schulz, T. Suckow, I. Todte, H. Trapp, and B. Wuntke for assistance in capturing birds and supporting the study; N. Hagen and C. Schmitt for invaluable assistance in blood sampling; and M. Müller and K. Hattermann for helpful discussions.
Financial support: This study was supported by the German Ministry of Health grant BMGS 115-1720-1/31. The work conducted by Daniel Schmidt was supported by the Deutsche Ornithologen-Gesellschaft DO-G.
* Address correspondence to Georg Pauli, Robert Koch-Institut, Nordufer 20, 13353 Berlin, Germany. E-mail: paulig{at}rki.de 
Authors’ addresses: Sonja Linke, Robert Koch-Institut, Nordufer 20, 13353 Berlin, Germany, Telephone: 49-30-1875-42244, Fax: 49-30-1875-42605, E-mail: linkes{at}rki.de. Matthias Niedrig, Robert Koch-Institut, Nordufer 20, 13353 Berlin, Germany, Telephone: 49-30-1875-42370, Fax: 049-30-1875-42625, E-mail: niedrigm{at}rki.de. Andreas Kaiser, Department V (Ecology), Institute for Zoology, University of Mainz, J.-J.-Becher-Weg 13, 55128 Mainz, Germany, Telephone: 49-6131-392-3856, Fax: 49-6131-392-3731, E-mail: dr.andreas.kaiser{at}t-online.de. Heinz Ellerbrok, Robert Koch-Institut, Nordufer 20, 13353 Berlin, Germany, Telephone: 49-30-1875-42258, Fax: 49-30-1875-42605, E-mail: ellerbrokh{at}rki.de. Kerstin Müller, Department of Veterinary Medicine Small Animal Clinic (WE20), Freie Universität Berlin, Oertzenweg 19b, 14163 Berlin, Germany, Telephone: 49-30-8386-2422, Fax: 49-30-8386-2521, E-mail: MuellerKerstin{at}gmx.de. Thomas Müller, Friedrich-Loeffler-Institute, Federal Research Institute for Animal Health, Institute for Epidemiology, Seestraße 55, 16868 Wusterhausen, Germany, Telephone: 49-33979-80186, Fax: 49-33979-80200, E-mail: thomas.mueller{at}fli.bund.de. Franz Josef Conraths, Friedrich-Loeffler-Institute, Federal Research Institute for Animal Health, Institute for Epidemiology, Seestraße 55, 16868 Wusterhausen, Germany, Telephone: 49-33979-80176, Fax: 49-33979-80200, E-mail: franz.conraths{at}fli.bund.de. Ralf-Udo Mühle, Ökologische Station Gülpe, Universität Potsdam, 14715 Gülpe, Germany, Telephone: 49-33875-30621, Fax: 49-33875-30752, E-mail: muehle{at}rz.uni-potsdam.de. Daniel Schmidt, NABU-Vogelschutzzentrum Mössingen, Ziegelhütte 21, 72116 Mössingen, Germany, Telephone: 49-7473-1022, Fax: 49-7473-21181, E-mail: schmidt{at}NABU-Vogelschutzzentrum.de. Ulrich Köppen, Landesamt für Umwelt, Naturschutz und Geologie Mecklenburg-Vorpommern, Beringungszentrale, Badenstraße 18, 18439 Stralsund, Germany, Telephone: 49-3831-696243, Fax: 49-3831-696249, E-mail: ulrich.koeppen{at}lung.mv-regierung.de. Franz Bairlein, Institute of Avian Research, Vogelwarte Helgoland, An der Vogelwarte 21, 26386 Wilhelmshaven, Germany, Telephone: 49-4421-96890, Fax: 49-4421-968955, E-mail: franz.bairlein{at}ifv.terramare.de. Peter Berthold, Max Planck Institute for Ornithology, Vogelwarte Radolfzell, Schlossallee 2, 78315 Radolfzell, Germany, Telephone: 49-7732-15010, Fax: 49-7732-150169, E-mail: berthold{at}orn.mpg.de. Georg Pauli, Robert Koch-Institut, Nordufer 20, 13353 Berlin, Germany, Telephone: 49-30-1875-42310, Fax: 49-30-1875-42605, E-mail: paulig{at}rki.de.
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TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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