West Nile Virus in Wild Birds

Overview and Taxonomy of West Nile Virus in Wild Birds

West Nile virus (WNV) is a single-stranded, positive-sense RNA virus belonging to the genus Orthoflavivirus within the family Flaviviridae, a taxonomic group that includes several other globally significant arthropod-borne pathogens such as Japanese encephalitis virus (JEV), St. Louis encephalitis virus (SLEV), Usutu virus (USUV), and dengue virus [6, 20, 37]. The virus is classified within the Japanese encephalitis serocomplex, a cluster of antigenically related flaviviruses that share ecological and structural similarities, particularly in their reliance on avian-mosquito transmission cycles [6, 21]. WNV was first isolated in 1937 from a febrile patient in the West Nile district of Uganda, and for decades it was considered a relatively obscure pathogen causing mild febrile illness in parts of Africa, the Middle East, and southern Europe [14, 37]. However, its emergence in the Western Hemisphere in 1999, when a lineage 1 strain was detected in New York City, dramatically altered its global status, transforming WNV into the most geographically widespread arbovirus affecting humans, horses, and a vast diversity of avian species [14, 21, 38]. The World Health Organization (WHO), the World Organisation for Animal Health (WOAH), and the Centers for Disease Control and Prevention (CDC) now recognize WNV as a major zoonotic threat with expanding geographic range, driven by climate change, land-use alterations, and the movements of its primary reservoir hosts: wild birds [1, 14, 18, 31].

The enzootic cycle of WNV is fundamentally dependent on wild birds, which serve as the principal amplifying hosts and reservoirs for the virus [3, 4, 14, 21, 38]. In this sylvatic cycle, ornithophilic mosquitoes, primarily of the Culex genus, particularly Culex pipiens, Culex quinquefasciatus, and Culex perexiguus, acquire the virus by feeding on viremic birds [5, 11, 23, 28]. Following an extrinsic incubation period within the mosquito, the virus is transmitted to subsequent avian hosts during subsequent blood meals, thereby perpetuating the transmission cycle [21, 37]. Birds are uniquely suited as reservoirs because many species develop high-titer viremias sufficient to infect feeding mosquitoes, often without exhibiting overt clinical signs [21, 38]. This asymptomatic carriage allows for silent amplification and widespread dissemination of the virus across vast geographic distances, particularly through migratory bird movements [3, 12, 27, 38]. The role of birds as the primary reservoir is so central that the WHO and WOAH emphasize avian surveillance as a cornerstone of early warning systems for WNV emergence in naïve regions [1, 32]. The virus is considered a naturally occurring focal disease, with its persistence in an area determined by the continuous interaction between competent avian hosts, competent mosquito vectors, and favorable environmental conditions [2, 8, 18, 30].

From a taxonomic perspective, WNV exhibits considerable genetic diversity, with nine distinct lineages described to date, though lineages 1 and 2 are responsible for the vast majority of epizootic and human disease globally [17, 37]. Lineage 1 is further subdivided into two major clades: lineage 1a, which includes strains from Europe, Africa, the Middle East, and the Americas, and lineage 1b (Kunjin virus), which circulates predominantly in Australasia [27, 33, 37]. The introduction of lineage 1 into North America in 1999, a strain belonging to the WN99 genotype, led to a catastrophic epizootic among wild birds, particularly corvids, and resulted in the rapid continental spread of the virus [13, 21]. In Europe, lineage 1 strains (particularly the Western Mediterranean-1 sub-cluster, WMed-1) have been circulating in the Mediterranean basin since the 1990s, causing recurrent outbreaks in horses and wild birds in countries such as Spain, Portugal, Italy, France, and Morocco [9, 27, 33, 39]. Phylogenetic analyses of lineage 1 strains from southern Spain have revealed a remarkable persistence and evolution of the WMed-1 sub-cluster, with evidence of long-term local circulation, co-circulation of multiple phylogenetic groups within the same province, and transcontinental spread between Europe and Africa, likely mediated by migratory birds [33]. The 2024 isolation of a lineage 1a strain from a Northern Goshawk (Astur gentilis) in Portugal further underscores the ongoing circulation of this lineage in the Iberian Peninsula and its connection to West African strains via Afro-Palearctic migratory flyways [27].

Lineage 2 of WNV, historically considered a less virulent African lineage, emerged as a major pathogen in Europe following its introduction into Hungary in 2004 [17, 24, 29, 36]. Since then, lineage 2 has spread explosively across the continent, becoming the dominant lineage in many European countries and causing large-scale outbreaks in humans, horses, and wild birds [8, 15, 17, 24, 29, 38]. Phylogenetic studies have demonstrated that the European lineage 2 strains belong to a single monophyletic clade that originated in Hungary and subsequently radiated into neighboring regions, including Austria, the Czech Republic, Slovakia, Serbia, Greece, Italy, Germany, and Belgium [17, 24, 29, 36]. Within this clade, multiple subclusters have been identified; for instance, subcluster 2.5.3.4.3c has become the dominant circulating variant in Germany, accounting for approximately 73% of sequenced samples from 2023–2024, while a second distinct variant within cluster 2.5.3.2 also co-circulates, indicating ongoing viral evolution and genetic diversification [17]. In Greece, lineage 2 strains isolated from wild birds during the 2010 outbreak, including a strain from a Eurasian Magpie (Pica pica), showed >99% sequence similarity to strains from Austria and Hungary, confirming the rapid southward expansion of this lineage [22, 36]. Similarly, in Serbia, phylogenetic analysis of WNV RNA-positive birds (including Northern Goshawks, White-tailed Eagles, and Hooded Crows) revealed at least two distinct lineage 2 clusters closely related to strains responsible for outbreaks in Greece, Hungary, and Italy [24]. The detection of lineage 2 in Belgium for the first time in 2025, in three corvids, and its subsequent identification in additional birds later that season, provides clear evidence of the continued northward and westward expansion of this lineage across Europe [15].

The molecular characterization of WNV strains from wild birds has been instrumental in understanding viral dispersal patterns, transmission dynamics, and the emergence of pathogenic variants. Whole genome sequencing (WGS) of avian isolates has enabled high-resolution phylogeographic analyses that trace the movement of the virus across continents and identify key geographic hubs for viral incursion and dissemination [17, 27, 29, 33]. For example, continuous phylogeographic reconstruction of Italian WNV-2 strains from 2011 to 2023 revealed that the Italian clade originated in the area between Emilia-Romagna and Lombardy around 2009, followed by an east-to-west spread during the 2022–2023 transmission seasons [29]. Furthermore, genomic surveillance has identified specific mutations associated with increased pathogenicity, such as the F49L substitution in the NS2A gene, which is linked to neuronal tropism, and the M184V substitution in the NS4B gene, which has been associated with enhanced virulence in avian and mammalian hosts [29]. The glycosylation status of the envelope (E) protein is another critical genetic determinant of pathogenicity; studies have demonstrated that glycosylation of the E protein enhances viral replication in peripheral tissues, increases viremic levels, and is associated with heightened virulence in bird models, particularly in young chicks [34]. This glycosylation also confers thermal stability to the virus, which may facilitate its persistence in environmental conditions and enhance transmission efficiency [34].

The taxonomic classification of WNV within the Japanese encephalitis serocomplex has important implications for serological surveillance in wild birds, as cross-reactivity among flaviviruses can complicate diagnostic interpretation [6, 7, 9, 20]. Birds infected with WNV may produce antibodies that cross-react with USUV, SLEV, or Bagaza virus (BAGV), necessitating the use of virus neutralization tests (VNT) for confirmation [7, 9, 19]. Experimental studies have demonstrated that prior infection with WNV can provide complete protection against subsequent challenge with SLEV genotype III and USUV in house sparrows (Passer domesticus), indicating that cross-protective immunity may shape the co-circulation dynamics of these flaviviruses in avian communities [6]. However, naturally exposed birds with prior WNV immunity have been shown to develop viremia upon challenge with heterologous flaviviruses, suggesting that the degree of cross-protection may vary under natural conditions and may be influenced by the intensity and duration of prior immunity [6]. This immunological interplay has profound implications for the emergence and displacement of flaviviruses in regions where multiple serocomplex members co-circulate, such as the United States (WNV and SLEV) and Europe (WNV and USUV) [6, 10, 16, 26].

The taxonomic diversity of WNV is also reflected in the differential susceptibility and reservoir competence of various avian species. Experimental infections have revealed that passeriform birds, particularly members of the families Corvidae (crows, jays, magpies), Turdidae (thrushes), and Passeridae (sparrows), are highly competent reservoirs, developing high-titer viremias that efficiently infect feeding mosquitoes [21, 38]. In contrast, raptors (Accipitriformes and Strigiformes) are highly susceptible to WNV infection and often develop severe neurological disease, but their role as amplifying hosts may be limited by high mortality rates [9, 25, 38]. Nevertheless, birds of prey are considered valuable sentinel species for WNV surveillance due to their high exposure rates and the ease of detecting clinical disease [9, 25, 38]. The reservoir potential of a given avian species is influenced by a complex interplay of eco-ethological traits, including migratory behavior, habitat use, gregariousness, and immunological factors [18, 21, 35]. A recent trait-based model applied to 150 European passerine species identified specific characteristics associated with higher WNV seroprevalence, such as insectivorous diet, ground-foraging behavior, and residency in wetland habitats, and used these traits to generate a WNV Reservoir Index (RI) that maps the spatial heterogeneity of reservoir potential across Europe [18]. This index showed a strong positive association with the number of years in which human WNV cases were notified at the NUTS administrative level, validating the utility of avian trait-based approaches for predicting zoonotic risk [18].

In summary, the taxonomy and overview of WNV in wild birds encompass a complex and dynamic interplay between viral genetics, avian host biology, and ecological factors that govern transmission at local, regional, and global scales. The recognition of multiple genetic lineages, particularly the globally dominant lineage 1 and the rapidly expanding European lineage 2, and the identification of key avian reservoir species have provided a foundational framework for understanding the epidemiology of this emerging zoonotic pathogen. Continued genomic surveillance of WNV in wild birds, coupled with ecological and immunological studies, remains essential for tracking viral evolution, predicting emergence events, and informing evidence-based control strategies under a One Health framework [1, 14, 17, 29, 32, 33].

Molecular Pathogenesis of West Nile Virus in Avian Hosts

The molecular pathogenesis of West Nile virus (WNV) in avian hosts represents a complex interplay between viral genetic determinants, host immune responses, and species-specific susceptibility factors that ultimately dictate the outcome of infection. As the primary amplifying hosts in the enzootic transmission cycle, birds are not merely passive reservoirs but rather active participants in viral evolution, selection, and dissemination. Understanding the molecular mechanisms underlying WNV infection in birds is critical for predicting emergence patterns, assessing zoonotic risk, and developing targeted surveillance strategies, as recognized by the World Health Organization (WHO) and the World Organisation for Animal Health (WOAH) in their global arbovirus monitoring frameworks.

Viral Entry and Cellular Tropism in Avian Tissues

The initial molecular events of WNV infection in birds commence with viral attachment to host cell receptors, a process mediated primarily by the envelope (E) glycoprotein. The E protein facilitates receptor-mediated endocytosis, with dendritic cell-specific ICAM-3-grabbing non-integrin (DC-SIGN) and integrins serving as putative entry receptors in avian cells. Following entry, the positive-sense single-stranded RNA genome is released into the cytoplasm, where it undergoes immediate translation into a single polyprotein that is subsequently cleaved by viral and host proteases into three structural proteins (capsid [C], pre-membrane [prM], and envelope [E]) and seven non-structural proteins (NS1, NS2A, NS2B, NS3, NS4A, NS4B, NS5). The non-structural proteins orchestrate genome replication, host immune evasion, and modulation of the cellular environment to favor viral propagation.

Tissue tropism in avian hosts is remarkably broad, reflecting the capacity of WNV to infect multiple organ systems. Experimental infections have demonstrated that the virus initially replicates in peripheral tissues, particularly in the skin and draining lymph nodes at the site of mosquito inoculation, before disseminating to visceral organs. The spleen, kidney, and heart are primary sites of viral amplification, with the central nervous system (CNS) representing a target for neuroinvasive strains. In birds of prey, which are particularly susceptible to WNV infection [38], viral RNA has been detected at high titers in the brain, spinal cord, and eyes, correlating with the severe neurological signs frequently observed in raptors. The molecular basis for this enhanced neurotropism in certain avian species remains an active area of investigation, though it likely involves differential expression of attachment factors and variations in blood-brain barrier integrity across taxonomic groups.

Genetic Determinants of Virulence and Pathogenicity

The molecular pathogenesis of WNV in birds is profoundly influenced by specific genetic determinants within the viral genome. The envelope protein glycosylation status has emerged as a critical virulence factor, with studies demonstrating that glycosylation at position N154 of the E protein enhances viral replication in peripheral organs and is associated with increased pathogenicity in avian models [34]. Glycosylated E proteins facilitate more efficient viral entry and dissemination, likely through improved interactions with cellular lectins and enhanced stability of the virion. In young chick infection models, glycosylated WNV strains exhibited heat-stable multiplication properties and achieved significantly higher viremic levels compared to non-glycosylated variants, directly correlating with increased mortality rates [34].

The NS3 helicase/protease and NS5 RNA-dependent RNA polymerase harbor additional determinants of avian virulence. A specific proline-to-histidine substitution at position 249 of the NS3 protein (H249P) has been identified as a molecular marker associated with enhanced pathogenicity in corvids, particularly American crows. This mutation increases the efficiency of viral RNA replication, leading to higher viral loads and accelerated disease progression. The presence of this mutation in lineage 2 strains responsible for the 2010 Greek outbreak [22] and subsequent European epizootics underscores the evolutionary pressure favoring variants with increased replicative fitness in avian hosts.

Recent genomic analyses have identified additional mutations linked to enhanced pathogenicity in birds. The F49L substitution in the NS2A gene has been associated with increased neuronal tropism, while the M184V mutation in the NS4B gene correlates with heightened pathogenicity in Italian WNV lineage 2 strains [29]. These mutations likely function through modulation of the host interferon response, as NS4B is a known antagonist of the JAK-STAT signaling pathway. The co-circulation of multiple genetic variants within avian populations, as demonstrated by the simultaneous detection of subclusters 2.5.3.4.3c and 2.5.3.2 in German birds [17], indicates ongoing viral evolution that may alter pathogenic potential over time.

Host Immune Responses and Immunopathogenesis

The avian immune response to WNV infection involves both innate and adaptive components, with the outcome of infection largely determined by the kinetics and magnitude of these responses. The innate immune system provides the first line of defense, with pattern recognition receptors such as RIG-I and MDA5 detecting viral RNA and triggering type I interferon (IFN) production. However, WNV has evolved sophisticated countermeasures to evade these responses. The NS1 protein inhibits complement activation by binding to factor H, while NS5 suppresses interferon signaling through STAT2 degradation. The efficiency of these immune evasion strategies varies across avian species, contributing to the marked differences in reservoir competence observed among bird taxa.

Humoral immunity plays a central role in controlling WNV infection, with neutralizing antibodies directed primarily against the E protein. The development of IgY antibodies, the avian equivalent of mammalian IgG, is critical for viral clearance and long-term protection. Serological studies have demonstrated that prior WNV infection provides robust cross-protection against closely related flaviviruses, including St. Louis encephalitis virus (SLEV) and Usutu virus (USUV). Experimental infections in house sparrows revealed that WNV-immune birds were completely protected against secondary challenge with SLEV genotype III and USUV, with no detectable viremia [6]. This cross-protection is mediated by cross-neutralizing antibodies that recognize conserved epitopes within the E protein, though the titers of these antibodies are often low, suggesting that additional cellular immune mechanisms contribute to protection.

The phenomenon of seroreversion, where previously seropositive birds become seronegative over time, has important implications for interpreting surveillance data and understanding population immunity. Longitudinal studies in common coots demonstrated that some birds seroreverted within one year of initial capture [42], indicating that antibody titers may wane below detectable levels in the absence of re-exposure. This finding suggests that seroprevalence surveys may underestimate the true extent of WNV circulation, as birds with prior exposure may test negative if sampled months or years after infection.

Species-Specific Susceptibility and Reservoir Competence

The molecular pathogenesis of WNV varies dramatically across avian species, with differences in susceptibility, viremia duration, and disease outcome reflecting underlying genetic and immunological variations. Passeriformes, particularly corvids, are among the most susceptible hosts, often developing high-titer viremia and fatal neurological disease. In contrast, many columbiforms (pigeons and doves) and galliforms (chickens and turkeys) exhibit lower viremia levels and minimal clinical signs, making them less efficient amplifying hosts. These differences have been quantified through experimental infections, which have identified viral and host factors that determine infection outcome [21].

The molecular basis for this differential susceptibility involves multiple factors, including variations in viral receptor expression, differences in interferon signaling efficiency, and polymorphisms in major histocompatibility complex (MHC) genes. Birds of prey, including Accipitriformes and Strigiformes, demonstrate particularly high susceptibility to WNV infection, with seroprevalence rates reaching 25.77% and 22.92%, respectively, in Spanish studies [9]. The elevated susceptibility of raptors may be partially attributable to their predatory behavior, which increases exposure through consumption of infected prey, as well as potential genetic factors that impair their ability to control viral replication.

Reservoir competence, defined as the ability of a host to transmit virus to feeding mosquitoes, is determined by the magnitude and duration of viremia. Species that develop high-titer viremia (>10^5 plaque-forming units/mL) for extended periods are considered competent reservoirs. Experimental studies have identified several passerine species, including house sparrows, common grackles, and blue jays, as highly competent reservoirs, while many non-passerine species exhibit lower competence. The molecular determinants of viremia magnitude include the efficiency of viral RNA replication, the capacity of the host immune system to control infection, and the ability of the virus to evade host defenses.

Viral Persistence and Overwintering Mechanisms

The molecular mechanisms underlying WNV persistence in avian hosts during winter months, when mosquito vectors are inactive, represent a critical aspect of viral ecology. Evidence of WNV RNA detection in birds during cold seasons, including December in Italy [40, 41] and February in Greece [8], suggests that birds may serve as overwintering reservoirs. The molecular basis for viral persistence likely involves establishment of low-level chronic infection in immunologically privileged sites, such as the central nervous system or lymphoid tissues, where the virus can evade immune clearance.

The detection of WNV RNA in tissues of apparently healthy birds months after initial infection indicates that viral persistence may be more common than previously recognized. In Italian studies, WNV RNA was detected in a little grebe during December [40], while Greek surveillance identified positive birds from February through November [8], suggesting year-round circulation. The molecular mechanisms facilitating persistence may include downregulation of viral replication to levels below the threshold for immune activation, selection of quasispecies with reduced cytopathogenicity, and exploitation of cellular stress responses that promote cell survival.

Co-infections and Interactions with Other Flaviviruses

The molecular pathogenesis of WNV in avian hosts is further complicated by co-circulation with other flaviviruses, particularly USUV and SLEV. The cross-reactivity of immune responses between these viruses has significant implications for disease dynamics and surveillance interpretation. Serological studies have documented widespread co-circulation of WNV and USUV in European bird populations, with differential virus neutralization tests required to distinguish between infections [9, 10]. The presence of cross-reactive antibodies complicates serosurveillance efforts, as ELISA-positive samples may reflect exposure to any of several flaviviruses.

Experimental evidence indicates that prior infection with one flavivirus can modulate the outcome of subsequent infection with another. WNV-immune birds challenged with USUV or SLEV genotype III showed complete protection from viremia [6], suggesting that heterologous immunity may limit the emergence of novel flaviviruses in regions with established WNV circulation. However, the observation that naturally WNV-exposed house sparrows developed viremia upon USUV challenge [6] indicates that the level and duration of cross-protection may be influenced by the intensity and recency of primary infection.

Molecular Epidemiology and Evolutionary Dynamics

The application of whole genome sequencing and phylogenetic analysis to avian WNV isolates has revealed complex patterns of viral evolution and dispersal. Studies in Germany demonstrated that WNV lineage 2 strains circulating in birds in 2023-2024 belonged predominantly to subcluster 2.5.3.4.3c, with approximately 27% of sequences clustering within group 2.5.3.2 [17], indicating co-circulation of genetically distinct variants. The emergence of these variants likely reflects ongoing adaptation to local avian hosts and mosquito vectors, with selection favoring mutations that enhance transmission efficiency.

Phylogeographic analyses have traced the movement of WNV between continents via migratory birds, with Spanish studies revealing that lineage 1 strains from southern Spain are phylogenetically related to strains from Senegal, Morocco, Italy, France, and Portugal [33]. The identification of Spain's southernmost province, Cádiz, as a hotspot for virus incursion from Africa [33] highlights the role of migratory birds in facilitating intercontinental viral traffic. Similarly, a 2024 Portuguese isolate from a goshawk was found to belong to lineage 1a, with phylodynamic analysis suggesting an origin in West Africa and introduction via the Cádiz coast [27], confirming the importance of Afro-Palearctic migratory flyways in WNV dispersal.

The molecular clock analysis of WNV genomes from Italian birds and mosquitoes indicates that the main European clade originated in Hungary around 2004, with introduction into Italy occurring between 2009 and 2010 [29]. The subsequent east-to-west spread across northern Italy during 2022-2023 demonstrates how avian movements facilitate viral expansion into new geographic areas. These findings underscore the importance of integrated genomic surveillance combining avian, mosquito, and human samples to track viral evolution and anticipate future emergence events.

Epidemiology and Transmission Dynamics of West Nile Virus in Wild Bird Populations

The perpetuation of West Nile virus (WNV) in nature is fundamentally dependent upon a complex sylvatic cycle involving ornithophilic mosquito vectors and susceptible avian reservoir hosts. Wild birds serve as the primary amplifying hosts, developing viremias of sufficient magnitude to infect feeding mosquitoes, thereby sustaining enzootic transmission [1, 4, 14, 21, 37, 38]. Understanding the epidemiological patterns and transmission dynamics within wild bird populations is paramount for predicting spillover risk to humans and horses, as recognized by global health authorities including the World Health Organization (WHO), the World Organisation for Animal Health (WOAH), and the Centers for Disease Control and Prevention (CDC).

The Enzootic Cycle and Reservoir Competence

The fundamental transmission unit for WNV is the mosquito-bird-mosquito cycle, primarily involving Culex spp. mosquitoes as vectors [11, 23, 28, 30]. Within this cycle, avian reservoir competence, the ability of a bird species to become infected and subsequently transmit the virus to a naïve vector, exhibits extraordinary interspecific variation. This variation is driven by viral and host factors, including the magnitude and duration of viremia, which is determined by host susceptibility, immune status, and viral genetic determinants such as envelope protein glycosylation [21, 34, 37]. Experimental infections have demonstrated that certain passerine species, particularly those in the families Corvidae, Turdidae, and Passeridae, can develop viremias exceeding 10⁷ plaque-forming units per milliliter of serum, levels sufficient to infect Culex vectors at high efficiency [21]. Conversely, species such as the barn swallow (Hirundo rustica) and various Piciformes appear to exhibit lower viremic potential or rapid viral clearance, as evidenced by serosurveys in Ukraine where no antibodies were detected in barn swallows despite co-circulation of WNV in sympatric species [2]. This heterogeneity in reservoir competence establishes a hierarchical structure within avian communities, where a minority of "super-spreader" species may disproportionately drive transmission intensity.

Longitudinal serosurveillance across multiple continents has confirmed that certain avian taxa consistently demonstrate elevated seroprevalence. In Spain, raptors (Accipitriformes and Strigiformes) and Pelecaniformes have shown seroprevalences of 25.77% and 33.33%, respectively, following epidemic outbreaks [9, 25]. In Germany, Eurasian goshawks (Astur gentilis) have emerged as the predominant species among WNV RNA-positive birds, comprising the majority of identified cases in nationwide surveillance [17]. This raptor affinity may be partly attributable to their trophic position: predation on infected passerines provides an additional oral route of exposure, potentially leading to higher infection prevalence than would be expected from vector-borne transmission alone [38]. The northern cardinal (Cardinalis cardinalis) has been identified as a particularly informative sentinel species in North America, exhibiting consistently high seroprevalence across urban gradients in both Atlanta and Chicago, and explaining more variance in seroprevalence than year or location [35].

Transmission Pathways and Vector-Bird Interactions

While the canonical transmission pathway involves mosquito inoculation, accumulating evidence suggests that alternative transmission routes may contribute to viral maintenance in bird populations. Direct bird-to-bird transmission via the fecal-oral route has been hypothesized, based on experimental detection of WNV RNA in feces of infected birds, though field studies attempting to detect WNV in fecal samples from wild birds in Nigeria yielded negative results, suggesting this route may be epidemiologically insignificant under natural conditions [45]. However, the detection of WNV RNA in multiple tissue types, including brain, myocardium, kidney, and spleen, in naturally infected birds during autumn and winter months in Italy indicates that the virus can persist in avian hosts beyond the typical transmission season, potentially facilitating overwintering mechanisms [46].

The interplay between mosquito community composition and avian infection risk is exquisitely nuanced. In southwestern Spain, the abundance of ornithophilic Culex perexiguus was positively correlated with WNV neutralizing antibody prevalence in house sparrows, whereas the abundance of mammophilic species such as Ochlerotatus caspius and Anopheles atroparvus was negatively associated [11]. This suggests that mosquito community structure exerts a deterministic influence on enzootic amplification, with ornithophilic species acting as bridge vectors that sustain transmission in avian reservoirs. Further supporting this, research in Louisiana demonstrated that exposure to Aedes albopictus bites, measured via IgY antibodies against mosquito salivary proteins, was more strongly associated with WNV exposure in northern cardinals than exposure to Culex quinquefasciatus [5]. This finding implies that Ae. albopictus may play a previously underappreciated role in early-season transmission, particularly among brooding females and hatchlings, while Cx. quinquefasciatus becomes more relevant later in the season as recaptured birds showed increased exposure to this species [5].

Factors Modulating Transmission Intensity: Climate, Biodiversity, and Land Use

Transmission dynamics are profoundly modulated by abiotic and biotic factors. Altitude, land use, and distance from water bodies have emerged as significant predictors of WNV occurrence in birds [8, 30]. Ecological niche modeling in the Peloponnese region of Greece identified low-altitude zones with specific land cover types as especially suitable for virus circulation [30]. Similarly, in Greece, population density and proximity to water sources were important factors associated with detection of WNV RNA in wild birds [8]. These landscape-level determinants likely operate through their influence on mosquito breeding habitat availability and bird community composition.

The dilution effect hypothesis proposes that high avian biodiversity dampens WNV transmission by reducing the relative abundance of competent reservoir hosts. Empirical support for this hypothesis was provided by studies in southwestern Spain, where higher phylogenetic diversity in bird communities correlated with lower WNV exposure rates [23]. Conversely, bird communities dominated by passerines, particularly in farm environments where wildlife-livestock interactions are intensified, exhibited elevated seroprevalence [43]. In these agricultural settings, the bird community composition shifted toward a predominance of competent passerine species, and exposure risk was maximized in the immediate vicinity of horse farms (9%) compared to surrounding natural areas (5.8%) [43]. The presence of equines, while not contributing to transmission as dead-end hosts, may inadvertently concentrate ornithophilic mosquitoes and avian reservoirs, creating transmission hotspots.

Climate change is increasingly recognized as a driver of WNV expansion into previously non-endemic areas. Temperature influences mosquito vector competence, extrinsic incubation period, and viral replication rates within the vector, while also affecting bird migration phenology and breeding success [14, 31]. In Germany, the northward expansion of WNV lineage 2 following its initial detection in 2018 has been attributed to increasingly favorable climatic conditions, with WNV RNA-positive birds detected in 13 federal states by 2024, compared to only six in 2023 [17]. Similarly, the detection of WNV lineage 1a in a goshawk from Portugal in 2024, with phylogeographic reconstruction indicating an origin in West Africa and spread via migratory routes through Senegal, Mauritania, and Morocco, underscores how changing climatic envelopes facilitate transcontinental viral movement [27].

Spatial Dynamics: Migration and Viral Dispersal

Migratory birds constitute a critical mechanism for the long-distance dispersal of WNV, connecting disparate geographic regions and facilitating viral introduction into naive populations [12, 22, 27, 33, 36, 44]. The Afro-Palearctic migratory flyway, which connects sub-Saharan Africa with Europe, has been implicated in the repeated introduction of WNV lineage 1 and lineage 2 into southern Europe [27, 33]. Phylogeographic analyses of WNV lineage 2 in Italy traced the origin of the main European clade to Hungary in 2004, with introduction into Italy between 2009 and 2010, subsequently followed by east-west spread within the country [29]. This pattern mirrors the migratory routes of numerous passerine species that breed in central and eastern Europe and winter in sub-Saharan Africa.

Species-specific migration traits significantly influence viral introduction and amplification dynamics. In Greece, serological evidence of WNV exposure in migratory birds upon their arrival during autumn migration, prior to the 2010 outbreak, implicated long-distance migrants as likely introduction vehicles [22]. The Eurasian blackcap (Sylvia atricapilla), a common and widespread migrant, represented the majority of seropositive birds on Cyprus, implying a high risk of WNV introduction throughout the island [44]. In Malaysia, phylogenetic analysis of 16 WNV isolates from resident and migratory birds revealed 99% similarity to strains from South Africa within lineage 2, suggesting that migratory birds connecting the Palearctic and Oriental regions are introducing novel viral variants into Southeast Asia [12].

The distinction between resident and migratory birds is critical for interpreting serosurveillance data. Resident birds, particularly juveniles, provide evidence of local, ongoing transmission, whereas seropositive migratory birds may reflect exposure acquired elsewhere along their migratory route or at their wintering grounds [7, 47]. In Madrid, two seropositive juvenile birds that could barely fly provided compelling evidence of local flavivirus transmission within the region [7]. Similarly, in Morocco, high WNV seroprevalence among juvenile resident birds during non-epidemic periods indicated sustained local circulation independent of migratory influx [47].

Temporal Patterns and Overwintering

Seasonality is a hallmark of WNV epidemiology, with transmission typically peaking during late summer and early autumn when mosquito populations are at their zenith. However, accumulating evidence challenges the assumption that transmission ceases entirely during winter months. In Northern Italy, WNV RNA was detected in wild birds during December, well outside the standard June-to-September monitoring period [46]. The detection of WNV RNA in an adult little grebe (Tachybaptus ruficollis) in December in Central Italy further supports the hypothesis that birds may serve as overwintering reservoirs, maintaining the virus through colder months when mosquito activity is minimal [40, 41]. This phenomenon has significant implications for surveillance design, as extending monitoring beyond the traditional vector season may enhance detection sensitivity and strengthen early-warning capacity [1, 46]. In Greece, WNV RNA was detected in wild birds from February through November, providing evidence of year-round circulation and possible overwintering [8]. In southern Spain, genomic analysis of WNV from mosquitoes collected in 2020 and 2021 demonstrated that the strain circulating in 2021 was highly related to the 2020 outbreak strain, suggesting that the virus overwintered in the region rather than being reintroduced annually [39].

Seroprevalence Patterns and Implications for Surveillance

Global seroprevalence in wild birds exhibits remarkable geographic variation, ranging from undetectable in some regions to exceeding 50% in others. In Ukraine, seropositivity rates varied dramatically by region, with Poltava and Khmelnytsky regions showing 86% and 67% seropositivity, respectively, compared to only 9% in Kyiv [2]. Species-specific seroprevalence in Ukraine was particularly striking: great tits, song thrushes, blackbirds, and several finch species exhibited seroprevalences between 60% and 100%, while house sparrows and field sparrows showed lower rates [2]. In Bangladesh, seroprevalence was 11.9% overall, with migratory tufted ducks showing 28.5% seropositivity compared to 12.5% in resident house crows [3]. In extreme cases, such as the Far Eastern region of Russia, WNV-specific antibodies were detected in 21 of 145 resident and migratory birds, indicating northward expansion of the virus into temperate Asia [34].

The strategic selection of sentinel species is crucial for cost-effective surveillance. Magpies (Pica pica) have been proposed as particularly valuable sentinels due to their high seroprevalence, sedentary nature, and abundance in peri-urban environments [8, 22, 36]. In Serbia, phylogenetic analysis of a WNV lineage 2 strain isolated from a Eurasian magpie indicated close similarity to strains responsible for human outbreaks in Greece, Hungary, and Italy, underscoring the sentinel value of this species [24]. Raptors, particularly goshawks and white-tailed eagles, have also been identified as effective sentinels due to their elevated susceptibility and tendency to develop detectable infections [24, 38].

It must be acknowledged that passive surveillance, such as sampling at wildlife rehabilitation centers, while offering logistical advantages, introduces inherent biases. The low detection rates in central Spain (0% WNV RNA over a nine-year period) may partly reflect the variability inherent to passive surveillance, heterogeneity of sample types, and the predominance of subclinical or non-viremic infections in sampled species [1]. Conversely, active surveillance programs that target specific high-risk species during periods of peak transmission yield substantially higher detection rates [9, 30]. The integration of both passive and active surveillance modalities, combined with molecular and serological diagnostics, provides the most comprehensive picture of WNV circulation in avian populations [10, 32]. The multidisciplinary approach employed in the Netherlands, combining wild bird surveys, mosquito monitoring, and sentinel chickens, successfully detected local WNV circulation 35 days before the first autochthonous human case was identified, demonstrating the critical role of integrated avian surveillance in public health preparedness [32].

Clinical Manifestations and Pathological Findings in Infected Wild Birds

The clinical and pathological outcomes of West Nile virus (WNV) infection in wild birds are extraordinarily heterogeneous, ranging from entirely subclinical infections with transient viremia to rapid, fulminant neurological disease and death. This spectrum is governed by a complex interplay of viral factors, including lineage, genotypic determinants of virulence, and envelope protein glycosylation status, and host factors such as species, age, immunocompetence, and prior flavivirus exposure history. Understanding these manifestations is critical not only for identifying infected birds in surveillance programs, but also for elucidating the mechanisms by which WNV achieves high-titer viremia sufficient to infect feeding mosquitoes, thereby perpetuating the enzootic cycle.

Neurological Manifestations and Central Nervous System Pathology

Neurological disease is the most clinically overt and frequently described manifestation of WNV infection in susceptible wild birds, although it is important to recognize that many infected individuals, particularly in competent reservoir species, remain asymptomatic. Among passerines, which are considered primary amplifying hosts, neurological signs are often subtle or absent, yet viremia levels may be extremely high [21, 38]. In contrast, raptors, including Eurasian goshawks (Astur gentilis), white-tailed eagles (Haliaeetus albicilla), and various owl species, are disproportionately susceptible to severe neuroinvasive disease, frequently exhibiting overt clinical signs and high mortality rates [24, 25, 38]. The heightened susceptibility of raptors is hypothesized to relate to their predatory feeding behavior, which may result in oral ingestion of infected prey, leading to direct viral entry into the central nervous system (CNS) via the olfactory or trigeminal nerves, bypassing the peripheral immune defenses that typically limit neuroinvasion in passerines [38].

Clinically, affected birds present with a constellation of neurological deficits that include ataxia, head tilt, torticollis, circling, tremors, proprioceptive deficits, paresis, paralysis, and seizures. Obtundation and stupor may progress to coma and death within hours to days of symptom onset [20, 40, 41, 50]. These signs reflect the predilection of WNV for neuronal tissues, particularly the brainstem, cerebellum, and spinal cord. Gross pathological examination of the CNS in acutely infected birds is frequently unremarkable, although mild congestion or edema may be noted [20]. Histopathological examination, however, reveals the hallmark lesions of WNV encephalitis: multifocal to diffuse lymphocytic meningoencephalitis, perivascular cuffing with mononuclear cells (primarily T-lymphocytes and macrophages), gliosis, microglial nodule formation, and neuronal necrosis (neuronophagia) [20, 40, 41]. In a notable case of WNV infection in a little grebe (Tachybaptus ruficollis) from central Italy, the predominant CNS lesions included CD3+ lymphocytic meningoencephalitis, consistent with a T-cell-mediated immunopathological response [40, 41]. The detection of WNV RNA in brain tissue by RT-PCR is a definitive diagnostic finding, and the presence of viral antigen can be confirmed via immunohistochemistry [20, 24].

Extraneural Pathology: Cardiac, Renal, and Hepatic Involvement

While neurotropism is a defining feature of WNV, the virus exhibits extensive tropism for extraneural tissues, and pathological changes outside the CNS are frequently observed and may contribute significantly to morbidity and mortality. The heart, kidney, and spleen are consistently implicated as sites of viral replication and pathological injury [20, 50].

Myocarditis is a common and potentially fatal extraneural manifestation. Histologically, affected hearts demonstrate multifocal myocyte degeneration, necrosis, and loss, accompanied by interstitial edema and a mixed inflammatory infiltrate dominated by lymphocytes (CD3+ T-cells) and macrophages [20, 40, 41]. In a detailed post-mortem study of WNV-infected birds from Italy, cardiomyocyte loss and interstitial edema were prominent findings in the little grebe, alongside the aforementioned meningoencephalitis [40, 41]. Such myocardial injury can compromise cardiac output and may contribute to acute death, particularly in birds under the physiological stress of migration or concurrent disease.

Renal pathology is another hallmark of WNV infection in birds. Grossly, the kidneys may appear pale and swollen. Microscopic examination reveals lymphocytic tubulo-interstitial nephritis, often with a predominance of CD3+ T-cells infiltrating the interstitium and surrounding tubules [20, 40, 41]. Tubular epithelial cell degeneration, necrosis, and sloughing into the tubular lumen are observed. The functional consequence of this renal injury may include reduced glomerular filtration and electrolyte imbalance. Importantly, the virus is shed in high concentrations in the urine, and the presence of WNV RNA in cloacal swabs is a well-documented phenomenon, reflecting viral replication within the urogenital tract [3, 13, 45]. This cloacal shedding is a critical mechanism for direct bird-to-bird transmission, potentially via fecal contamination of water sources or shared feeding sites, independent of mosquito vectors [45].

Hepatic and splenic involvement is also frequently documented. The spleen, a primary lymphoid organ in birds, often exhibits lymphoid depletion, necrosis, and the presence of virus-laden macrophages. The liver may show multifocal lymphocytic hepatitis, with scattered foci of hepatocellular necrosis and Kupffer cell hyperplasia [20]. While not typically the direct cause of death, hepatic and splenic damage compromises the bird's immune response and overall physiological resilience.

Species-Specific Variation in Clinical and Pathological Outcomes

The clinical and pathological response to WNV infection is profoundly species-dependent, and this variation is the cornerstone of understanding reservoir competence and outbreak dynamics. Passerine species such as house sparrows (Passer domesticus), Northern cardinals (Cardinalis cardinalis), and various thrushes (e.g., Turdus merula) are highly competent reservoirs, typically developing high-titer, prolonged viremia sufficient to infect feeding Culex mosquitoes, yet often displaying minimal or no clinical signs [6, 11, 21, 35]. These species are the engine of WNV amplification in nature. For instance, experimental infections of house sparrows with WNV lineage 1 and 2 strains consistently produce viremia lasting 4–7 days, with peak titers exceeding (10^8) plaque-forming units (PFU)/mL of serum, even in the absence of clinical illness [21]. In contrast, infection in many raptor species, including goshawks, eagle owls (Bubo bubo), and great horned owls, is far more pathogenic, often resulting in rapid neurological decline and death [24, 38]. This high mortality in raptors is paradoxical because these birds are not considered primary reservoirs (competent hosts) due to their lower population density, but they may serve as sentinels, and their carcasses are frequently the first indicator of WNV incursion into a new area [38].

Even within the same species, age is a critical determinant. Juvenile birds, particularly nestlings and fledglings, are more susceptible to severe disease and death than adults, likely due to their immature immune systems. Seroconversion in adult birds, however, occurs at high rates, and adults that survive infection develop long-lasting immunity [25, 35, 42]. In a longitudinal study of common coots (Fulica atra) in Spain, antibody prevalence fluctuated between years, and some birds seroreverted within 12 months, indicating waning humoral immunity, though cellular immunity likely persists [42].

Subclinical Infection, Seroconversion, and the Role of Non-Lethal Infections

The vast majority of WNV infections in wild birds are subclinical, meaning that infected birds show no overt signs of illness but mount a detectable antibody response. This phenomenon is the basis for serosurveillance programs that measure WNV-specific antibodies (IgY) in wild bird populations to determine the history and intensity of local virus circulation [2, 3, 7, 35, 44, 47-49]. For example, serosurveys in Ukraine, Bangladesh, Morocco, and Spain have demonstrated widespread seropositivity in multiple passerine and non-passerine species, even when no clinical cases are detected [2, 3, 7, 47]. In these scenarios, the birds are infected, develop viremia, mount an immune response, and subsequently clear the virus, often without ever entering a rehabilitation center or being reported dead. The presence of neutralizing antibodies (NAb) at titers >1:10 indicates past infection and serves as a reliable marker of exposure [7, 47, 48].

Seroconversion, the first detection of antibodies in a previously naive bird, is a powerful indicator of recent, local WNV transmission. Studies in Spain, Greece, and Italy have documented seroconversion in juvenile birds captured during the late summer and early autumn, immediately following the peak mosquito season, providing clear evidence of local virus amplification [7, 8, 42, 48]. Furthermore, cross-protection studies have shown that birds with pre-existing WNV immunity are completely protected against secondary challenge with closely related flaviviruses, such as St. Louis encephalitis virus (SLEV) and Usutu virus (USUV) [6]. This immune cross-protection has profound ecological implications, as it may limit the co-circulation and emergence of these related viruses in areas where WNV is endemic [6].

Pathological Findings in Asymptomatic Yet Viremic Birds

Even in birds that appear clinically healthy, pathological changes may be present upon necropsy. In a comprehensive German surveillance study involving hundreds of wild birds, many individuals that tested positive for WNV RNA by RT-PCR in organ tissues showed no gross lesions [51]. However, histopathological examination of these same tissues frequently revealed mild lymphocytic infiltrates in the heart, kidney, and brain, indicating subclinical inflammation. These findings underscore that WNV can cause tissue damage even in the absence of clinical signs, and that the absence of overt disease does not preclude the bird from being an effective amplifying host. In these sublethally infected birds, the virus replicates to high titers in multiple organs, and the bird is capable of shedding virus in oral and cloacal secretions, contributing to transmission [45, 50].

Overwintering and Chronic Infection: The Enigmatic Role of Avian Reservoirs

One of the most intriguing aspects of WNV epidemiology is its ability to overwinter in temperate regions, persisting through the winter months when mosquito vectors are inactive. The detection of WNV RNA in a little grebe during December in central Italy provides compelling evidence that wild birds may serve as overwintering reservoirs [40, 41]. Similarly, in Greece, WNV RNA was detected in wild birds from February to November, indicating year-round circulation, with the virus likely persisting in a subset of infected birds that maintain a chronic, low-level infection [8]. Chronic infection in birds, while not definitively proven for WNV, is hypothesized to involve viral persistence in lymphoid or neural tissues, with intermittent reactivation under periods of stress (e.g., migration, food scarcity, concurrent infection). The observation that WNV RNA could be detected in the organs of birds months after the end of the mosquito season suggests that these birds may be responsible for re-introducing the virus to the vector population the following spring [8, 40, 46]. This overwintering mechanism is critical for the long-term maintenance of WNV in areas with harsh winters and underscores the need for year-round surveillance of avian populations, not just during the vector season [8].

Diagnostic Approaches for West Nile Virus Detection in Wild Birds

The accurate and timely detection of West Nile virus (WNV) in wild bird populations is fundamental to understanding the epizootiology of this globally significant zoonotic arbovirus. As the primary amplifying hosts in the enzootic transmission cycle, wild birds serve as critical sentinels for virus introduction, circulation, and potential spillover to humans and equids [14, 18, 38]. Given the expanding geographic range of WNV in Europe, the Americas, Africa, and Asia, driven in part by climate change and land-use alterations [8, 14, 31], robust diagnostic surveillance frameworks are indispensable. The World Organisation for Animal Health (WOAH) and the World Health Organization (WHO) emphasize the need for integrated, multi-species surveillance under a One Health umbrella, wherein bird monitoring provides an early warning system for human and veterinary public health [1, 32]. Diagnostic approaches for WNV in avian reservoirs have evolved substantially, encompassing direct viral detection through molecular and virological methods and indirect detection via serological assays. Each modality presents distinct advantages, limitations, and optimal applications depending on the epidemiological objectives, sample types available, and the timing of sampling relative to infection. This section provides a comprehensive, critical analysis of the diagnostic armamentarium employed for WNV detection in wild birds, with an emphasis on methodological rigor, interpretative challenges, and strategic deployment in surveillance programs.

Molecular Detection of Viral RNA

The direct detection of WNV RNA in avian tissues and exudates remains the gold standard for confirming active infection and is indispensable for characterizing circulating viral lineages and genotypes [4, 15, 17, 24]. Reverse transcription polymerase chain reaction (RT-PCR) and its quantitative variant (RT-qPCR) are the most widely employed molecular techniques, offering high sensitivity, specificity, and rapid turnaround times [8, 30, 46]. These assays typically target conserved regions of the viral genome, including the envelope (E) gene, the non-structural protein 3 (NS3) gene, the 5′ untranslated region (UTR), the 3′ non-coding region (3′NC), or the NS2A gene [1, 4, 8, 50]. The choice of target can influence assay performance; for instance, the 3′NC region is highly conserved among flaviviruses, making it suitable for broad flavivirus screening, whereas E-gene or NS3-targeted assays offer greater specificity for WNV lineage discrimination [1, 4, 8].

Sample selection and collection critically impact molecular detection success. The temporal window for detecting WNV RNA in wild birds is relatively narrow, as viremia peaks 2–6 days post-infection and may persist for only 7–10 days in many passerine species [21]. Consequently, sampling during the acute phase of infection is essential. Tissue tropism studies have identified the brain, kidney, spleen, and heart as organs with the highest viral loads in naturally and experimentally infected birds, even in cases where blood viremia has waned [20, 46, 50]. In active surveillance of live birds, whole blood, plasma, or oropharyngeal and cloacal swabs are commonly collected [1, 3, 12, 13]. Oropharyngeal swabs have demonstrated utility for detecting viral shedding, particularly in species where oral excretion is pronounced [1, 12]. Cloacal swabs, while less sensitive than tissue samples, can detect viral RNA shed in feces and may be useful for non-invasive sampling in captive or rehabilitating birds [1, 3]. For dead birds, pooled tissue samples from visceral and central nervous system (CNS) compartments are recommended, as this approach maximizes the probability of detection even in decomposed carcasses [50]. In a post-mortem study of 576 birds from Italian rescue centers, WNV RNA was detected in a single little grebe (Ct value 34.36) using RT-PCR on pooled tissue samples, illustrating that even low-level viremia can be captured with appropriate sampling [40, 41].

Real-time quantitative PCR (RT-qPCR) has supplanted conventional RT-PCR in most modern surveillance programs due to its ability to provide a quantitative measure of viral load via cycle threshold (Ct) values. Lower Ct values correlate with higher viral RNA copy numbers and may indicate acute infection with intense viral replication, whereas high Ct values near the limit of detection require careful interpretation and confirmatory testing [40, 41]. Multiplex RT-qPCR assays capable of simultaneously detecting WNV, Usutu virus (USUV), dengue virus, and Zika virus have been developed, enhancing surveillance efficiency in regions where multiple flaviviruses co-circulate [28]. In Kosovo, a WNV-DENV-ZIKV-specific multiplex RT-qPCR was used to screen mosquito pools, identifying WNV lineage 2 for the first time in the country, demonstrating the power of multiplexing for arbovirus discovery [28].

Whole genome sequencing (WGS) has become an increasingly critical tool for detailed molecular characterization of WNV strains, particularly in the context of emerging lineages and transboundary spread [15, 17, 29, 33]. WGS provides complete genetic information, enabling phylogenetic and phylogeographic analyses that trace virus dispersal pathways, identify evolutionary lineages, and detect mutations associated with virulence or vector competence [17, 29, 33, 37]. Recent studies have leveraged WGS to document the co-circulation of multiple WNV lineage 2 subclusters in Germany (e.g., subcluster 2.5.3.4.3c and cluster 2.5.3.2) [17], to reconstruct the introduction of WNV lineage 1a from Africa into Portugal via Spain [27], and to demonstrate the long-term persistence and spatial expansion of WNV lineage 2 in Italy [29]. Sequencing of WNV from wild birds has also revealed mutations with functional significance, such as F49L in NS2A (linked to neuronal tropism) and M184V in NS4B (associated with increased pathogenicity) [29]. These genomic insights are invaluable for real-time risk assessment and for informing vaccine and therapeutic development. The use of portable sequencing platforms like MinION has facilitated field-based WGS, enabling rapid characterization even in resource-limited settings [17].

Conventional RT-PCR followed by amplicon sequencing remains a useful approach in regions where advanced sequencing infrastructure is lacking. By amplifying and sequencing a portion of the E gene or NS3 gene, researchers can assign viral lineage and genotype with reasonable confidence. For example, in Greece, partial NS3 sequencing of WNV-positive bird samples confirmed all isolates as lineage 2, closely related to previous outbreak-causing Greek strains [8]. Similarly, in Malaysia, sequencing of 16 WNV isolates from wild birds revealed 99% similarity to South African lineage 2 strains, indicating transcontinental introduction via migratory flyways [12]. However, conventional sequencing provides only a partial genetic snapshot and may miss recombination events or minority variants that WGS can capture.

Critical considerations for molecular detection include the potential for false negatives due to low viral load, sample degradation, or inappropriate sample storage. RNA is labile, and samples should be preserved in RNA-stabilizing solutions or stored at −80°C to maintain integrity [1]. PCR inhibitors present in blood, feces, or tissue homogenates can also compromise assay performance; the use of internal controls and optimized extraction protocols is mandatory [3]. Furthermore, in regions where WNV circulates at very low prevalence, such as in central Spain where a nine-year surveillance program across 1,024 birds yielded zero WNV-positive samples by RT-PCR, the sample size and sampling strategy must be carefully designed to achieve adequate statistical power [1]. Passive surveillance at wildlife rehabilitation centers, while logistically convenient, may introduce bias toward sick or injured birds and may not reflect background circulation in healthy populations [1, 40, 41]. Active surveillance of free-ranging birds, targeting key species such as corvids, raptors, and passerines, is essential for unbiased prevalence estimation [8, 30, 38].

Serological Detection of Anti-WNV Antibodies

Serological detection of antibodies against WNV provides evidence of past or current infection and is particularly valuable for assessing cumulative exposure within avian populations, especially during inter-epidemic periods when viral RNA is rarely detectable [2, 7, 22, 42, 49]. Unlike molecular methods, which capture an instantaneous snapshot of active infection, serology reveals historical exposure, offering insights into transmission intensity, seasonal patterns, and population-level immunity [11, 18, 35, 48]. The primary serological tools for WNV detection in wild birds are enzyme-linked immunosorbent assays (ELISA) and virus neutralization tests (VNT), each with distinct performance characteristics and interpretative limitations.

Competitive enzyme-linked immunosorbent assay (cELISA) is the most commonly employed screening tool for WNV antibodies in wild birds due to its species-independent nature, high throughput, and commercial availability (e.g., ID Screen West Nile Competition Multi-species ELISA, IDvet, France) [2, 3, 7, 12, 19, 25, 43, 49]. The cELISA platform utilizes a competitive format where antibodies in the test serum compete with a labeled monoclonal antibody for binding to immobilized WNV antigen. A reduction in signal indicates the presence of anti-WNV antibodies. This assay detects immunoglobulin G (IgG) antibodies, which appear approximately 5–10 days post-infection and can persist for months to years, making it suitable for serosurveys [2, 3, 35].

The widespread application of cELISA in wild bird surveillance has yielded seroprevalence estimates ranging from 1.3% in Cyprus [44] to over 30% in Spain following the 2020 epidemic [25]. In Bangladesh, cELISA revealed 11.9% seropositivity among 184 wild birds, with higher rates in migratory tufted ducks (28.5%) compared to resident house crows (12.5%) [3]. In Ukraine, seroprevalence as measured by commercial WNV ELISA reached 100% in several passerine species, including chaffinch, goldfinch, and house sparrow, with notable regional variation (9% in Kyiv to 86% in Poltava) [2]. However, a critical limitation of cELISA is its inability to discriminate between WNV and other closely related flaviviruses, such as Usutu virus (USUV), St. Louis encephalitis virus (SLEV), Japanese encephalitis virus (JEV), and Bagaza virus (BAGV), due to cross-reactivity of antibodies against conserved epitopes [6, 7, 9, 12, 19, 43, 49]. This is particularly problematic in regions where multiple flaviviruses co-circulate, such as Europe and the Mediterranean basin, where USUV and WNV share overlapping avian and mosquito hosts [6, 9, 10, 16, 26]. In a Spanish study, 8.2% of wild birds tested positive by WNV cELISA, but confirmatory VNT indicated that only 2.5% were truly WNV-positive, while 2.5% had antibodies to an undetermined flavivirus, likely USUV or BAGV [7]. Similarly, in Extremadura, Spain, differential VNT revealed USUV-specific antibodies in 1.04% of birds and undetermined flavivirus reactivity in 4.17%, underscoring the need for confirmatory testing [9].

Virus neutralization test (VNT) , including micro-virus neutralization test (micro-VNT) and plaque reduction neutralization test (PRNT), is considered the serological gold standard for WNV confirmation due to its high specificity [7, 9, 11, 19, 22, 25, 26, 49, 52]. VNT measures the ability of antibodies in the test serum to neutralize infectious WNV in cell culture, typically using Vero cells, and requires BSL-3 containment facilities [7, 11]. The test is performed by incubating serial dilutions of heat-inactivated serum with a standardized dose of WNV before inoculating cell monolayers. The endpoint titer is defined as the highest dilution that reduces plaque formation or cytopathic effect by 90% (PRNT90) or 50% (PRNT50) [7, 11, 22, 49, 52]. VNT is particularly valuable for discriminating between WNV and USUV or other flavivirus infections, as neutralizing antibodies often exhibit higher titers against the homologous virus [6, 9, 10, 26]. Differential VNT involves parallel testing against multiple flaviviruses (e.g., WNV, USUV, JEV, SLEV) and comparing endpoint titers; a four-fold or greater difference in titer is typically used to assign specificity [6, 9, 10, 26].

In practice, a two-step serological algorithm is widely adopted: cELISA screening followed by VNT confirmation of all positive and doubtful samples [7, 9, 25, 49]. This approach balances throughput with specificity. For example, in a study of wild birds in Bangladesh, 11.9% were cELISA-positive, but confirmatory VNT was not performed, leaving the possibility that some positives represented cross-reactions with JEV or other flaviviruses endemic to the region [3]. In contrast, in Greece, WNV neutralizing antibodies were confirmed by VNT in 53 of 295 hunter-harvested birds, providing robust evidence of local enzootic circulation [22]. The specificity of VNT is not absolute, as low-level cross-neutralization between WNV and USUV or SLEV can occur, particularly in birds with high antibody titers or after secondary flavivirus exposure [6]. Experimental studies in house sparrows have shown that prior WNV infection confers complete protection against subsequent SLEV genotype III and USUV challenge, yet low levels of cross-neutralizing antibodies were detected, suggesting that immunological memory rather than sterile immunity may be responsible [6]. This finding has important implications for interpreting VNT results in naturally exposed birds where sequential flavivirus infections may occur.

IgM antibody detection can differentiate recent from past infection, as IgM appears earlier than IgG (2–5 days post-infection) and wanes within a few weeks to months [25]. Commercial IgM capture ELISAs specific for WNV are available for some species but have not been extensively validated in wild birds. In a Spanish study, no IgM antibodies were detected in any of 243 unvaccinated equids sampled post-outbreak, indicating that all infections were likely historical [25]. While IgM detection could theoretically identify recent infections in birds, its utility is limited by the narrow detection window and the need for species-specific reagents.

Novel serological approaches include the detection of antibodies against mosquito salivary proteins as a proxy for exposure intensity [5]. In

Surveillance Strategies and One Health Implications of West Nile Virus in Wild Birds

The integrated surveillance of West Nile virus (WNV) in wild birds represents a cornerstone of modern zoonotic disease monitoring, bridging the disciplines of wildlife ecology, vector biology, virology, and public health. Given the complex sylvatic cycle involving ornithophilic mosquito vectors and avian amplifying hosts, the design of effective surveillance systems must account for the ecological heterogeneity of transmission, the temporal dynamics of viral circulation, and the varying susceptibility and competence of different avian species [14, 18, 21]. Surveillance strategies can be broadly categorized into passive (opportunistic) and active (targeted) approaches, each with distinct advantages, limitations, and optimal applications within a One Health framework. The World Health Organization (WHO) and the World Organisation for Animal Health (WOAH) have recognized the importance of integrating animal and human surveillance to detect emerging arboviral threats, and wild bird monitoring programs have proven instrumental in providing early warning signals for impending human outbreaks [15, 32].

Passive Surveillance: Leveraging Wildlife Rehabilitation Centers and Opportunistic Sampling

Passive surveillance relies on the collection and testing of samples from birds that are found dead, moribund, or admitted to wildlife rehabilitation centers (WRCs). This approach capitalizes on the natural filtering effect of disease-induced mortality, which tends to concentrate sampling efforts on individuals that have succumbed to acute infection or those showing clinical signs compatible with WNV neuroinvasive disease [1, 40, 41]. WRCs are particularly valuable sentinel systems because they concentrate a diverse array of avian species from surrounding peri-urban and urban environments, often including those species most likely to encounter bridge vectors that transmit WNV to humans and equids [1, 9, 38]. Long-term passive surveillance in central Spain over a nine-year period (2013–2022) involving 1024 bird samples from WRCs failed to detect any WNV RNA, despite testing oropharyngeal and cloacal swabs and tissue samples by real-time PCR targeting the 3′ non-coding region [1]. The authors rightly noted that the variability inherent to passive surveillance, including heterogeneity in sample type, timing of collection relative to infection, and the condition of carcasses, can substantially reduce detection sensitivity [1]. Nevertheless, the continued monitoring of WRCs remains essential, particularly for detecting spillover events from enzootic cycles into high-risk avian species and for assessing the occupational exposure risk to personnel handling infected birds [1, 40].

Post-mortem monitoring studies in central Italy, spanning November 2017 to October 2020, tested 576 wild birds from five WRCs for WNV and Usutu virus (USUV) RNA [40, 41]. Only one bird, an adult little grebe (Tachybaptus ruficollis), tested positive for WNV RNA with a high cycle threshold (Ct = 34.36), and no USUV RNA was detected [40, 41]. The presence of WNV RNA in December, a month well outside the typical mosquito vector season, raises important questions about possible overwintering mechanisms in birds, a phenomenon that has been suggested by other European studies as well [8, 46]. The pathological examination of the positive little grebe revealed mild CD3+ lymphocytic tubulo-interstitial nephritis, meningoencephalitis, and myocardial lesions, consistent with acute flaviviral infection [40, 41]. This case underscores the value of combining molecular diagnostics with histopathological assessments in passive surveillance, as the detection of viral RNA in brain or kidney tissue may persist longer than in blood, thereby extending the diagnostic window [20, 40, 41]. Similarly, in Extremadura, western Spain, passive surveillance of 391 birds from two WRCs between 2017 and 2019 detected WNV-specific antibodies in 18.23% of birds across 18 species, with high seroprevalences in Pelecaniformes (33.33%), Accipitriformes (25.77%), and Strigiformes (22.92%) [9]. Notably, this study provided the first European detection of WNV antibodies in a black stork (Ciconia nigra), and sequential virus neutralization tests (VNT) revealed USUV-specific antibodies in four birds, indicating the co-circulation of multiple flaviviruses in the region [9].

The value of passive surveillance for detecting emerging WNV foci is illustrated by the first detection of WNV in Belgium in August 2025 through a wild bird monitoring program [15]. Three corvids tested positive by RT-qPCR, with four additional infected birds identified in September and October. Whole genome sequencing classified the strain as WNV lineage 2, consistent with strains circulating in other European countries, and this detection provided critical evidence of local WNV circulation with direct implications for public health preparedness [15]. In contrast, passive surveillance in South Korea from 2005 to 2008, testing dead wild birds nationwide, found no evidence of WNV activity, suggesting that the virus had not yet been introduced into the Korean peninsula during that period [55]. These contrasting outcomes highlight the importance of geographic context: passive surveillance is most effective in regions where WNV is known or suspected to be circulating, but may lack sensitivity for detecting early incursions if mortality rates are low or if affected species are not readily encountered by the surveillance system [54, 55].

One significant limitation of passive surveillance is its inherent bias toward clinically affected birds, which may represent only a small fraction of infected individuals. Many avian species, particularly passerines, can sustain high-titer viremias sufficient to infect feeding mosquitoes without showing any overt signs of disease [21, 38]. Furthermore, the detection of WNV RNA in dead birds may be compromised by rapid RNA degradation post-mortem, especially in warm climates [45]. Despite these limitations, passive surveillance remains a cost-effective approach for detecting the presence of highly pathogenic viral strains and for identifying species that are particularly susceptible to severe disease, such as raptors and corvids [13, 38, 50].

Active Surveillance: Targeted Trapping, Serosurveys, and Longitudinal Monitoring

Active surveillance involves the systematic capture, sampling, and release of wild birds, often targeting specific species, age classes, or ecological guilds to assess the prevalence of WNV infection or exposure [2, 7, 10, 43]. This approach provides a more representative picture of viral circulation within avian communities, as it samples both asymptomatic and symptomatic individuals and allows for the estimation of seroprevalence, seroconversion rates, and the identification of demographic or ecological risk factors [2, 7, 10, 11, 23]. A landmark active surveillance program in Ukraine during 2023–2024 collected 268 blood samples and 9 egg yolks from wild Passeriformes and Piciformes across five regions [2]. Using a commercial competitive ELISA (ID Screen West Nile), the study found that seroprevalence varied dramatically by species and region: Great Tits showed seroprevalences ranging from 20% to 100%, Blackbirds from 93% to 100%, and House Sparrows 100% in some locations [2]. Among the highest seroprevalences were found in the Poltava region (86%) and Khmelnytsky region (67%), while the lowest were in Kyiv (9%) and Odesa (17.1%) [2]. Importantly, seropositivity increased from 27.4% in 2023 to 50.5% in 2024, suggesting a rapidly intensifying transmission cycle that closely paralleled the increase in human cases and fatalities reported in Ukraine during 2024 [2]. No antibodies were detected in swallows, sedge warblers, or woodpeckers, indicating that not all avian species are equally exposed or susceptible [2].

In Bangladesh, a study of 184 wild birds (19 species, nine families) collected between 2012 and 2016 found an overall seroprevalence of 11.9% by c-ELISA, with higher rates in migratory birds (15.9%) than in resident birds (10.7%) [3]. The migratory tufted duck (Aythya fuligula) showed 28.5% seropositivity, while resident house crows (Corvus splendens) had 12.5% seropositivity [3]. None of the tracheal or cloacal swabs tested positive for flavivirus RNA, suggesting that active viral shedding is transient and that detection of nucleic acid requires careful timing relative to the peak of viremia [3]. Another larger study in Bangladesh (888 birds) found a lower overall seroprevalence of 5.4%, but confirmed that both resident and migratory species are exposed to WNV [53]. In Malaysia, active surveillance of 260 wild birds (163 migratory, 97 resident) in the West Coast of Peninsular Malaysia detected WNV antibodies in 18.71% of tested sera and molecular evidence of WNV lineage 2 RNA in 15.2% of oropharyngeal swabs [12]. Phylogenetic analysis of the 16 isolates revealed 99% similarity to strains from South Africa, suggesting that migratory birds may have introduced WNV lineage 2 into Malaysia [12]. Critically, the detection of WNV RNA in resident birds indicates that local transmission cycles have become established, with potential for spillover into the human population [12].

Active surveillance programs that incorporate serological and molecular testing of the same individuals provide the most comprehensive assessment of WNV exposure and infection. In Germany, a nationwide wild bird surveillance network monitored migratory and resident birds from 2014 to 2016 [54]. No WNV-specific RNA was detected in 1900+ blood samples, but WNV neutralizing antibodies were found predominantly in long-distance, partial, and short-distance migrants, while USUV neutralizing antibodies were found mainly in resident species [54]. Subsequent monitoring in 2017–2018 revealed the first signs of local WNV circulation in eastern Germany, where high WNV-specific neutralizing antibody titers were found in resident and short-distance migratory birds [10]. By 2019–2020, WNV seroprevalence in eastern Germany had reached 14.77–16.15%, with a clearly observable westward and southward expansion over time [51]. This temporal progression, documented through systematic active surveillance, has been instrumental in tracking the invasion and establishment of WNV in a previously naive region [10, 51, 54]. The emergence of WNV in Germany was confirmed in 2023–2024, when whole genome sequencing of 86 positive bird samples revealed that approximately 73% belonged to subcluster 2.5.3.4.3c, the dominant circulating variant, while the remaining 27% grouped within cluster 2.5.3.2, demonstrating the co-circulation of at least two genetically distinct WNV-2 variants [17]. These findings highlight the ongoing geographic expansion and genetic diversification of WNV in Germany, seven years after its initial detection [17].

Longitudinal active surveillance studies that recapture individual birds over multiple seasons provide unique insights into the dynamics of WNV exposure, including seroconversion, seroreversion, and the duration of protective immunity. A study of common coots (Fulica atra) in Doñana, Spain, conducted from 2003 to 2005, found that antibody prevalence was highest in 2003, intermediate in 2004, and lowest in 2005, with some birds seroreverting to negative status within one year of first capture [42]. This observation suggests that WNV antibody titers in some avian species may wane over time, potentially rendering previously exposed birds susceptible to reinfection [42]. Similarly, a multi-year comparison of WNV antibody prevalence in birds from Atlanta, Georgia, and Chicago, Illinois (2005–2016), found that seroprevalence was highest in year-round and summer-resident species compared with migrants, and that Northern cardinals (Cardinalis cardinalis) were the species most likely to test positive in both cities [35]. Seroprevalence was higher in Atlanta (13.7%) than in Chicago (5%), and year-to-year variation was more pronounced in Chicago, suggesting that environmental and ecological factors exert strong influences on transmission intensity at the local scale [35]. These longitudinal data are crucial for parameterizing mathematical models that predict the risk of human outbreaks based on preceding avian seroprevalence levels [7, 11, 18].

Sentinel Species Selection: Raptors, Corvids, and Ecological Correlates of WNV Exposure

The selection of appropriate sentinel species is critical for the cost-effectiveness and predictive power of WNV surveillance programs. Not all birds contribute equally to WNV amplification or serve as reliable indicators of local transmission. Raptors (birds of prey) have emerged as particularly sensitive sentinels because they often develop high-titer viremias, exhibit clinical signs that bring them to the attention of wildlife rehabilitators, and occupy high trophic levels that may expose them to infected prey [9, 38, 51]. In Spain, following the major 2020 WNV outbreak, IgG antibodies against flaviviruses were detected in 32.7% of 171 wild birds by bELISA, with VNT-confirmed WNV seroprevalence of 19.3% [25]. Species group (raptors), age (greater than one year old), and body size (large) were identified as the main risk factors for seropositivity, emphasizing the utility of raptors as sentinels [25]. In Serbia, molecular surveillance of 133 wild birds during 2012 identified WNV RNA in nine birds, including Northern Goshawks (Accipiter gentilis), White-tailed Eagles (Haliaeetus albicilla), and a Hooded Crow (Corvus cornix) [24]. The detection of WNV lineage 2 in these raptors signaled the establishment of an enzootic cycle that subsequently led to human outbreaks in the region [24]. Similarly, in Greece, molecular detection of WNV lineage 2 in a hunter-harvested Eurasian Magpie (Pica pica) in 2010 provided the first evidence of WNV in sedentary wild birds during a large human outbreak, and the virus was subsequently isolated from goshawks and other raptors [8, 22, 36]. Magpies have been proposed as particularly useful sentinels in Mediterranean ecosystems, as they are common, sedentary, and frequently exposed to Culex mosquitoes [8].

Corvids (crows, magpies, jays) are widely recognized as highly competent amplifying hosts for WNV, often developing exceptionally high viremias that efficiently infect feeding mosquitoes [15, 24, 50]. In Israel, testing of 136 frozen wild bird carcasses in the months preceding the 2018 human outbreak detected WNV RNA in 11.03% of tissue pools, with all positive samples belonging to WNV lineage 1, cluster 2 eastern European [50]. The majority of infected birds were local corvid species, suggesting that local enzootic amplification, rather than introduction by migrants, was the primary driver of the outbreak [50]. In Belgium, the first detection of WNV in 2025 occurred in corvids, underscoring their role as early warning sentinels [15]. The Dutch multidisciplinary surveillance program, which successfully identified the first autochthonous WNV cases in the Netherlands in 2020, relied heavily on wild live bird research surveys targeting corvids and other passerines, followed by mosquito research and sentinel chicken monitoring [32]. The Netherlands program highlighted that real-time testing of birds provided an early warning of 35 days before the first human case was identified, demonstrating the practical utility of avian sentinel surveillance [32].

Species-specific differences in exposure and competence are influenced by ecological and behavioral traits. A comprehensive study of the bird reservoir potential for WNV across Europe utilized a bird traits-based model to estimate WNV seroprevalence in 150 European passerine species, generating a WNV Reservoir Index (RI) that was mapped across the continent [18]. The RI showed strong spatial heterogeneity, with high values in Central and Eastern Europe, and was positively associated with the number of years with notified human WNV cases at the NUTS administrative region scale [18]. Eco-ethological characteristics associated with higher seroprevalence included habitat preferences (wetlands, agricultural areas), gregariousness, and exposure to ornithophilic mosquitoes [18]. In southwestern Spain, analysis of 1194 serum samples from 44 avian species found that migratory birds had higher WNV exposure likelihood than native and exotic species, and that higher phylogenetic diversity in bird communities correlated with lower exposure rates, a pattern consistent with the dilution effect hypothesis [23]. Moreover, birds with WNV antibodies were positively associated with the abundance of competent vectors (Culex pipiens s.l. and the Univittatus subgroup) but negatively associated with overall mosquito species richness, suggesting that vector community composition is a strong driver of transmission risk [11, 23].

Molecular and Serological Tools: Enhancing Detection Sensitivity and Specificity

The choice of diagnostic tools profoundly influences the sensitivity, specificity, and interpretability of WNV surveillance data. Molecular detection by reverse transcription quantitative PCR (RT-qPCR) targeting conserved regions of the flaviviral genome (e.g., the 3′ non-coding region, the envelope gene, or NS2A/NS3 regions) allows for the direct detection of viral RNA, providing evidence of active infection [1, 4, 8, 12]. However, the window for RNA detection in blood is brief, typically lasting only a few days during peak viremia, after which the virus may be cleared or sequestered in tissues [3, 21]. In Bangladesh, none of 184 swab samples tested positive for flavivirus RNA despite a seroprevalence of 11.9%, illustrating the challenge of detecting acute infections in cross-sectional sampling [3]. Conversely, testing of visceral tissues (e.g., spleen, kidney, brain, myocardium) from dead birds can extend the detection window, as viral RNA may persist for weeks in these compartments [40, 41, 46]. In northern Italy, sampling of brain, myocardium, kidney, and spleen from 164 wild birds during the off-season (October to May) revealed WNV RNA in 4 birds (2.44%) and USUV RNA in 5 birds (3.05%), demonstrating that tissue-based molecular surveillance can detect viral persistence even during periods of low vector activity [46].

Serological assays, including enzyme-linked immunosorbent assays (ELISA) and virus neutralization tests (VNT), provide evidence of past exposure and are particularly useful for estimating cumulative incidence and identifying high-risk species [2, 7, 9, 19, 49]. The ID Screen West Nile Competition Multi-species ELISA is widely used for screening because it detects antibodies against flaviviral envelope proteins across a broad range of avian species [2, 7, 25]. However, cross-reactivity among flaviviruses (WNV, USUV, St. Louis encephalitis virus [SLEV], Japanese

References

[1] Ayllón T, Martínez I, Ortiz-Díez G, Navarro A, Fúster F, Iriso A, et al.. Long-Term Surveillance of Chlamydia psittaci and West Nile Virus in Wild Birds from Central Spain (2013–2022). Microorganisms. 2025. DOI: https://doi.org/10.3390/microorganisms14010048

[2] Muzyka D, Popova A. Serological studies on the presence of antibodies against West Nile virus in wild birds of the order Passeriformes in Ukraine. Veterinary Medicine: inter-departmental subject scientific collection. 2024. DOI: https://doi.org/10.36016/vm-2024-110-3

[3] Islam A, Islam S, Hossain ME, Ferdous J, Abedin J, Rahman MZ, et al.. Serological Evidence of West Nile Virus in Wild Birds in Bangladesh. Veterinary Sciences. 2020. DOI: https://doi.org/10.3390/vetsci7040164

[4] Nyamwaya D, Wang'ondu V, Amimo J, Michuki G, Ogugo M, Ontiri E, et al.. Detection of West Nile virus in wild birds in Tana River and Garissa Counties, Kenya. BMC Infectious Diseases. 2016. DOI: https://doi.org/10.1186/s12879-016-2019-8

[5] Schwinn A, Harris S, Jacobs Z, Verges JEd, Jameson SB, Wesson DM, et al.. Immune Responses Against West Nile Virus and Mosquito Salivary Proteins in Wild Birds from St. Tammany Parish, Louisiana. Zoonotic Diseases. 2025. DOI: https://doi.org/10.3390/zoonoticdis5020011

[6] Bosco-Lauth A, Kooi K, Hawks SA, Duggal NK. Cross-Protection between West Nile Virus and Emerging Flaviviruses in Wild Birds. American Journal of Tropical Medicine and Hygiene. 2024. DOI: https://doi.org/10.4269/ajtmh.24-0363

[7] Williams RAJ, Valencia HAC, Márquez IL, González FG, Llorente F, Jiménez-Clavero MÁ, et al.. West Nile Virus Seroprevalence in Wild Birds and Equines in Madrid Province, Spain. Veterinary Sciences. 2024. DOI: https://doi.org/10.3390/vetsci11060259

[8] Athanasakopoulou Z, Sofia M, Skampardonis V, Giannakopoulos A, Birtsas P, Tsolakos K, et al.. Indication of West Nile Virus (WNV) Lineage 2 Overwintering among Wild Birds in the Regions of Peloponnese and Western Greece. Veterinary Sciences. 2023. DOI: https://doi.org/10.3390/vetsci10110661

[9] Bravo-Barriga D, Aguilera-Sepúlveda P, Guerrero-Carvajal F, Llorente F, Reina D, Pérez-Martín JE, et al.. West Nile and Usutu virus infections in wild birds admitted to rehabilitation centres in Extremadura, western Spain, 2017-2019.. Veterinary Microbiology. 2021. DOI: https://doi.org/10.1016/j.vetmic.2021.109020

[10] Michel F, Sieg M, Fischer D, Keller M, Eiden M, Reuschel M, et al.. Evidence for West Nile Virus and Usutu Virus Infections in Wild and Resident Birds in Germany, 2017 and 2018. Viruses. 2019. DOI: https://doi.org/10.3390/v11070674

[11] Puente JMl, Ferraguti M, Ruíz S, Roiz D, Llorente F, Pérez-Ramírez E, et al.. Mosquito community influences West Nile virus seroprevalence in wild birds: implications for the risk of spillover into human populations. Scientific Reports. 2018. DOI: https://doi.org/10.1038/s41598-018-20825-z

[12] Ain-Najwa MY, Yasmin AR, Omar A, Arshad S, Abu J, Mohammed H, et al.. Evidence of West Nile virus infection in migratory and resident wild birds in west coast of peninsular Malaysia. One Health. 2020. DOI: https://doi.org/10.1016/j.onehlt.2020.100134

[13] Barbachano-Guerrero A, Vásquez-Aguilar A, Aguirre AA, Zavala‐Norzagaray A, Gonzalez EC, Terrazas AL, et al.. West Nile Virus Prevalence in Wild Birds from Mexico. Journal of Wildlife Diseases. 2019. DOI: https://doi.org/10.7589/2018-03-065

[14] Bruno L, Nappo M, Frontoso R, Perrotta MG, Lecce RD, Guarnieri C, et al.. West Nile Virus (WNV): One-Health and Eco-Health Global Risks. Veterinary Sciences. 2025. DOI: https://doi.org/10.3390/vetsci12030288

[15] Sohier C, Breman F, Vervaeke M, Sikkema R, Boter M, Munnink BOO, et al.. First detection of West Nile virus in Belgium through wild bird surveillance, Belgium, 2025. Euro surveillance : bulletin Europeen sur les maladies transmissibles = European communicable disease bulletin. 2026. DOI: https://doi.org/10.2807/1560-7917.ES.2026.31.4.2600049

[16] Sohier C, Breman F, Vervaeke M, Regge ND. West Nile Virus Monitoring in Flanders (Belgium) During 2022–2023 Reveals Endemic Usutu Virus Circulation in Birds. Transboundary and Emerging Diseases. 2024. DOI: https://doi.org/10.1155/tbed/4146156

[17] Schwarzer A, Schopf F, Groschup M, Bock S, Heenemann K, Herms L, et al.. Circulation dynamics of West Nile virus in Germany, 2023 and 2024. Virology Journal. 2025. DOI: https://doi.org/10.1186/s12985-025-03043-8

[18] Bastard J, Metras R, Durand B. Mapping the bird reservoir potential for West Nile virus in Europe and its relationship with disease occurrence in humans. medRxiv. 2025. DOI: https://doi.org/10.1101/2025.02.24.25322781

[19] Niczyporuk J, Samorek-Salamonowicz E, Lecollinet S, Pancewicz S, Kozdruń W, Czekaj H. Occurrence of West Nile Virus Antibodies in Wild Birds, Horses, and Humans in Poland. BioMed Research International. 2015. DOI: https://doi.org/10.1155/2015/234181

[20] Agliani G, Giglia G, Marshall EM, Gröne A, Rockx B, Brand JVDvd. Pathological features of West Nile and Usutu virus natural infections in wild and domestic animals and in humans: A comparative review. One Health. 2023. DOI: https://doi.org/10.1016/j.onehlt.2023.100525

[21] Pérez-Ramírez E, Llorente F, Jiménez-Clavero MÁ. Experimental Infections of Wild Birds with West Nile Virus. Viruses. 2014. DOI: https://doi.org/10.3390/v6020752

[22] Valiakos G, Touloudi A, Athanasiou L, Giannakopoulos A, Iacovakis C, Birtsas P, et al.. Serological and molecular investigation into the role of wild birds in the epidemiology of West Nile virus in Greece. Virology Journal. 2012. DOI: https://doi.org/10.1186/1743-422X-9-266

[23] Ferraguti M, Magallanes S, Mora-Rubio C, Bravo-Barriga D, Marzal A, Hernandez-Caballero I, et al.. Implications of migratory and exotic birds and the mosquito community on West Nile virus transmission. Infectious Diseases. 2023. DOI: https://doi.org/10.1080/23744235.2023.2288614

[24] Petrović T, Blázquez A, Lupulović D, Lazić G, Escribano-Romero E, Fabijan D, et al.. Monitoring West Nile virus (WNV) infection in wild birds in Serbia during 2012: first isolation and characterisation of WNV strains from Serbia.. Euro surveillance : bulletin Europeen sur les maladies transmissibles = European communicable disease bulletin. 2013. DOI: https://doi.org/10.2807/1560-7917.ES2013.18.44.20622

[25] García‐Bocanegra I, Franco JJ, León CI, Barbero-Moyano J, García-Miña MV, Fernández-Molera V, et al.. High exposure of West Nile virus in equid and wild bird populations in Spain following the epidemic outbreak in 2020. Transboundary and Emerging Diseases. 2022. DOI: https://doi.org/10.1111/tbed.14733

[26] Bergmann F, Schmoock-Wellhausen M, Fast C, Holicki CM, Michel F, Wysocki P, et al.. Longitudinal Study of the Occurrence of Usutu Virus and West Nile Virus Infections in Birds in a Zoological Garden in Northern Germany. Pathogens. 2023. DOI: https://doi.org/10.3390/pathogens12060753

[27] Maroco D, Parreira R, Santos FAd, Lopes Â, Simões F, Orge L, et al.. Tracking the Pathways of West Nile Virus: Phylogenetic and Phylogeographic Analysis of a 2024 Isolate from Portugal. Microorganisms. 2025. DOI: https://doi.org/10.3390/microorganisms13030585

[28] Hoxha I, Xhekaj B, Muja-Bajraktari N, Sekulin K, Unterköfler MS, Schlamadinger L, et al.. First Detection of West Nile Virus (WNV) Lineage 2 in Mosquitoes in the Republic of Kosovo. Transboundary and Emerging Diseases. 2025. DOI: https://doi.org/10.1155/tbed/3208806

[29] Ventura Cd, Carrera M, Defilippo F, Lelli D, Nogarol C, Mandola M, et al.. Genomic epidemiology and phylogeographic reconstruction of West Nile virus 2 in Italy from 2011 to 2023. One Health. 2025. DOI: https://doi.org/10.1016/j.onehlt.2025.101310

[30] Sofia M, Giannakopoulos A, Giantsis I, Touloudi A, Birtsas P, Papageorgiou K, et al.. West Nile Virus Occurrence and Ecological Niche Modeling in Wild Bird Species and Mosquito Vectors: An Active Surveillance Program in the Peloponnese Region of Greece. Microorganisms. 2022. DOI: https://doi.org/10.3390/microorganisms10071328

[31] Rajeendran A, Ballakrishnan N, Tulis AN, Zameer M, Selvia L, Shakawi A, et al.. Prevalence of West Nile Virus in Zoonotic Animal Species in Asia (2000–2024): A Systematic Review and Meta-Analysis. Journal of Tropical Resources and Sustainable Science (JTRSS). 2026. DOI: https://doi.org/10.47253/jtrss.v14i1.1621

[32] Best PAd, Braks M, Timen A, Sikkema R, Koopmans M. A multidisciplinary approach to the detection of and response to West Nile virus in the Netherlands between 2020 and 2023: best practices, challenges and opportunities. Euro surveillance : bulletin Europeen sur les maladies transmissibles = European communicable disease bulletin. 2026. DOI: https://doi.org/10.2807/1560-7917.ES.2026.31.10.2500276

[33] Aguilera-Sepúlveda P, Cano-Gómez C, Villalba R, Borges V, Agüero M, Bravo-Barriga D, et al.. The key role of Spain in the traffic of West Nile virus lineage 1 strains between Europe and Africa. Infectious Diseases. 2024. DOI: https://doi.org/10.1080/23744235.2024.2348633

[34] Kariwa H, Murata R, Totani M, Yoshii K, Takashima I. Increased Pathogenicity of West Nile Virus (WNV) by Glycosylation of Envelope Protein and Seroprevalence of WNV in Wild Birds in Far Eastern Russia. International Journal of Environmental Research and Public Health. 2013. DOI: https://doi.org/10.3390/ijerph10127144

[35] McMillan JR, Hamer G, Levine R, Mead D, Waller L, Goldberg T, et al.. Multi-Year Comparison of Community- and Species-Level West Nile Virus Antibody Prevalence in Birds from Atlanta, Georgia and Chicago, Illinois, 2005–2016. American Journal of Tropical Medicine and Hygiene. 2022. DOI: https://doi.org/10.4269/ajtmh.21-1086

[36] Valiakos G, Valiakos G, Touloudi A, Touloudi A, Iacovakis C, Iacovakis C, et al.. Molecular detection and phylogenetic analysis of West Nile virus lineage 2 in sedentary wild birds (Eurasian magpie), Greece, 2010.. Euro surveillance : bulletin Europeen sur les maladies transmissibles = European communicable disease bulletin. 2011. DOI: https://doi.org/10.2807/ESE.16.18.19862-EN

[37] Fiacre L, Nougairède A, Migné C, Bayet M, Cochin M, Dumarest M, et al.. Different viral genes modulate virulence in model mammal hosts and Culex pipiens vector competence in Mediterranean basin lineage 1 West Nile virus strains. Frontiers in Microbiology. 2024. DOI: https://doi.org/10.3389/fmicb.2023.1324069

[38] Vidaña B, Busquets N, Napp S, Pérez-Ramírez E, Jiménez-Clavero MÁ, Johnson N. The Role of Birds of Prey in West Nile Virus Epidemiology. Vaccines. 2020. DOI: https://doi.org/10.3390/vaccines8030550

[39] Ruiz-López M, Muñoz-Chimeno M, Figuerola J, Gavilán A, Varona S, Cuesta I, et al.. Genomic Analysis of West Nile Virus Lineage 1 Detected in Mosquitoes during the 2020–2021 Outbreaks in Andalusia, Spain. Viruses. 2023. DOI: https://doi.org/10.3390/v15020266

[40] Giglia G, Mencattelli G, Lepri E, Agliani G, Gobbi M, Gröne A, et al.. West Nile virus and Usutu virus in wild birds from Rescue Centers, a post-mortem monitoring study from Central Italy. bioRxiv. 2022. DOI: https://doi.org/10.1101/2022.07.19.500416

[41] Giglia G, Mencattelli G, Lepri E, Agliani G, Gobbi M, Gröne A, et al.. West Nile Virus and Usutu Virus: A Post-Mortem Monitoring Study in Wild Birds from Rescue Centers, Central Italy. Viruses. 2022. DOI: https://doi.org/10.3390/v14091994

[42] Figuerola J, Soriguer R, Rojo G, Tejedor CG, Jiménez-Clavero MÁ. Seroconversion in Wild Birds and Local Circulation of West Nile Virus, Spain. Emerging Infectious Diseases. 2007. DOI: https://doi.org/10.3201/eid1312.070343

[43] Casades-Martí L, Holgado-Martín R, Aguilera-Sepúlveda P, Llorente F, Pérez-Ramírez E, Jiménez-Clavero MÁ, et al.. Risk Factors for Exposure of Wild Birds to West Nile Virus in A Gradient of Wildlife-Livestock Interaction. Pathogens. 2023. DOI: https://doi.org/10.3390/pathogens12010083

[44] Pallari C, Efstathiou A, Moysi M, Papanikolas N, Christodoulou V, Mazeris A, et al.. Evidence of West Nile virus seropositivity in wild birds on the island of Cyprus.. Comparative Immunology, Microbiology & Infectious Diseases. 2020. DOI: https://doi.org/10.1016/j.cimid.2020.101592

[45] Fasuan OS, Faneye DO, Adeyanju T. Attempted Detection of West Nile Virus from Wild and Peridomestic Birds within Ibadan Metropolis in Nigeria. International Journal of Pathogen Research. 2019. DOI: https://doi.org/10.9734/ijpr/2019/v2i129625

[46] Musto C, Tamba M, Calzolari M, Rossi A, Grisendi A, Marzani K, et al.. Detection of West Nile and Usutu Virus RNA in Autumn Season in Wild Avian Hosts in Northern Italy. Viruses. 2023. DOI: https://doi.org/10.3390/v15081771

[47] Figuerola J, Baouab RE, Soriguer R, Fassi-Fihri O, Llorente F, Jiménez-Clavero MÁ. West Nile Virus Antibodies in Wild Birds, Morocco, 2008. Emerging Infectious Diseases. 2009. DOI: https://doi.org/10.3201/eid1510.090340

[48] Csank T, Korytár L, Pošiváková T, Bakonyi T, Pistl J, Csanády A. Surveillance on antibodies against West Nile virus, Usutu virus, tick-borne encephalitis virus and Tribeč virus in wild birds in Drienovská wetland, Slovakia. Biologia. 2019. DOI: https://doi.org/10.2478/s11756-019-00211-4

[49] Medrouh B, Lafri I, Beck C, Leulmi H, Akkou M, Abbad L, et al.. First serological evidence of West Nile virus infection in wild birds in Northern Algeria.. Comparative Immunology, Microbiology & Infectious Diseases. 2020. DOI: https://doi.org/10.1016/j.cimid.2020.101415

[50] Schvartz G, Tirosh-Levy S, Bider S, Lublin A, Farnoushi Y, Erster O, et al.. West Nile Virus in Common Wild Avian Species in Israel. Pathogens. 2022. DOI: https://doi.org/10.3390/pathogens11010107

[51] Ziegler U, Bergmann F, Fischer D, Müller K, Holicki CM, Sadeghi B, et al.. Spread of West Nile Virus and Usutu Virus in the German Bird Population, 2019–2020. Microorganisms. 2022. DOI: https://doi.org/10.3390/microorganisms10040807

[52] Vasić A, Oșlobanu L, Marinov M, Crivei L, Rățoi I, Aniță A, et al.. Evidence of West Nile Virus (WNV) Circulation in Wild Birds and WNV RNA Negativity in Mosquitoes of the Danube Delta Biosphere Reserve, Romania, 2016. Tropical Medicine and Infectious Disease. 2019. DOI: https://doi.org/10.3390/tropicalmed4030116

[53] Islam A, Hossain ME, Rahman MH, Islam A, Rahman A, Paul S, et al.. Sero-prevalence of West Nile virus in Wild Birds in Bangladesh. . 2020. DOI: https://doi.org/10.22541/au.159707702.21777108

[54] Michel F, Fischer D, Eiden M, Fast C, Reuschel M, Müller K, et al.. West Nile Virus and Usutu Virus Monitoring of Wild Birds in Germany. International Journal of Environmental Research and Public Health. 2018. DOI: https://doi.org/10.3390/ijerph15010171

[55] Yeh J, Kim H, Nah J, Lee H, Kim Y, Moon J, et al.. Surveillance for West Nile Virus in Dead Wild Birds, South Korea, 2005–2008. Emerging Infectious Diseases. 2011. DOI: https://doi.org/10.3201/eid1702.100551