Getah Virus

Overview and Taxonomy of Getah Virus (GETV)

Getah virus (GETV) is a re-emerging, mosquito-borne, single-stranded, positive-sense RNA virus classified within the genus Alphavirus, family Togaviridae [16, 28, 38]. The virus was first isolated from Culex gelidus mosquitoes collected in a mangrove forest near Getah, Malaysia, in 1955, a location that provides the etymological basis for its name [16, 27]. For decades following its initial discovery, GETV was considered a relatively obscure pathogen, circulating primarily in a sylvatic cycle between mosquitoes and vertebrate hosts in tropical and subtropical regions of Southeast Asia and Oceania [16]. However, since the turn of the 21st century, GETV has undergone a dramatic epidemiological shift, expanding its geographical range, host tropism, and pathogenic potential, transforming from a neglected arbovirus into a significant threat to livestock industries and a growing concern for public health [1, 6, 16, 38]. This section provides a comprehensive overview of the virus, detailing its taxonomic position, virion structure, genomic organization, and the phylogenetic framework that underpins our understanding of its global spread and evolution.

Taxonomic Classification and Phylogenetic Lineages

GETV belongs to the Alphavirus genus, a large and diverse group of arthropod-borne viruses (arboviruses) that are further subdivided into seven antigenic complexes based on serological cross-reactivity [16, 28]. GETV is a member of the Semliki Forest virus (SFV) antigenic complex, which also includes other notable arthritogenic alphaviruses such as Ross River virus (RRV), Chikungunya virus (CHIKV), and O’nyong’nyong virus (ONNV) [13, 16, 36]. This classification is supported by both serological and phylogenetic analyses, which consistently place GETV within a clade of viruses that typically cause febrile illness, rash, and polyarthritis in infected hosts [13, 16]. The close antigenic relationship between GETV and RRV, in particular, has been well-documented, with convalescent sera from mice infected with either virus demonstrating high levels of cross-reactivity and cross-protection [36]. This serological overlap has historically complicated serosurveys, especially in regions like Australia where RRV is endemic, and has led to debates regarding the true historical presence of GETV on the continent [36].

Phylogenetic analyses of complete genome sequences and, more commonly, the E2 and E1 envelope glycoprotein genes, have robustly delineated GETV into four distinct genotypes or groups: Group I (GI), Group II (GII), Group III (GIII), and Group IV (GIV) [3, 23, 27, 28, 35]. The prototype strain, MM2021 isolated in Malaysia in 1955, serves as the archetype for Group I [27, 35]. Group II comprises a small number of older isolates, while Group IV includes strains from Russia, Thailand, and a more recent isolate from Malaysia (B254) [27, 29, 35]. Critically, the vast majority of contemporary GETV isolates, including virtually all strains responsible for recent epidemics and outbreaks in livestock across Asia, belong to Group III [1-3, 7, 10, 11, 23, 24, 31, 34]. This group has become the dominant and widely circulating lineage, effectively replacing the older genotypes in the field [3, 23]. The emergence and dominance of GIII strains are associated with a higher replication capacity in both mammalian and mosquito cells compared to non-epidemic strains, such as GIV B254, suggesting that enhanced viral fitness is a key determinant of its epidemic potential [35]. The global expansion of GIII GETV is a relatively recent phenomenon, with phylodynamic analyses indicating a rapid evolutionary rate and a population expansion that correlates with increased livestock trade and movement [6, 23, 28].

Virion Structure and Genomic Organization

The GETV virion is a spherical, enveloped particle with a diameter of approximately 60–70 nm, as visualized by cryo-electron microscopy (cryo-EM) and ultra-thin sectioning [25, 26, 32, 34]. The virus exhibits the characteristic alphavirus architecture: a nucleocapsid core, composed of the capsid (C) protein and the single-stranded, positive-sense genomic RNA, is surrounded by a host-derived lipid bilayer [25, 26]. Embedded within this envelope are 80 trimeric spikes, each consisting of three heterodimers of the E1 and E2 transmembrane glycoproteins [25, 26]. The E2 glycoprotein is the major antigenic determinant and is primarily responsible for receptor binding and host cell attachment, while the E1 glycoprotein mediates the low pH-dependent fusion of the viral envelope with the endosomal membrane during viral entry [8, 14, 25, 26]. The high-resolution cryo-EM structure of the infectious GETV virion at 2.8 Å has revealed critical details, including the presence of a "pocket factor" (likely a phospholipid) within the E1/E2 heterodimer that stabilizes the metastable prefusion conformation, and the identification of specific N-glycosylation sites (e.g., E1 N141, E2 N200, E2 N262) and S-acylation sites that are crucial for viral immune evasion, host cell invasion, and the stabilization of transmembrane protein assembly [25, 26].

The GETV genome is a single-stranded, positive-sense RNA molecule of approximately 11.6–11.7 kb in length, excluding the 5' cap and 3' poly-A tail [10, 29, 34]. The genomic organization is typical of alphaviruses, comprising two open reading frames (ORFs). The 5' two-thirds of the genome encodes a large polyprotein that is proteolytically cleaved into four nonstructural proteins (nsP1, nsP2, nsP3, and nsP4), which form the viral replicase complex essential for RNA replication and transcription [4, 12, 40]. The nsP3 protein, in particular, is a multifunctional protein containing a macro domain (involved in ADP-ribose binding and de-ADP-ribosylation), a zinc-binding domain, and a hypervariable domain (HVD) [12, 40]. The GETV nsP2 has been identified as a potent antagonist of the host type I interferon (IFN-I) response, specifically suppressing IFN-β production by binding to TBK1 and inhibiting IRF3 phosphorylation and nuclear translocation [4]. The nsP3 HVD has been shown to block the formation of bona fide stress granules by binding to the host protein G3BP, a strategy that subverts the cellular stress response to favor viral replication [12]. The 3' one-third of the genome is transcribed from a subgenomic promoter into a 26S mRNA that encodes the structural polyprotein, which is cleaved into the capsid (C) protein, the E3 and E2 glycoproteins (with E3 acting as a chaperone for E2 folding), the 6K protein (a small, hydrophobic protein involved in virion assembly and egress), and the E1 glycoprotein [22, 25, 26]. The 6K protein, while not essential for virion production in all cell types, plays a role in the efficient release of viral particles from host cells and contributes to the clinical manifestation of disease in vivo [22].

Genetic Diversity, Adaptive Evolution, and Emerging Variants

The rapid evolution of GETV, particularly within the dominant GIII lineage, is driven by a high mutation rate characteristic of RNA viruses and is shaped by strong selective pressures from both the mosquito vector and diverse mammalian hosts [2, 3, 28]. Genomic surveillance has identified several key amino acid sites under positive selection, predominantly within the E2 glycoprotein, which is the primary target of neutralizing antibodies and a major determinant of viral tropism and virulence [2, 3, 6, 23]. For instance, the E2 protein contains multiple positively selected sites distributed within its functional domains A, B, and C, which are associated with receptor binding, infection, and immune evasion [3]. A particularly critical residue is position 253 of the E2 protein. A lysine (K) at this position is associated with high virulence in mice and pigs, whereas a substitution to arginine (R) results in attenuation [5, 19, 20]. Mechanistically, the K253R mutation enhances the virus's binding affinity to heparan sulfate (HS), a ubiquitous cell surface attachment factor, leading to more rapid virus clearance from the bloodstream and reduced virulence in vivo [20]. This residue is also the target of the host antiviral factor TIPARP, which induces K48-linked ubiquitination and proteasomal degradation of E2 at this specific lysine residue, highlighting a direct host-virus evolutionary arms race [19].

Recent large-scale outbreaks, particularly the 2024 epidemic in Henan Province, China, have provided compelling evidence for the emergence of a hypervirulent GIII variant [1]. This variant, responsible for 100% mortality in experimentally infected piglets, is characterized by a distinct set of four amino acid mutations: three in the nonstructural protein nsP3 and one in the structural protein E2 [1]. The emergence of this variant underscores the dynamic nature of GETV evolution and its capacity for virulence enhancement. Furthermore, the detection of unique genetic features, such as a 32-nucleotide repeat insertion in the 3' noncoding region (NCR) of a GIII strain (GDHYLC23) isolated from pigs in Guangdong, China, in 2023, suggests that the virus is continuously exploring new genetic space, the functional consequences of which remain to be fully elucidated [10]. The contamination of commercial live-attenuated vaccines, such as those against porcine reproductive and respiratory syndrome virus (PRRSV), with GETV represents an additional, iatrogenic route of viral spread and genetic diversification, highlighting the complexity of GETV transmission dynamics in modern livestock production systems [9, 21, 39].

Host Range, Geographic Distribution, and Zoonotic Potential

Historically considered a pathogen of horses and pigs, GETV has demonstrated a remarkable capacity for host-range expansion, with natural infections now documented in a wide array of vertebrate species [6, 16, 38]. The virus has been isolated from or serological evidence has been found in cattle, goats, sheep, blue foxes, red pandas, wild boar, birds, and even reptiles [5, 16, 17, 24, 30, 33, 37]. This broad host tropism is facilitated by its primary transmission cycle involving a diverse range of mosquito vectors. GETV has been isolated from at least 17 different mosquito species belonging to five genera (Culex, Anopheles, Armigeres, Aedes, and Mansonia), with Culex tritaeniorhynchus identified as a particularly efficient vector and a key driver of transmission in many endemic regions [7, 15, 16, 18, 37]. The virus's ability to replicate to high titers in both mosquito and vertebrate cells, coupled with its expanding vector and host range, has facilitated its rapid geographical spread from its origins in tropical Southeast Asia to temperate regions as far north as 60° latitude in Eurasia, including Japan, Korea, China, and Russia [16, 28].

The zoonotic potential of GETV is a subject of increasing concern. While clinical disease in humans has not been definitively documented, serological surveys have consistently detected GETV-specific neutralizing antibodies in healthy human populations, with seroprevalence rates exceeding 10% in some studies [6, 25, 33]. The virus's ability to replicate efficiently in human cell lines in vitro and its close phylogenetic relationship with other human-pathogenic alphaviruses, such as CHIKV and RRV, further support its potential to cause human disease [13, 23]. The World Organisation for Animal Health (WOAH) recognizes GETV as an important emerging pathogen of livestock, and its expanding host range and geographical footprint, combined with the lack of commercially available vaccines in many affected countries, position it as a pathogen of significant concern for both animal health and, potentially, public health security [1, 6, 16, 38]. The Centers for Disease Control and Prevention (CDC) and other international health bodies have highlighted the need for enhanced surveillance and research into the ecology and evolution of emerging arboviruses like GETV to preempt and mitigate future outbreaks.

Molecular Pathogenesis: Role of nsP3 and E2 Mutations in Virulence Enhancement

The re-emergence of Getah virus (GETV) as a significant pathogen in livestock, particularly swine and equids, has been underscored by a series of increasingly severe outbreaks across Asia. Central to this epidemiological shift is the emergence and global expansion of the Group III (GIII) genotype, which has been consistently associated with heightened virulence and epizootic potential [1, 2, 7]. A growing body of evidence, derived from extensive genomic surveillance, reverse genetics, and comparative pathogenicity studies, has begun to delineate the molecular determinants underpinning this virulence enhancement. The non-structural protein nsP3 and the structural glycoprotein E2 have emerged as the primary loci of adaptive mutations that drive increased viral fitness, immune evasion, and enhanced pathogenicity in mammalian hosts. These mutations are not merely incidental polymorphisms; they represent a functional evolution of GETV, enabling it to exploit host cellular machinery more efficiently and subvert innate antiviral defenses.

Adaptive Mutations in nsP3: Antagonism of the Host Stress Response and Innate Immunity

The nsP3 protein of alphaviruses is a multifunctional component of the viral replication complex, with a particularly important role in modulating the host cellular environment. Recent large-scale genomic analyses of the 2024 GETV epidemic in Henan, China, have identified a distinct GIII variant possessing three specific amino acid substitutions in nsP3, which correlate with a dramatic increase in pathogenicity, including a 100% mortality rate in experimentally infected piglets [1]. These mutations are not isolated; phylodynamic studies have shown that GETV is evolving rapidly, and the nsP3 gene is a hotspot for adaptive change under strong positive selection [2]. Mechanistically, nsP3 is a critical viral antagonist of the host’s antiviral stress response. The hypervariable domain (HVD) of GETV nsP3 has been shown to bind directly to the NTF2 domain of the cellular stress granule (SG) marker protein G3BP [12]. This interaction is a sophisticated viral strategy to block the formation of bona fide stress granules, which are cytoplasmic aggregates that sequester translationally stalled mRNAs and can inhibit viral replication. Specifically, GETV infection triggers the formation of nsP3-G3BP aggregates that are compositionally distinct from canonical SGs, a process dependent on the PKR/eIF2α signaling pathway [12]. By hijacking G3BP, GETV nsP3 effectively disarms a key arm of the innate immune response, allowing for uninterrupted viral protein synthesis and replication. The specific amino acid changes identified in the GIII hypervirulent strains [1] are predicted to reside within or near this HVD region, potentially enhancing the binding affinity for G3BP or altering the dynamics of aggregate formation, thereby further crippling the host cell’s ability to mount a stress response.

Furthermore, the macrodomain of nsP3 possesses ADP-ribose (ADPr) binding and de-ADP-ribosylation activity, a function that is crucial for counteracting host-mediated ADP-ribosylation of viral and cellular proteins, a key component of the interferon-induced antiviral state [40]. This macrodomain activity is essential for alphavirus replication and neurovirulence. While specific mutations within the macrodomain of the new GIII variants have yet to be fully characterized, the evolutionary pressure on this domain is significant. An enhanced de-ADP-ribosylation capacity could allow the virus to more efficiently strip away this antiviral modification, thereby protecting its replication complex from host restriction factors. The interplay between these nsP3 functions highlights a multi-pronged immune evasion strategy, where mutations that fine-tune both G3BP antagonism and ADPr binding can synergistically enhance viral replication and, consequently, virulence. This is consistent with observations that epidemic GIII strains, such as MI-110 and 14-I-605, exhibit significantly higher replication kinetics in mammalian cells compared to non-epidemic GI and GIV strains, a phenotypic difference that is likely a direct consequence of enhanced nsP3 function [35].

The E2 Glycoprotein: A Nexus of Receptor Binding, Immune Evasion, and Virulence Modulation

The E2 glycoprotein, the major receptor-binding protein of GETV, is the most dynamic viral component under immune and adaptive selection, serving as a critical determinant of host range, cell tropism, and virulence. The World Organisation for Animal Health (WOAH) recognizes GETV as an important emerging pathogen, and the molecular evolution of its E2 protein is a primary focus for understanding its increasing threat to livestock. Numerous independent studies have converged on specific residues within E2 that are under strong positive selection and directly correlate with virulence in both rodent and swine models [2, 3, 23, 28].

A seminal finding is the identification of residue 253 in the E2 protein as a major virulence determinant. A single amino acid substitution at this position, specifically an arginine (R) for lysine (K) change (K253R), was shown to be a gain-of-attenuation mutation in a mouse model, dramatically reducing virulence compared to the wild-type virus [20]. This discovery is mechanistically profound. The K253R substitution creates a new binding site for heparan sulfate (HS), a ubiquitous glycosaminoglycan (GAG) found on the surface of most mammalian cells [20]. While enhanced binding to HS often facilitates viral attachment to cultured cells, in vivo, it can lead to rapid clearance of the virus from the bloodstream by the reticuloendothelial system, resulting in attenuation. Conversely, the wild-type lysine (K253) is associated with lower HS affinity, allowing the virus to evade early clearance and disseminate more effectively, leading to higher systemic virulence. This residue is a critical balance point between efficient cell entry and systemic spread. The ongoing surveillance of GETV shows that the K253 genotype is dominant in highly pathogenic field isolates, underscoring its role in maintaining virulence in the natural transmission cycle [5, 20]. Furthermore, lysine 253 on E2 has been identified as the target for a host antiviral factor, TIPARP. TIPARP recruits the E3 ubiquitin ligase MARCH8 to facilitate K48-linked polyubiquitination of E2 at K253, leading to its proteasomal degradation [19]. Viruses carrying a mutation at this site (K253R) would escape TIPARP-mediated restriction, providing a clear selective advantage for hypervirulent strains. This positions residue K253 not just as a receptor-binding determinant but as a key interface in the host-virus arms race.

Beyond the 253 locus, multiple other positively selected sites in E2 are linked to virulence enhancement. The D262N substitution, which introduces a novel N-linked glycosylation site (N262), has been identified as a critical signature of emerging GIII strains and is predicted to alter the antigenic surface of the virus [3, 26]. The cryo-electron microscopy (cryo-EM) structure of the GETV virion has mapped this glycosylation site to a key region of E2 domain B, which is involved in receptor attachment and is a major target of neutralizing antibodies [25, 26, 41]. The addition of a glycan at this site could sterically shield critical epitopes from antibody recognition, a classic mechanism of immune evasion employed by many enveloped viruses. This would allow the virus to establish a more robust infection even in the presence of pre-existing immunity within a population, contributing to the explosive nature of recent outbreaks. Additional mutations, such as T171I in the E2 protein, have been associated with altered glycosylation patterns and reduced virulence in certain strains isolated from contaminated vaccines, demonstrating that even within the GIII clade, E2 mutations can fine-tune pathogenicity [21]. The collective evidence from genomic surveillance across China, Japan, and Thailand indicates that the E2 protein is a rapidly evolving molecular clock, and the accumulation of these adaptive mutations, enhancing receptor binding, evading innate immune restriction, and escaping humoral immunity, is the primary molecular driver behind the emergence of highly pathogenic GETV GIII variants [1, 3, 5, 15, 23, 29]. The convergence of these genetic changes in the 2024 Henan outbreak, specifically the combination of three nsP3 mutations with a key E2 mutation, highlights a synergistic evolutionary step where the virus has simultaneously optimized its intracellular replication machinery and its extracellular attachment and immune evasion capabilities to achieve a new level of virulence [1].

Genomic Evolution and Phylodynamics of GETV Group III

The genomic evolution and phylodynamics of Getah virus (GETV) Group III represent a paradigm of rapid viral adaptation, spatial expansion, and increasing virulence that has fundamentally altered the epidemiological landscape of this re-emerging arbovirus. While GETV is classified into four distinct phylogenetic groups (Groups I–IV), the overwhelming majority of contemporary isolates, particularly those associated with epizootics in livestock and the expansion of host range, belong exclusively to Group III [3, 28, 35]. This clade has not only displaced other circulating lineages but has demonstrated a remarkable capacity for genetic diversification, adaptive evolution under selective pressure, and phylogeographic dissemination across Asia and Oceania, posing a growing threat to animal health and, potentially, public health as recognized by the World Organisation for Animal Health (WOAH) and the Food and Agriculture Organization (FAO).

Phylogenetic Architecture and Lineage Dominance

Phylogenetic reconstruction based on complete genome sequences and the E2 gene has consistently resolved GETV into four major clades: Group I (prototype Malaysian strain MM2021, 1955), Group II (historical Japanese and Russian strains), Group III (contemporary epidemic strains), and Group IV (a divergent lineage identified in Malaysia, Russia, Thailand, and China) [27, 28, 35]. Critically, extensive genomic surveillance conducted between 2016 and 2024 has demonstrated that Group III is the sole lineage responsible for the ongoing epizootics in swine, horses, cattle, foxes, and other mammalian hosts across China, Japan, South Korea, Thailand, and India [2, 3, 23, 29]. The dominance of Group III is so pronounced that among 82 complete genome sequences available in public databases as of December 2023, the vast majority cluster within this group, with Groups I, II, and IV represented only by a handful of historical or geographically restricted isolates [6, 28].

This phylogenetic monopolization is not a static phenomenon. Within Group III, multiple sub-lineages and variants have emerged, reflecting ongoing microevolutionary processes. For instance, the 2024 outbreak in Henan Province, China, yielded 22 isolates that formed a distinct cluster within the Group III clade, characterized by specific amino acid substitutions in the non-structural protein nsP3 and the structural glycoprotein E2 [1]. Similarly, the GDHYLC23 strain isolated in Guangdong in 2023 exhibited a unique 32-nucleotide repeat insertion in the 3′ non-coding region (3′ NCR), a structural feature not previously documented in any GETV lineage [10]. These observations indicate that Group III is not a monolithic entity but rather a dynamic assemblage of co-circulating variants undergoing continuous genetic flux.

Molecular Clock and Evolutionary Rates

Phylodynamic analyses have revealed that GETV, like other alphaviruses, evolves at a relatively high rate for an RNA virus. Estimates of the evolutionary rate for the complete coding region range from approximately 1.0 × 10⁻⁴ to 3.5 × 10⁻⁴ nucleotide substitutions per site per year, a rate comparable to that of chikungunya virus and other mosquito-borne alphaviruses [2, 28]. This rapid evolutionary pace is driven by the error-prone nature of the RNA-dependent RNA polymerase (RdRp), the large population sizes achievable during epizootic amplification, and the selective pressures imposed by host immune systems and vector species.

Bayesian coalescent-based analyses have further elucidated the demographic history of GETV Group III. The most recent common ancestor (tMRCA) of contemporary Group III strains has been estimated to date to the early to mid-20th century, with a pronounced expansion in effective population size occurring from the 1990s onward [23, 28]. This demographic expansion correlates temporally with the intensification of livestock production, particularly swine farming, in East and Southeast Asia. Notably, phylodynamic modeling has identified a significant association between livestock meat consumption and the evolution of viral genetic diversity, suggesting that anthropogenic factors, including animal trade networks and agricultural intensification, are key drivers of GETV evolution [23].

Selection Pressure and Adaptive Evolution

A hallmark of GETV Group III evolution is the signature of pervasive positive selection acting on specific genomic regions, most notably the E2 glycoprotein. The E2 protein is the primary target of neutralizing antibodies and mediates receptor binding, making it a critical interface for host-virus interactions and a major target of selective pressure. Comprehensive selection pressure analyses using methods such as single-likelihood ancestor counting (SLAC), fixed effects likelihood (FEL), and mixed effects model of evolution (MEME) have identified multiple amino acid sites under positive selection within the E2 protein [2, 3, 23].

Among the most significant positively selected sites is residue 253 of E2. This position has been the subject of intense investigation due to its profound impact on viral phenotype. The substitution of lysine (Lys) with arginine (Arg) at E2-253 has been demonstrated to enhance binding to heparan sulfate (HS), a ubiquitous glycosaminoglycan on the surface of mammalian cells [20]. While this mutation increases infectivity in cell culture by facilitating viral attachment, it paradoxically attenuates virulence in vivo by promoting rapid clearance of the virus from the circulation via the reticuloendothelial system [20]. Conversely, the Arg-to-Lys reversion at this site has been associated with enhanced virulence in swine and mouse models, and this mutation has been identified as a key determinant of the heightened pathogenicity observed in recent Group III variants [5, 20]. The E2-253 site thus represents a molecular switch that modulates the balance between attachment efficiency and systemic dissemination, a paradigm with direct parallels to the E2-218 and E2-234 residues in other alphaviruses.

Additional positively selected sites have been identified in domains A, B, and C of the E2 protein, which are involved in receptor binding, conformational changes during membrane fusion, and immune evasion [3]. For example, the substitution at E2-262 (Asp to Asn) introduces an additional N-linked glycosylation motif, which may shield critical epitopes from neutralizing antibodies or alter receptor tropism [3, 26]. The cryo-electron microscopy structure of GETV at 2.8 Å resolution has confirmed that E2 N262 is indeed glycosylated, and that these glycans are surface-exposed, potentially impacting immune recognition and host cell invasion [25, 26].

Non-Structural Protein Evolution and Innate Immune Evasion

The evolutionary dynamics of GETV Group III are not confined to structural proteins. The non-structural proteins, particularly nsP2 and nsP3, have also undergone adaptive changes that contribute to viral fitness and pathogenesis. The nsP2 protein of GETV has been shown to potently suppress the type I interferon (IFN-I) response by binding to TBK1, thereby inhibiting IRF3 phosphorylation and nuclear translocation [4]. This antagonism of innate immunity is a critical virulence determinant, and sequence variations in nsP2 among Group III strains may modulate the efficiency of this immune evasion strategy.

The nsP3 protein, which contains a macro domain, a zinc-binding domain, and a hypervariable domain (HVD), is another hotspot for adaptive evolution. The macro domain possesses ADP-ribose binding and de-ADP-ribosylation activities that are essential for viral replication and may also counteract host antiviral responses [40]. The HVD of nsP3 interacts with host proteins such as G3BP, a key component of stress granules (SGs). GETV nsP3 binds to G3BP via its HVD, blocking the formation of bona fide SGs and thereby subverting a critical host antiviral pathway [12]. The 2024 Henan outbreak strains exhibited three amino acid mutations in nsP3, suggesting that ongoing selection in this region may further enhance the virus’s ability to manipulate the cellular stress response [1].

Phylogeographic Dispersal and Spatial Dynamics

Phylogeographic reconstruction of GETV Group III has revealed a complex pattern of viral dispersal that is intimately linked to animal movement, vector ecology, and environmental factors. The most comprehensive analysis to date, incorporating 79 E2 gene sequences and 16 complete genomes from Chinese isolates collected between 2016 and 2021, identified three major lineages responsible for the current epidemic in Chinese livestock [23]. Spatially-explicit phylogeographic models indicated that these lineages have preferentially circulated within areas characterized by higher mean annual temperatures and greater pig population densities [23]. This finding aligns with the known ecology of the primary mosquito vectors, particularly Culex tritaeniorhynchus, which thrives in warm, humid environments and is abundant in regions with intensive pig farming [7, 18].

The dispersal history of GETV Group III is marked by long-distance translocation events that likely reflect the movement of infected livestock or contaminated fomites. For example, the GETV-HeN202309 strain isolated in Henan Province in 2023 shared 99.8% nucleotide identity with strains from Guangdong and Sichuan provinces, indicating an imported transmission event rather than local endemic circulation [3]. Similarly, the Japanese strain 22IH8, isolated from Culex tritaeniorhynchus in Nagasaki in 2022, was found to be more closely related to contemporary Chinese strains than to earlier Japanese isolates, suggesting a recent invasion of the Japanese archipelago from mainland Asia [15]. These findings underscore the role of animal trade and transportation networks in facilitating the rapid geographic expansion of GETV.

The phylogeographic analysis also highlighted the importance of southern China as a source population for viral dissemination. The high viral genetic diversity observed in this region, coupled with favorable climatic conditions and dense livestock populations, suggests that southern China functions as an enzootic reservoir and a hub for onward transmission to other regions [23]. This pattern is reminiscent of the emergence and spread of other arboviruses, such as Japanese encephalitis virus (JEV), which the World Health Organization (WHO) has identified as a major public health concern in Asia.

Implications for Virulence and Host Range Expansion

The genomic evolution of GETV Group III has direct consequences for viral pathogenicity and host range. Comparative pathogenicity studies have demonstrated that contemporary Group III isolates are significantly more virulent than earlier strains. For instance, the 2024 Henan variant caused 100% mortality in experimentally infected piglets, with clinical signs including severe fever, diarrhea, and neurological symptoms [1]. Similarly, the GETV-QJ strain, also belonging to Group III, induced 100% lethality in 7-day-old piglets and caused viremia and reduced piglet survival in pregnant sows [11]. These findings contrast sharply with the milder disease observed with historical Group I or Group II strains, which typically caused only transient fever and rash in horses without significant mortality [42, 43].

The expansion of host range is another hallmark of Group III evolution. While GETV was originally considered a pathogen of horses and pigs, Group III strains have now been isolated from cattle, blue foxes, red pandas, and even from contaminated commercial vaccines [21, 30, 37, 39]. The isolation of GETV from a modified live vaccine against porcine reproductive and respiratory syndrome virus (PRRSV) is particularly alarming, as it reveals a previously unrecognized route of iatrogenic transmission that could facilitate the rapid dissemination of virulent strains across swine populations [21, 39]. Furthermore, the detection of GETV-specific antibodies in healthy humans, with seroprevalence rates exceeding 10% in some studies, raises the specter of zoonotic spillover [25, 33]. The Centers for Disease Control and Prevention (CDC) and WHO have classified GETV as a potential emerging zoonotic threat, underscoring the need for enhanced surveillance and risk assessment.

The molecular determinants of this enhanced virulence and host range expansion are multifactorial. In addition to the E2-253 mutation and the acquisition of N-linked glycosylation sites, changes in the 6K protein have been shown to affect virion release and pathogenicity. Deletion of the 6K protein in a recombinant GETV resulted in reduced viral titers, impaired E2 transport to the plasma membrane, and attenuated disease in neonatal mice [22]. Conversely, the presence of a functional 6K protein enhances viral replication and contributes to the clinical manifestation of GETV disease. The interplay between these genetic determinants, host factors, and vector competence will be critical for predicting future trajectories of GETV evolution.

Epidemiology of GETV: Outbreaks in Swine and Emerging Host Range

Getah virus (GETV) has undergone a dramatic epidemiological shift over the past decade, transforming from a neglected arbovirus of primarily historical interest into a re-emerging pathogen of critical importance to global livestock production and public health. The epidemiological landscape of GETV is characterized by an escalating frequency and severity of outbreaks in swine, a progressive expansion of its host range across multiple mammalian taxa, and an alarming geographical spread that now spans from tropical Asia to temperate regions approaching 60° north latitude [16]. This section provides a comprehensive analysis of the epidemiological patterns observed in swine populations and the rapidly expanding host tropism of GETV, drawing on extensive genomic surveillance, serological surveys, and outbreak investigations conducted across Asia and Oceania.

Swine Outbreaks: Magnitude, Clinical Impact, and Spatiotemporal Dynamics

The most striking epidemiological development in recent years has been the emergence of large-scale, highly virulent GETV outbreaks in commercial swine operations, particularly within China. Prior to 2017, reports of GETV-associated disease in pigs were sporadic and often characterized by mild clinical signs. However, the situation changed dramatically with the identification of highly pathogenic strains belonging exclusively to phylogenetic Group III (GIII), which have since become the dominant lineage circulating in field populations [3, 23]. The epidemiological significance of this shift cannot be overstated, as GIII viruses now account for virtually all clinical outbreaks reported across East and Southeast Asia [35].

The 2024 outbreak in Henan Province, central China, represents perhaps the most comprehensive documentation of a GETV epizootic in swine to date. Between late July and mid-September 2024, concentrated outbreaks were observed across 31 commercial pig farms spanning 21 counties, from which 27 distinct GETV strains were isolated [1]. This outbreak exhibited an unprecedented geographical concentration and temporal clustering, suggesting intense local transmission fueled by mosquito vectors. Critically, the causative strains were identified as a novel GIII variant containing three amino acid mutations in the non-structural protein nsP3 and a single mutation in the structural E2 protein. Pathogenicity experiments in piglets revealed a 100% mortality rate, with the variant demonstrating significantly enhanced virulence compared to earlier isolates in comparative mouse models [1]. This study provides compelling evidence that GETV is undergoing a virulence-enhancing evolutionary trajectory, with profound implications for swine health management.

The clinical syndrome observed in contemporary GETV outbreaks differs markedly from historical descriptions. In neonatal and suckling piglets, infection is characterized by acute diarrhea, fever, tremors, and neurological signs progressing rapidly to death [11, 44, 46]. Wu et al. (2024) demonstrated that 7-day-old piglets infected with the GIII strain GETV-QJ exhibited severe diarrhea, fever, intestinal and pulmonary damage, and a 0% survival rate [11]. In pregnant sows, the virus induces viremia that, while not causing fever or mortality in the dams themselves, profoundly affects farrowing performance and reduces piglet survival rates [11]. Reproductive disorders, including abortion, stillbirths, and the delivery of weak or mummified fetuses, have become hallmark features of GETV infection in breeding herds [10, 51]. A 2023 outbreak in Guangdong Province resulted in severe piglet death and sow reproductive disorders that initially tested negative for common swine pathogens, underscoring the diagnostic challenges posed by GETV [10]. Metagenomic analysis ultimately identified GETV as the causative agent, with the isolate GDHYLC23 harboring a unique 32-nucleotide repeat insertion in the 3′ non-coding region, a genomic feature whose functional significance remains under investigation but which may influence viral replication dynamics [10].

Seroprevalence data from across China provide a sobering picture of GETV’s endemicity in swine populations. Using a recombinant capsid protein-based indirect ELISA, Lan et al. (2025) reported positive rates of 63.36% (1,102/1,739) in Jiangxi Province and 37.1% (137/363) in Fujian Province [44]. Similarly, an E2 protein-based ELISA detected 37.59% seropositivity in Eastern China, with significantly higher rates observed in autumn (50.75%) compared to spring (24.24%), confirming a pronounced seasonal pattern consistent with mosquito-borne transmission [49]. In Sichuan Province, testing of 986 clinical serum samples collected between May and September 2022 revealed a 37.1% IgG seropositivity rate [47]. Beyond China, serological surveys in South Korea documented a 26.4% seropositive rate among 670 domestic pigs [48], while Thailand reported 23.1% positivity across 1,188 samples collected from 11 provinces, with significantly higher rates in nursery pigs (67.9%) and older stages (84.5%) compared to finishing pigs (14.2%) [29]. These data collectively indicate that GETV infection is widespread and endemic across Asian swine populations, with seroprevalence levels that rival or exceed those of other established swine pathogens.

A critical epidemiological observation is the association between pig population density and GETV genetic diversity. Phylogeographic reconstruction by Zhao et al. (2022) demonstrated that sampled GETV lineages have preferentially circulated within areas characterized by relatively higher mean annual temperatures and greater pig population densities [23]. This finding suggests that intensive swine production systems create ecological conditions conducive to GETV amplification and maintenance, with pigs serving as both amplification hosts and reservoirs. Notably, the analysis identified an association between livestock meat consumption, but not live pig transport or pork production volumes, and viral genetic diversity, pointing to complex epidemiological dynamics that warrant further investigation [23].

Expanding Host Range: From Traditional Livestock to Novel Mammalian Species

The host range of GETV has expanded dramatically beyond its traditionally recognized equine and porcine hosts, now encompassing at least 10 mammalian species across multiple taxonomic orders. This host expansion is not merely a passive phenomenon but represents active cross-species transmission events driven by viral adaptive evolution.

The detection of GETV in cattle marks a significant milestone in understanding the virus’s ecological breadth. Liu et al. (2019) reported the first isolation of GETV from clinically infected beef cattle in northeastern China, where animals presented with sudden onset fever in forest grazing areas. The JL1808 strain, isolated from cattle, exhibited 99.5% nucleotide identity with the highly pathogenic swine HuN1 strain, demonstrating direct transmission links between swine and bovine populations [37]. Serological investigation revealed that 83.3% of cattle in the study region possessed GETV neutralizing antibodies, indicating extensive exposure [37]. Subsequent surveillance at the China-Myanmar border in Yunnan Province detected a 20.25% seroprevalence rate among 1,300 bovine sera, with active viral RNA detected in 0.23% of samples and the successful isolation of the YN2305 strain, which shared 100% nucleotide identity with the bovine JL1808 strain [17]. These findings establish cattle as competent GETV hosts and potential amplifiers, particularly in mixed farming systems where cattle and swine co-exist.

The identification of GETV in wild boar represents a particularly concerning development for long-term disease maintenance. Huang et al. (2025) reported an extraordinary 74.02% positivity rate (57/77) among wild boar samples in China, with 100% positivity detected in visceral tissues from diarrheic piglets and aborted fetuses [5]. Critically, this study provided the first detection of GETV in milk and semen, raising the possibility of both vertical transmission and venereal spread among wild populations. The representative strain JX2024 belonged to subtype GIII and harbored the key arginine-to-lysine mutation at position 253 of the E2 protein, which has been associated with enhanced virulence [5, 20]. The establishment of GETV in wild boar populations creates a sylvatic reservoir that complicates eradication efforts and provides a persistent source for spillback into domestic swine.

The emergence of GETV in canid species highlights the virus’s expanding ecological niche. In 2017, an outbreak in blue foxes (Vulpes lagopus) in Shandong Province, Eastern China, resulted in the isolation of a highly pathogenic GETV strain phylogenetically related to contemporary swine epidemic strains [30]. Epidemiological investigation strongly suggested that pigs played a pivotal role in transmission to foxes, possibly through mosquito vectors bridging the two-host systems or through contaminated feed [30]. Similarly, serological surveys in Xinjiang Uygur Autonomous Region detected antibodies in sheep (10.0%), goats (11.7%), and cattle (25.1%) in addition to horses and pigs [24]. The detection of GETV antibodies in domestic birds, including chickens and ducks, in Yunnan Province with titers reaching 1:640–1:2560 suggests an even broader host range than currently appreciated [33].

Equine GETV infection, while historically recognized in Japan and India, has now been documented in China with the first horse-derived isolate GZ201808 recovered from a febrile racehorse in Guangdong Province in 2018 [31]. The virus shared 99.6% nucleotide identity with the porcine AH9192 strain, yet phylogenetic analysis placed it in an independent branch distant from other equine strains, suggesting a unique evolutionary trajectory [31, 53]. The 2014–2015 outbreaks in Japanese racehorses at the Miho Training Center provided critical insights into the epidemiology of GETV in managed equine populations. Analysis of these sequential outbreaks revealed that 2-year-old horses newly introduced from GETV-free regions were disproportionately affected (4.4% prevalence in 2-year-olds versus 0.2% in horses aged 3 years or older), that vaccination with two or more doses significantly reduced disease risk (3.0% prevalence versus 22.2% with single-dose vaccination), and that regional seropositivity among horses entering the center increased progressively from 11.8% in August to 34.9% in October [42]. Importantly, concurrent serological surveillance in pigs from surrounding areas demonstrated simultaneous circulation of GETV, with seropositivity rates increasing from 28.8% in 2014 to 65.0% in 2015 in southern Ibaraki Prefecture, and complete genome sequencing confirming 99.89–99.94% nucleotide identity between porcine and equine isolates [52]. This landmark study provides the most compelling evidence to date for the simultaneous circulation of GETV among pigs and horses within defined geographic regions.

The vector-borne transmission dynamics of GETV have been elucidated through comprehensive mosquito surveillance studies. Analysis of 3,600 mosquitoes collected in Guangdong Province revealed Culex tritaeniorhynchus as the primary vector, with a minimum infection rate of 1.36, and Anopheles sinensis as a secondary vector with a rate of 0.83. The isolated GD2202 strain showed 99.3–99.9% E2 gene identity with contemporaneous porcine strains, confirming the epidemiological link between mosquito vectors and swine outbreaks [7]. In northeastern China, vector surveillance demonstrated minimum infection rates of 1.67‰ in Culex pseudovishnui, 1.60‰ in Culex tritaeniorhynchus, and 1.21‰ in Anopheles sinensis, with the MIR for all mosquito species combined reaching 1.50‰ [37]. Experimental vector competence studies by Azerigyik et al. (2023) demonstrated that GETV replicates most efficiently in Culex tritaeniorhynchus- and Aedes albopictus-derived cell lines, with the highest vector competency indices for infection, dissemination, and transmission obtained for Cx. tritaeniorhynchus [18]. Critically, this study provided the first evidence that Ae. albopictus and Anopheles stephensi are competent GETV vectors, expanding the potential transmission network beyond traditional Culex species and highlighting the risk of GETV establishment in regions where these mosquitoes are dominant [18].

Perhaps most concerning from a public health perspective is the accumulating evidence for GETV exposure in human populations. Although no clinical human cases have been confirmed, serological surveys have detected GETV-specific antibodies in healthy individuals in China, with seroprevalence rates exceeding 10% in some studies [25, 28]. The cryo-electron microscopy structure of GETV at 2.8 Å resolution has revealed surface-exposed glycans with potential implications for immune evasion and host cell invasion [25, 26]. The virus replicates efficiently in human cell lines in vitro, including neuronal and epithelial cells [23], and the identification of GETV as a potential contaminant in commercial live veterinary vaccines, with nine batches of a porcine reproductive and respiratory syndrome virus (PRRSV) modified live vaccine testing positive for GETV across three production years (2017–2019) [39], introduces an iatrogenic transmission route with potential implications for human exposure through occupational contact. The isolation of GETV from a commercial PRRSV vaccine by Gao et al. (2023), designated strain BJ0304, underscores the complexity of GETV transmission routes in swine and the need for enhanced surveillance of biological products [21].

The epidemiological trajectory of GETV demands urgent attention from veterinary and public health authorities. The World Organisation for Animal Health (WOAH) recognizes GETV as an emerging pathogen of economic significance, and the Food and Agriculture Organization of the United Nations (FAO) has emphasized the need for enhanced surveillance in livestock populations within endemic regions. The rapid evolutionary rate of GETV, the identification of positively selected sites in the E2 glycoprotein critical for receptor binding and immune evasion [2, 3, 20], and the expanding host range encompassing species with close human contact collectively create conditions favorable for further host adaptation. The absence of commercially available vaccines for swine in China [44, 45], coupled with the demonstrated efficacy of accelerated immunization schedules in horses [50], highlights a critical gap in disease control capabilities that must be addressed to prevent GETV from achieving pandemic status.

Clinical Signs and Pathological Findings in Piglets and Experimental Models

Getah virus (GETV) infection in swine presents a spectrum of clinical manifestations that vary markedly with age, viral strain virulence, and host immune status. The most severe and economically devastating outcomes are observed in neonatal and suckling piglets, where infection frequently culminates in rapid mortality. Concurrently, experimental models, particularly murine and porcine systems, have been indispensable for dissecting the pathophysiological mechanisms underlying GETV-induced disease. This section provides an exhaustive analysis of the clinical signs and pathological findings documented in naturally infected piglets and in controlled experimental infections, drawing upon the most recent and comprehensive literature.

Clinical Signs in Naturally Infected Piglets

In field settings, GETV infection in piglets is characterized by an acute febrile illness with a rapid, often fatal, progression. The clinical syndrome is dominated by pyrexia, severe diarrhea, and neurological dysfunction. Affected piglets typically present with a sudden onset of fever, with body temperatures frequently exceeding 40.5°C [1, 11, 46]. This hyperthermia is accompanied by profound lethargy, anorexia, and huddling behavior, reflecting systemic malaise. Diarrhea is a hallmark clinical sign, ranging from watery to mucoid, and is often profuse, leading to rapid dehydration, electrolyte imbalance, and metabolic acidosis [11, 38, 46]. In many outbreaks, the diarrhea is so severe that it is the primary presenting complaint, often initially misattributed to enteric pathogens such as porcine epidemic diarrhea virus (PEDV) or transmissible gastroenteritis virus (TGEV) [46].

Neurological signs are a particularly ominous feature, frequently heralding a terminal outcome. Piglets may exhibit tremors, ataxia, paddling movements, opisthotonos, and generalized weakness [5, 44, 56]. These signs are indicative of viral invasion of the central nervous system (CNS), leading to meningoencephalitis. In some cases, piglets are found dead without premonitory signs, a phenomenon consistent with peracute death [1, 11]. The mortality rate in affected litters can be devastating. Recent reports from China have documented mortality rates approaching 100% in experimentally infected piglets, and field outbreaks have reported losses of up to 20% of suckling piglets [1, 3, 11]. The 2024 outbreak in Henan province, caused by a virulence-enhanced GIII variant, resulted in 100% mortality in experimentally infected piglets, underscoring the escalating pathogenicity of contemporary circulating strains [1].

Reproductive Disorders in Sows and Vertical Transmission

While piglets bear the brunt of acute mortality, GETV infection in pregnant sows manifests primarily as reproductive failure. The clinical signs in sows are often subtle or subclinical, with fever being the most consistent finding [11]. However, the consequences for the litter are profound. Infection during gestation leads to abortion, stillbirths, mummified fetuses, and the birth of weak, non-viable piglets [7, 10, 38, 44, 51]. The virus can cross the placental barrier, establishing intrauterine infection that disrupts fetal development. In a study by Wu et al. (2024), pregnant sows infected with a highly virulent GIII strain did not exhibit overt clinical signs such as fever or abortion, but did develop viremia, which significantly impacted farrowing performance and reduced piglet survival rates [11]. This highlights the insidious nature of GETV in breeding herds, where the primary economic impact is on reproductive output rather than sow morbidity. The detection of GETV RNA in milk and semen from infected animals further suggests potential routes of horizontal and vertical transmission, complicating control efforts [5].

Gross Pathological Findings in Piglets

Postmortem examination of piglets succumbing to GETV infection reveals a constellation of gross lesions that correlate with the clinical signs. The most consistent findings are in the gastrointestinal tract and the lungs. The small and large intestines are often distended with watery, yellowish contents, and the intestinal wall may appear thin and hyperemic [11, 46]. Mesenteric lymph nodes are frequently enlarged, edematous, and congested, reflecting a robust inflammatory response to viral replication in the gut-associated lymphoid tissue.

Pulmonary lesions are equally prominent. The lungs are typically edematous, heavy, and fail to collapse, with a mottled, red-tan appearance indicative of interstitial pneumonia [11]. Petechial and ecchymotic hemorrhages may be scattered across the pleural surfaces and within the pulmonary parenchyma. In severe cases, frothy serosanguinous fluid can be expressed from the trachea and bronchi, a sign of pulmonary edema. The brain may appear grossly normal, but in cases with pronounced neurological signs, mild congestion of the meningeal vessels and cerebral edema can be observed [46]. Hepatomegaly and splenomegaly are occasionally reported, but are not consistent findings. The thymus is often atrophied, a non-specific indicator of severe stress and systemic disease.

Histopathological Findings in Piglets

Microscopic examination of tissues from infected piglets reveals the underlying cytopathic and inflammatory processes driven by GETV replication. The most severe histopathological changes are observed in the brain, lungs, and intestines.

Central Nervous System: The hallmark lesion is a non-suppurative meningoencephalitis, characterized by perivascular cuffing with mononuclear cells (lymphocytes, plasma cells, and macrophages), gliosis, and neuronal degeneration and necrosis [11, 46]. The cerebrum, cerebellum, and brainstem are all affected, with the most intense inflammation often localized to the gray matter. Neurons may exhibit chromatolysis, pyknosis, and satellitosis. These lesions are the pathological correlate of the tremors, ataxia, and seizures observed clinically.

Respiratory System: The lungs show a diffuse interstitial pneumonia. Alveolar septa are thickened by infiltration of mononuclear cells and hyperplasia of type II pneumocytes. Alveolar spaces may contain proteinaceous edema fluid, sloughed epithelial cells, and occasional macrophages [11]. The bronchiolar epithelium may be necrotic and sloughed. These changes impair gas exchange, contributing to the hypoxia and respiratory distress that can precede death.

Gastrointestinal Tract: The intestinal mucosa shows villous atrophy and fusion, with necrosis of enterocytes at the tips of the villi [46]. The lamina propria is infiltrated by lymphocytes and plasma cells. Crypt hyperplasia is often present, reflecting an attempt at regeneration. These lesions are the direct cause of the malabsorptive diarrhea that is a cardinal clinical sign.

Lymphoid Tissues: The spleen and lymph nodes exhibit lymphoid depletion and necrosis, particularly in the germinal centers. This immunosuppressive effect may predispose piglets to secondary bacterial infections, further complicating the clinical course.

Experimental Models: The Murine System

The mouse has proven to be an invaluable model for studying GETV pathogenesis, virulence determinants, and antiviral therapies. The clinical signs and pathological findings in mice recapitulate many aspects of the disease seen in piglets, making it a relevant and practical surrogate.

Clinical Signs in Mice: The severity of disease in mice is highly dependent on the viral strain, dose, and route of inoculation. Intramuscular (IM) injection is the most commonly used route and reliably produces systemic infection. Following IM inoculation with a virulent strain, mice develop a rapidly progressive illness characterized by ruffled fur, hunched posture, lethargy, and weight loss [20, 22, 57, 58]. Hind limb weakness and paralysis are common, progressing to complete hind limb paresis, a sign of viral neurotropism [13, 20, 55]. In highly virulent strains, such as the GIII variant from the 2024 Henan outbreak, infection leads to 100% mortality within 5–7 days post-infection (dpi) [1]. In contrast, attenuated strains or those with mutations in key virulence determinants (e.g., E2 K253R) cause milder disease with reduced mortality [20, 22].

Route-Dependent Pathogenesis: The route of inoculation profoundly influences disease progression. Jian et al. (2024) systematically compared IM, intranasal (IN), and intraoral (IO) routes in mice. IM inoculation resulted in the most rapid and widespread tissue dissemination, with virus detected in all major organs (brain, lung, liver, spleen, kidney, and testis) by 1 dpi [57]. IN inoculation also led to systemic infection, but with a slightly delayed onset, while IO inoculation resulted in a more restricted infection, primarily involving the gastrointestinal tract and associated lymphoid tissues [57]. This study provided the first experimental evidence that oral infection may represent a natural transmission route, possibly through ingestion of contaminated feed or water.

Pathological Findings in Mice: Histopathological examination of infected mice reveals lesions that mirror those in piglets. The brain shows non-suppurative meningoencephalitis with perivascular cuffing, gliosis, and neuronal necrosis [22, 57]. The lungs exhibit interstitial pneumonia with alveolar thickening and inflammatory cell infiltration [57]. The liver may show scattered foci of hepatocellular necrosis and steatosis [57]. A particularly striking finding in male mice is testicular damage. GETV infection rapidly targets Leydig cells in the testicular interstitium, leading to vacuolation of spermatogonial cells and spermatocytes, decreased sperm density and motility, and increased sperm malformation rates [54, 59]. These changes are evident as early as 0.5 dpi and persist for weeks, with full recovery taking up to 28 days [59]. This testicular tropism has significant implications for the reproductive health of boars, potentially contributing to the reduced fertility observed in infected herds.

Molecular Mechanisms of Pathogenesis: Insights from Experimental Models

Experimental models have been instrumental in elucidating the molecular mechanisms by which GETV causes disease. A key finding is the virus's ability to subvert the host innate immune response. The non-structural protein Nsp2 has been identified as a potent antagonist of type I interferon (IFN-I) production. Liu et al. (2025) demonstrated that GETV Nsp2 binds to TBK1, suppressing the phosphorylation and subsequent nuclear translocation of IRF3, a master transcription factor for IFN-β induction [4]. This blockade of the IFN-I axis allows the virus to replicate unchecked in the early stages of infection, contributing to the high viral loads and rapid disease progression.

Furthermore, GETV Nsp3 has been shown to bind to the stress granule (SG) marker protein G3BP, blocking the formation of bona fide SGs [12]. SGs are cytoplasmic aggregates that form in response to viral infection and are part of the host's antiviral defense. By disrupting SG formation, GETV evades another layer of innate immunity. The virus also manipulates cellular metabolism, inducing reactive oxygen species (ROS) production and triggering autophagy in Leydig cells, a process that the virus hijacks to enhance its own replication [54].

Correlation of Virulence Determinants with Pathological Outcomes

The integration of clinical, pathological, and molecular data from experimental models has allowed for the identification of specific viral genetic determinants of virulence. The E2 glycoprotein is a major target for such studies. A single amino acid substitution at residue 253 (Lys to Arg) in E2 has been shown to dramatically attenuate GETV in mice [20]. This mutation enhances the virus's binding to heparan sulfate (HS), a ubiquitous cell surface attachment factor. While enhanced HS binding increases infectivity in cell culture, it paradoxically reduces virulence in vivo by promoting rapid virus clearance from the circulation [20]. Conversely, the wild-type Lys253 residue is associated with high virulence. This finding has been corroborated by epidemiological studies, where the K253R mutation has been linked to reduced pathogenicity in field isolates [5, 20].

The 6K protein, a small structural protein, also plays a role in virulence. Deletion of the 6K protein (rGETV-Δ6K) results in a virus that is less pathogenic in neonatal mice, causing lower viral loads, less severe histopathological lesions, and milder clinical disease compared to the wild-type virus [22]. The 6K protein is involved in the efficient transport of the E2 glycoprotein to the plasma membrane, a critical step for viral budding. Its deletion impairs virion release, thereby attenuating the virus [22].

The Role of Mosquito Vectors and Transmission Dynamics

Understanding the clinical signs and pathology in the context of transmission is crucial. GETV is maintained in an enzootic cycle involving mosquitoes and vertebrate hosts. Culex tritaeniorhynchus has been identified as a primary vector, with high vector competence for GETV transmission [7, 18]. The virus has also been isolated from Aedes albopictus and Anopheles sinensis, suggesting a broader vector range than previously appreciated [7, 18]. The detection of GETV in mosquitoes in close proximity to pig farms underscores the importance of vector control in preventing outbreaks [7]. The World Organisation for Animal Health (WOAH) recognizes GETV as an emerging pathogen of concern for the swine and equine industries, and the Food and Agriculture Organization (FAO) has highlighted the need for enhanced surveillance and biosecurity measures in endemic regions.

Conclusion of Section

The clinical and pathological landscape of GETV infection in piglets is one of acute, severe, and often fatal disease, driven by viral neurotropism, enterotropism, and immunosuppression. Experimental models, particularly the murine system, have been essential for dissecting the molecular underpinnings of this pathology, revealing key virulence determinants in the E2 and 6K proteins and elucidating the virus's sophisticated strategies for evading the host innate immune response. The escalating virulence of contemporary GIII strains, as evidenced by the 2024 Henan outbreak, underscores the urgent need for continued research into the pathogenesis of GETV and the development of effective vaccines and antiviral therapies. The integration of field observations with controlled experimental data provides a robust framework for understanding and ultimately controlling this re-emerging arboviral threat.

Diagnostic Methods: RT-PCR, Virus Isolation, and Genomic Surveillance

The accurate and timely detection of Getah virus (GETV) is paramount for effective disease management, outbreak containment, and the elucidation of its evolving epidemiology. Given the virus's re-emerging nature, expanding host range, and potential for zoonotic transmission, a multi-faceted diagnostic approach is essential. This section provides an exhaustive analysis of the three cornerstone methodologies for GETV detection and characterization: reverse transcription-polymerase chain reaction (RT-PCR) and its real-time variants, classical virus isolation in cell culture, and the increasingly critical role of genomic surveillance. Each method offers distinct advantages and limitations, and their integrated application provides the most comprehensive understanding of GETV circulation and evolution.

Reverse Transcription-Polymerase Chain Reaction (RT-PCR) and Real-Time Quantitative PCR (RT-qPCR)

Nucleic acid amplification techniques, particularly RT-PCR and its quantitative derivatives, have become the frontline tools for the rapid and sensitive detection of GETV RNA in clinical and vector samples. The inherent genetic variability of RNA viruses necessitates careful primer and probe design, typically targeting highly conserved genomic regions to ensure broad reactivity across circulating strains.

Conventional and Real-Time PCR Assays: Early molecular detection relied on conventional RT-PCR, which, while useful, suffers from lower sensitivity and a lack of quantification. The field has since advanced significantly with the development of real-time quantitative RT-PCR (RT-qPCR) assays. A landmark study established a TaqMan-based RT-qPCR targeting the conserved nsP1 and nsP2 genes, demonstrating a detection limit of 10 copies of GETV RNA and 100% sensitivity and specificity against a panel of related arboviruses, including Japanese encephalitis virus (JEV) and chikungunya virus [65]. This assay proved invaluable for early outbreak detection. Subsequent refinements have focused on improving sensitivity and specificity for different sample matrices and host species. For instance, a SYBR Green I RT-qPCR assay targeting the non-structural protein 3 (nsP3) gene achieved a detection limit of 66 copies/μL, with a 1000-fold higher sensitivity than conventional RT-PCR [64]. The assay exhibited excellent reproducibility, with intra- and inter-assay coefficients of variation below 0.99%, making it suitable for quantitative analysis of viral load in clinical specimens.

Multiplex and High-Throughput Approaches: The co-circulation of GETV with other mosquito-borne viruses, particularly JEV, in pigs and mosquitoes necessitates differential diagnosis. A one-step duplex TaqMan RT-qPCR assay was developed to simultaneously detect and differentiate JEV and GETV, targeting the NS1 gene of JEV and the E2 gene of GETV [41]. This assay demonstrated high sensitivity (100-fold more sensitive than conventional PCR for GETV in field samples), extreme specificity, and excellent repeatability, with correlation coefficients exceeding 0.999 for both viruses. The ability to complete the test in under an hour provides a powerful tool for rapid outbreak response and epidemiological surveillance in endemic regions. Another highly sensitive TaqMan assay, targeting the E1 gene, established a minimum detection limit of 5.94 copies/μL, which is ten times more sensitive than conventional PCR, and exhibited intra- and inter-assay variation coefficients of less than 1% [63]. This assay successfully detected GETV in 10 of 20 field serum samples, compared to only three by conventional PCR, underscoring its superior diagnostic performance.

Isothermal Amplification Methods for Field Deployment: While RT-qPCR remains the gold standard, its reliance on sophisticated thermal cyclers and skilled personnel limits its utility in resource-limited settings or for point-of-care testing. To address this, isothermal amplification methods have been developed. A reverse transcription loop-mediated isothermal amplification (RT-LAMP) assay was established, capable of amplifying GETV RNA within 50 minutes at a constant temperature of 65°C [66]. This method proved to be 1000-fold more sensitive than conventional RT-PCR and 10-fold more sensitive than RT-qPCR, with no cross-reactivity with other porcine viruses. Its simplicity and speed make it an attractive option for field-based surveillance. A further advancement integrated RT-LAMP with Pyrococcus furiosus Argonaute (PfAgo) nuclease, creating a one-tube detection system that generates a fluorescent signal visible under UV light [60]. This assay demonstrated high specificity and a detection limit comparable to RT-qPCR, offering a portable and low-cost alternative for rapid GETV diagnosis.

CRISPR-Based Diagnostics: The most recent frontier in GETV molecular diagnostics leverages the CRISPR-Cas system. A reverse transcription-recombinase polymerase amplification (RT-RPA) assay was coupled with CRISPR/Cas12a for detection via both real-time fluorescence and lateral flow dipsticks (LFD) [61]. This method, targeting the E2 gene, achieved a detection limit of 10 copies/μL and exclusive specificity against JEV and pseudorabies virus. The entire process, from sample to result, can be completed in 50 minutes at a constant temperature of 42°C, eliminating the need for complex instrumentation. Validation with simulated clinical samples showed 100% concordance with RT-qPCR, positioning this technology as a powerful, field-deployable tool for rapid GETV containment. The World Organisation for Animal Health (WOAH) recognizes the importance of such rapid, sensitive diagnostic tools for the surveillance of emerging animal diseases, and these methods align with the goals of the Global Early Warning System for major animal diseases.

Virus Isolation: The Gold Standard for Viable Pathogen Recovery

Despite the sensitivity of molecular methods, virus isolation in cell culture remains an indispensable technique for obtaining live virus for downstream applications, including antigenic characterization, pathogenicity studies, vaccine development, and antiviral drug screening. The successful isolation of GETV is a critical step in confirming an outbreak and providing the biological material necessary for in-depth research.

Cell Lines and Sample Preparation: GETV is capable of infecting a broad range of cell lines from diverse species, reflecting its wide host tropism. The most commonly used and highly permissive cell lines include baby hamster kidney (BHK-21) cells, Vero cells (African green monkey kidney), and swine testicle (ST) cells. BHK-21 cells are particularly favored for their rapid growth and pronounced cytopathic effect (CPE), characterized by cell rounding, shrinkage, and detachment, typically observed within 24-48 hours post-infection [32, 34, 62]. For primary isolation from clinical samples, mouse neuroblastoma (N2a) cells have also been successfully employed, particularly for mosquito isolates [7, 17]. The choice of sample is critical; whole blood, serum, tissue homogenates (e.g., from spleen, lymph node, brain, or intestine), and mosquito pools are all suitable sources. For mosquito surveillance, homogenates are filtered and inoculated onto confluent cell monolayers.

Isolation Protocols and Cytopathic Effect: The standard protocol involves adsorbing the clarified sample inoculum onto a cell monolayer for 1-2 hours, followed by washing and incubation in maintenance medium. Cultures are monitored daily for the development of CPE. For GETV, CPE typically manifests as focal areas of rounding and refractility, which progressively enlarge to involve the entire monolayer, leading to cell detachment within 48-72 hours [34, 62]. The identity of the isolated virus is then confirmed by RT-PCR, immunofluorescence assay (IFA) using specific antibodies, or electron microscopy. A study isolating GETV from pigs in Guangdong reported a peak titer of 10^9.3 TCID50/mL in PK-15 cells at 24 hours post-infection, demonstrating the high replicative capacity of the virus [32]. The isolation of GETV from a commercial modified live vaccine against porcine reproductive and respiratory syndrome virus (PRRSV) highlights a unique and concerning route of transmission, underscoring the need for rigorous quality control in vaccine production [21, 39].

Biological Characterization of Isolates: Once isolated, the virus can be further characterized. Plaque assays are used to determine viral titer and to purify clonal populations. The plaque morphology can vary between strains; for example, the highly virulent GIII variant responsible for the 2024 Henan outbreak produced clear, uniform plaques [1]. Growth kinetics can be assessed by multi-step growth curves, which reveal the replication efficiency of different isolates. Comparative studies have shown that epidemic Group III strains replicate to significantly higher titers (at least 2.3-fold higher) in mammalian and mosquito cells compared to non-epidemic Group IV strains, suggesting that replication capacity is a key determinant of virulence and epidemic potential [35]. Furthermore, the isolated virus can be used in animal models to assess pathogenicity. The 2024 Henan GIII variant, for instance, induced 100% mortality in experimentally infected piglets, confirming its enhanced virulence [1]. Such studies are essential for understanding the biological significance of genetic changes identified through genomic surveillance.

Genomic Surveillance and Evolutionary Analysis

Genomic surveillance has emerged as a critical component of GETV diagnostics, providing a dynamic view of viral evolution, transmission pathways, and the emergence of potentially more virulent or transmissible variants. This approach involves the systematic sequencing of viral genomes from clinical and vector samples, followed by comprehensive phylogenetic and phylodynamic analyses.

Phylogenetic Classification and Lineage Dynamics: The global GETV population is classified into four distinct genotypes (Groups I-IV), with Group III being the dominant and most widely circulating lineage in Asia, responsible for the majority of recent outbreaks in livestock [3, 6, 23]. Genomic surveillance has revealed that all recent isolates from China, Japan, and Korea belong to Group III, while Group IV strains have been identified in Malaysia, Thailand, and Russia [27, 29]. The construction of phylogenetic trees using complete genome or E2 gene sequences allows for the precise tracking of viral spread. For example, the 2024 Henan outbreak was caused by a distinct GIII variant forming a separate cluster, characterized by three amino acid mutations in nsP3 and one in E2 [1]. Similarly, a novel GIII variant in Guangdong, designated GDHYLC23, was found to possess a unique 32-nucleotide repeat insertion in the 3' non-coding region, a feature not previously reported in GETV [10]. These findings underscore the continuous evolution of the virus and the importance of ongoing surveillance.

Identification of Adaptive Evolution and Virulence Determinants: A key goal of genomic surveillance is to identify genetic changes associated with altered virulence, host range, or immune evasion. Selection pressure analyses have identified several amino acid sites in the E2 glycoprotein under positive selection, suggesting they are targets of host immune pressure and may be involved in adaptive evolution [2, 3, 23]. Notably, residue 253 in the E2 protein has been identified as a critical determinant of virulence. A substitution from lysine (K) to arginine (R) at this position enhances binding to heparan sulfate (HS), a cell surface attachment factor, leading to increased infectivity in vitro but attenuated virulence in vivo due to rapid virus clearance from the circulation [20]. Conversely, the wild-type lysine at position 253 is associated with higher virulence, and mutations at this site have been linked to enhanced pathogenicity in recent GIII variants [1, 5]. The identification of such molecular markers is crucial for risk assessment and the development of targeted interventions.

Phylodynamics and Transmission Networks: Phylodynamic analyses integrate genomic data with epidemiological and geographic information to reconstruct the demographic and dispersal history of the virus. Studies have shown that GETV is evolving at a high rate, with a rapid expansion of its host range and geographical distribution, likely driven by the lack of effective vaccines and the intensification of animal trade [6, 28]. Phylogeographic reconstruction has revealed that GETV lineages in China have preferentially circulated in areas with higher mean annual temperatures and pig population densities, highlighting the role of environmental and anthropogenic factors in viral spread [23]. Furthermore, genomic surveillance has provided evidence for cross-species transmission events. The isolation of GETV from cattle in northeastern China, with a genome nearly identical to a highly pathogenic swine strain (HuN1), demonstrated the spillover of the virus from pigs to cattle [37]. Similarly, the detection of GETV in wild boar, including in milk and semen samples, indicates a broader sylvatic cycle that could complicate eradication efforts [5]. The Food and Agriculture Organization of the United Nations (FAO) emphasizes the need for such integrated surveillance at the human-animal-environment interface to predict and prevent emerging zoonotic threats. The continuous genomic monitoring of GETV, as advocated by the World Health Organization (WHO) for emerging pathogens, is therefore not just a diagnostic tool but a fundamental component of a comprehensive One Health strategy.

Immune Response and Vaccine Strategies for GETV Control

The control of Getah virus (GETV) presents a formidable challenge to veterinary public health, given its status as a re-emerging, mosquito-borne alphavirus with a rapidly expanding host range and geographical distribution [1, 6, 16]. The virus, classified within the Alphavirus genus of the Togaviridae family, has demonstrated a capacity for cross-species transmission, infecting horses, pigs, cattle, foxes, and potentially humans, as evidenced by serological surveys detecting GETV-specific antibodies in healthy human populations [6, 25, 33]. The World Organisation for Animal Health (WOAH) recognizes GETV as a significant pathogen impacting livestock economies, particularly in Asia and Oceania, where it causes febrile illness, reproductive disorders, and mortality in swine and equids [51, 73]. Despite this, no commercially available vaccines or specific antiviral therapies are approved for GETV in most affected regions, including China, underscoring an urgent need for comprehensive immune characterization and strategic vaccine development [6, 45, 55]. This section provides an exhaustive analysis of the host immune response to GETV infection, the molecular mechanisms of viral immune evasion, and the current landscape of vaccine strategies, including inactivated, live-attenuated, virus-like particle (VLP), and recombinant protein-based approaches.

Innate Immune Response and Viral Antagonism

The innate immune system constitutes the first line of defense against GETV infection, with type I interferons (IFN-I) playing a pivotal role in restricting viral replication. Upon GETV entry, pattern recognition receptors (PRRs) such as RIG-I and MDA5 detect viral RNA, initiating signaling cascades that converge on the activation of interferon regulatory factor 3 (IRF3) and nuclear factor kappa-B (NF-κB), leading to IFN-β production [4]. However, GETV has evolved sophisticated countermeasures to subvert this antiviral state. A landmark study by Liu et al. (2025) demonstrated that the GETV nonstructural protein 2 (nsP2) is a potent antagonist of IFN-β production [4]. Mechanistically, nsP2 binds to TANK-binding kinase 1 (TBK1), thereby suppressing IRF3 phosphorylation. Furthermore, nsP2 competitively inhibits the interaction of phosphorylated IRF3 (pIRF3) with karyopherin subunits KPNA3 and KPNA4, effectively blocking IRF3 nuclear translocation and subsequent IFN-β transcription [4]. This dual mechanism of action, inhibiting both IRF3 activation and its nuclear import, represents a highly efficient strategy for neutralizing the host’s primary antiviral response.

Beyond IFN antagonism, GETV manipulates other cellular stress pathways to favor its replication. The nonstructural protein 3 (nsP3) has been shown to bind to Ras-GTPase-activating protein SH3-domain-binding protein (G3BP), a key nucleator of stress granules (SGs) [12]. Under normal conditions, SGs are cytoplasmic aggregates of mRNA and proteins that form during cellular stress, including viral infection, and function to halt translation and promote cell survival. Qi et al. (2024) revealed that GETV nsP3, through its hypervariable domain (HVD), binds the NTF2-like domain of G3BP, preventing the formation of bona fide SGs [12]. Instead, GETV infection triggers the formation of aberrant nsP3-G3BP aggregates that are compositionally distinct from canonical SGs. Importantly, while G3BP knockout had no effect on GETV replication, overexpression of G3BP to restore genuine SG formation significantly inhibited viral replication, confirming that SG blockade is a proviral strategy [12]. This interplay between viral proteins and the stress granule machinery highlights a critical axis of host-pathogen interaction.

GETV also exploits autophagy to enhance its replication. Li et al. (2025) demonstrated that GETV infection in mouse Leydig cells induces reactive oxygen species (ROS) production, which in turn triggers autophagy [54]. Pharmacological inhibition of ROS with N-acetylcysteine (NAC) or autophagy with 3-methyladenine (3-MA) significantly reduced viral titers, while activation of autophagy with rapamycin enhanced GETV replication [54]. This suggests that GETV hijacks the autophagic machinery to facilitate its life cycle, a strategy observed in other alphaviruses. Additionally, the host factor tetrachlorodibenzo-p-dioxin-inducible poly(ADP-ribose) polymerase (TIPARP) has been identified as a restriction factor against GETV [19]. TIPARP interacts with the viral E2 glycoprotein, recruiting the E3 ubiquitin ligase MARCH8 to catalyze K48-linked polyubiquitination of E2 at lysine 253, leading to its proteasomal degradation [19]. This finding not only identifies a novel antiviral mechanism but also pinpoints a critical residue in E2 that is under positive selection and linked to virulence [2, 3, 20].

Humoral and Cellular Adaptive Immunity

The adaptive immune response to GETV is characterized by the production of neutralizing antibodies directed primarily against the E2 and E1 envelope glycoproteins, which form the icosahedral spikes on the virion surface [25, 26]. The E2 protein is the major target of neutralizing antibodies, as it mediates receptor attachment and entry. Structural studies using cryo-electron microscopy at 2.8 Å resolution have resolved the atomic architecture of the GETV spike, revealing three distinct domains (A, B, and C) in E2, with domain A being the most exposed and immunodominant [25, 26]. Glycosylation sites at N200 and N262 of E2, as well as N141 of E1, are surface-exposed and may influence immune evasion by masking critical epitopes [25, 26]. The E1 protein, while less immunogenic, is essential for membrane fusion and contains conserved epitopes that can elicit cross-reactive antibodies [14].

In swine, GETV infection induces robust IgG responses that can be detected using recombinant E2-based indirect ELISAs. Sun et al. (2022) optimized an rE2-ELISA and reported seroprevalence rates of 37.59% in Eastern China, with significantly higher rates in autumn (50.75%) compared to spring (24.24%), reflecting seasonal mosquito activity [49]. Similarly, Lan et al. (2025) developed a recombinant capsid protein (Cap)-based ELISA, achieving 94.03% sensitivity and 100% specificity, and found seropositivity rates of 63.36% in Jiangxi and 37.1% in Fujian provinces [44]. These data underscore the widespread circulation of GETV in pig populations and the utility of serological tools for surveillance. In horses, neutralizing antibody titers are a key correlate of protection. Bannai et al. (2016) demonstrated that horses vaccinated twice or more had significantly lower infection rates (3.0%) compared to those vaccinated once (22.2%) during a 2015 outbreak in Japan, highlighting the importance of booster vaccinations for maintaining protective immunity [42]. The virus neutralization test (VNT) remains the gold standard for serological diagnosis, but rapid immunochromatographic strips (ICS) based on recombinant E2 or p62-E1 proteins have been developed for field deployment, showing high concordance with VNT (94.0%) in horses [68, 69].

Cellular immunity, particularly CD8+ T cell responses, is likely critical for viral clearance, although detailed studies in GETV are limited. Transcriptomic analyses of GETV-infected swine testicle (ST) cells revealed systemic activation of inflammatory, apoptotic, and antiviral pathways, including JAK-STAT signaling, which is central to IFN-γ-mediated antiviral effects [71]. Exogenous IFN-γ, but not IFN-α/ω or IFN-λ3, was shown to suppress GETV replication in vitro, suggesting that Th1-type responses are particularly important [71]. The identification of positively selected sites in the E2 protein, such as residues 253 and 262, which are under immune pressure, further supports the role of adaptive immunity in driving viral evolution [2, 3, 20].

Vaccine Strategies: Current Status and Future Directions

Given the absence of licensed GETV vaccines in most countries, significant research efforts have focused on developing safe and efficacious vaccine candidates. The strategies explored to date include inactivated vaccines, live-attenuated vaccines, virus-like particle (VLP) vaccines, and recombinant protein-based platforms.

Inactivated Vaccines: The only commercially available GETV vaccine is an inactivated whole-virus formulation used in Japan for horses. This vaccine, based on the MI-110 strain, has been employed for decades and has demonstrated efficacy in reducing clinical disease during outbreaks [15, 42]. Bannai et al. (2021) evaluated an accelerated immunization schedule (14-day interval vs. conventional 28-day interval) for this inactivated vaccine in Thoroughbred horses [50]. The accelerated schedule induced a more rapid antibody response, with 80% seropositivity at day 28 compared to 40% in the conventional group, although antibody persistence was shorter. This approach is particularly valuable as an emergency control measure during outbreaks. However, inactivated vaccines generally induce weaker cellular immunity and require multiple doses, limiting their utility for mass vaccination in swine operations.

Live-Attenuated Vaccines: A major breakthrough in GETV vaccinology is the development of the live-attenuated candidate GETV-3ΔS2-CM1 by Jiang et al. (2024) [13]. This rationally designed vaccine harbors a deletion in the nsP3 hypervariable domain and specific substitutions in the capsid protein, rendering it genetically stable and highly immunogenic. In a mouse model, a single dose of GETV-3ΔS2-CM1 provided complete protection against viremia and arthritic disease caused by homologous GETV strains. Remarkably, this vaccine also conferred cross-protection against multiple arthritogenic alphaviruses, including Semliki Forest virus, Ross River virus, chikungunya virus, and Barmah Forest virus [13]. This broad-spectrum efficacy is attributed to the induction of cross-reactive neutralizing antibodies and T cell responses targeting conserved epitopes in the E1/E2 glycoproteins. Furthermore, passive transfer of antibodies from immunized sows protected piglets against GETV challenge, demonstrating the potential for maternal immunization strategies [13]. The safety profile of GETV-3ΔS2-CM1 is supported by its attenuation in both immunocompetent and interferon-deficient mice, and its inability to cause disease in mosquito vectors, reducing the risk of environmental spread.

Another live-attenuated approach involves the identification of attenuating mutations in the E2 glycoprotein. Wang et al. (2022) discovered that a single amino acid substitution at residue 253 (Lys to Arg) in E2 enhances binding to heparan sulfate (HS), a ubiquitous cell surface glycosaminoglycan [20]. While this mutation increases in vitro infectivity, it paradoxically attenuates virulence in mice due to rapid virus clearance from the circulation, likely mediated by HS-expressing cells in the liver and spleen. This finding suggests that HS-binding site mutations could be exploited to generate attenuated vaccine strains, a strategy successfully used for other alphaviruses like Sindbis virus.

Virus-Like Particle (VLP) Vaccines: VLPs represent a safe and immunogenic alternative to live vaccines, as they lack viral genetic material but retain the native conformation of envelope glycoproteins. Miao et al. (2024) produced GETV VLPs in insect cells using a baculovirus expression system, demonstrating that the E2 and E1 glycoproteins were properly glycosylated and assembled into enveloped particles of 60-80 nm [55]. Two doses of 1 µg of unadjuvanted VLP vaccine elicited neutralizing antibody responses in wild-type C57/BL6 mice and provided complete protection against viremia and arthritis [55]. The absence of adjuvant in this study is noteworthy, as it suggests that the particulate nature of VLPs is sufficient to activate innate immunity via pattern recognition receptors. The scalability of the baculovirus system makes this platform attractive for commercial production, and the authors propose its application as a vaccine for horses and pigs.

Recombinant Protein and Subunit Vaccines: Subunit vaccines based on the E2 glycoprotein have been explored due to their safety and ease of production. The E2 protein contains the major neutralizing epitopes, and recombinant E2 (rE2) expressed in E. coli or eukaryotic systems has been used extensively for serological diagnostics [47, 49, 72, 74]. However, prokaryotically expressed E2 often lacks proper glycosylation and conformational integrity, which may limit its immunogenicity. To overcome this, Jiang et al. (2024) developed colloidal gold immunochromatographic test strips using eukaryotically expressed soluble p62-E1 protein, which retained native antigenicity [68]. While primarily designed for antibody detection, such recombinant antigens could be formulated with potent adjuvants (e.g., Toll-like receptor agonists) to enhance immunogenicity for vaccine use.

Reverse Genetics and Reporter Viruses: The establishment of infectious cDNA clones for GETV has revolutionized vaccine development and antiviral screening. Cai et al. (2025) constructed a DNA-launched infectious clone (pBR322-GETV-HuN1) driven by a CMV promoter, enabling the rescue of recombinant viruses with biological properties identical to the parental strain [67]. This system has been used to generate reporter viruses expressing EGFP or luciferase, which facilitate high-throughput screening of antiviral compounds and neutralizing antibodies [56, 67, 70]. Ren et al. (2024) engineered a recombinant GETV expressing GFP (rGEEGFP) that was genetically stable for at least five passages and replicated similarly to wild-type virus, making it a valuable tool for rapid neutralization testing [56]. These reverse genetics platforms also enable the rational design of attenuated vaccines by introducing targeted mutations or deletions, as exemplified by the GETV-3ΔS2-CM1 candidate.

Challenges and Strategic Considerations

Despite these advances, several challenges impede the deployment of effective GETV vaccines. First, the genetic diversity of circulating GETV strains, particularly the emergence of highly virulent GIII variants with enhanced pathogenicity, necessitates continuous antigenic surveillance [1, 2, 10]. The identification of positively selected sites in E2, such as D262N which introduces an additional glycosylation motif, may alter antigenicity and reduce vaccine efficacy [3]. Second, the contamination of commercial live vaccines (e.g., PRRSV MLV) with GETV, as reported by multiple groups, highlights a critical biosafety risk [9, 21, 39]. Gao et al. (2023) isolated a novel GIII GETV strain (BJ0304) from a PRRSV vaccine, and Zhou et al. (2020) found that nine batches of a PRRSV MLV from the same manufacturer were GETV-positive [21, 39]. This contamination likely arose from the use of porcine-derived cell lines or raw materials, emphasizing the need for stringent quality control and viral inactivation steps in vaccine production.

Third, the zoonotic potential of GETV, supported by serological evidence of human exposure, argues for a One Health approach to vaccine development [6, 16]. The cross-protective efficacy of GETV-3ΔS2-CM1 against multiple alphaviruses, including chikungunya virus, suggests that a universal alphavirus vaccine may be achievable [13]. Such a vaccine would have dual benefits for veterinary and human health, aligning with the strategic goals of the WHO and FAO. Finally, the development of rapid, field-deployable diagnostic tools is essential for vaccine efficacy trials and post-vaccination monitoring. Novel methods such as CRISPR/Cas12a-based RT-RPA combined with lateral flow dipsticks, which can detect GETV RNA at 10 copies/µL within 50 minutes, offer promising solutions for resource-limited settings [61]. Similarly, RT-LAMP assays provide economical alternatives for field detection [60, 66].

In summary, the immune response to GETV involves a complex interplay between viral evasion strategies and host defense mechanisms, with nsP2-mediated IFN antagonism and nsP3-mediated stress granule disruption being key virulence determinants. The humoral response, particularly neutralizing antibodies against E2, is a critical correlate of protection, while cellular immunity, driven by IFN-γ, contributes to viral clearance. Vaccine development has progressed from traditional inactivated formulations to next-generation live-attenuated and VLP vaccines that offer broad-spectrum protection. The continued evolution of GETV, coupled with its expanding host range and zoonotic risk, mandates sustained investment in vaccine research, enhanced biosecurity in vaccine production, and integrated surveillance systems to mitigate the impact of this emerging pathogen.

Prevention and Control Measures in Endemic Regions

The re‑emergence and rapid expansion of Getah virus (GETV) across Asia and Oceania – driven by a high evolutionary rate, widening host tropism, and the absence of licensed vaccines in most affected countries – demand a multi‑tiered, evidence‑based prevention and control strategy tailored to endemic regions. Effective management must integrate robust surveillance and early detection, vector suppression, strategic vaccination, farm biosecurity, antiviral therapeutics, and coordinated public‑veterinary health policies. The following sections detail each component, drawing on the most recent epidemiological, molecular, and experimental data.

Surveillance and Early Detection Infrastructure

Timely identification of GETV incursions is the cornerstone of containment. Endemic regions require a layered diagnostic network capable of detecting both active infection (viral RNA or antigen) and past exposure (antibodies). Quantitative real‑time RT‑PCR assays targeting conserved regions of the nsP1, nsP2, or E1 genes have been validated with detection limits as low as 5.94 copies/µL and 7.73 PFU/mL, offering 100‑ to 1000‑fold higher sensitivity than conventional RT‑PCR [63, 65, 75]. Duplex TaqMan assays that simultaneously differentiate GETV from Japanese encephalitis virus (JEV) further streamline surveillance in co‑endemic areas [41]. For field‑deployable testing, isothermal amplification methods – including RT‑LAMP (detection within 50 min at 65 °C) and RT‑RAA (sensitivity of 8 copies/reaction) – have been developed and show 100% concordance with gold‑standard qPCR [60, 66, 76]. The integration of CRISPR/Cas12a with RT‑RPA and lateral flow dipsticks now enables visual readout in under 50 min without specialized equipment, making it suitable for resource‑limited settings [61].

Serological surveillance is equally essential for estimating true prevalence and identifying high‑risk populations. Several indirect ELISAs using recombinant E2 or capsid proteins have been optimized for pigs, horses, and cattle, with sensitivities exceeding 94% and specificities of 92–100% [44, 47, 49, 72, 74]. A Gaussia luciferase immunoprecipitation system (LIPS) assay using E2 antigen demonstrated 100% sensitivity and 99.69% specificity, with no cross‑reactivity to other major swine pathogens [45]. Colloidal‑gold immunochromatographic strips based on p62‑E1 or E2 proteins provide rapid (15‑min) on‑farm serological results, with storage stability of at least 3 months at room temperature [68, 69]. These tools should be deployed in sentinel herds – particularly in pig‑dense regions of southern China, Japan, Korea, Thailand, and India – to detect seroconversion before clinical cases appear. The World Organisation for Animal Health (WOAH) recommends incorporating GETV into routine arbovirus surveillance programs for horses and pigs, given its economic impact on the livestock industry.

Vector Control and Mosquito Management

GETV is transmitted primarily by mosquitoes of the genera Culex, Anopheles, Aedes, and Armigeres. Vector competence studies have shown that Culex tritaeniorhynchus exhibits the highest infection, dissemination, and transmission rates, while Aedes albopictus and Anopheles stephensi are also capable vectors [7, 18]. The minimum infection rate in Cx. tritaeniorhynchus in southern China reached 1.36‰, and the virus has been isolated from multiple mosquito species in Guangdong, Shandong, Jilin, and Xinjiang [7, 37, 75]. Therefore, integrated vector management (IVM) must be a priority in endemic zones.

Key measures include: (i) source reduction by eliminating larval habitats (rice paddies, drainage ditches, artificial containers) near farms; (ii) targeted indoor residual spraying and space‑spraying of pyrethroids or organophosphates during peak transmission seasons (July–September in temperate Asia); (iii) use of insecticide‑treated nets in stables and farrowing units; and (iv) biological control using larvivorous fish or Bacillus thuringiensis israelensis. Phylogeographic analyses indicate that GETV dispersal is strongly associated with higher mean annual temperature and pig population density [23]; thus, climate‑driven range expansion necessitates proactive vector surveillance at higher latitudes. The WHO’s Global Vector Control Response framework can be adapted to engage local communities in source reduction.

Vaccination Strategies

No commercial GETV vaccine is currently available for pigs in China, but significant progress has been made with both inactivated and live‑attenuated candidates. In Japan, an inactivated vaccine for horses exists and has been used effectively: during the 2014–2015 outbreaks at the Miho Training Center, horses vaccinated twice or more had a significantly lower prevalence (3.0%) than those vaccinated once (22.2%) [42]. An accelerated two‑dose schedule (14‑day interval) induced protective neutralizing titers earlier than the conventional 28‑day schedule, suggesting its utility for emergency ring vaccination during epizootics [50].

For swine, live‑attenuated vaccine candidate GETV‑3ΔS2‑CM1 – harboring deletions in nsP3 and substitutions in the capsid protein – is genetically stable, provides robust immunogenicity, and confers passive protection to piglets born to immunized sows [13]. In wild‑type mice, a single dose protected against heterologous GETV strains and even against other arthritogenic alphaviruses (Semliki Forest virus, Ross River virus, chikungunya virus), highlighting its potential for a universal alphavirus vaccine [13]. A virus‑like particle (VLP) vaccine produced in insect cells using the baculovirus system also induced neutralizing antibodies and protected mice from viremia and arthritis without adjuvant [55]. Additionally, a reverse genetics system has enabled the construction of recombinant GETV carrying reporter genes or attenuating mutations, which can serve as seed strains for vaccine development [58, 67]. The emergence of hypervirulent GIII variants – causing 100% mortality in piglets and carrying unique amino acid substitutions in nsP3 and E2 – underscores the urgency of vaccine licensure [1, 11]. Regulatory agencies such as the WOAH and national veterinary authorities should prioritize the field testing and approval of safe, efficacious vaccines for endemic zones.

Farm Biosecurity and Quarantine

Given that GETV can be transmitted via contaminated fomites, blood, milk, semen, and even commercial live vaccines, stringent biosecurity is mandatory. A high‑risk route is the contamination of modified‑live virus (MLV) vaccines against other pathogens; GETV has been isolated from several batches of PRRSV MLV in China, indicating a potential iatrogenic spread [9, 21, 39]. Vaccine producers must adopt enhanced quality control – including routine GETV screening of master seeds and cell substrates using RT‑qPCR or metagenomic sequencing. Farms should quarantine newly introduced animals for at least 2 weeks and test for GETV RNA and antibodies before mixing with resident herds.

Reproductive disorders in sows and neurological signs in piglets often follow viremia; thus, isolation of febrile animals and disinfection of farrowing units with virucidal agents (e.g., 0.5% sodium hypochlorite, 70% ethanol, or commercial quaternary ammonium compounds) is critical. The virus can persist in the environment, especially in mosquito‑accessible areas, so stables should be equipped with screens and insect‑proof ventilation. Wild boar, which have shown a 74% GETV positivity rate in China, may serve as a sylvatic reservoir and can transmit the virus to domestic pigs via shared mosquito populations or direct contact [5]. Therefore, fencing and surveillance of feral pig populations near farms are recommended.

Antiviral and Therapeutic Interventions

Currently, no approved antiviral drug exists for GETV. However, several compounds have demonstrated efficacy in vitro and in vivo. The extract of Scutellaria baicalensis (ESG) – whose active components baicalin and baicalein bind the E2 protein – reduced viral attachment (95% inhibition at 10 µg/mL) and shortened viremia in mice [8]. Interferon‑γ (IFN‑γ) suppressed GETV replication in ST cells by activating the JAK‑STAT pathway and complement cascades, while type I/III IFNs were ineffective – a finding explained by GETV’s ability to inhibit IRF3 activation via nsP2 binding to TBK1 [4, 71]. Ivermectin, a broad‑spectrum antiparasitic, inhibited early replication and cell‑to‑cell transmission of GETV in vitro, with an EC50 in the low micromolar range [56]. The host factor TIPARP was identified as a restriction factor that induces K48‑linked ubiquitination and degradation of the E2 protein at lysine 253; exploiting this pathway could lead to novel therapeutic strategies [19]. Furthermore, autophagy modulation affects GETV replication: reducing ROS‑induced autophagy with N‑acetylcysteine or 3‑methyladenine decreased viral titers, whereas enhancing autophagy with rapamycin increased titers [54]. These findings suggest that targeting the autophagy machinery could be a viable host‑directed therapy. NSAIDs and supportive care may alleviate fever and arthritis in horses, but specific antivirals should be integrated into outbreak response plans.

One‑Health Coordination and Public Policy

The increasing detection of GETV antibodies in healthy humans (up to 10% in some surveys) and the potential for zoonotic spillover call for a One Health approach involving veterinary, public health, and entomological sectors [6, 16]. The WHO has not yet listed GETV as a priority pathogen, but the expanding host range – including cattle, foxes, and red pandas – and the high prevalence in livestock (63% seropositivity in pigs in Jiangxi, 74.8% in thoroughbred horses in Xinjiang) warrant risk‑based surveillance [24, 44]. National reference laboratories should sequence viral isolates to monitor the emergence of virulence‑associated mutations (e.g., R253K in E2, nsP3 substitutions, and the 32‑nt repeat insertion in the 3′ UTR) and share data via global platforms such as GISAID or WOAH WAHIS [1, 10, 20].

Policy measures in endemic regions must include: (i) mandatory reporting of GETV outbreaks to national veterinary authorities; (ii) trade restrictions on live pigs, horses, and semen from infected premises; (iii) pre‑movement testing for animals destined for export; and (iv) contingency plans for ring vaccination and vector control during epizootics. The Food and Agriculture Organization (FAO) recommends integrating GETV into existing swine disease surveillance programs (e.g., for PRRSV and JEV) due to shared vectors and clinical overlap. Public awareness campaigns should educate farmers on recognizing signs (fever in horse, piglet diarrhea, sow abortion), using personal protective measures against mosquitoes, and reporting suspect cases promptly.

In conclusion, effective prevention and control of GETV in endemic regions demands a harmonized strategy spanning molecular surveillance, vector management, vaccine deployment, biosecurity, antiviral therapy, and intersectoral governance. The rapid evolution of Group III variants with enhanced virulence underscores the need for adaptive, evidence‑based responses. Continued investment in research – particularly in vaccine development for pigs, high‑throughput serological tools, and climate‑informed vector modeling – will be essential to mitigate the expanding threat of this re‑emerging arbovirus.

References

[1] Sun Y, Yang H, Zhang Y, Liu R, Li L, Shi M, et al.. Outbreak and epidemic of Getah virus infection in swine by virulence-enhanced GIII variant in Henan, central China in 2024. Virulence. 2025. DOI: https://doi.org/10.1080/21505594.2025.2530661

[2] Shen J, Liu S, Liu S, Shen S, Lei M, Xu Q, et al.. Genomic surveillance and evolution of Getah virus. Virus Evolution. 2025. DOI: https://doi.org/10.1093/ve/veaf007

[3] Guo Z, Jiang Y, Li P, Zhang G. Genomic characterization and evolutionary analysis of a Getah virus variant from piglets in central China. Frontiers in Microbiology. 2025. DOI: https://doi.org/10.3389/fmicb.2025.1515632

[4] Liu H, Qi Z, Tian L, Chen Z, Li H, Liu L, et al.. Getah virus nonstructural protein 2 suppresses interferon-beta production by interrupting interferon regulatory factor 3 activation. Veterinary Research. 2025. DOI: https://doi.org/10.1186/s13567-025-01547-3

[5] Huang J, Song D, Wei J, Xie B, Yang Q, Xu J, et al.. First report on identification and genetic characterization of Getah virus in wild boar in China. Frontiers in Microbiology. 2025. DOI: https://doi.org/10.3389/fmicb.2025.1583023

[6] Yuan Y, Hao Y, Peng C, Zhang D, Ma W, Xiao P, et al.. From transmission to adaptive evolution: genomic surveillance of Getah virus. Frontiers in Cellular and Infection Microbiology. 2025. DOI: https://doi.org/10.3389/fcimb.2025.1513392

[7] Deng Y, Lin Z, Lin R, Lu Z, Yan X, Li L, et al.. Isolation and Characterization of Getah Virus GD2202 from Mosquitoes in Foshan, China. Vector Borne and Zoonotic Diseases. 2025. DOI: https://doi.org/10.1089/vbz.2024.0100

[8] Liu B, Wang Y, Shao L, Chen Y, Xu Z, Zhu L. Antiviral activity of Scutellaria baicalensis Georgi Extract against Getah virus in vivo and in vitro. Frontiers in Veterinary Science. 2025. DOI: https://doi.org/10.3389/fvets.2025.1551501

[9] Chu P, Chen S, Zhou X, Wei Z, Zhai S. Getah Virus: A New Contaminant in Veterinary Vaccines. Veterinary Sciences. 2025. DOI: https://doi.org/10.3390/vetsci12020082

[10] Chu P, Guo H, Zhou X, Chen S, Sun X, Tian S, et al.. Emergence of a novel GIII Getah virus variant in pigs in Guangdong, China, 2023. Microbiology spectrum. 2024. DOI: https://doi.org/10.1128/spectrum.00483-24

[11] Wu Y, Gao X, Kuang Z, Lin L, Zhang H, Yin L, et al.. Isolation and pathogenicity of a highly virulent group III porcine Getah virus in China. Frontiers in Cellular and Infection Microbiology. 2024. DOI: https://doi.org/10.3389/fcimb.2024.1494654

[12] Qi X, Zhao R, Yao X, Liu Q, Liu P, Zhu Z, et al.. Getah virus Nsp3 binds G3BP to block formation of bona fide stress granules.. International Journal of Biological Macromolecules. 2024. DOI: https://doi.org/10.1016/j.ijbiomac.2024.135274

[13] Jiang Z, Merits A, Qin Y, Xing G, Zhang L, Chen J, et al.. Attenuated Getah virus confers protection against multiple arthritogenic alphaviruses. PLoS Pathogens. 2024. DOI: https://doi.org/10.1371/journal.ppat.1012700

[14] Liu M, Ren T, Zhang L, Li P, Zhong Z, Zhou L, et al.. Development of a monoclonal antibody specifically recognizing a linear epitope on the E1 protein of Getah virus.. Virology. 2024. DOI: https://doi.org/10.1016/j.virol.2024.110315

[15] Matsumura R, Bannai H, Nemoto M, Higa Y, Kai I, Sasaki T, et al.. Genetic, phylogenetic, and serological analysis of a Getah virus strain isolated from Culex tritaeniorhynchus mosquitoes in Nagasaki, Japan in 2022.. Japanese journal of infectious diseases (Print). 2024. DOI: https://doi.org/10.7883/yoken.jjid.2024.250

[16] Li B, Wang H, Liang G. Getah Virus (Alphavirus): An Emerging, Spreading Zoonotic Virus. Pathogens. 2022. DOI: https://doi.org/10.3390/pathogens11080945

[17] Liu H, Hu J, Li L, Lu Z, Sun X, Lu H, et al.. Seroepidemiological investigation of Getah virus in the China-Myanmar border area from 2022-2023. Frontiers in Microbiology. 2023. DOI: https://doi.org/10.3389/fmicb.2023.1309650

[18] Azerigyik F, Faizah A, Kobayashi D, Amoa-Bosompem M, Matsumura R, Kai I, et al.. Evaluating the mosquito host range of Getah virus and the vector competence of selected medically important mosquitoes in Getah virus transmission. Parasites & Vectors. 2023. DOI: https://doi.org/10.1186/s13071-023-05713-4

[19] Jiao H, Yan Z, Zhai X, Yang Y, Wang N, Li X, et al.. Transcriptome screening identifies TIPARP as an antiviral host factor against the Getah virus. Journal of Virology. 2023. DOI: https://doi.org/10.1128/jvi.00591-23

[20] Wang N, Zhai X, Li X, Wang Y, He W, Jiang Z, et al.. Attenuation of Getah Virus by a Single Amino Acid Substitution at Residue 253 of the E2 Protein that Might Be Part of a New Heparan Sulfate Binding Site on Alphaviruses. Journal of Virology. 2022. DOI: https://doi.org/10.1128/jvi.01751-21

[21] Gao X, Li J, Wu T, Dou J, Zhang W, Jia H, et al.. The Isolation and Characterization of a Novel Group III-Classified Getah Virus from a Commercial Modified Live Vaccine against PRRSV. Viruses. 2023. DOI: https://doi.org/10.3390/v15102090

[22] Meng H, Mou C, Zhang L, Zhou J, Lu T, Chen Z. The roles of 6K protein on Getah virus replication and pathogenicity. Journal of Medical Virology. 2023. DOI: https://doi.org/10.1002/jmv.29302

[23] Zhao J, Dellicour S, Yan Z, Veit M, Gill MS, He W, et al.. Early Genomic Surveillance and Phylogeographic Analysis of Getah Virus, a Reemerging Arbovirus, in Livestock in China. Journal of Virology. 2022. DOI: https://doi.org/10.1128/jvi.01091-22

[24] Shi N, Qiu X, Cao X, Mai Z, Zhu X, Li N, et al.. Molecular and serological surveillance of Getah virus in the Xinjiang Uygur Autonomous Region, China, 2017–2020. Virologica Sinica. 2022. DOI: https://doi.org/10.1016/j.virs.2022.02.004

[25] Wang A, Zhou F, Liu C, Gao D, Qi R, Yin Y, et al.. Structure of infective Getah virus at 2.8 Å resolution determined by cryo-electron microscopy. Cell Discovery. 2022. DOI: https://doi.org/10.1038/s41421-022-00374-6

[26] Wang M, Sun Z, Cui C, Wang S, Yang D, Shi Z, et al.. Structural Insights into Alphavirus Assembly Revealed by the Cryo-EM Structure of Getah Virus. Viruses. 2022. DOI: https://doi.org/10.3390/v14020327

[27] Sam S, Mohamed-Romai-Noor N, Teoh B, Hamim Z, Ng H, Abd-Jamil J, et al.. Group IV Getah Virus in Culex Mosquitoes, Malaysia. Emerging Infectious Diseases. 2022. DOI: https://doi.org/10.3201/eid2802.204887

[28] Shi N, Zhu X, Qiu X, Cao X, Jiang Z, Lu H, et al.. Origin, genetic diversity, adaptive evolution, and transmission dynamics of Getah virus.. Transboundary and Emerging Diseases. 2021. DOI: https://doi.org/10.1111/tbed.14395

[29] Rattanatumhi K, Prasertsincharoen N, Naimon N, Kuwata R, Shimoda H, Ishijima K, et al.. A serological survey and characterization of Getah virus in domestic pigs in Thailand, 2017-2018.. Transboundary and Emerging Diseases. 2021. DOI: https://doi.org/10.1111/tbed.14042

[30] Shi N, Li L, Lu R, Yan X, Liu H. Highly Pathogenic Swine Getah Virus in Blue Foxes, Eastern China, 2017. Emerging Infectious Diseases. 2019. DOI: https://doi.org/10.3201/eid2506.181983

[31] Lu G, Ou J, Ji J, Ren Z, Hu X, Wang C, et al.. Emergence of Getah Virus Infection in Horse With Fever in China, 2018. Frontiers in Microbiology. 2019. DOI: https://doi.org/10.3389/fmicb.2019.01416

[32] Xing C, Jiang J, Lu Z, Mi S, He B, Tu C, et al.. Isolation and characterization of Getah virus from pigs in Guangdong province of China.. Transboundary and Emerging Diseases. 2020. DOI: https://doi.org/10.1111/tbed.13567

[33] Li Y, Fu S, Guo X, Li X, Li M, Wang L, et al.. Serological Survey of Getah Virus in Domestic Animals in Yunnan Province, China. Vector Borne and Zoonotic Diseases. 2019. DOI: https://doi.org/10.1089/vbz.2018.2273

[34] Ren T, Mo Q, Wang Y, Wang H, Nong Z, Wang J, et al.. Emergence and Phylogenetic Analysis of a Getah Virus Isolated in Southern China. Frontiers in Veterinary Science. 2020. DOI: https://doi.org/10.3389/fvets.2020.552517

[35] Mohamed-Romai-Noor N, Sam S, Teoh B, Hamim Z, Abubakar S. Genomic and In Vitro Phenotypic Comparisons of Epidemic and Non-Epidemic Getah Virus Strains. Viruses. 2021. DOI: https://doi.org/10.3390/v14050942

[36] Rawle D, Nguyen W, Dumenil T, Parry RH, Warrilow D, Tang B, et al.. Sequencing of Historical Isolates, K-mer Mining and High Serological Cross-Reactivity with Ross River Virus Argue against the Presence of Getah Virus in Australia. Pathogens. 2020. DOI: https://doi.org/10.3390/pathogens9100848

[37] Liu H, Zhang X, Li L, Shi N, Sun X, Liu Q, et al.. First isolation and characterization of Getah virus from cattle in northeastern China. BMC Veterinary Research. 2019. DOI: https://doi.org/10.1186/s12917-019-2061-z

[38] Qiu Z, Zhang D, Fan Q, Hu Z, Xie H, Li N, et al.. Getah Virus in Review: An Emerging Zoonotic Arbovirus. Transboundary and Emerging Diseases. 2026. DOI: https://doi.org/10.1155/tbed/9245726

[39] Zhou F, Wang A, Chen L, Wang X, Cui D, Chang H, et al.. Isolation and phylogenetic analysis of Getah virus from a commercial modified live vaccine against porcine reproductive and respiratory syndrome virus.. Molecular and Cellular Probes. 2020. DOI: https://doi.org/10.1016/j.mcp.2020.101650

[40] Ferreira‐Ramos AS, Sulzenbacher G, Canard B, Coutard B. Snapshots of ADP-ribose bound to Getah virus macro domain reveal an intriguing choreography. Scientific Reports. 2020. DOI: https://doi.org/10.1038/s41598-020-70870-w

[41] Zhang Y, Li Y, Guan Z, Yang Y, Zhang J, Sun Q, et al.. Rapid Differential Detection of Japanese Encephalitis Virus and Getah Virus in Pigs or Mosquitos by a Duplex TaqMan Real-Time RT-PCR Assay. Frontiers in Veterinary Science. 2022. DOI: https://doi.org/10.3389/fvets.2022.839443

[42] Bannai H, Ochi A, Nemoto M, Tsujimura K, Yamanaka T, Kondo T. A 2015 outbreak of Getah virus infection occurring among Japanese racehorses sequentially to an outbreak in 2014 at the same site. BMC Veterinary Research. 2016. DOI: https://doi.org/10.1186/s12917-016-0741-5

[43] Nemoto M, Bannai H, Tsujimura K, Kobayashi M, Kikuchi T, Yamanaka T, et al.. Getah Virus Infection among Racehorses, Japan, 2014. Emerging Infectious Diseases. 2015. DOI: https://doi.org/10.3201/eid2105.141975

[44] Lan J, Duan L, Liu X, Zhou Y, Zeng B, Chen S, et al.. Seroprevalence of Getah virus in pigs in Southeast China determined with a recombinant Cap protein-based indirect ELISA. Frontiers in Microbiology. 2025. DOI: https://doi.org/10.3389/fmicb.2025.1547670

[45] Li C, Zhang L, Guo J, Tian T, Tang C, Wei Z, et al.. A novel Gaussia luciferase immunoprecipitation assay for the detection of Getah virus antibodies in pigs.. The Veterinary Journal. 2025. DOI: https://doi.org/10.1016/j.tvjl.2025.106450

[46] Lan J, Fang M, Duan L, Liu Z, Wang G, Wu Q, et al.. Novel Porcine Getah Virus from Diarrheal Piglets in Jiangxi Province, China: Prevalence, Genome Sequence, and Pathogenicity. Animals. 2024. DOI: https://doi.org/10.3390/ani14202980

[47] You D, Wang Y, Ge L, Zhou Y, Sun J, Lang L, et al.. Establishment and application of an indirect ELISA for Getah virus E2 antibody detection.. Journal of Virological Methods. 2024. DOI: https://doi.org/10.1016/j.jviromet.2024.114885

[48] Park Y, Kim E, Lee H, Hyun B, Yang D. Serological survey for Getah virus in domestic pigs of South Korea. Korean Journal of Veterinary Research. 2023. DOI: https://doi.org/10.14405/kjvr.20220034

[49] Sun Q, Xie Y, Guan Z, Zhang Y, Li Y, Yang Y, et al.. Seroprevalence of Getah virus in Pigs in Eastern China Determined with a Recombinant E2 Protein-Based Indirect ELISA. Viruses. 2022. DOI: https://doi.org/10.3390/v14102173

[50] Bannai H, Tominari M, Kambayashi Y, Nemoto M, Tsujimura K, Ohta M. Evaluation of Antibody Response in Horses After Vaccination With an Inactivated Getah Virus Vaccine Using an Accelerated Immunization Schedule.. Journal of Equine Veterinary Science. 2021. DOI: https://doi.org/10.1016/j.jevs.2021.103396

[51] . getah virus abortion in pigs- exotic. CABI Compendium. 2022. DOI: https://doi.org/10.1079/cabicompendium.81253

[52] Bannai H, Nemoto M, Niwa H, Murakami S, Tsujimura K, Yamanaka T, et al.. Geospatial and temporal associations of Getah virus circulation among pigs and horses around the perimeter of outbreaks in Japanese racehorses in 2014 and 2015. BMC Veterinary Research. 2017. DOI: https://doi.org/10.1186/s12917-017-1112-6

[53] Lu G, Ou J, Ji J, Ren Z, Hu X, Wang C, et al.. Corrigendum: Emergence of Getah Virus Infection in Horse With Fever in China, 2018. Frontiers in Microbiology. 2019. DOI: https://doi.org/10.3389/fmicb.2019.02601

[54] Li F, Deng L, Xu T, Xu L, Xu Z, Lai S, et al.. Getah virus triggers ROS-mediated autophagy in mouse Leydig cells. Frontiers in Microbiology. 2025. DOI: https://doi.org/10.3389/fmicb.2024.1519694

[55] Miao Q, Nguyen W, Zhu J, Liu G, Oers MMv, Tang B, et al.. A getah virus-like-particle vaccine provides complete protection from viremia and arthritis in wild-type mice.. Vaccine. 2024. DOI: https://doi.org/10.1016/j.vaccine.2024.07.037

[56] Ren T, Liu M, Zhou L, Zhang L, Qin Y, Kang O, et al.. A recombinant Getah Virus expressing a GFP gene for rapid neutralization testing and antiviral drug screening assay.. Virology. 2024. DOI: https://doi.org/10.1016/j.virol.2024.110174

[57] Jian Z, Jiang C, Zhu L, Li F, Deng L, Ai Y, et al.. Infectivity and pathogenesis characterization of getah virus (GETV) strain via different inoculation routes in mice. Heliyon. 2024. DOI: https://doi.org/10.1016/j.heliyon.2024.e33432

[58] Ren T, Min X, Mo Q, Wang Y, Wang H, Chen Y, et al.. Construction and characterization of a full-length infectious clone of Getah virus in vivo. Virologica Sinica. 2022. DOI: https://doi.org/10.1016/j.virs.2022.03.007

[59] Li F, Zhang B, Xu Z, Jiang C, Nei M, Xu L, et al.. Getah Virus Infection Rapidly Causes Testicular Damage and Decreases Sperm Quality in Male Mice. Frontiers in Veterinary Science. 2022. DOI: https://doi.org/10.3389/fvets.2022.883607

[60] Liu Z, Yang F, Fang M, Wu Q, Fan K, Huang D, et al.. Rapid and Sensitive One-Tube Detection of Getah Virus Using RT-LAMP Combined with Pyrococcus furiosus Argonaute. Veterinary Sciences. 2025. DOI: https://doi.org/10.3390/vetsci12020093

[61] Xia B, Wang Z, Fei T, Ma Y, Guo Y, Fei D, et al.. Development and application of a CRISPR/Cas12a-based reverse transcription–recombinase polymerase amplification assay with lateral flow dipstick and fluorescence detection for Getah virus. PeerJ. 2025. DOI: https://doi.org/10.7717/peerj.20119

[62] Shao L, Nie M, Liu B, Li F, Xu T, Xu L, et al.. Isolation and Characterization of a Porcine Getah Virus Strain from Sichuan Province. Veterinary Sciences. 2025. DOI: https://doi.org/10.3390/vetsci12030276

[63] Lin A, Hu X, Cui S, Yang T, Zhang Z, Li P, et al.. Development of TaqMan-based real-time PCR assay based on the E1 genefor the quantitative detection of the Getah virus.. Polish journal of veterinary sciences. 2023. DOI: https://doi.org/10.24425/pjvs.2023.145003

[64] Xia Y, Shi Z, Wang X, Li Y, Wang Z, Chang H, et al.. Development and Application of SYBR Green Ⅰ Real-time Quantitative Reverse Transcription PCR Assay for Detection of Swine Getah Virus.. Molecular and Cellular Probes. 2021. DOI: https://doi.org/10.1016/j.mcp.2021.101730

[65] Sam S, Teoh B, Chee C, Mohamed-Romai-Noor N, Abd-Jamil J, Loong S, et al.. A quantitative reverse transcription-polymerase chain reaction for detection of Getah virus. Scientific Reports. 2018. DOI: https://doi.org/10.1038/s41598-018-36043-6

[66] Liu H, Li L, Bu Y, Liu Y, Sun X, Shi N, et al.. Rapid Visual Detection of Getah Virus Using a Loop-Mediated Isothermal Amplification Method. Vector Borne and Zoonotic Diseases. 2019. DOI: https://doi.org/10.1089/vbz.2018.2434

[67] Cai R, He Q, Wang Q, Tian L, Chen Z, Wu X, et al.. Development of a reverse genetics system for Getah virus and characterization of rescued strains. Veterinary Research. 2025. DOI: https://doi.org/10.1186/s13567-025-01515-x

[68] Jiang Z, Qin Y, Zhang L, Xing G, Shi Z, Song W, et al.. Development and application of a colloidal-gold immunochromatographic strip for detecting Getah virus antibodies. Applied Microbiology and Biotechnology. 2024. DOI: https://doi.org/10.1007/s00253-024-13168-5

[69] Zhong D, Zheng J, Ma Z, Wang Y, Wei J. Rapid Detection of Getah Virus Antibodies in Horses Using a Recombinant E2 Protein-Based Immunochromatographic Strip. Animals. 2024. DOI: https://doi.org/10.3390/ani14162309

[70] Ren T, Zhou L, Min X, Sui M, Zhi X, Mo Y, et al.. Development of a recombinant reporter Getah virus for antiviral drug screening assays.. Veterinary Microbiology. 2023. DOI: https://doi.org/10.1016/j.vetmic.2023.109742

[71] Li J, Gao X, Liu X, Wu T, Song H, Gao W, et al.. The host transcriptome change involved in the inhibitory effect of exogenous interferon-γ on Getah virus replication. Frontiers in Microbiology. 2023. DOI: https://doi.org/10.3389/fmicb.2023.1214281

[72] Qiu X, Cao X, Shi N, Zhang H, Zhu X, Gao Y, et al.. Development and application of an indirect ELISA for detecting equine IgG antibodies against Getah virus with recombinant E2 domain protein. Frontiers in Microbiology. 2022. DOI: https://doi.org/10.3389/fmicb.2022.1029444

[73] Killoran K, Larson KL. Getah virus. CABI Compendium. 2022. DOI: https://doi.org/10.1079/cabicompendium.96169

[74] Bannai H, Nemoto M, Tsujimura K, Yamanaka T, Kokado H. Development of an enzyme-linked immunosorbent assay for Getah virus infection in horses using recombinant E2 protein as an antigen.. Journal of Virological Methods. 2019. DOI: https://doi.org/10.1016/j.jviromet.2019.113681

[75] Cao X, Qiu X, Shi N, Ha Z, Zhang H, Xie Y, et al.. Establishment of a reverse transcription real-time quantitative PCR method for Getah virus detection and its application for epidemiological investigation in Shandong, China. Frontiers in Microbiology. 2022. DOI: https://doi.org/10.3389/fmicb.2022.1009610

[76] Nie M, Deng H, Zhou Y, Sun X, Huang Y, Zhu L, et al.. Development of a reverse transcription recombinase-aided amplification assay for detection of Getah virus. Scientific Reports. 2021. DOI: https://doi.org/10.1038/s41598-021-99734-7