Spring Viremia of Carp Virus

Overview and Taxonomy of Spring Viremia of Carp Virus

Taxonomic Classification and Phylogenetic Position

Spring viremia of carp virus (SVCV) is the etiological agent of spring viremia of carp (SVC), a devastating hemorrhagic disease affecting numerous cyprinid species. Classified within the family Rhabdoviridae, SVCV belongs to the genus Sprivivirus, a taxonomic reassignment that distinguishes it from the archetypal vesiculoviruses such as vesicular stomatitis virus (VSV) [15, 20]. This reclassification, based on phylogenetic analyses of the glycoprotein (G) and nucleoprotein (N) genes, reflects the unique evolutionary trajectory and host adaptation of SVCV within aquatic ecosystems [20]. The virus is a notifiable pathogen to the World Organisation for Animal Health (WOAH), underscoring its transboundary significance and the stringent regulatory frameworks governing its detection and control in international aquaculture trade [7, 15, 18].

The Sprivivirus genus currently encompasses SVCV and pike fry sprivivirus (PFSV), with SVCV being the most economically important member. Comparative genomic analyses reveal that SVCV shares a common ancestry with other rhabdoviruses but has evolved distinct molecular strategies for immune evasion and host cell manipulation [20]. The virus is genetically diverse, with isolates classified into four major genotypes (Ia–Id) that correlate strongly with geographic origin: genotype Ia encompasses isolates from Asia, genotype Ib and Ic are associated with Eastern Europe, and genotype Id predominates in Central Europe [11, 25]. This genotypic partitioning reflects historical viral spread and contemporary epizootic patterns, with Asian isolates (particularly from China) demonstrating a wide range of virulence phenotypes in susceptible hosts [11]. Phylogenetic studies using complete genome sequences and individual gene analyses (most commonly the G and P genes) have provided robust frameworks for tracing viral movements. For instance, Korean SVCV isolates from 2016 were found to cluster within the Asian Ia genogroup, sharing over 98% sequence similarity with Chinese isolates, suggesting a likely introduction from China [25]. Similarly, the first reported SVCV infection in Percocypris pingi in China was classified into genogroup Ia, further confirming the dominance of this lineage in Asian aquaculture systems [26].

Genotypic Diversity and Global Distribution

The global expansion of SVCV has been a matter of significant concern since the late 1990s, with the virus spreading beyond its original European epicenters to establish endemicity in North America and Asia [11, 13]. Genotype Ia isolates are now widely distributed across China, Korea, and other Asian countries, representing a major threat to intensive carp farming operations [11, 25]. In contrast, European genotypes Ib, Ic, and Id exhibit more restricted geographical ranges, though they have been detected in wild and cultured populations across Eastern and Central Europe [11]. The introduction of SVCV into the United States, particularly within the upper Mississippi River basin, has raised concerns about viral persistence in wild fish populations and the potential for spillover into naive cyprinid communities [13]. Notably, SVCV was isolated from quillback (Carpiodes cyprinus), a previously unreported host species, indicating a broader host range than initially recognized and highlighting the potential for viral maintenance in non-cyprinid reservoirs [13].

Comparative virulence studies using koi (Cyprinus carpio koi) as a model host have revealed substantial variation among genotypes. Genotype Ia isolates, while sharing high nucleotide identity, display a wide spectrum of virulence, with mortality rates ranging from 0% to 94% depending on the specific isolate and host variety [11]. European genotypes Ib and Ic exhibit moderate virulence (38–56% mortality), whereas the Id genotype shows variable pathogenicity that can be influenced by host morphology and scale pattern [11]. These findings emphasize that viral genotype, rather than host intraspecific variation, is the predominant determinant of disease outcome, though host factors such as age, immune status, and environmental temperature critically modulate susceptibility [3, 24]. The existence of low-virulence isolates capable of establishing persistent infections (lasting at least 167 days post-exposure) further complicates epidemiological dynamics, as such strains can circulate undetected within populations and serve as reservoirs for future outbreaks [15].

Morphological and Genomic Architecture

SVCV particles exhibit the classic bullet-shaped morphology characteristic of rhabdoviruses, with a length of approximately 100–200 nm and a diameter of 50–80 nm. The viral genome consists of a single-stranded, negative-sense RNA molecule of approximately 11,034 nucleotides, organized into five canonical genes in the order: 3′-N-P-M-G-L-5′ [20, 25]. Each gene encodes a structural or functional protein essential for viral replication, assembly, and pathogenesis. The nucleoprotein (N) encapsulates the genomic RNA, forming the ribonucleoprotein complex that serves as the template for transcription and replication. The phosphoprotein (P) functions as a polymerase cofactor and plays a critical role in both viral RNA synthesis and immune evasion [20]. The matrix protein (M) facilitates virion assembly by bridging the nucleocapsid with the lipid envelope and, as recent studies have revealed, exerts non-structural functions by antagonizing host interferon (IFN) responses [9]. The glycoprotein (G) forms trimeric spikes on the viral surface and mediates attachment to host cell receptors and subsequent membrane fusion. Finally, the large protein (L) is the RNA-dependent RNA polymerase (RdRp), responsible for genome replication and mRNA transcription [20, 22].

High-resolution structural biology has provided exquisite detail into the molecular architecture of SVCV proteins. The central domain of the phosphoprotein (PCD) has been crystallized and resolved at 1.5 Å, revealing a dimeric structure composed of two β-sheets, an α-helix, and an additional two β-sheets per monomer [20]. The dimerization interface, formed by hydrophobic surface residues, is essential for the P protein’s capacity to negatively regulate host IFN production. Mutations that disrupt dimer formation impair the P protein’s immunosuppressive function, highlighting the structural basis of a key virulence mechanism [20]. This detailed structural information provides potential targets for rational drug design aimed at disrupting protein–protein interactions critical for viral replication and immune evasion.

Host Range and Pathogenesis

SVCV exhibits a broad host spectrum within the family Cyprinidae, with common carp (Cyprinus carpio) and koi being the most economically significant and highly susceptible species [1, 6, 11]. However, the virus can infect a diverse array of cyprinids, including goldfish (Carassius auratus), grass carp (Ctenopharyngodon idellus), and zebrafish (Danio rerio), the latter serving as a powerful laboratory model for dissecting host–pathogen interactions [3, 17, 21]. Non-cyprinid hosts have also been documented, including largemouth bass (Micropterus salmoides) and quillback, suggesting an expanding host range that may be facilitated by ecological overlap and viral adaptation [13, 25]. The disease predominantly affects juvenile fish, with mortality rates reaching up to 90% in epizootics, though age-dependent resistance develops, with fish older than 9 months often exhibiting complete resistance to lethal challenge [3, 24].

Pathogenesis is initiated when SVCV gains entry through mucosal surfaces, particularly the gills, fins, and intestinal epithelium. Using recombinant SVCV expressing fluorescent proteins in zebrafish larvae, fins were identified as the primary initial replication sites following bath immersion, a route mimicking natural infection [3]. From these entry points, the virus spreads systemically via the bloodstream, targeting the kidney, spleen, liver, and other internal organs, where it induces severe pathology, including petechial hemorrhages, edema, necrosis, and widespread inflammation [1-3]. The incubation period is temperature-dependent, typically ranging from 1 to 30 days, with disease outbreaks classically occurring in spring when water temperatures rise above 10°C, hence the disease name [25]. At the cellular level, SVCV triggers a multifaceted host response that includes pyroptosis, apoptosis, and autophagy, often concurrently [1, 5]. The virus activates the NLRP3 inflammasome via the viral glycoprotein, leading to gasdermin Eb-dependent pyroptotic cell death, which is characterized by plasma membrane rupture and release of pro-inflammatory cytokines such as interleukin-1β [1]. Concurrently, SVCV infection induces endoplasmic reticulum (ER) stress, which serves as a nexus for both autophagic and apoptotic pathways, with caspase-3, -9, and BCL-2 family members being upregulated [5].

Economic and Regulatory Impact

SVC is considered one of the most serious viral diseases affecting global cyprinid aquaculture, causing substantial economic losses through direct mortality, reduced growth performance, and trade restrictions [1, 2, 7]. The disease is listed as notifiable by WOAH, and its detection triggers immediate quarantine measures, movement controls, and, in many cases, stamping-out policies to prevent international spread [7, 18]. The Ministry of Agriculture and Rural Affairs of China classifies SVC as a Class 2 animal disease, reflecting its high pathogenicity and potential for large-scale outbreaks [2]. Despite decades of research, no commercial vaccine is universally available, though experimental DNA vaccines, subunit vaccines, and probiotic-based oral vaccines have shown promise in laboratory settings [10, 16, 24]. The lack of efficacious antiviral therapeutics further exacerbates the economic burden, driving ongoing efforts to identify natural compounds (e.g., coumarin derivatives, phenylpropanoids, and β-glucans) and repurposed pharmaceuticals (e.g., ribavirin) that can inhibit SVCV replication or enhance host antiviral immunity [4, 8, 12, 14, 19]. The development of validated, high-sensitivity molecular diagnostic tools, such as the WOAH-validated RT-qPCR assay targeting the L gene, has improved surveillance capabilities, enabling early detection and rapid containment of outbreaks [7]. Alternative isothermal amplification methods, including recombinase polymerase amplification (RPA), offer field-deployable options for resource-limited settings [23]. The emergence of SVCV in new geographic regions and host species, coupled with the potential for climate change to expand the thermal window for viral transmission, underscores the need for continued vigilance and investment in prophylactic and therapeutic strategies.

Evolutionary Dynamics and Taxonomic Considerations

The evolutionary trajectory of SVCV is shaped by a combination of genetic drift, reassortment (though rare in non-segmented RNA viruses), and strong selective pressures exerted by host immune systems and environmental factors. The four recognized genotypes likely diverged in allopatry, with subsequent long-distance dispersal facilitated by international trade of live fish and contaminated equipment [11, 25]. Whole-genome comparisons have confirmed that Asian isolates form a monophyletic clade within genotype Ia, while European isolates constitute distinct evolutionary lineages [11]. The recent identification of SVCV in North American wild fish populations, including from the upper Mississippi River, suggests ongoing viral expansion and highlights the inadequacy of current biosecurity measures [13]. Notably, the Korean outbreak isolates (ADC-SVC2016-1 and ADC-SVC2016-3) shared identical genome sequences despite being recovered from different host species (common carp and largemouth bass), indicating a single introduction event and subsequent host jump without significant genetic adaptation [25]. These findings emphasize the importance of phylogenetic monitoring to track viral movements and inform risk assessment.

From a taxonomic perspective, the separation of SVCV from the genus Vesiculovirus into the novel genus Sprivivirus was justified based on substantial genetic divergence, distinct ecological niches, and unique molecular features [15]. The genus name itself is derived from “spring viremia of carp,” reflecting the disease’s most prominent clinical and epidemiological characteristics. Future taxonomic revisions may be warranted as new spriviviruses are discovered and as genomic data from diverse rhabdoviruses accumulate. For now, the classification of SVCV within Sprivivirus provides a coherent framework for understanding its biology and developing targeted control measures.

Epidemiology and Transmission of SVCV

Spring Viremia of Carp Virus (SVCV) is a highly contagious pathogen of major economic and ecological significance, classified as a notifiable disease by the World Organisation for Animal Health (WOAH) due to its capacity for rapid international spread and severe impacts on cyprinid aquaculture [7, 15]. The virus, a member of the genus Sprivivirus within the family Rhabdoviridae, was first identified following epizootics in European carp populations during the 1960s, but has since undergone a dramatic global expansion, emerging as a persistent threat in Asia, North America, and beyond [11, 25]. Understanding the intricate epidemiology and transmission dynamics of SVCV is fundamental for designing effective surveillance, control, and biosecurity strategies.

Host Range and Geographic Distribution

The host spectrum of SVCV is remarkably broad, encompassing a wide range of cyprinid species, most notably the common carp (Cyprinus carpio) and its ornamental variety, koi [11, 15]. However, the virus is not restricted to these primary hosts. Natural infections have been documented in a growing list of species, including grass carp (Ctenopharyngodon idellus), silver carp, bighead carp, and orfe, as well as in non-cyprinid fish such as largemouth bass (Micropterus salmoides) in Korea and Nile tilapia (Oreochromis niloticus), where SVCV infection elicits a measurable innate immune response [17, 25, 30]. Particularly concerning is the identification of new host species, such as the Quillback (Carpiodes cyprinus) in the upper Mississippi River basin, suggesting that SVCV may be exploiting native, non-cyprinid fish as reservoirs or vectors, which could facilitate undetected viral persistence and transmission [13].

Genotyping of SVCV isolates based on the glycoprotein (G) and phosphoprotein (P) genes has delineated four major genotypes that correlate strongly with geographic origin: Ia (Asia), Ib and Ic (Eastern Europe), and Id (Central Europe) [11, 25]. The Asian genotype (Ia) is particularly diverse and virulent, with Chinese isolates exhibiting a wide range of pathogenicity in experimental challenges, causing mortalities from 0% to 94% depending on the specific strain [11]. The global spread is exemplified by the emergence of SVCV in South Korea, where isolates from common carp and largemouth bass clustered within the Asian Ia genogroup, sharing over 98% sequence similarity with Chinese isolates, indicating a likely introduction via trade or movement of infected fish [25]. Similarly, the detection of SVCV in the USA, including the new host Quillback, underscores the virus's capacity for transcontinental dispersal and establishment in novel ecosystems [13].

Environmental Determinants and Transmission Dynamics

SVCV is a quintessential seasonal pathogen, with disease outbreaks typically occurring in the spring when water temperatures range from 10°C to 17°C [24, 25]. This temperature dependency is a defining characteristic of SVC epidemiology. The virus replicates efficiently at cooler temperatures, and optimal virulence is observed at approximately 15°C, while elevated summer temperatures (>20°C) often correlate with a reduction in clinical disease, although the virus may persist in a covert or carrier state [15, 25]. The natural route of infection is horizontal, primarily via the aquatic environment. Waterborne transmission occurs through direct contact with infected individuals or exposure to contaminated water containing shed virus. The virus enters the host through the gills, skin, and gastrointestinal tract. Crucially, high-resolution imaging using recombinant SVCV expressing fluorescent proteins in zebrafish larvae has revealed that the fins are the primary initial sites of viral replication, followed by a systemic spread to internal organs [3]. This finding highlights the importance of mucosal surfaces and the skin as critical portals of entry.

The infection process is initiated by the viral glycoprotein (G protein), which mediates attachment and entry into host cells. The host protein prohibitin (PHB) has been identified as a critical cellular receptor for SVCV; its interaction with the G protein is essential for both attachment and internalization, and overexpression of PHB can confer susceptibility to otherwise non-permissive cells [28]. Other host factors, such as 14-3-3β/α-A, also positively regulate viral entry by interacting with the G protein, further delineating the molecular machinery of infection [31]. Once inside the cell, SVCV triggers a complex cascade of cellular stress responses, including endoplasmic reticulum (ER) stress, autophagy, apoptosis, and pyroptosis, all of which influence the efficiency of viral replication and dissemination [1, 5].

Persistence, Subclinical Carriers, and Mixed Infections

A critical aspect of SVCV epidemiology is its ability to establish persistent infections in survivors of an outbreak. Detailed longitudinal studies in koi have demonstrated that SVCV can be detected for at least 167 days following experimental exposure, with viral titer, prevalence, and replicative rate being strongly correlated with the virulence of the infecting strain [15]. High-virulence isolates not only cause higher acute mortality but also reach higher titers more quickly and persist for longer periods in surviving fish. This carrier state is epidemiologically significant, as seemingly healthy fish can serve as a source of virus for naïve cohorts, particularly during subsequent spring temperature rises. The virus's ability to exploit host antiviral mechanisms for its advantage is evident in its induction of interleukin-10 (IL-10), a potent immunosuppressive cytokine, through the JAK-STAT, NF-κB, and p38 MAPK pathways, thereby dampening the host's antiviral response and facilitating viral persistence [29].

Furthermore, SVCV ecology is complicated by interactions with other aquatic pathogens. A seminal study demonstrated viral interference between SVCV and Infectious Pancreatic Necrosis Virus (IPNV) in zebrafish. A prior infection with IPNV, which causes an asymptomatic infection in this model, significantly increased the survival rate of fish subsequently challenged with SVCV, correlating with inhibition of SVCV RNA synthesis. Conversely, co-infection could sometimes enhance IPNV replication relative to single infections [32]. This phenomenon underscores the complexity of multi-pathogen environments in aquaculture and natural waters, where prior or concurrent infections with other viruses may modulate SVCV transmission and disease outcome.

Transmission in Aquaculture and Implications for Control

In intensive aquaculture settings, the high stocking density and stressful conditions (e.g., handling, temperature fluctuations) amplify transmission rates. The virus can be shed in feces, urine, and body fluids, leading to rapid horizontal spread within tanks or ponds. Control measures are hampered by the lack of a widely available commercial vaccine, although DNA vaccines encoding the G protein have shown remarkable efficacy in experimental settings, conferring up to 100% protection in juvenile carp via intramuscular injection [24]. The route of exposure is crucial for disease progression; bath immersion infection triggers a persistent pro-inflammatory response dominated by IL-1β, whereas intravenous injection primarily activates an antiviral IFN signaling pathway [3]. This distinction has profound implications for vaccine testing and understanding natural pathogenesis.

Robust surveillance and early detection are paramount for containment. WOAH recommends diagnostic methods including virus isolation and RT-PCR. Advanced molecular tools, such as a one-step semi-nested RT-PCR, have demonstrated a 1000-fold improvement in sensitivity over the conventional two-step method, enhancing detection rates in clinical samples [27]. Similarly, a newly developed, rigorously validated RT-qPCR assay targeting the L gene exhibits a limit of detection of 1.28 copies/μL and 100% diagnostic sensitivity for cell-culture isolates, making it a powerful tool for international surveillance and certification of SVCV-free status [7]. The use of such highly sensitive assays is crucial for identifying subclinical carriers, the silent disseminators of SVCV, which represent the greatest challenge for eradication from endemic regions.

Molecular Pathogenesis: Inflammasome Activation and Pyroptosis Induction

Spring viremia of carp virus (SVCV) infection in cyprinids is characterized by profound tissue necrosis, petechial hemorrhages, and a dysregulated inflammatory response that culminates in high mortality, particularly among juvenile fish [1, 3]. While the clinical and histopathological features of SVC have been well-documented, the molecular underpinnings of the host inflammatory pathology have remained elusive until recently. A paradigm-shifting advance in our understanding of SVCV pathogenesis has emerged from the recognition that the virus triggers a specific, pro-inflammatory form of programmed cell death known as pyroptosis. This process, mediated by the activation of canonical inflammasome complexes and the subsequent cleavage of gasdermin family proteins, is now understood to be a central driver of the tissue damage and systemic inflammation observed during acute SVCV infection [1]. The World Organisation for Animal Health (WOAH) lists SVC as a notifiable disease, underscoring the critical need to understand its pathogenic mechanisms to develop effective control strategies [7, 27]. The elucidation of the inflammasome-pyroptosis axis in fish represents a major conceptual leap, revealing a conserved yet host-specific mechanism of viral pathogenesis that bridges the innate immune response with cellular demise.

The NLRP3 Inflammasome: A Central Hub for SVCV-Induced Inflammation

The initiation of pyroptosis during SVCV infection is fundamentally dependent on the assembly and activation of the NLRP3 inflammasome. In mammalian systems, NLRP3 is a well-characterized pattern recognition receptor (PRR) that responds to a diverse array of pathogen-associated molecular patterns (PAMPs) and damage-associated molecular patterns (DAMPs). Li et al. (2025) provided the seminal demonstration that SVCV infection in fish cells activates the NLRP3 inflammasome, leading to the recruitment and cleavage of pro-caspase-B (the fish functional ortholog of mammalian caspase-1) into its active form, caspase-B (p20) [1]. This activation is not a passive consequence of viral replication but is a specific, virus-driven process. The critical role of NLRP3 was confirmed through pharmacological intervention; treatment with a specific NLRP3 inhibitor effectively abrogated SVCV-induced pyroptosis, as evidenced by the suppression of caspase-B activation, mature interleukin-1β (IL-1β) production, and lactate dehydrogenase (LDH) release [1]. This finding firmly establishes the NLRP3 inflammasome as the primary molecular platform through which SVCV initiates the inflammatory cascade.

The mechanism by which SVCV triggers NLRP3 activation is multifaceted and involves the stabilization of the NLRP3 protein itself. A key discovery is the role of the host mitochondrial protein, voltage-dependent anion channel 2 (VDAC2). VDAC2, traditionally known for its function in mitochondrial metabolite transport, was identified as an essential host factor for NLRP3 inflammasome activation during SVCV infection [1]. Mechanistically, VDAC2 interacts with NLRP3 and maintains its protein stability, preventing its degradation and ensuring a sufficient pool of NLRP3 is available for inflammasome assembly upon viral challenge. This dependency was elegantly demonstrated in vivo: zebrafish larvae with VDAC2 gene knockdown exhibited significantly reduced cellular damage and increased survival following SVCV infection, correlating with diminished NLRP3 inflammasome activation [1]. Furthermore, the pharmacological inhibition of VDAC2 with DIDS (4,4′-diisothiocyanatostilbene-2,2′-disulfonic acid) in zebrafish reduced SVCV-induced pyroptosis and alleviated tissue inflammation [1]. This positions VDAC2 not merely as a passive bystander but as a critical upstream regulator of the NLRP3 inflammasome, linking mitochondrial homeostasis to the inflammatory response against SVCV.

Gasdermin Eb-Dependent Pyroptosis: The Execution Phase

The terminal step in the pyroptotic pathway is the execution of cell lysis, which is mediated by the gasdermin family of proteins. Following NLRP3 inflammasome activation, active caspase-B cleaves the pore-forming protein gasdermin Eb (GSDMEb), liberating its N-terminal domain. This domain then translocates to the plasma membrane, where it oligomerizes to form large, non-selective pores [1]. The formation of these pores is the defining event of pyroptosis, leading to a loss of membrane integrity, cell swelling, and the release of pro-inflammatory intracellular contents, including LDH and mature IL-1β. Li et al. (2025) demonstrated that SVCV infection specifically induces GSDMEb-dependent pyroptosis in fish cells, a process that is completely blocked by inhibiting either NLRP3 or caspase-B [1]. This confirms that the entire signaling cascade, from NLRP3 activation to GSDMEb pore formation, is a linear and essential pathway for SVCV-induced inflammatory cell death.

The consequences of this pyroptotic cell death are profound for the host. While pyroptosis can serve as a mechanism to eliminate the replicative niche of intracellular pathogens, the massive and uncontrolled release of DAMPs and cytokines, particularly IL-1β, fuels a hyperinflammatory state. In the context of SVCV, this is directly linked to the severe tissue necrosis observed in infected fish. The persistent and robust expression of IL-1β has been documented in multiple SVCV infection models, including bath immersion of zebrafish larvae, where it correlates with neutrophil recruitment and uncontrolled inflammation leading to host death [3]. This IL-1β-driven inflammation is a hallmark of SVCV pathogenesis, and its production is directly tied to the pyroptotic machinery. The virus itself appears to exploit this pathway; the SVCV glycoprotein (G protein) was identified as the primary viral determinant responsible for inducing pyroptosis [1]. This suggests that the virus has evolved to actively trigger this inflammatory cell death program, potentially to facilitate viral dissemination, evade intracellular antiviral defenses, or create a favorable environment for replication.

Viral Countermeasures and the Regulation of Inflammatory Cell Death

The host-pathogen arms race is vividly illustrated by the complex interplay between SVCV and the host’s pyroptotic and inflammatory pathways. While SVCV actively induces NLRP3-dependent pyroptosis, it simultaneously engages in sophisticated strategies to modulate the host immune response, often with conflicting outcomes. For instance, the virus can induce a state of profound immunosuppression to counteract the antiviral effects of the interferon (IFN) system. The SVCV phosphoprotein (P protein) and matrix protein (M protein) are potent antagonists of the IFN response, targeting key signaling molecules such as MAVS, TRAF3, and STING to suppress IFN production [9, 34, 35]. This creates a paradoxical environment where the virus promotes a potent inflammatory cell death pathway (pyroptosis) while simultaneously dampening the type I IFN response, which is crucial for establishing an antiviral state. This dichotomy suggests that SVCV’s primary pathogenic strategy is to drive a maladaptive, pro-inflammatory response that causes tissue damage, rather than to simply replicate unchecked.

Further complicating the picture is the virus’s ability to manipulate other forms of programmed cell death, including autophagy and apoptosis, which are intimately connected to inflammasome regulation. SVCV infection is known to induce endoplasmic reticulum (ER) stress, which in turn can trigger both autophagy and apoptosis [5]. The interplay between these pathways and pyroptosis is critical. For example, the autophagy receptor optineurin (OPTN) is upregulated by SVCV and promotes mitophagy, a process that can dampen NLRP3 inflammasome activation by removing damaged mitochondria, a source of ROS and mtDNA [33]. Conversely, the virus also utilizes the autophagic machinery for immune evasion, as seen with the SVCV N protein degrading STING via the autophagy-lysosome pathway [34]. The balance between these processes, pyroptosis, apoptosis, and autophagy, ultimately dictates the outcome of infection. The engagement of gcFKBP5 by SVCV to promote host cell apoptosis for supporting viral replication further highlights the virus’s ability to co-opt cell death pathways for its own benefit [17]. This intricate network of interactions underscores that SVCV pathogenesis is not a simple, linear process but a dynamic and highly regulated battle for control over the host cell’s fate.

In Vivo Consequences and Therapeutic Implications

The relevance of the NLRP3-GSDMEb pyroptosis axis extends beyond in vitro observations and is a critical determinant of disease outcome in vivo. In zebrafish models, SVCV infection leads to a persistent pro-inflammatory response, characterized by sustained IL-1β expression and neutrophil infiltration at sites of initial viral replication, such as the fins [3]. This uncontrolled inflammation is a direct driver of the tissue damage and mortality seen in infected larvae. The protective effect of VDAC2 knockdown or pharmacological inhibition with DIDS in zebrafish provides compelling evidence that targeting this pathway can mitigate disease [1]. By reducing NLRP3 inflammasome activation and subsequent pyroptosis, these interventions alleviate tissue inflammation and improve survival, even in the face of ongoing viral replication. This suggests that the host’s own inflammatory response, rather than the virus itself, is a primary cause of pathology.

These findings have significant therapeutic implications. The identification of VDAC2 as a druggable target for controlling SVCV-induced inflammation opens new avenues for intervention. While direct antiviral drugs are being explored, such as ribavirin and various natural compounds like rhein and coumarin derivatives [4, 12, 36, 38], targeting the host inflammatory response could provide a complementary strategy to reduce disease severity. The concept of “immunomodulation” rather than direct viral inhibition is particularly attractive for managing SVCV outbreaks, as it may be less prone to the development of viral resistance. Furthermore, the intricate relationship between pyroptosis and other cellular processes like mitophagy, which is manipulated by the virus via OPTN [33], suggests that multi-targeted approaches may be necessary. For instance, combining an inhibitor of NLRP3 or VDAC2 with an agent that restores IFN signaling could potentially provide a synergistic benefit, curbing both the inflammatory damage and enhancing antiviral immunity. The development of such host-directed therapies, alongside advances in vaccine technology using DNA vaccines or nanoparticle delivery systems [10, 24, 37, 39, 40], represents the future of SVCV control, moving beyond a purely virus-centric view to one that considers the complex host-pathogen interface.

Host Antiviral Response: Role of pIgRL4.2 and Interferon Signaling

The host defense against Spring Viremia of Carp Virus (SVCV) is orchestrated through a multifaceted network of innate immune pathways, with the interferon (IFN) system constituting the central antiviral axis. The World Organization for Animal Health (WOAH) lists SVC as a notifiable disease, underscoring the economic and ecological threat posed by this rhabdovirus to global cyprinid aquaculture [7, 15, 27]. Understanding the molecular architecture of the teleost IFN response is therefore paramount for developing effective prophylactic and therapeutic strategies. Within this intricate signaling landscape, the polymeric immunoglobulin receptor-like protein 4.2 (pIgRL4.2) has emerged as a critical positive regulator, functioning at the nexus of mucosal immunity and the type I IFN cascade. This section dissects the mechanistic role of pIgRL4.2, integrates its function within the broader framework of IFN induction and signaling, and contrasts these antiviral mechanisms with the sophisticated countermeasures deployed by SVCV to subvert host immunity.

The pIgRL4.2 Molecule: A Mucosal Sentinel Amplifying IFN Signaling

The mucosal immune system of teleosts represents the first line of defense against waterborne pathogens, including SVCV [2]. The polymeric immunoglobulin receptor (pIgR) is a cornerstone of mucosal immunity, mediating the transcytosis of polymeric immunoglobulins across epithelial barriers. In zebrafish, a diversified family of pIgR-like (pIgRL) molecules exists, and among these, pIgRL4.2 is distinguished by its high constitutive expression in immune-relevant organs, suggesting a specialized, non-redundant role in host defense [41]. Critically, SVCV infection itself significantly downregulates pIgRL4.2 expression, a phenomenon hypothesized to represent a viral strategy to dismantle a key component of the host's antiviral arsenal [41]. This negative regulation by the virus implicitly underscores the molecule's importance in restraining viral pathogenesis.

Functional investigations revealed that ectopic overexpression of pIgRL4.2 in epithelioma papulosum cyprini (EPC) cells dramatically delays the onset of SVCV-induced cytopathic effects and potently suppresses viral replication [41]. The antiviral mechanism of pIgRL4.2 is inextricably linked to its role as a positive regulator of the IFN system. Overexpression of pIgRL4.2 synergistically enhances IFN promoter activation induced by both the double-stranded RNA mimic poly(I:C) and by live SVCV infection [41]. This amplification of IFN signaling is not merely correlative; it is functionally required for its antiviral activity. The generation of pIgRL4.2-null zebrafish via CRISPR/Cas9 gene editing provided definitive in vivo evidence for this role. These knockout animals exhibit significantly reduced survival rates following SVCV challenge, accompanied by a consistent and profound downregulation of antiviral responsive genes, including critical components of the IFN signaling cascade, and a concomitant increase in SVCV replication [41]. Thus, pIgRL4.2 functions as a pivotal molecular scaffold or adaptor that tunes the magnitude of the IFN response, effectively translating a mucosal signal into a robust systemic antiviral state.

The Interferon Signaling Axis: Convergence of Multiple PRRs and Adaptors

The IFN system in cyprinids is activated by pattern recognition receptors (PRRs) that detect SVCV-associated molecular patterns. The RIG-I-like receptor (RLR) family, particularly RIG-I and MDA5, are primary sensors of SVCV genomic RNA. Transcriptomic and functional analyses in common carp have demonstrated that SVCV infection profoundly enriches the RLR signaling pathway, with RIG-I and MDA5 eliciting potent antiviral responses by inducing TBK1 and IRF3 phosphorylation, culminating in the expression of ifn-1, viperin, isg15, and mx [43, 44]. The crucial role of the E3 ubiquitin ligase TRIM25 in this context has been elucidated, where CcTRIM25 (common carp TRIM25) interacts with the CARD domain of CcRIG-I and facilitates its K63-linked polyubiquitination, a post-translational modification essential for RIG-I activation and downstream signal propagation [43]. Similarly, the laboratory of genetics and physiology 2 (LGP2) has been shown to act as a positive regulator of MDA5-mediated signaling during SVCV infection, colocalizing and interacting with MDA5 to enhance IFN-β promoter activation [44].

Downstream of the RLRs, the mitochondrial antiviral-signaling protein (MAVS) and the stimulator of interferon genes (STING) serve as critical adaptor platforms. While STING is traditionally associated with DNA virus sensing, its role in the response to the RNA virus SVCV is now well-established. Common carp STING (CcSTING) is localized to the endoplasmic reticulum and its expression is strongly induced by SVCV and poly(I:C) in immune tissues such as the gills and head kidney [42]. Overexpression of CcSTING profoundly suppresses SVCV replication by enhancing IFN-1 and ISG expression, a process that involves the phosphorylation of TBK1 and p65, thereby linking the STING pathway to both IRF3 and NF-κB activation [42]. The convergence of these pathways dictates the magnitude and kinetics of the type I IFN response, and it is at this juncture that pIgRL4.2 is hypothesized to exert its regulatory influence, likely by stabilizing signaling complexes or promoting the post-translational activation of key kinases such as TBK1 or IKKε.

The Antagonistic Battlefield: Viral Subversion of the IFN Response

The critical importance of the IFN axis is underscored by the multiple, layered strategies SVCV has evolved to dismantle it. SVCV employs a multipronged offensive aimed at every level of the IFN cascade, from PRR signaling to the activity of ISG products. The viral N protein has been identified as a potent IFN antagonist that targets STING for degradation. Specifically, the SVCV N protein interacts with STING and targets it for degradation via the autophagy-lysosome pathway, a process that is dependent on Beclin1 [34]. This effectively removes a central signaling hub, preventing IFN induction. Furthermore, the SVCV matrix protein (M protein) orchestrates a sophisticated molecular burglary by competitively recruiting TRAF3 away from the MAVS signaling complex. The M protein exhibits a higher binding affinity for TRAF3 than MAVS, physically displacing the adaptor and inhibiting the K63-linked polyubiquitination of TRAF3, a prerequisite for downstream IRF3 activation [9]. This effectively silences the RLR-MAVS axis.

The SVCV phosphoprotein (P protein) is another multifunctional antagonist. Beyond its essential role in viral replication, the P protein acts as a cofactor for the negative regulator IRF2a, which suppresses the IFN response. SVCV-P interacts with IRF2a, inhibits its K48-linked ubiquitination to prevent its degradation, and promotes its nuclear translocation [35]. Once in the nucleus, the IRF2a-P protein complex collaborates to destabilize STAT1a and block its nuclear translocation, thereby crippling the JAK-STAT signaling pathway that is essential for ISG expression [35]. The crystal structure of the P protein central domain (PCD) revealed that its ability to form dimers through hydrophobic interactions is directly linked to its capacity to negatively regulate the host IFN response, underscoring the structural basis of its antagonistic function [20]. Additionally, SVCV has been shown to degrade the potent splice variant viperin_sv1, a direct ISG product, through the ubiquitin-proteasome pathway mediated by its N protein, providing a mechanism to eliminate a powerful antiviral effector [45, 46]. This intricate interplay between host antiviral factors (like pIgRL4.2, TRIM25, STING, and MAP2K7) and viral antagonists (N, M, and P proteins) defines the delicate balance between pathogenesis and survival.

Route of Infection and the Dichotomy of the Inflammatory Response

The route of SVCV entry profoundly shapes the nature of the host response, a factor that must be considered when interpreting in vivo studies. Using a bath immersion infection model with a recombinant SVCV expressing mCherry, it was demonstrated that the fins are the primary initial replication sites [3]. Crucially, this natural route of infection triggers a persistent pro-inflammatory response, characterized by sustained neutrophil recruitment and robust expression of the pro-inflammatory cytokine IL-1β, rather than a strong activation of the antiviral IFN pathway [3]. This contrasts sharply with infection models using intravenous injection, which predominantly activate IFN signaling, highlighting the critical importance of the mucosal interface in shaping the innate immune trajectory [3]. The tissue damage observed at these initial infection sites is likely a consequence of both virus-induced cytopathology and the ensuing uncontrolled inflammatory response. The ability of SVCV to induce pyroptosis through the NLRP3 inflammasome and gasdermin Eb, leading to the release of IL-1β and LDH, further amplifies this pathogenic inflammatory milieu [1]. This suggests that the host's response to SVCV, particularly at mucosal surfaces, is a double-edged sword: the IFN response is essential for viral clearance, but an exaggerated or misdirected pro-inflammatory response can contribute to tissue pathology and mortality. The strategic downregulation of pIgRL4.2 by SVCV may therefore serve to dampen the protective IFN arm while simultaneously allowing unchecked viral replication and the propagation of a damaging inflammatory state.

Diagnostic Strategies for SVCV Detection and Surveillance

The accurate and timely detection of Spring Viremia of Carp Virus (SVCV) constitutes the cornerstone of effective disease management, quarantine protocols, and international trade in cyprinid fish. As a notifiable pathogen under the World Organisation for Animal Health (WOAH) Aquatic Animal Health Code, SVCV demands diagnostic approaches that are not only exquisitely sensitive and specific but also robust enough to be deployed across diverse laboratory settings and epidemiological contexts [7, 15]. The diagnostic landscape for SVCV has evolved considerably from traditional virus isolation and conventional endpoint PCR, driven by the imperative to detect subclinical, carrier-state infections and to differentiate between viral genotypes with varying pathogenic potential [11, 15]. The selection of an appropriate diagnostic strategy is therefore a multi-factorial decision, influenced by the purpose of testing (e.g., routine surveillance, confirmatory diagnosis, or pre-movement screening), the stage of infection, the tissues sampled, and the required throughput.

Molecular Diagnostics: The Cornerstone of Modern Detection

Reverse transcription polymerase chain reaction (RT-PCR) and its quantitative variant (RT-qPCR) have become the workhorses of SVCV diagnostics, offering unparalleled analytical sensitivity and rapid turnaround times compared to traditional cell culture. The WOAH-recommended method has historically been a two-step semi-nested RT-PCR, but contemporary research has systematically refined these protocols to overcome inherent limitations.

Evolution of RT-PCR Methodologies

The conventional two-step semi-nested RT-PCR, while functional, is labor-intensive, consumes significant reagent volumes, and presents a substantial risk of carry-over contamination due to the multiple open-tube handling steps required for the nested reaction. A seminal comparative analysis demonstrated that substituting this process with a one-step semi-nested RT-PCR, where reverse transcription and primary amplification are combined in a single tube, followed by a single secondary nested PCR step, yields a dramatic improvement in analytical sensitivity [27]. Specifically, the one-step method detected viral RNA at a 10⁻⁵ dilution, representing a 1000-fold increase in sensitivity over the two-step method, which detected only up to a 10⁻² dilution [27]. This enhanced sensitivity translated directly into improved clinical detection rates, increasing from 84.2% to 91.7% in field-derived fish tissue samples [27]. The operational advantages of this approach, reduced processing time, minimized handling steps, and lower contamination risk, make it a compelling candidate for adoption as a high-throughput diagnostic tool and potential revision of future WOAH guidelines [27].

Real-Time Quantitative RT-PCR (RT-qPCR) Platforms

For definitive diagnosis, quantification of viral load, and high-throughput surveillance, RT-qPCR assays provide the gold standard. The development of these assays requires rigorous design, targeting highly conserved genomic regions to ensure detection of all known SVCV genotypes (Ia through Id) while maximizing specificity.

A landmark study by Zhu et al. conducted a whole-genome comparison of 24 representative SVCV strains and designed a novel primer-probe set (Cefas AR) targeting a highly conserved region of the L (RNA-dependent RNA polymerase) gene [7]. This assay underwent exhaustive validation in accordance with the WOAH Manual of Diagnostic Tests for Aquatic Animals. The performance metrics are exemplary: a limit of detection (LOD) of 1.28 copies/μL, a diagnostic sensitivity of 100% for cell-culture isolates and 96.6% for tissue samples, and a diagnostic specificity of 100% [7]. The assay demonstrated strong reproducibility and consistency across a multi-laboratory ring trial involving nine independent facilities, underscoring its robustness for international reference applications [7].

Concurrently, Clouthier et al. developed and analytically validated two alternative RT-qPCR tests targeting the nucleoprotein (Q2N) and glycoprotein (Q1G) genes [15]. These tests were designed to be pan-specific for SVCV genogroups Ia through Id. The Q2N assay demonstrated broader analytical specificity, although it showed occasional detection of pike fry sprivivirus (PFSV), while the Q1G assay failed to detect one specific SVCV isolate (20120450), highlighting the critical importance of multi-target or carefully validated single-target approaches [15]. Both assays, however, were highly sensitive, with 50% LODs at three plasmid copies and high repeatability [15].

The diagnostic accuracy of these RT-qPCR methods has been rigorously evaluated against the traditional gold standard of virus isolation (VI) using Bayesian latent class models, which circumvent the absence of a perfect reference standard. In a study involving koi populations with varying SVCV prevalence, the Q1G and Q2N assays demonstrated diagnostic sensitivity (DSe) estimates exceeding 96% and 98%, respectively, whereas VI showed a DSe of only approximately 60% [18]. Diagnostic specificity (DSp) estimates were comparable across all three tests, ranging from 79-97% [18]. Further precision analysis revealed that both Q1G and Q2N tests exhibited high repeatability (within-laboratory agreement) and reproducibility (between-laboratory agreement), with Gwet's AC1 values consistently above 0.74 when using kidney tissue [47]. Critically, this work demonstrated that tissue selection is paramount; precision estimates dropped significantly when using brain tissue or kidney-brain tissue pairs, reinforcing the recommendation to use kidney tissue for RT-qPCR-based SVCV surveillance [47].

Alternative Molecular Methods: Isothermal Amplification

For field-deployable or point-of-care diagnostics where qPCR thermocyclers are unavailable, isothermal amplification technologies offer a compelling alternative. A real-time reverse transcription recombinase polymerase amplification (RT-RPA) assay targeting the M gene has been developed. This assay operates at a constant 39°C, providing a result in just 20 minutes [23]. It demonstrated a detection limit of 10² copies per reaction, no cross-reactivity with other major fish viruses, and perfect concordance with real-time RT-PCR when testing 65 clinical samples [23]. The simplicity and speed of RT-RPA make it an ideal candidate for on-site screening at aquaculture facilities or in resource-limited settings.

Traditional and Complementary Diagnostic Approaches

Despite the ascendency of molecular methods, virus isolation in cell culture remains an important tool, particularly for virus characterization, archiving, and virulence studies. The standard cell line for SVCV isolation is Epithelioma Papulosum Cyprini (EPC), which supports robust cytopathic effect (CPE) development, typically characterized by rounding, detachment, and syncytia formation [25, 26]. However, as the Bayesian analysis revealed, the diagnostic sensitivity of VI is substantially lower than RT-qPCR, likely due to the requirement for infectious virus and interference from the host immune response or tissue toxicity [18]. VI also suffers from lower repeatability and reproducibility compared to molecular tests [47].

Histopathological and immunohistochemical examinations provide supportive evidence but are not primary diagnostic modalities. Characteristic lesions, including hemorrhages, necrosis in hematopoietic tissues (kidney and spleen), and enteritis, are hallmarks of SVCV infection [2, 26]. Advanced techniques such as in situ hybridization or immunofluorescence using antibodies against SVCV proteins (e.g., G or N) can confirm infection in formalin-fixed tissues and are valuable for pathogenesis studies [3, 28].

Integrated Surveillance Strategy

An effective surveillance program for SVCV cannot rely on a single test. The biology of SVCV, including its ability to establish persistent, subclinical infections, necessitates a layered diagnostic strategy. RT-qPCR, particularly the highly validated Cefas AR assay, is the optimal first-line screening tool due to its superior sensitivity and high throughput [7]. A positive RT-qPCR result, especially in a healthy-looking population, should be interpreted in context. Given the possibility of detecting non-infectious viral RNA, follow-up testing with virus isolation is recommended for regulatory confirmation or to obtain a live isolate for further characterization [18]. The recent discovery that high-virulence SVCV isolates can persist in koi for at least 167 days post-exposure, with higher titers and prevalence than low-virulence strains, adds further nuance [15]. This suggests that RT-qPCR surveillance may detect not only acute infections but also carriers, which is critical for controlling the long-distance spread of the virus via movement of apparently healthy fish.

The diagnostic approach must also account for host species and age. While common carp and koi are primary targets, SVCV has a broad host range, including species such as quillback (Carpiodes cyprinus) and Percocypris pingi [13, 26]. Any surveillance strategy should therefore be inclusive of all susceptible species within a water body. The route of infection also influences pathology; bath immersion, mimicking natural exposure, triggers a persistent pro-inflammatory response, while injection models may yield different quantitative results in terms of viral RNA load [3]. Ultimately, a harmonized diagnostic framework that combines highly sensitive molecular screening (RT-qPCR) with confirmatory culture, performed on the optimal tissue (kidney), and interpreted in light of prevalence and host population dynamics, is essential for the global control of this economically devastating pathogen.

In Vivo and In Vitro Models for Studying SVCV Pathogenesis

The multifaceted pathogenesis of Spring Viremia of Carp Virus (SVCV) necessitates a diverse arsenal of experimental models, ranging from established immortalized cell lines to sophisticated genetically modified whole-organism systems. These models are indispensable for dissecting the molecular mechanisms of viral entry, replication, immune evasion, and the host’s innate and adaptive antiviral responses. The selection of a particular model must be guided by the specific biological question under investigation, as each system offers distinct advantages and inherent limitations. Collectively, the integration of in vitro and in vivo approaches has yielded paradigm-shifting insights into the molecular and cellular underpinnings of SVCV-induced pathology, informing both fundamental virology and the development of prophylactic and therapeutic interventions for this economically critical pathogen, which is listed as a notifiable disease by the World Organisation for Animal Health (WOAH).

In Vitro Models: The Cell Culture Foundation for Mechanistic Dissection

In vitro cell culture models serve as the cornerstone for high-resolution mechanistic studies, enabling the precise control of experimental variables and the detailed characterization of virus-host interactions at the molecular level. The epithelioma papulosum cyprini (EPC) cell line, derived from carp epidermal herpesvirus-induced hyperplasia, remains the gold standard for SVCV propagation and titration, as recommended by WOAH diagnostic manuals [7, 27]. EPC cells are highly permissive to SVCV infection, developing a characteristic and reproducible cytopathic effect (CPE) that allows for quantification of viral infectivity via TCID₅₀ and plaque assays [7, 25]. Beyond simple propagation, EPC cells have been instrumental in elucidating critical host-pathogen interactions. For instance, studies utilizing EPC cells have demonstrated that SVCV infection triggers a complex interplay of cellular stress responses, including the induction of both autophagy and apoptosis through endoplasmic reticulum (ER) stress pathways [5]. Furthermore, EPC models were pivotal in defining the role of the viral glycoprotein (G) in inducing pyroptosis, a pro-inflammatory form of programmed cell death. Mechanistic investigations in EPC cells revealed that the SVCV-G protein activates the NLRP3 inflammasome, leading to gasdermin Eb-dependent pyroptosis, a process dependent on the host protein voltage-dependent anion channel 2 (VDAC2) [1]. The tractability of EPC cells for transfection and gene silencing has also enabled detailed mapping of antiviral signaling cascades. Overexpression studies in EPC cells have confirmed the role of key pattern recognition receptors (PRRs) and signaling adaptors, demonstrating that molecules such as common carp STING (CcSTING), MDA5, and LGP2 mediate potent antiviral responses by inducing type I interferon (IFN) and downstream interferon-stimulated genes (ISGs), thereby restricting SVCV replication [42, 44]. Similarly, the EPC system has been indispensable for evaluating the antiviral efficacy of novel compounds, such as the phenylpropanoid derivative N6 and the coumarin derivative D5, providing the first critical evidence of their ability to inhibit viral protein expression and reduce apoptotic cell death [19, 38].

Other teleost cell lines, including the fathead minnow (FHM) and zebrafish embryonic fibroblast (ZF4) lines, offer complementary advantages. FHM cells have been particularly valuable for studying the early stages of infection, notably viral entry. Tandem affinity purification (TAP) coupled with mass spectrometry using FHM cells identified prohibitin (PHB) and 14-3-3β/α-A as critical host factors that interact with the SVCV-G protein to facilitate viral attachment and entry [28, 31]. Knockdown or inhibition of PHB in FHM cells markedly reduced SVCV binding, while overexpression conferred susceptibility to otherwise non-permissive cell types, establishing a functional role for this protein in viral tropism [28]. ZF4 cells, derived from zebrafish embryos, provide a bridge between in vitro mechanistic studies and the powerful in vivo genetic toolkit of the zebrafish model. Studies in ZF4 cells have been employed to confirm the antiviral activity of host factors like IRF7, STING, and MAP2K7 [42, 51], and to investigate the proviral role of host proteins such as the autophagy receptor optineurin (OPTN), which promotes viral replication by enhancing mitophagy and simultaneously suppressing the interferon response [33]. Moreover, the use of primary cell cultures, such as carp head kidney (cHK) primary cells, provides a more physiologically relevant ex vivo system to study immune cell-specific responses, as demonstrated by the analysis of SVCV-induced IL-10 expression in these cells [29].

In Vivo Models: From Natural Hosts to Genetically Tractable Systems

While in vitro models are essential for mechanistic clarity, they cannot recapitulate the complex, multi-systemic interactions of a whole organism, including tissue-specific tropism, immune cell trafficking, and systemic physiological responses. Therefore, in vivo models are critical for understanding SVCV pathogenesis in its full complexity. The natural host, the common carp (Cyprinus carpio), remains the most relevant model for studying disease progression, transmission, and vaccine efficacy against SVCV. Challenges with SVCV in common carp can be performed via intraperitoneal (i.p.) injection, bath immersion, or cohabitation to model different routes of natural infection [6, 24]. These models have been instrumental in defining the age-dependent susceptibility of carp, with young fish (e.g., 3 months) exhibiting high mortality, while older fish (e.g., 9 months) become resistant [24]. The common carp model has also been central to vaccine development, where DNA vaccines encoding the SVCV-G protein have been shown to confer up to 100% protection against subsequent lethal bath challenge, inducing robust virus-specific B and T cell responses [24]. Furthermore, multi-omics analyses using common carp have revealed a trade-off between growth performance and disease resistance, with fast-growing populations exhibiting lower survival rates upon SVCV challenge, likely due to differential expression of genes related to nutrient metabolism, detoxification, and immune regulation [6]. Histopathological examination of infected carp tissues has confirmed that SVCV causes severe damage to mucosal barriers, compromising the architecture of gill and intestinal tissues and stimulating mucous cell proliferation [2]. The common carp model is also indispensable for evaluating the efficacy of antiviral drugs and immunostimulants, with compounds like the phenylpropanoid derivative S3 and yeast β-glucan demonstrating significant reductions in viral load and fish mortality [8, 14].

The zebrafish (Danio rerio) has emerged as a transformative model for SVCV research, offering a unique combination of genetic tractability, optical transparency for advanced imaging, and a fully sequenced genome. Zebrafish are highly susceptible to SVCV infection, which can be established via bath immersion of larvae or i.p. injection of adults [3, 21]. The generation of a reverse genetics system for SVCV, leading to recombinant viruses (rSVCV) expressing fluorescent proteins like mCherry, has been a landmark achievement. This tool has allowed for the direct visualization of viral pathogenesis in vivo, revealing that the fins are the primary sites of initial replication and entry following bath immersion [3]. This model has also uncovered the critical role of neutrophils in the antiviral response, showing that they are rapidly recruited to infection foci but their persistent activation contributes to uncontrolled inflammation, tissue damage, and ultimately death [3]. This persistent pro-inflammatory response, characterized by high IL-1β expression, was shown to be dependent on the route of infection, with bath immersion triggering a more robust inflammatory profile than i.p. injection [3].

The genetic tractability of zebrafish has allowed for the functional validation of numerous host factors identified in in vitro screens. CRISPR/Cas9-mediated knockout and morpholino-based knockdown approaches have conclusively demonstrated the in vivo role of specific genes. For example, VDAC2 knockdown in zebrafish larvae reduced SVCV-induced pyroptosis and tissue damage, resulting in increased survival, directly corroborating findings from EPC cells [1]. Similarly, zebrafish lacking maoc1 or trim2b were significantly more susceptible to SVCV infection, exhibiting higher viral loads and mortality, confirming the antiviral roles of these proteins [48, 50]. Conversely, knockout of negative regulators like irf2a in zebrafish conferred enhanced resistance to SVCV, validating its role as a suppressor of the interferon response [35]. The zebrafish model also allows for the investigation of viral immune evasion strategies at the organismal level. It was shown that SVCV infection stabilizes hypoxia-inducible factor 1α (hif1α) by interacting with the G protein, thereby activating a hypoxia response and promoting glycolysis, a state that favors viral replication; pharmacological inhibition of HIF1α with the inhibitor PX478 enhanced antiviral immunity in zebrafish [49]. Furthermore, the use of transgenic zebrafish lines with fluorescently labeled immune cells has provided unprecedented insights into the spatiotemporal dynamics of the host response, demonstrating the recruitment of neutrophils and the persistence of inflammation [3]. The zebrafish has also proven valuable as a high-throughput platform for screening antiviral compounds, with compounds like the imidazole coumarin derivative D5 and red elemental selenium showing significant protective efficacy by reducing mortality and viral titers [12, 52].

References

[1] Li C, Zhao W, Gao Y, Lu Y, Ye J, Liu X. Pivotal role of voltage-dependent anion channel 2 in pyroptosis induced by spring viremia of carp virus in fish cells.. Journal of Immunology. 2025. DOI: https://doi.org/10.1093/jimmun/vkaf154

[2] Ping O, Li Q, Liu S, Li Y, Li S, Zhou Y, et al.. Histopathology and transcriptome profiling reveal features of immune responses in gills and intestine induced by Spring viremia of carp virus.. Fish and Shellfish Immunology. 2024. DOI: https://doi.org/10.1016/j.fsi.2024.109726

[3] Souto S, Lama R, Mérour E, Mehraz M, Bernard J, Lamoureux A, et al.. In vivo multiscale analyses of spring viremia of carp virus (SVCV) infection: From model organism to target species. PLoS Pathogens. 2024. DOI: https://doi.org/10.1371/journal.ppat.1012328

[4] Baek E, Jeong Y, Kim G, Kim MJ, Kim K. Effects on viral suppression and the early-immune expression of ribavirin against spring viremia of carp virus in vitro.. Developmental and Comparative Immunology. 2024. DOI: https://doi.org/10.1016/j.dci.2024.105145

[5] Jiao X, Lu Y, Wang B, Guo Z, Qian A, Li Y. Infection of epithelioma papulosum cyprini (EPC) cells with spring viremia of carp virus (SVCV) induces autophagy and apoptosis through endoplasmic reticulum stress.. Microbial Pathogenesis. 2023. DOI: https://doi.org/10.1016/j.micpath.2023.106293

[6] Lei K, Li Q, Zhou J, Deng Y, Ping O, Feng Y, et al.. Mechanistic investigation into the differences in growth performance and resistance to spring viremia of carp virus in common carp. Frontiers in Immunology. 2026. DOI: https://doi.org/10.3389/fimmu.2026.1721974

[7] Zhu P, Sun J, Liao L, Zuo Z, Rice A, Gui S, et al.. Development and partial validation of an RT-qPCR assay for the rapid detection of spring viremia of carp virus (SVCV). Frontiers in Microbiology. 2026. DOI: https://doi.org/10.3389/fmicb.2025.1726705

[8] Liang H, Li Y, Li M, Zhou W, Chen J, Zhang Z, et al.. The effect and underlying mechanism of yeast β-glucan on antiviral resistance of zebrafish against spring viremia of carp virus infection. Frontiers in Immunology. 2022. DOI: https://doi.org/10.3389/fimmu.2022.1031962

[9] Wang Y, Chen Y, Ji J, Fan D, Lin A, Xiang L, et al.. Negative Regulatory Role of the Spring Viremia of Carp Virus Matrix Protein in the Host Interferon Response by Targeting the MAVS/TRAF3 Signaling Axis. Journal of Virology. 2022. DOI: https://doi.org/10.1128/jvi.00791-22

[10] Zhang C, Zhang P, Guo S, Zhao Z, Wang G, Zhu B. Dual-Targeting Polymer Nanoparticles Efficiently Deliver DNA Vaccine and Induce Robust Prophylactic Immunity against Spring Viremia of Carp Virus Infection. Microbiology spectrum. 2022. DOI: https://doi.org/10.1128/spectrum.03085-22

[11] Emmenegger E, Bueren EK, Jia P, Hendrix N, Liu H. Comparative virulence of spring viremia of carp virus (SVCV) genotypes in two koi varieties.. Diseases of Aquatic Organisms. 2022. DOI: https://doi.org/10.3354/dao03650

[12] Liu L, Hu Y, Lu J, Wang G. An imidazole coumarin derivative enhances the antiviral response to spring viremia of carp virus infection in zebrafish.. Virus Research. 2019. DOI: https://doi.org/10.1016/j.virusres.2019.01.009

[13] Katona R, Standish I, McCann R, Dziki S, Bailey J, Puzach C, et al.. Isolations of the Spring Viremia of Carp Virus in the Upper Mississippi River (USA), Including a New Host, the Quillback.. Journal of Aquatic Animal Health. 2022. DOI: https://doi.org/10.1002/aah.10153

[14] Song D, Liu L, Fu X, Liu G, Hu Y, Chen J. Immune responses and protective efficacy on 4-(2-methoxyphenyl)-3,4-dihydro-2H-chromeno[4,3-d] pyrimidine-2,5 (1H)-dione against spring viremia of carp virus in vivo. Aquaculture. 2021. DOI: https://doi.org/10.1016/J.AQUACULTURE.2021.736694

[15] Clouthier S, Schroeder T, Bueren EK, Anderson ED, Emmenegger E. Analytical validation of two RT-qPCR tests and detection of spring viremia of carp virus (SVCV) in persistently infected koi Cyprinus carpio.. Diseases of Aquatic Organisms. 2021. DOI: https://doi.org/10.3354/dao03564

[16] Jia S, Zhou K, Pan R, Wei J, Liu Z, Xu Y. Oral immunization of carps with chitosan-alginate microcapsule containing probiotic expressing spring viremia of carp virus (SVCV) G protein provides effective protection against SVCV infection.. Fish and Shellfish Immunology. 2020. DOI: https://doi.org/10.1016/j.fsi.2020.07.052

[17] Chen Y, Zhao M, Fan X, Zhu P, Jiang Z, Li F, et al.. Engagement of gcFKBP5/TRAF2 by spring viremia of carp virus to promote host cell apoptosis for supporting viral replication in grass carp.. Developmental and Comparative Immunology. 2021. DOI: https://doi.org/10.1016/j.dci.2021.104291

[18] Clouthier S, McClure C, Schroeder T, Anderson ED. Bayesian latent class model estimates of diagnostic accuracy for three test methods designed to detect spring viremia of carp virus.. Preventive Veterinary Medicine. 2021. DOI: https://doi.org/10.1016/j.prevetmed.2021.105338

[19] Song D, Liu L, Shan L, Qiu T, Chen J, Chen J. Rhabdoviral clearance effect of a phenylpropanoid medicine against spring viremia of carp virus infection in vitro and in vivo. Aquaculture. 2020. DOI: https://doi.org/10.1016/j.aquaculture.2020.735412

[20] Wang Z, Liu S, Guan H, Lu L, Tu J, Ouyang S, et al.. Structural and Functional Characterization of the Phosphoprotein Central Domain of Spring Viremia of Carp Virus. Journal of Virology. 2020. DOI: https://doi.org/10.1128/JVI.00855-20

[21] Guttula P, Rather M. Network analysis of Differentially Expressed Genes (DEGs) identified in zebrafish after infection with Spring viremia of carp virus (SVCV) – an in silico approach. bioRxiv. 2022. DOI: https://doi.org/10.1101/2022.03.01.482441

[22] Fouad A, Soliman H, Abdallah E, Ibrahim S, El-Matbouli M, Elkamel A. In-vitro inhibition of spring viremia of carp virus replication by RNA interference targeting the RNA-dependent RNA polymerase gene.. Journal of Virological Methods. 2019. DOI: https://doi.org/10.1016/j.jviromet.2018.10.008

[23] Cong F, Zeng F, Wu M, Wang J, Huang B, Wang Y, et al.. Development of a real-time reverse transcription recombinase polymerase amplification assay for rapid detection of spring viremia of carp virus.. Molecular and Cellular Probes. 2019. DOI: https://doi.org/10.1016/j.mcp.2019.101494

[24] Embregts C, Rigaudeau D, Veselý T, Pokorová D, Lorenzen N, Petit J, et al.. Intramuscular DNA Vaccination of Juvenile Carp against Spring Viremia of Carp Virus Induces Full Protection and Establishes a Virus-Specific B and T Cell Response. Frontiers in Immunology. 2017. DOI: https://doi.org/10.3389/fimmu.2017.01340

[25] Godahewa GI, Lee S, Kim J, Perera N, Kim M, Kwon M, et al.. Analysis of complete genome and pathogenicity studies of the spring viremia of carp virus isolated from common carp (Cyprinus carpio carpio) and largemouth bass (Micropterus salmoides): An indication of SVC disease threat in Korea.. Virus Research. 2018. DOI: https://doi.org/10.1016/j.virusres.2018.06.015

[26] Zheng L, Geng Y, Yu Z, Wang K, Ou Y, Chen D, et al.. First report of spring viremia of carp virus in Percocypris pingi in China. Aquaculture. 2018. DOI: https://doi.org/10.1016/J.AQUACULTURE.2018.04.056

[27] Park J, Shin I, Kim H, Lee ES, Choi E, Kim H, et al.. Improved Detection Sensitivity of Spring Viremia of Carp Virus by Substituting a Two-Step with a One-Step Nested Reverse Transcription Polymerase Chain Reaction Method. Microorganisms. 2025. DOI: https://doi.org/10.3390/microorganisms13122727

[28] Li C, Zhang W, Shi L, Lu Y, Ye J, Liu X. Prohibitin mediates the cellular invasion of spring viremia of the carp virus.. Fish and Shellfish Immunology. 2023. DOI: https://doi.org/10.1016/j.fsi.2023.108689

[29] Ping O, Tao Y, Wei W, Li Q, Liu S, Ren Y, et al.. Spring Viremia of Carp Virus Infection Induces Carp IL-10 Expression, Both In Vitro and In Vivo. Microorganisms. 2023. DOI: https://doi.org/10.3390/microorganisms11112812

[30] Romeih N, Abdallah E, Mahmoud M, Elkamel A, Fouad A. EXPRESSION PROFILE OF TUMOR NECROSIS FACTOR ALPHA DURING SPRING VIREMIA OF CARP VIRUS INFECTION IN NILE TILAPIA. Assiut veterinary medical journal. 2023. DOI: https://doi.org/10.21608/avmj.2023.190678.1121

[31] Chen B, Li C, Wang Y, Lu Y, Wang F, Liu X. 14‐3‐3&bgr;/&agr;‐A interacts with glycoprotein of spring viremia of carp virus and positively affects viral entry. Fish and Shellfish Immunology. 2018. DOI: https://doi.org/10.1016/j.fsi.2018.04.031

[32] Bello-Perez M, Medina-Gali R, Coll J, Perez L. Viral interference between infectious pancreatic necrosis virus and spring viremia of carp virus in zebrafish. Aquaculture. 2019. DOI: https://doi.org/10.1016/J.AQUACULTURE.2018.10.039

[33] Zhang Y, Li C, Zhang M, Qi S, Kong X. Autophagy receptor optineurin promotes spring viremia of carp virus replication via mitophagy and innate immune pathways in Cyprinus carpio.. International Journal of Biological Macromolecules. 2025. DOI: https://doi.org/10.1016/j.ijbiomac.2025.144309

[34] Wang X, Li Z, Zhang C, Jiang J, Han K, Tong J, et al.. Spring Viremia of Carp Virus N Protein Negatively Regulates IFN Induction through Autophagy-Lysosome-Dependent Degradation of STING.. Journal of Immunology. 2022. DOI: https://doi.org/10.4049/jimmunol.2200477

[35] Huang W, Zhao X, Ji N, Guo J, Feng J, Chen K, et al.. IRF2 Cooperates with Phosphoprotein of Spring Viremia of Carp Virus to Suppress Antiviral Response in Zebrafish. Journal of Virology. 2022. DOI: https://doi.org/10.1128/jvi.01314-22

[36] Zhou Y, Qiu T, Wang H, Hu L, Liu L, Chen J. Application of rhein as an immunostimulant controls spring viremia of carp virus infection.. Fish and Shellfish Immunology. 2023. DOI: https://doi.org/10.1016/j.fsi.2023.109128

[37] Zhao Z, Jiang F, Zhou G, Duan H, Xia J, Zhu B. Protective immunity against spring viremia of carp virus by mannose modified chitosan loaded DNA vaccine.. Virus Research. 2022. DOI: https://doi.org/10.1016/j.virusres.2022.198896

[38] Liu G, Wang C, Wang H, Zhu L, Zhang H, Wang Y, et al.. Antiviral efficiency of a coumarin derivative on spring viremia of carp virus in vivo.. Virus Research. 2019. DOI: https://doi.org/10.1016/j.virusres.2019.05.007

[39] Zhang C, Zhao Z, Liu G, Li J, Wang G, Zhu B. Immune response and protective effect against spring viremia of carp virus induced by intramuscular vaccination with a SWCNTs‐DNA vaccine encoding matrix protein. Fish and Shellfish Immunology. 2018. DOI: https://doi.org/10.1016/j.fsi.2018.05.029

[40] Zhang C, Li L, Wang J, Zhao Z, Li J, Tu X, et al.. Enhanced protective immunity against spring viremia of carp virus infection can be induced by recombinant subunit vaccine conjugated to single-walled carbon nanotubes.. Vaccine. 2018. DOI: https://doi.org/10.1016/j.vaccine.2018.08.003

[41] Chen X, Cai C, Li S, Shi Y, Zhang Q, Cheng G, et al.. pIgR-like4.2 enhances the antiviral immune response of zebrafish against spring viremia of carp virus.. Fish and Shellfish Immunology. 2025. DOI: https://doi.org/10.1016/j.fsi.2025.110350

[42] Liu R, Meng F, Li X, Li H, Yang G, Shan S. Characterization of STING from common carp (Cyprinus carpio L.) involved in spring viremia of carp virus infection.. Fish and Shellfish Immunology. 2023. DOI: https://doi.org/10.1016/j.fsi.2023.109164

[43] Liu R, Li H, Liu X, Liang B, Qi Y, Meng F, et al.. TRIM25 inhibits spring viremia of carp virus replication by positively regulating RIG-I signaling pathway in common carp (Cyprinus carpio L.).. Fish and Shellfish Immunology. 2022. DOI: https://doi.org/10.1016/j.fsi.2022.06.033

[44] Liu R, Niu Y, Qi Y, Li H, Yang G, Shan S. Transcriptome analysis identifies LGP2 as an MDA5-mediated signaling activator following spring viremia of carp virus infection in common carp (Cyprinus carpio L.). Frontiers in Immunology. 2022. DOI: https://doi.org/10.3389/fimmu.2022.1019872

[45] Wang F, Jiao H, Liu W, Chen B, Wang Y, Chen B, et al.. The antiviral mechanism of viperin and its splice variant in spring viremia of carp virus infected fathead minnow cells. Fish and Shellfish Immunology. 2019. DOI: https://doi.org/10.1016/j.fsi.2018.12.012

[46] Gao Y, Xiang Y, Wang F, Ye J, Lu Y, Ashraf U, et al.. The N protein of spring viremia of carp virus promotes the ubiquitination and degradation of viperin_sv1 to escape from the fish innate immunity. Aquaculture. 2021. DOI: https://doi.org/10.1016/J.AQUACULTURE.2021.736583

[47] Clouthier S, McClure C, Schroeder T, Aldous S, Allen J, Collette-Belliveau C, et al.. Measures of diagnostic precision (repeatability and reproducibility) for three test methods designed to detect spring viremia of carp virus.. Preventive Veterinary Medicine. 2021. DOI: https://doi.org/10.1016/j.prevetmed.2021.105288

[48] Fang H, Wu XM, Zheng S, Chang M. Tripartite motif 2b (trim2b) restricts spring viremia of carp virus by degrading viral proteins and negative regulators NLRP12-like receptors. Journal of Virology. 2024. DOI: https://doi.org/10.1128/jvi.00158-24

[49] Wang Z, Zhu C, Sun X, Deng H, Liu W, Jia S, et al.. Spring viremia of carp virus infection induces hypoxia response in zebrafish by stabilizing hif1α. Journal of Virology. 2024. DOI: https://doi.org/10.1128/jvi.01491-24

[50] Song Y, Fan S, Zhang D, Li J, Li Z, Li Z, et al.. Zebrafish maoc1 Attenuates Spring Viremia of Carp Virus Propagation by Promoting Autophagy-Lysosome-Dependent Degradation of Viral Phosphoprotein. Journal of Virology. 2023. DOI: https://doi.org/10.1128/jvi.01338-22

[51] Zhang C, Lu L, Li Z, Han K, Wang X, Chen D, et al.. Zebrafish MAP2K7 Simultaneously Enhances Host IRF7 Stability and Degrades Spring Viremia of Carp Virus P Protein via Ubiquitination Pathway. Journal of Virology. 2023. DOI: https://doi.org/10.1128/jvi.00532-23

[52] Tian J, Zhang Y, Zhu R, Wu Y, Liu X, Wang X. Red elemental selenium (Se0 ) improves the immunoactivities of EPC cells, crucian carp and zebrafish against spring viremia of carp virus (SVCV).. Journal of Fish Biology. 2020. DOI: https://doi.org/10.1111/jfb.14571