Tilapia Lake Virus

Taxonomy and Genomic Organization of Tilapia Lake Virus

Taxonomic Classification and Phylogenetic Placement

Tilapia lake virus (TiLV), also referred to as Tilapinevirus tilapiae, represents a highly divergent and emerging viral pathogen that has been taxonomically classified within the family Amnoonviridae, order Articulavirales [4, 10, 18, 27]. This classification places TiLV in a distinct phylogenetic lineage that is distantly related to the well-characterized orthomyxoviruses, such as influenza A virus, yet sufficiently divergent to warrant its own monotypic genus, Tilapinevirus [11, 18, 22, 27, 29]. The virus was first identified and characterized following outbreaks in Israel in 2014, and subsequent genomic and phylogenetic analyses have confirmed its unique position within the articulaviral clade [16, 18, 29]. The World Organisation for Animal Health (WOAH) has formally recognized TiLV as a listed disease, underscoring its significance as a transboundary pathogen of major economic and food security concern [24, 28]. The taxonomic assignment to the Amnoonviridae family is supported by the virus's fundamental genomic architecture, a segmented, negative-sense, single-stranded RNA genome, and by structural studies of its nucleoprotein (NP), which, despite being considerably smaller and lacking sequence homology to influenza NP, maintains a conserved topology comprising head and body domains with a positively charged RNA-binding groove [4, 10]. This structural conservation across the Articulavirales order suggests a common evolutionary origin for the core replication machinery, even as the accessory proteins have diverged dramatically [4, 10, 19].

Genomic Architecture: Segmented Negative-Sense RNA Genome

The TiLV genome is composed of ten discrete segments of linear, negative-sense, single-stranded RNA, a configuration that is characteristic of orthomyxo-like viruses but is notably smaller in total genomic size compared to influenza viruses [2, 13, 18, 27]. Each segment is encapsidated by the viral nucleoprotein (NP, encoded by segment 4) to form ribonucleoprotein (RNP) complexes, which serve as the functional templates for transcription and replication [4, 10, 19]. The total genome length is approximately 10.3–10.5 kb, with individual segments ranging in size from approximately 400 to 1,600 nucleotides [13, 18]. The segmented nature of the genome has profound implications for viral evolution, as it facilitates genetic reassortment during co-infection of a single cell with multiple TiLV strains, a phenomenon that has been documented in several epidemiological studies [16, 23]. Reassortment events have been detected in segment 3 and other genomic segments, indicating that mixed infections occur in farmed and wild tilapia populations and that this mechanism contributes to the generation of genetic diversity and potentially to the emergence of novel strains with altered virulence or host range [16, 23].

Segment-Specific Protein Coding and Functional Annotation

Each of the ten genomic segments encodes at least one major protein, and recent evidence has identified an 11th protein, designated S9-F3, which is translated from an alternative open reading frame (ORF) within segment 9 [13]. This discovery expands the known coding capacity of TiLV and highlights the complexity of its gene expression strategy. The functions of the proteins encoded by segments 1 through 10 have been partially elucidated, though the majority remain uncharacterized due to a lack of sequence homology to known viral or cellular proteins [13, 19].

• Segments 1, 2, and 3: These three segments encode the components of the viral RNA-dependent RNA polymerase (RdRp) complex. Segment 1 encodes the putative polymerase basic 1 (PB1) subunit, which contains conserved motifs characteristic of orthomyxoviral polymerases and is essential for RNA synthesis [16, 22]. Segments 2 and 3 encode additional polymerase subunits (PB2 and PA-like proteins), and together they form the heterotrimeric polymerase complex responsible for both transcription of viral mRNAs and replication of the genomic RNA [16, 31]. Phylogenetic analyses based on concatenated ORFs 1, 3, and 5 have been shown to provide the most reliable and fully supported tree topology for TiLV, underscoring the evolutionary conservation of these core replication proteins [16].

• Segment 4: This segment encodes the nucleoprotein (NP), which is the most extensively characterized TiLV protein at the structural level [4, 10, 19]. Cryo-electron microscopy (cryo-EM) studies have resolved the atomic structure of TiLV NP bound to RNA, revealing that it forms ring-like, pseudo-symmetrical oligomers that encapsidate the viral genome [4, 10]. The NP is composed of head and body domains with a positively charged groove that accommodates single-stranded RNA, and an oligomerization loop that inserts into a hydrophobic pocket of the adjacent NP monomer, allowing flexibility in the orientation of neighboring subunits [4, 10]. This structural organization is conserved across the Articulavirales order, despite the lack of sequence similarity between TiLV NP and influenza NP [4, 10]. Functional studies have confirmed that NP binds RNA in a sequence-independent manner and is essential for viral replication, and it also undergoes CRM1-dependent nuclear export, suggesting a role in trafficking viral RNPs between the nucleus and cytoplasm [19].

• Segment 5: The protein encoded by segment 5 (S5) has been localized predominantly to the cytoplasm of infected cells, but its precise function remains unknown [13]. Bioinformatic analyses have not revealed significant homology to any characterized proteins, and it is considered an accessory or structural protein of undetermined role [13].

• Segments 6 and 7: These segments encode proteins that have been implicated in the induction of protective immune responses. Recombinant proteins derived from segments 5 (specifically the S5196–272 region) and 6 (S6200–317) have been shown to elicit synergistic antibody responses and confer partial protection against TiLV challenge in vaccinated tilapia [8]. The S6 protein, in particular, appears to be a target of neutralizing antibodies, and co-immunization with both S5 and S6 fragments resulted in higher virus-neutralizing activity and relative percent survival (RPS) compared to single-segment vaccines [8].

• Segment 8: The S8 protein is predominantly cytoplasmic and has been shown to interact with the host cellular receptor coxsackievirus and adenovirus receptor (CAR) [9]. This interaction is believed to play a role in viral entry or intracellular trafficking, as overexpression of OnCAR in tilapia brain (TiB) cells significantly inhibited TiLV replication, genome copy number, and cytopathic effect [9]. The S8 protein, along with S10, appears to be a target of the host antiviral response mediated by CAR [9].

• Segment 9: This segment is notable for encoding two distinct proteins: the primary ORF product (S9) and an alternative reading frame product, S9-F3, which represents the 11th identified TiLV protein [13]. S9-F3 contains a predicted nuclear export signal (NES) and undergoes CRM1-dependent nuclear export, suggesting a role in regulating the subcellular localization of viral components or in modulating host nuclear functions [13]. Evolutionary analyses indicate that both S9 and S9-F3 are under selective pressure, and a homologue of S9-F3 has been identified in a TiLV-like virus infecting guppies, suggesting functional conservation [13].

• Segment 10: The S10 protein is predominantly nuclear in localization and, like S8, interacts with the host CAR protein [9, 13]. The S10 protein has been targeted in several vaccine development efforts, and DNA and recombinant protein vaccines based on S10 have demonstrated protective efficacy, particularly when combined with S9-based vaccines [21]. The TaqMan quantitative reverse transcription PCR (RT-qPCR) assay targeting a conserved region of segment 10 has been validated as a highly sensitive and specific diagnostic tool, with a limit of detection of 10 RNA copies per reaction and no cross-reactivity with other fish viruses [30].

Genetic Diversity, Phylogenetic Relationships, and Reassortment

Phylogenetic analyses of TiLV isolates from diverse geographic regions have revealed a moderate to high degree of genetic diversity, with isolates clustering into distinct clades that often correlate with geographic origin [5, 16, 23, 25]. Whole-genome sequencing using multiplexed RT-PCR and nanopore technology has enabled the recovery of complete TiLV genomes from clinical samples, water concentrates, and cell culture isolates, providing unprecedented resolution of the virus's evolutionary dynamics [25]. Phylogenetic trees constructed using concatenated ORFs 1, 3, and 5 have been shown to yield the most robust and fully supported topologies, and these analyses have confirmed that TiLV isolates from Israel, Thailand, India, Vietnam, and other countries form distinct lineages [5, 16, 23].

Reassortment is a major driver of genetic diversity in TiLV, as evidenced by the detection of multiple reassortment events among field isolates [16, 23]. For example, the Vietnamese isolate RIA2-VN-2019 was found to be genetically distant from other TiLV strains, and reassortment analysis identified seven potential events among 23 TiLV genomes, indicating that co-infection with multiple strains is common in aquaculture settings [23]. The ability of TiLV to undergo reassortment has important implications for vaccine development and disease control, as it could lead to the emergence of strains that escape vaccine-induced immunity or exhibit altered virulence [16, 23].

Host Genetic Factors and Quantitative Trait Loci

The genomic organization of TiLV is not limited to the viral genome itself; host genetic factors also play a critical role in determining susceptibility to infection. Genome-wide association studies (GWAS) in Nile tilapia (Oreochromis niloticus) have identified a major quantitative trait locus (QTL) on chromosome Oni22 that is strongly associated with resistance to TiLV [7, 26]. The most significant single nucleotide polymorphism (SNP) within this QTL has a substitution effect of 0.15, and tilapia homozygous for the resistance allele exhibit a mortality rate of only 11%, compared to 43% for those homozygous for the susceptibility allele [26]. Fine mapping and functional annotation of this QTL have identified several candidate genes, including lgals17, vps52, trim29, proteosome subunit beta type-9a, and ha1f, which are involved in host response to viral infection [7, 26]. These findings suggest that the genetic architecture of TiLV resistance is oligogenic, with a major QTL on Oni22 and additional moderate-effect QTLs on other chromosomes, such as Oni09 [7]. The identification of these genetic markers opens the door for marker-assisted selection (MAS) to breed TiLV-resistant tilapia strains, offering a sustainable, long-term strategy for disease management [7, 26].

Implications for Diagnostics, Vaccine Development, and Disease Control

The detailed understanding of TiLV taxonomy and genomic organization has direct translational applications. The conservation of segment 4 (nucleoprotein) and segment 10 across all known TiLV isolates has enabled the development of highly sensitive and specific molecular diagnostic assays, including RT-qPCR, droplet digital PCR (ddPCR), reverse transcription recombinase polymerase amplification (RT-RPA), and CRISPR-based detection platforms [1, 3, 6, 15, 30]. The multiplex PCR assay targeting segments 2, 3, and 8 minimizes the risk of false negatives due to genetic variability or reassortment [14]. Furthermore, the identification of protective epitopes within segments 1, 5, 6, 9, and 10 has guided the design of subunit vaccines, DNA vaccines, and nanoparticle-based immersion vaccines that have shown promising efficacy in laboratory and field trials [2, 8, 12, 17, 20, 21]. The structural resolution of the NP-RNA complex provides a foundation for structure-based antiviral drug design, targeting the conserved RNA-binding groove or oligomerization interface [4, 10]. Collectively, the genomic and taxonomic insights into TiLV are essential for developing robust surveillance programs, effective vaccines, and genetically resistant tilapia strains to mitigate the impact of this devastating pathogen on global aquaculture and food security.

Molecular Pathogenesis and Host–Virus Interactions of TiLV

The pathogenesis of Tilapia lake virus (TiLV) is a multifactorial process orchestrated by a sophisticated interplay between viral determinants of pathogenicity and the host’s innate and adaptive immune systems. As a member of the Amnoonviridae family within the Articulavirales order, TiLV is a segmented, negative-sense single-stranded RNA virus that shares fundamental replicative strategies with orthomyxoviruses like influenza, yet exhibits a unique set of molecular traits that drive its systemic and highly lethal pathology in tilapia [4, 10, 11]. Understanding the molecular basis of TiLV infection, from cellular entry and genome replication to immune subversion and tissue destruction, is critical for developing effective vaccines and antiviral strategies, particularly given the virus’s listing by the World Organisation for Animal Health (WOAH) as a significant transboundary pathogen affecting global food security [18, 24, 28, 42].

Viral Entry and Cellular Tropism

The initial step of TiLV infection is a cholesterol- and dynamin-dependent endocytic process that is independent of clathrin and endosomal acidification, distinguishing it from the classical entry mechanism of influenza A virus [27]. Studies using pharmacological inhibitors have demonstrated that dynamin activity is essential for productive TiLV entry, while depletion of membrane cholesterol with methyl-β-cyclodextrin severely abrogates viral protein synthesis and infectious particle production [27]. This pathway suggests that TiLV hijacks a specific endocytic route that may involve lipid rafts and cytoskeletal rearrangements, as inhibitors of actin and microtubule polymerization only partially blocked infection [27]. The virus does not require endosomal low pH for uncoating, a feature more akin to entry mechanisms used by some paramyxoviruses than by orthomyxoviruses [27].

Cellular tropism is a hallmark of TiLV pathogenesis, with the virus displaying a clear predilection for certain cell types that facilitates systemic dissemination. In vivo, immunofluorescence and flow cytometry studies have identified granulocytes as the primary target cells in peripheral blood, with infection rates reaching 46% at 5 days post-infection, while lymphocytes show profound depletion due to virus-induced apoptosis [34]. This tropism for granulocytes, cells typically associated with innate antibacterial defense, is paradoxical; despite high infection rates, granulocytes exhibit minimal apoptotic changes, suggesting they may serve as viral reservoirs or vehicles for systemic spread [34]. Conversely, lymphocytes are the most depleted leukocyte subpopulation, correlating with the severe lymphopenia that underlies TiLV-induced immunosuppression [34]. This differential tropism is a key pathogenic strategy that undermines both innate granulocyte functions and adaptive lymphocyte-mediated immunity.

At the organ level, TiLV exhibits broad tissue distribution, targeting the liver, brain, kidney, spleen, gills, and heart, with viral loads peaking in the liver during the acute phase of infection [40, 44, 46]. The liver is a primary site of pathology, where syncytial hepatocyte formation, a pathognomonic lesion, and multifocal necrotic hepatitis are consistently observed [18, 37, 40, 41, 46]. The brain is also heavily affected, leading to neurological signs such as lethargy and abnormal swimming behavior [40]. Quantitative PCR studies demonstrate that viral loads in target organs increase exponentially by day 3 post-infection and peak around 72 hours, followed by a decline in peripheral viral load despite ongoing mortality, indicating that immune dysregulation and tissue damage, rather than uncontrolled viral replication per se, drive lethality [44].

The cell surface coxsackievirus and adenovirus receptor (OnCAR) has been identified as a potential host restriction factor against TiLV. Overexpression of OnCAR in tilapia brain cells significantly inhibits viral replication, genome copy number, and cytopathic effect, largely through direct interaction with viral proteins encoded by segments 8 and 10 [9]. This suggests that OnCAR is a cellular surface molecule capable of interfering with TiLV entry or early replication, representing the first described anti-TiLV molecule with direct virus-binding properties [9]. The dynamic expression of OnCAR during TiLV infection in the liver and muscle further implies a host antiviral response that TiLV must overcome to establish infection [9].

Genome Replication, Nucleoprotein Structure, and the Viral Replication Complex

Central to TiLV replication is the nucleoprotein (NP), encoded by segment 4, which encapsidates the negative-sense RNA genome into protective ribonucleoprotein (RNP) complexes. Cryo-electron microscopy structures of TiLV NP bound to RNA have revealed a surprising degree of structural conservation with orthomyxoviral NPs, despite a lack of sequence homology [4, 10]. The TiLV NP is composed of a head and body domain separated by a positively charged groove that accommodates single-stranded RNA [4, 10]. An oligomerization loop inserts into a hydrophobic pocket of the neighboring NP, with flexible hinges allowing variable relative orientation between adjacent subunits [4]. Importantly, focused cryo-EM maps at up to 2.9 Å resolution unambiguously defined the 5′ to 3′ direction of bound RNA, and double-stranded A-form RNA regions were observed extruding from NP–NP interfaces, providing the first detailed description of RNA binding to an articulaviral NP [4, 10].

The viral RNA-dependent RNA polymerase is a heterotrimeric complex encoded by segments 1, 2, and 3, representing the PB1, PB2, and PA-like subunits, respectively [11, 19, 22]. Segment 3, encoding the PA-like subunit, is highly conserved and serves as a common target for molecular diagnostics [3, 5]. The polymerase complex is responsible for both transcription of viral mRNAs and replication of the genome. Recent proteomic characterization has identified an 11th protein, S9-F3, translated from an alternative reading frame of segment 9 [13]. Bioinformatic analysis and functional assays revealed that S9-F3 contains a nuclear export signal (NES) and undergoes CRM1-dependent nuclear export, a mechanism typical of many RNA viruses that shuttle viral RNP components between the nucleus and cytoplasm to coordinate replication [13]. The identification of an S9-F3 homologue in a TiLV-like guppy virus suggests that this alternative translation product is evolutionarily conserved and likely plays a critical role in the viral life cycle [13].

Viral proteins exhibit distinct subcellular localizations that reflect their functions. GFP-tagged constructs expressed in fish cells showed that segments 2 (polymerase subunit) and 10 are predominantly nuclear, while segments 1, 3, 5, 8, and S9-F3 are mostly cytoplasmic [13]. This nuclear-cytoplasmic shuttling equilibrium is critical for viral replication; the nucleoprotein itself, as demonstrated by immunofluorescence, can shuttle between the cytoplasm and nucleus, and inhibition of CRM1 leads to increased nuclear accumulation, suggesting that TiLV RNA is exported from the nucleus via a CRM1-dependent pathway [19]. This process likely ensures that newly replicated viral genomes are efficiently transported to the cytoplasm for packaging into progeny virions.

Mitochondrial Dysfunction and Cellular Cytopathicity

One of the most striking molecular findings in TiLV pathogenesis is the pronounced disruption of mitochondrial integrity and function. Transmission electron microscopy of E-11 cells infected with TiLV revealed the presence of viral particles in the cytoplasm as early as 1 hour post-infection, followed by progressive swelling of mitochondria and ultrastructural damage at 1, 3, and 6 days post-infection [45]. Functional analyses using JC-1 staining and MitoTracker Red probes confirmed a significant loss of mitochondrial mass and membrane potential (MMP) as early as day 1 post-infection [45]. The MTT assay further supported that cellular death during TiLV infection is mechanistically linked to mitochondrial disruption [45]. This mitochondrial damage is likely a major contributor to the extensive cell death observed in infected tissues, as the release of pro-apoptotic factors from damaged mitochondria initiates both apoptotic and necrotic pathways [45].

The mitochondrion is also a central hub for antiviral signaling, particularly via the interferon-inducible protein PKR and the retinoic acid-inducible gene I (RIG-I)-like receptor pathways. TiLV-mediated mitochondrial dysfunction could simultaneously cripple cellular energy production and short-circuit early antiviral signaling, allowing the virus to replicate rapidly before the host mounts an effective interferon response [11, 45]. Indeed, the temporal dynamics of viral load and innate immune gene expression show that immune activation (upregulation of MYD88, MCP, TNF-α) is initially responsive to viral replication but is transiently suppressed during peak viral replication, with a late-phase surge that correlates with host mortality rather than viral clearance [44]. This suggests that TiLV actively manipulates mitochondrial function to evade innate sensing.

Host Innate Immune Responses and Interferon Signaling

The innate immune system of tilapia is the first line of defense against TiLV, and the interferon (IFN) system plays a decisive role in restricting viral replication. The interferon regulatory factor 6 (OnIRF6) has been characterized as a positive regulator of type I IFN, specifically IFN-a3, in Nile tilapia. OnIRF6 is highly expressed in the gill, spleen, and head kidney, key immune organs, and is activated by both poly(I:C) and TiLV, leading to its translocation from the cytoplasm to the nucleus [35]. Overexpression of OnIRF6 in E11 cells significantly inhibited TiLV propagation, while in vivo silencing of OnIRF6 increased mortality, increased viral loads in tissues, and downregulated multiple components of the IFN and JAK-STAT pathways, including ISGs [35]. This clearly establishes OnIRF6 as a critical transcription factor in the anti-TiLV immune response [35].

Downstream of interferon induction, several interferon-stimulated genes (ISGs) are upregulated and contribute to the antiviral state. The Mx protein, a classical IFN-inducible dynamin-like GTPase, has been shown to be highly expressed in TiLV-resistant tilapia strains and correlates with lower viral loads and reduced mortality [49]. The radical s-adenosyl methionine domain containing 2 (RSAD2, also known as viperin) is another key antiviral effector that is upregulated in infected fish and acts by disrupting viral membrane integrity and replication [11, 40, 43]. Protein kinase R (PKR), activated by double-stranded RNA intermediates produced during viral replication, phosphorylates the translation initiation factor eIF2α, leading to global shutdown of host cap-dependent translation and inhibition of viral protein synthesis [11]. This pathway is a conserved component of the innate immune response in fish [11].

The interplay between viral replication and host innate immunity is also governed by pattern recognition receptors (PRRs). Toll-like receptors (TLRs), particularly TLR3 (which recognizes dsRNA), TLR5 (flagellin), and TLR9 (CpG DNA), and their signaling adaptor MyD88, are expressed in TiLV-infected fish and likely contribute to the induction of pro-inflammatory cytokines and type I IFNs [43]. The upregulation of TLR3 and myD88 in the spleen and liver of infected fish suggests that the virus triggers both MyD88-dependent and TRIF-dependent pathways [43]. Pro-inflammatory cytokines such as interleukin-8 (IL-8) and tumor necrosis factor-alpha (TNF-α) are also induced, but their expression profile is biphasic, early upregulation followed by suppression during peak viral replication, which may reflect viral immunomodulation [40, 44]. Co-infection with Aeromonas hydrophila further complicates the immune landscape, as feed additives that modulate immune responses can partially restore antiviral gene expression and improve survival [38].

Adaptive Immunity and Humoral Responses

Protective immunity against TiLV is achievable, as demonstrated by the survival of fish following natural outbreaks and the generation of TiLV-specific antibodies upon vaccination [8, 12, 33, 48]. The nucleoprotein (segment 4) is a major immunogen, and indirect ELISA using recombinant S4 protein can detect anti-TiLV IgM in serum and mucus, providing a tool for serosurveillance [33, 47]. Pooled serum testing with this ELISA offers a cost-effective approach for monitoring herd immunity at the farm level, although sensitivity is reduced when only one seropositive fish is present in a pool [33].

Passive immunization studies have convincingly demonstrated that antibodies alone can confer protection: sera from TiLV-vaccinated broodstock, when transferred to naïve juveniles, provided 85–90% protection against challenge [48]. This indicates that neutralizing antibodies are a critical component of protective immunity. Passive immunization with IgY antibodies generated in hens and targeting the S4 protein also showed dose-dependent neutralization activity, reducing TiLV infectivity from 5.01 × 10⁶ TCID50/mL to as low as 5.01 × 10⁴ TCID50/mL [32]. Immunofluorescence assays confirmed a significant reduction in TiLV antigen levels in IgY-treated cells, highlighting the potential of this approach as a therapeutic strategy [32].

Several major antigenic sites have been identified. The epitope S1³⁹⁹–⁴¹⁰ (TYTTRNEDFLPT) on segment 1, identified by phage display and structural analysis, induced robust antibody responses and afforded 81.8% survival in challenge studies [12]. Co-immunization with recombinant S5¹⁹⁶–²⁷² and S6²⁰⁰–³¹⁷ proteins synergistically enhanced S6-specific antibody production and neutralization, resulting in a relative percent survival (RPS) of 57.14%, whereas individual immunization provided no protection [8]. Similarly, combining segments 9 and 10 in DNA or recombinant protein vaccines yielded RPS values of 61% and 55.56%, respectively, compared to 16.67–61.11% for single-segment vaccines [21]. These findings indicate that multivalent vaccination strategies targeting multiple structural and non-structural proteins can elicit broader, more effective humoral responses [8, 21].

Maternally derived immunity also plays a role in protecting early life stages. Vaccination of tilapia broodstock with heat-killed or formalin-killed TiLV vaccines induced anti-TiLV IgM that was transferred to fertilized eggs and larvae [48]. Maternal antibodies were detectable in eggs and larvae, with higher levels observed after heat-killed vaccination, but these antibodies persisted only for 1–3 days post-hatching and were not detectable by day 7–14, indicating a narrow window of passive protection [48].

Immune Evasion and Subversion Strategies

TiLV employs multiple molecular strategies to evade host immune responses. The preferential infection and survival of granulocytes, which act as viral reservoirs, is a key evasion tactic that allows persistent virus replication in the face of innate immune activation [34]. The lack of apoptosis in infected granulocytes suggests that TiLV actively blocks programmed cell death in these cells, enabling their continued function as vehicles for systemic dissemination [34]. Concurrently, the induction of apoptosis in lymphocytes represents a direct immunosuppressive mechanism that depletes the adaptive immune system, delaying or preventing the development of effective cytotoxic and humoral responses [34].

Transcriptomic analysis of the liver following TiLV infection has revealed a complex pattern of gene expression that illustrates the host–virus arms race. Critical immune pathways are activated, including antigen processing, MAPK, apoptosis, chemokine signaling, and JAK-STAT pathways, yet the virus successfully subverts these responses [50]. For example, the upregulation of SOCS (suppressor of cytokine signaling), PI3K, and AKT in the JAK-STAT pathway, along with the downregulation of IFIT1 and TRIM25, suggests that TiLV can inhibit interferon signaling at multiple nodes [50]. NF-κB pathway modulation, including upregulation of NFKBIA (IκBα), further dampens the pro-inflammatory response, reducing the recruitment of immune cells to sites of infection [50].

The discovery that TiLV segment 9 encodes an alternative protein, S9-F3, that undergoes CRM1-dependent nuclear export, adds another layer of complexity to our understanding of viral immune evasion [13]. Nuclear export of viral proteins is often required to prevent the detection of viral nucleic acids in the nucleus by cytosolic sensors. By actively shuttling viral components out of the nucleus, TiLV may avoid activation of nuclear PRRs such as IFI16 or additional DNA/RNA sensors, thereby avoiding a branch of the innate immune response that is particularly important in fish [13]. The presence of selective pressure on S9 and S9-F3 across different TiLV isolates suggests that these sequences are critical for viral fitness and adaptation to the host immune environment [13].

Genetic Determinants of Host Resistance

Host genetics play a substantial role in determining the outcome of TiLV infection, with significant variation observed both within and between tilapia strains. A major quantitative trait locus (QTL) on chromosome Oni22 has been confirmed by multiple independent genome-wide association studies (GWAS) [7, 26]. This QTL explains a large proportion of the genetic variance for resistance to TiLV: fish homozygous for the resistance allele at the most significant SNP showed an 11% mortality rate compared to 43% for those with the susceptibility allele [26]. Fine mapping and functional annotation have narrowed the QTL region to a 10 Mb window containing 74 top markers, with candidate genes including proteasome subunit beta type-9a and ha1f, both involved in antigen processing and presentation [7]. Additional QTLs on chromosome Oni09 were also identified, suggesting an oligogenic architecture with both moderate- and small-effect loci [7].

Phenotypic resistance studies in different tilapia strains confirm that genetic background profoundly influences susceptibility. The ELM strain (originating from Lake Turkana) exhibited near-complete survival (100%) following cohabitation challenge, while the MAN and DRE strains suffered mortalities of 93.3% and 70.7%, respectively [49]. The resistance of the ELM strain correlated with lower viral loads in both mucosal and internal tissues, which was associated with a stronger mx1 antiviral response early in infection and a tempered pro-inflammatory response [49]. Similarly, among six Indonesian strains, the Gesit strain showed the highest resistance to TiLV, followed by Best and Red/Albino, while the Srikandi strain was most susceptible [39]. These strain-specific differences underscore the potential for marker-assisted selection (MAS) to improve TiLV resistance in breeding programs, offering a sustainable genetic solution to complement vaccination and biosecurity [7, 26].

Temperature also modulates host resistance. Mortality is exacerbated at higher water temperatures, with mass mortality peaking at 30°C, which is consistent with the optimal replication temperature for TiLV [36, 39]. Daily temperature fluctuations further increased virus-induced mortality among all tested strains, likely through stress-induced immunosuppression [39]. This temperature-dependent pathogenesis has implications for the geographic distribution and seasonality of TiLVD outbreaks.

Conclusion of the Pathogenesis Section

In summary, the molecular pathogenesis of TiLV involves a coordinated cascade of events: entry via a cholesterol- and dynamin-dependent, pH-independent endocytic route; replication mediated by an articulaviral polymerase and a structurally conserved nucleoprotein; targeted tropism for granulocytes and hepatocytes; mitochondrial destruction as a trigger for cell death; and sophisticated immune evasion through apoptosis of lymphocytes, suppression of interferon signaling via SOCS and NF-κB modulation, and CRM1-dependent nuclear export of viral proteins. The host fights back through IRF6-mediated interferon induction, Mx and RSAD-2 antiviral effectors, and neutralizing antibodies, but the virus’s ability to subvert these mechanisms frequently results in systemic disease and high mortality. The discovery of strain-specific resistance alleles and major QTLs provides a roadmap for genetic improvement, while ongoing advances in nanotechnology-based vaccines and immunostimulants offer practical countermeasures against this devastating pathogen.

Global Epidemiology and Transmission Dynamics of TiLV

The emergence of Tilapia lake virus (TiLV) represents one of the most significant infectious disease challenges to global aquaculture in the twenty-first century. Since its initial identification in 2014, this highly contagious pathogen has rapidly expanded its geographic footprint, establishing itself as a transboundary disease of critical importance to food security and economic stability in numerous developing nations. The World Organisation for Animal Health (WOAH) has formally recognized TiLV as a listed disease, underscoring its capacity to cause substantial morbidity and mortality in both farmed and wild tilapia populations [18, 24, 57]. The epidemiological trajectory of TiLV, characterized by intercontinental dissemination, persistent viral circulation in reservoir hosts, and complex transmission dynamics, demands rigorous scientific scrutiny to inform effective control strategies.

Historical Emergence and Geographic Expansion

The first documented outbreak of TiLV occurred in Israel in 2014, where mass mortality events in farmed tilapia prompted intensive diagnostic investigations that ultimately led to the discovery of a novel orthomyxo-like virus [18, 29, 42]. This initial report was rapidly followed by confirmations across multiple continents, with Ecuador and Colombia reporting outbreaks in South America, Egypt in Africa, and Thailand, Chinese Taipei, India, Malaysia, and Bangladesh in Asia [18, 29]. By 2023, TiLV had been officially reported in 17 countries, with evidence suggesting that the actual geographic distribution is substantially broader due to underreporting and limited surveillance capacity in many tilapia-producing regions [42, 52]. The rapid global spread of TiLV is attributable to several interconnected factors, including the extensive international trade of live tilapia for aquaculture, the movement of infected broodstock and fry, and the virus’s ability to establish subclinical infections in carrier fish that evade standard detection protocols [28, 29].

The epidemiological significance of TiLV is further magnified by its documented ability to infect multiple cichlid species and even non-cichlid hosts. While Nile tilapia (Oreochromis niloticus) and red hybrid tilapia (Oreochromis spp.) are the primary species affected, TiLV has been detected in wild tilapia populations and in tinfoil barbs (Barbonymus schwanenfeldii), giant gourami (Osphronemus goramy), and river barb, indicating a broader host range than initially appreciated [18, 24, 58]. This expanded host tropism has profound implications for viral persistence in aquatic ecosystems, as non-target species may serve as asymptomatic reservoirs that maintain viral circulation even in the absence of clinical disease in tilapia populations [58].

Prevalence and Incidence Across Endemic Regions

Comprehensive surveillance studies have revealed striking variability in TiLV prevalence across different geographic regions and production systems. In a massive surveillance effort spanning 11 districts of Tamil Nadu, India, molecular screening of 102 tilapia samples from 34 sites using semi-nested RT-PCR targeting segment 3 demonstrated an overall TiLV prevalence of 67.64% (69/102), with positive detections in both outbreak farms (10 of 13 sites with elevated mortality) and apparently healthy populations (13 of 21 asymptomatic sites) [5]. This finding is particularly concerning as it indicates widespread subclinical circulation of TiLV in the absence of overt disease, complicating efforts to estimate the true burden of infection and implement targeted control measures.

In the Indian Sundarbans, a UNESCO World Heritage site characterized by ecologically fragile aquatic ecosystems, surveillance of farmed tilapia revealed a more nuanced epidemiological picture. Among 51 samples collected from 6 disease cases and routine farm surveys, semi-nested PCR detected TiLV in 5.9% of samples in the first amplification step and 37.3% in the second step, with TiLV detected exclusively in O. niloticus (46.3% of this species) and in 37.3% of surveyed ponds [37]. The two-step semi-nested PCR demonstrated superior sensitivity compared to single-step SYBR green-based qPCR, highlighting the importance of diagnostic methodology in accurately estimating prevalence [37]. Histopathological examination confirmed the presence of syncytial hepatocytes, the pathognomonic lesion of TiLV infection, in diseased fish, providing definitive evidence of active viral replication and tissue damage [37].

In Indonesia, one of the world’s largest tilapia producers, TiLV detection rates have varied considerably across regions. In East Kalimantan Province, molecular analysis of 60 tilapia samples from three major aquaculture areas using conventional PCR revealed an overall prevalence of only 1.67%, with a single positive sample detected in Penajam Paser Utara [54]. This low prevalence may reflect genuine geographic variation in viral circulation, differences in diagnostic sensitivity, or the effectiveness of biosecurity measures in certain regions. Conversely, in East Java, semi-nested RT-PCR detected TiLV in 1 of 4 samples from pond-cultured tilapia, with the 250 bp amplicon confirming viral RNA presence [55]. In Yogyakarta, all tilapia exhibiting clinical signs consistent with TiLV infection tested positive by semi-nested PCR, suggesting that when clinical disease is apparent, viral detection rates are high [56]. A separate study in Indonesia detected TiLV in 2 of 3 fish (66.6%) showing suspected symptoms, further corroborating the association between clinical presentation and molecular confirmation [36].

Bangladesh presents a particularly instructive case study in TiLV epidemiology. Despite being designated by the Food and Agriculture Organization (FAO) in 2017 as being in a high-risk zone for TiLVD spread, a comprehensive investigation of 102 pooled organ samples from 34 ponds across 25 farms in 12 Upazilas conducted from May to October 2019 yielded no molecular evidence of TiLV by either conventional or real-time PCR [51]. Histopathological analysis also failed to identify pathognomonic TiLV lesions. This negative result may reflect genuine absence of the virus in the sampled regions, temporal variation in viral circulation, or limitations in sampling strategy. The authors appropriately recommended large-scale studies employing viral metagenomics to resolve the true TiLV status of Bangladeshi tilapia populations [51].

Transmission Pathways and Mechanisms

TiLV transmission occurs through both horizontal and vertical routes, each with distinct epidemiological implications for disease spread and persistence. Horizontal transmission, the primary mechanism of viral dissemination within and between populations, occurs through direct contact between infected and susceptible fish, exposure to contaminated water, and ingestion of infected tissues [18, 29]. The virus is shed in high concentrations from infected fish into the surrounding aquatic environment, where it can remain infectious for extended periods, particularly in closed or recirculating systems where viral accumulation is amplified [53].

Experimental cohabitation studies have provided critical insights into the efficiency of horizontal transmission. In controlled laboratory settings, naive fish placed in direct contact with TiLV-infected individuals rapidly acquire infection, with mortality rates varying substantially depending on viral dose, fish strain, and environmental conditions [39, 49]. The cohabitation model more accurately reflects natural transmission dynamics than intraperitoneal injection, as it encompasses the full spectrum of exposure routes, including waterborne transmission and direct contact [49]. In studies evaluating six Indonesian tilapia strains, cohabitation resulted in mortality ranging from 1.67% to 28.33%, compared to 16.67% to 61.67% following intraperitoneal injection, demonstrating that natural transmission is less efficient than direct inoculation but still capable of causing substantial mortality [39].

The role of water temperature in modulating transmission dynamics is increasingly recognized as a critical epidemiological factor. TiLV infection is most severe at temperatures above 25°C, with peak mortality occurring at approximately 30°C, where death rates can reach 50% [36]. Daily temperature fluctuations have been shown to significantly exacerbate virus-induced mortality patterns across different tilapia strains, suggesting that thermal stress may compromise host immune defenses and enhance viral replication [39]. These temperature-dependent effects have important implications for seasonal disease patterns and for predicting outbreak risk in different climatic zones.

Vertical transmission, the transfer of virus from infected broodstock to progeny, represents a particularly insidious mechanism for viral persistence and dissemination. TiLV has been detected in fertilized eggs and larvae derived from infected broodstock, confirming that the virus can be transmitted through the reproductive route [48]. Immunization of broodstock with inactivated TiLV vaccines elicits protective antibody responses that are passively transferred to eggs and larvae, with heat-killed vaccines demonstrating superior efficacy compared to formalin-killed preparations [48]. The presence of maternal antibodies in fertilized eggs and 1–3 day old larvae, but not in 7–14 day old larvae, indicates that passive immunity is short-lived, leaving early life stages vulnerable to infection once maternal antibody titers wane [48]. This window of susceptibility has critical implications for the timing of vaccination strategies and biosecurity interventions in hatchery settings.

Viral Shedding and Environmental Persistence

Quantification of viral shedding dynamics has provided essential data for understanding transmission risk and designing control measures. In recirculating aquaculture systems (RAS), TiLV concentrations in liver samples and rearing water are significantly higher during outbreak conditions compared to non-outbreak periods, with viral loads in water positively correlated with fish mortality [53]. This relationship between waterborne viral concentration and disease severity underscores the potential for environmental monitoring as a non-invasive surveillance tool. Droplet digital PCR (ddPCR) has demonstrated exceptional sensitivity for detecting TiLV in environmental samples, with the ability to quantify as few as 7.9 ± 0.99 copies per reaction in water samples, enabling detection of low-level viral contamination that may precede clinical outbreaks [15].

The persistence of TiLV in aquatic environments is influenced by multiple factors, including water temperature, pH, organic load, and the presence of UV radiation. In RAS operations, continuous water recirculation allows viral accumulation over time, leading to prolonged outbreaks and higher cumulative mortality compared to non-RAS systems where regular water discharge reduces viral loads and facilitates faster recovery [53]. This finding has direct practical implications for farm management, suggesting that water exchange protocols and disinfection strategies should be tailored to the specific production system to minimize viral amplification.

Reservoir Hosts and Subclinical Carriage

The identification of asymptomatic carriers represents one of the most challenging aspects of TiLV epidemiology. Wild tilapia and tinfoil barbs sampled from a lake in Malaysia that experienced a massive die-off event in 2017 continued to harbor TiLV for at least two years following the initial outbreak, with moderate to high prevalence of infection in the absence of clinical signs or mortality [58]. This persistent, subclinical carriage has profound implications for disease control, as apparently healthy fish may serve as mobile reservoirs that introduce virus into naive populations during translocation or natural movement. The isolation of TiLV from these wild fish using cell culture, with demonstration of cytopathic effects, confirmed that the detected viral RNA represents infectious virus capable of replication and transmission [58].

The phenomenon of subclinical infection is not limited to wild populations. In farmed tilapia, a substantial proportion of TiLV-positive fish may lack overt clinical signs, as demonstrated by the detection of TiLV in 13 of 21 asymptomatic farm sites in Tamil Nadu [5]. This subclinical carriage complicates efforts to estimate true prevalence and poses significant risks for disease introduction through the movement of apparently healthy stock. The development of a semi-quantitative lesion scoring system has enabled differentiation between clinical and subclinical infection, with clinical cases exhibiting significantly higher gross lesion scores and subclinical cases showing more pronounced histopathological alterations [46]. This scoring system provides a valuable tool for assessing disease severity and predicting outcomes in field settings.

Genetic Diversity and Reassortment

The segmented nature of the TiLV genome, comprising 10 negative-sense RNA segments, provides substantial capacity for genetic diversification through reassortment, a mechanism well-documented in other orthomyxoviruses such as influenza A virus. Phylogenetic analyses of TiLV isolates from diverse geographic regions have revealed substantial genetic diversity, with evidence of multiple reassortment events that may contribute to viral evolution and adaptation [16, 23]. A comprehensive phylogenetic study employing a multifactorial bioinformatics approach identified that concatenated open reading frames 1, 3, and 5 provide the most reliable and fully supported tree topology for phylogenetic inference, and confirmed reassortment events in segment 3 of an Israeli isolate [16].

The genetic relationship between TiLV isolates from different geographic regions provides insights into transmission pathways and viral movement. Phylogenetic analysis of TiLV segment 3 sequences from Tamil Nadu revealed close clustering with each other and with Israeli isolates, suggesting a potential transmission link between these geographically distant regions [5]. This finding is consistent with the hypothesis that international trade in live tilapia has facilitated the global dissemination of TiLV, with Israel potentially serving as an early source of viral introduction to South Asia.

Vietnamese TiLV isolates have demonstrated particularly high genetic diversity, with seven potential reassortment events detected among 23 TiLV genomes [23]. The Southern Vietnamese isolate RIA2-VN-2019 was found to be genetically distantly related to 21 reference isolates from other countries, while the Northern isolate HB196-VN-2020 showed closer affinity to international strains, suggesting multiple independent introductions of TiLV into Vietnam [23]. This pattern of multiple introductions and subsequent reassortment highlights the complexity of TiLV molecular epidemiology and the need for comprehensive genomic surveillance to track viral movement and evolution.

Epidemiological Modeling and Transmission Parameters

Quantitative epidemiological modeling has provided valuable insights into TiLV transmission dynamics and the potential impact of control interventions. A susceptible-infected-mortality disease model, parameterized using viral load dynamics data from experimental infections, estimated key epidemiological parameters including the transmission rate and basic reproduction number (R₀) [59]. Probiotic supplementation was found to significantly enhance immune responses (approximately 30% improvement), reduce disease transmission (approximately 80% reduction), and decrease R₀ (approximately 70% reduction), resulting in a substantially higher tolerance of farming densities (approximately 400-fold increase) in aquaculture populations [59]. These modeling approaches provide a mechanistic framework for predicting the population-level effects of interventions and optimizing management strategies.

The temporal dynamics of viral load and host immune responses have been characterized in experimental infection models. Following intraperitoneal injection of tilapia fingerlings with TiLV homogenate, systemic viral dissemination was confirmed across major organs, with viral load peaking at 72 hours post-infection before declining at 96 hours despite increasing mortality [44]. This paradoxical pattern, where viral load decreases while mortality continues to rise, suggests that immune dysregulation and immunopathology, rather than direct viral cytopathology alone, contribute significantly to disease pathogenesis [44]. The strong positive correlation between viral load and expression of innate immune genes (MYD88, MCP, TNF-α; r = 0.75–0.84) indicates that while immune activation is responsive to viral replication, it may be insufficient to prevent disease progression and may even contribute to tissue damage through excessive inflammation [44].

Risk Factors and Epidemiological Determinants

Multiple host, pathogen, and environmental factors modulate the epidemiology of TiLV infection. Host genetic background exerts a profound influence on susceptibility, with certain tilapia strains demonstrating markedly higher resistance to TiLV-induced disease. In a comparative study of six Indonesian tilapia strains, the Gesit strain exhibited the highest resistance to TiLVD, followed by Best, Red/Albino, Nirwana, Non-specific, and Srikandi [39]. The genetic basis of resistance has been further elucidated through genome-wide association studies, which identified a major quantitative trait locus (QTL) on chromosome Oni22 that explains a substantial proportion of the genetic variance in TiLV resistance [7, 26]. Fish homozygous for the resistance allele at the most significant SNP within this QTL had an average mortality rate of only 11%, compared to 43% for fish homozygous for the susceptibility allele [26]. This finding represents a rare example of a major QTL affecting a trait of critical importance in a farmed animal species and provides a foundation for marker-assisted selection programs aimed at enhancing TiLV resistance.

The identification of additional QTLs on chromosome Oni09 and other chromosomes suggests an oligogenic architecture underlying TiLV resistance, with several loci of moderate effect and many with small effect contributing to the overall phenotype [7]. Transcriptomic analyses have identified proteasome subunit beta type-9a and ha1f as potential causal genes within the Oni22 QTL, providing mechanistic insights into the molecular basis of resistance [7]. The interferon regulatory factor 6 gene (OnIRF6) has also been implicated in TiLV resistance, with in vivo silencing of OnIRF6 increasing mortality in TiLV-infected tilapia and downregulating multiple components of the interferon and JAK-STAT pathways [35].

Fish size and age represent additional risk factors, with smaller fish demonstrating greater susceptibility to TiLV infection. In a Vietnamese study, red hybrid tilapia weighing 4.5 ± 1.98 g experienced 92.5% mortality following TiLV challenge, compared to only 12.5% mortality in larger fish weighing 20.8 ± 7.5 g [23]. This age-dependent susceptibility has important implications for farm management, suggesting that juvenile fish require enhanced biosecurity protection and that vaccination strategies should target early life stages to maximize impact.

Implications for Surveillance and Control

The epidemiological patterns described above have direct implications for the design and implementation of TiLV surveillance and control programs. The high prevalence of subclinical infection in both farmed and wild populations necessitates the use of sensitive diagnostic tools capable of detecting low-level viral carriage. The development of rapid, field-deployable diagnostic assays, including RT-RPA combined with lateral flow dipsticks [1], RT-RAA-CRISPR-Cas12a systems [3], and one-pot RT-LAMP CRISPR/Cas12b platforms [6], represents a significant advance in surveillance capacity, enabling real-time detection of TiLV in resource-limited settings. These assays achieve detection limits as low as 9.10 copies per reaction [3] and 79.6 copies [6], with diagnostic sensitivities of 92% and specificities of 100% [6].

Pooled serum testing using indirect ELISA based on recombinant segment 4 protein offers a cost-effective approach for population-level surveillance, with pooling of five serum samples proving effective for detecting TiLV-specific antibodies when multiple seropositive individuals are present in the pool [33]. This strategy reduces testing costs while maintaining acceptable diagnostic performance, facilitating large-scale monitoring of immune status in tilapia populations.

The demonstration that vaccination can reduce both fish mortality and viral shedding into the environment [53] provides a compelling rationale for integrating vaccination into comprehensive control programs. Vaccinated fish showed substantially lower cumulative mortality (16.7%) compared to unvaccinated controls (37.7%), with a relative percent survival of 55.6%, and TiLV concentrations in pond water were significantly lower in vaccinated groups [53]. This reduction in environmental viral load has population-level benefits, decreasing infection pressure across the entire production system and potentially reducing the risk of outbreaks in neighboring farms.

The persistent detection of TiLV in wild fish populations [58] underscores the importance of biosecurity measures that prevent contact between farmed and wild fish, and highlights the limitations of farm-level control in the absence of coordinated regional and national strategies. The implementation of strict quarantine protocols, movement controls, and surveillance programs, as recommended by WOAH and FAO, remains essential for preventing the

Clinical Manifestations and Pathological Features of Tilapia Lake Virus Disease

Tilapia Lake Virus Disease (TiLVD) presents as a systemic, peracute to acute viral septicemia with a constellation of clinical signs, gross pathological lesions, and histopathological alterations that collectively define a pathognomonic disease profile. The severity and progression of these manifestations are governed by a complex interplay of viral virulence, host genetic susceptibility, developmental stage, environmental conditions, and the presence of concurrent infections. A comprehensive understanding of these features is not merely an academic exercise but a critical prerequisite for accurate field diagnosis, the development of effective control strategies, and the implementation of informed biosecurity measures.

Gross Clinical Manifestations and Behavioral Alterations

The clinical progression of TiLVD is remarkably consistent across reported outbreaks, though the intensity and speed of onset can vary. Infected fish typically exhibit a progression from subtle behavioral changes to overt signs of morbidity and mortality. Initial indicators include lethargy, anorexia, and a tendency to congregate near the water surface or at the pond edges, often accompanied by a "gasping" behavior suggestive of respiratory distress [5, 29]. Affected fish display abnormal swimming patterns, including loss of equilibrium, corkscrewing, and a generally sluggish or disoriented movement, which are strongly indicative of central nervous system involvement [24, 29]. Exophthalmia (pop-eye), a pronounced darkening or discoloration of the skin, and congestion of the gills are frequently reported external signs [5, 24]. As the disease progresses, more severe external lesions may become apparent, including open wounds, skin ulcers, and fraying or thinning of the fins [5, 24]. In many cases, the most striking clinical finding is the presence of severe anemia, manifesting as profoundly pale gills and a generalized pallor of the body [40].

Mortality is the hallmark of TiLVD, with rates often reaching 50-90% in affected populations, particularly among juveniles and fry [5, 18, 52, 56]. The onset of mortality can be explosive, occurring within days of the first clinical signs [29]. The specific mortality pattern is influenced by a multitude of factors. The infecting viral dose is critical; experimental studies have shown that higher titers lead to more rapid and complete mortality [23]. Water temperature is another key environmental modulator, with mass mortality peaking at approximately 30°C and outbreaks being more severe under conditions of daily temperature fluctuation [36, 39]. Importantly, the genetic constitution of the host plays a decisive role. Specific strains of Oreochromis niloticus, such as the ELM strain from Lake Turkana, have demonstrated near-complete resistance to clinical disease and mortality under cohabitation challenge, whereas other strains like MAN may experience mortality rates exceeding 90% [49]. This differential susceptibility is linked to a major quantitative trait locus (QTL) on chromosome Oni22, where fish homozygous for the resistance allele show dramatically lower mortality (11%) compared to those homozygous for the susceptibility allele (43%), underscoring the profound influence of host genetics on clinical outcome [7, 26]. Furthermore, fish size is inversely correlated with susceptibility, with smaller, younger fish being markedly more vulnerable to lethal infection than larger adults [23].

Gross Pathological Findings: The Macroscopic Signature of Systemic Disease

Necropsy of moribund or recently deceased fish reveals a consistent pattern of gross pathological lesions, primarily affecting organs of the reticuloendothelial and nervous systems. The liver is the most consistently and severely affected visceral organ. It is typically described as pale, friable, and often mottled, indicating widespread hepatocellular necrosis and lipid depletion [18, 40, 46]. The spleen is frequently dark, shrunken, and congested, reflecting the profound hematopoietic and lymphoid depletion that characterizes the disease [18, 40]. The gastrointestinal tract often exhibits a pale, fluid-filled intestine with catarrhal or mucoid content, suggesting enteritis and impaired absorptive function [40]. The kidney may appear swollen and congested. The brain, while not always showing obvious macroscopic lesions, is a primary target, and careful examination may reveal congestion or softening of the tissue [24]. The overall picture is one of a severe, multisystemic disease affecting the organs responsible for metabolism, immunity, and neural function.

Histopathological Features: The Pathognomonic Hallmark of TiLVD

The microscopic pathology of TiLVD is defined by a characteristic triad of lesions: severe necrotizing hepatitis with syncytial cell formation, lymphoid depletion in hematopoietic organs, and meningoencephalitis. These histopathological findings are so consistent and specific that they are considered the gold standard for confirming a clinical diagnosis of TiLVD.

Syncytial Hepatitis: The most pathognomonic lesion of TiLVD is found in the liver. Histological examination reveals a severe, multifocal to coalescing necrotizing hepatitis. The key finding is the presence of large, multinucleated syncytial cells (also known as fusion cells) formed by the fusion of adjacent hepatocytes [5, 18, 37, 40, 41, 46, 55]. These syncytia contain numerous, often pyknotic or karyolytic nuclei, and are surrounded by areas of massive hepatocellular necrosis and lysis. The loss of hepatocytes is profound, leading to a breakdown of normal hepatic architecture. In experimentally infected fish, these lesions are mild at early time points (e.g., 3 days post-challenge) but become severe and widespread by 7-14 days post-infection, correlating with peak viral loads and mortality [40]. The severity of these hepatic lesions has been systematically graded, providing a semi-quantitative tool for assessing pathogenicity and clinical stage [46].

Lymphoid Depletion and Splenic Pathology: TiLV infection results in a profound and systemic depletion of lymphocytes. In the spleen and head kidney, the primary hematopoietic and lymphoid organs of teleosts, histopathology reveals a marked loss of lymphoid tissue, a reduction in the number of red blood cells (indicating anemia), and a significant increase in the number and size of melanomacrophage centers [18, 34, 40]. Melanomacrophage centers are aggregates of pigmented macrophages that accumulate cellular debris, and their proliferation is a hallmark of chronic inflammation and immune system activation or exhaustion. This lymphoid depletion is a direct consequence of massive virus-induced apoptosis, particularly of lymphocytes, which contributes to the severe leukopenia observed in infected fish [34].

Meningoencephalitis and Brain Pathology: The brain is a major target organ for TiLV, consistent with the neurological signs observed clinically. Histological sections of the brain demonstrate a severe, non-suppurative meningoencephalitis. This is characterized by perivascular cuffing (infiltration of lymphocytes and other mononuclear cells around blood vessels), gliosis (proliferation of glial cells), and the presence of necrotic foci within the brain parenchyma [29, 41, 46, 55]. In some cases, syncytial cell formation, analogous to that seen in the liver, has also been observed within the brain tissue, further confirming the virus's ability to induce cell-cell fusion in neural cells [41]. The presence of virus in the brain is likely a major driver of the disorientation and abnormal swimming behaviors seen in moribund fish.

Pathology in Other Organs: The systemic nature of TiLV infection is further underscored by histopathological changes in other tissues. The gills often show congestion, lamellar fusion, and epithelial hyperplasia, which correlates with the observed respiratory distress and gasping behavior [46, 55]. The heart may show evidence of myocarditis, and the kidney may exhibit interstitial nephritis and tubular necrosis [46, 55]. The pancreas can also be affected, showing necrotic changes [29]. These findings confirm that TiLV is a pantropic virus capable of infecting and damaging a wide range of cell types beyond its primary targets in the liver, brain, and hematopoietic tissues.

Cellular and Ultrastructural Pathology: The Mechanism of Cell Death

At the cellular level, TiLV-induced pathology is intimately linked to the destruction of mitochondria. Transmission electron microscopy of TiLV-infected cells has revealed a progressive and severe disruption of mitochondrial architecture. Within hours of infection, mitochondria begin to swell, lose their internal cristae structure, and eventually become completely disrupted [45]. This mitochondrial damage is accompanied by a loss of mitochondrial mass and a collapse of the mitochondrial membrane potential, early hallmarks of the intrinsic apoptotic pathway [45]. The consequent depletion of cellular ATP and the release of pro-apoptotic factors from damaged mitochondria are believed to be the primary drivers of the widespread cell death observed in infected tissues, especially in the liver [44, 45]. This mechanism explains the rapid and extensive necrosis that distinguishes TiLVD from many other viral infections in fish.

Immunopathogenesis and Hematological Alterations

TiLV infection triggers a complex and often dysregulated host immune response that contributes significantly to disease pathology. The infection leads to a pronounced leukopenia, with a dramatic reduction in total white blood cell counts, driven primarily by the massive depletion of lymphocytes through virus-induced apoptosis [34]. This lymphopenia effectively cripples the adaptive immune response, facilitating uncontrolled viral replication and systemic dissemination. Paradoxically, while lymphocytes are being destroyed, other leukocyte populations are differentially affected. Flow cytometry studies have revealed that granulocytes are the primary cellular targets for TiLV infection, with up to 46% of these cells harboring the virus at peak infection [34]. Despite a high infection rate, these granulocytes show minimal signs of apoptosis, suggesting they may serve as viral reservoirs and vehicles for systemic dissemination [34]. This preferential tropism for granulocytes, coupled with the destruction of lymphocytes, reveals a sophisticated viral strategy for immune evasion. Hematological analysis consistently demonstrates severe anemia, characterized by significant reductions in hematocrit, hemoglobin concentration, and red blood cell counts [34, 40]. This anemia is a direct contributor to the pale appearance of the fish and likely exacerbates tissue hypoxia and organ failure.

The host transcriptional response to TiLV further illuminates the pathogenic process. Transcriptome analysis of the liver reveals the upregulation of thousands of genes involved in key immune pathways, including antigen processing and presentation, interferon signaling, and the JAK-STAT pathway, representing a robust attempt by the host to resist infection [50]. However, the virus actively subverts this response. The virus upregulates suppressors of cytokine signaling (SOCS) and downregulates critical antiviral effectors like IFIT1, effectively neutralizing the host's interferon-mediated antiviral state [50]. This manipulation of the host's immune signaling network allows for unchecked viral replication and the progression of pathological lesions. The expression of pro-inflammatory cytokines like IL-8 and interferon regulatory factors is strongly correlated with the severity of pathological changes and viral load, suggesting that the host's own inflammatory response, while intended to be protective, may ultimately contribute to tissue damage and disease severity [40, 44].

Pathological Scoring and Host Susceptibility

The clinical and pathological picture of TiLVD is not uniform. A semi-quantitative lesion scoring system has been developed to differentiate between clinical and subclinical infections [46]. Subclinically infected fish, while positive for TiLV by molecular methods, may show minimal or no external clinical signs and only mild histopathological lesions. In contrast, clinically affected fish display severe, high-grade lesions, particularly in the liver and brain, which are strongly correlated with high viral loads and mortality [46]. The existence of subclinical carriers is a major concern for disease management, as these fish can act as silent reservoirs for virus transmission within and between farms.

The genetic background of the host is a powerful determinant of pathological outcome. The identification of a major QTL for TiLV resistance on chromosome Oni22 provides a mechanistic basis for the observed strain-specific differences in disease severity [7, 26]. Resistant genotypes, such as the ELM strain, exhibit a superior ability to control viral replication, characterized by a more robust and rapid mx1-based antiviral response and a lower pro-inflammatory response, thereby minimizing immunopathological damage [49]. Conversely, susceptible genotypes experience uncontrolled viral replication, leading to the severe pathological lesions described above. This genetic variation underscores the potential for marker-assisted selection as a long-term strategy for improving disease resistance in farmed tilapia populations. The Food and Agriculture Organization of the United Nations (FAO) has recognized TiLV as a significant transboundary disease, and the World Organisation for Animal Health (WOAH) has included it in its list of notifiable diseases, highlighting its global importance for aquatic animal health and food security [24, 28].

Pathological Considerations in Coinfections and Natural Reservoirs

The pathological picture of TiLVD can be further complicated by the presence of coinfecting pathogens. Experimental coinfection models with Aeromonas hydrophila have demonstrated that TiLV-infected fish are more susceptible to secondary bacterial infections, and that such coinfections can exacerbate mortality and tissue damage [38]. Histological examination of such cases may reveal a combined pathology, including the characteristic TiLV-induced syncytial hepatitis alongside lesions of bacterial septicemia. The role of other fish species as reservoirs for TiLV is also an important pathological consideration. TiLV has been detected in asymptomatic wild tilapia and tinfoil barbs (Barbonymus schwanenfeldii) long after initial outbreaks, indicating that these species can serve as persistent carriers without showing clinical disease [58]. The pathological features in such carriers are likely minimal to absent, but their role in maintaining the virus in the environment is critical for the epidemiology of the disease.

Advances in Diagnostic Technologies: From RT-PCR to Field-Deployable Assays

The emergence of Tilapia Lake Virus (TiLV) as a transboundary pathogen of global significance has catalyzed an unprecedented acceleration in the development of diagnostic technologies. The World Organisation for Animal Health (WOAH) has formally recognized TiLV as a listed disease, underscoring the critical need for robust, sensitive, and accessible detection methods to support surveillance, outbreak management, and international trade of tilapia and their products [18, 24]. The diagnostic landscape for TiLV has evolved dramatically since the virus was first described in 2014, progressing from conventional molecular techniques to sophisticated, field-deployable platforms that promise to democratize disease detection across resource-limited settings. This section provides an exhaustive analysis of these technological advances, examining their underlying principles, analytical performance, operational requirements, and suitability for various surveillance contexts.

Foundational Molecular Diagnostics: Conventional and Real-Time RT-PCR

The cornerstone of TiLV detection has been reverse transcription polymerase chain reaction (RT-PCR), which targets the virus’s segmented negative-sense RNA genome. Early diagnostic efforts focused on amplifying conserved regions of segment 3, which encodes a component of the viral RNA-dependent RNA polymerase complex [18, 29]. This segment has proven remarkably stable across geographic isolates, making it an ideal target for broad-spectrum detection. The conventional RT-PCR assay, particularly the semi-nested format targeting a 250-base pair amplicon, has been widely adopted for surveillance and outbreak confirmation across Asia, Africa, and the Americas [36, 55, 56, 58]. The semi-nested approach, which employs two rounds of amplification with an internal primer set, substantially enhances analytical sensitivity compared to single-round PCR, enabling detection of low viral loads that might otherwise escape identification [37]. Indeed, studies comparing one-step and two-step semi-nested PCR have consistently demonstrated that the nested format detects TiLV in a significantly higher proportion of field samples, with one investigation reporting positivity rates of 5.9% in the first round versus 37.3% in the second round among farmed tilapia in the Indian Sundarbans [37].

The transition to real-time quantitative RT-PCR (RT-qPCR) represented a major advancement, offering quantification of viral load, enhanced throughput, and reduced risk of amplicon contamination through closed-tube detection [30, 31]. Multiple RT-qPCR assays have been developed targeting various genomic segments, with segments 1, 2, 3, 4, and 10 all demonstrating utility [30, 31]. Comparative evaluation of primer sets across all ten segments revealed that those targeting segments 1 through 4 exhibited the highest detection sensitivity, equivalent to 100.3 TCID₅₀/mL, while maintaining amplification efficiencies between 93% and 98% and excellent reproducibility, with inter-assay coefficients of variation below 5.9% [31]. A TaqMan-based RT-qPCR assay targeting segment 10 achieved a limit of detection of 10 RNA copies per reaction and demonstrated 96.7% diagnostic sensitivity and 100% diagnostic specificity across a validation panel of 91 proven-positive and 185 proven-negative samples [30]. Importantly, this assay detected TiLV strains from diverse geographic origins, including North America, South America, Africa, and Asia, confirming its suitability for global surveillance networks [30]. These quantitative assays have proven invaluable for understanding viral dynamics, revealing that TiLV loads peak at approximately 72 hours post-infection in experimentally challenged fish and correlate strongly with disease severity and mortality [30, 44].

Despite their robustness, both conventional and real-time RT-PCR assays present limitations for large-scale surveillance. They require expensive thermal cycling equipment, skilled personnel, stable electricity supplies, and cold chain maintenance for reagents. These constraints have motivated the development of alternative platforms better suited to field conditions.

Enhanced Sensitivity and Specificity: Droplet Digital PCR and Multiplex Approaches

The demand for absolute quantification of low-abundance viral targets, particularly in carrier fish and environmental samples, has driven the development of droplet digital PCR (ddPCR) for TiLV detection [15]. This partitioning-based technology compartmentalizes the sample into thousands of nanoliter-sized droplets, each undergoing independent PCR amplification. By counting the proportion of positive droplets at the endpoint, ddPCR provides absolute quantification without the need for standard curves. A TiLV-specific ddPCR assay demonstrated a detection limit of 3.3 copies per reaction, outperforming a comparator RT-qPCR by approximately 10-fold [15]. The assay exhibited perfect diagnostic sensitivity and specificity (100% for both) and showed no cross-reactivity with other tilapia pathogens, including Tilapia parvovirus, Infectious spleen and kidney necrosis virus, and several bacterial species [15]. Critically, ddPCR detected TiLV in water samples with viral loads as low as 7.9 copies per reaction, highlighting its potential for environmental surveillance and early warning systems [15].

Another significant advancement is the development of multiplex PCR assays targeting multiple genomic segments simultaneously [14]. This strategy addresses a critical vulnerability of single-target assays: the potential for false-negative results arising from genetic variability or viral reassortment, which is well-documented in TiLV epidemiology [16, 23]. A multiplex PCR targeting segments 2, 3, and 8 demonstrated consistent amplification of all three targets in a single reaction, with a detection limit of 100 pg/μL of TiLV cDNA and no cross-reactivity with non-TiLV pathogens [14]. Receiver operating characteristic analysis yielded a perfect area under the curve of 1.0, confirming flawless sensitivity and specificity [14]. This multi-segment approach provides an internal control against genetic drift and reassortment, as the probability of simultaneous mutation or loss of three conserved regions is vanishingly small. The assay’s design also facilitates genotypic characterization, as differential amplification patterns may indicate the emergence of novel variants [14].

Isothermal Amplification Technologies: RT-LAMP and RT-RPA

The need for rapid, equipment-independent diagnostics suitable for on-site deployment has stimulated intense interest in isothermal amplification technologies. These methods amplify nucleic acids at constant temperatures, eliminating the requirement for thermal cyclers and significantly reducing time to result. Loop-mediated isothermal amplification (LAMP) has been adapted for TiLV detection through reverse transcription LAMP (RT-LAMP), which amplifies viral RNA at a single temperature (typically 60-65°C) within 30-60 minutes [6, 18]. The assay targeting a conserved region of segment 4 achieves detection within 75 minutes at 62°C, with results visualized using a portable fluorescence viewer or through colorimetric indicators suitable for naked-eye interpretation [6]. Validation across 261 samples from nine source populations demonstrated 92% diagnostic sensitivity and 100% diagnostic specificity [6]. The 95% limit of detection was 79.6 copies, with no cross-reactivity to other fish RNA or DNA viruses [6]. The assay’s compatibility with non-lethal sampling using gill tissue, which showed reliable detection despite lower viral loads compared to internal organs, represents a significant welfare advantage for repeated monitoring of valuable broodstock [6].

Recombinase polymerase amplification (RPA) represents an even more rapid alternative, with reverse transcription RPA (RT-RPA) achieving amplification at temperatures between 37-42°C in as little as 20 minutes [1]. The combination of RT-RPA with lateral flow dipstick (LFD) readout yields a complete sample-to-result workflow that requires only a heat block or water bath and produces visual results on a paper strip, analogous to a pregnancy test [1]. This RT-RPA-LFD assay demonstrated approximately 100-fold greater sensitivity than a comparator RT-PCR method, a remarkable improvement attributable to the RPA enzyme’s high processivity and tolerance of inhibitors present in crude tissue lysates [1]. The assay’s simplicity makes it accessible to fish farmers and aquatic animal health workers with minimal training, enabling decentralized testing at the point of care [1]. The major limitation of RPA, its relatively short amplicon length, which constrains multiplexing and sequence confirmation, is offset by its unparalleled speed and operational simplicity.

CRISPR-Based Diagnostics: Precision and Programmability

The convergence of isothermal amplification with CRISPR-Cas systems has produced a new generation of diagnostics that combine the sensitivity of nucleic acid amplification with the sequence-specific recognition capabilities of CRISPR effectors. Two independent platforms have been developed for TiLV detection, both employing CRISPR-Cas12a or Cas12b nucleases that cleave single-stranded DNA reporters upon recognizing their target sequence [3, 6]. The first approach couples reverse transcription recombinase-aided amplification (RT-RAA) with CRISPR-Cas12a detection, achieving a limit of detection of 9.10 copies per reaction for plasmid standards and 91.82 fg/μL for TiLV RNA [3]. The assay targets segment 3 and showed 100% concordance with standard fluorescence-based methods when testing clinical samples, with no cross-reactivity to other fish pathogens [3]. The second platform integrates RT-LAMP directly with CRISPR-Cas12b in a one-pot reaction, reducing hands-on time and minimizing contamination risk [6]. This one-pot system operates at 62°C and provides real-time or endpoint fluorescence signals detectable with portable readers [6].

The mechanistic elegance of CRISPR-based diagnostics lies in their dual-specificity: the isothermal amplification step provides exponential signal generation, while the CRISPR nuclease adds a second layer of sequence verification that virtually eliminates false positives from non-specific amplification. The collateral cleavage activity of Cas12/13 enzymes, which indiscriminately cleaves nearby single-stranded DNA or RNA reporters after target recognition, amplifies the signal further and enables detection at attomolar concentrations [3]. These platforms are inherently programmable; redesigning the CRISPR RNA (crRNA) allows rapid adaptation to detect emerging TiLV variants or other pathogens, offering a flexible diagnostic platform for aquaculture health management.

Serological and Protein-Based Detection Methods

Complementary to nucleic acid-based tests, serological assays provide information about past exposure and immune status, which is essential for epidemiological studies and vaccine efficacy assessment. The development of an indirect enzyme-linked immunosorbent assay (ELISA) using recombinant TiLV segment 4 (nucleoprotein) antigen has enabled high-throughput screening of anti-TiLV antibodies in tilapia populations [33]. This assay, optimized through checkerboard titration, demonstrates intra- and inter-assay coefficients of variation below 10%, meeting accepted standards for diagnostic serology [33]. A key operational innovation is the validation of pooled serum testing, where combining five individual serum samples before analysis provides a cost-effective strategy for population-level surveillance without substantially compromising sensitivity, particularly when multiple seropositive individuals are present in the pool [33].

Antigen-capture ELISA systems have also been developed for direct detection of TiLV proteins in tissue homogenates and non-lethally collected mucus samples [47]. Using polyclonal antisera raised against purified TiLV, this assay achieved positive and negative predictive values of 76.19% and 65.62%, respectively, with an overall accuracy of 73.28% [47]. Notably, the highest detection rate (92.3%) was observed in mucus samples, supporting the feasibility of non-invasive screening for TiLV in asymptomatic carrier fish [47]. This capability is particularly valuable for broodstock screening programs aimed at preventing vertical transmission, which has been demonstrated for TiLV [23, 48].

The production of specific immunoglobulins for diagnostic and therapeutic applications has benefited from advances in recombinant protein expression. The TiLV nucleoprotein (segment 4) has been successfully expressed in Escherichia coli using codon-optimized constructs, yielding purified protein at concentrations of approximately 250 mg/L that retains antigenicity when tested with sera from infected fish [61]. Polyclonal antibodies generated against this recombinant protein in rabbits and chickens (IgY) show high specificity in Western blot and immunofluorescence assays, with no cross-reactivity to non-target proteins [19, 32, 63]. These antibodies serve as critical reagents for immunohistochemistry, flow cytometry, and neutralization assays, enabling detailed investigation of TiLV pathogenesis at the cellular level [34, 60].

RNA Extraction Innovations: Enabling Field Diagnostics

The pre-analytical step of RNA extraction represents a significant bottleneck for molecular diagnostics in field settings. Conventional methods using guanidinium thiocyanate-phenol-chloroform (Trizol) or commercial silica membrane kits require multiple centrifugation steps, hazardous organic solvents, and cold storage. A novel citrate buffer-based RNA extraction method addresses these limitations by eliminating the need for organic solvents and DNase treatment [62]. This approach uses mild acidic conditions to selectively precipitate RNA while DNA remains in solution, followed by simple centrifugation and washing steps. The resulting RNA is of sufficient quality and quantity for downstream RT-PCR and RT-qPCR, with performance comparable to the Trizol method [62]. The citrate method reduces reagent costs by approximately 70%, eliminates biohazard waste streams from organic solvents, and can be performed with basic laboratory equipment, making it ideally suited for decentralized testing in regional diagnostic laboratories [62].

Genomic Surveillance and High-Throughput Sequencing

Beyond detection and quantification, the characterization of TiLV genetic diversity is essential for understanding transmission pathways, identifying reassortment events, and tracking the emergence of variants with altered pathogenicity or antigenicity. A multiplexed RT-PCR assay capable of amplifying all ten complete genomic segments in a single reaction has been developed for nanopore-based sequencing [25]. This amplicon sequencing approach generates near-complete TiLV genomes from total RNA extracted from infected tissues, concentrated rearing water, or virus isolates cultured in E-11 cells [25]. The assay has enabled the recovery of 10 complete genomes from diverse sample types, contributing to a growing database of over 36 TiLV genomes that has revealed substantial genetic diversity across geographic regions [25]. Phylogenetic analyses using concatenated open reading frames from segments 1, 3, and 5 provide the most reliable tree topology for understanding TiLV evolution and phylogeography [16].

The genomic resolution afforded by these sequencing approaches has documented multiple reassortment events among TiLV isolates, indicating that co-infection with distinct viral strains occurs in aquaculture settings and may generate novel genotypes with unpredictable biological properties [16, 23]. The detection of genetically distant Vietnamese isolates suggests that TiLV diversity in Southeast Asia may be greater than previously appreciated, with implications for vaccine design and diagnostic target selection [23]. The integration of nanopore sequencing into routine surveillance programs offers the potential for real-time genomic epidemiology, enabling rapid identification of transmission sources and assessment of intervention effectiveness.

Summary of Diagnostic Performance and Operational Characteristics

The expanding toolkit for TiLV detection spans a continuum from high-throughput central laboratory methods to rapid point-of-care tests. RT-qPCR and ddPCR remain the gold standards for sensitivity and quantification, with ddPCR offering advantages for detecting low viral loads in carrier fish and environmental samples [15]. Semi-nested RT-PCR provides a cost-effective alternative for confirmatory testing in regional laboratories [37, 56]. Isothermal amplification methods, particularly RT-LAMP and RT-RPA with lateral flow readout, achieve clinically relevant sensitivity with minimal equipment requirements, making them suitable for on-farm use by trained personnel [1, 6]. CRISPR-based platforms add an additional layer of specificity and are poised for further development as integrated sample-to-result devices [3, 6]. Serological assays, while less sensitive for acute infection, are indispensable for seroprevalence studies and vaccine monitoring [33, 47].

The choice of diagnostic method must consider the specific application: outbreak confirmation demands high specificity to avoid false positives; surveillance requires high throughput and low cost; field deployment necessitates robustness and simplicity. The ideal diagnostic ecosystem for TiLV control will integrate multiple complementary technologies, leveraging central laboratory capacity for definitive diagnosis and genomic characterization while deploying rapid tests for front-line screening in production systems. Continued innovation in sample preparation, signal amplification, and device miniaturization will further lower barriers to widespread diagnostic access, supporting global efforts to mitigate the impact of this devastating pathogen.

Vaccine Development Strategies: Subunit, Nanoparticle, and Passive Immunization Approaches

The development of effective prophylactic interventions against Tilapia Lake Virus (TiLV) remains a paramount objective for the global aquaculture industry, given its status as a highly contagious pathogen causing mortality rates exceeding 90% in severe outbreaks and its classification as a notifiable disease by the World Organisation for Animal Health (WOAH) [18, 29, 57]. Despite the urgent need, no commercial vaccine is currently available, a gap that underscores the multifaceted challenges posed by this emerging orthomyxo-like virus, including its segmented RNA genome, limited understanding of protein function, and the absence of a fully optimized in vitro culture system for large-scale antigen production [29, 42, 52]. However, the demonstrated ability of tilapia to develop protective immunity following natural exposure provides a strong rationale for vaccination, and recent years have witnessed a surge in innovative vaccine strategies targeting this pathogen [52, 64]. Three primary categories of intervention, subunit vaccines, nanoparticle-based delivery systems, and passive immunization, have been explored with varying degrees of success, each leveraging distinct immunological principles and offering unique advantages for the specific constraints of tilapia aquaculture.

Subunit and Recombinant Protein Vaccine Approaches

The subunit vaccine strategy for TiLV relies on the identification and production of specific viral proteins capable of eliciting a protective immune response, circumventing the need for live virus handling and enabling precise targeting of immunogenic epitopes. Given the segmented nature of the TiLV genome, encoding at least ten major proteins and an eleventh alternative reading frame product (S9-F3), a critical first step has been the selection of appropriate antigenic targets [13, 52]. Segment 4 (S4), encoding the nucleoprotein (NP), has been a primary focus. The NP is a highly conserved structural protein essential for viral replication, as it encapsulates the viral RNA into ribonucleoprotein complexes, and the recent cryo-EM structure of TiLV-NP bound to RNA has provided exquisite detail on its head-body domain architecture and RNA-binding groove, validating its role as a core viral component [4, 10, 19]. Recombinant S4 protein has been successfully expressed in Escherichia coli systems, with optimized protocols using codon optimization and specific induction conditions (0.1 mM galactose at 37°C) achieving high yields exceeding 250 mg/L of purified protein, demonstrating the scalability potential for industrial production [61]. The immunogenicity of this recombinant S4 has been confirmed through dot blot assays showing consistent interaction with sera from TiLV-infected fish and rabbit anti-TiLV antibodies, establishing it as a viable antigen for further development [61].

Beyond S4, other genomic segments have been investigated as subunit vaccine candidates. The surface glycoproteins encoded by segments 5 (S5) and 6 (S6) are predicted to be involved in host-cell attachment and entry, making them logical targets for neutralizing antibody induction [52]. A seminal study evaluating co-immunization with recombinant S5 and S6 proteins demonstrated a synergistic effect, where combined administration resulted in significantly higher production of S6-specific antibodies compared to S6 alone. This co-immunization strategy yielded a relative percent survival (RPS) of 57.14% in a viral challenge model, while individual immunization with either protein provided negligible protection [8]. This finding highlights the potential importance of multivalent subunit formulations to compensate for the low immunogenicity often associated with single recombinant proteins. Similarly, segments 9 (S9) and 10 (S10) have been explored as DNA and recombinant protein vaccines. Immunization with combined Tis9 and Tis10 recombinant proteins in hybrid red tilapia conferred up to 55.56% RPS, with in silico analysis revealing multiple B-cell epitopes in their predicted coil structures, which may account for their immunogenicity [21]. The S9-F3 protein, an alternative translation product of segment 9 that undergoes CRM1-dependent nuclear export, adds a further layer of antigenic complexity that could be exploited [13].

An elegant approach to epitope discovery has been the combination of phage display technology with in silico bioinformatics. By panning a Ph.D.-12 phage library against sera from TiLV-surviving tilapia, researchers identified a mimotope (Pep3) that provided 57.6% protection. Computational alignment and structural analysis of the target protein mapped this mimotope to a natural antigenic site on segment 1 (S1), specifically the S1^399-410 epitope. Vaccination with a KLH-conjugated version of this epitope (KLH-S1^399-410) elicited a robust and durable antibody response, achieving a remarkable 81.8% survival rate in challenge studies, with antibody depletion experiments confirming that anti-S1^399-410 antibodies were essential for neutralization [12]. This study provides a powerful proof-of-concept for epitope-based vaccines. Immunoinformatic approaches have further advanced the field by enabling the rational design of multi-epitope subunit vaccines. Using the putative polymerase basic 1 (PB1) gene, researchers have identified immunodominant cytotoxic T lymphocyte (CTL), helper T lymphocyte (HTL), and linear B lymphocyte (LBL) epitopes, constructing a vaccine predicted to be antigenic, non-allergenic, and capable of inducing real-life immune responses in computer simulations [22]. The baculovirus expression vector system (BEVS) has also been employed to produce recombinant S4 protein in insect cells and larvae without the need for antibiotic resistance markers, generating polyclonal anti-S4 antibodies in rabbits, further validating the utility of this platform for diagnostic and vaccine antigen production [63].

Nanoparticle-Based Vaccine Delivery Systems

While subunit vaccines demonstrate antigenicity, their efficacy is often constrained by poor immunogenicity and rapid degradation in the biological milieu. Nanoparticle-based delivery systems address these limitations by protecting the antigen, enhancing uptake by antigen-presenting cells, and providing a depot effect for sustained immune stimulation. The unique physiological characteristics of fish, particularly the mucosal surfaces of the gills, skin, and intestine, make immersion-based nanoparticle vaccines an especially attractive strategy for mass vaccination of juvenile tilapia, avoiding the stress and labor costs associated with intraperitoneal injection [2, 20, 52].

Chitosan, a biocompatible and mucoadhesive polysaccharide derived from chitin, has emerged as a leading nanoparticle platform for TiLV vaccines. Its positive zeta potential facilitates electrostatic interaction with negatively charged mucosal surfaces, enhancing antigen retention and uptake. An early formulation involved encapsulating whole, inactivated TiLV (killed virus, KV) into chitosan nanoparticles (CN-KV). This combination yielded a nanovaccine with a spherical morphology and a diameter of approximately 120.4 nm, which was significantly smaller than the free inactivated virus [20]. Confocal microscopy and immunohistochemistry confirmed that the CN-KV nanovaccine was mucoadhesive to fish gills and intestines, with antigen deposition occurring within 30 minutes of immersion [2, 20]. Under a cohabitation challenge model, the CN-KV nanovaccine achieved an RPS of 68.17%, a dramatic improvement over the 25.01% RPS observed for the free KV alone. This protection correlated with significantly higher TiLV-specific antibody levels. Crucially, this effectiveness translated to field conditions, where vaccinated fish exhibited an RPS of 52.2% compared to non-vaccinated controls, demonstrating real-world applicability [20].

Building on this success, the approach was refined by encapsulating recombinant TiLV segment 4 protein into chitosan nanoparticles, termed nanoTiLV-S4 (CNS4). This formulation addressed the challenge of producing large quantities of inactivated virus, instead relying on a defined, recombinant antigen. The CNS4 particles had a size of 284 ± 9.2 nm with a positive zeta potential of 17.7 ± 0.7 mV. The nano-encapsulation process reduced the particle size from over 2.2 μm to the nanoscale, which is critical for efficient cellular uptake [2]. Similar to the CN-KV vaccine, the CNS4 antigen was rapidly deposited on gills and intestines. However, while the CNS4 vaccine demonstrated lower efficacy compared to the whole-virus nanovaccine in laboratory (RPS 25%) and field (RPS 31.88%) settings, it still provided significant protection over unvaccinated controls and represents a safer, more defined product that can be standardized for manufacturing [2]. The higher efficacy of the whole-virus nanovaccine likely reflects the presentation of multiple conserved epitopes, whereas the single-protein subunit nano-vaccine may benefit from the inclusion of additional antigens or adjuvants.

A distinct biomimetic approach involves the use of erythrocyte membrane-coated nanoparticles. This strategy leverages the inherent biocompatibility and long-circulating properties of red blood cell membranes. A system termed Cs-S2@M-M, based on chitosan nanoparticles loaded with TiLV S2 protein and cloaked in tilapia erythrocyte membrane, achieved a particle size of ~100 nm and an encapsulation efficiency of approximately 79.15% [17]. This biomimetic nanovaccine significantly upregulated immune gene expression and enhanced the activity of non-specific immune-related enzymes. Following TiLV challenge, it elevated the RPS by 17.4% to 26.1% compared to controls, demonstrating that cell membrane cloaking can augment the immune response to subunit antigens [17]. The success of these nanoparticle platforms is partly attributable to their ability to target and be internalized by antigen-presenting cells. The granulocyte tropism of TiLV, where these cells harbor the highest infection rates (46%) and may act as viral reservoirs for systemic dissemination, presents both a challenge and an opportunity; nanoparticle vaccines could be designed to specifically target these cells to block viral propagation [34].

Passive Immunization Strategies

Complementing active vaccination, passive immunization offers a rapid, non-replicative means of conferring immediate protection, which is particularly valuable for controlling acute outbreaks and protecting high-value broodstock or larvae during critical early life stages. This strategy relies on the administration of pre-formed antibodies, bypassing the lag period required for the host to mount its own adaptive immune response [32, 48, 52]. For TiLV, two primary sources of passive antibodies have been investigated: immunoglobulin Y (IgY) derived from egg yolks of immunized hens, and maternal antibodies transferred from vaccinated broodstock to offspring.

IgY technology presents a cost-effective and scalable approach for mass antibody production. Laying hens immunized with recombinant TiLV segment 4 protein produced specific IgY antibodies, which were extracted and purified from egg yolks using polyethylene glycol precipitation. Western blot analysis confirmed the specificity of the anti-TiLV IgY, showing no cross-reactivity with non-target proteins [32]. In vitro neutralization assays demonstrated a clear dose-dependent reduction in TiLV infectivity, with viral titers declining from 5.01 × 10⁶ TCID50/mL to between 5.01 × 10⁴ and 1.26 × 10⁵ TCID50/mL. The highest neutralizing efficacy was observed at a 1:2 dilution of IgY. Immunofluorescence assays further confirmed a significant reduction in TiLV antigen levels in treated RHTiB cells, and the IgY effectively inhibited virus-induced cytopathic effects [32]. This specificity and neutralizing capacity position IgY as a promising prophylactic, potentially suitable for administration via feed or immersion to provide broad, short-term protection across a population, especially in field settings where vaccination programs are challenging to implement.

A second vital passive immunization route involves the maternal transfer of immunity. This approach is particularly logical for TiLV, given evidence of vertical transmission [18, 48]. Broodstock tilapia were immunized with either heat-killed TiLV vaccine (HKV) or formalin-killed TiLV vaccine (FKV). Both vaccines elicited specific anti-TiLV IgM in the majority of immunized male and female broodstock. Critically, these maternal antibodies were transferred to fertilized eggs and detected in larvae. The HKV group showed higher levels of maternal antibody transfer to eggs compared to the FKV group. However, these passive antibodies were short-lived, detectable in 1-3-day-old larvae but absent by 7-14 days, indicating a finite window of protection [48]. To confirm the functionality of these antibodies, passive immunization experiments using sera from immunized female broodstock were performed. Administration of this immune serum to naïve juvenile tilapia conferred remarkably high protection, ranging from 85% to 90% against TiLV challenge [48]. This finding strongly supports the concept that maternal vaccination can protect offspring during their most vulnerable early stages, bridging the gap until they are old enough for active vaccination with immersion-based nanovaccines. The synergistic potential of combining passive maternal immunization with active nanoparticle vaccination for fry is a promising avenue for integrated TiLV control. Furthermore, the development and validation of high-throughput serological tools, such as an indirect ELISA using recombinant S4 protein for pooled serum analysis, are essential for monitoring the effectiveness of these vaccination strategies at the farm level, enabling cost-effective surveillance of population immunity [31, 33, 47, 50].

Control and Prevention Measures for TiLV in Aquaculture Systems

The control and prevention of Tilapia Lake Virus (TiLV) disease represents one of the most formidable challenges facing global tilapia aquaculture, necessitating a multi-faceted, integrated strategy that combines rigorous biosecurity, advanced diagnostic surveillance, strategic vaccination, genetic selection for resistance, and environmental management. As TiLV has been declared a notifiable disease by the World Organisation for Animal Health (WOAH) [24], the imperative for implementing comprehensive control measures has never been more urgent, particularly given the virus's capacity to cause mortality rates exceeding 90% in affected populations [18, 28, 29, 52] and its documented presence across 17 countries on multiple continents [29, 42, 65]. The following analysis provides an exhaustive examination of the current and emerging strategies for TiLV control, grounded in the latest research findings and field observations.

Biosecurity and Surveillance as Foundational Control Pillars

The cornerstone of any effective TiLV control program is the implementation of stringent biosecurity protocols combined with robust surveillance systems capable of detecting viral presence before clinical outbreaks occur. The virus's ability to persist in both wild and feral fish populations, as demonstrated by persistent detection of TiLV in wild tilapia and tinfoil barbs in Malaysian lakes for up to two years following initial outbreaks, without obvious clinical signs [58], underscores the critical importance of preventing viral introduction into naive populations. Biosecurity measures must encompass multiple layers of protection, including strict quarantine protocols for introduced fish, disinfection of equipment and water sources, control of movement between farms, and elimination of potential vectors. The identification of TiLV in giant gourami (Osphronemus goramy) in East Java with genetic identity reaching 96.8% similarity to the Israeli isolate [24], coupled with documented susceptibility of river barb and other species [18], expands the range of potential reservoir hosts that must be considered in biosecurity planning.

Surveillance strategies must be adaptive and sensitive enough to detect subclinical infections, which are now recognized as a major mechanism for viral spread. The development and validation of pooled serum ELISA testing for monitoring farm-level immunity offers a cost-effective approach for large-scale surveillance, with pooling of five serum samples proving effective for detecting TiLV-specific antibodies, though sensitivity may be reduced when only one seropositive sample is present in the pool due to dilution effects [33]. This serological approach complements molecular diagnostic methods, providing a comprehensive picture of both current infections and historical exposure. The use of non-invasive sampling methods, particularly mucus, has emerged as a practical alternative for surveillance programs, with indirect ELISA demonstrating positive rates of 92.3% (36/39) in mucus samples compared to other tissues [47], while droplet digital PCR (ddPCR) has shown remarkable sensitivity in detecting TiLV in mucus, water, and infected tissue samples with a detection limit as low as 3.3 copies of TiLV [15]. The development of a semi-quantitative lesion scoring system for differentiating clinical from subclinical infection provides an additional tool for field-based surveillance, allowing for prognostic assessment of disease severity [46].

Advanced Diagnostic Tools for Early Detection and Outbreak Management

The rapid evolution of diagnostic technologies has revolutionized the capacity for early TiLV detection, enabling proactive intervention before outbreaks escalate. The advent of isothermal amplification methods has been particularly transformative for field applications, eliminating the dependency on sophisticated laboratory infrastructure. The reverse transcriptase recombinase polymerase amplification combined with lateral flow dipstick (RT-RPA-LFD) assay achieves detection within approximately 20 minutes at constant temperature without complex instrumentation, demonstrating sensitivity approximately 100 times greater than comparable RT-PCR methods [1]. Similarly, the one-pot RT-LAMP CRISPR/Cas12b platform provides results within 75 minutes at 62°C with a 95% limit of detection of 79.6 copies, and has been partially validated against 261 samples from 9 source populations, achieving 92% diagnostic sensitivity and 100% diagnostic specificity [6]. The CRISPR/Cas12a system combined with reverse transcription recombinase aided amplification (RT-RAA) offers another powerful option, with a detection limit of 9.10 copies per reaction for recombinant plasmid standards and 91.82 fg/μL for TiLV RNA, showing 100% concordance with standard fluorescence methods [3].

For quantitative applications, the TaqMan RT-qPCR assay targeting segment 10 has been rigorously validated, demonstrating a limit of detection of 10 RNA viral copies per reaction with 96.7% diagnostic sensitivity and 100% diagnostic specificity across 91 proven-positive and 185 proven-negative samples, while successfully detecting TiLV strains from North America, South America, Africa, and Asia [30]. The droplet digital PCR (ddPCR) assay offers absolute quantification with 10-fold higher sensitivity than RT-qPCR, capable of detecting TiLV in water samples at concentrations as low as 7.9 ± 0.99 copies per reaction, making it particularly valuable for environmental surveillance [15]. The development of multiplex PCR targeting three conserved genome segments (2, 3, and 8) addresses the critical issue of genetic variability and potential reassortment, with receiver operating characteristic analysis yielding an area under the curve value of 1.0, indicating perfect sensitivity and specificity [14]. For comprehensive genomic surveillance, the multiplexed RT-PCR assay coupled with Nanopore sequencing enables whole genome recovery from infected fish tissues, concentrated rearing water, and cell culture, providing essential data for tracking viral evolution and reassortment events [25].

The practical implementation of these diagnostic tools in field settings has been facilitated by the development of cost-effective RNA extraction methods, such as the citrate buffer-based approach that eliminates the need for harsh organic solvents and DNase treatment while yielding high-quality RNA suitable for downstream applications [62]. The availability of species-specific cell lines, including the RHTiB cell line derived from red hybrid tilapia brain supporting TiLV propagation with maximum concentrations of 10^7.82 ± 0.22 viral copies/400 ng cDNA after 9 days of incubation [60], and the characterization of ten fish cell lines with varying susceptibilities, with the highest viral titers generated on TiB (tilapia brain), MSF (largemouth bass), CAMK (hybrid snakehead), and SS (perch) cells [69], provides essential tools for virus isolation and characterization. The establishment of the RBT-H cell line from rainbow trout heart, which is refractory to TiLV infection [66], offers a negative control system for specificity testing in diagnostic applications.

Vaccination Strategies: Current Advances and Remaining Challenges

The development of effective vaccines against TiLV has progressed rapidly, driven by the recognition that tilapia exposed to TiLV can develop protective immunity, suggesting that vaccination is achievable [52]. Several vaccine platforms have been evaluated, with varying degrees of success, and the field is moving toward combination approaches that target multiple viral proteins to enhance protective efficacy.

Inactivated and Subunit Vaccines

The chitosan nanoparticle-based immersion vaccine has emerged as a particularly promising platform, addressing the practical challenges of mass vaccination in aquaculture settings. The nanovaccine targeting segment 4 (S4) demonstrated a particle size of 284 ± 9.2 nm with positive zeta potential of 17.7 ± 0.7 mV, enabling mucoadhesive properties that facilitate antigen uptake through gills and intestines within 30 minutes of immersion [2]. Under laboratory cohabitation challenge conditions, this vaccine achieved a relative percent survival (RPS) of 25%, while field trials yielded an RPS of 31.88% [2]. Earlier work with the chitosan nanoparticle inactivated vaccine (CN-KV) showed more substantial protection, with RPS of 68.17% under laboratory conditions and 52.2% in field trials, accompanied by significantly higher TiLV-specific antibody responses compared to both the inactivated virus alone and control groups [20]. The superiority of the nanoparticle formulation over the traditional inactivated vaccine (KV, RPS 25.01%) highlights the importance of delivery systems in enhancing immunogenicity.

The development of a biomimetic nanovaccine based on erythrocyte membrane encapsulation (Cs-S2@M-M) represents an innovative approach to improving subunit vaccine efficacy. This system, with a particle size of approximately 100 nm and encapsulation efficiency of 79.15%, significantly upregulated immune gene expression, enhanced non-specific immune-related enzyme activity, and improved RPS by 17.4-26.1% following intramuscular injection [17]. The recognition that co-immunization can produce synergistic effects has been demonstrated with recombinant S5^196–272 and S6^200–317 proteins, where combined administration resulted in higher production of S6-specific antibodies than individual immunization, with the highest virus-neutralizing effect observed at 87.22% viability and an RPS of 57.14% compared to no protection from individual immunization [8].

DNA and Recombinant Protein Vaccines

DNA vaccine approaches targeting segments 9 and 10 have shown particular promise, especially when used in combination. The pcDNA-Tis9 + pcDNA-Tis10 combination achieved the highest RPS of 61.11 ± 9.62%, while the recombinant protein combination Tis9 + Tis10 yielded 55.56 ± 9.62% protection [21]. The superiority of segment 10-based vaccines over segment 9-based vaccines (Tis10 RPS higher than Tis9) suggests that careful antigen selection is critical for optimizing immune responses. The use of extended host-range baculovirus expression vector systems for producing recombinant S4 protein in insect cells and larvae offers a scalable manufacturing platform, with the expressed protein successfully inducing rabbit polyclonal antibodies capable of recognizing native S4 protein [63].

Epitope-Based Vaccines and Immunoinformatic Approaches

The identification of protective epitopes through combined experimental and computational approaches has opened new avenues for rational vaccine design. Using phage display technology with serum from TiLV survivors, researchers identified the mimotope Pep3 (TYTTRMHITLPI), which provided protection with an RPS of 57.6% following prime-boost vaccination [12]. Further bioinformatic analysis identified the natural epitope S1^399–410 (TYTTRNEDFLPT410) located on segment 1, and the corresponding KLH-conjugated epitope vaccine induced a robust antibody response with survival rates reaching 81.8% [12]. Antibody depletion studies confirmed that anti-S1^399–410-specific antibodies were essential for neutralizing TiLV, validating this epitope as a critical target for vaccine development. Immunoinformatic approaches have also been employed to design multi-epitope subunit vaccines based on the putative polymerase basic 1 (PB1) gene, identifying cytotoxic T lymphocyte (CTL), helper T lymphocyte (HTL), and linear B lymphocyte (LBL) epitopes that were antigenic, non-allergenic, and demonstrated structural compactness and binding stability through molecular dynamics simulation [22].

Broodstock Vaccination and Maternal Immunity

The demonstration of vertical transmission of TiLV [18, 48] has prompted investigation into broodstock vaccination as a strategy for protecting offspring. Immunization of Nile tilapia broodstock with heat-killed TiLV vaccine (HKV) or formalin-killed TiLV vaccine (FKV) resulted in production of anti-TiLV IgM in both males and females, with maternal antibodies detected in fertilized eggs and larvae from vaccinated broodstock [48]. Notably, HKV induced higher levels of maternal antibody in fertilized eggs compared to FKV, and passive immunization experiments revealed that antibodies elicited by TiLV vaccination conferred 85% to 90% protection against TiLV challenge in naive juvenile tilapia [48]. However, the persistence of maternal antibodies was limited, detectable in 1-3 day old larvae but undetectable by 7-14 days post-hatch, indicating that broodstock vaccination may require timed administration to align with peak susceptibility periods.

Passive Immunization Approaches

Given the challenges of vaccine development and the need for immediate therapeutic options, passive immunization using immunoglobulin Y (IgY) antibodies raised against recombinant TiLV segment 4 protein in laying hens has emerged as an alternative strategy. In vitro neutralization assays demonstrated dose-dependent reduction in TiLV infectivity, declining from 5.01 × 10^6 TCID50/mL to 5.01 × 10^4–1.26 × 10^5 TCID50/mL, with the highest efficacy at a 1:2 dilution [32]. Immunofluorescence assays confirmed significant reduction in TiLV antigen levels in IgY-treated RHTiB cells, suggesting that IgY-based passive immunization could provide a rapid intervention tool during outbreaks, particularly in high-value broodstock or vulnerable juvenile stages.

Genetic Selection for TiLV Resistance

The identification of genetic markers associated with TiLV resistance offers a sustainable, long-term approach to disease management that complements biosecurity and vaccination. Genome-wide association studies have revealed a major quantitative trait locus (QTL) on chromosome Oni22 in Nile tilapia, where the average mortality rate of fish homozygous for the resistance allele at the most significant SNP was 11% compared to 43% for those homozygous for the susceptibility allele [26]. Candidate genes within this QTL region include lgals17, vps52, and trim29, which are involved in host response to viral infection [26]. Fine mapping and functional annotation have narrowed this QTL to the proximal end of Oni22, identifying 74 out of the top 99 markers associated with binary survival within a 10 Mb window, with the marker explaining the highest genetic variance located at 1.7 Mb with a substitution effect of 0.15 [7]. Additional SNPs on other chromosomes, particularly two separate regions of Oni09, suggest an oligogenic architecture underlying TiLV resistance, with several QTLs of moderate effect and many with small effect [7]. Transcriptomic analyses identified proteosome subunit beta type-9a and ha1f as potential causal genes within the Oni22 QTL, providing targets for marker-assisted selection [7].

The practical application of genetic selection has been demonstrated through comparative evaluation of tilapia strains. Among six Indonesian tilapia strains evaluated for TiLV resistance, the Gesit strain showed highest resistance, followed by Best, Red/Albino, Nirwana, Non-specific, and Srikandi, with mortality ranges of 1.67%–28.33% for cohabitation and 16.67%–61.67% for intraperitoneal injection [39]. Research at the University of Göttingen identified a strain originating from Lake Turkana (ELM) that did not develop clinical signs following cohabitation challenge, achieving nearly 100% survival compared to 29.3% in the DRE strain and 6.7% in the MAN strain [49]. This resistance correlated with lower viral loads in both mucosal and internal tissues, potentially mediated by higher magnitude of mx1-based antiviral response and lower pro-inflammatory responses [49]. However, the observation that resistant strains may retain significant virus loads in liver and brain and thus could become persistent carriers [49] emphasizes that genetic selection must be integrated with other control measures rather than relied upon exclusively.

Immunomodulation Through Feed Additives and Probiotics

The modulation of host immune responses through nutritional interventions represents a practical, scalable strategy for enhancing TiLV resistance in commercial aquaculture operations. Dietary supplementation with β-glucan derived from the marine diatom Chaetoceros muelleri at 0.1% of feed for 14 days prior to viral challenge resulted in significantly lower cumulative mortality (26.67%) compared to controls (55.56%) [43]. This protection was associated with coordinated upregulation of multiple immune-related genes, including cytolytic (NCCRP-1), stress-related (Hsp70), antimicrobial (C-lysozyme), cytokine (IL-8), pattern recognition receptors (TLR3, TLR5, TLR9), signaling adaptor (myD88), antiviral effectors (Mx, RSAD-2), and adaptive immunity markers (IgM, CD4) [43]. Gene co-expression network analyses identified Mx, RSAD-2, and CD4 as central hub genes associated with antiviral defense pathways, providing mechanistic insights into β-glucan-mediated protection [43].

Probiotic interventions have demonstrated significant potential for reducing disease transmission and enhancing immune responses in TiLV-infected tilapia populations. Mechanistic modeling of probiotic effects revealed that the most marked benefits are associated with immune system enhancements of approximately 30%, reductions in disease transmission of approximately 80%, and reductions in the basic reproduction number (R0) of approximately 70% [59]. These effects translate to a substantially higher tolerance of farming densities, approximately 400-fold increase, in aquaculture systems [59]. Specific feed additive strategies have been evaluated in coinfection models with Aeromonas hydrophila, demonstrating that targeted dietary supplementation can improve survival and immune competence against simultaneous viral and bacterial challenges [38].

The discovery that the coxsackievirus and adenovirus receptor (CAR) in tilapia (OnCAR) inhibits TiLV infection by binding to viral segment 8 and 10 encoded proteins [9] opens the possibility of developing compounds that enhance this natural antiviral mechanism. OnCAR overexpression in TiB cells significantly decreased viral gene transcripts, genome copy number, cytopathic effect, and viral internalization, identifying a cellular receptor that could be targeted for therapeutic intervention [9]. Similarly, the characterization of interferon regulatory factor 6 (IRF6) from Nile tilapia as a positive regulator of type I interferon-a3 promoter activity, with silencing experiments demonstrating increased mortality and viral load in TiLV-infected tilapia [35], suggests that immunostimulatory strategies targeting the IRF/IFN axis could enhance host resistance.

Environmental Management and Culture System Design

The dynamics of TiLV transmission within aquaculture systems are profoundly influenced by environmental conditions and system design, providing opportunities for intervention through environmental management. A comprehensive field study comparing recirculating aquaculture systems (RAS) with non-RAS operations revealed that continuous water recirculation allows the virus to accumulate in the system, leading to more prolonged outbreaks, while non-RAS conditions with regular water discharge showed lower viral loads and faster recovery [53]. In RAS, TiLV concentrations in pond water were significantly higher in outbreak conditions and positively correlated with increased fish mortality, whereas non-RAS operations demonstrated limited viral accumulation and shorter disease outbreak duration [53]. Importantly, vaccinated fish in RAS showed substantially lower cumulative mortality (16.7%) compared to unvaccinated controls (37.7%), with an RPS of 55.6%, and TiLV concentrations in pond water of the vaccinated group were significantly lower, indicating reduced environmental viral shedding [53].

Temperature manipulation represents another environmental management tool, as TiLV has been shown to cause mass mortality peaking at 30°C, reaching 50% death rates, with infection occurring in waters above 25°C [36]. Daily temperature variation results in significantly higher virus-induced mortality patterns among tilapia strains [39], suggesting that maintaining stable, optimal temperatures below 30°C could reduce disease severity. Water quality parameters also influence TiLV dynamics, with moderate negative correlation between water pH and TiLV presence (R = -0.4472; p < 0.05) [58], indicating that pH management may have ancillary benefits for disease control.

The use of environmental sampling as a non-invasive monitoring tool has been validated through ddPCR detection of TiLV in water samples with copy numbers as low as 7.9 ± 0.99 copies/reaction [15], enabling early warning systems that can detect viral presence before clinical signs emerge. This approach is particularly valuable in RAS where water recirculation facilitates viral accumulation and detection. The integration of molecular monitoring of rearing water with vaccination and good water management practices has been proposed as a practical, non-invasive strategy for detecting and controlling TiLV outbreaks in intensive farming systems [53].

Quarantine and Movement Control Measures

Given the documented role of fish movement in the international and regional spread of TiLV, strict quarantine and movement control measures are essential components of prevention strategies. The detection of TiLV in tilapia being transported through quarantine facilities in Surabaya, Indonesia, where 1 of 35 samples tested positive using conventional nested PCR [68], demonstrates the importance of screening at border checkpoints. Similarly, surveillance in the Indian Sundarbans revealed that TiLV was detected in 37.3% of surveyed ponds, with a two-step semi-nested PCR showing higher sensitivity than single-step SYBR green-based qPCR, emphasizing the need for sensitive diagnostic methods in quarantine applications [37]. The development and implementation of strict quarantine measures are necessary to combat the outbreak and spread of TiLV in ecologically sensitive areas [37].

The recognition that TiLV can be shed into water and persists in the environment [53, 58] necessitates comprehensive disinfection protocols for equipment, transport vehicles, and facilities. The virus's ability to infect multiple fish species beyond tilapia, including giant gourami [24] and tinfoil barbs [58], expands the scope of quarantine requirements to include all potentially susceptible species. Country-level risk assessments, such as the declaration by FAO that Bangladesh was in a high-risk zone for TiLV spread [51], should inform national surveillance and import control policies.

Antiviral Drug Development

The identification of potential antiviral compounds through computational drug design approaches has opened new avenues for therapeutic intervention. Targeting the CRM1 protein, which plays a critical function in the development and spreading of animal viruses, researchers have identified promising phytochemicals from tropical mangrove plants Heritiera fomes and Ceriops candolleana [67]. Molecular docking, ADME analysis, toxicity assessment, and molecular dynamics simulation identified top candidate compounds, CID107876 (-8.3), CID12795736 (-8.2), and CID12303662 (-7.9), with favorable binding affinities and stability profiles [67]. While these compounds require further in vivo validation, they represent a potential avenue for antiviral drug development. The structural resolution of the TiLV nucleoprotein bound to RNA through cryo-electron microscopy at up to 2.9 Å resolution [4, 10] provides atomic-level detail of the viral replication machinery, enabling structure-based drug design targeting the nucleoprotein's RNA-binding groove. The characterization of TiLV entry pathways, revealing dependence on dynamin activity and cholesterol but independence from endosomal acidification [27], identifies specific cellular targets for entry inhibitors that could block viral infection at an early stage.

Future Directions and Research Gaps in TiLV Biology and Management

The trajectory of Tilapia Lake Virus (TiLV) research, while having advanced significantly since its initial identification in 2014, reveals profound and persistent gaps that must be addressed to safeguard global tilapia aquaculture. Despite the declaration of TiLV disease (TiLVD) as a notifiable pathogen by the World Organisation for Animal Health (WOAH) and its recognition as a transboundary disease by the Food and Agriculture Organization (FAO), the scientific community remains in a reactive rather than a predictive posture. The following analysis delineates the critical frontiers for future investigation, structured around fundamental virology, host-pathogen dynamics, diagnostic evolution, vaccinology, genetic resistance, and ecological management.

### Unraveling the Molecular Virology and Pathogenesis of TiLV

A foundational impediment to rational therapeutic and vaccine design is the incomplete understanding of TiLV’s molecular biology. While the cryo-electron microscopy (cryo-EM) structure of the nucleoprotein (NP, encoded by segment 4) has been resolved, revealing a conserved articulaviral fold with a positively charged RNA-binding groove and an oligomerization loop [4, 10], the functions of the majority of the ten (and potentially eleven) encoded proteins remain enigmatic. The recent identification of an 11th protein, S9-F3, translated from an alternative reading frame of segment 9 and subject to CRM1-dependent nuclear export, underscores the complexity of the viral proteome and suggests that additional, undiscovered polypeptides or regulatory elements may exist [13]. Future work must systematically characterize the function of each viral protein, particularly those encoded by segments 5 through 10, which lack homology to known sequences [19]. This includes elucidating their roles in immune evasion, host shutoff, and modulation of cellular signaling pathways.

The entry pathway of TiLV has been partially mapped, demonstrating a dependence on dynamin activity, cholesterol, and the actin cytoskeleton, while being independent of clathrin and endosomal acidification [27]. This is a striking divergence from the prototypical orthomyxovirus, influenza A virus, and suggests a unique fusion mechanism. Future research must identify the specific cellular receptor(s) for TiLV. The coxsackievirus and adenovirus receptor (CAR) has been identified as a potential anti-TiLV factor that binds to viral segments 8 and 10, but its role as a primary entry receptor is not established [9]. Identifying the bona fide receptor is a prerequisite for understanding tissue tropism and developing entry-blocking therapeutics. Furthermore, the mechanism of viral egress and the role of the recently characterized nuclear export signal in S9-F3 in the viral life cycle require detailed investigation [13]. The observation that TiLV causes profound mitochondrial damage, leading to loss of membrane potential and ATP depletion, provides a mechanistic basis for the extensive cell death observed in infected tissues [45]. Future studies should explore whether this mitochondrial dysfunction is a direct consequence of viral protein activity or a secondary effect of the host immune response, and whether targeting mitochondrial integrity could offer a therapeutic avenue.

### Deciphering Host-Pathogen Interactions and Immunopathogenesis

The host immune response to TiLV is a double-edged sword, characterized by both protective antiviral mechanisms and immunopathogenic processes that contribute to mortality. The discovery that granulocytes are the primary target of TiLV infection, with 46% of these cells harboring the virus while exhibiting minimal apoptosis, suggests a novel strategy for systemic dissemination, potentially using these innate immune cells as “Trojan horses” [34]. This finding challenges the conventional view of granulocytes as purely antiviral effectors and opens a new research avenue into the role of these cells as viral reservoirs. Concurrently, the profound depletion of lymphocytes, driven by virus-induced apoptosis, explains the severe leukopenia and immunosuppression observed in infected fish [34, 40]. The interplay between these two phenomena, granulocyte tropism and lymphocyte depletion, is a critical area for future investigation.

The innate immune response, particularly the interferon (IFN) system, is central to host defense. The interferon regulatory factor 6 (IRF6) has been identified as a positive regulator of type I IFN in tilapia, and its silencing increases susceptibility to TiLV [35]. However, the virus appears to have evolved mechanisms to subvert this response. Transcriptomic analyses reveal that TiLV infection downregulates key antiviral genes such as IFIT1 and TRIM25 while upregulating suppressors of cytokine signaling (SOCS), indicating a sophisticated strategy of immune evasion [50]. Future research must map the specific viral proteins responsible for this subversion and identify the host factors that are critical for restriction. The role of the Mx protein, a classic IFN-stimulated gene (ISG), appears to be a correlate of resistance in certain tilapia strains, with higher baseline mx expression linked to survival [49]. However, the direct antiviral activity of tilapia Mx proteins against TiLV has not been rigorously demonstrated. The development of a zebrafish model for TiLV infection, as suggested by recent work [11], could accelerate the functional characterization of host genes through forward and reverse genetic screens.

### Advancing Diagnostic Capabilities for Surveillance and Control

While significant progress has been made in developing molecular diagnostics, critical gaps remain in their deployment and validation for field use. The current gold standard, RT-qPCR, is sensitive and specific but requires sophisticated laboratory infrastructure [30, 31]. The emergence of isothermal amplification technologies, such as reverse transcription recombinase polymerase amplification (RT-RPA) combined with lateral flow dipsticks (LFD) and CRISPR-based assays (RT-RAA-CRISPR/Cas12a and RT-LAMP-CRISPR/Cas12b), represents a paradigm shift towards point-of-care (POC) diagnostics [1, 3, 6]. These assays can deliver results in under 75 minutes with minimal equipment, making them suitable for on-farm use in resource-limited settings. However, the diagnostic sensitivity and specificity of these novel assays must be rigorously validated against large, geographically diverse panels of clinical samples, not just laboratory isolates. The RT-LAMP-CRISPR/Cas12b assay, for example, showed 92% diagnostic sensitivity and 100% specificity on a panel of 261 samples, which is promising but requires further validation across different production systems and seasons [6].

A major research gap is the development of robust, non-lethal sampling strategies for surveillance. The detection of TiLV in mucus and water samples offers a promising alternative to destructive tissue sampling [15, 47]. The droplet digital PCR (ddPCR) assay has demonstrated the ability to detect as few as 3.3 copies of TiLV in water samples, providing an ultra-sensitive tool for environmental surveillance [15]. Future work should focus on standardizing protocols for environmental DNA (eDNA) sampling, determining the correlation between water viral load and fish infection status, and establishing threshold levels that trigger management interventions. Furthermore, the development of serological assays, such as the indirect ELISA using recombinant segment 4 protein, is crucial for assessing herd immunity and identifying past exposure [33, 47]. The pooling of serum samples for ELISA testing has been shown to be a cost-effective strategy for population-level surveillance, but its sensitivity is reduced when seroprevalence is low [33]. Future studies should optimize pooling strategies and validate these assays for use in different tilapia strains and age classes. The development of a multiplex PCR targeting multiple genome segments (2, 3, and 8) is a critical advancement to mitigate the risk of false negatives due to genetic variability or reassortment, which is a known phenomenon in TiLV [14, 16, 23]. This multi-target approach should become the standard for confirmatory diagnostics.

### Accelerating Vaccine Development and Immunoprophylaxis

The absence of a licensed commercial vaccine remains the most significant obstacle to controlling TiLVD. While numerous vaccine platforms have been explored, their efficacy is highly variable and often insufficient for field application. Inactivated vaccines, DNA vaccines, recombinant protein vaccines, and nanoparticle-based delivery systems have all been tested, with relative percent survival (RPS) values ranging from 16% to 82% [2, 8, 12, 17, 20, 21, 52]. The most promising results have been achieved with epitope-based vaccines, where a peptide derived from segment 1 (S1₃₉₉–₄₁₀) elicited an RPS of 81.8% in a laboratory challenge [12]. This approach, combining phage display technology with in silico epitope prediction, represents a rational design strategy that should be prioritized for further development.

A critical research gap is the lack of understanding of the correlates of protection. While neutralizing antibodies are likely important, their role has not been definitively established. The co-immunization of S5 and S6 proteins induced a synergistic antibody response and provided an RPS of 57.14%, suggesting that targeting multiple viral proteins may be superior to single-antigen vaccines [8]. The chitosan nanoparticle-based immersion vaccine targeting segment 4 (CNS4) showed only modest protection (RPS of 25-31%) in laboratory and field trials, indicating that the antigen itself or the delivery system requires optimization [2]. The use of biomimetic nanovaccines, such as erythrocyte membrane-coated nanoparticles, has shown promise in enhancing immune responses and improving RPS by 17-26% over the uncoated antigen [17]. Future research should systematically compare different adjuvants, delivery systems (e.g., chitosan, alginate, liposomes), and administration routes (immersion, oral, injection) to identify the most effective and practical vaccination strategy for different production scales. The development of a maternal vaccination strategy, where broodstock are immunized to passively transfer protective antibodies to offspring, is a particularly attractive approach for early-life protection [48]. However, the short persistence of maternal antibodies (undetectable by 7-14 days post-hatch) limits its utility, and strategies to extend this window, such as booster immunizations of fry, need exploration. The use of egg yolk-derived immunoglobulin Y (IgY) for passive immunization represents an alternative, cost-effective strategy that warrants further in vivo testing [32].

### Harnessing Genetic Resistance and Selective Breeding

The discovery of a major quantitative trait locus (QTL) on chromosome Oni22 that explains a substantial proportion of the genetic variance for TiLV resistance is a landmark achievement [7, 26]. This QTL, which contains candidate genes such as lgals17, vps52, and trim29, offers a direct route to marker-assisted selection (MAS) to improve resistance in breeding programs. The identification of additional QTLs on chromosome Oni09 and elsewhere suggests an oligogenic architecture, with multiple loci of moderate effect contributing to resistance [7]. Future research must fine-map these QTLs to identify the causal mutations and understand the underlying biological mechanisms. The differential expression of proteosome subunit beta type-9a and ha1f between resistant and susceptible genotypes provides a starting point for functional validation [7].

The existence of naturally resistant tilapia strains, such as the ELM strain from Lake Turkana, which showed nearly 100% survival in cohabitation challenges, highlights the potential for exploiting standing genetic variation [49]. However, a critical caveat is that resistant strains can still harbor significant viral loads in tissues like the liver and brain, potentially acting as asymptomatic carriers that perpetuate the virus in the environment [49]. Therefore, selective breeding for resistance must be coupled with strategies to reduce viral shedding, such as vaccination or culling of carriers. The Indonesian Gesit strain has also been identified as relatively resistant compared to other local strains, but its resistance is not absolute, and temperature fluctuations can exacerbate mortality [39]. Future breeding programs should aim to combine multiple resistance alleles from different strains (e.g., ELM and Gesit) to achieve more robust and durable resistance. The integration of genomic selection, using whole-genome resequencing data, will accelerate genetic gain by capturing the effects of both major and minor QTLs [7].

### Ecological and Management Strategies for Disease Mitigation

The dynamics of TiLV transmission in different aquaculture systems are poorly understood, yet this knowledge is essential for designing effective biosecurity protocols. A recent field study in recirculating aquaculture systems (RAS) demonstrated that continuous water recirculation leads to viral accumulation in the water column, prolonged outbreaks, and higher mortality compared to flow-through systems [53]. Critically, vaccination not only reduced mortality in the vaccinated fish (RPS of 55.6%) but also significantly lowered TiLV concentrations in the rearing water, thereby reducing infection pressure on unvaccinated cohorts [53]. This finding has profound implications for disease management, suggesting that vaccination can provide herd immunity in intensive systems. Future research should model the transmission dynamics of TiLV in different production systems (ponds, cages, RAS) to identify critical control points and optimize interventions such as water exchange rates, disinfection protocols, and stocking densities.

The role of the gut microbiome in modulating susceptibility to TiLV is an emerging area of research. Infection induces life-stage-dependent shifts in the gut microbiota, with fingerlings showing an expansion of Akkermansia and adults showing dominance of Acinetobacter [70]. The functional implications of these shifts are unknown, but they may influence immune maturation and antiviral defense. Probiotic supplementation has shown promise in reducing TiLV transmission by up to 80% and enhancing immune responses, potentially by modulating the gut-liver axis [59]. Similarly, dietary β-glucan derived from marine diatoms has been shown to enhance resistance, reducing cumulative mortality from 55.6% to 26.7% [43]. Future research should focus on identifying specific probiotic strains and prebiotic compounds that consistently enhance antiviral immunity, and on understanding the mechanisms by which the gut microbiota influences systemic antiviral responses. The use of feed additives, such as those tested in co-infection models with Aeromonas hydrophila, should be further explored as a practical, low-cost intervention for farmers [38].

Finally, the role of wild fish as reservoirs for TiLV is a critical but understudied aspect of disease ecology. Persistent detection of TiLV in wild tilapia and tinfoil barbs for years after an initial outbreak, without clinical signs, suggests that wild populations can serve as long-term viral reservoirs [58]. This complicates eradication efforts and necessitates a landscape-level approach to disease management. Future surveillance programs must include wild fish populations and environmental samples to understand the true geographic distribution and persistence of TiLV. The development of a structured phylogenetic framework, using concatenated ORFs 1, 3, and 5, is essential for tracking the global spread and evolution of the virus, including the detection of reassortment events that could lead to increased virulence or host range [16, 23]. The recent detection of TiLV in giant gourami (Osphronemus goramy) in Indonesia [24] and the identification of an S9-F3 homologue in a TiLV-like guppy virus [13] raise concerns about the potential for host-switching events, which must be monitored through ongoing genomic surveillance.

References

[1] Srivorakul S, Guntawang T, Sittisak T, Gordsueb T, Boonsri K, Khattiya R, et al.. Development of Reverse Transcriptase Recombinase Polymerase Amplification Combined with Lateral Flow Dipstick for Rapid Detection of Tilapia Lake Virus (TiLV): Pilot Study. Veterinary Sciences. 2025. DOI: https://doi.org/10.3390/vetsci12090845

[2] Thanapasuk C, Tattiyapong P, Yamkasem J, Kitiyodom S, Setthawong P, Lertwanakarn T, et al.. Development and Efficacy of a Chitosan Nanoparticle-Based Immersion Vaccine Targeting Segment 4 of Tilapia Lake Virus.. Journal of Fish Diseases. 2025. DOI: https://doi.org/10.1111/jfd.14106

[3] Zheng X, Xu H, Huang Y, Liu X, Zhu S, Liu H, et al.. Development of an RT-RAA-CRISPR-Cas12a Assay for Rapid, Sensitive and Visual Detection of Tilapia Lake Virus (TiLV).. Journal of Virological Methods. 2025. DOI: https://doi.org/10.1016/j.jviromet.2025.115266

[4] Arragain B, Pelosse M, Huard K, Cusack S. Structure of the tilapia lake virus nucleoprotein bound to RNA. Nucleic Acids Research. 2025. DOI: https://doi.org/10.1093/nar/gkaf112

[5] Rebecca G, Uma A, Shanmugam S, Tirumurugaan K. Massive Surveillance of Tilapia Lake Virus (TiLV) in Tilapia Farms and Feral from Various Districts of Tamil Nadu, India. Indian Journal of Animal Research. 2025. DOI: https://doi.org/10.18805/ijar.b-5558

[6] Megarani D, Yang LG, Siler HJ, Cardé EMQ, Martyniuk CJ, Hick P, et al.. One-Pot RT-LAMP CRISPR/Cas12b Platform for Rapid Detection of Tilapia Lake Virus.. Journal of Fish Diseases. 2025. DOI: https://doi.org/10.1111/jfd.70087

[7] Barría A, Nunticha P, Trịnh TQ, Mahmuddin M, Peñaloza C, Papadopoulou A, et al.. Fine mapping and functional annotation of a QTL for resistance to tilapia lake virus in Nile tilapia (Oreochromis niloticus). G3. 2025. DOI: https://doi.org/10.1093/g3journal/jkaf276

[8] Sangsuriya P, Plysup B, Lueangyangyuen A, Khrisanapant P, Senapin S, Rattanarojpong T, et al.. Co-immunization with recombinant S5196–272 and S6200–317 proteins for enhanced protective antibody response against Tilapia lake virus in Nile tilapia, Oreochromis niloticus (Linnaeus, 1758). Agriculture and Natural Resources. 2025. DOI: https://doi.org/10.34044/j.anres.2025.59.5.14

[9] Wen J, Ke Z, Wang Y, Li Y, Zhang D, Mo X, et al.. Coxsackievirus and adenovirus receptor inhibits tilapia lake virus infection via binding to viral segment 8 and 10 encoded protein.. Fish and Shellfish Immunology. 2024. DOI: https://doi.org/10.1016/j.fsi.2024.109438

[10] Arragain B, Pelosse M, Huard K, Cusack S. Structure of the Tilapia lake virus nucleoprotein bound to RNA. bioRxiv. 2024. DOI: https://doi.org/10.1101/2024.11.26.625387

[11] Saranya S, Sudhakaran R. A mini review of Tilapia Lake Virus (TiLV): Disease pathogenesis and host immune response against virus. Research journal of biotechnology. 2024. DOI: https://doi.org/10.25303/1908rjbt1050111

[12] Gong Y, Wei X, Zheng Y, Li Y, Yu Q, Li P, et al.. Combining Phage Display Technology with In Silico-Designed Epitope Vaccine to Elicit Robust Antibody Responses against Emerging Pathogen Tilapia Lake Virus. Journal of Virology. 2023. DOI: https://doi.org/10.1128/jvi.00050-23

[13] Pankaew N, Kurian D, Angelis FD, Del-Pozo J, Houston RD, Gratacap RL, et al.. Characterisation of the tilapia lake virus proteome and identification of an 11th protein, S9-F3. npj Viruses. 2026. DOI: https://doi.org/10.1038/s44298-025-00167-0

[14] Ponsrinivasan A, Uma A, Chidambaram P, Antony C, Sudhagar A, Selvaraj S, et al.. Multiplex PCR Targeting Multiple Genome Segments for Enhanced Detection of Tilapia Lake Virus (TiLV) in Tilapia. Indian Journal of Animal Research. 2026. DOI: https://doi.org/10.18805/ijar.b-5687

[15] Shahi N, Prasartset T, Surachetpong W. A specific and sensitive droplet digital polymerase chain reaction assay for the detection of tilapia lake virus in fish tissue and environmental samples.. Journal of Fish Diseases. 2023. DOI: https://doi.org/10.1111/jfd.13816

[16] Abbadi M, Basso A, Biasini L, Quartesan R, Buratin A, Davidovich N, et al.. Tilapia lake virus: A structured phylogenetic approach. Frontiers in Genetics. 2023. DOI: https://doi.org/10.3389/fgene.2023.1069300

[17] Wei X, Gong Y, Xia J, Liu M, Li P, Wang G, et al.. Biomimetic nanovaccine based on erythrocyte membrane enhances immune response and protection against tilapia lake virus.. Virology. 2023. DOI: https://doi.org/10.1016/j.virol.2023.01.014

[18] Aich N, Paul A, Choudhury TG, Saha H. Tilapia Lake Virus (TiLV) disease: Current status of understanding. Aquaculture and Fisheries. 2021. DOI: https://doi.org/10.1016/J.AAF.2021.04.007

[19] Rass RA, Kustin T, Zamostiano R, Smorodinsky N, Meir DB, Feder D, et al.. Inferring Protein Function in an Emerging Virus: Detection of the Nucleoprotein in Tilapia Lake Virus. Journal of Virology. 2022. DOI: https://doi.org/10.1128/jvi.01757-21

[20] Tattiyapong P, Kitiyodom S, Yata T, Jantharadej K, Adamek M, Surachetpong W. Chitosan nanoparticle immersion vaccine offers protection against tilapia lake virus in laboratory and field studies.. Fish and Shellfish Immunology. 2022. DOI: https://doi.org/10.1016/j.fsi.2022.10.063

[21] Chamtim P, Suwan E, Dong H, Sirisuay S, Areechon N, Wangkahart E, et al.. Combining segments 9 and 10 in DNA and recombinant protein vaccines conferred superior protection against tilapia lake virus in hybrid red tilapia (oreochromis sp.) compared to single segment vaccines. Frontiers in Immunology. 2022. DOI: https://doi.org/10.3389/fimmu.2022.935480

[22] Islam SI, Mahfuj S, Alam MA, Ara Y, Sanjida S, Mou M. Immunoinformatic Approaches to Identify Immune Epitopes and Design an Epitope-Based Subunit Vaccine against Emerging Tilapia Lake Virus (TiLV). Aquaculture Journal. 2022. DOI: https://doi.org/10.3390/aquacj2020010

[23] Tran T, Nguyen VT, Bui H, Tran YT, Tran H, Le TT, et al.. Tilapia Lake Virus (TiLV) from Vietnam is genetically distantly related to TiLV strains from other countries.. Journal of Fish Diseases. 2022. DOI: https://doi.org/10.1111/jfd.13669

[24] Jumria A, Amin M, Mufasirin M. Analisis Molekuler Tilapia Lake Virus (TiLV) di Sentra Budidaya Benih Ikan Gurami (Osphronemus goramy) Jawa Timur. Bioscientist. 2023. DOI: https://doi.org/10.33394/bioscientist.v11i2.9515

[25] Delamare‐Deboutteville J, Meemetta W, Pimsannil K, Sangpo P, Gan H, Mohan C, et al.. A multiplexed RT-PCR assay for nanopore whole genome sequencing of Tilapia lake virus (TiLV). bioRxiv. 2023. DOI: https://doi.org/10.1101/2023.04.24.537954

[26] Barría A, Trịnh TQ, Mahmuddin M, Peñaloza C, Papadopoulou A, Gervais O, et al.. A major quantitative trait locus affecting resistance to Tilapia Lake Virus in farmed Nile tilapia (Oreochromis niloticus). Heredity. 2021. DOI: https://doi.org/10.1038/s41437-021-00447-4

[27] Rass RA, Kembou-Ringert JE, Zamostiano R, Eldar A, Ehrlich M, Bacharach E. Mapping of Tilapia Lake Virus entry pathways with inhibitors reveals dependence on dynamin activity and cholesterol but not endosomal acidification. Frontiers in Cell and Developmental Biology. 2022. DOI: https://doi.org/10.3389/fcell.2022.1075364

[28] Jansen MD, Dong H, Mohan C. Tilapia lake virus: a threat to the global tilapia industry?. Reviews in Aquaculture. 2019. DOI: https://doi.org/10.1111/RAQ.12254

[29] Surachetpong W, Roy S, Nicholson P. Tilapia lake virus: The story so far.. Journal of Fish Diseases. 2020. DOI: https://doi.org/10.1111/jfd.13237

[30] Megarani DV, Al-Hussinee L, Subramaniam K, Sriwanayos P, Imnoi K, Keleher B, et al.. Development of a TaqMan quantitative reverse transcription PCR assay to detect tilapia lake virus.. Diseases of Aquatic Organisms. 2022. DOI: https://doi.org/10.3354/dao03700

[31] Chengula A, Mugimba KK, Tal S, Levi RT, Dubey S, Mutoloki S, et al.. Efficiency, sensitivity and specificity of a quantitative real-time PCR assay for tilapia lake virus (TiLV).. Journal of Virological Methods. 2022. DOI: https://doi.org/10.1016/j.jviromet.2022.114567

[32] Setthawong P, Yamkasem J, Khemthong M, Tattiyapong P, Metheenukul P, Prasertsincharoen N, et al.. Development of IgY-Based Passive Immunization Against Tilapia Lake Virus: Development and In Vitro Neutralization Assays. Viruses. 2025. DOI: https://doi.org/10.3390/v17030448

[33] Yamkasem J, Tattiyapong P, Gardner IA, Surachetpong W. Assessment and Performance of Pooled Serum Samples for Monitoring Farm-Level Immunity in Tilapia Infected with Tilapia Lake Virus. Viruses. 2025. DOI: https://doi.org/10.3390/v17070877

[34] Youngnoi N, Yamkasem J, Khemthong M, Setthawong P, Nedumpun T, Surachetpong W, et al.. Granulocyte tropism and lymphocyte depletion highlight the immunopathogenesis of tilapia lake virus infection in Nile tilapia.. Fish and Shellfish Immunology. 2025. DOI: https://doi.org/10.1016/j.fsi.2025.110410

[35] Feng M, Liu Q, Wang J, Li B, Lu H, Hu J, et al.. Interferon regulatory factor 6 gene from Oreochromis niloticus is involved in resistance against tilapia lake virus.. Fish and Shellfish Immunology. 2025. DOI: https://doi.org/10.1016/j.fsi.2025.111096

[36] Fitria D, Fathonah D, Afrianti R, Turrohmah A, Mujaidiah, Destiana AL, et al.. Pemeriksaan TiLV (Tilapia Lake Virus) Pada Ikan Nila (Oreochromis niloticus) Budidaya. PENDIPA Journal of Science Education. 2025. DOI: https://doi.org/10.33369/pendipa.9.2.310-317

[37] Karmokar S, Das S, Priyadarsani L, Dash G, Abraham T. Surveillance of tilapia lake virus (TiLV) in diverse aquaculture systems of the Indian Sundarbans and adjoining regions in West Bengal, India. Indian Journal of Fisheries. 2025. DOI: https://doi.org/10.21077/ijf.2025.72.3.166556-15

[38] Mohamad A, Yamkasem J, Paimeeka S, Khemthong M, Lertwanakarn T, Setthawong P, et al.. Efficacy of Feed Additives on Immune Modulation and Disease Resistance in Tilapia in Coinfection Model with Tilapia Lake Virus and Aeromonas hydrophila. Biology. 2024. DOI: https://doi.org/10.3390/biology13110938

[39] Taukhid T, Lusiastuti A, Novita H, Caruso D, Avarre J, Sugiani D, et al.. Resistance of Indonesian tilapia strains, Oreochromis niloticus to Tilapia Lake Virus Disease (TiLVD). Bulletin of The European Association of Fish Pathologists. 2024. DOI: https://doi.org/10.48045/001c.118454

[40] Turner J, Sakulpolwat S, Sukdanon S, Lertwanakarn T, Waiyamitra P, Piewbang C, et al.. Tilapia lake virus (TiLV) causes severe anaemia and systemic disease in tilapia.. Journal of Fish Diseases. 2023. DOI: https://doi.org/10.1111/jfd.13775

[41] Sihananto BS, Wijayanti E. INTRASITOPLASMIC INCLUSION BODIES OF HEPATOSIT, SYNCYTIAL CELLS IN BRAIN AND LIVER AS KEY TO IDENTIFICATION OF TILAPIA LAKE VIRUS (TiLV) BY USING HISTOPATOLOGICAL METHODS IN TILAPIA (Oreochromis niloticus) WITH MOTILE AEROMONAS SEPTICEMIA (MAS) SECONDARY. Fisheries Science. 2024. DOI: https://doi.org/10.20527/fishscientiae.v14i1.235

[42] Megarani D, Martyniuk CJ, Yanong R, Pierezan F, Surachetpong W, Subramaniam K. Tracing a Decade of Tilapia Lake Virus Research: Breakthroughs, Challenges, and Projections. Reviews in Aquaculture. 2026. DOI: https://doi.org/10.1111/raq.70148

[43] Madyod S, Pholmai S, Prombanchong T, Laksana-aut P, Unajak S, Hirono I, et al.. Dietary β‐Glucan From the Marine Diatom Chaetoceros muelleri Enhances Immune Responses and Resistance of Nile Tilapia to Tilapia Lake Virus Infection. Journal of Fish Diseases. 2026. DOI: https://doi.org/10.1111/jfd.70184

[44] Saranya S, Sudhakaran R. Temporal Dynamics of Viral Load and Innate Immune Gene Expression in Tilapia (Oreochromis niloticus) Experimentally Infected with Tilapia Lake Virus (TiLV). Research journal of biotechnology. 2026. DOI: https://doi.org/10.25303/216rjbt015024

[45] Raksaseri P, Lertwanakarn T, Tattiyapong P, Kijtawornrat A, Klomkleaw W, Surachetpong W. Tilapia lake virus causes mitochondrial damage: a proposed mechanism that leads to extensive death in fish cells. PeerJ. 2023. DOI: https://doi.org/10.7717/peerj.16190

[46] Valsalam A, Bedekar M, Kezhedath J, Sood N, Poojary N, Rajendran KV, et al.. Isolation, in vitro, and in vivo pathogenicity test of Tilapia lake virus (TiLV) and development of a prognostic semi-quantitative lesion scoring system for differentiating clinical/subclinical infection in farmed tilapia (Oreochromis niloticus L.).. Microbial Pathogenesis. 2023. DOI: https://doi.org/10.1016/j.micpath.2023.106475

[47] Valsalam A, Rajendran K, Kezhedath J, Godavarikar A, Sood N, Bedekar M. Development of an indirect ELISA test for the detection of Tilapia lake virus (TiLV) in fish tissue and mucus samples.. Journal of Virological Methods. 2023. DOI: https://doi.org/10.1016/j.jviromet.2023.114707

[48] Mai T, Kayansamruaj P, Soontara C, Kerddee P, Nguyen D, Senapin S, et al.. Immunization of Nile Tilapia (Oreochromis niloticus) Broodstock with Tilapia Lake Virus (TiLV) Inactivated Vaccines Elicits Protective Antibody and Passive Maternal Antibody Transfer. Vaccines. 2022. DOI: https://doi.org/10.3390/vaccines10020167

[49] Adamek M, Rebl A, Matras M, Lodder C, Rahman SE, Stachnik M, et al.. Immunological insights into the resistance of Nile tilapia strains to an infection with tilapia lake virus.. Fish and Shellfish Immunology. 2022. DOI: https://doi.org/10.1016/j.fsi.2022.03.027

[50] Sood N, Verma DK, Paria A, Yadav SC, Yadav M, Bedekar M, et al.. Transcriptome analysis of liver elucidates key immune-related pathways in Nile tilapia Oreochromis niloticus following infection with tilapia lake virus.. Fish and Shellfish Immunology. 2021. DOI: https://doi.org/10.1016/j.fsi.2021.02.005

[51] Akter M, Punom NJ, Eshik ME, Ahmmed S, Rabbane M, Rahman MS. Investigation of Tilapia (Oreochromis niloticus) Mortality Events Targeting Tilapia Lake Virus Disease (TiLVD) in Bangladesh. Veterinary Medicine and Science. 2025. DOI: https://doi.org/10.1002/vms3.70262

[52] Kembou-Ringert JE, Steinhagen D, Readman JAJ, Daly J, Adamek M. Tilapia Lake Virus Vaccine Development: A Review on the Recent Advances. Vaccines. 2023. DOI: https://doi.org/10.3390/vaccines11020251

[53] Sresung M, Yamkasem J, Theplhar S, Setthawong P, Rattanakul S, Mongkolsuk S, et al.. Dynamics of Tilapia Lake Virus in Recirculating Aquaculture Systems and the Impact of Vaccination on Outbreak Control. Viruses. 2026. DOI: https://doi.org/10.3390/v18010096

[54] Noerhidayanti F, Jafar FYA, Fitriyana F. Deteksi Molekuler Tilapia Lake Virus (TiLV) pada Ikan Nila (Oreochromis niloticus) di Provinsi Kalimantan Timur, Indonesia. Inovasi Kesehatan Global. 2026. DOI: https://doi.org/10.62383/ikg.v3i1.2756

[55] Dita S, Laela N, Putri D, Laminem, Anita T, Laili R, et al.. Detection of Tilapia lake virus (TiLV) using semi-nested RT-PCR method in farmed Tilapia (Oreochromis niloticus) from ponds East Java, Indonesia. IOP Conference Series: Earth and Environment. 2023. DOI: https://doi.org/10.1088/1755-1315/1273/1/012076

[56] Bimantara A, Anindita NS, Amin TRA, Wantonoro W, Setianah H. Detection of Tilapia Lake Virus (TiLV) in Tilapia (Oreochromis niloticus) by Semi-Nested Reverse Transcriptase Polymerase Chain Reaction (RT-PCR) Method. International Journal of Health Science and Technology. 2023. DOI: https://doi.org/10.31101/ijhst.v5i2.3316

[57] . Tilapia lake virus. CABI Compendium. 2022. DOI: https://doi.org/10.1079/cabicompendium.15961926

[58] Abdullah A, Pazai AMM, Ridzuan M, Sudirwan F, Hashim S, Abas A, et al.. Persistent detection of Tilapia lake virus in wild tilapia and tinfoil barbs. Veterinary World. 2022. DOI: https://doi.org/10.14202/vetworld.2022.1097-1106

[59] Yang Y, Yamkasem J, Surachetpong W, Lin Y, You S, Lu T, et al.. Assessing the effect of probiotics on tilapia lake virus-infected tilapia: Transmission and immune response.. Journal of Fish Diseases. 2022. DOI: https://doi.org/10.1111/jfd.13635

[60] Mohamad A, Khemthong M, Trongwongsa P, Lertwanakarn T, Setthawong P, Surachetpong W. A New Cell Line from the Brain of Red Hybrid Tilapia (Oreochromis spp.) for Tilapia Lake Virus Propagation. Animals. 2024. DOI: https://doi.org/10.3390/ani14111522

[61] Sanyalukruechai C, Watthanasakphuban N, Khemthong M, Surachetpong W, Rattanaporn K. Expression and purification of recombinant tilapia lake virus segment 4 protein and its in-vitro biological activity for potential use in vaccine development. Scientific Reports. 2024. DOI: https://doi.org/10.1038/s41598-024-83293-8

[62] Saranya S, Sudhakaran R. Sensitive and cost-effective citrate-based RNA extraction procedure for isolation of RNA from Tilapia Lake Virus-infected fish. Journal of Applied Biology & Biotechnology. 2024. DOI: https://doi.org/10.7324/jabb.2024.196791

[63] Cabal ABS, Choresca C, Hsieh F, Wu T. In-Vitro and In-Vivo Recombinant Expression of Tilapia Lake Virus S4 Protein Using Extended Host-Range Baculovirus Expression Vector and Generation of Polyclonal Anti-S4 Antibodies. Processes. 2026. DOI: https://doi.org/10.3390/pr14101642

[64] Kembou-Ringert JE, Steinhagen D, Thompson K, Daly J, Adamek M. Immune responses to Tilapia lake virus infection: what we know and what we don’t know. Frontiers in Immunology. 2023. DOI: https://doi.org/10.3389/fimmu.2023.1240094

[65] Barde I, Ishaku LE, Melba B. Mapping the research landscape: A bibliometric analysis of Tilapia Lake Virus disease worldwide. Global Journal of Fisheries Science. 2024. DOI: https://doi.org/10.31248/gjfs2024.058

[66] Pande A, Thakuria D, Kushwaha B, Kumar R, S M, Rastogi A, et al.. A cell line derived from heart of rainbow trout is refractory to Tilapia lake virus. Cell Biology International. 2024. DOI: https://doi.org/10.1002/cbin.12125

[67] Sumon MAA, Asseri AH, Molla MHR, Aljahdali MO, Hasan MR, Rahman M, et al.. Identification of natural antiviral drug candidates against Tilapia Lake Virus: Computational drug design approaches. bioRxiv. 2023. DOI: https://doi.org/10.1371/journal.pone.0287944

[68] Alisukma ISP. Deteksi Molekuler Tilv (Tilapia Lake Virus) pada Ikan Nila (Oreochromisniloticus) yang dilalulintaskan di Balai Karantina Ikan, Pengendalian Mutu dan Keamanan Hasil Perikanan Surabaya I, Jawa Timur. Media Kedokteran Hewan. 2023. DOI: https://doi.org/10.20473/mkh.v34i1.2023.1-12

[69] Li B, Zheng S, Wang Y, Wang Q, Li Y, Yin J, et al.. Susceptibilities of ten fish cell lines to infection with Tilapia lake virus.. Microbial Pathogenesis. 2022. DOI: https://doi.org/10.1016/j.micpath.2022.105510

[70] Gopan A, Kollanoor RJ. Gut Microbiome Dynamics in Farmed Tilapia: A Comparative Analysis of Fingerlings and Adults Infected with Tilapia Lake Virus. Fishery technology. 2025. DOI: https://doi.org/10.56093/ft.v62i4.166640