Scale Drop Disease Virus: Aquaculture Reference

Overview and Taxonomy of Scale Drop Disease Virus: A Megalocytivirus Threat to Global Teleost Aquaculture

Scale drop disease virus (SDDV) represents a distinct and emerging etiological agent within the family Iridoviridae, a lineage of large, double-stranded DNA (dsDNA) viruses renowned for their impact on aquatic ectothermic vertebrates. Since its initial characterization in 2015, SDDV has been unequivocally established as the causative agent of scale drop syndrome (SDS), a disease that has precipitated significant economic losses and high mortality in global finfish aquaculture, particularly within the Asia-Pacific region [4, 14]. The taxonomic placement of SDDV has been refined through comprehensive genomic and phylogenetic analyses, solidifying its position as a unique member of the genus Megalocytivirus, a group whose significance is underscored by the listing of certain members as notifiable to the World Organisation for Animal Health (WOAH) [16]. Understanding the precise taxonomic identity and biological characteristics of SDDV is paramount not only for accurate diagnostic efforts but also for elucidating its pathogenic mechanisms, host range, and evolutionary dynamics within the complex aquaculture virome.

Taxonomic Classification and Phylogenetic Position

The family Iridoviridae is subdivided into two subfamilies: Alphairidovirinae, which infects vertebrates (fish, amphibians, and reptiles), and Betairidovirinae, which primarily infects arthropods and annelids [16]. Within the subfamily Alphairidovirinae, the genus Megalocytivirus has emerged as one of the most economically significant groups of piscine pathogens. Traditional classification has recognized several distinct genotypes, including infectious spleen and kidney necrosis virus (ISKNV), red sea bream iridovirus (RSIV), turbot reddish body iridovirus (TRBIV), threespine stickleback iridovirus (TSIV), and scale drop disease virus (SDDV) [15]. However, a recent and authoritative reclassification by the International Committee on Taxonomy of Viruses (ICTV), as reflected in the literature, has consolidated these genotypes into two officially recognized viral species within the genus Megalocytivirus. The first is Megalocytivirus pagrus 1, which encompasses the ISKNV, RSIV, and TRBIV genogroups. The second, and critically for this review, is Megalocytivirus lates 1, which comprises a single genotype: SDDV [16]. This designation highlights the distinct evolutionary trajectory of SDDV relative to the other megalocytiviruses.

Phylogenetic analyses based on core viral genes, most notably the major capsid protein (MCP) and the ATPase gene, consistently demonstrate that SDDV isolates form a robust, monophyletic clade that is clearly delineated from other Megalocytivirus lineages [4, 12, 13]. The MCP gene of SDDV, for instance, exhibits high sequence homology among isolates (often exceeding 99% at the nucleotide level), yet it clusters separately from ISKNV and RSIV homologs, confirming its status as a distinct viral species [7, 8, 14]. This genetic distance is further supported by whole-genome comparative analyses, which have revealed that while the general genome organization is conserved among megalocytiviruses, SDDV possesses unique regions and lineage-specific open reading frames (ORFs) that are not present in M. pagrus 1 genotypes. For example, the genome of SDDV isolate TH2019 was found to harbor a unique 7,695-bp genomic region encoding six hypothetical proteins, a feature that is absent in ISKNV and RSIV genomes [10]. Such genomic signatures provide robust molecular markers for distinguishing SDDV infections from other megalocytiviral diseases.

Genomic Architecture and Genetic Diversity

The SDDV genome is a large, linear dsDNA molecule, typical of the Iridoviridae. Genomic characterization of multiple isolates from different geographical locations and host species has revealed a genome size ranging from approximately 131 kb to 132 kb, with a predicted complement of 134 to 143 putative open reading frames (ORFs) [4, 7, 12, 13]. The complete genome of the Singaporean reference strain, along with genomes from Thailand (e.g., TH7_2019), Malaysia, and China (e.g., ZH-06/20, SDDV-ZH918/23), have been sequenced, providing a rich foundation for comparative genomics [7, 10, 12, 14]. These analyses have consistently shown a high degree of nucleotide and amino acid identity across all SDDV isolates, often exceeding 99%, indicating a high level of genomic conservation despite geographic separation [4, 7, 10]. For instance, the SDDV TH7_2019 genome from Thailand shared 99.97% nucleotide identity with the Singaporean reference genome within the aligned regions [10].

Despite this overall conservation, detailed scrutiny has identified specific genetic variations that may serve as biomarkers for epidemiological tracking and strain classification. Among the 134 predicted ORFs, missense, frameshift, insertion, and deletion mutations have been documented in several core genes [10]. Notably, a consistent deletion of four deduced amino acids in the product of ORF_030L was identified in Thai SDDV isolates, suggesting this could be a potential biomarker for strain differentiation within Southeast Asia [10]. Furthermore, a more recent study documented, for the first time, a 152 bp deletion mutant within two SDDV isolates from yellowfin seabream in China, demonstrating that the viral genome is subject to dynamic mutational events, including the formation of partial deletion mutants [7]. The biological implications of these deletions, whether they affect virulence, transmissibility, or host range, remain a critical area for future investigation. The majority of annotated ORFs encode hypothetical proteins with unknown functions, but key structural and enzymatic components have been identified, including the MCP, the ATPase, and a suite of proteins potentially involved in manipulating host apoptosis and immune responses [4]. The MCP itself is of particular interest, as it has been demonstrated to directly interact with the host entry receptor, transferrin receptor 1 (TfR1), to mediate viral internalization [1].

Virion Structure and Morphology

The physical architecture of SDDV is consistent with that of other iridoviruses. Electron microscopy studies performed on infected cell cultures, primarily using the MFF-1 cell line derived from mandarin fish (Siniperca chuatsi), have revealed icosahedral virions with a diameter of approximately 140 nm [12, 14]. These virions are observed within the cytoplasm of infected cells, accumulating in paracrystalline arrays, a hallmark of iridovirus replication [14]. The virion itself is non-enveloped, though an external lipid membrane is often associated with the capsid, facilitating entry via endocytic pathways. The icosahedral capsid is composed of repeated subunits of the MCP, which provides structural stability and serves as the primary antigenic target for host antibody responses [3, 14]. The identification of 113 viral proteins within the purified virion proteome of the ZH-06/20 isolate underscores the complexity of the SDDV particle, which includes not only structural components but also enzymes required for the initiation of replication within the host cell [12].

Pathobiological Significance and Host Range

SDDV is recognized as a significant threat to global teleost aquaculture, with its emergence linked to severe disease outbreaks across Southeast Asia and now extending into Southern China [16]. The primary host is the Asian seabass (Lates calcarifer), also known as barramundi, in which the virus causes the characteristic and namesake scale drop disease [14]. Clinical signs in this species include lethargy, severe scale loss (the hallmark of the disease), darkened dorsal pigmentation, reddening and hemorrhaging on the ventral body, and ascites in some cases [8, 14]. Mortality rates can be exceedingly high, with laboratory challenges and field outbreaks reporting mortality approaching 60–100% in juvenile fish [3, 12, 14].

Critically, the host range of SDDV has expanded beyond Asian seabass. Recent virological surveys have identified SDDV as the causative agent of a distinct disease, yellowfin seabream (Acanthopagrus latus) ascites disease (YFSBAD), in Southern China. Outbreaks in this species were characterized by severe ascites and high mortality, but notably, the typical scale drop symptom was absent, suggesting host-dependent pathophysiological variation [12]. This discovery highlights the phenotypic plasticity of SDDV infections and underscores the necessity for robust diagnostic methods that do not rely solely on clinical signs. The virus has also been molecularly detected in freshwater-reared barramundi, often in co-infections with other pathogens like Flavobacterium columnare, indicating a broader ecological niche and potential adaptation to diverse environmental conditions [9]. Furthermore, SDDV has been detected in co-infections with Lates calcarifer herpes virus (LCHV) in commercial mariculture settings, where co-infection leads to more severe immunosuppression and necrotic pathology, particularly in the spleen and kidney [11].

The molecular basis for SDDV's pathogenesis is an area of vigorous research. A landmark study demonstrated that SDDV utilizes the host transferrin receptor 1 (TfR1) as an entry receptor, binding to it via the viral MCP and entering host cells through clathrin-mediated endocytosis [1]. This interaction is critical for establishing infection, as blocking TfR1 with antibodies or the inhibitor ferristatin II significantly reduces viral entry. Furthermore, SDDV has been shown to exploit host iron metabolism. Infection upregulates TfR1 expression, leading to iron overload, reactive oxygen species (ROS) accumulation, lipid peroxidation, and glutathione depletion, culminating in a non-apoptotic, iron-dependent form of cell death known as ferroptosis [5, 6]. This process is not merely a bystander effect but actively facilitates viral replication, as inhibition of ferroptosis with Ferrostatin-1 or iron chelators significantly reduces viral titers and improves host survival [5, 6]. The virus also appears to manipulate host antioxidant responses; for instance, Vitamin C (VC) has been shown to inhibit SDDV infectivity by activating the Nrf2 pathway, which counteracts ferroptosis, suggesting a potential therapeutic avenue [2]. These findings collectively position SDDV as a highly adapted pathogen that subverts fundamental cellular processes, including iron homeostasis and cell death pathways, to its advantage. Understanding these mechanisms is not only fundamental virology but also provides actionable targets for developing antiviral strategies, such as receptor mimics or ferroptosis inhibitors, to mitigate the devastating impact of this virus on global aquaculture.

Molecular Pathogenesis of SDDV: Transferrin Receptor 1-Mediated Entry and Clathrin-Dependent Endocytosis

The initial step in any viral infection, the binding of the virion to a specific host cell receptor and subsequent internalization, is a critical determinant of host range, tissue tropism, and ultimately, pathogenicity. For scale drop disease virus (SDDV), a distinct and emerging member of the genus Megalocytivirus within the family Iridoviridae, the molecular details of this process remained entirely enigmatic until very recently [1, 4, 14]. The identification of the mandarin fish transferrin receptor 1 (mfTfR1) as the primary entry receptor for SDDV, and the subsequent elucidation of a clathrin-dependent endocytic pathway, represents a paradigm-shifting advancement in our understanding of megalocytiviral pathogenesis [1]. This discovery not only provides a mechanistic framework for SDDV’s ability to establish lethal systemic infections but also reveals a striking evolutionary convergence with the entry strategies employed by certain mammalian and avian viruses, highlighting TfR1 as a conserved Achilles' heel across diverse host taxa [1].

The Identification of Transferrin Receptor 1 (TfR1) as a Critical Host Factor

The journey to identifying mfTfR1 began with proteomic analyses of purified SDDV virions, which unexpectedly revealed a high abundance of this host transmembrane protein associated with the viral particle [1]. This initial observation suggested a specific and potentially functional interaction between SDDV and TfR1, rather than mere passive incorporation. The physiological role of TfR1 is to mediate the cellular uptake of iron-bound transferrin (holo-transferrin) via clathrin-mediated endocytosis, a process essential for iron homeostasis and cellular metabolism [1]. The presence of TfR1 on the virion surface hinted at a viral strategy to hijack this well-characterized internalization machinery.

Functional validation of TfR1’s role in SDDV infection was achieved through a series of complementary experiments. First, the expression of mfTfR1 was shown to be dynamically regulated in response to SDDV infection, both in vivo in mandarin fish and in vitro in MFF-1 cells, suggesting a virus-host interplay at the transcriptional level [1, 5]. More compellingly, genetic manipulation of TfR1 expression levels directly correlated with viral permissivity. Overexpression of mfTfR1 in fathead minnow (FHM) cells, which are naturally low-permissive to SDDV, dramatically enhanced viral replication, particularly during the early stages of binding and entry [1]. Conversely, blocking the availability of TfR1, either through specific antibodies or the pharmacological inhibitor ferristatin II, potently suppressed SDDV entry and subsequent infection [1]. These loss- and gain-of-function experiments provided unequivocal evidence that mfTfR1 is not merely a bystander but an indispensable molecular gateway for SDDV invasion.

Molecular Interaction: The MCP-TfR1 Axis and the Helical Domain Binding Site

The specificity of the virus-receptor interaction was further refined by demonstrating a direct physical association between the SDDV major capsid protein (MCP) and mfTfR1 [1]. The MCP is the most abundant structural protein of iridoviruses, forming the icosahedral capsid that encapsulates the viral genome [14, 16]. Co-immunoprecipitation and pull-down assays confirmed that the MCP of SDDV specifically binds to mfTfR1, establishing the molecular basis for viral attachment [1].

The precise docking site on the receptor was mapped to the helical domain of mfTfR1 [1]. This is a functionally significant finding, as the helical domain of TfR1 in other species is known to be involved in receptor dimerization and interaction with transferrin. By targeting this domain, SDDV may have evolved to engage a structurally conserved and functionally critical region of the receptor, potentially minimizing the ability of the host to evade infection through simple point mutations. The identification of the specific binding interface within this helical domain opens the door for the rational design of competitive inhibitors or peptide-based therapeutics that could disrupt the MCP-TfR1 interaction [1]. Indeed, disrupting this interaction through site-directed mutagenesis or competitive peptides was shown to significantly reduce viral entry and, critically, host mortality in challenge experiments, underscoring the in vivo relevance of this binding event [1].

Hijacking the Endocytic Machinery: Clathrin-Mediated Endocytosis and Src Kinase Signaling

Following receptor binding, SDDV must traverse the plasma membrane to deliver its genome into the host cell cytoplasm. The data strongly support a model in which SDDV, like its physiological ligand transferrin, exploits the clathrin-mediated endocytosis (CME) pathway [1]. This is a highly regulated process involving the recruitment of clathrin triskelions to the inner leaflet of the plasma membrane, forming coated pits that invaginate and pinch off to form clathrin-coated vesicles.

The involvement of CME in SDDV entry was confirmed through multiple lines of evidence. Pharmacological inhibition of key components of the CME pathway, such as with chlorpromazine or pitstop 2, significantly blocked SDDV infection [1]. Furthermore, the internalization process was shown to be dependent on the activation of Src family kinases. SDDV infection was found to trigger Src kinase-mediated tyrosine phosphorylation of mfTfR1 [1]. This phosphorylation event is a critical signaling cue that enhances the internalization of the receptor-ligand complex, effectively accelerating the rate of CME. This mechanism is analogous to that employed by some other viruses that use TfR1, where receptor phosphorylation acts as a molecular switch to promote endocytosis [1]. The activation of Src signaling by SDDV represents a sophisticated viral strategy to actively commandeer the host cell’s endocytic apparatus, ensuring efficient and rapid internalization of the viral particle.

Implications for Pathogenesis and Tissue Tropism

The utilization of TfR1 as an entry receptor has profound implications for SDDV pathogenesis. TfR1 is ubiquitously expressed on the surface of actively dividing and metabolically active cells, particularly in tissues with high iron demands, such as the hematopoietic organs (spleen and kidney), the liver, and the intestinal epithelium. This broad expression pattern provides a mechanistic explanation for the systemic nature of SDDV infection. Histopathological studies consistently demonstrate that SDDV targets a wide range of organs, including the spleen, kidney, liver, gills, brain, and heart, leading to severe necrosis and inflammation [8, 11, 12, 17]. The presence of TfR1 on these diverse cell types allows SDDV to disseminate rapidly throughout the host, causing the multi-organ failure characteristic of scale drop disease.

Furthermore, the link between TfR1 and iron metabolism creates a direct nexus between viral entry and the induction of ferroptosis, a non-apoptotic form of cell death driven by iron-dependent lipid peroxidation [2, 5, 6]. SDDV infection has been shown to upregulate TfR1 expression, leading to iron overload, accumulation of reactive oxygen species (ROS), and depletion of glutathione peroxidase 4 (GPX4), all hallmarks of ferroptosis [5, 6]. This is not a passive consequence of infection but an active process that facilitates viral replication. Inhibiting ferroptosis with ferrostatin-1 or iron chelators significantly reduces viral titers and improves survival in mandarin fish [5, 6]. Thus, the TfR1-mediated entry pathway is intimately linked to the downstream induction of ferroptosis, creating a vicious cycle where the virus uses the receptor for entry, upregulates its expression to disrupt iron homeostasis, and triggers a cell death pathway that ultimately aids in viral release and spread. This dual role of TfR1, as both an entry portal and a driver of pathogenic cell death, positions it as a central node in the SDDV-host interaction network.

Evolutionary and Comparative Virology Context

The discovery that SDDV, a large double-stranded DNA virus of fish, uses TfR1 for entry is remarkable from a comparative virology perspective. TfR1 has been co-opted as an entry receptor by a diverse array of viruses, including several mammalian RNA viruses (e.g., some arenaviruses and retroviruses) and small DNA viruses (e.g., canine parvovirus and mouse mammary tumor virus) [1]. The fact that SDDV, an iridovirus with a vastly different genome size and replication strategy, has convergently evolved to target the same receptor underscores the fundamental importance of TfR1 as a viral entry portal across the animal kingdom. This conservation suggests that TfR1 possesses intrinsic properties, such as high surface expression, constitutive recycling via CME, and a structurally stable extracellular domain, that make it an attractive target for viral exploitation.

This finding also distinguishes SDDV from other megalocytiviruses. While the entry mechanisms for infectious spleen and kidney necrosis virus (ISKNV) and red sea bream iridovirus (RSIV) are less well-defined, the specific reliance on TfR1 may be a unique feature of Megalocytivirus lates 1 (SDDV) [16]. In contrast, other iridoviruses like decapod iridescent virus 1 (DIV1) have been shown to enter cells via caveola-mediated endocytosis, a distinct pathway from the clathrin-dependent route used by SDDV [18]. This divergence in entry mechanisms highlights the evolutionary plasticity within the Iridoviridae family and has significant implications for the development of broad-spectrum versus virus-specific antiviral strategies. The World Organisation for Animal Health (WOAH) lists ISKNV as a notifiable pathogen, and the growing understanding of SDDV’s unique entry mechanism may warrant similar surveillance and targeted intervention efforts for this emerging pathogen [16].

Epidemiology and Host Range of Scale Drop Disease Virus in Aquaculture Systems

The emergence of scale drop disease virus (SDDV) as a distinct and highly pathogenic member of the genus Megalocytivirus within the family Iridoviridae represents a paradigm-shifting challenge for global teleost aquaculture. Unlike its better-characterized congeners, Megalocytivirus pagrus 1 (encompassing infectious spleen and kidney necrosis virus [ISKNV], red sea bream iridovirus [RSIV], and turbot reddish body iridovirus [TRBIV]), SDDV (classified as Megalocytivirus lates 1) exhibits a unique ecological niche, a rapidly expanding geographic footprint, and a profoundly concerning capacity for eliciting lethal disease in an expanding roster of commercially vital fish species [16]. The epidemiology of SDDV is not a static picture but a dynamic, unfolding crisis driven by intensive aquaculture practices, international movement of live fish and germplasm, and the virus’s sophisticated molecular strategies for subverting host defenses. A comprehensive understanding of its distribution, host range, transmission dynamics, and the ecological co-factors that precipitate outbreaks is therefore not merely an academic exercise, it is an absolute prerequisite for designing evidence-based biosecurity protocols, risk-based surveillance systems, and effective vaccine deployment strategies.

Global Dissemination and Geographic Distribution

The first formal description of SDDV as the aetiological agent of scale drop syndrome (SDS) in Asian seabass (Lates calcarifer) was a landmark study from Singapore in 2015, which fulfilled Koch’s postulates and identified the virus as a novel megalocytivirus [14]. However, retrospective analyses and subsequent molecular surveillance have revealed that SDDV had been circulating in Southeast Asian aquaculture systems for at least two decades prior, with clinical observations of a “previously unknown illness” characterized by scale loss reported as early as 1992 [14]. The virus is now firmly established as an endemic pathogen across the major mariculture hubs of Southeast Asia. Following the initial Singaporean description, SDDV was rapidly confirmed in Thailand, where it was detected in both marine and, critically, freshwater-reared barramundi, expanding the perceived risk envelope [9]. The incursion into Indonesia was documented in floating net cages in Batam, where a mass mortality event affecting fish of market size (0.3–2.0 kg) was retrospectively linked to SDDV via PCR and sequencing, with the Indonesian isolate sharing 100% nucleotide identity with the Singaporean strain [22]. The Malaysian outbreak, reported in 2020, occurred in marine cage-cultured adult Asian seabass in September 2019 and was marked by significant mortality, further cementing the virus’s pan-regional presence [8]. Phylogenetic analysis based on the major capsid protein (MCP) gene confirmed the high similarity between SDDV isolates from Malaysia, Thailand, and Singapore, suggesting a common source or rapid, unimpeded circulation of a single, highly fit viral lineage [8]. More recently, the geographic boundaries of SDDV have expanded dramatically northward into East Asia. The virus was identified as the causative agent of a novel ascitic disease in farmed yellowfin seabream (Acanthopagrus latus) in Zhuhai, Guangdong Province, Southern China, marking the first scientific confirmation in this species and this country [12]. This finding is particularly alarming as it demonstrates host jumps and adaptation to temperate or subtropical Chinese aquaculture systems. Subsequent molecular surveillance from 2021 to 2023 in the same region confirmed the persistent presence of SDDV in yellowfin seabream ponds, with 31 out of 73 batches of naturally diseased fish testing positive by conventional PCR [7]. Whole-genome sequencing of the Chinese isolate (SDDV-ZH918/23) revealed a genome of 131,741 bp with high collinearity to other Asian isolates, but also uncovered a 152 bp deletion mutant in two separate isolates, the first documentation of a naturally occurring genomic deletion in SDDV, hinting at ongoing microevolution and potential adaptation [7]. The detection of SDDV in clinically healthy fish from farms with no prior history of disease, using highly sensitive qPCR and RPA-Cas12a assays, strongly suggests that subclinical carriers are widespread and act as cryptic reservoirs for viral maintenance and dissemination [19, 20]. This subclinical carriage, combined with the ability to detect the virus in non-destructive samples like blood and even in ectoparasites (Copepoda: Lernanthropus sp. and Monogenea: Diplectanum sp.), underscores the complexity of its transmission ecology and the inadequacy of relying solely on visual signs for disease surveillance [21]. The virus has not yet been reported in the Americas, Europe, Africa, or Oceania, but the extensive global trade in live aquatic animals and frozen products from endemic regions poses a significant risk of incursion. Critically, the closely related ISKNV has been demonstrated to remain infectious in frozen fillets stored at -20°C for seven days, a finding with profound implications for risk assessment of international commodity trade; it is highly plausible that SDDV shares similar cryo-resistance properties [24].

Host Range and Species Susceptibility

The host range of SDDV, while narrower than that of ISKNV which infects over 100 fish species, is expanding at an alarming rate and is of critical concern [16]. The primary and most economically important host remains the Asian seabass (barramundi), where infection results in the archetypal disease presentation of scale drop, cutaneous hemorrhage, lethargy, and high mortality [14]. The virus has been associated with mortality in both juvenile and adult fish in marine net-pens in Singapore, Thailand, Malaysia, and Indonesia [8, 14, 22]. A critical epidemiological finding that alters risk perception for the industry is the confirmation of SDDV infection in freshwater-reared barramundi in Thailand. This case involved a concurrent infection with Flavobacterium columnare, leading to severe gill necrosis and atypical “saddleback” lesions, demonstrating that SDDV pathogenesis can be modified by polymicrobial interactions and that freshwater environments do not constitute a biosecure barrier [9]. This event also illustrated that SDDV should be considered in the differential diagnosis of mortality events in freshwater systems, not just marine ones.

The most significant host range expansion documented to date is the adaptation of SDDV to yellowfin seabream (Acanthopagrus latus), a species of major economic importance in China and other parts of East Asia [12]. Remarkably, the clinical presentation in this novel host differs drastically from that in Asian seabass. Yellowfin seabream do not exhibit scale drop; instead, the dominant clinical sign is severe ascites (abdominal fluid accumulation), leading to the description “yellowfin seabream ascitic disease” (YFSBAD) [7, 12]. This suggests that the pathological effects of SDDV, and potentially the tissue tropism, are highly host-dependent. Experimental challenge confirmed that the SDDV isolate ZH-06/20 caused 100% mortality in juvenile yellowfin seabream via an ascitic disease pathway, definitively fulfilling Koch’s postulates for this new host-pathogen pairing [12]. This phenomenon of serovar- or strain-specific pathology in different hosts is reminiscent of other aquatic pathogens and necessitates a re-evaluation of diagnostic criteria in different aquaculture settings. Beyond these two major hosts, the potential host range is likely much broader. Epidemiological studies on the Megalocytivirus genus have shown that megalocytiviruses can infect a vast range of freshwater, brackish, and marine species, including gourami, tilapia, mandarin fish, and various ornamental species [15, 16, 23]. Given the conserved entry mechanism of SDDV, which utilizes transferrin receptor 1 (TfR1), a highly conserved transmembrane protein across diverse teleosts, the theoretical host range is vast [1]. Indeed, the demonstration that mandarin fish (Siniperca chuatsi) are permissive to SDDV infection and that mfTfR1 functions as its entry receptor suggests that any fish species expressing a structurally compatible TfR1 could be a potential target [1, 5]. This underscores the urgent need for active surveillance efforts beyond the known susceptible hosts to proactively identify future host jumps.

Transmission Dynamics and Co-infection Patterns

The mechanisms of SDDV transmission in aquaculture systems are multifactorial and poorly characterized, yet essential for developing containment strategies. Horizontal transmission via the waterborne route is considered the primary mode of spread within a farm. The virus is shed in large quantities from the skin lesions, feces, and ascitic fluid of infected fish, creating a high-concentration pathogen load in the surrounding water column [12, 14]. The detection of SDDV DNA in blood, mucus, and fin tissue confirms that any breach of the integument (e.g., handling, netting, or ectoparasite damage) can facilitate viral release and entry [21]. The high-density stocking practices typical of commercial aquaculture dramatically accelerate the effective contact rate and infectious dose per fish. The role of ectoparasites as mechanical vectors is a compelling new dimension to SDDV epidemiology. The detection of SDDV-specific PCR amplicons from copepods (Lernanthropus sp.) and monogeneans (Diplectanum sp.) collected from infected Asian seabass suggests that these parasites can harbor the virus on their surface or within their gut [21]. Whether they act as true biological vectors (where the virus replicates within the parasite) or simple mechanical carriers (fomites) remains unknown, but either scenario enhances viral dissemination within and between fish populations.

A hallmark of SDDV epidemiology in the field is its frequent occurrence as part of polymicrobial infections, which can dramatically worsen disease outcome. Co-infection of SDDV with Lates calcarifer herpes virus (LCHV) in commercial farming conditions in Singapore was found to be highly prevalent in both clinically sick and apparently healthy fish [11]. However, RNA sequencing of sick vs. healthy fish revealed that high SDDV loads were strongly associated with the “sick” phenotype and with severe histopathological lesions, particularly necrotizing splenitis and nephritis [11]. Critically, gene expression analysis from these co-infected fish showed that immune-relevant pathways, including phagosome, cytokine-cytokine receptor interaction, and Toll-like receptor signaling, were massively and uniformly downregulated in sick fish [11]. This suggests that the combined viral burden of SDDV and LCHV induces a state of profound systemic immunosuppression, facilitating uncontrolled viral replication and fulminant tissue necrosis. Similarly, the aforementioned co-infection of SDDV with F. columnare in freshwater barramundi led to a clinical presentation that was more severe and included atypical pathologies (gill necrosis) that would not be expected from SDDV alone [9]. The epidemiology of SDDV, therefore, cannot be understood in isolation; it operates within a complex pathobiome where the presence of other pathogens can significantly modify disease expression, severity, and transmission potential. This has major implications for vaccine development, as a vaccine effective against SDDV alone might offer only partial protection in a polymicrobial disease complex.

Molecular Epidemiology: Receptor Tropism and Virulence Determinants

A revolution in understanding SDDV epidemiology has come from the elucidation of its cellular entry mechanism. The identification of mandarin fish transferrin receptor 1 (mfTfR1) as the primary entry receptor for SDDV provides a molecular basis for host specificity and pathogenesis [1]. The study by Chen et al. demonstrated that SDDV directly interacts with mfTfR1 via its major capsid protein (MCP), and that the helical domain of the receptor is the critical docking site [1]. This interaction triggers clathrin-mediated endocytosis, a pathway exploited by many mammalian viruses but novel for a megalocytivirus in fish [1]. The expression level of TfR1 is not static; SDDV infection itself upregulates TfR1 expression, effectively increasing the number of entry portals available for viral invasion in a positive feedback loop [1, 5, 6]. This is linked to the virus’s ability to induce ferroptosis, an iron-dependent, non-apoptotic form of cell death. SDDV infection triggers iron overload, lipid peroxidation, and glutathione depletion, hallmarks of ferroptosis, by upregulating TfR1 to disrupt cellular iron homeostasis [5, 6]. The virus then exploits this pro-death environment to facilitate its own replication and egress. Inhibiting ferroptosis, either with the specific inhibitor Ferrostatin-1 or by iron chelation therapy, significantly reduces SDDV replication in cell culture and improves survival in mandarin fish [2, 5]. Furthermore, vitamin C has been shown to potently inhibit SDDV infectivity by targeting the Nrf2 pathway, reducing oxidative stress and blocking ferroptosis [2]. This molecular understanding provides powerful new tools for epidemiology. The conservation of TfR1 across teleosts implies a broad potential host range, while subtle interspecies variations in the helical domain of TfR1 could dictate the binding efficiency of the SDDV MCP and thus correlate with differential susceptibility. Genomic epidemiology studies can now leverage this knowledge by screening SDDV isolates for mutations in the MCP gene that might alter receptor affinity or specificity, potentially explaining host jumps like the one to yellowfin seabream [7]. The discovery of a 152 bp deletion mutant in Chinese SDDV isolates further emphasizes that the viral genome is actively evolving, and such deletions could be linked to changes in virulence, host range, or transmissibility [7].

Diagnostic Surveillance as the Backbone of Epidemiology

Effective epidemiological investigation of SDDV is entirely dependent on the availability of robust, sensitive, and field-deployable diagnostic tools. The reliance on clinical signs for diagnosis is patently insufficient given the existence of asymptomatic carriers, the variable clinical presentation in different hosts (ascites vs. scale drop), and the confounding effects of co-infections. A suite of molecular tools has been developed to fill this gap, enabling large-scale surveillance. The gold

Clinical Signs and Pathological Manifestations of SDDV Infection in Teleosts

Scale drop disease virus (SDDV), a distinct member of the genus Megalocytivirus within the family Iridoviridae, represents an emergent and increasingly significant threat to global teleost aquaculture [1, 4, 16]. Since its first formal description in 2015, SDDV has been identified as the etiological agent of a devastating syndrome characterized by profound dermal and systemic pathology, predominantly impacting Asian seabass (Lates calcarifer) and, more recently, yellowfin seabream (Acanthopagrus latus) [12, 14]. The clinical presentation and underlying pathological manifestations of SDDV infection are complex, reflecting a systemic viral disease with a particular tropism for hematopoietic tissues, endothelial cells, and the integumentary system. Understanding these manifestations in exhaustive detail is paramount for accurate diagnosis, effective surveillance, and the development of targeted therapeutic interventions.

Gross Clinical Signs: A Spectrum of Integumentary and Systemic Dysfunction

The most conspicuous and pathognomonic clinical sign of SDDV infection, from which the virus derives its name, is the extensive loss of scales, or “scale drop,” observed in affected Asian seabass. This is not a subtle finding; infected fish display large, denuded patches of skin where scales have been shed, leaving the underlying epidermis exposed and vulnerable [8, 14]. This profound scale loss is often accompanied by severe cutaneous hemorrhages and erythema, particularly on the ventral aspect of the body, presenting as a vivid reddening of the skin [8, 9, 23]. In many cases, the dorsal body surface becomes markedly darkened or discolored, contrasting sharply with the hemorrhagic ventral regions [8]. Affected fish are consistently reported to be lethargic, exhibiting a reduced responsiveness to stimuli and a tendency to swim near the water surface, a behavioral change indicative of profound physiological distress [8]. In freshwater-reared barramundi, concurrent infections with Flavobacterium columnare can complicate the clinical picture, leading to additional signs such as severe gill necrosis and characteristic “saddleback” lesions at the base of the dorsal fin, highlighting the potential for polymicrobial disease presentations [9].

Critically, the clinical presentation of SDDV is not uniform across all teleost hosts. In a remarkable divergence from the syndrome observed in Asian seabass, SDDV infection in farmed yellowfin seabream (Acanthopagrus latus) in Southern China presents a dramatically different clinical picture. Instead of scale loss, the predominant clinical sign is severe ascites, manifesting as a pronounced, fluid-filled distension of the abdominal cavity, leading to what has been described as yellowfin seabream ascitic diseases (YFSBAD) [7, 12]. This starkly different phenotype underscores the influence of host species on viral pathogenesis and clinical expression. In both species, however, the disease is associated with high morbidity and mortality, with outbreaks in juvenile yellowfin seabream resulting in 100% mortality in controlled challenges, and significant die-offs in market-sized Asian seabass in marine cage systems [7, 8, 12, 22].

Systemic Pathology and Gross Lesions: The Silent Spread

Post-mortem examination of SDDV-infected fish reveals a spectrum of gross pathological changes across multiple organ systems, confirming the systemic nature of the infection. The spleen is frequently identified as the most severely affected organ. In clinically sick barramundi, the spleen presents with severe multifocal and coalescing necrosis, rendering the organ friable and discolored [11]. The kidneys are also consistently affected, displaying moderate to severe inflammation, glomerular necrosis, and tubular degeneration, often accompanied by blood congestion [8, 11]. The liver exhibits notable pathology, ranging from moderate to severe abnormal fat accumulation (hepatic lipidosis) in fish under commercial farming conditions to vacuolated hepatocyte cytoplasm and, in some cases, the presence of basophilic hypertrophied cells and intracytoplasmic inclusion bodies [8, 11, 14]. Splenomegaly and renomegaly (enlargement of the spleen and kidney) are common findings, reflecting the intense inflammatory and necrotic processes within these hematopoietic organs [10, 16]. Congestion and hemorrhage are frequently observed in the spleen and brain, indicating a breakdown of vascular integrity that contributes to the overall pathology [8]. The severity of these lesions is strongly correlated with SDDV viral loads, with sick fish harboring significantly higher copy numbers than their apparently healthy counterparts, confirming the direct role of the virus in inducing this profound tissue damage [11]. In yellowfin seabream, the collection of ascitic fluid is a key gross finding, and this fluid itself is infectious, serving as a source for viral isolation in cell culture [12].

Histopathological Hallmarks: A Cellular Portrait of Viral Cytopathology

At the microscopic level, SDDV induces a characteristic suite of histopathological changes that are critical for definitive diagnosis. The hallmark lesion, common to megalocytiviruses, is the presence of basophilic hypertrophied cells or megalocytes. These are grossly enlarged cells, often found in the spleen, kidney, liver, and heart, whose cytoplasm is packed with large, basophilic, intracytoplasmic inclusion bodies representing viral assembly sites or “virus factories” [8, 9, 14, 16, 17]. In Asian seabass, these inclusion bodies are occasionally observed in the cytoplasm of infected cells, while in yellowfin seabream and other species, they represent a more consistent diagnostic feature [12, 17]. Transmission electron microscopy (TEM) confirms that these cytoplasmic regions contain numerous icosahedral viral particles, approximately 140-150 nm in diameter, arranged in paracrystalline arrays, a morphology characteristic of the Iridoviridae family [8, 12, 14, 15].

Beyond the formation of inclusion bodies, SDDV infection drives extensive necrosis of infected tissues. In the spleen and kidney, the primary hematopoietic organs, there is widespread necrosis of the parenchyma, leading to the destruction of erythrocytes, lymphocytes, and other immune cells [11, 16, 23]. This splenic and renal necrosis is a primary driver of the immunosuppression and anemia observed in advanced disease. In the liver, histopathology reveals extensive necrosis of hepatocytes, with fibrous serous exudation and inflammatory cell infiltration [8, 28]. Gill pathology includes necrosis of the gill filaments, which can be exacerbated by secondary bacterial infections [9]. In the brain, blood congestion and hemorrhage are noted, and while viral particles have been detected in neural tissues, the cellular pathology is less pronounced than in the hematopoietic organs [8, 20]. The severe lymphocidal effects, including atrophy and depletion of lymphocytes in the spleen and kidney, are reminiscent of the pathology induced by other highly virulent viruses, such as velogenic Newcastle disease virus in poultry, though the underlying mechanisms differ [29].

Molecular and Cellular Pathophysiology: Ferroptosis and the Iron Nexus

Recent groundbreaking research has elucidated the molecular underpinnings of SDDV-induced pathology, revealing that the virus hijacks fundamental cellular processes to drive pathogenesis. A critical pathway identified is the induction of ferroptosis, a non-apoptotic, iron-dependent form of regulated cell death characterized by the lethal accumulation of lipid peroxides [2, 5, 6]. SDDV infection triggers a cascade of events that culminate in ferroptosis both in vivo and in vitro. Infected cells display hallmark features of ferroptosis, including profound iron overload, massive accumulation of lipid peroxides and reactive oxygen species (ROS), depletion of the critical antioxidant glutathione (GSH), and downregulation of glutathione peroxidase 4 (GPX4), the key enzyme responsible for detoxifying lipid peroxides [5, 6]. Mitochondria in SDDV-infected MFF-1 cells exhibit shrinkage and increased membrane density, a classic ultrastructural sign of ferroptosis [5].

The mechanism by which SDDV induces ferroptosis is intricately linked to the viral entry receptor. SDDV utilizes host transferrin receptor 1 (TfR1) to gain entry into cells via clathrin-mediated endocytosis [1]. Critically, SDDV infection was shown to upregulate the expression of TfR1, thereby disrupting cellular iron homeostasis [5, 6]. This increased iron import directly fuels the Fenton reaction, generating highly reactive hydroxyl radicals that drive lipid peroxidation and initiate ferroptosis. The virus thus not only uses TfR1 for entry but also manipulates the host's iron metabolism to create a permissive environment for its own replication by triggering this pro-inflammatory, destructive form of cell death. The pathological significance is profound: administering the potent ferroptosis inhibitor Ferrostatin-1 significantly attenuates SDDV replication in vitro and, importantly, improves the survival of mandarin fish (Siniperca chuatsi) upon SDDV challenge, directly linking this cell death pathway to lethality [5, 6]. Furthermore, treatment with the iron chelator deferoxamine also mitigates ferroptosis and reduces mortality, confirming the iron-dependent nature of this pathway [5, 6]. This discovery positions ferroptosis not merely as a consequence of infection but as a central pathogenic mechanism exploited by SDDV to facilitate its spread and cause systemic damage.

Immune Pathophysiology and Cellular Landscape

The massive tissue necrosis driven by ferroptosis and direct viral cytopathicity precipitates a severe systemic inflammatory response. Single-cell transcriptome profiling of convalescent Asian seabass has revealed a complex immune cell landscape, with elevated proportions of B cells, granulocytes, and T cells, including CD4+ T helper and CD8+ cytotoxic T cells, indicating an adaptive immune response is mounted [25]. However, in clinically sick fish, transcriptomic analyses of the spleen and kidney paint a picture of immune failure. RNA sequencing of sick vs. healthy barramundi reveals the downregulation of numerous key immune-relevant pathways, including phagosome, cytokine-cytokine receptor interaction, Toll-like receptor (TLR), and complement pathways [11]. This widespread suppression of both innate and adaptive immune genes in the face of high viral loads suggests that SDDV actively evades or subverts the host immune response, or that the severe tissue necrosis itself leads to a functional collapse of the immune system, preventing the fish from controlling the infection. This immune dysregulation, characterized by a failure to upregulate critical antiviral mediators, is a key pathological feature of end-stage disease.

Tissue Tropism and Subclinical Carriage

SDDV exhibits a broad tissue tropism, confirming its systemic nature. Quantitative PCR (qPCR) assays have detected viral DNA in a wide array of tissues, with high viral loads (ranging from 8.0 × 10² to 6.8 × 10⁴ copies per 200 ng DNA) found in the liver, spleen, kidney, brain, eyes, gills, fin, heart, intestine, muscle, and skin [20, 26, 27]. The virus is also readily detectable in blood, mucus, and ascitic fluid, enabling non-destructive diagnostic approaches [12, 21]. A critical and challenging epidemiological aspect of SDDV is the existence of subclinical carriers. Using highly sensitive qPCR assays, the virus has been detected in a significant proportion (58%) of overtly healthy fish on farms where SDDV had not previously been identified [20, 27]. These subclinical carriers, with low-level persistent infections, likely serve as a cryptic reservoir for viral maintenance and transmission, complicating control efforts and highlighting the need for sensitive surveillance methods like droplet digital PCR (ddPCR) and field-deployable CRISPR-based diagnostics [19, 26, 27]. The detection of SDDV in ectoparasites like Lernanthropus sp. and Diplectanum sp. from infected fish further suggests potential mechanical vectors for transmission, adding another layer of complexity to the epizootiology of this disease [21].

Diagnostic Strategies for Scale Drop Disease Virus: Molecular, Serological, and Histopathological Approaches

The accurate and timely diagnosis of Scale Drop Disease Virus (SDDV) is paramount for effective disease management, surveillance, and the implementation of control strategies in affected aquaculture systems. As a distinct member of the genus Megalocytivirus within the family Iridoviridae, SDDV presents unique diagnostic challenges that necessitate a multi-faceted approach integrating molecular, serological, and histopathological techniques [1, 16]. The World Organisation for Animal Health (WOAH) recognizes the significant economic and ecological threat posed by megalocytiviruses, underscoring the need for robust, validated diagnostic protocols to monitor and contain outbreaks [16, 24]. Given the virus’s emergence as a pathogen of global concern in teleost aquaculture, including Asian seabass (Lates calcarifer), mandarin fish (Siniperca chuatsi), and yellowfin seabream (Acanthopagrus latus), a comprehensive diagnostic strategy must be both highly sensitive for detecting subclinical carriers and specific enough to differentiate SDDV from other closely related iridoviruses [7, 12, 14].

Molecular Detection Methods: The Cornerstone of SDDV Diagnosis

Nucleic acid amplification techniques (NAATs) form the bedrock of SDDV diagnostics, owing to their superior sensitivity and specificity. The initial characterization and subsequent field surveillance of SDDV have been profoundly advanced by polymerase chain reaction (PCR)-based methods. Conventional PCR and semi-nested PCR (snPCR) assays, often targeting the highly conserved major capsid protein (MCP) gene or the viral adenosine triphosphatase (ATPase) gene, were among the first molecular tools developed [8, 32]. The snPCR assay, for instance, demonstrated a 100-fold higher sensitivity compared to single-step PCR, detecting as few as 100 viral copies per microliter of template, and proved invaluable for confirming SDDV in clinically diseased fish across Southeast Asia [32]. This method was crucial in the first confirmation of SDDV in Malaysian farmed Asian seabass, where histopathological findings of basophilic hypertrophied cells and intracytoplasmic inclusion bodies were corroborated by PCR and sequence analysis [8].

The advent of real-time quantitative PCR (qPCR) has significantly enhanced diagnostic capabilities, enabling not only detection but also precise quantification of viral loads. A highly sensitive SYBR Green-based qPCR assay targeting a 135-bp fragment of the SDDV ATPase gene was developed, achieving a limit of detection of just two viral copies per reaction [20, 27]. This assay exhibited no cross-reactivity with a panel of 12 other common aquatic viral and bacterial pathogens, confirming its high specificity [27]. Critically, this qPCR assay demonstrated a substantially higher detection rate in subclinically infected fish, those harboring low-level, chronic infections without overt clinical signs, compared to snPCR, making it an indispensable tool for active surveillance and early screening programs [20, 27]. The assay revealed that SDDV DNA loads varied dramatically across tissue types, from 8.0 × 10² to 6.8 × 10⁴ copies per 200 ng of total DNA, reinforcing the recommendation to test multiple tissues (e.g., spleen, kidney, liver, and blood) for reliable diagnosis, especially in carrier fish [20].

For applications requiring absolute quantification without the need for standard curves, droplet digital PCR (ddPCR) has emerged as a powerful alternative. A ddPCR method developed for SDDV detection in yellowfin seabream demonstrated exceptional sensitivity, with a limit of detection of 1.4–1.7 copies/μL, and outstanding reproducibility (relative standard deviation of 0.77%) [26]. This high precision makes ddPCR particularly suited for detecting low-level infections, monitoring viral clearance in therapeutic trials, and serving as a reference standard for calibrating other molecular assays [26]. The tissue distribution of SDDV, as determined by ddPCR, confirmed systemic infection across multiple organs including liver, spleen, kidney, heart, intestine, brain, blood, muscle, and skin, with results consistent with clinical findings [26].

Isothermal and Point-of-Care Molecular Diagnostics

The deployment of molecular diagnostics in field settings, particularly in resource-limited aquaculture farms, has been a major hurdle. Conventional PCR requires expensive thermal cycling equipment and skilled personnel. To address this, several isothermal amplification platforms have been developed for SDDV, offering rapid, cost-effective, and field-deployable solutions.

Loop-mediated isothermal amplification (LAMP) has been adapted for SDDV detection with a colorimetric readout using xylenol orange (XO), allowing for naked-eye visualization of positive reactions [31]. This LAMP-XO assay demonstrated a sensitivity of 100 picograms of total DNA extracted from infected fish tissue and, when coupled with a rapid DNA extraction method, provided a simple workflow suitable for on-site application [31]. Similarly, a cross-priming amplification (CPA) assay incorporating uracil-DNA glycosylase (UDG) to prevent carryover contamination was developed targeting the MCP gene [30]. This UCPA method, combined with hydroxynaphthol blue (HNB) or lateral flow dipstick (LFD) readouts, achieved a sensitivity of 100 viral copies/µL and 10 pg of total DNA, outperforming conventional PCR by tenfold [30]. Notably, the UCPA-LFD format provides a user-friendly, instrument-free detection system ideal for farm-level diagnostics [30].

Perhaps the most significant advancement in field-deployable molecular diagnostics for SDDV is the development of a CRISPR-Cas12a-based platform coupled with recombinase polymerase amplification (RPA) [19]. This RPA-Cas12a assay targets the SDDV ATPase gene and achieves a detection limit of 40 copies per reaction, with high specificity for SDDV over other pathogens [19]. The entire workflow operates at a constant 37°C and can be completed within one hour, eliminating the need for thermal cyclers. The assay provides results via fluorescence or lateral flow readouts, enabling straightforward interpretation. Critically, when evaluated with field samples, the RPA-Cas12a assay was more sensitive than semi-nested PCR and could detect SDDV in asymptomatic fish, highlighting its potential for proactive surveillance at the point of need [19].

Metagenomic sequencing has also proven to be a powerful tool for the discovery, genomic characterization, and epidemiological tracking of SDDV. This approach has enabled the recovery of complete SDDV genomes directly from infected fish tissues, revealing genomic organization, repeat sequences, and mutations that may serve as biomarkers for strain differentiation [4, 10]. For instance, analysis of the SDDV TH2019 genome identified a unique 7,695-bp genomic region containing six hypothetical protein-encoding genes, illustrating the power of metagenomics to uncover genomic diversity that might be missed by targeted PCR approaches [10]. This method has been instrumental in confirming the presence of SDDV in new host species, such as yellowfin seabream, and in detecting co-infections with other pathogens like Lates calcarifer herpes virus (LCHV) [11, 12].

Non-Destructive Sampling and Sample Matrices

A critical consideration for surveillance and broodstock management is the feasibility of non-destructive sampling. Research has demonstrated that SDDV DNA is detectable in blood, mucus, and fin clips of infected Asian seabass, with blood proving to be the most reliable non-destructive sample source [21]. Blood samples yielded 100% SDDV-positive results from both clinically sick and subclinically infected fish, providing a sterile sample that minimizes the risk of contamination [21]. This finding has profound implications for routine health monitoring, as it avoids the need to sacrifice valuable broodstock or experimental fish. Furthermore, the detection of SDDV in ectoparasites such as Lernanthropus sp. and Diplectanum sp. collected from infected fish suggests that these parasites could serve as sentinel samplers for environmental surveillance of the virus [21].

Histopathological and Ultrastructural Examination

Histopathology remains a cornerstone for initial disease screening and provides invaluable insights into the pathogenesis of SDDV infection. The hallmark histopathological lesion of SDDV, and indeed all megalocytiviruses, is the presence of markedly enlarged, basophilic hypertrophied cells, often referred to as “megalocytes,” in the cytoplasm of infected cells [8, 17]. These inclusions are most consistently observed in the spleen, kidney, liver, and gills, reflecting the systemic nature of the infection [8, 9, 11]. In a study of Malaysian farmed Asian seabass, histopathological examination of multiple organs revealed vacuolated cytoplasm of hepatocytes, necrotic kidney tubules, atrophied glomeruli, and blood congestion with hemorrhage in the spleen and brain [8]. The presence of intracytoplasmic inclusion bodies, particularly in the liver, was a key indicator of viral etiology [8].

The severity and distribution of histopathological changes correlate strongly with disease progression and viral load. In co-infection scenarios with Flavobacterium columnare, histopathology revealed extensive necrosis of skeletal muscle and gill filaments alongside SDDV-specific basophilic inclusion bodies and megalocytosis in muscle, gill, liver, and kidney [9]. During severe outbreaks in barramundi under commercial farming conditions, the spleen was the most severely affected organ, presenting severe multifocal and coalescing necrosis, while the kidneys exhibited moderate to severe inflammation and glomerular necrosis [11]. These histopathological findings were strongly associated with high SDDV viral loads, as determined by qPCR, confirming the primary role of SDDV in the observed pathology [11].

Interestingly, SDDV infection in yellowfin seabream presents a distinct clinical and pathological picture compared to that in Asian seabass. Instead of the characteristic scale loss, the predominant symptom is severe ascites [12]. Histopathological examination of these cases is crucial for differential diagnosis, particularly to distinguish SDDV from other viral and bacterial causes of ascites. The presence of megalocytes in the spleen and kidney of ascitic fish, together with molecular confirmation, was pivotal in identifying SDDV as the etiological agent of yellowfin seabream ascites disease [12].

At the ultrastructural level, transmission electron microscopy (TEM) provides definitive evidence of SDDV infection by revealing the presence of non-enveloped, icosahedral viral particles approximately 140–200 nm in diameter within the cytoplasm of infected cells [8, 12, 14]. These virions, characteristic of the Iridoviridae family, can be observed either free in the cytoplasm or within paracrystalline arrays [14]. TEM is particularly valuable for distinguishing SDDV from other viral pathogens that may produce similar histopathological lesions, such as infectious spleen and kidney necrosis virus (ISKNV), which also produces polygonal nucleocapsids of a similar size (115–125 nm) [15]. The precise measurement of virion diameter and the observation of the unique hexagonal profile of the icosahedral capsid are critical for accurate differentiation [14].

Serological Approaches and Their Limitations

While molecular and histopathological methods are well-established for SDDV detection, serological approaches are still in their infancy but hold significant promise for large-scale surveillance. The development of serological assays, such as enzyme-linked immunosorbent assays (ELISA), has been driven by the need for high-throughput, cost-effective methods to assess population-level exposure and immunity.

Research on formalin-inactivated SDDV vaccines (SDDV-FIV) has provided a platform for understanding the humoral immune response to SDDV. Vaccination trials in Asian seabass demonstrated that SDDV-FIV elicits both SDDV-specific IgM and total IgM production [3, 13]. A booster dose significantly prolonged the duration of the antibody response, with elevated IgM levels maintained until 70 days post-vaccination [3]. This work establishes the feasibility of detecting anti-SDDV antibodies in fish serum, forming the basis for developing indirect ELISAs to monitor natural exposure or vaccine efficacy. However, the lack of standardized, commercially available SDDV-specific monoclonal antibodies for use in capture or competitive ELISAs remains a significant bottleneck. Research has successfully produced monoclonal antibodies against SDDV, validated by immunofluorescence assays on infected cell cultures, which could be instrumental in developing robust serodiagnostic tools [7].

The challenge of monitoring immunity in large fish populations has led to the exploration of pooled serum testing strategies, an approach successfully applied for other fish viruses like tilapia lake virus (TiLV) [33]. Pooling samples reduces cost and labor, making large-scale surveillance feasible. However, this strategy must be validated for SDDV, as the dilution effect from pooling can reduce sensitivity, particularly when only a small proportion of individuals are seropositive [33]. The development of a validated, WOAH-compliant serological assay for SDDV would be a major step forward for the aquaculture industry, enabling retrospective studies, trade certification, and epidemiological investigations without the need for lethal sampling. Currently, the combination of molecular detection from non-lethal samples like blood and histopathological confirmation from deceased or moribund fish provides the most comprehensive diagnostic picture [21].

The integrated application of these diagnostic strategies, from highly sensitive molecular assays for early detection, to isothermal platforms for field use, to histopathological and ultrastructural characterization for confirmatory diagnosis and pathogenesis studies, is essential for combating SDDV. The continuous refinement of these tools, informed by genomic insights into the receptor binding and entry mechanisms of SDDV through TfR1, will be crucial for the sustainable management of this devastating aquaculture pathogen [1, 5].

Immunological Responses and Vaccine Development Strategies Against SDDV

The host immune response to scale drop disease virus (SDDV) infection is a complex, multifaceted interplay between viral pathogenesis and the teleost immune system, encompassing both innate and adaptive arms. Understanding these immunological dynamics is paramount for the rational design of effective prophylactic and therapeutic interventions. Concurrently, the development of robust vaccine platforms against SDDV has emerged as a critical priority for sustaining global aquaculture, particularly for Asian seabass (Lates calcarifer) and other susceptible species. This section provides an exhaustive analysis of the immunological responses elicited by SDDV and the current landscape of vaccine development strategies, drawing upon the most recent and foundational research in the field.

Innate Immune Responses and the Induction of Ferroptosis

The initial host-virus interaction is governed by the innate immune system, which serves as the first line of defense. However, SDDV has evolved sophisticated mechanisms to subvert these defenses, often exploiting host cellular processes to its advantage. A landmark discovery in this context is the identification of mandarin fish transferrin receptor 1 (mfTfR1) as a critical entry receptor for SDDV [1]. This receptor, a type II transmembrane glycoprotein, directly interacts with the viral major capsid protein (MCP), facilitating clathrin-mediated endocytosis [1]. This entry mechanism is not merely a passive event; it is an active process where SDDV infection upregulates TfR1 expression, thereby disrupting cellular iron homeostasis [5, 6]. The resulting iron overload is a key trigger for ferroptosis, a non-apoptotic, iron-dependent form of regulated cell death characterized by the accumulation of lipid peroxides, reactive oxygen species (ROS), and glutathione depletion [2, 5, 6]. This is a critical immunological paradox: while ferroptosis is a host cell death mechanism that can limit pathogen replication, SDDV actively induces it to facilitate its own infection and dissemination [5, 6]. The virus’s ability to upregulate TfR1, leading to iron dysregulation and subsequent ferroptosis, represents a novel pathogenic strategy among megalocytiviruses [5, 6]. The downstream consequences of this process include the upregulation of genes related to iron, oxidative stress, and lipid metabolism, creating a cellular environment conducive to viral replication [2]. This intricate link between viral entry, iron metabolism, and cell death provides a rich target for therapeutic intervention. For instance, the ferroptosis inhibitor Ferrostatin-1 and iron chelators have been shown to significantly attenuate SDDV replication in vitro and improve survival in mandarin fish challenge models, underscoring the therapeutic potential of targeting this pathway [5, 6].

Beyond ferroptosis, SDDV infection elicits a broader innate response. Studies using single-cell RNA sequencing (scRNA-seq) in convalescent Asian seabass have revealed a dynamic shift in the immune cell landscape. Infection leads to elevated proportions of granulocytes, which are key effectors of the innate immune response, alongside B and T cells [25]. The analysis of macrophage populations in these studies indicates an upregulation of genes linked to inflammatory processes, suggesting a robust, albeit potentially dysregulated, innate inflammatory response [25]. Furthermore, transcriptomic analyses of barramundi co-infected with SDDV and Lates calcarifer herpesvirus (LCHV) under commercial farming conditions have shown that clinically sick fish exhibit a significant downregulation of key innate immune pathways, including those involving toll-like receptors (TLRs), the complement system, and cytokine-cytokine receptor interactions [11]. This suggests that in severe, advanced stages of disease, SDDV may actively suppress or evade the innate immune response, leading to a failure of the host to control the infection. The downregulation of these pathways, coupled with the induction of ferroptosis, paints a picture of a virus that manipulates host cell death and immune signaling to establish a productive infection.

Adaptive Humoral and Cellular Immune Responses

The adaptive immune system, comprising both humoral (antibody-mediated) and cellular (T cell-mediated) components, is essential for long-term protection and immunological memory against SDDV. The humoral response, primarily involving immunoglobulin M (IgM), is a key correlate of vaccine-induced protection. Vaccination with a formalin-inactivated SDDV vaccine (SDDV-FIV) has been shown to trigger robust production of both SDDV-specific IgM and total IgM in Asian seabass [3, 13]. A single dose of the vaccine elicited detectable SDDV-specific IgM levels that persisted until 28 days post-vaccination (dpv), while a booster dose significantly extended this response, maintaining elevated antibody levels until at least 70 dpv [3]. This prolonged antibody response is crucial for protecting fish during high-risk periods and demonstrates the importance of booster vaccination strategies for sustaining humoral immunity. The protective role of these antibodies was further evidenced by the higher survival rates observed in the booster-vaccinated group following a viral challenge at 42 dpv [3, 13].

The cellular immune response, mediated by T lymphocytes, is equally critical. The scRNA-seq data from convalescent Asian seabass has provided unprecedented detail into the T cell landscape following SDDV exposure. Distinct clusters of T cells expressing cd4-1, cd8a, perforin-1, and il-2rβ have been identified, indicating the presence of both CD4+ T helper (Th) cells and CD8+ cytotoxic T (Tc) cells [25]. The presence of Tc1 cells, which are typically associated with a Th1-like response and are crucial for killing virus-infected cells, was particularly notable [25]. This cellular response is further corroborated by gene expression studies following vaccination. Vaccination with SDDV-FIV significantly upregulated the expression of key immune-related genes in the head kidney and peripheral blood leukocytes (PBLs), including cd4, cd8, ifng (interferon-gamma), and mx (a myxovirus resistance protein induced by type I interferons) [3, 13]. The upregulation of ifng is particularly significant, as it is a master regulator of the Th1-type cellular immune response, promoting the activation of macrophages and cytotoxic T cells. The expression of mx indicates the induction of an antiviral state, a hallmark of a robust interferon-mediated response. Furthermore, the upregulation of ighm (the gene for the secreted form of IgM) and dcst (a dendritic cell-specific transcript) confirms the activation of both humoral and antigen-presenting cell pathways [3]. These findings collectively demonstrate that effective vaccination against SDDV elicits a comprehensive, mixed Th1/Th2-like response, characterized by both potent antibody production and a strong cytotoxic T cell component, which is essential for clearing the virus and establishing long-term memory.

Vaccine Development Strategies: From Inactivated to Novel Platforms

The development of a safe and efficacious vaccine is the cornerstone of long-term SDDV control. The foundational work in this area was established by Groof et al. (2015), who fulfilled Koch’s postulates for SDDV and demonstrated that vaccines prepared from binary ethylenimine (BEI)- and formalin-inactivated virus, as well as an E. coli-produced recombinant major capsid protein (MCP), provided efficacious protection against scale drop disease [14]. This seminal study laid the groundwork for all subsequent vaccine development efforts.

Inactivated Whole-Virus Vaccines: The most extensively studied vaccine platform to date is the formalin-inactivated SDDV vaccine (SDDV-FIV). As detailed above, this vaccine, administered via the intraperitoneal route, has been shown to stimulate both humoral and cellular immune responses, with a prime-boost strategy significantly enhancing protection [3, 13]. The booster group demonstrated a 60% survival rate compared to 20% in the control group following a challenge at 42 dpv [3]. However, the protection waned over time, as the survival benefit was not statistically significant at 70 dpv, although a trend towards improved survival was observed [3]. This highlights a key challenge for inactivated vaccines: the need for multiple doses to maintain protective immunity over the long production cycle of fish. Despite this, the SDDV-FIV remains a strong candidate for commercial development, as it is a well-established technology that is relatively straightforward to produce and has a proven safety profile.

Subunit and Recombinant Protein Vaccines: The use of recombinant proteins, particularly the major capsid protein (MCP), offers a safer and more defined alternative to whole-virus vaccines. The early success of an E. coli-produced MCP vaccine [14] validated this approach. The MCP is an ideal target because it is the most abundant structural protein and is directly involved in host cell entry via interaction with TfR1 [1]. A subunit vaccine based on the MCP or other immunogenic viral proteins could be produced in large quantities without the need for high-containment facilities required for live virus culture. Furthermore, the identification of the helical domain of TfR1 as the crucial docking site for MCP [1] opens the door for structure-based vaccine design, where specific epitopes can be targeted to elicit neutralizing antibodies that block this critical interaction.

Novel and Emerging Platforms: The field is also exploring more advanced vaccine technologies to overcome the limitations of traditional vaccines. The use of virus-like particles (VLPs), which are self-assembling, non-infectious structures that mimic the native virus, is a promising avenue. While not yet reported for SDDV, the successful application of VLP technology against nervous necrosis virus (NNV) in Asian seabass, using Lactococcus lactis as an oral delivery vehicle, provides a powerful proof-of-concept [34]. This approach is particularly attractive for aquaculture because oral vaccination is non-invasive, stress-free, and suitable for mass immunization of large fish populations. The encapsulation of SDDV VLPs in a food-grade bacterium like L. lactis could revolutionize vaccine delivery in the industry. Another cutting-edge strategy involves RNA-based biopesticides, which utilize double-stranded RNA (dsRNA) to trigger RNA interference (RNAi) and silence essential viral genes [36]. While primarily discussed for pest control, this technology could be adapted to target SDDV genes, offering a highly specific and rapidly deployable antiviral strategy. The species-specific targeting and rapid environmental degradation of dsRNA are significant advantages [36].

Adjuvants and Immunostimulants: The efficacy of any vaccine can be significantly enhanced by the inclusion of appropriate adjuvants. Beyond traditional adjuvants, there is growing interest in using natural compounds as immunostimulants. For instance, Vitamin C (VC) has been shown to inhibit SDDV infectivity by activating the Nrf2 pathway, which in turn reduces ferroptosis by decreasing ROS, lipid peroxidation, and iron levels while enhancing glutathione peroxidase 4 (GPX4) expression [2]. This finding suggests that VC could be used as a dietary supplement or an adjuvant in vaccine formulations to boost the host’s antiviral state and reduce the pathological damage caused by ferroptosis. The use of such nutraceuticals represents a cost-effective and safe strategy to complement vaccination efforts.

Challenges and Future Directions

Despite significant progress, several challenges remain in the development and deployment of SDDV vaccines. The waning immunity observed with the inactivated vaccine [3] necessitates the development of vaccines that can induce more durable, long-term memory. This could be achieved through the use of more potent adjuvants, novel delivery systems (e.g., slow-release implants or nanoparticles), or the development of live-attenuated vaccines, which typically induce stronger and longer-lasting immunity than inactivated vaccines. However, the safety of live-attenuated vaccines must be rigorously evaluated to prevent reversion to virulence.

Another critical challenge is the lack of standardized correlates of protection. While SDDV-specific IgM levels are a useful indicator, a clear threshold for protective immunity has not been established. The development of standardized assays, such as virus neutralization tests (VNTs) and ELISpot assays for T cell responses, is essential for evaluating vaccine efficacy in a reproducible manner across different laboratories and studies. The recent development of a droplet digital PCR (ddPCR) method for SDDV detection, with a limit of detection of 1.4–1.7 copies/μL [26], and a CRISPR-Cas12a-based platform for field-deployable detection [19] provide powerful tools for monitoring viral loads in vaccinated and challenged fish, which is crucial for assessing vaccine efficacy.

Finally, the economic and practical constraints of aquaculture must be considered. Injectable vaccines, while effective, are labor-intensive and stressful for the fish. The development of effective immersion and oral vaccines is a top priority for the industry [35]. The use of L. lactis as an oral delivery vehicle for VLPs [34] represents a significant step in this direction. Furthermore, the high cost of vaccine production, particularly for recombinant proteins, can be a barrier to adoption, especially for small-scale farmers. The World Organisation for Animal Health (WOAH) recognizes the threat of megalocytiviruses, and the development of affordable, easy-to-administer vaccines is critical for global food security and the sustainability of the aquaculture sector. Future research should focus on optimizing vaccine formulations, exploring novel delivery platforms, and conducting large-scale field trials to validate the efficacy and cost-effectiveness of these interventions under real-world farming conditions.

Prevention, Control, and Biosecurity Measures for Scale Drop Disease in Aquaculture

The management of Scale Drop Disease (SDD) caused by Scale Drop Disease Virus (SDDV) represents a formidable challenge for global aquaculture, particularly for the culture of Asian seabass (Lates calcarifer) and, more recently, yellowfin seabream (Acanthopagrus latus). As a distinct member of the genus Megalocytivirus within the Iridoviridae family, SDDV exhibits a complex pathogenesis involving host receptor-mediated entry, iron-dependent ferroptosis, and profound immunosuppression [1, 5, 11]. Consequently, effective prevention and control necessitate a multi-layered, integrated strategy that combines rigorous biosecurity protocols, advanced surveillance diagnostics, strategic vaccination, and emerging host-directed therapeutic interventions. The World Organisation for Animal Health (WOAH) recognizes the threat posed by megalocytiviruses, and while SDDV (Megalocytivirus lates 1) is not yet universally listed as notifiable in all member countries, its devastating economic impact and potential for transboundary spread mandate a proactive, science-based approach to containment [16].

### Biosecurity: The First Line of Defense

Biosecurity remains the cornerstone of SDDV prevention, particularly given the absence of commercially available, widely adopted vaccines in many endemic regions. The primary goal is to prevent the introduction of SDDV into naïve populations and to contain its spread within infected facilities. The virus’s ability to establish subclinical infections, as evidenced by the detection of SDDV in clinically healthy fish via highly sensitive qPCR, underscores the critical need for stringent biosecurity measures even in the absence of overt disease [20, 27].

Source Control and Stock Management: The most effective biosecurity measure is the procurement of SDDV-free seed stock. The first evidence of SDDV in Malaysia highlighted how the movement of infected broodstock and fingerlings can introduce the virus to new geographic areas [8]. Similarly, the detection of SDDV in Indonesia was linked to stock originating from Singapore and Batam, leading to recommendations against using broodstock and seed from these regions to prevent nationwide spread [22]. Quarantine protocols for all new introductions are non-negotiable. Incoming fish should be held in isolated facilities for a minimum of 4-6 weeks and subjected to diagnostic testing using highly sensitive methods, such as the SYBR Green qPCR assay targeting the ATPase gene, which can detect as few as 2 viral copies per reaction and identify subclinical carriers that would be missed by conventional PCR [20, 27]. The use of non-destructive sampling, particularly blood, has been validated as an effective method for screening broodstock without sacrificing valuable genetic material, offering a practical advantage for biosecurity programs [21].

Facility Design and Operational Protocols: Physical barriers are essential to prevent horizontal transmission. SDDV, like other iridoviruses, is shed in water and can be transmitted via fomites. Dedicated equipment for each production unit, footbaths with effective disinfectants (e.g., iodophors, chlorine compounds), and strict protocols for personnel movement (e.g., changing boots and coveralls between ponds or tanks) are fundamental. The risk of pathogen spread via contaminated equipment is a well-recognized challenge in aquaculture, and the high viral loads detected in multiple organs, including skin and muscle, of infected fish make carcass disposal a critical control point [20, 27]. Dead and moribund fish must be promptly removed and disposed of through incineration, deep burial with lime, or alkaline hydrolysis to prevent scavenging and waterborne contamination.

Environmental Management and Stress Reduction: Environmental stressors, particularly temperature fluctuations, can significantly impact disease outcome. A study in a recirculating aquaculture system (RAS) in Singapore demonstrated that heat treatment, while intended to reduce pathogens, could induce an overreaction to temperature stress, leading to dysbiosis of the skin microbiota and increased mortality in moribund fish [37]. This finding suggests that while thermal manipulation may have a role, it must be applied with extreme caution and precise control. Conversely, surviving fish after heat treatment showed a reduction in SDDV load, indicating that a carefully managed, moderate heat stress might enhance antiviral responses [37]. Maintaining optimal water quality parameters, stable temperature, adequate dissolved oxygen, and low ammonia, is paramount, as any deviation can compromise the fish’s immune system and increase susceptibility to SDDV. The implementation of IoT-enabled water quality monitoring systems, which can autonomously maintain optimal conditions and provide real-time alerts, represents a significant technological advancement for biosecurity in intensive aquaculture [38, 40].

### Advanced Surveillance and Diagnostics: The Cornerstone of Early Detection

The ability to detect SDDV rapidly and accurately, even in pre-clinical stages, is fundamental to effective control. The development of a suite of diagnostic tools, from laboratory-based qPCR to field-deployable isothermal amplification assays, has revolutionized our capacity for surveillance.

Molecular Diagnostics for High-Throughput Surveillance: The validated semi-nested PCR (snPCR) and the highly sensitive SYBR Green qPCR assays have become the gold standards for laboratory confirmation [20, 27, 32]. The qPCR assay, in particular, is indispensable for active surveillance programs, as it can detect the virus in asymptomatic fish with low viral loads, revealing a hidden reservoir of infection that can perpetuate outbreaks [20, 27]. The recent development of a droplet digital PCR (ddPCR) method for SDDV in yellowfin seabream offers even greater precision and sensitivity, with a limit of detection of 1.4–1.7 copies/μL, making it an ideal tool for quantifying viral loads in research and for validating the efficacy of control measures [26]. For genomic surveillance and epidemiological tracking, metagenomic approaches have proven invaluable for recovering complete SDDV genomes directly from infected tissues, allowing for the characterization of strain diversity, identification of deletion mutants (e.g., the 152 bp deletion found in some Chinese isolates), and tracking of viral evolution [4, 7, 10].

Field-Deployable Point-of-Care Testing: The logistical challenges of transporting samples to centralized laboratories and the time delay inherent in PCR-based methods have spurred the development of rapid, field-deployable diagnostics. A CRISPR-Cas12a-based platform combined with recombinase polymerase amplification (RPA-Cas12a) represents a major breakthrough. This assay can detect as few as 40 copies of the SDDV ATPase gene per reaction at a constant temperature of 37°C within one hour, with results visualized by fluorescence or lateral flow dipsticks [19]. This technology is more sensitive than semi-nested PCR and can detect SDDV in asymptomatic fish, making it a powerful tool for on-farm screening and rapid response during outbreaks [19]. Similarly, loop-mediated isothermal amplification (LAMP) assays, using colorimetric indicators like xylenol orange (LAMP-XO) or hydroxynaphthol blue (UCPA-HNB), provide simple, naked-eye detection of SDDV with sensitivity comparable to or exceeding conventional PCR, and are ideally suited for resource-limited settings [30, 31]. The development of cross-priming amplification (CPA) combined with lateral flow dipsticks (UCPA-LFD) further enhances the user-friendliness and specificity of point-of-care diagnostics [30].

Non-Destructive Sampling for Broodstock Management: The validation of blood, mucus, and fin clips as non-destructive sample sources for SDDV detection is a critical advancement for biosecurity in broodstock facilities [21]. Blood sampling, in particular, yielded 100% positive detection in both sick and clinically healthy fish, providing a reliable method for screening valuable genetic stock without mortality [21]. This approach allows for the establishment and maintenance of SDDV-free broodstock populations, which is the most sustainable long-term strategy for disease prevention.

### Vaccination Strategies: Inducing Protective Immunity

Vaccination is the most promising long-term strategy for sustainable SDDV control. Research has progressed from proof-of-concept studies to detailed evaluations of vaccine efficacy, immune mechanisms, and optimized delivery strategies.

Inactivated Whole-Virus Vaccines: The most extensively characterized vaccine candidate is the formalin-inactivated SDDV vaccine (SDDV-FIV). Early work demonstrated that both BEI- and formalin-inactivated vaccines, as well as a recombinant major capsid protein (MCP) vaccine, could provide efficacious protection against scale drop disease [14]. A comprehensive study on SDDV-FIV in Asian seabass evaluated prime and prime-booster vaccination strategies [3, 13]. The results demonstrated that SDDV-FIV effectively stimulates both humoral and cellular immune responses. The booster vaccination strategy significantly prolonged the duration of SDDV-specific IgM antibodies, maintaining elevated levels until 70 days post-vaccination (dpv), compared to 28 dpv in the single-vaccination group [3]. Crucially, the booster group showed a significantly higher relative percent survival (RPS) of 60% compared to 20% in controls following a challenge at 42 dpv [3]. While protection waned by 70 dpv (42% vs. 25% survival, not statistically significant), the study clearly established the necessity of a booster dose for robust, long-term protection. The immune response was characterized by the upregulation of key immune-related genes in the head kidney and peripheral blood leukocytes, including cd4, cd8, ifng, mx, ighm, and il8, confirming the activation of both adaptive and innate antiviral pathways [3, 13].

Route of Administration and Future Vaccine Platforms: The intraperitoneal (IP) injection route used in these studies is effective but labor-intensive and not practical for large-scale, small-fingerling vaccination. The development of oral or immersion vaccines is a critical research priority. While IP injection of purified virus-like particles (VLPs) has shown promise in other viral systems like NNV, oral delivery remains challenging due to antigen degradation in the gut [34]. However, encapsulation of antigens in Lactococcus lactis has shown potential for oral delivery, and this platform could be adapted for SDDV antigens [34]. The identification of the major capsid protein (MCP) as the viral ligand that interacts with the host entry receptor, transferrin receptor 1 (TfR1), provides a rational basis for designing subunit vaccines based on the MCP protein or its receptor-binding domain [1]. Such a targeted subunit vaccine could potentially induce neutralizing antibodies that block viral entry at the earliest stage of infection.

### Host-Directed Therapeutic and Prophylactic Interventions

A paradigm shift in antiviral strategy involves targeting host cellular pathways that are hijacked by the virus, rather than the virus itself. This approach reduces the likelihood of developing drug-resistant viral mutants. Recent discoveries regarding SDDV pathogenesis have revealed several promising host-directed targets.

Targeting the Viral Entry Receptor (TfR1): The identification of mandarin fish transferrin receptor 1 (mfTfR1) as the essential entry receptor for SDDV is a landmark discovery [1]. SDDV MCP directly binds to the helical domain of mfTfR1, initiating clathrin-mediated endocytosis. This finding opens the door to receptor-targeted therapies. For instance, the TfR1 inhibitor ferristatin II was shown to significantly suppress SDDV entry in vitro [1]. Furthermore, disrupting the specific interaction between MCP and the helical domain of TfR1 reduced viral entry and host mortality in experimental models [1]. While the use of systemic TfR1 inhibitors in food fish requires extensive safety evaluation, this pathway represents a high-value target for the development of entry-blocking peptides or small molecules.

Inhibiting Ferroptosis: SDDV infection triggers ferroptosis, an iron-dependent form of regulated cell death, to facilitate its own replication [5, 6]. Infected cells exhibit iron overload, massive lipid peroxidation, glutathione depletion, and downregulation of glutathione peroxidase 4 (GPX4) [5, 6]. Crucially, treatment with the ferroptosis inhibitor Ferrostatin-1 significantly attenuated SDDV replication in vitro and improved the survival of mandarin fish upon SDDV challenge [5]. Similarly, treatment with an iron chelator (e.g., deferoxamine) mitigated ferroptosis and reduced mortality, confirming the iron-dependent nature of this process [5, 6]. These findings suggest that pharmacological inhibition of ferroptosis could be a viable therapeutic strategy.

Nutritional Immunomodulation with Vitamin C: Vitamin C (VC) has emerged as a potent, safe, and cost-effective prophylactic and therapeutic agent against SDDV. A study demonstrated that VC treatment reduced SDDV-induced mortality in mandarin fish by 37.5% [2]. Mechanistically, VC acts by activating the nuclear factor erythroid 2-related factor 2 (Nrf2) pathway, which promotes the nuclear translocation of Nrf2 and the subsequent expression of antioxidant genes [2]. This counteracts the SDDV-induced oxidative stress, reduces reactive oxygen species (ROS) and lipid peroxidation (LPO), lowers intracellular iron levels, and restores GPX4 expression [2]. The antiviral effect of VC was reversed by the Nrf2 inhibitor ML-385, confirming the pathway’s necessity [2]. This work provides a strong scientific basis for the use of dietary VC supplementation as a practical, low-cost intervention to bolster host resistance against SDDV in aquaculture settings.

### Integrated Control and Future Directions

A holistic, integrated health management plan is essential. This plan must combine the biosecurity measures, advanced diagnostics, vaccination, and host-directed therapies discussed above. The recent detection of SDDV in new hosts like yellowfin seabream, where it presents with ascites rather than scale loss, highlights the virus’s expanding host range and the need for broad surveillance across multiple species [7, 12]. Furthermore, the frequent occurrence of coinfections, such as SDDV with Flavobacterium columnare or Lates calcarifer herpes virus (LCHV), complicates diagnosis and disease outcome, as coinfections can downregulate immune pathways and exacerbate pathology [9, 11]. Therefore, diagnostic protocols should include screening for multiple pathogens.

Looking forward, the application of genomic selection to breed SDDV-resistant lines of Asian seabass, as has been successfully applied to other aquaculture species for disease resistance, represents a powerful, long-term genetic solution [39, 41]. The development of RNA-based biopesticides, using double-stranded RNA to specifically silence essential SDDV genes, also holds promise as a future precision therapy [36]. Ultimately, the sustainable control of SDDV will depend on the translation of these cutting-edge scientific discoveries into practical, affordable, and accessible tools for fish farmers, supported by robust regulatory frameworks and international cooperation to prevent the transboundary spread of this devastating pathogen.

9. Future Perspectives: Antiviral Targets and Therapeutic Interventions for SDDV

The emergence of scale drop disease virus (SDDV) as a distinct member of the genus Megalocytivirus (family Iridoviridae) represents a paradigm-shifting challenge for global teleost aquaculture, particularly for the economically vital Asian seabass (Lates calcarifer) and yellowfin seabream (Acanthopagrus latus) industries [4, 7, 12, 16]. Unlike other megalocytiviruses such as infectious spleen and kidney necrosis virus (ISKNV) and red sea bream iridovirus (RSIV), SDDV, classified as Megalocytivirus lates 1, exhibits a unique pathobiology characterized by profound scale loss, severe ascites, and culminating in mortality rates that can approach 100% in juvenile fish [12, 14, 16]. The World Organisation for Animal Health (WOAH) lists Megalocytivirus pagrus 1 (which includes ISKNV, RSIV, and TRBIV) as a notifiable disease, and while SDDV is not currently separately listed, its expanding geographic footprint across Southeast Asia and into southern China underscores an urgent need for targeted antiviral strategies [8, 16, 22]. Critically, no licensed antiviral drugs exist for any finfish iridovirus, and management relies heavily on biosecurity and nascent vaccine development [3, 43]. Therefore, future therapeutic interventions must pivot from empirical approaches toward a mechanistic understanding of SDDV’s molecular vulnerabilities. Recent breakthroughs in elucidating the virus’s entry machinery, its subversion of host cell death pathways, and its immunomodulatory strategies have illuminated a constellation of high-value antiviral targets. This section provides an exhaustive analysis of these emerging targets and the therapeutic paradigms, ranging from small-molecule inhibitors and host-directed therapies to advanced biologics and RNA-based interventions, that promise to transform SDDV disease management.

9.1 Targeting the TfR1-MCP Axis: A Portal for Intervention

The single most transformative discovery for SDDV antiviral development is the identification of mandarin fish transferrin receptor 1 (mfTfR1) as the primary entry receptor [1]. This finding, which builds upon a well-established paradigm in mammalian virology where TfR1 serves as a receptor for arenaviruses, circoviruses, and parvoviruses, represents the first characterization of a TfR1-dependent entry mechanism for a large DNA virus in teleosts [1]. Chen et al. (2025) demonstrated that the major capsid protein (MCP) of SDDV directly binds to the helical domain of mfTfR1, and that this interaction is critical for viral internalization, which proceeds via Src kinase-mediated tyrosine phosphorylation of TfR1 and subsequent clathrin-mediated endocytosis [1]. This mechanistically rich pathway offers multiple druggable nodes. The TfR1 inhibitor ferristatin II, which blocks iron uptake and receptor recycling, was shown to significantly suppress SDDV entry in vitro [1]. This positions ferristatin II and its analogues as immediate repurposing candidates, though careful toxicological profiling in finfish species is required, as iron homeostasis is critical for erythropoiesis and immune function. Beyond small molecules, the receptor-ligand interface itself, specifically the MCP-helical domain interaction, represents a compelling target for biologics. Monoclonal antibodies targeting the TfR1 helical domain could sterically hinder MCP docking, while recombinant soluble TfR1 decoys could sequester viral particles before cell contact. The latter approach has proven successful in mitigating arenavirus infections in mammalian models and could be adapted for SDDV as an injectable or orally delivered immunotherapeutic, potentially using Lactococcus lactis as a mucosal delivery vehicle [34]. Furthermore, the downstream signaling cascade offers additional targets. Since Src kinase activation is required for TfR1 internalization and SDDV entry, selective Src family kinase inhibitors, such as dasatinib or saracatinib, warrant investigation. These compounds are well-characterized in human oncology and have been used experimentally in fish models, presenting a viable pathway for drug repurposing. However, the reliance on a single entry receptor raises the possibility of viral escape mutants. While TfR1 is a highly conserved host protein, the MCP sequence, though somewhat conserved among SDDV strains, may tolerate mutations that alter binding affinity [4]. Consequently, a multi-pronged strategy that simultaneously targets entry and post-entry replication steps will be essential to forestall resistance.

9.2 The Ferroptosis Nexus: From Vulnerability to Therapeutic Opportunity

A second major therapeutic frontier has emerged from the discovery that SDDV infection actively triggers ferroptosis, an iron-dependent, non-apoptotic cell death pathway, both in vivo and in vitro [5, 6]. Chen et al. demonstrated that SDDV infection leads to hallmark ferroptotic features: iron overload, massive lipid peroxide accumulation due to unchecked lipoxygenase activity, glutathione depletion, and profound downregulation of glutathione peroxidase 4 (GPX4), the master suppressor of ferroptosis [5, 6]. Critically, SDDV-induced ferroptosis is mechanistically linked to TfR1 upregulation. By enhancing TfR1 expression on the host cell surface, SDDV increases iron import, disrupts cellular redox homeostasis, and creates a permissive environment for viral replication [5]. This virus-driven manipulation of iron metabolism, termed a “ferroptotic hijack”, represents both a pathogenic mechanism and an Achilles’ heel. Treatment with the potent ferroptosis inhibitor Ferrostatin-1 significantly attenuated SDDV replication in MFF-1 cells and, more importantly, improved survival in mandarin fish following SDDV challenge [5]. Similarly, iron chelation with agents like desferrioxamine reduced mortality, confirming that iron dependency is the critical driver of SDDV-induced pathology [5]. These findings establish ferroptosis inhibition as a bona fide therapeutic strategy. Beyond Ferrostatin-1, the toolbox of ferroptosis inhibitors is expanding rapidly. Liproxstatin-1, which blocks lipid peroxidation by inhibiting lipoxygenases, and the GPX4 mimetic ebselen are prime candidates for evaluation in SDDV infection models. Moreover, the natural antioxidant vitamin C (ascorbic acid) has shown remarkable efficacy in this context. Chen et al. (2025) reported that vitamin C treatment reduced SDDV-induced mortality in mandarin fish by 37.5%, suppressed viral replication, decreased ROS and lipid peroxidation, and restored GPX4 expression [2]. Mechanistically, vitamin C acts through an Nrf2-dependent pathway, promoting Nrf2 nuclear translocation and the upregulation of a battery of antioxidant response element (ARE)-driven genes [2]. This is particularly elegant, as it targets the host’s endogenous cytoprotective machinery rather than a viral protein, reducing the probability of resistance. The Nrf2 inhibitor ML-385 completely reversed vitamin C’s antiviral effects, confirming the specificity of this pathway [2]. Given that vitamin C is inexpensive, widely available, and generally recognized as safe for aquatic species, it represents an immediately deployable therapeutic. However, dose optimization, administration route (feed-based vs. injection), and pharmacokinetics in warm-water fish require rigorous standardization. Future research should explore whether combining vitamin C with a ferroptosis inhibitor like Ferrostatin-1 or an iron chelator provides additive or synergistic protection. The intersection of iron metabolism, lipid peroxidation, and Nrf2 signaling is therefore one of the most promising nexus points for SDDV therapeutic intervention.

9.3 Immunomodulation and Vaccine-Integrated Prophylaxis

While antiviral drugs are critical for outbreak response, sustainable long-term control of SDDV will rely on effective vaccination. The formalin-inactivated SDDV vaccine (SDDV-FIV) has shown significant promise. Chokmangmeepisarn et al. (2024) demonstrated that a prime-boost strategy with SDDV-FIV elicited robust humoral immunity, with SDDV-specific IgM levels persisting up to 70 days post-vaccination, and significantly upregulated immune-related genes (cd4, cd8, ifng, mx, ighm) in head kidney and peripheral blood lymphocytes [3]. Booster vaccination conferred a 60% survival rate upon homologous challenge at 42 days post-vaccination, compared to 20% in controls, although protection waned by 70 days, suggesting the need for booster frequency optimization [3]. These findings validate the concept of whole-virus inactivated vaccines, but several limitations must be addressed. First, SDDV-FIV production requires cell culture amplification of live virulent virus, which poses biocontainment challenges and is costly for large-scale production. Second, the durability of protection against heterologous SDDV strains, which show genetic variation in repeat sequences and core genes, is unknown [4, 10]. Third, injectable vaccines are labor-intensive and stress-inducing for small fish. Future vaccine development should pivot toward next-generation platforms. Subunit vaccines based on recombinant MCP, which was previously shown to provide efficacious protection in a landmark study by de Groof et al. (2015) [14], merit further optimization. The identification of the MCP helical domain as the TfR1-binding site [1] provides a structural basis for designing protective epitopes. A rationally designed subunit vaccine targeting this receptor-binding domain could elicit neutralizing antibodies that block viral entry at the first step of infection. Virus-like particles (VLPs) produced in bacterial or yeast systems offer a safer, more immunogenic alternative to inactivated virus. The success of VLP-based vaccines for other aquatic viruses, such as nervous necrosis virus (NNV), where oral vaccination with VLPs encapsulated in hypochlorite-inactivated L. lactis induced significant antibody responses and reduced viral load [34], provides a proven roadmap. Adapting this platform for SDDV by expressing the MCP and potentially the capsid components of the inner membrane (e.g., the putative myristoylated proteins) could yield a multivalent VLP vaccine suitable for oral administration, enabling mass immunization of fry without handling stress. Furthermore, the RNA-seq data from convalescent Asian seabass [25] and the transcriptional profiling of vaccinated fish [3] offer a rich dataset to identify correlates of protection. The single-cell transcriptome profiling of SDDV-convalescent Asian seabass revealed elevated proportions of B cells, activated antibody-secreting cells, and cytotoxic CD8+ T cells, suggesting that both humoral and cell-mediated immunity are crucial for viral clearance [25]. Future vaccines should be designed to activate both arms of the adaptive immune system. This could be achieved by incorporating a CD8+ T-cell epitope from SDDV proteins (e.g., MCP or ATPase) alongside B-cell epitopes. Adjuvant development for fish vaccines is severely underdeveloped; however, the discovery that SDDV triggers Nrf2 activation [2] raises the possibility that Nrf2 agonists (e.g., sulforaphane) could serve as vaccine adjuvants by enhancing the innate immune microenvironment.

9.4 Emerging Modalities: RNA Interference, CRISPR, and Host-Directed Therapies

Beyond conventional small molecules and vaccines, several transformative technologies are poised to redefine SDDV therapeutics. RNA interference (RNAi) technology offers a sequence-specific approach to silence viral genes. For SDDV, the ATPase gene (required for viral packaging and maturation) and the MCP gene are prime RNAi targets. The development of double-stranded RNA (dsRNA) or short interfering RNA (siRNA) formulations that are stable in the aquatic environment and can be delivered orally via bioencapsulation in Artemia or rotifers could provide a scalable prophylactic strategy. However, the challenge of dsRNA stability and potential off-target effects in the host must be addressed. RNA-based biopesticides are gaining regulatory traction in China for agricultural pests, and this framework could be adapted for aquaculture [36]. A particularly innovative approach is the use of CRISPR-Cas13a systems for direct RNA degradation of SDDV transcripts. While originally developed as a diagnostic tool, as demonstrated by Sukonta et al. (2021) for SDDV detection using RPA-Cas12a [19], the Cas13 effector can be reprogrammed for therapeutic RNA knockdown. Delivery of Cas13a ribonucleoproteins via lipid nanoparticles or engineered probiotics could provide a potent, sequence-based antiviral tool. Yet, this application for finfish remains highly experimental and faces substantial hurdles in delivery, cost, and off-target editing. Host-directed therapy (HDT) represents perhaps the most resistance-proof strategy. By targeting host factors that are essential for viral replication but non-essential for the host, HDTs present a high genetic barrier to viral escape. The TfR1 receptor itself is a prime HDT target, but its essential role in iron delivery for erythrocyte production raises safety concerns [1]. However, short-term, low-dose TfR1 inhibition during an outbreak peak may be tolerable. A more attractive HDT target is the host Src kinase that phosphorylates TfR1 [1]. Src inhibitors are well-tolerated in mammals and could be screened for efficacy in SDDV-infected MFF-1 cells. Similarly, inhibitors of clathrin-mediated endocytosis, such as pitstop 2, have demonstrated antiviral activity against a range of viruses and could be evaluated for SDDV [1, 18]. The recent demonstration that decapod iridescent virus 1 (DIV1), another iridovirus, enters cells via caveola-mediated endocytosis [18], highlights the diversity of entry mechanisms within the Iridoviridae and underscores the need for target validation studies specific to SDDV. Finally, the application of compound library screening, as has been performed for white spot syndrome virus (WSSV) using a library of 303 bioactive compounds targeting specific host signaling pathways (e.g., Hedgehog, NF-κB) [42], should be urgently applied to SDDV. Such screens could rapidly identify FDA-approved drugs with repurposing potential, accelerating the path from bench to pond-side application. The convergence of these diverse modalities, entry blockers, ferroptosis inhibitors, RNA-based therapeutics, and host-directed strategies, offers a multi-layered defense against SDDV. The imperative now is to translate these mechanistic insights from the laboratory to rigorous field trials, while simultaneously developing the regulatory frameworks necessary to authorize the use of novel antiviral agents in food fish species.

References

[1] Chen J, Fu Y, Li Y, Weng S, Wang H, He J, et al.. Transferrin receptor 1 (TfR1) functions as an entry receptor for scale drop disease virus to invade the host cell via clathrin-mediated endocytosis. bioRxiv. 2025. DOI: https://doi.org/10.1128/jvi.00671-25

[2] Chen J, Fu Y, Weng S, He J, Dong C. Vitamin C Inhibits Scale Drop Disease Virus Infectivity by Targeting Nrf2 to Reduce Ferroptosis. Antioxidants. 2025. DOI: https://doi.org/10.3390/antiox14050576

[3] Chokmangmeepisarn P, Senapin S, Taengphu S, Thompson KD, Srisapoome P, Uchuwittayakul A, et al.. Protective efficiency and immune responses to single and booster doses of formalin-inactivated scale drop disease virus (SDDV) vaccine in Asian seabass (Lates calcarifer). BMC Veterinary Research. 2024. DOI: https://doi.org/10.1186/s12917-024-04132-6

[4] Chokmangmeepisarn P, Azmai MNA, Domingos J, Aerle Rv, Bass D, Prukbenjakul P, et al.. Genome Characterization and Phylogenetic Analysis of Scale Drop Disease Virus Isolated from Asian Seabass (Lates calcarifer). Animals. 2024. DOI: https://doi.org/10.3390/ani14142097

[5] Chen J, Fu Y, Chen S, Weng S, He J, Dong C. Scale drop disease virus (SDDV) triggers ferroptosis both in mandarin fish (Siniperca chuatsi) and MFF-1 cells to facilitate virus infection via linking to transferrin receptor 1 (Tfr1).. Fish and Shellfish Immunology. 2025. DOI: https://doi.org/10.1016/j.fsi.2025.110416

[6] Chen J, Fu Y, Chen S, Weng S, He J, Dong C. Scale drop disease virus (SDDV) triggering ferroptosis both in vivo and in vitro facilitates virus infection via targeting transferrin receptor 1 (TfR1). bioRxiv. 2025. DOI: https://doi.org/10.1101/2025.03.18.643978

[7] Fu Y, Mao C, Wan J, Sun Q, Yu F, Sun J, et al.. Detection and Genomic Characterisation of Scale Drop Disease Virus (SDDV) in Farmed Yellowfin Seabream (Acanthopagrus latus) in Southern China.. Journal of Fish Diseases. 2025. DOI: https://doi.org/10.1111/jfd.14161

[8] Nurliyana M, Lukman B, Ina-Salwany M, Zamri-Saad M, Annas S, Dong HT, et al.. First evidence of scale drop disease virus in farmed Asian seabass (Lates calcarifer) in Malaysia. Aquaculture. 2020. DOI: https://doi.org/10.1016/j.aquaculture.2020.735600

[9] Kerddee P, Dong H, Chokmangmeepisarn P, Rodkhum C, Srisapoome P, Areechon N, et al.. Simultaneous detection of scale drop disease virus and Flavobacterium columnare from diseased freshwater-reared barramundi Lates calcarifer.. Diseases of Aquatic Organisms. 2020. DOI: https://doi.org/10.3354/dao03500

[10] Kayansamruaj P, Soontara C, Dong H, Phiwsaiya K, Senapin S. Draft genome sequence of scale drop disease virus (SDDV) retrieved from metagenomic investigation of infected barramundi, Lates calcarifer (Bloch, 1790).. Journal of Fish Diseases. 2020. DOI: https://doi.org/10.1111/jfd.13240

[11] Domingos J, Shen X, Terence C, Senapin S, Dong H, Tan MR, et al.. Scale Drop Disease Virus (SDDV) and Lates calcarifer Herpes Virus (LCHV) Coinfection Downregulate Immune-Relevant Pathways and Cause Splenic and Kidney Necrosis in Barramundi Under Commercial Farming Conditions. Frontiers in Genetics. 2021. DOI: https://doi.org/10.3389/fgene.2021.666897

[12] Fu Y, Li Y, Fu W, Su H, Zhang L, Huang C, et al.. Scale Drop Disease Virus Associated Yellowfin Seabream (Acanthopagrus latus) Ascites Diseases, Zhuhai, Guangdong, Southern China: The First Description. Viruses. 2021. DOI: https://doi.org/10.3390/v13081617

[13] Chokmangmeepisarn P. Genome characterization and development of inactivated vaccine against scale drop disease virus (SDDV) in asian sea bass (lates calcarifer). . None. DOI: https://doi.org/10.58837/chula.the.2023.1081

[14] Groof Ad, Guelen L, Deijs M, Wal YAvd, Miyata M, Ng KS, et al.. A Novel Virus Causes Scale Drop Disease in Lates calcarifer. PLoS Pathogens. 2015. DOI: https://doi.org/10.1371/journal.ppat.1005074

[15] Swaminathan T, Raj NS, Preena PG, Pradhan PK, Sood N, Kumar R, et al.. Infectious spleen and kidney necrosis virus-associated large-scale mortality in farmed giant gourami, Osphronemus goramy, in India.. Journal of Fish Diseases. 2021. DOI: https://doi.org/10.1111/jfd.13519

[16] Guo C, He J, Xu X, Weng S, He J. Megalocytivirus: A Review of Epidemiology, Pathogenicity, Immune Evasion, and Prevention Strategies. Reviews in Aquaculture. 2025. DOI: https://doi.org/10.1111/raq.70025

[17] Kurapati Rb, Ramena G, Wanjala H, Gudapati S, Ramena Y. Viral Disease Histopathology in Aquaculture Finfish: Organ‐Specific Pathological Changes and Diagnostic Insights, Referencing the World Organisation for Animal Health: A Review. Reviews in Aquaculture. 2026. DOI: https://doi.org/10.1111/raq.70129

[18] Zheng Q, Chen X, Zhao F, Zhang J, Chen J. Decapod iridescent virus 1 (DIV1) enters hematopoietic Cherax quadricarinatus cells via caveola-mediated endocytosis in a pH-dependent manner. Journal of Virology. 2026. DOI: https://doi.org/10.1128/jvi.01681-25

[19] Sukonta T, Senapin S, Meemetta W, Chaijarasphong T. CRISPR-based platform for rapid, sensitive and field-deployable detection of scale drop disease virus in Asian sea bass (Lates calcarifer).. Journal of Fish Diseases. 2021. DOI: https://doi.org/10.1111/jfd.13541

[20] Sriisan S, Boonchird C, Thitamadee S, Sonthi M, Dong HT, Senapin S. A sensitive and specific SYBR Green-based qPCR assay for detecting scale drop disease virus (SDDV) in Asian sea bass.. Diseases of Aquatic Organisms. 2020. DOI: https://doi.org/10.3354/dao03484

[21] Charoenwai O, Senapin S, Dong H, Sonthi M. Detection of scale drop disease virus from non-destructive samples and ectoparasites of Asian sea bass, Lates calcarifer.. Journal of Fish Diseases. 2020. DOI: https://doi.org/10.1111/jfd.13290

[22] Koesharyani I, Sunarto A, Sugama K. DETEKSI PENYAKIT SCALE DROP PADA IKAN KAKAP PUTIH Lates calcarifer BLOCH. Jurnal Riset Akuakultur. 2020. DOI: https://doi.org/10.15578/JRA.15.3.2020.195-204

[23] Dinh-Hung N, Dong HT, Phiwsaiya K, Taengphu S, Linh NV, Chatchaiphan S, et al.. Natural Infection of Two Gourami Species (Trichopodus spp.) With Infectious Spleen and Kidney Necrosis Virus (ISKNV): Clinical, Molecular and Histopathological Findings.. Journal of Fish Diseases. 2025. DOI: https://doi.org/10.1111/jfd.14110

[24] Becker JA, Hick P, Megarani D, Siler HJ, Pierezan F, Gray SN, et al.. Risk of Spread of Megalocytivirus pagrus1 (Infectious Spleen and Kidney Necrosis Virus) From Frozen Fillets. Journal of Fish Diseases. 2025. DOI: https://doi.org/10.1111/jfd.70086

[25] Loh Z, Lim TW, Howland S, Awate S, Rénia L, Chen J, et al.. Single-Cell Transcriptome Profiling of Scale Drop Disease Virus-Infected Asian Seabass (Lates calcarifer). Aquaculture Journal. 2024. DOI: https://doi.org/10.3390/aquacj4020003

[26] Yin B, Mao C, Yu F, Li W, Pan R, Feng W, et al.. A droplet digital PCR method for the detection of scale drop disease virus in yellowfin seabream (Acanthopagrus latus). Frontiers in Microbiology. 2024. DOI: https://doi.org/10.3389/fmicb.2024.1444235

[27] Sriisan S, Boonchird C, Thitamadee S, Sonthi M, Dong H, Senapin S. A highly sensitive and specific SYBR Green quantitative polymerase chain reaction (qPCR) method for rapid detection of scale drop disease virus in Asian sea bass, Lates calcarifer. bioRxiv. 2019. DOI: https://doi.org/10.1101/849661

[28] Zeng Y, Zhang H, Zhang H. Isolation, Identification, and Whole Genome Analysis of Chicken Infectious Anemia Virus in an Outbreak of Disease in Adult Layer Hens. Veterinary Sciences. 2023. DOI: https://doi.org/10.3390/vetsci10070481

[29] Okoye J, Igwe AO. La Sota Vaccination Does Not Offer Satisfactory Protection Against Velogenic Newcastle Disease Virus Infection In Chickens- A Review. SPG BioMed. 2019. DOI: https://doi.org/10.32392/BIOMED.38

[30] Prasitporn T, Senapin S, Vaniksampanna A, Longyant S, Chaivisuthangkura P. Development of cross-priming amplification (CPA) combined with colorimetric and lateral flow dipstick visualization for scale drop disease virus (SDDV) detection.. Journal of Fish Diseases. 2021. DOI: https://doi.org/10.1111/jfd.13448

[31] Dangtip S, Kampeera J, Suvannakad R, Khumwan P, Jaroenram W, Sonthi M, et al.. Colorimetric detection of scale drop disease virus in Asian sea bass using loop-mediated isothermal amplification with xylenol orange. Aquaculture. 2019. DOI: https://doi.org/10.1016/J.AQUACULTURE.2019.05.071

[32] Charoenwai O, Meemetta W, Sonthi M, Dong H, Senapin S. A validated semi-nested PCR for rapid detection of scale drop disease virus (SDDV) in Asian sea bass (Lates calcarifer).. Journal of Virological Methods. 2019. DOI: https://doi.org/10.1016/j.jviromet.2019.03.007

[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] Hong HY, Carmen LCP, Chee PX, Ying LX, Wong ZW, Chan J, et al.. Oral vaccination via virus-like particles encapsulated in Lactococcus lactis.. Fish and Shellfish Immunology. 2026. DOI: https://doi.org/10.1016/j.fsi.2025.111013

[35] Lan NGT, Dong H, Shinn AP, Vinh NT, Senapin S, Salin KR, et al.. Review of current perspectives and future outlook on bacterial disease prevention through vaccination in Asian seabass (Lates calcarifer).. Journal of Fish Diseases. 2024. DOI: https://doi.org/10.1111/jfd.13964

[36] Huang Y, Dai Y, Huang Z, Zhang M, Xiu L, Zhang X, et al.. RNA‐Based Biopesticides: Pioneering Precision Solutions for Sustainable Aquaculture in China. Animal Research and One Health. 2025. DOI: https://doi.org/10.1002/aro2.70000

[37] Ng TH, M S, Chew XZ, Nair T, Chow JW, Low A, et al.. Dissecting the impact of heat stress on heat-shock response and skin microbiota in farmed fish in a recirculating aquaculture system in Singapore. Microbiology spectrum. 2025. DOI: https://doi.org/10.1128/spectrum.00568-24

[38] Acero MRC, Tacda EA, Labalan KKRJM, Pangisban BD. IoT-Enabled Autonomous Water-Surface Drone for Real-Time Water Quality Monitoring, Forecast-Driven Decision Support, and Automated Aeration in Small-Scale Fishponds. 2025 9th International Artificial Intelligence and Data Processing Symposium (IDAP). 2025. DOI: https://doi.org/10.1109/IDAP68205.2025.11222219

[39] Zenger K, Khatkar M, Jones DB, Khalilisamani N, Jerry D, Raadsma H. Genomic Selection in Aquaculture: Application, Limitations and Opportunities With Special Reference to Marine Shrimp and Pearl Oysters. Frontiers in Genetics. 2019. DOI: https://doi.org/10.3389/fgene.2018.00693

[40] Campomanes F, Pabelico JL, Recto E, Soriano R, Deligencia J, Zacarias M, et al.. Precision Ulang Aquaculture: A Sustainable Approach Leveraging Atmega2560-based Automation and Data-Driven Control. QCU The Star. 2025. DOI: https://doi.org/10.64807/ghxect90

[41] 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

[42] Lin W, Guo G, Zou C, Shi H, Ruan L. Large–scale screening of molecules involved in virus–host interaction by specific compounds in Cherax quadricarinatus hematopoietic tissue cells. Aquaculture. 2020. DOI: https://doi.org/10.1016/j.aquaculture.2020.735435

[43] Lakshmi B, Syed S, Buddolla V. Current Advances in the Protection of Viral Diseases in Aquaculture With Special Reference to Vaccination. Recent Developments in Applied Microbiology and Biochemistry. 2019. DOI: https://doi.org/10.1016/B978-0-12-816328-3.00010-6