Tilapia is the world’s second most cultivated freshwater fish and has become an important source of dietary protein as well as a substantial economic resource in developing nations (FAO, 2022; Hounmanou et al., 2018). Global tilapia production, including other cichlids, reached 6.1 million metric tons in 2020, with China, Thailand, Ecuador, Egypt and Indonesia as major contributors (Harvey, 2016). Despite their relative resistance, farmed tilapia are susceptible to a variety of disease-causing organisms (Machimbirike et al., 2019; Surachetpong et al., 2020; Kaviarasu et al., 2022).
       
Diseases of viral origin are considered among the most significant threats in tilapia aquaculture and Tilapia Lake Virus (TiLV) has emerged as one of the most important pathogens in recent years (Eyngor et al., 2014; Ferguson et al., 2014). TiLV, also known as Tilapia tilapinevirus, is an enveloped virus with a 10-segment, negative-sense RNA genome encoding 14 predicted proteins belonging to the genus Tilapine virus within the family Amnoonviridae (Koonin et al., 2023) and is listed as an emerging finfish disease by the World Organisation for Animal Health (WOAH, 2022).
       
Detection of TiLV generally relies on molecular techniques that identify viral genetic material in clinical specimens. Several diagnostic assays have been developed, including RT-PCR (Eyngor et al., 2014), nested RT-PCR (Kembou et al., 2017), semi-nested RT-PCR (Dong et al., 2017), RT-qPCR (Tattiyapong et al., 2018; Waiyamitra et al., 2018; Taengphu et al., 2020; Taengphu et al., 2022), RT-LAMP (Phusantisampan et al., 2019; Yin et al., 2019; Kampeera et al., 2021) and nanopore-based PCR amplicon sequencing (Delamare-Deboutteville et al., 2021). However, most of these assays target a single genome segment, which may reduce diagnostic reliability when a probable genetic variation occurs in the primer-binding region.
       
Growing evidence indicates that TiLV undergoes frequent genetic reassortment, which is a major driver of its genomic diversity (Chaput et al., 2020; Thawornwattana et al., 2021; Verma et al., 2022). As a result, assays that amplify only one segment may fail to detect variants carrying mutations in that locus (Kembou et al., 2017; Dong et al., 2017; Waiyamitra et al., 2018; Yin et al., 2019). This problem has become more evident with reports of TiLV-negative fish that nevertheless display typical syncytial hepatitis, one of the hallmark lesions of TiLV infection indicating that the viral variants may escape detection due to mutations or reassortment in the targeted genome region (Taengphu et al., 2020).
               
These concerns highlight the need for diagnostic strategies that simultaneously target multiple conserved genomic regions to improve robustness and reduce the likelihood of false negatives. Therefore, the present study aimed to develop a mPCR assay targeting TiLV segments 2, 3 and 8. By amplifying three conserved regions in a single reaction, this multi-segment approach provides a more reliable and comprehensive detection tool for TiLV in tilapia aquaculture.

Sample collection
 
Samples used in this study were obtained from a previously published TiLV surveillance investigation conducted across 15 tilapia farms in Tamil Nadu, India (Ponsrinivasan et al., 2025). The original surveillance included both clinically affected and apparently healthy fish, providing a broad representation of farm-level infection status. For the present study conducted during 2023 and 2024, a subset of TiLV-positive (n = 100) and TiLV-negative (n = 100) samples previously confirmed by semi-nested RT-PCR targeting segment 3 (Dong et al., 2017) was randomly selected ensuring unbiased representation of the broader dataset. All laboratory analyses in the present study were performed at the State Referral Laboratory for Aquatic Animal Health (AAH), Madhavaram, Chennai.
 
RNA extraction and cDNA synthesis
 
Total RNA was extracted from the eye, liver, brain, gills, kid­ney and spleen tissues using RNAiso Plus reagent (Takara Bio) according to the manufacturer’s protocol. RNA quality and concentration were assessed using a Nanodrop ND-1000 spectrophotometer (Thermo Fischer, USA). The RNA was then transcribed to cDNA using a first-strand cDNA synthe­sis kit (QIAGEN, Germany, Thermo Scientific) and stored at -80°C.
 
Primer designing
 
Primers were designed to target conserved regions of TiLV genome segments 2, 3 and 8. Representative sequences from multiple countries (Table 1) were retrieved from GenBank and aligned using Clustal Omega (https://www.ebi.ac.uk/jdispatcher/msa/clustalo) to identify conserved regions. Primers were then designed using Primer-BLAST (https://www.ncbi.nlm.nih.gov/tools/primer-blast/) using the following criteria: GC content: 40-60%, Melting temperature (Tm): 52-62°C, Low self-complementarity; avoidance of hairpins and dimers, High specificity to TiLV with no predicted cross-reactivity. The designed primers were examined for multiplex compatibility and synthesized by Eurofins Technologies (Bangalore, India). Primer sequences and corresponding amplicon sizes are listed in Table 2.




 
PCR amplification and optimization
 
PCR amplification was first optimized in uniplex format to evaluate the performance of each primer pair targeting TiLV genome segments 2, 3 and 8. Each 25 µL uniplex reaction contained 12.5 µL of 2× Red PCR Master Mix (Amplicon, Denmark), 1 µL each of forward and reverse primers (10 µM; final concentration 0.4 µM), 1 µL of cDNA template and nuclease-free water to a final volume of 25 µL. Thermal cycling conditions consisted of an initial denaturation at 95°C for 5 min, followed by 35 cycles of 95°C for 30 s, annealing at a 52-60°C gradient for 30 s and extension at 72°C for 45 s, with a final extension at 72°C for 5 min. Gradient optimization showed that 58°C was the most suitable annealing temperature across all three segments. This temperature produced strong, clear and distinct amplicons for segments 2, 3 and 8, eliminated non-specific bands and ensured balanced amplification efficiency among primer sets.
       
For mPCR, all three primer pairs were combined in a single 25 µL reaction using the same reagent composition as the uniplex assays, with each primer maintained at a final concentration of 0.4 µM. Cycling conditions were identical to the optimized uniplex protocol, using the standardized 58°C annealing temperature. PCR amplicons were resolved on 1.5% agarose gels prepared in 1× TAE buffer, stained with 0.5 µg/mL ethidium bromide and visualized using a Bio-Rad gel documentation system (Bio- Rad, Germany). Both uniplex and multiplex assays showed consistent and reproducible amplification profiles across three independent runs.
 
Analytical sensitivity and specificity
 
Analytical sensitivity was evaluated using 10-fold serial dilutions of TiLV-positive cDNA, starting from 100 ng per reaction. All assays were carried out in three independent runs in triplicates to ensure consistency and the detection limit was defined as the lowest dilution at which all three TiLV target segments produced visible amplicons. Gel densitometry was performed using ImageJ 1.x to objectively quantify band intensities (Schneider et al., 2012). Each PCR lane was selected with the rectangular ROI tool and lane profiles were generated using Analyse → Gels → Select First Lane/Select Next Lane → Plot Lanes. Peak detection was carried out on the resulting intensity profiles. A band was considered as detectable when a corresponding peak was present above baseline noise. Peak area (integrated optical density) and relative intensity were used to compare detection across template dilutions. This analysis allowed objective determination of the lowest template concentration at which at least one amplicon peak remained detectable.
       
Assay specificity was assessed using DNA/cDNA from non-TiLV aquatic viral and bacterial pathogens, such as Tilapia parvovirus, Lymphocystis disease virus, Carp edema virus, Aeromonas veronii, Streptococcus agalactiae, Enterococcus sp. and Pseudomonas sp. Specificity testing was also conducted in three independent runs in triplicates and no amplification was observed for any non-TiLV organisms.
 
Diagnostic accuracy assessment
 
Diagnostic performance of the mPCR assay was compared with the reference semi-nested RT-PCR method (Dong et al., 2017) using 200 field samples. Samples were classified as true positives, true negatives, false positives, or false negatives. Receiver Operating Characteristic (ROC) analysis was carried out in R version 4.3.0 using the pROC package (R Core Team, 2021). Band presence/absence (binary scoring) was used for diagnostic classification of the positive and negative samples. The area under the curve (AUC) was calculated to assess overall diagnostic accuracy.

Sample validation
 
Tilapia Lake Virus (TiLV) has emerged as a major transboundary pathogen in tilapia aquaculture, with reports from several countries including Colombia, Chinese Taipei, Egypt, Thailand, Malaysia (Jansen et al., 2019). In India, TiLV has been recently confirmed in farmed tilapia, (Rao et al., 2024; Rebecca et al., 2025) underlining the urgent need for sensitive and reliable diagnostic tools to support surveillance and early intervention strategies. To address this issue, we conducted an on-field surveillance as part of our research work and collected samples with visual clinical signs of exophthalmia, skin hemorrhages, fin erosion and abdominal distension (Fig 1). Samples selected from the previous surveillance study (Ponsrinivasan et al., 2025) were reconfirmed using semi-nested RT-PCR targeting segment 3, yielding the expected 415 bp and 250 bp amplicons (Fig 2). This confirmation ensured that only verified positive and negative samples were used for subsequent assay validation.




 
PCR optimization
 
Uniplex PCR assays were performed to evaluate the performance of primers targeting segments 2, 3 and 8 across annealing temperatures from 52°C to 60°C. Each primer pairs produced clear amplicons of 724 bp, 553 bp and 331 bp, respectively (Fig 3). The amplification profiles were reproducible across three independent runs, with each primer pair yielding amplicons of identical size with no nonspecific bands. Gradient analysis showed that 58°C yielded the strongest, most distinct bands for all three segments and this temperature was selected for multiplex optimization. Using the optimized annealing temperature of 58°C, the mPCR successfully amplified all three segments simultaneously, generating distinct bands of the expected sizes without cross-reactivity or primer interference (Fig 4). The ability to amplify three conserved genomic regions in a single reaction demonstrates the robust primer compatibility and enhances diagnostic confidence by reducing the risk of false negatives.




       
The choice of these segments was guided by their diagnostic and evolutionary relevance, as segment 2 is evolutionarily stable and less prone to reassortment, making it a reliable diagnostic marker (Chaput et al., 2020). Segment 3 has been widely used in TiLV detection and phylogenetic studies (Verma et al., 2022; Tattiyapong et al., 2018; Waiyamitra et al., 2018; Taengphu et al., 2022), while segment 8 is highly conserved with minimal reassortment (Thawornwattana et al., 2021). Notably, studies on TiLV genome dynamics indicate that reassortment is a dominant evolutionary force, highlighting the importance of multi-target assays to capture genetic diversity and ensure reliable detection (Verma et al., 2022; Tran et al., 2022). By targeting a combination of segments with different evolutionary stability, the assay enhances detection accuracy across genetically diverse isolates and reduces the risk of false negatives caused by mutations or segment reassortment.
 
Sensitivity and specificity
 
The analytical sensitivity of the optimized mPCR was evaluated using serial ten-fold dilutions of TiLV template cDNA ranging from 100 (nanogram) ng to 10 pg. It revealed that the mPCR could detect TiLV concentrations as low as 100pg/µL (Fig 5). Sensitivity testing was repeated across three independent runs and the detection limit remained consistent, confirming assay reliability. To provide an objective measure of band detectability, densitometric peak analysis was performed in ImageJ. Lane profiling produced three clear peaks corresponding to the multiplex amplicons in the 100 ng, 10 ng and 1 ng lanes. At 100 pg, a two small peak remained detectable, while the remaining target fell below the peak detection threshold. No measurable peaks were detected in the 10 pg reaction, consistent with the absence of visible bands on the gel (Fig 6). Together, these objective measurements establish an analytical sensitivity of 100 pg for the mPCR assay, with no detectable amplification observed at 10 pg. Although primer competition contributed to reduced band intensity at low template concentrations, detection across all targets indicates the assay’s suitability for surveillance and early-stage infection detection.




       
Most published TiLV assays report detection limits in viral copy number, whereas our study reports sensitivity in template mass (100 pg/µL of cDNA). Because these units are not directly comparable without copy-number calibration, direct numerical comparison is not possible. Nested and semi-nested RT-PCR assays have demonstrated low-copy detection limits, including ~7 copies per reaction (Dong et al., 2017; Kembou et al., 2017). qRT-PCR assays remain the most sensitive, detecting as few as 2-62 copies per reaction (Tattiyapong et al., 2018; Megarani et al., 2022; Chengula et al., 2022), while iron-flocculation-coupled qPCR detects ~10 copies/µL in environmental samples (Taengphu et al., 2022). In contrast, the mPCR described here emphasizes diagnostic robustness through multi-segment detection rather than maximal analytical sensitivity, reducing the likelihood of false negatives due to segment-specific mutations or reassortment.
       
DNA from non-TiLV aquatic pathogens including Tilapia parvovirus, Lymphocystis disease virus, Carp edema virus, Aeromonas veronii, Streptococcus agalactiae, Enterococcus  sp. and Pseudomonas sp. did not yield any amplification (Fig 7). Specificity testing was conducted in three independent runs for each pathogen and all runs produced identical results. These findings confirm that the assay specifically targets TiLV without cross-amplification of co-infecting bacterial or viral pathogens that commonly affect tilapia in aquaculture systems.


 
ROC curve
 
Diagnostic performance was evaluated by comparing the developed mPCR with the semi-nested RT PCR assay of Dong et al., (2017) using 200 field samples (TiLV-positive and TiLV-negative). ROC curve analysis revealed an AUC value of 1.0, indicating perfect concordance with the reference method and demonstrating 100% diagnostic sensitivity (Fig 8). A reaction was scored positive only if a band appeared at the expected size and exceeded background noise under uniform exposure conditions. While this high AUC demonstrates excellent diagnostic discrimination, we acknowledge that perfect performance likely reflects the well-characterized nature of the validation set. Additional testing on a broader panel of field samples and diverse TiLV genotypes would further substantiate assay robustness.


 
Application and limitations
 
The developed mPCR provides a balanced compromise between sensitivity, specificity and operational accessibility. While qRT-PCR remains the most analytically sensitive method but requires real-time platforms, fluorescent chemistries and skilled personnel limiting its use in many routine diagnostic settings. LAMP assays are rapid and equipment-light but vulnerable to nonspecific amplification and aerosol contamination if strict workflow controls are not followed. This risk is particularly pronounced in non-segregated workspaces or high-template environments where aerosolised product can seed unintended amplification and lead to false positives. Although the mPCR requires a thermocycler, electrophoresis system and gel documentation unit, it is more accessible than qRT-PCR and more robust than single-segment PCR assays, particularly in the context of TiLV genome reassortment. Therefore, the assay is most suitable for regional diagnostic facilities rather than on-farm use and its practical role is within surveillance networks where farms can submit samples to nearby laboratories for routine screening.
       
Targeting three conserved segments may  reduce the risk of false negatives, yet it is also important to know that highly divergent TiLV variants could still potentially affect primer binding. Broader validation and periodical update to primer sets using isolates from different regions and genetic backgrounds will be important for confirming its robustness. The assay was validated using tissue-derived cDNA from clinically affected fish, particularly from ponds experiencing clinical signs and mortality, where viral loads are expected to be sufficient for reliable detection; its application low-viral-load samples or alternative matrices warrants future evaluation.

The optimized mPCR assay targeting TiLV segments 2, 3 and 8 represents a reliable, sensitive and specific diagnostic platform. By addressing genomic variability and minimizing false negatives, it enhances detection accuracy and provides a robust tool for effective TiLV surveillance and disease management in tilapia aquaculture. Given the ongoing global threat posed by TiLV, the adoption of this multi-segment diagnostic strategy has the potential to significantly strengthen disease management frameworks, safeguard production and support the sustainability of tilapia aquaculture.

The authors acknowledge the research facilities extended by Tamil Nadu Dr. J. Jayalalithaa Fisheries University to carry out this research work.
 
Disclaimers
 
The views and conclusions expressed in this article are solely those of the authors and do not necessarily represent the views of their affiliated institutions. The authors are responsible for the accuracy and completeness of the information provided, but do not accept any liability for any direct or indirect losses resulting from the use of this content.
 
Informed consent
 
All animal procedures for experiments were approved by the Committee of Experimental Animal care and handling techniques were approved by the University of Animal Care Committee.

The authors declare that there are no conflicts of interest regarding the publication of this article. No funding or sponsorship influenced the design of the study, data collection, analysis, decision to publish, or preparation of the manuscript.