All animal experiments were reviewed and approved by the Institutional Animal Care and Use Committee of Universiti Putra Malaysia (IACUC) (Approval No. UPM/IACUC/AUP-U016/2023). Chickens were handled by trained personnel, housed under controlled biosafety conditions, and monitored daily to ensure animal welfare. All procedures complied with institutional policies and internationally recognized guidelines for the ethical use of animals in research.

This study was conducted between July 2023 and February 2024 at the Biosafety Level 2 Animal Research Facility, Virology Laboratory, and Histopathology Laboratory, all located within the Faculty of Veterinary Medicine, Universiti Putra Malaysia, and at the Abadiah Laboratory, MTDC Technology Center III, Universiti Putra Malaysia.

Sixty 21-day-old SPF chickens were housed in stainless steel wire cages within a Biosafety Level-2 Research Facility maintained at 26°C, 63% relative humidity, and continuous lighting, with feed and water provided ad libitum. The chickens were randomly allocated into Group A (nVarIBDV-inoculated, n = 28) and Group B (control, n = 32). Each group was maintained in separate rooms that were cleaned daily, with personnel movement from the control to the inoculated group and the use of disposable PPE and strict biosecurity measures.

At 0 dpi (21 days of age), chickens in Group A were inoculated with 1.0 mL of pathogenic field strain nVarIBDV (106.75 EID₅₀/mL) via combined ocular (0.1 mL) and oral (0.9 mL) routes, while Group B remained uninoculated. Clinical signs were monitored at least twice daily, and any moribund chickens were humanely sacrificed for sampling. Four chickens from Group B were sacrificed at 0 dpi for baseline evaluation. Subsequently, four chickens from each group were randomly selected and sacrificed at 1, 3, 5, 7, 10, 14, and 21 dpi.

Prior to sacrifice, body weight was recorded, and blood samples were collected for the determination of IBD antibody titers using ELISA. Gross lesions were recorded at necropsy. Bursa of Fabricius and spleen weights were measured, and bursa-to-body weight and spleen-to-body weight ratios were calculated. Bursa samples were fixed in Formalin solution (10% neutral buffered formalin) (SysterM, Shah Alam, Malaysia) for histopathology and lesion scoring. Additional samples from the bursa, spleen, cecal tonsil, thymus, and bone marrow were collected for the detection of nVarIBDV using RT-qPCR (Figure 1).

The nVarIBDV isolate (UPM1432/2019) was obtained from a commercial broiler farm in Selangor, Malaysia [3]. The isolate was confirmed as nVarIBDV by PCR and sequencing (GenBank accession number MT431217). Bursa tissues from affected chickens were processed, and the viral supernatant was inoculated into embryonated SPF chicken eggs. The CAM from infected eggs was harvested, mixed with PBS (Sigma-Aldrich, Missouri, USA) at a 1:2 ratio (CAM:PBS), manually homogenized with a mortar and pestle, and centrifuged at 252 × g for 5 min. The clarified supernatant was filtered through syringe filters (Membrane Solutions, Shanghai, China) to obtain the viral inoculum [19].

Viral titration was performed using the Reed and Muench method to determine EID₅₀/mL [20]. The inoculum was stored at −80°C and used within 48 h after thawing.

Ten-fold serial dilutions of nVarIBDV were inoculated into 9–11-day-old SPF embryonated chicken eggs, with five eggs per dilution, via the CAM route. Eggs were incubated at 37°C for 7 days and candled daily. Mortality within 24 h post-inoculation was regarded as nonspecific contamination and excluded from analysis; contaminated eggs were discarded [21]. Viral titers were calculated and expressed as EID₅₀/mL using the Reed and Muench method [20].

Serum samples were analyzed for IBD antibody using a commercial IBDV antibody ELISA kit (BioChek BV, Reeuwijk, Netherlands). Antigen-coated plates were equilibrated to room temperature (22°C–27°C). Negative and positive controls (100 µL each) and 1:500 (v/v) diluted serum samples (100 µL) were added to the wells and incubated for 30 min at room temperature. Plates were washed four times with wash solution (350 µL/well), followed by the addition of 100 µL conjugate reagent (anti-chicken IgG conjugated with alkaline phosphatase) and incubation for 30 min.

After washing, 100 µL of substrate buffer was added to each well and incubated in the dark for 15 min at room temperature (22°C–27°C). The reaction was stopped by adding 100 µL stop solution. Absorbance was measured at 405 nm using a microplate reader, MR7000 (Dynatech Laboratories, Denkendorf, Germany), and IBD antibody titers were calculated using BioChek 2000 software.

Pooled tissue samples from the bursa of Fabricius, spleen, thymus, bone marrow, and cecal tonsil were analyzed for viral detection using RT-qPCR in compliance with MIQE guidelines. RNA was extracted using a commercial RNA/DNA purification kit (Kylt®, Höltinghausen, Germany), with 20 mg of tissue per extraction. Lysis, binding, washing, and elution steps were performed according to the manufacturer’s instructions.

RT-qPCR was conducted using a SensiFAST™ Probe No-ROX One-Step Kit (Bioline, London, United Kingdom). Reverse transcription and amplification were performed using specific primers and probes (Table 1) [11]. Viral genome copy numbers were quantified using a Bio-Rad CFX Opus 96 real-time PCR system (BIO-RAD, Hercules, California, USA).

Bursa of Fabricius samples were fixed in 10% buffered formalin (SysterM) for at least 24 h, trimmed to 5 mm thickness, and processed using an automated tissue processor (Leica ASP300, Wetzlar, Germany). Paraffin-embedded tissues (Leica EG1160) were sectioned at 4 µm and stained with hematoxylin and eosin (Leica). Histopathological changes were evaluated under light microscopy (Leica DM LB2).

Bursal lesion severity was scored in a blinded manner on a scale of 0–5 as follows: 0 = normal, 1 = mild, 2 = mild to moderate, 3 = moderate, 4 = moderate to severe, and 5 = severe [22].

Data were analyzed using Student’s t-test in IBM SPSS Statistics version 23.0 (IBM Corp., Armonk, NY, USA). Results were expressed at a 95% confidence level, and differences were considered statistically significant when p < 0.05.

No clinical signs were observed in Group B chickens throughout the study period. In contrast, chickens in Group A exhibited whitish to yellowish watery droppings from 3 to 12 dpi (Supplementary Figure 1), brownish to reddish watery droppings on 8, 9, and 18 dpi (Supplementary Figure 2), and ruffled feathers from 13 to 21 dpi (Supplementary Figure 3).

Body weight increased progressively in both groups from 0 to 21 dpi, with no statistically significant difference between groups at any time point (p > 0.05). However, chickens in Group A showed higher body weight at 1, 3, and 5 dpi, whereas chickens in Group B exhibited higher body weight at 7, 10, and 14 dpi (Figure 2).

The bursa weight of Group B increased steadily from 0 to 21 dpi. In contrast, Group A showed a marked reduction in bursa weight from 1 to 5 dpi, followed by stabilization from 5 to 21 dpi. No significant difference was observed between groups at 1 and 3 dpi (p > 0.05). However, from 5 to 21 dpi, bursa weight in Group A was significantly lower than in Group B (p < 0.05) (Supplementary Figure 4).

No statistically significant difference was detected in the bursa-to-body weight ratio between groups at 1 dpi (p > 0.05). From 3 to 21 dpi, the ratio in Group A was significantly lower than in Group B (p < 0.05). In Group A, the bursa-to-body weight ratio declined sharply from 1 to 5 dpi and decreased more gradually from 5 to 21 dpi (Figure 3).

Spleen weight increased progressively in both groups from 1 to 21 dpi. At 5 and 7 dpi, spleen weight in Group A was significantly higher than in Group B (p < 0.05). At 21 dpi, spleen weight in Group A remained higher than in Group B, although the difference was not statistically significant (p > 0.05) (Supplementary Figure 5).

The spleen-to-body weight ratio was higher in Group A than in Group B at 1, 3, 10, 14, and 21 dpi; however, these differences were not statistically significant (p > 0.05). In contrast, at 5 and 7 dpi, the ratio was significantly higher in Group A compared with Group B (p < 0.05) (Figure 4).

IBD antibodies became detectable from 5 dpi. Antibody titers increased rapidly between 5 and 7 dpi and gradually declined from 7 to 21 dpi. No statistically significant difference was observed between groups from 1 to 5 dpi (p > 0.05). From 7 to 21 dpi, antibody titers in Group A were significantly higher than those in Group B (p < 0.05) (Figure 10).

The highest nVarIBDV load was detected in the bursa, whereas the lowest viral load was observed in the bone marrow. Viral loads in the bursa and thymus peaked at 3 dpi and gradually declined thereafter. Viral loads in the spleen and bone marrow peaked at 5 dpi, while the cecal tonsil reached the highest viral load at 7 dpi. All organs exhibited peak viral loads between 3 and 7 dpi. Viral RNA was not detected in the bone marrow after 7 dpi (Figure 11; Table 2).

Since its initial description, nVarIBDV has spread widely across multiple regions and has been associated with immunosuppression and substantial economic losses in poultry production systems [17, 23]. In the present study, the pathogenicity and immunogenicity of nVarIBDV were comprehensively evaluated using 11 biological, pathological, immunological, and molecular parameters. The controlled experimental infection of SPF chickens using a defined inoculum dose (106.75 EID₅₀/mL), dual inoculation routes, and systematic sampling intervals provides a robust and reproducible model for studying nVarIBDV infection dynamics.

Infected chickens exhibited ruffled feathers, watery diarrhea ranging from whitish–yellowish to brownish–red, depression, insensitivity, and reduced feed intake persisting up to 21 dpi, without recorded mortality. These findings indicate that nVarIBDV does not directly induce mortality but can cause prolonged clinical illness that may translate into significant economic losses due to reduced performance and increased management costs [4, 11, 23].

Although no statistically significant difference in body weight was observed between groups throughout the study, chickens in the control group had higher body weight at 7, 10, and 14 dpi. This suggests that nVarIBDV transiently impaired growth performance in Group A until 14 dpi, followed by recovery at 21 dpi. These findings differ from reports by Huang et al. [23] and Fan et al. [10], who observed significant body weight reduction following nVarIBDV infection. This discrepancy may be attributed to the use of SPF layer-type chickens in the present study, which inherently exhibit slower growth rates than commercial broilers.

A significant reduction in bursa weight was observed in nVarIBDV-infected chickens from 5 to 21 dpi, indicating progressive bursal atrophy. These findings are consistent with previous reports showing that nVarIBDV induces marked bursal damage beginning at 5 dpi, with sustained effects [4–6, 11]. The bursa-to-body weight ratio further confirmed the occurrence and severity of bursal atrophy and served as a more reliable indicator of bursal damage. Bursal atrophy is a hallmark of IBDV-induced immunosuppression and predisposes infected chickens to secondary infections and vaccination failure. Indeed, nVarIBDV has been reported to reduce the efficacy of ND and AI vaccination [4, 10].

Splenomegaly was observed in nVarIBDV-infected chickens at 5 and 7 dpi, in agreement with previous findings [4]. However, other studies have reported no significant differences in spleen-to-body weight ratio between infected and control chickens [11, 23]. Histological alterations in the spleen may impair immune function by affecting lymphoid architecture and splenocyte activity, thereby contributing to overall immunosuppression [11].

Severe gross lesions, including bursal atrophy, yellowish discoloration, reduced folds, and firm consistency, were evident from 3 to 21 dpi, indicating extensive bursal damage following nVarIBDV infection. Histopatho-logically, moderate to severe lesions persisted throughout the same period, consistent with earlier reports [4, 11]. The observed lesions closely resemble those induced experimentally by vvIBDV [4, 11, 24]. Although vvIBDV is typically associated with high mortality, the comparable degree of bursal destruction observed with nVarIBDV highlights that immunosuppression alone can be equally detrimental to flock health and productivity.

The humoral immune response was characterized by detectable IBD antibody production beginning at 5 dpi, with a rapid increase peaking at 7 dpi, coinciding with the active phase of viral replication. Immunosuppression following IBDV infection is primarily mediated by viral targeting of B lymphocytes and macrophages [25]. Additionally, effects on T cells [26] and splenocytes [27] have been reported, which may further compromise vaccine-induced immunity. Despite these immunosuppressive effects, high antibody titers (13,000–14,000) were maintained from 7 to 21 dpi, consistent with previous studies [11, 28, 29], indicating strong humoral immunogenicity and confirming the pathogenic potential of the isolate.

Following oral entry, nVarIBDV initially replicates in GALT before entering the bloodstream and reaching the bursa, where extensive replication occurs, followed by secondary viremia and dissemination to other lymphoid organs [30]. Detection of viral RNA in the bursa, spleen, cecal tonsil, thymus, and bone marrow up to 21 dpi underscores the strong immunocompromising potential of nVarIBDV. To the best of our knowledge, this is the first report documenting nVarIBDV persistence in these organs for up to 21 dpi. Previous studies reported viral detection up to 14 dpi [31] or limited to the bursa up to 7 dpi [11], while other variant strains were described as short-lived and undetectable after 7 dpi [12, 32]. The sustained lesion severity observed from 3 dpi onward, coinciding with peak viral load in the bursa, indicates limited tissue recovery. Persistent infection of multiple immune organs may disrupt their structural integrity and function, facilitate viral shedding, and promote disease spread, thereby informing containment strategies and vaccination programs for ND and AI. Prolonged viral persistence may also exacerbate long-term immunosuppression and production losses, highlighting the importance of optimizing vaccination timing.

Given the ability of nVarIBDV to escape existing IBD vaccine immunity [3], the development of vaccines specifically targeting nVarIBDV is imperative. Potential approaches include whole-virus inactivated vaccines or recombinant vaccines expressing the VP2 protein using platforms such as HVT. Future studies should investigate the relationship between viral persistence and unique VP2 motifs, replication kinetics, and host immune responses. Molecular profiling of immune markers and cytokine expression, along with larger sample sizes, would further strengthen understanding of nVarIBDV pathogenesis. As viral RNA was detected up to 21 dpi, extended persistence studies may provide critical insights for refining containment measures and vaccine design.