This cross-sectional experimental study was conducted from March 2024 to March 2025. A total of 48 healthy neonatal Sprague-Dawley rats (7 days old, both sexes) were obtained from the institutional breeding colony. Pups were randomly assigned to one of three groups (16 pups per group): Control, BE and BE + DHA. Each litter was distributed across groups to minimize confounders.

We employed a neonatal bilirubin encephalopathy (BE) model based on intraperitoneal administration of unconjugated bilirubin in postnatal day 7 (PND7) Sprague–Dawley rat pups. This experimental paradigm produces acute bilirubin neurotoxicity by directly elevating circulating unconjugated bilirubin levels and has been widely used to reproduce the neuropathological features of kernicterus in neonatal rodents. In contrast to hemolytic models induced by agents such as phenylhydrazine (PHZ), which generate hyperbilirubinemia through erythrocyte destruction, the bilirubin injection model allows controlled elevation of bilirubin concentrations and precise timing of neurotoxic exposure. Therefore, it is particularly suitable for investigating the mechanisms of bilirubin-induced neuronal injury and evaluating potential neuroprotective interventions (8, 14).

DHA (100 mg/kg/day, oral gavage) was administered on PND7, PND8 and PND9, beginning ∼1 h after bilirubin injection on PND7. Behavioral testing on PND9 and PND11 was performed before daily gavage to avoid acute effects. Controls and BE group pups received equivalent vehicle gavage (8).

Neurobehavioral assessments were conducted to evaluate sensorimotor development and neurological function in neonatal rats. Righting reflex testing was performed on PND9, whereas negative geotaxis testing was performed on PND11, after completion of the DHA treatment period.

The experimental timeline and group allocation are summarized in Figure 1. Additional methodological details are provided in Supplementary Figure 1. Bilirubin injection was performed on postnatal day 7 (PND7), which was defined as time 0. Subsequent measurements at 24 h, 48 h, and 72 h therefore corresponded to PND8, PND9, and PND10, respectively. DHA treatment was administered daily from PND7 to PND9. Behavioral testing was conducted on PND9 (righting reflex) and PND11 (negative geotaxis). Blood samples for bilirubin measurement were collected at 24, 48, and 72 h after bilirubin injection, and subsets of animals were euthanized at 72 h (PND10) for biochemical, molecular, and histopathological analyses (Figure 1).

Brain tissue bilirubin content was quantified by a solvent extraction method: cerebrum samples were weighed, homogenized, and mixed with acetone to extract bilirubin, which was then measured via an automatic biochemical analyzer at 450 nm. Results were expressed as micrograms of bilirubin per gram of brain tissue (μg/g). Oxidative stress markers were evaluated in brain cortex homogenates collected at 72 h. Malondialdehyde (MDA), an end-product of lipid peroxidation, was measured by thiobarbituric acid reactive substances assay (using a commercial kit, Sigma) and normalized to protein content (nmol MDA per mg protein). We also measured reactive oxygen species (ROS) levels in cortical cells using a 2′,7′-dichlorofluorescein diacetate (DCFDA) fluorescence assay. Briefly, fresh cortex tissue was minced and incubated with DCFDA; the fluorescence intensity (excitation 485 nm, emission 535 nm) was recorded on a microplate reader, reflecting ROS accumulation. In addition, we examined expression of key proteins involved in oxidative stress and ferroptosis pathways, including GPX4, PTGS2 (COX-2), and ACSL4, by Western blot.

Experimental timeline and group allocation of neonatal rat model of BE. On postnatal day 7 (PND7), BE and BE + DHA pups received intraperitoneal bilirubin (120 mg/kg); controls received vehicle. DHA was administered orally on PND7–PND9. Righting reflex and negative geotaxis testing were performed on postnatal days PND9 and PND11, respectively. DHA treatment was administered from PND7 to PND9; therefore, the PND11 behavioral assessment was conducted after completion of the treatment period.

Serum bilirubin levels were quantified using a micro-volume spectrophotometric bilirubinometer calibrated for neonatal rat serum samples. This approach differs from the clinical biochemical analyzer used in our previous study, which was optimized for human samples. Brain tissue bilirubin levels were determined using an acetone-based solvent extraction method followed by spectrophotometric detection at 450 nm, allowing quantification of bilirubin deposition within neural tissue.

Neonatal Sprague–Dawley rats were randomly assigned to Control, BE, and BE + DHA groups (n = 16 per group). Bilirubin injection was performed on postnatal day 7 (PND7), defined as time 0. DHA was administered orally once daily from PND7 to PND9. Blood samples for serum bilirubin measurement were collected at 24 h (PND8), 48 h (PND9), and 72 h (PND10) after bilirubin injection. Behavioral testing was conducted on PND9 (righting reflex) and PND11 (negative geotaxis). At 72 h post-injection (PND10), subsets of animals were euthanized for biochemical, molecular, and histopathological analyses.

Neonatal Sprague–Dawley rats were randomly assigned to Control, BE, or BE + DHA groups (n = 16 per group). Bilirubin injection was performed on postnatal day 7 (PND7), defined as time 0. DHA was administered orally once daily from PND7 to PND9. Blood samples for serum bilirubin measurement were collected at 24 h (PND8), 48 h (PND9), and 72 h (PND10) after bilirubin injection. Behavioral testing was conducted on PND9 (righting reflex) and PND11 (negative geotaxis). At 72 h post-injection (PND10), subsets of animals were euthanized for biochemical, molecular, and histopathological analyses. The diagram illustrates the allocation of animals and timing of experimental procedures.

To determine whether miR-155-5p directly targets Kdm5a, a luciferase reporter assay was performed in HEK293 cells. Cells were co-transfected with a pmirGLO reporter plasmid containing the Kdm5a 3′UTR, a miR-155-5p mimic or inhibitor, and/or a CTBP1 overexpression plasmid using Lipofectamine 3,000 (Thermo Fisher). After 48 h, firefly and Renilla luciferase activities were measured using the Dual-Luciferase Reporter Assay System (Promega). Firefly luciferase activity was normalized to Renilla luciferase activity.

At 72 h post-injection, rats (n = 6 per group) were anesthetized and decapitated for cortical tissue collection. Total RNA was extracted from cerebral cortex using TRIzol (Invitrogen, United States). cDNA synthesis was performed with the RevertAid kit (Thermo, United States) for mRNA and the miScript kit (Qiagen, Germany) for miRNA. Quantitative RT-qPCR was performed on an ABI Prism system using SYBR Green Mastermix. We quantified mRNA levels of Ctbp1 and Kdm5a, and miRNA levels of miR-155-5p normalized to U6 small RNA. Primers were designed based on rat sequences (miR-155-5p assay ID: rno-miR-155-5p). Relative expression was calculated using the 2^−ΔΔCt method, with the Control group as calibrator (fold change = 1.0).

Cortical tissue samples were lysed in RIPA buffer containing protease inhibitors. Protein concentrations were determined using the bicinchoninic acid (BCA) assay. Thirty micrograms (30 μg) of total protein per sample were separated by SDS–polyacrylamide gel electrophoresis and transferred onto polyvinylidene fluoride (PVDF) membranes.

Membranes were blocked with 5% non-fat milk in Tris-buffered saline with 0.1% Tween-20 (TBST) for 1 h at room temperature and then incubated overnight at 4°C with primary antibodies against CTBP1, KDM5A, GPX4, ACSL4, and COX-2 (PTGS2). After washing with TBST, membranes were incubated with horseradish peroxidase–conjugated secondary antibodies for 1 h at room temperature.

Protein bands were visualized using enhanced chemiluminescence and captured with a digital imaging system. Band intensities were quantified by densitometric analysis and normalized to β-actin, which served as the loading control.

To specifically test whether miR-155-5p directly targets Kdm5a and whether CTBP1 modulates this interaction, we performed a luciferase reporter assay in HEK293 cells (ATCC). This in vitro system was independent of the animal model and is therefore described separately. Cells were maintained in DMEM supplemented with 10% FBS. HEK293 cells were co-transfected with a pmirGLO reporter containing the 3′UTR of rat Kdm5a (with the predicted miR-155-5p binding site), a miR-155-5p mimic or inhibitor, and/or a CTBP1 overexpression plasmid using Lipofectamine 3,000 (Thermo Fisher). After 48 h, firefly and Renilla luciferase activities were measured using the Dual-Luciferase Reporter Assay System (Promega). The ratio of firefly/Renilla luciferase was normalized to control transfections to determine miR-155-5p–Kdm5a interactions.

For biochemical assays with low-abundance markers (e.g., oxidative stress and ferroptosis indices), whole-brain homogenates were used to obtain sufficient tissue mass for accurate measurement. For histopathological, immunohistochemical, and Western blot analyses, cortical tissue was selected because it represents one of the primary targets of bilirubin neurotoxicity and showed the most pronounced neuropathological changes in our pilot work. In selected assays, the hippocampus and basal ganglia were also analyzed because these regions are known to be vulnerable to bilirubin toxicity and are clinically relevant to kernicterus (auditory and motor sequelae). This tiered approach allowed us to balance analytical sensitivity with region-specific vulnerability.

All histopathological analyses were conducted on brains collected at postnatal day 10 (PND10) to capture acute neuropathological changes before behavioral testing on PND11. For histology, at 72 h after bilirubin injection a subset of pups (n = 4 per group) was perfused transcardially with phosphate-buffered saline followed by 4% paraformaldehyde. Brains were post-fixed, embedded in paraffin, and coronal sections (5 μm) were cut at the level of the hippocampus and basal ganglia. Sections were stained with hematoxylin and eosin (H&E) following standard protocols.

An experienced neuropathologist blinded to group examined the sections under light microscopy. We qualitatively assessed neuronal morphology (pyknosis, necrosis), presence of bilirubin pigment (yellow discoloration in tissue), gliosis, and any hemorrhages. We also performed Nissl staining on adjacent sections to evaluate neuronal density in hippocampal CA regions and cortex. For apoptosis detection in situ, a TUNEL (terminal deoxynucleotidyl transferase dUTP nick-end labeling) assay was performed on some sections (ApopTag kit, Millipore). TUNEL-positive nuclei were counted in five random high-power fields (HPF) per section to quantify apoptotic cell percentage.

For histopathological quantification, three coronal sections per brain were analyzed. For each section, five random high-power fields (400 × magnification) were examined. Normal-appearing cortical neurons were defined as neurons with intact morphology, visible nuclei, and preserved Nissl substance. Neuronal vacuolation was assessed using a semi-quantitative scoring system (0–3) based on the proportion of neurons exhibiting cytoplasmic vacuolization. GFAP-positive astrocytes were scored using a 0–3 scale reflecting the intensity and distribution of astrocytic activation. All analyses were performed by a neuropathologist blinded to experimental group allocation.

To examine the temporal pattern of Cyclin D protein expression following bilirubin-induced injury, an additional independent cohort of neonatal rats was used for time-course immunohistochemical analysis. Animals were subjected to the same bilirubin injection protocol described above and were sacrificed at 1, 7, 14, and 28 days post-injury. Brain tissues were processed using the same fixation and sectioning procedures as described for the primary cohort. Immunohistochemical staining for Cyclin D1, Cyclin D2, and Cyclin D3 was performed on cortical sections. For each animal, three coronal sections were analyzed and five random high-power fields per section were evaluated under 400 × magnification. The number of Cyclin-positive cells was assessed semi-quantitatively by an investigator blinded to group allocation to evaluate the temporal progression of cell-cycle protein expression following bilirubin exposure.

All data were tested for normality using the Shapiro–Wilk test and for homogeneity of variance using Levene’s test. Parametric data were analyzed using one-way ANOVA followed by Tukey’s post-hoc test. Non-parametric data were analyzed using Kruskal–Wallis tests followed by Dunn’s post-hoc test with Bonferroni correction. For time-course analyses (e.g., serum bilirubin levels over time), repeated-measures ANOVA (or mixed-effects models where appropriate) was employed to account for within-subject correlations. Statistical significance was set at p < 0.05 after correction for multiple comparisons.

All animal procedures were approved by the Institutional Animal Care and Use Committee, and conducted in strict accordance with National Research Council guidelines for the care of laboratory animals. The study was also approved by the institutional research committee bearing Approval No. FMU/SHJ/2024/797.

Because the present study measured both serum and brain tissue bilirubin using different analytical methods than those used in our previous report, the absolute values and temporal kinetics are not directly comparable. However, the overall trend of bilirubin elevation following injection and subsequent reduction with DHA treatment remained consistent with the neuroprotective effects previously reported.

All bilirubin-injected rats developed visible jaundice within 12 h, confirming the induction of hyperbilirubinemia. Untreated BE rats appeared lethargic with reduced suckling and episodes of opisthotonic posturing by 48 h, indicative of acute bilirubin neurotoxicity. In contrast, BE rats treated with DHA maintained relatively normal feeding behavior and less frequent opisthotonus. No adverse effects of DHA were observed. Final body weights at day 11 were slightly lower in the BE group (mean 18.3 ± 1.2 g) compared to Control (19.5 ± 1.1 g, p < 0.05), whereas the DHA-treated group had intermediate weights (19.0 ± 1.3 g, not significant vs. Control).

All bilirubin-injected rats developed visible jaundice within 12 h after bilirubin administration, confirming successful induction of hyperbilirubinemia. By 48 h post-injection, untreated BE rats displayed neurological signs consistent with acute bilirubin neurotoxicity, including lethargy, reduced suckling activity, and episodes of opisthotonic posturing.

In contrast, rats in the BE + DHA group exhibited milder clinical manifestations, maintaining relatively normal feeding behavior and demonstrating fewer episodes of opisthotonus. These clinical observations corresponded with the quantitative neurological scoring results described below (Figure 2). No adverse effects related to DHA administration were observed.

Final body weights measured on postnatal day 11 (PND11) were slightly lower in the BE group (18.3 ± 1.2 g) compared with Control animals (19.5 ± 1.1 g, p < 0.05), whereas the DHA-treated group showed intermediate values (19.0 ± 1.3 g), which were not significantly different from controls.

To quantitatively evaluate neurological impairment, neurological status was assessed using the scoring system described in the Methods section. In the BE group, half of the animals (8/16, 50%) exhibited neurological scores ≥ 2, indicating moderate neurological impairment. In contrast, animals in the BE + DHA group predominantly displayed lower scores (0–1), consistent with milder neurological dysfunction and partial neuroprotection following DHA treatment (Figure 2).

Distribution of neurological scores among animals in each group at 72 h post-bilirubin injection. Although the scoring system allows a maximum value of 8, the observed scores in this experiment ranged from 0 to 3, indicating predominantly mild to moderate neurological impairment.

Neurobehavioral testing revealed significant sensorimotor impairment in untreated BE pups compared with controls.

For the righting reflex test (PND9), BE animals showed markedly prolonged response times (8.7 ± 2.1 s) relative to Control animals (3.2 ± 0.8 s, p < 0.01). DHA treatment significantly improved performance (4.5 ± 1.2 s, p < 0.05 vs. BE), approaching control levels.

Similarly, in the negative geotaxis test (PND11), BE pups required significantly more time to reorient on the inclined plane (15.4 ± 4.5 s) compared with Control animals (6.1 ± 1.9 s, p < 0.01). DHA treatment improved this response (8.0 ± 2.3 s, p < 0.05 vs. BE), indicating partial recovery of vestibular-motor coordination.

These findings demonstrate that bilirubin exposure impaired neurobehavioral development, while DHA treatment significantly improved behavioral performance (Figure 3).

Serum bilirubin levels were measured at 24, 48, and 72 h after bilirubin injection (Figure 4a and Table 1). Baseline bilirubin concentrations were similar across all groups prior to injection (approximately 5.0 μmol/L).

In the BE group, serum bilirubin increased markedly to 15.0 ± 1.3 μmol/L at 24 h, followed by a gradual decline to 12.0 ± 1.1 μmol/L at 48 h and 10.1 ± 1.0 μmol/L at 72 h. In contrast, the BE + DHA group showed a similar peak at 24 h (14.0 ± 1.5 μmol/L) but demonstrated a more rapid decline thereafter, reaching 8.0 ± 0.9 μmol/L at 48 h and 5.2 ± 0.8 μmol/L at 72 h.

By 72 h, serum bilirubin concentrations in the BE + DHA group were significantly lower than in the untreated BE group (p < 0.01).

Brain bilirubin accumulation measured at 72 h is shown in Figure 4b. Control animals exhibited minimal brain bilirubin levels (0.3 ± 0.1 μg/g tissue), whereas the BE group showed substantially elevated levels (8.0 ± 1.2 μg/g tissue). DHA treatment significantly reduced brain bilirubin accumulation to 3.1 ± 0.8 μg/g tissue (p < 0.01 vs. BE).

These findings indicate that bilirubin exposure resulted in marked increases in circulating and brain bilirubin levels, whereas DHA treatment was associated with lower bilirubin levels at later time points (Figure 4).

DHA-treated animals showed a faster decline in serum bilirubin levels over time compared with untreated BE animals.

While all groups started with similar baseline bilirubin levels (5.0 μmol/L), the BE group exhibited a sharp increase, peaking at 15.0 μmol/L at 24 h and gradually decreasing to 10.1 μmol/L by 72 h. In contrast, DHA-treated rats (BE + DHA) showed a faster reduction in bilirubin levels, declining to near-normal levels (5.2 μmol/L) by 72 h. This observation suggests that DHA may facilitate bilirubin clearance or redistribution in this experimental setting (Table 1).

Serum neuron-specific enolase (NSE) levels were measured at 72 h after bilirubin injection (Figure 4c and Table 2). NSE concentrations were significantly higher in the BE group (15.3 ± 2.4 ng/mL) compared with the Control group (5.2 ± 1.1 ng/mL, p < 0.001).

In the BE + DHA group, NSE levels were significantly lower than in the BE group (7.4 ± 1.5 ng/mL, p < 0.001) but remained slightly higher than Control values (p < 0.05).

Apoptotic cell death in cortical tissue was evaluated using TUNEL staining (Figure 4d). Control brains showed a low proportion of TUNEL-positive cells (∼5%). In contrast, the BE group exhibited a significantly higher percentage of apoptotic cells (25 ± 4%). DHA treatment significantly reduced the proportion of TUNEL-positive cells to 10 ± 3% (p < 0.01 vs. BE).

Immunohistochemical analysis of apoptosis-related proteins showed increased Bax and cleaved caspase-3 staining and reduced Bcl-2 expression in the BE group compared with Controls. In the BE + DHA group, Bax and cleaved caspase-3 staining were reduced, whereas Bcl-2 expression was increased relative to the BE group.

Bilirubin-induced encephalopathy leads to dysregulation of key cell cycle proteins in the neonatal brain. Western blot analysis revealed significant upregulation of CyclinD1, CDK4, and CyclinE1 in the BE group compared to controls, indicating aberrant reactivation of the G1/S cell cycle transition in post-mitotic neurons, a process linked to apoptosis and neurodegeneration. Expression of cell-cycle–related markers differed significantly across groups (Figure 5). Compared with controls, BE cortex showed increased Cyclin D1, CDK4, and Cyclin E1 levels and reduced P21, suggesting aberrant cell-cycle re-entry. DHA treatment significantly reduced Cyclin D1, CDK4, and Cyclin E1 expression compared with the BE group; however, only CDK4 approached control levels, while Cyclin E1 remained moderately higher. P21 expression increased relative to BE but did not fully return to control values.

To examine the temporal progression of Cyclin D family protein expression following bilirubin exposure, an independent cohort of neonatal rats was analyzed at multiple time points after injury (1, 7, 14, and 28 days post-injection) (Figure 6).

At 1 day post-injury, only a small number of Cyclin D–positive cells were observed in the cortex. The number and staining intensity of positive cells increased at 7 days, became more pronounced at 14 days, and remained elevated at 28 days, suggesting sustained activation of cell-cycle–related pathways during the subacute and chronic phases following bilirubin exposure.

H&E-stained sections from BE pups at PND10 revealed classic kernicterus features: neuronal shrinkage, vacuolation and yellow pigment deposition. Quantitatively, the BE group showed a 32 ± 5% reduction in normal-appearing cortical neurons per high-power field compared to controls (p < 0.01). DHA treatment preserved cortical neuron counts (89 ± 6% of control, p < 0.05 vs. BE) and reduced vacuolation scores by ∼50%.

Nissl staining corroborated these findings: hippocampal CA1 neurons decreased by 28 ± 4% in BE rats, whereas DHA rats showed only a 10% decrease vs. control (p < 0.05).

TUNEL assay results (already partly described in your Figure 4d) showed 25 ± 4% apoptotic cortical cells in BE vs. 10 ± 3% in DHA-treated pups (p < 0.01).

Gliosis (GFAP immunoreactivity) was rated on a 0–3 scale: BE brains scored 2.5 ± 0.3 vs. 1.2 ± 0.2 in DHA-treated pups (p < 0.01).

Bilirubin pigment was detected in 65 ± 8% of BE cortical fields but in only 15 ± 5% of DHA-treated fields (p < 0.01).

Markers of oxidative stress were evaluated in cortical tissue at 72 h post-injection (Tables 2, 3).

Malondialdehyde (MDA), an indicator of lipid peroxidation, was significantly elevated in the BE group (5.6 ± 0.6 nmol/mg) compared with the Control group (2.0 ± 0.4 nmol/mg, p < 0.001) (Table 2). In the BE + DHA group, MDA levels were significantly lower than in the BE group (3.0 ± 0.5 nmol/mg, p < 0.01 vs. BE).

Reactive oxygen species (ROS) levels measured using a DCFDA fluorescence assay were also elevated in the BE group (220 ± 30 arbitrary units) compared with Control animals (100 ± 20 arbitrary units, p < 0.01) (Table 3). ROS levels in the BE + DHA group were lower than in the BE group (120 ± 25 arbitrary units, p < 0.01 vs. BE).

Expression of proteins associated with ferroptosis-related pathways, including glutathione peroxidase 4 (GPX4), prostaglandin-endoperoxide synthase 2 (PTGS2/COX-2), and acyl-CoA synthetase long-chain 4 (ACSL4), was analyzed by Western blot (Table 4). In the BE group, GPX4 expression was reduced (∼0.40 relative to β-actin), whereas COX-2/PTGS2 (∼3.0-fold) and ACSL4 (∼1.8-fold) were increased compared with Control animals. In the BE + DHA group, GPX4 expression was higher than in the BE group (∼0.85 relative expression), while COX-2/PTGS2 and ACSL4 levels were lower than in the BE group (∼1.2- and ∼1.1-fold, respectively) (Table 4).

The expression of ferroptosis-related proteins (GPX4, COX-2/PTGS2, and ACSL4) was evaluated by Western blot to assess the effects of bilirubin exposure and DHA treatment. In the BE group, ferroptotic activity is evident with a marked decrease in GPX4 (0.40) and increased expression of pro-ferroptotic markers COX-2 (3.00) and ACSL4 (1.80), compared to control levels. DHA treatment partially restored GPX4 expression (0.85) and reduced COX-2 and ACSL4 levels to near-control values (1.20 and 1.10, respectively), indicating that DHA suppresses ferroptosis and contributes to neural protection by stabilizing oxidative homeostasis (Table 4).

RT-qPCR analysis demonstrated significant dysregulation of the CTBP1/miR-155-5p/KDM5A pathway in cortical tissue following bilirubin exposure. Compared with the Control group, the BE group showed significant downregulation of CTBP1 (0.52 ± 0.10-fold) and KDM5A (0.60 ± 0.08-fold), whereas miR-155-5p expression increased to 3.0 ± 0.5-fold (p < 0.01).

In the BE + DHA group, miR-155-5p expression decreased to 1.4 ± 0.3-fold, while CTBP1 and KDM5A expression increased to 0.94 ± 0.15-fold and 1.12 ± 0.12 fold, respectively (p < 0.01 vs. BE).

These results indicate that bilirubin exposure disrupted the CTBP1/miR-155-5p/KDM5A regulatory axis, whereas DHA treatment partially restored the expression of CTBP1 and KDM5A while suppressing miR-155-5p upregulation (Figure 7 and Table 5).

To verify whether miR-155-5p directly targets KDM5A, a luciferase reporter assay was performed in HEK293 cells. Transfection with a miR-155-5p mimic significantly reduced luciferase activity (∼40%) compared with controls (p < 0.01). In contrast, inhibition of miR-155-5p increased reporter activity, indicating reduced repression of the KDM5A 3′UTR.

Co-transfection with a CTBP1 overexpression plasmid partially restored luciferase activity, suggesting that CTBP1 modulates the miR-155-5p–KDM5A interaction.

Gene-expression changes are illustrated in Figure 7, while Table 5 provides the quantitative RT-qPCR values. The luciferase reporter assay results are shown in Figure 8, and Table 6 summarizes the overall behavioral, biochemical, and molecular outcomes across experimental groups.

Table 6 provides a comprehensive comparison of behavioral, biochemical, and molecular parameters among the study groups. The BE group exhibited significant impairments, including delayed reflexes (righting reflex: 8.18 ± 1.97 s), elevated neuronal injury markers (NSE: 15.96 ± 2.77 ng/mL), high brain bilirubin (8.22 ± 1.05 μg/g), oxidative stress (MDA: 5.73 ± 0.58 nmol/mg), and dysregulation of the CTBP1/miR-155-5p/KDM5A axis. DHA treatment markedly improved these parameters, restoring reflex performance, lowering NSE and MDA, reducing brain bilirubin, and normalizing gene expression profiles. All differences were statistically significant (p < 0.0001), confirming DHA’s robust neuroprotective effects in bilirubin-induced encephalopathy (Table 6).

Our results showed that DHA intervention substantially improved neurobehavioral outcomes and reduced neuropathology in neonatal rats with bilirubin-induced encephalopathy. These benefits were associated with: (1) enhanced clearance of bilirubin from serum and brain, (2) reduction of neuronal injury markers and apoptosis, (3) attenuation of oxidative stress and ferroptosis, and (4) modulation of the CTBP1–miR-155–KDM5A molecular pathway toward a neuroprotective profile. No significant differences were observed between the DHA-treated group and controls in many measures (e.g., by 72 h, DHA group’s bilirubin, ROS, CTBP1/KDM5A levels were statistically indistinguishable from controls), underscoring DHA’s efficacy in mitigating the damage.

While our study provides important insights, several limitations warrant consideration. First, although our animal model is well established, it may not fully recapitulate the timing and complexity of human bilirubin encephalopathy, so translational caution is needed. Second, our experiments were limited to 72 h and did not assess long-term neurodevelopmental outcomes, phototherapy or exchange transfusion interactions, or alternate DHA dosing regimens. Third, our mechanistic findings on the CTBP1/miR-155-5p/KDM5A pathway are associative rather than causative and require validation with genetic or pharmacologic manipulation. Finally, our biochemical measures capture overall oxidative stress but not cell-type specificity. Despite these limitations, our findings offer a solid foundation for future studies and support the potential of DHA as an adjunctive therapy in neonatal bilirubin encephalopathy. Although both male and female pups were included, our study was not powered to evaluate sex differences; future studies will be needed to determine whether DHA’s effects vary by sex. Because our study did not include direct manipulation of CTBP1, miR-155-5p, or KDM5A, we cannot confirm a causal relationship. These findings should be interpreted as associations, and future studies using knockdown, overexpression, or pharmacologic modulation will be necessary to establish causality.

Future studies should clarify DHA’s effects on microglial activation and miRNA regulation (including miR-155, miR-122, and miR-125b) and test whether combining DHA with ferroptosis inhibitors yields additive neuroprotection. Translational work using preterm or genetically jaundiced models, as well as evaluating brain-targeted DHA formulations such as lysophosphatidylcholine, could further optimize its therapeutic potential. Collectively, these investigations will refine DHA-based strategies for preventing neonatal bilirubin encephalopathy.

Our findings suggest that DHA, already used as a nutritional supplement in premature infants, could be repurposed as an adjunct neuroprotective therapy for neonates at risk of bilirubin-induced brain injury. By modulating the CTBP1/miR-155-5p/KDM5A pathway and reducing oxidative stress, DHA may promote cortical repair and attenuate progression to acute bilirubin encephalopathy. Given its established safety profile, enteral DHA administration alongside conventional treatments warrants clinical investigation to improve neurodevelopmental outcomes and reduce kernicterus-related disabilities.