Integrative multi-omics profiling reveals an association between lipid metabolism dysregulation and bortezomib-induced cerebral hemorrhage in zebrafish

Abstract

Bortezomib (BTZ), a first-generation proteasome inhibitor, serves as a cornerstone chemotherapeutic agent for Multiple Myeloma (MM), with its clinical application in oncology expanding rapidly in recent years. Using vascular-specific transgenic zebrafish models, we systematically evaluated BTZ-induced toxicity at 4–5 mg/L gradient concentrations and its underlying molecular mechanisms. We found 72-hour BTZ exposure triggered significant intracerebral hemorrhage in zebrafish larvae with concentration-dependent severity, accompanied by cerebrovascular disruption, tight junction gene downregulation, lipid metabolism disorder, ROS accumulation and antioxidant enzyme response, and abnormal nitric oxide activation. This study first reveals that BTZ-induced intracerebral hemorrhage is closely associated with lipid metabolism dysregulation, highlighting lipid disturbance as a potential contributor to cerebrovascular injury.

Supplementary Information

The online version contains supplementary material available at https://doi.org/10.1186/s12987-026-00867-6.

Introduction

As a first-generation proteasome inhibitor, bortezomib (BTZ) primarily acts by reversibly inhibiting the chymotrypsin-like activity of the 26S proteasome in mammalian cells, thereby blocking the activation of key signalling pathways such as nuclear factor κB (NF-κB) and other key signalling pathways, ultimately inducing apoptosis in tumour cells [1–3]. Since its approval by the US Food and Drug Administration (FDA) in 2003 for the treatment of multiple myeloma (MM) and mantle cell lymphoma (MCL) [4–6], BTZ has become a cornerstone drug in chemotherapy regimens for haematological malignancies, significantly improving patient prognosis and prolonging survival [7, 8]. With the gradual expansion of clinical indications and its exploratory use in oncology, the frequency of BTZ use in patients has increased year on year, and its potential safety risks in developing populations have increasingly become a focus of attention in both clinical practice and research.

The clinical use of BTZ is associated with a variety of adverse reactions, of which peripheral neuropathy (PN) is the most common; approximately 30% to 60% of patients experience symptoms of neurotoxicity to varying degrees during treatment, which in severe cases may lead to treatment discontinuation [9–11]. Furthermore, clinical studies have demonstrated that BTZ can cause abnormal changes in patients’ lipid profiles [12, 13], and these changes are associated with an increased risk of cardiovascular events. The cardiotoxicity of BTZ is associated with abnormalities in the regulation of endothelial nitric oxide synthase (eNOS) activity and nitric oxide (NO) levels [14], which can indirectly affect vascular function by increasing apoptosis in endothelial progenitor cells and reducing their proliferation and migration capacity [15]. However, research on the effects of BTZ on the developing vascular system remains scarce.

Due to its unique advantages—transparent embryos that develop externally and remain optically clear throughout embryogenesis, together with a genetic homology to humans of up to 87%—the zebrafish (Danio rerio) has become a widely used model organism in the fields of developmental toxicology and drug safety assessment [16–18]. As the organ system that forms earliest and performs physiological functions in vertebrates, the cardiovascular system plays a crucial regulatory role not only in embryonic survival but also in the formation and functional maturation of subsequent organ systems [19–22]. In recent years, the application of transgenic zebrafish lines (such as the vascular endothelial-specific marker Tg(kdrl:mCherry) and the erythrocyte-specific marker Tg(gata1:DsRed)), has made it possible to observe in real time and dynamically monitor the effects of drugs on the integrity of the cerebral vascular network and erythrocyte distribution in vivo, providing intuitive experimental evidence for the precise identification of vascular injury phenotypes such as intracerebral haemorrhage (ICH) [23–25]. Previous studies have confirmed that statin-class lipid-lowering drugs can induce ICH in zebrafish larvae [26], and that this effect is closely associated with the inhibition of cerebral vascular development and lipid metabolism disorders, suggesting that drugs targeting intracellular metabolic pathways may pose a potential risk of damage to the vascular system during development [27, 28]. However, there are currently no reports, either domestically or internationally, regarding whether BTZ, a proteasome inhibitor, exhibits similar developmental vascular toxicity.

The cytotoxic mechanism of BTZ is multifaceted; in addition to inhibiting the proteasome-mediated protein degradation pathway, it also plays a role in regulating various biological processes, including oxidative stress and endoplasmic reticulum stress [29–31]. Clinical studies have shown that BTZ treatment can lead to abnormal fluctuations in lipid parameters such as triglycerides (TG), low-density lipoprotein cholesterol (LDL-C) in patients [32–35]. Based on the above research background, we propose the following hypothesis: BTZ may disrupt the normal developmental process of cerebral blood vessels during development by interfering with lipid metabolic homeostasis [36], thereby contributing to the pathogenesis of ICH in zebrafish embryos.

In this study, zebrafish embryos at 5–6 hours post-fertilisation (hpf) were acutely exposed to BTZ until 72 hpf, utilising the Tg(fli:GFP) and Tg(gata1:DsRed) double-transgenic zebrafish lines to observe cerebral vascular morphology and red blood cell distribution characteristics, respectively, in order to elucidate the mechanisms underlying ICH and its intrinsic association with cerebral vascular damage; Concurrently, non-targeted metabolomics techniques were employed to systematically analyse changes in the lipid metabolic profile of zebrafish larvae following BTZ exposure. The aim was to characterize the phenotypic and multi-omic features associated with of BTZ-induced ICH from a developmental biological perspective, thereby providing new experimental evidence and research insights for the safety assessment of clinical use and the prevention and control of adverse reactions.

Materials and methods

Transgenic zebrafish lines and imaging protocols

All zebrafish experiments were conducted in accordance with the guidelines of the Laboratory Animal Welfare and Ethics Committee of Jinggangshan University, which approved this study (approval certificate available upon request). All experimental procedures adhered to the 3 R principles (Replacement, Reduction, Refinement) to minimize animal use and ensure humane endpoints. Wild-type AB and transgenic lines—Tg(gata1:DsRed) (erythrocyte-specific), Tg(kdrl:mCherry) and Tg(fli:GFP) (vascular endothelium-specific), Tg(elavl3:EGPF)—were maintained at 28 °C under a 14:10 h light/dark cycle in an automated circulation system (Haisheng, China). Healthy embryos (5–6 hpf) were collected after natural mating (2 females:1–2 males) and transferred to embryo medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl₂, 0.33 mM MgSO₄). For display purposes, EGFP and GFP signals were pseudocolored cyan, while DsRed and mCherry signals were pseudocolored magenta. Color scale bars on each image map the original grayscale intensity to the respective lookup table (LUT), with lighter shades indicating lower intensity and darker shades indicating higher intensity. All pseudocoloring was applied identically across experimental groups.

Chemicals and reagents

Bortezomib (CAS 179,324-69–7, HPLC grade, ≥98% purity) was purchased from Macklin Biochemical (Shanghai, China). Prepare the working solution (10 mg/mL) in 100% dimethyl sulfoxide (DMSO), use immediately after preparation. N-Phenylthiourea (PTU; CAS 103–85-5) was purchased from Sigma-Aldrich (USA). 3,3’-Dimethoxybenzidine (≥97% purity, CAS 119–90-4) was purchased from Aladdin Reagent (Shanghai, China).

BTZ exposure and experimental design

BTZ was dissolved in dimethyl sulfoxide (DMSO) to prepare a working solution at a concentration of 10 mg/mL, which was used immediately upon preparation. Subsequently, 0.003% 1-phenyl-2-thiourea (PTU) was supplemented into the embryo medium to inhibit melanogenesis, and PTU was exclusively applied in experiments requiring fluorescence imaging and staining. Zebrafish embryos were exposed to BTZ at gradient concentrations ranging from 4 mg/L to 5 mg/L. After 72 h of continuous BTZ exposure, phenotypic observations and image acquisition were performed using a Leica M205 FA stereomicroscope (Leica Microsystems, Wetzlar, Germany). All experiments for each concentration treatment group were conducted in triplicate with three independent biological replicates.

In clinical settings, BTZ is administered intravenously at 1.3 mg/m², yielding peak plasma concentrations (Cmax) of approximately 100–200 ng/mL (0.1–0.2 mg/L) in patients with multiple myeloma [37]. Due to the dilutional effect of the aqueous environment and drug absorption barriers across the skin and gills in the zebrafish immersion model, the nominal exposure concentration of 4–5 mg/L is substantially higher than clinical plasma levels. While the actual internal tissue concentration achieved in zebrafish larvae has not been directly measured in this study, the observed concentration-dependent biological responses—ICH, vascular disruption, and lipid metabolism dysregulation—suggest that the effective exposure level at target tissues is within a pharmacologically relevant range. Notably, BTZ is known to accumulate in lipid-rich tissues and endothelial cells, and repeated clinical dosing may result in cumulative tissue exposure exceeding transient plasma Cmax. Therefore, the concentrations used here may partially model aspects of prolonged or repeated clinical exposure scenarios. Nevertheless, direct quantitative extrapolation of these findings to clinical dosing regimens should be approached with caution, and future studies incorporating pharmacokinetic modeling and internal dosimetry in the zebrafish model would strengthen translational interpretation. This concentration range can produce observable biological effects in zebrafish models without causing non-specific toxicity.

Behavioral analysis

To evaluate the neurobehavioral effects of BTZ exposure on zebrafish embryos, we performed a locomotor behavioral assessment assay. Six larvae were randomly selected from each concentration treatment group and individually placed into separate wells of a 24-well plate, with 1 mL of embryo medium added to each well. The locomotor activity of hatched larvae was monitored using the DanioVision Observation System (Noldus Information Technology, Wageningen, the Netherlands). A 30-minute light-dark stimulation paradigm was adopted in the experiment with an alternating light-dark cycle protocol: after a 10-minute acclimation period for larvae to adapt to the system environment, the recording program was initiated with a repeating cycle consisting of 5 minutes of continuous light illumination followed by 5 minutes of dark phase. This cycle was repeated three times, with a total recording duration of 30 minutes. All experiments for each treatment group were conducted in triplicate with three independent biological replicates.

Chemical staining

After 72 hours of drug treatment, wash the larvae three times with pure water, add the o-anisidine staining solution, and incubate in the dark at 37.5 °C for 30 minutes. After incubation, use a Leica M205 FA stereomicroscope (Germany) for imaging observation.The experiment was repeated three times.

DCFH-DA probe (Nanjing Jiancheng Bioengineering Institute, Nanjing, Jiangsu, China) was used for the detection of reactive oxygen species (ROS) in zebrafish larvae. Larvae at 72 hpf were incubated with 2 μM DCFH-DA for 30 min at 28.5 °C in the dark, washed, and imaged by fluorescence microscopy. For display, DCFH-DA (green) fluorescence was pseudocolored cyan, following the same LUT mapping as described for transgenic lines.

After 72 h of BTZ exposure, zebrafish larvae were collected and washed three times with ice-cold phosphate buffered saline (PBS). Subsequently, Acridine Orange (AO) reagent (Beyotime Biotechnology, Shanghai, China) was prepared for the detection of cellular apoptosis in brain tissue. The samples were stained with the freshly prepared AO working solution for 20–30 minutes in the dark. After staining, excess residual dye on the body surface of larvae was thoroughly removed by rinsing with fresh PBS. Finally, 20 zebrafish larvae from each group were immobilized in 1% low-melting point agarose, and fluorescence imaging was performed using a Leica TCS SP8 laser scanning confocal microscope. The imaging magnification was kept consistent across all experimental groups. For display, acridine orange (green) fluorescence was pseudocolored cyan, following the same LUT mapping as described for transgenic lines.

Cerebrovascular changes following cerebral haemorrhage in zebrafish

Tg(gata1:DsRed;fli:GPF) double-transgenic line was used to label red blood cells and vascular endothelial cells. Zebrafish embryos carrying the dual transgenic line were obtained. Following 72 hours of drug treatment, the fish were anaesthetised with 0.02% tricaine. Under a Zeiss fluorescence microscope, juvenile zebrafish exhibiting both fluorescence signals were selected, fixed using 1% low-melting-point agarose, and imaged using a Leica confocal microscope. The AngioTool software was used to analyse and process the cerebral vasculature of the zebrafish larvae. The experiment was repeated three times.

Untargeted metabolomics analyses

After 72 h of continuous BTZ exposure, approximately 400 zebrafish larvae were collected from the high-concentration BTZ treatment group and the control group respectively for liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis, with three independent biological replicates set for each group. Raw LC-MS data were subjected to baseline filtering, peak detection, integration, retention time alignment, and normalization using XCMS software (Nonlinear Dynamics, Newcastle, UK). Features present in fewer than 50% of samples within any group were excluded to reduce noise from sporadically detected ions. Missing values were imputed using the k-nearest neighbors (kNN) algorithm (k = 10) implemented in the preprocessCore R package. Data were log2-transformed and Pareto-scaled prior to multivariate analysis. Compound identification was performed by matching accurate mass, MS/MS fragmentation patterns, and retention time against public databases, including MassBank, Fiehn HILIC, Respect, and the Global Natural Products Social Molecular Networking (GNPS), with a mass tolerance of ± 10 ppm. Partial least squares-discriminant analysis (PLS-DA) and principal component analysis (PCA) were performed using MetaboAnalyst (version 5.0); PLS-DA model validation was conducted using 10-fold cross-validation and 2000 permutation tests, with model quality assessed by R² Y and Q [2] values. Metabolites with a fold change ≥ 1.5 and a Benjamini-Hochberg adjusted p-value < 0.05 (one-way ANOVA with Fisher’s LSD post-hoc test, SPSS 27.0) were defined as differentially expressed metabolites (DEMs). Pathway enrichment analysis of DEMs was performed using the hypergeometric test against the KEGG and GO databases, with significantly enriched pathways identified at a Benjamini-Hochberg adjusted p-value < 0.05. The raw data can be obtained from the public repository (project number: MTBLS14230).

Nitric oxide (NO) content assay and measurement of enzyme activity

Thirty larvae per biological replicate were homogenized in ice-cold lysis buffer, centrifuged (12,000 × g, 15 min, 4 °C), and total NO was measured via nitrate reductase method (Nitric Oxide Assay Kit, BC1475, Solarbio) [38, 39]. Absorbance at 550 nm was read on a SpectraMax iD3 plate reader. NO content was normalized to total protein (BCA kit, Beyotime). For enzyme and biomarker assays, approximately 100 larvae per group were homogenized in 400 μL normal saline, centrifuged (15,000 × g, 15 min, 4 °C), and supernatants were analyzed using commercial kits for MDA, CAT, SOD, HDL-C, LDL-C, T-CHO, and TG. All assays used three biological replicates with three technical replicates each.

Transcriptome analysis

To evaluate the changes in gene expression of zebrafish embryos following BTZ exposure, 600 embryos at 72 hpf were divided into the control group and the BTZ treatment group. Transcriptomic profiling was performed via high-throughput RNA sequencing (RNA-seq) on the DNBSEQ platform. To ensure the reliability of sequencing data, raw sequencing reads with low quality (Phred quality score < 20), adapter contamination, or an excessive proportion of unknown bases (N content > 5%) were discarded. On average, 44.40 million and 44.20 million high-quality clean reads were obtained for each sample in the control group and BTZ treatment group, respectively, ultimately yielding 6.64 Gb of valid sequencing data per sample. Clean reads were aligned to the zebrafish reference genome (GRCz11) using Bowtie2 software, with an average mapping rate of 64.78%, and a total of 25,608 expressed genes were detected across all samples.

Raw FASTQ files were preprocessed using SOAPnuke (version 2.3, filtering parameters: -l 20; -q 0.4; -n 0.01) to remove low-quality reads (Phred quality score < 20), adapter-contaminated sequences, and reads with >5% unknown bases (N). For differential gene expression analysis, clean reads were aligned to the reference gene set using HISAT2 (version 2.1.4, alignment parameters: –sensitive –no-discordant –no-mixed; -I 1; -X 1000; -p 8). Clean reads were also aligned to the reference gene set using Bowtie2 (version 2.3.4.3). Gene expression levels were quantified using RSEM (version 1.3.1) software, and a clustering heatmap of gene expression profiles across different samples was plotted using the pheatmap package (version 1.0.8) in R. Differential gene expression analysis was performed using DESeq2 (version 1.4.5), and genes with |log2 fold change| ≥1 and a Benjamini-Hochberg adjusted p-value (Q value) ≤0.05 were defined as significantly differentially expressed genes (DEGs). Genes with a mean normalized count < 10 across all samples were excluded from differential analysis to reduce noise from low-expression transcripts. For functional enrichment analysis, the hypergeometric test was performed using the Phyper function in R, with all expressed genes as the background set. Significantly enriched GO terms and KEGG pathways were identified at a Benjamini-Hochberg adjusted p-value ≤ 0.05, and a minimum of 5 overlapping genes per term was required.

To further explore the gene functions associated with the observed phenotypic changes, hierarchical clustering analysis of DEGs was performed based on the hypergeometric test using the Phyper function (https://en.wikipedia.org/wiki/Hypergeometric_distribution), and Pearson correlation coefficients were calculated for pairwise comparisons between all samples. Functional enrichment analysis was conducted using the Gene Ontology (GO; http://geneontology.org/) and Kyoto Encyclopedia of Genes and Genomes (KEGG; https://www.kegg.jp/) databases. Significantly enriched GO terms and KEGG pathways were identified with a threshold of Q value ≤ 0.05. The raw data can be obtained from the public repository (project number: GSE327263).

Real-time-quantitative polymerase chain reaction (RT-qPCR) analysis of gene expression

Sixty larvae per group were rinsed with ice-cold PBS. Caudal tissue (posterior one-third) was dissected, and total RNA was extracted using TransZol UP Reagent (TransGen Biotech). RNA purity and concentration were assessed by NanoDrop One (A260/A280 and A260/A230). cDNA was synthesized using TransScript One-Step gDNA Removal and cDNA Synthesis SuperMix (TransGen Biotech). RT-qPCR was performed with PerfectStart Green qPCR Supermix on a StepOne Plus system (Applied Biosystems), with the following cycling: 95 °C for 10 min; 40 cycles of 95 °C for 30 s, 60 °C for 30 s. β-actin served as the internal reference; relative expression was calculated by the 2^(-ΔΔCt) method. Primers are listed in Supplementary Table S1.

Statistical analysis

All statistical analyses were performed using GraphPad Prism 9.5.1 version. All image data measurements are done using ImageJ 1.53e. The Student’s One-way ANOVA followed by LSD post-hoc test was used to determine statistical significance, normality was tested using the Shapiro-Wilk test, homogeneity of variance using Bartlett’s test. All data represents the mean ± S.D. standard deviation * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001, **** p ≤ 0.0001.

Results

Toxicity of BTZ to zebrafish larvae after 72 hours of exposure

Bright-field microscopy showed that 72 h of BTZ exposure induced severe morphological abnormalities in zebrafish larvae, including intracranial hemorrhage (ICH), brain injury (BI), pericardial edema (PE), and yolk sac edema (YSE), while control larvae remained normal (Fig. 1A). Quantitative analysis revealed a concentration-dependent increase in the incidence of all four malformation subtypes, with the highest rates observed in the 5 mg/L treatment group (Fig. 1B).

Fig. 1.

Toxic effects of bortezomib (BTZ) on zebrafish larvae after 72 h exposure.(A) Representative bright-field images of zebrafish larvae at 72 hpf following continuous BTZ exposure (4.0, 4.5, and 5.0 mg/L). Red arrowheads indicate intracerebral hemorrhage (ICH). Scale bar, 50 μm. (B) Quantification of the four malformation subtypes. (C) Median lethal concentration (LD50) of zebrafish embryos. (D) Hatching rate. (E) Heart rate. (F) Representative o-dianisidine hemoglobin staining of 72-hpf larvae. Red arrowheads indicate multifocal intracranial hemoglobin deposits absent in controls. Scale bar, 50 μm. n = 3 independent biological replicates (20 larvae per replicate). (G) Ratio of larvae with ICH alone and those with ICH plus other malformations in each group. (H) ICH severity grading by hemorrhagic brain area proportion: normal (C0, 0%), slight (C1, <10%), moderate (C2, 10–20%), severe (C3, >20%). Grading criteria are provided in Supplementary Figure S1. n = 3 independent biological replicates (20 larvae per replicate). Data are presented as the mean ± S.D. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

Acute toxicity assays demonstrated time- and concentration-dependent lethality, with LD₅₀ values of 15.0 mg/L (24 hpf), 12.4 mg/L (48 hpf), and 9.9 mg/L (72 hpf), indicating enhanced toxicity with prolonged exposure (Fig. 1C). Concurrently, BTZ exposure significantly reduced embryonic hatchability in a concentration-dependent manner (Fig. 1D). Cardiotoxicity was confirmed by a concentration-dependent decrease in larval heart rate (Fig. 1E). To characterize ICH in detail, we labeled larval fish hemoglobin using o-Diazetidine, which revealed that BTZ-induced ICH exhibited a multifocal and patchy hemorrhagic pattern intracranially, which was absent in controls (Fig. 1F). Chemical exposure-induced ICH during the early developmental stages of zebrafish suggests the presence of vascular developmental toxicity [40]. New blood vessels consist solely of endothelial cells and lack pericytic support; their fragile nature makes them highly susceptible. Therefore, the effects of BTZ-induced ICH on aquatic organisms require further clarification.

Collectively, these results demonstrate that BTZ exerts potent concentration- and time-dependent toxic effects on zebrafish embryos, characterized by elevated malformations, increased mortality, impaired hatching, bradycardia, and intracranial hemorrhage.

BTZ causes neuronal loss and impairs locomotor activity in zebrafish larvae

To quantify the severity of ICH in embryos, we categorized the proportion of hemorrhagic area in brain tissue into four grades: normal (C0), mild (C1, less than 10%), moderate (C2, between 10 and 20%), and severe (C3, more than 20%, more details in Supplementary Figure S1). BTZ exposure led to a concentration-dependent increase in ICH severity, with 5 mg/L BTZ inducing the highest proportion of moderate (21%) and severe (41%) hemorrhage, while control larvae remained unaffected (Fig. 1H). We further quantified the proportion of larvae exhibiting isolated ICH versus ICH combined with other malformations. All BTZ-treated groups showed a high incidence of ICH, with a notable fraction of larvae displaying only the ICH phenotype, indicating that BTZ at 4–5 mg/L induces intracerebral hemorrhage even in the absence of severe systemic malformations (Fig. 1G).

To directly determine whether the observed ICH causes neuronal damage, we imaged the brains of BTZ-treated larvae using the pan-neuronal transgenic line Tg(elavl3:EGFP), which labels all differentiated neurons throughout the central nervous system (Fig. 2A–B). At 72 hpf, control larvae exhibited dense, continuous EGFP fluorescence across the forebrain, midbrain, and hindbrain, consistent with intact neuronal architecture. Morphological observation revealed that BTZ-treated larvae exhibited concentration-dependent disruption of neuronal integrity, characterized by discrete hypointense hollow areas within brain tissue (white arrows, Fig. 2A) and regional loss of neuronal signal (white dashed lines, Fig. 2B), providing direct morphological evidence that BTZ-induced ICH is accompanied by regional neuronal loss and brain tissue damage. To determine whether BTZ-induced ICH persists or resolves after the cessation of exposure, we performed a recovery experiment: after 72 h of continuous BTZ treatment, zebrafish embryos were transferred to drug-free embryo medium and maintained for an additional 24 h, then re-examined for intracranial hemorrhage. Strikingly, the hemorrhagic foci that were prominent at 72 hpf were largely resolved after 24 h of recovery in clean medium, indicating that BTZ-induced ICH is partially reversible upon drug withdrawal (Supplementary Figure S3).

Fig. 2.

BTZ-induced damage to brain tissue and impaired locomotor activity in zebrafish larvae. (A) Representative confocal images of Tg(elavl3:EGFP) larval heads at 72 hpf, shown in pseudocolor (cyan) for EGFP signal. White arrows indicate discrete hypointense hollow areas. The color scale bar represents relative fluorescence intensity (light, low intensity; dark, high intensity). Scale bar, 100 μm. *n* = 3 independent biological replicates (8 larvae imaged per replicate). (B) Lateral brain view using Tg(elavl3:EGFP), also displayed in pseudocolor (cyan). The white dashed line and white arrow point to the hollow area. Scale bar, 50 μm. (C) Representative movement heatmaps of zebrafish larvae in each treatment group. The color marks indicate the area through which the zebrafish passed, and the depth of color represents the cumulative frequency of the tracking path. (D) Swimming behavior of larvae under 30 min light–dark stimulation. (E–G) quantitative analysis of larval locomotion parameters.For panels C–G, *n* = 3 independent biological replicates (6 larvae per replicate; total *n* = 20 larvae per group). Each larva was recorded individually in a separate well. Data are presented as the mean ± S.D. *p* <0.05, **p* <0.01, ***p* <0.001, ****p* <0.0001

Given the brain’s critical role in behavioral regulation [41], we monitored locomotor activity in each treatment group 6 dpf later. Notably, all BTZ-treated juvenile fish exhibited a significant reduction in total distance traveled during the open-field test (Fig. 2C), suggesting that the suppression of locomotor behavior was not solely attributable to ICH; the pharmacological properties of BTZ itself also caused severe impairment of the juveniles’ behavior. Quantitative assessment confirmed significant decreases in distance traveled (Fig. 2D), movement time (Fig. 2F), locomotor activity (Fig. 2G), and average swimming speed (Fig. 2E).

Collectively, these results demonstrate that BTZ at 4–5 mg/L specifically induces ICH in a concentration-dependent manner, and that the accompanying local neuronal loss occurs at the same time as the ICH.

BTZ induces cerebrovascular disruption and structural damage in zebrafish brain after 72 hours of exposure

Vascular development in zebrafish occurs during 48 to 96 hpf [42, 43], a critical developmental window characterized by three core sequential biological processes: vasculogenesis, angiogenesis, and vascular pruning. The structural framework of the vascular network tends to be fully stabilized at approximately 5 to 7 dpf [44], and the BBB does not acquire full functional competence until approximately 5 dpf [45]. Notably, the peak incidence of BTZ-induced intracerebral hemorrhage occurs at 72 hpf, which exactly falls within this critical window of vascular development, suggesting that BTZ exposure may disrupt the physiological maturation of the cerebrovascular system.

To observe the damage to the cerebrovascular system in zebrafish, we performed drug exposure experiments using the transgenic zebrafish lines Tg(fli:GFP), Tg(kdrl:mCherry) and Tg(gata1:DsRed) (Figs. 3A and 4A). Control larvae exhibited intact, well-organized cerebral vasculature and normal blood distribution, while BTZ-treated larvae showed concentration-dependent vascular disruption, with 5 mg/L BTZ causing severe vascular fragmentation and abnormal blood accumulation, consistent with ICH pathology. To quantify the characteristics of the cerebrovascular skeleton (Fig. 3B), we used AngioTool to analyze structural alterations of the vascular skeleton (Fig. 3D), including vascular area, total number of junctions, junction density, total vessel length, and total number of endpoints (Fig. 3C). Quantitative assessment revealed that BTZ exposure led to a concentration-dependent reduction in vessel area (Fig. 3E), total number of junctions (Fig. 3F), junction density (Fig. 3G), and total vessel length (Fig. 3H), while the total number of endpoints increased significantly (Fig. 3I), confirming progressive vascular fragmentation and loss of network complexity. These structural changes indicate that, regardless of whether gross hemorrhage resolves, BTZ exposure causes persistent structural damage to the cerebral vasculature. The potential cerebrovascular pathogenesis underlying BTZ-induced intracranial hemorrhage is illustrated in the schematic diagram (Fig. 4B). To investigate the molecular mechanisms underlying this disruption, we examined the transcriptional levels of genes associated with endothelial tight junctions (cldn5a, the BBB-specific claudin paralog in zebrafish; oclna), inflammatory factors (il-1β, tnf-α), and glial cell-derived neurotrophic factor (bdnf) (Fig. 4D). The expression levels of these genes were measured at 72 hpf, the time point of peak bleeding.

Fig. 3.

Effects of BTZ on cerebral vascular morphology and structural parameters in zebrafish larvae. (A) Vascular morphology and red blood cell status visualized by fluorescence microscopy using Tg(fli1:EGFP) and Tg(gata1:DsRed). Images are shown in pseudocolor: cyan (EGFP, vasculature) and magenta (DsRed, red blood cells). Color scale bars indicate relative fluorescence intensity (light, low; dark, high). Scale bar, 100 μm. *n* = 3 independent biological replicates (8 larvae imaged per replicate). (B) Schematic diagram of blood vessels. Red indicates blood vessels, orange indicates branch points. (C) Illustration of vascular parameter definitions. (D) Cerebral vascular skeleton analyzed by AngioTool, dorsal view of the larval brain. Vascular outlines are shown in yellow, vascular skeletons in red, and branch points in blue. Scale bar, 100 μm. (E–I) quantitative analysis of vascular structural indexes. Zebrafish were treated with control, 4, 4.5, and 5 mg/L BTZ for 72 h. Each dot represents one larva; *n* = 15–20 larvae per group pooled from 3 independent biological replicates. Data are presented as the mean ± S.D. *p* <0.05, **p* <0.01, ***p* <0.001, ****p* <0.0001

Fig. 4.

BTZ regulates the expression of cerebrovascular-related genes in zebrafish larvae. (A) Fluorescence images of cerebral vascular morphology in zebrafish larvae labeled with Tg(fli:GFP) and Tg(kdrl:mCherry). Images are shown in pseudocolor: cyan (GFP) and magenta (mCherry). Color scale bars indicate relative fluorescence intensity (light, low; dark, high). Scale bar, 100 μm. *n* = 3 independent biological replicates (8 larvae imaged per replicate). (B) Schematic diagram of the potential cerebrovascular pathogenesis underlying BTZ-induced intracranial hemorrhage. (C–D) relative expression levels of vascular function and pro-inflammatory genes determined by qPCR. Zebrafish were treated with control and 5 mg/L BTZ for 72 h. Data are presented as the mean ± S.D. *p* <0.05, **p* <0.01, ***p* <0.001, ****p* <0.0001

qPCR analysis revealed dysregulation of genes associated with vascular development and stability (Fig. 4C–D). BTZ exposure significantly down-regulated the expression of key vascular markers, including VEGFa, KLF2a, ANGPT1, VEGFr2, KRIT1, and TIE1. The observed downregulation of tight junction genes at 72 hpf suggests impaired maturation of the developing cerebrovascular barrier. Notably, knockdown of cldn5a, the zebrafish BBB-specific tight junction gene, has been shown to disrupt blood-neural barrier function during zebrafish development [46]. In mammalian models, reduced expression of cldn5a and occludin leads to increased vascular permeability [47]. The blood‑brain barrier (BBB) in zebrafish embryos is not fully established before 72 hpf. Nevertheless, we detected reduced (though non‑significant) transcript levels of cldn5a, oclna and tjp1a in embryos at this critical developmental stage (Fig. 4D). These findings still indicate that BTZ disrupts the normal assembly and stabilization of endothelial tight junctions, which may increase the permeability of the developing vasculature and trigger subsequent hemorrhage.

A significant upregulation of pro-inflammatory factors (il-1β, tnf-α) was observed as early as 72 hpf, and this trend persisted during the resolution phase of inflammation (Fig. 4D). This persistent neuroinflammatory response closely matches the pattern observed in zebrafish and mammalian hemorrhage models—where disruption of the endothelial barrier triggers cytokine overexpression [48, 49]. Decreased expression levels of tight junction-associated genes and bdnf were observed concurrently with the onset of the inflammatory response (Fig. 4D). bdnfa is a critical neurotrophin that supports neuronal survival, synaptic plasticity, and cognitive function. In the context of brain injury, reduced bdnfa levels are associated with impaired neurogenesis, diminished neuronal resilience, and poorer functional recovery [50]. Conversely, elevation of bdnfa expression through various interventions has been widely validated to facilitate neurogenesis, protect neurons against excitotoxic and oxidative damage, and improve neurological outcomes in multiple models of central nervous system injury and disease [51]. Furthermore, bdnfa has been shown to exert anti-inflammatory effects and to modulate the crosstalk between the immune and nervous systems. Therefore, in the present model, the combination of persistent inflammatory response and reduced bdnfa expression may reflect a failure in the vascular-neural repair cascade, exacerbating the damage initiated by BTZ-induced vascular disruption.

Collectively, these findings demonstrate that 72 h of BTZ exposure impairs the normal developmental program of the zebrafish cerebrovascular system by altering the expression of vascular development and tight junction genes. This early disruption of barrier maturation, together with persistent neuroinflammation and reduced neurotrophic support, coincides with progressive structural deterioration of the cerebral vasculature—characterized by reduced vascular density, impaired branching, and vascular fragmentation—which ultimately contributes to the compromised vascular integrity and intracerebral hemorrhage observed in BTZ-treated larvae.

Integrated multi-omics reveals an association between lipid metabolism dysregulation and BTZ-Induced ICH

During our experiments, we observed that exposure to BTZ induces intracranial hemorrhage in zebrafish larvae, accompanied by a phenotype of disrupted lipid metabolism. To investigate the underlying molecular alterations associated with this phenomenon, we performed integrated transcriptomic and metabolomic analyses on BTZ-treated zebrafish larvae. As illustrated in Fig. 5A (Details are listed in Supplementary Table S2), Venn diagram analysis of differentially expressed genes (DEGs) and differentially expressed metabolites (DEMs) related to lipid metabolism identified 79 overlapping molecules between the two omics datasets, indicating moderate consistency in lipid-associated molecular changes at both the transcriptional and metabolic levels.

Fig. 5.

Correlation analysis of BTZ-induced intracranial hemorrhage and lipid metabolism disorder in zebrafish larvae. (A) Venn diagram of differentially expressed genes and metabolites related to lipid metabolism disorder. (B–D) combined transcriptomic and metabolomic analysis exploring the potential mechanism of lipid metabolism disorder involved in BTZ-induced intracranial hemorrhage. (E) Representative Oil red O staining of larval heads at 72 hpf. White arrowheads indicate lipid-positive deposits in cerebral vessels. Scale bar, 250 μm. n = 3 independent biological replicates (15 larvae stained per replicate). (F–J) quantitative detection of lipid metabolism-related gene expression and enzyme activities. Zebrafish were treated with control and 5 mg/L BTZ for 72 h. Data are presented as the mean ± S.D. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

Joint KEGG pathway enrichment analysis of these overlapping molecules (Fig. 5B. Details are listed in Supplementary Table S3) revealed predominant involvement of the phospholipase D signaling pathway, gap junctions, and bile secretion pathway. Gap junctions are critical structures for maintaining BBB integrity, and their dysfunction is tightly linked to intracranial hemorrhage—consistent with the hemorrhagic phenotype observed in our experiments. Subsequent transcriptomic pathway enrichment analysis of DEGs (Fig. 5C. Details are listed in Supplementary Table S5) showed significant enrichment in the Hippo, Rap1, Wnt, and phospholipase D signaling pathways, as well as ECM-receptor interaction. All of these pathways are closely implicated in vascular development and the maintenance of vascular homeostasis.

At the metabolomic level (Fig. 5D Details are listed in Supplementary Table S4), differential metabolites were markedly enriched in multiple lipid metabolism pathways, including the phospholipase D signaling pathway, bile secretion, arachidonic acid metabolism, glycerophospholipid metabolism, ether lipid metabolism, and retinol metabolism. This aligns with the disrupted lipid metabolism phenotype detected in our assays. Notably, the phospholipase D signaling pathway was enriched at both the transcriptomic and metabolomic layers. This pathway exerts vital regulatory functions in both lipid metabolism and vascular development, and its dual enrichment is highly consistent with the dual phenotypic outcomes of our experiments (disordered lipid metabolism plus intracranial hemorrhage). Collectively, these multi-omics results indicate that BTZ exposure induces coordinated dysregulation at both the transcriptional and metabolic levels, affecting lipid and vascular developmental signaling. To validate these observations, we next examined the expression of key lipid metabolism-related genes by qPCR. Results showed (Fig. 5F) that the mRNA expression level of the fatty acid synthesis regulator PPARA1 exhibited a significant downward trend with increasing BTZ concentration (*p ≤ 0.05), while the mRNA expression levels of the lipid transport-related factor LPL (**p ≤ 0.01) and the lipid catabolism-related factor RBP4 both exhibited a significant downward trend with increasing BTZ concentration (***p ≤ 0.001), reaching a peak in the 5 mg/L group. This suggests that BTZ exposure disrupts the expression balance of lipid metabolism-related factors in zebrafish larvae by negatively regulating lipid catabolism and transport factors.

To investigate the effects of BTZ exposure on lipid metabolism in the brains of zebrafish larvae, Oil Red O staining was used to examine lipid deposition and hemorrhagic phenotypes in cerebral blood vessels. As shown in Fig. 5E, no significant Oil Red O-positive staining was observed in the cerebral vascular regions of the control group; in contrast, the BTZ-treated groups exhibited significant Oil Red O-positive staining signals (indicated by white arrows), with the extent and intensity of positive staining increasing in a dose-dependent manner with rising BTZ concentrations (more details in Supplementary Figure S4). These results indicate that the severity of BTZ-induced cerebral hemorrhage in zebrafish larvae is closely associated with abnormal lipid deposition in the cerebral vascular region, suggesting that lipid metabolism disorders may be involved in the BTZ-induced cerebral hemorrhage process.The results of key lipid parameter measurements in brain tissue and peripheral blood (Fig. 5G–J) showed that, compared with the control group, total cholesterol (T-CHO), triglycerides (TG), and high-density lipoprotein cholesterol (HDL-C) were significantly elevated in a concentration-dependent manner. Specifically, in the 4.5 mg/L group, T-CHO (***p ≤ 0.001), TG (****p ≤ 0.0001), and HDL-C (**p ≤ 0.01) were significantly elevated, while low-density lipoprotein cholesterol (LDL-C) levels in all treatment groups were significantly lower than those in the control group (***p ≤ 0.001~****p ≤ 0.0001). These results show a dissociation from the overall cholesterol accumulation phenotype characterized by elevated total cholesterol (T-CHO) levels, suggesting that BTZ exposure not only induces an imbalance in the overall cholesterol homeostasis of zebrafish larvae but also significantly disrupts the processes of lipoprotein assembly, secretion, and metabolism. This indicates abnormal accumulation of lipid substances in the larvae, leading to an imbalance in lipid metabolic homeostasis.

BTZ exposure induces cerebral ROS accumulation, antioxidant enzyme response, apoptosis and cerebrovascular injury in zebrafish larvae

Lipid metabolism dysregulation can further promote oxidative stress and endothelial dysfunction [52], which prompted us to examine ROS, NO, and apoptosis in the cerebrovascular system. To explore the mechanism of BTZ-induced intracerebral hemorrhage, we detected reactive oxygen species (ROS) levels, redox status, cell apoptosis, and nitric oxide (NO) content in larval brains using vascular-specific transgenic zebrafish Tg (kdrl:mCherry), fluorescence staining, and biochemical assays (Fig. 6). As shown in Fig. 6A, the control group exhibited only weak basal DCFH-DA fluorescence in the brain. In contrast, BTZ treatment induced strong cerebrovascular-associated ROS fluorescence signals that overlapped with vascular endothelial signals, indicating that ROS accumulation was concentrated in the cerebrovascular system. Quantitative analysis confirmed that cerebral ROS levels were significantly elevated in a concentration-dependent manner (****p ≤ 0.0001, Fig. 6B).

Fig. 6.

BTZ exposure induces oxidative stress and apoptosis in zebrafish larval brain. (A) Representative confocal images of DCFH-DA (ROS probe) and Tg(kdrl:mCherry) in larval heads at 72 hpf. Images are shown in pseudocolor: cyan (DCFH-DA, ROS) and magenta (kdrl:mcherry, vascular endothelium). Color scale bars indicate relative fluorescence intensity (light, low; dark, high). Scale bar, 100 μm. *n* = 3 independent biological replicates (5 larvae imaged per replicate). (B) Quantitative analysis of cerebral ROS fluorescence intensity. (C–E) detection of antioxidant enzyme activities (SOD, CAT) and MDA content in zebrafish larvae. (F) Acridine orange staining for neuronal apoptosis, merged with vascular fluorescence signals. Both signals are displayed in pseudocolor: cyan (acridine orange, apoptotic cells) and magenta (kdrl:mcherry, vasculature). Color scale bars as in (A). Yellow arrows indicate apoptotic cells. Scale bar, 100 μm. *n* = 3 independent biological replicates (8 larvae imaged per replicate). (G) Statistical analysis of apoptotic cell number in the larval brain. (H) Detection of NO content in brain tissue. Zebrafish were treated with control, 4, 4.5, and 5 mg/L BTZ for 72 h. Data are presented as the mean ± S.D. *p* <0.05, **p* <0.01, ***p* <0.001, ****p* <0.0001

To characterize oxidative stress responses, we measured lipid peroxidation and antioxidant enzyme activities. Malondialdehyde (MDA), a key product of lipid peroxidation, was significantly increased in BTZ-treated larvae relative to the control (Fig. 6C), suggesting severe oxidative damage to vascular endothelial membranes. Meanwhile, catalase (CAT) and superoxide dismutase (SOD) activities were significantly upregulated in a concentration-dependent manner (Fig. 6D, E), indicating a compensatory activation of the antioxidant system. However, this response was insufficient to counteract excessive ROS accumulation, leading to persistent oxidative stress injury.

Acridine Orange (AO) staining revealed that BTZ treatment markedly increased the number of apoptotic cells in the brain, intensifying with increasing BTZ concentration (Fig. 6F, G). To further evaluate apoptosis in the brain tissue, we performed TUNEL staining on BTZ-treated larvae (Supplementary Figure S2). TUNEL-positive signals were detected in the brain regions of BTZ-treated groups, with the number of apoptotic cells increasing in a concentration-dependent manner, consistent with the AO staining results. These findings indicate that BTZ exposure induces significant apoptosis in the brain.

Furthermore, we detected total NO content in larval brains to evaluate vascular endothelial function. BTZ exposure significantly increased NO levels in a concentration-dependent manner (****p ≤ 0.0001, Fig. 6H). Excessive ROS and abnormal NO elevation synergistically aggravated cerebrovascular oxidative damage and endothelial dysfunction, increased vascular permeability, and promoted the disruption of tight junctions. Together, these results indicate that ROS overproduction, insufficient antioxidant compensation, and NO dysregulation together correlate with BTZ-induced cerebrovascular damage and subsequent intracerebral hemorrhage.

Conclusion

This study demonstrates that BTZ exposure during early zebrafish development induces concentration-dependent ICH accompanied by cerebrovascular disruption, tight junction gene downregulation, and persistent neuroinflammation. Integrated transcriptomic and metabolomic analyses identified 45 molecules at the intersection of lipid metabolism-related DEGs and DEMs, with abnormal lipid deposition in cerebral vessels and disrupted cholesterol homeostasis further supporting a lipid disturbance–cerebrovascular injury association. ROS accumulation, insufficient antioxidant compensation, and NO dysregulation together constituted a pathological cascade correlated with cerebrovascular impairment. These findings identify lipid metabolism dysregulation as a potential contributor to BTZ-induced ICH and provide a multi-omics framework for evaluating developmental vascular toxicity of proteasome inhibitors.

These observations suggest that lipid metabolism disorder may act as a contributing factor that facilitates oxidative stress, NO imbalance, and vascular dysfunction. However, direct functional validation (e.g., lipid pathway rescue or inhibition experiments) will be required to establish a definitive causal relationship. Collectively, our findings indicate that BTZ exposure is associated with a multi-faceted pathological state involving lipid metabolism disorder, oxidative stress, NO dysregulation, and impaired cerebrovascular barrier maturation, which together correlate with the development of intracerebral hemorrhage in zebrafish larvae.

Electronic supplementary material

Below is the link to the electronic supplementary material.

Author contributions

Yulian Li: Writing original draft, Writing-review & editing, Conceptualization. Si Dong: Visualization, Data Curation. Hao Cheng: Methodology. Yunyun Du: Methodology, Formal analysis. Shanshan Zhu: Methodology. Fasheng Liu: Methodology. Yongliang Zheng: Methodology,Conceptualization. Ting Ding: Methodology, Conceptualization. Wenfeng He: Visualization, Conceptualization. Xiaoping Wang: Methodology,Conceptualization. Zigang Cao:Writing manuscript, Methodology. Junquan Zeng: Writing manuscript, Methodology, Conceptualization, Supervision.

Funding

This work was supported by Natural Science Foundation of Jiangxi Province (20242BAB20374; 20252BAC240523; S202671732). Jiangxi Provincial Degree and Postgraduate Education and Teaching Reform Research Project, General Project (JXYJG-2024-131). Ji’an City Science and Technology Program Special Project (20255-061723; No.20255-102195). Jinggangshan University Research Fund Special Project (JFD2506).

Data availability

The raw data can be obtained from the public repository (project number: GSE327263). The raw data can be obtained from the public repository (project number: MTBLS14230).

Declarations

Competing interests

The authors declare no competing interests.