INTRODUCTION
Sargassum fusiforme, a traditional edible seaweed in East Asia, is particularly notable for its rich fucoidan content and balanced nutritional composition, containing various bioactive components such as essential amino acids, minerals, and sterols (Chen et al. 2020). Among them, fucoidan is a sulfated polysaccharide primarily isolated from brown algae that has significant anti-inflammatory, antioxidant, and antithrombotic activities (Zhang et al. 2020, Zhuang et al. 2024). Its polysaccharide component S. fusiforme polysaccharides (SFPS) has shown potential in supporting neurovascular health, making it a candidate worthy of further investigation in cerebral ischemia research (Wang et al. 2021).
Ischemic stroke (IS), a major cerebrovascular disease characterized by cerebral artery occlusion and subsequent hypoxic injury, poses significant challenges to global healthcare systems (Fraser et al. 2023). Current treatment strategies remain limited. Although tissue plasminogen activator is the only Food and Drug Administration-approved pharmacological agent for acute IS, its clinical application is limited by the risks of hemorrhagic transformation and reperfusion injury (Wendelboe and Raskob 2016). Limitations highlight the urgent need to explore novel multitarget therapeutic agents from natural sources that possess cerebrovascular protective and neural repair functions. In this context, bioactive components derived from marine algae have emerged as promising candidates because of their structural diversity and biological functional properties (Haggag et al. 2023).
The hypoxia-inducible factor 1α (HIF-1α) signaling pathway serves as a core regulator of adaptive responses to oxygen deprivation, and plays a key role in postischemic recovery processes such as angiogenesis and neurogenesis (Ni et al. 2022). As an important downstream effector, vascular endothelial growth factor A (VEGFA) not only promotes vascular regeneration but also performs direct neurotrophic functions (Chen et al. 2019, Hu et al. 2022). Given the structural similarity between SFPS and known bioactive fucoidans, we hypothesize that SFPS may exert protective effects in IS by modulating the HIF-1α/VEGF axis.
To systematically test this hypothesis, an integrated experimental strategy combining a CoCl2-induced hypoxic injury model in human cerebral microvascular endothelial cells (hCMEC/D3) with a zebrafish model of cerebral ischemia was used in this study. The zebrafish system offers unique advantages for high-throughput neuropharmacological screening, including genetic homology with humans, embryonic optical transparency, and highly conserved brain structures (Saputra et al. 2025). Through this combined in vitro and in vivo approach, we aim to provide new theoretical foundations for the application of marine algal polysaccharides in cerebrovascular disease treatment, while establishing a basis for the development of neuroprotective agents based on bioactive algal components.
MATERIALS AND METHODS
Extraction and separation of fucoidan from Sargassum fusiforme
Brown seaweed S. fusiforme (voucher No. PEY0030068; synonymous with Hizikia fusiformis) was collected in May 2024 from Zhejiang Province, China (CH: 27°50′27.9″ N, 121°02′53.9″ E). The extraction method followed our prior protocol (Dai et al. 2019). Fresh algae were rinsed with tap water, lyophilized, and washed with 1% citric acid solution for 30 min. The pH was adjusted to 7.0 using distilled water. The samples underwent thermal treatment at 90°C for 10 min in deionized water to remove heavy metals (e.g., arsenic), followed by secondary lyophilization and pulverization. Powdered biomass was homogenized with distilled water at 50°C for 24 h with continuous agitation. The crude extracts were concentrated under vacuum at 37°C, mixed with 95% ethanol, and centrifuged for 15 min. The precipitates were redissolved in distilled water and dialyzed against distilled water at 4°C for 72 h with exchanges every 12 h.
Molecular weight distribution
Molecular weight distribution was assessed through agarose gel electrophoresis on the basis of previously documented procedures (Wang et al. 2019). The samples were resolved on 1% agarose gels at 100 V for 20 min, with dextran sulfate standards (5–160 kDa, D8906; Sigma, St. Louis, MO, USA) serving as molecular weight markers. Postelectrophoresis processing included 4 h of fixation in 0.1% cetrimonium bromide, 30 min of staining with 0.1% toluidine blue, and destaining in acetic acid/ethanol/water (1:50:49).
Structural characterization
Fourier-transform infrared (FT-IR) spectra were recorded on a Thermo Scientific Nicolet 6700 spectrometer (Waltham, MA, USA). Spectral data were collected across the 500–4,000 cm−1 range to characterize the functional groups present in the polysaccharide samples.
Analysis of monosaccharide composition
The monosaccharide profile of the samples was determined following precolumn derivatization with 1-phenyl-3-methyl-5-pyrazolone (PMP). Analysis was conducted using an Agilent 1260 HPLC system (Agilent Technologies, Santa Clara, CA, USA) according to established methodology (Tao et al. 2023), with both reference standards and experimental samples processed identically.
In vitro experiments
Cell culture
hCMEC/D3 cells were obtained from Wuhan Procell Biotechnology Co., Ltd. (Wuhan, China). The cells were maintained in endothelial cell growth medium supplemented with 10% fetal bovine serum, 100 mg L−1 penicillin, and 100 mg L−1 streptomycin. The cultures were grown in a humidified incubator at 37°C with 5% CO2 and 95% relative humidity. The culture medium was replaced with fresh medium every 2–3 days, and subculturing was performed when the cells reached 80% confluence. Cells in the logarithmic growth phase were collected for subsequent studies. For the experimental protocols, hCMEC/D3 cells were divided into the following five groups: a control group, a hypoxia model group (exposed to 160 μM CoCl2 for 24 h); and three intervention groups treated with 160 μM CoCl2 combined with graded concentrations of SFPS (12.5, 25, and 50 μg mL−1) for 24 h.
Cell viability assay
hCMEC/D3 cells were seeded in 96-well plates at a density of 2 × 104 cells per well. Following a 24 h incubation period, the cells were treated with complete medium containing different concentrations of CoCl2 (40, 80, 160, 320, and 400 μM) and SFPS (12.5, 25, 50, 100, 150, and 300 μg mL−1). After another 24 h of incubation, the supernatant was removed, and 100 μl of a diluted solution from the Cell Counting Kit-8 (CCK-8) (Yuanye, Shanghai, China) was added to each well. The cells were then incubated at 37°C. After 2 h of incubation, the cell viability was quantified by measuring the absorbance at 450 nm using an infinite M200 pro multifunctional microplate reader (Tecan, Mannedorf, Switzerland). After screening the concentrations of CoCl2 and SFPS, the cells were divided into five treatment groups. After 24 h of treatment, the same method was used to determine cell viability across all five groups.
Cell apoptosis assay
To comprehensively evaluate cellular apoptosis, a combined approach utilizing fluorescence staining and flow cytometry was employed. Briefly, hCMEC/D3 cells were seeded in 24-well plates and subjected to experimental treatments. For morphological analysis, cells were first stained with 50 μL of Hoechst 33342 solution (Beyotime, Shanghai, China) and incubated at 37°C under 5% CO2 for 30 min. Subsequently, an acridine orange (AO)/ethidium bromide (EB) dual-staining working solution was freshly prepared at a volumetric ratio of 1:1:8 (solutions A:B:C) according to the manufacturer’s instructions (Yuanye) and applied to the cells for 15 min at room temperature in the dark. Fluorescence signals were visualized and captured using an inverted microscope (Nikon, Tokyo, Japan), with Hoechst 33342 detected in the blue channel and AO/EB under 488 nm excitation. For quantitative analysis of apoptosis, an Annexin V-FITC/PI detection kit (Yuanye) was used. Cells were harvested and dual-stained with FITC-conjugated Annexin V and propidium iodide (PI) in binding buffer for 20 min at room temperature in the dark. Data acquisition and analysis via flow cytometry were completed within 1 h after staining to ensure accuracy.
Nitric oxide staining
Nitric oxide (NO) staining (Yuanye) was performed in 24-well plates. Media containing different concentrations of drugs were added to the cells, followed by incubation for 24 h. Then, 1 mL of 5 μM 4-amino-5-methylamino-2′,7′-difluorofluorescein diacetate (DAF-FM DA) solution was added to each well. The intracellular content of NO was examined in hCMEC/D3 cells loaded with the DAF-FM DA NO-sensitive fluorescent probe DAF-FM DA (5 μM) at 37°C for 20 min. The cells were subsequently washed three times with phosphate-buffered saline (PBS), and the cells were observed under a fluorescence microscope (Nikon).
Scratch test
hCMEC/D3 cells (2×105 cells well−1) were cultured in 6-well plates. Upon reaching >95% confluency, a scratch wound was generated using a 200 μL pipette tip, followed by two washes with PBS to remove detached cells. The cells were then treated with various concentrations of SFPS. Images of the scratch area were acquired at 0, 12, and 24 h post-scratch using a microscope (Olympus, Tokyo, Japan). Migratory activity was quantified by measuring scratch closure with ImageJ software (NIH, Bethesda, MD, USA).
In vitro tube formation assays
The tube-forming capacity of hCMEC/D3 cells was assessed using a basement membrane matrix (BMM)-based assay. Briefly, BMM (Corning Incorporated, Corning, NY, USA) was coated onto 96-well plates and polymerized at 37°C for 30 min. The cells (1 × 104 cells well−1) were seeded in conditioned medium containing 1% fetal bovine serum, treated with graded concentrations of SFPS, and maintained under standard culture conditions (37°C and 5% CO2) for 6 h. After incubation, the tubular networks were stained with 2 μM calcein-AM (Yuanye) for 20 min at 37°C. Angiogenic parameters were quantified using ImageJ software (NIH) with the Angiogenesis Analyzer plugin.
Experimental procedures for in vivo zebrafish assays
Zebrafish animal maintenance and embryo collection
Adult male and female zebrafish (wild-type AB) were purchased from the Chinese Zebrafish Resource Center (Wuhan, China). The zebrafish were maintained in a recirculating water system with a light/dark cycle (14 h of light/10 h of dark) and were kept in 20 L glass fish tanks at the standard temperature of the species (28°C). The zebrafish were fed brine shrimp twice a day with brine shrimp. Healthy, sexually mature zebrafish were placed into the spawning pool at a male-to-female ratio of 1:2, and the embryos were collected the next day. All the embryos were cultivated in E3 medium after methylene blue immersion. E3 medium (NaCl, KCl, CaCl2, and MgCl2 at pH 7.2) served as the embryonic environment. All zebrafish were used with permission from the Laboratory Animal Ethics Committee of Changchun University of Chinese Medicine (Approval No. 2025022).
Modeling of hypoxia and pharmacological treatment of zebrafish
Prior to pharmacologic treatment, the effects of CoCl2 as an inducer of hypoxia was evaluated by performing a concentration gradient analysis 2 days post-fertilization (dpf). A total of 20 normal larvae were placed in each well of a 6-well plate, and embryos at 2 dpf were treated with various doses of CoCl2 at concentrations ranging from 2.5 to 40 mM. Vitality was monitored every 2 h, and nonviable embryos were removed to ensure water quality. The numbers were recorded. The efficacious induction concentration criterion was defined as a post-24-h stimulation survival rate exceeding 80%. The experimental design included the following six groups: the control group, model group (5 mM CoCl2), positive control group (5 mM CoCl2 with 60 μM aspirin (Yuanye), and three treatment groups (5 mM CoCl2 with 12.5, 25, and 50 μg mL−1 SFPS).
O-dianisidine staining
On the basis of previous studies, the antithrombotic activity of SFPS was quantitatively assessed in zebrafish by measuring cardiac erythrocyte staining intensity using o-dianisidine. The staining solution contained 2 mL of o-dianisidine solution, 0.5 mL of 0.1 M sodium acetate, 2 mL of E3 medium, and 0.1 mL of 30% H2O2. At the experimental endpoint, six zebrafish per group were randomly selected, incubated with freshly prepared o-dianisidine solution for 5 min, and rinsed three times with PBS. The specimens were fixed in 4% paraformaldehyde on double-concave slides, and the cardiac regions were imaged using a stereomicroscope (CKX53; Olympus). ImageJ software (NIH) was used for quantitative analysis of therapeutic outcomes.
AO staining
After the pharmacologic treatment for 24 h, AO (Solarbio, Beijing, China) staining was performed. AO is a dye that penetrates cells with intact cell membranes and embeds in nuclear DNA, allowing the identification of apoptotic cells. AO staining solution (5 μg mL−1) were added to the zebrafish larvae at 3 dpf, which were subsequently incubated for 1.5 h, respectively, in the dark. The larvae were subsequently rinsed three times in E3 medium. The larvae were observed and imaged using a fluorescence microscope (Nikon).
Dynamic observation
At 2 dpf, the zebrafish larvae were randomly allocated into six groups (n = 10/group) and transferred to a 96-well plate. Owing to low baseline locomotor activity, 0.015% acetic acid solution was administered to all groups except the control group, to stimulate movement. Locomotion was quantified using the ViewPoint behavioral analysis system (MicroZebraLab, AnimaLab, Poznań, Poland), and total distance was analyzed as the primary endpoint.
Histopathological examination
Zebrafish samples were fixed in 4% paraformaldehyde for 24 h, dehydrated through a series of graded ethanol solutions, and embedded in paraffin following standard protocols. Tissue sections (4 μm thick) were stained with hematoxylin and eosin (H&E) (Baiqiandu, Wuhan, China) and Nissl solution (Baiqiandu), and histopathological features were examined under a microscope for image acquisition and analysis.
Immunohistochemistry
Zebrafish embryos were fixed overnight in 4% paraformaldehyde and dehydrated through a series of graded ethanol solutions (70, 80, and 100%). Fixed embryos were embedded in paraffin and cut into 4 μm-thick sections. Endogenous peroxidase activity was quenched by incubating the sections in 3% hydrogen peroxide at room temperature for 25 min under light-protected conditions. The sections were subsequently blocked with 3% bovine serum albumin solution for 2 h. The sections were incubated at 4°C overnight with an anti-endothelial nitric oxide synthase (eNOS) primary antibody (diluted in blocking buffer), followed by incubation with the horseradish peroxidase-conjugated goat anti-rabbit IgG secondary antibody (1:500) for 50 min at 37°C. Chromogenic visualization was achieved using a DAB substrate kit (Shitai Experimental Equipment Co., Ltd., Jiangsu, China) according to the manufacturer’s protocols.
Measurement of blood flow
To evaluate the hemodynamic status of the zebrafish larvae, 2 dpf larvae were randomly allocated to 6-well plates (10 larvae per well). Following the induction of the hypoxia model, the larvae were treated with graded concentrations of SFPS. Dorsal aorta blood flow was analyzed in three randomly selected larvae per group using the ViewPoint ZebraBlood module (MicroZebraLab, AnimaLab, Poland). Volumetric flow rate (in nL s−1) was calculated by the software through frame-by-frame tracking of erythrocyte movement within a defined ROI encompassing the vessel lumen, using a minimum of 30 frames per acquisition for stable measurement.
Western blotting analysis
Western blot analysis was performed to evaluate protein expression in both the hCMEC/D3 cell and zebrafish models. Cells were seeded at 2× 105 cells mL−1 and treated with varying concentrations of SFPS alongside 160 μM CoCl2 for 24 h, with an untreated group serving as a control. Following treatment, the cells were lysed usinglysis buffer (Biosharp, Hefei, China) and the protein concentrations were determined via a bicinchoninic acid assay.
Twenty micrograms of protein samples were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis and subsequently transferred to nitrocellulose membranes (Merck, Darmstadt, Germany). The membranes were blocked with 5% skim milk and incubated overnight at 4°C with primary antibodies, followed by a 3-h incubation with the corresponding secondary antibodies at room temperature. The same protocol was applied for zebrafish brain protein analysis using 128 hours post fertilization (hpf) larval samples.
The following primary antibodies were used: anti-caspase-3 (AF6311; Affinity Biosciences, Changzhou, China), anti-Bcl-2 (AF6139; Affinity Biosciences, Changzhou, China), anti-Bax (AF0120; Affinity Biosciences, Changzhou, China), anti-eNOS (27120-1-AP; Proteintech, Rosemont, IL, USA), and anti-VEGFA (19003-1-AP; Proteintech), with anti-β-actin (AF7018; Affinity Biosciences, Changzhou, China) and anti-glyceraldehyde 3-phosphate dehydrogenase (GAPDH, AF7021; Affinity Biosciences, Changzhou, China) antibodies serving as loading controls. Protein bands were visualized using chemiluminescent substrates (Cyanagen, Bologna, Italy) and imaged with a FUSION SOLO Vilber Lourmat system. Densitometric analysis was performed using ImageJ software (NIH).
Statistical analysis
Statistical analyses were conducted using GraphPad Prism 8.0 (GraphPad Software Inc., San Diego, CA, USA), and the data are expressed as the means ± standard deviations. For multiple group comparisons, one-way ANOVA was employed, followed by Tukey’s post hoc test for detailed intergroup comparisons. Differences were considered significant at p < 0.05. A graphical summary of the results of this study was generated using BioGDP (https://biogdp.com).
RESULTS
Chemical composition and structural characterization
According to the molecular weight assay, the molecular weight distribution curve of SFPS exhibited a continuous, unimodal profile, primarily ranging from 1,000 to 100,000 Da, with a peak molecular weight (Mp) at approximately 10,000 Da (Fig. 1A). This distribution indicates that SFPS is a polysaccharide component with a relatively broad molecular weight distribution. Its functional groups were identified by FT-IR. Figure 1B shows the broad absorption peak at 3,440 cm−1 is attributed to O-H stretching vibrations, while the strong absorption peak at 1,240 cm−1 is the characteristic signal of the S=O asymmetric stretching vibration of the sulfate ester group (-OSO3−). Furthermore, the absorption peak at 846 cm−1 corresponds to the C-O-S stretching vibration, indicating that the sulfate groups are attached to the sugar ring in an equatorial configuration. Characteristic peaks collectively confirm that SFPS is a typical sulfated polysaccharide. Finally, its monosaccharide composition was analyzed by acid hydrolysis combined with PMP pre-column derivatization and high-performance liquid chromatography. As shown in Table 1 and Fig. 1C, SFPS is composed of various monosaccharides, including a substantial amount of fucose and certain proportions of standard monosaccharides. This clearly demonstrates that SFPS is a complex sulfated fucan with fucose as its primary structural unit.
In vitro protective effects of SFPS on CoCl2-induced hCMEC/D3 cells
SFPS affects the survival and proliferation of hypoxic hCMEC/D3 cells
Fucoidan, a sulfated polysaccharide rich in fucose moieties, was isolated from S. fusiforme. First, the cytotoxicity of CoCl2 in hCMEC/D3 cells was determined by a CCK-8 assay. As shown in Fig. 2B, hCMEC/D3 cell viability decreased with increasing CoCl2 concentration. A concentration of 160 μM CoCl2, which reduced cell viability to approximately 60%, was selected to establish an in vitro hypoxic injury model. The optimal intervention doses of 12.5, 25, and 50 μg mL−1 for SFPS are shown in Fig. 2C. To explore the effect of SFPS on angiogenesis, CoCl2-induced hCMEC/D3 cells were used to assess whether SFPS exerts an anti-ischemic effect in a model of IS-associated hypoxic injury. Following hypoxic injury, the CCK-8 results revealed a decrease in cell viability, which was reversed in a concentration-dependent manner after 24 h of treatment with different concentrations of SFPS (Fig. 2D).
SFPS affects hypoxic hCMEC/D3 cell apoptosis
Following AO/EB staining, the normal group exhibited uniform green fluorescence, with occasional orange-yellow fluorescence observed in spontaneously apoptotic or necrotic cells. Compared with that of the CoCl2 group, a dose-dependent reduction in orange fluorescence intensity was detected with increasing drug concentration, indicating the progressive suppression of apoptosis (Fig. 2E & F).
Hoechst 33342, a cell membrane-permeable blue fluorescent dye, selectively stains nuclei. Fluorescence microscopy revealed that the control group nuclei displayed homogeneous blue fluorescence. In contrast, the CoCl2 group exhibited a marked increase in the number of bright blue fluorescent nuclei, suggesting the presence of chromatin condensation and nuclear pyknosis. SFPS treatment decreased the nuclear fluorescence intensity in a dose-dependent manner, which correlated with a reduction in the number of apoptotic cells. NO, an endothelial cell-dependent vascular modulator, plays a crucial role in promoting angiogenesis. NO staining revealed that the green fluorescence intensity increased in the SFPS-treated groups in a dose-dependent manner (Fig. 2G–I).
Annexin V-FITC/PI dual staining revealed a significant increase in the percentage of apoptotic cells in the CoCl2 group compared with the control group. SFPS treatment partially abrogated these effects, as evidenced by decreased proportions of apoptotic cells (Fig. 2J).
SFPS promotes the migration and tubular formation of hCMEC/D3 cells in vitro.
Scratch wound-healing assays revealed that the migration distance of SFPS-treated cells was significantly greater than that of control cells within the same timeframe (Fig. 3A & B), further confirming that SFPS enhances the migratory capacity of hCMEC/D3 cells in a time- and dose-dependent manner. To evaluate the effect of SFPS on angiogenesis in hCMEC/D3 cells, an in vitro angiogenesis assay was performed. The results showed a marked increase in tubular structure formation, in SFPS-treated cells compared with control cells (Fig. 3C). Treatment with 25 or 50 μg mL−1 SFPS significantly increased the number of junction points and total segment length (Fig. 3D & E).
SFPS exerts antiapoptotic and proangiogenic effects via the HIF/VEGF pathway
To investigate the SFPS-mediated regulation of the HIF/VEGF pathway, the protein expression profiles of HIF-1α, VEGFA, and eNOS were analyzed. Compared with normoxic conditions, CoCl2-induced hypoxic conditions increased HIF-1α and VEGFA protein levels. Compared with CoCl2 treatment, SFPS treatment increased VEGFA and eNOS protein levels (Fig. 3F & G). In addition, CoCl2 stimulation markedly induced apoptosis in hCMEC/D3 cells, whereas SFPS treatment significantly mitigated CoCl2-triggered apoptotic responses. The exposure of hCMEC/D3 cells to CoCl2 significantly upregulated the expression of the proapoptotic proteins Bax and caspase 3 but downregulated the expression of the antiapoptotic protein Bcl-2; SFPS intervention reversed CoCl2-induced apoptotic alterations (Fig. 3H & I). findings collectively indicated that SFPS promotes angiogenesis and inhibits apoptosis through the HIF-1α/VEGF pathway.
Protective effects of SFPS in a zebrafish hypoxia model
The protective effects of SFPS against hypoxia-induced damage in zebrafish larvae were investigated in a CoCl2-mediated model (Fig. 4A). Systematic evaluation of zebrafish larval viability, morphological integrity, and behavioral patterns across graded CoCl2 concentrations (0–40 mM) and induction intervals (4, 12, 16, 20, 24, and 28 h; 96–128 hpf) revealed that 5 mM CoCl2 was the optimal concentration for establishing hypoxia, effectively mitigating the decrease in survival rate (Fig. 4B).
To validate whether SFPS modulates cardiac perfusion in zebrafish larvae following CoCl2-induced IS, o-dianisidine staining was used to assess cardiac blood flow dynamics. SFPS exhibited antithrombotic efficacy comparable to that of aspirin, as evidenced by increased cardiac hemoglobin staining intensity (Fig. 4C).
To investigate apoptotic activation in response to CoCl2-induced hypoxic injury, zebrafish embryos were subjected to AO staining. CoCl2 exposure markedly increased AO fluorescence intensity, which indicated hypoxia-mediated apoptosis, whereas subsequent SFPS treatment restored fluorescence to baseline levels (Fig. 4D).
Cerebral hemodynamics were assessed across experimental groups using ZebraBlood software (ViewPoint, Lyon, France) to analyze erythrocyte movement within tracked regions (Fig. 4E & F). Control zebrafish presented robust, well-organized cerebral blood flow, whereas CoCl22-treated zebrafish presented markedly decelerated flow with complete or partial vascular occlusion. SFPS administration induced progressive restoration of the blood flow velocity, similar to the baseline hemodynamic parameters observed in the controls.
Behavioral analyses revealed a significant reduction in locomotor distance following hypoxia induction compared with that in the normoxic control group (p < 0.001). Both 50 μg mL−1 SFPS and aspirin markedly restored locomotor activity in hypoxic larvae (Fig. 4G & H).
Histopathological evaluation via H&E staining revealed distinct cerebral alterations. Compared with the control group, the hypoxia model group presented a disorganized cellular architecture, cytoplasmic rarefaction, and focal vacuolation. In contrast, treatment with aspirin and graded SFPS concentrations (12.5–50 μg mL−1) ameliorated cerebral histopathology, manifested as compact cellular alignment, reduced cytoplasmic voids, and attenuated vacuolation (Fig. 4I). Nissl staining selectively highlights Nissl bodies within the neuronal cytoplasm, enabling ultrastructural characterization of neurons and assessment of neuronal integrity through Nissl body morphology. Histochemical analysis revealed intact neuronal morphology with densely packed Nissl bodies in the control group, whereas the ischemic model group presented a marked reduction in Nissl body density. Notably, the therapeutic and positive control groups demonstrated robust restoration of Nissl body distribution in the peri-infarct neurons. Compared with those in the model group, the number of Nissl bodies in SFPS-treated cerebral tissues was significantly greater, which indicated that SFPS-mediated neuronal cytoarchitectural preservation (Fig. 4J & K).
Immunohistochemical analysis of eNOS expression revealed distinct spatial and quantitative alterations in NO biosynthesis in zebrafish brain tissues. In the control group, robust eNOS immunoreactivity was observed in cerebral microvascular endothelial cells, as characterized by intense cytoplasmic staining. In contrast, the hypoxia model group presented a significant reduction in eNOS-positive signals, which indicated impaired NO synthesis under ischemic conditions. Notably, SFPS intervention restored eNOS expression in a dose-dependent manner, with the high-dose SFPS group demonstrating near-physiological staining intensity (Fig. 4J). Findings suggest that SFPS increases NO bioavailability via eNOS upregulation, potentially contributing to its neurovascular protective effects through vasodilation and antiapoptotic mechanisms.
The expression of apoptosis-related genes, including Bax, Bcl-2, and caspase 3, was detected. In CoCl2-treated embryos, the expression of proapoptotic genes was upregulated, whereas that of the antiapoptotic gene Bcl-2 was downregulated, which demonstrated the cytoprotective efficacy of SFPS against hypoxia-driven apoptosis (Fig. 4L & M). Further mechanistic analysis revealed SFPS-mediated modulation of angiogenic regulators, with upregulated VEGF and eNOS expression in 128 hpf larvae, substantiating the protective efficacy of SFPS against hypoxia-mediated damage (Fig. 4N & O).
DISCUSSION
This investigation reveals the dual neurovascular protective capacity of SFPS through complementary in vitro and in vivo platforms, demonstrating its coordinated regulation of the HIF-1α/VEGF signaling cascade. Our results indicate that SFPS concurrently enhances angiogenic processes while inhibiting endothelial apoptosis, establishing a comprehensive pharmacological profile for this marine-derived compound. The convergence of evidence from the hCMEC/D3 cellular model and zebrafish models provides robust support for the therapeutic potential of SFPS in IS pathology marked by cerebrovascular impairment and angiogenesis dysregulation.
The multifaceted bioactivity of SFPS should be interpreted within the framework of algal biotechnology. Sargassum fusiforme cultivation represents a sustainable platform for polysaccharide production, with emerging aquaculture methods demonstrating increased biomass yield and fucoidan content (Roostaei et al. 2018). Marine algae efficiently absorb dissolved inorganic carbon from seawater through photosynthesis, converting it into organic biomass. This process not only mitigates ocean acidification but also sequesters atmospheric carbon dioxide, forming a “blue carbon sink” (Chaiklahan et al. 2022). In particular, structural polysaccharides such as fucoidan exhibit complex configurations that are resistant to microbial degradation, enabling long-term persistence in marine environments and facilitating extended carbon sequestration (Shitu et al. 2024). Integrated multitrophic aquaculture systems increase production efficiency while minimizing environmental impact through nutrient recycling (Roostaei et al. 2018). The scalability of SFPS production merits particular consideration, as current extraction methods can be refined via enzyme-assisted and subcritical water extraction technologies that optimize yield while preserving bioactivity (Wu et al. 2025). From a biochemical perspective, advances in understanding fucoidan biosynthesis in brown algae have revealed conserved sulfotransferase enzymes that govern structural heterogeneity (Mabate et al. 2021). Our findings contribute to this expanding knowledge domain by correlating specific polysaccharide characteristics with neurovascular protective efficacy.
Hypoxia, a pathological stress state characterized by diminished oxygen tension, serves as a critical mediator of cerebrovascular pathogenesis, including IS (Hong et al. 2019). Under hypoxic stress, organisms activate compensatory neovascularization mechanisms centered on the HIF-1α/VEGF axis to reestablish oxygen homeostasis. This cascade initiates with HIF-1α stabilization, driving the transcriptional upregulation of VEGF and its receptor system while concurrently increasing vascular permeability to facilitate plasma protein extravasation for provisional matrix formation (Dong et al. 2016, Song et al. 2023). Notably, the angiopoietin system exhibits dynamic reciprocity; Ang-1 reinforces vascular integrity through VE-cadherin/αβ-integrin-mediated endothelial adhesion, whereas Ang-2 induces vascular destabilization via Tie2 receptor antagonism. Paradoxically, Ang-2/VEGF coactivation overcomes homeostatic constraints to initiate vascular sprouting (Solecki et al. 2019). Subsequently, matrix metalloproteinases (MMP2/MMP9) mediate basement membrane degradation, creating migratory pathways for endothelial tip cell invasion and lumenogenesis. During vascular maturation, pericytes engage through platelet-derived growth factor receptor β signaling, collaborating with Ang-1/PDGF-BB complexes to reconstitute functional vessel walls. This multistage regulatory paradigm not only deciphers the molecular logic of hypoxia-adaptive angiogenesis but also informs therapeutic strategies targeting vascular homeostasis (Fig. 5). Moreover, the reconstruction of the functional vascular network after IS depends not only on morphogenesis during neovascularization but also on a multistage dynamic remodeling process of endothelial cell survival, directional differentiation, and lumen maturation (Zong et al. 2020). Increased endothelial viability critically suppresses hypoxia-induced vasculopathy while maintaining barrier integrity for 3D microvascular network reestablishment. Mechanistically, eNOS governs early-stage angiogenesis through NO-mediated regulation of endothelial proliferation and basement membrane remodeling, with its catalytic activity determining lumenogenesis efficiency (Liu et al. 2019). The present experimental validation using CoCl2-induced hCMEC/D3 cellular hypoxia models revealed that SFPS intervention significantly elevates intracellular NO levels, confirming the centrality of eNOS/NO signaling in therapeutic angiogenesis. The findings align with this article reported an eNOS phosphorylation mechanism, reinforcing the translational potential of SFPS in cerebrovascular repair (Jin et al. 2021).
The mechanistic insights from this study have significant implications for marine algal bioproduct development. Activation of the HIF-1α/VEGF pathway by SFPS not only clarifies its pharmacological mechanisms but also establishes critical quality benchmarks for standardizing algal polysaccharide preparations. This molecular-level understanding enables more precise cultivation and processing strategies to maximize target bioactivities (Shao and Duan 2022). Furthermore, the demonstrated effects on endothelial function and angiogenesis suggest that SFPS is a promising candidate for functional food and nutraceutical applications beyond pharmaceutical development.
Our research validates zebrafish as a robust model for evaluating algal compounds, combining physiological relevance with screening efficiency. The conservation of neurovascular responses between zebrafish and mammalian systems, coupled with embryonic transparency and genetic tractability, creates an optimal platform for the rapid assessment of marine natural products (Wang et al. 2024). This approach substantially accelerates the discovery pipeline for algae-based therapeutics.
In summary, this work advances our understanding of marine algal polysaccharides by delineating the mechanisms of action of SFPS while establishing its relevance to algal biotechnology. The dual emphasis on mechanistic pharmacology and sustainable bioproduct development provides a framework for future investigations bridging fundamental phycology with therapeutic applications. Subsequent research should prioritize structure-activity relationship analyses, production process optimization, and clinical translation to fully realize the potential of SFPS as a prototype algae-derived therapeutic agent.
Collectively, the results of this investigation reveal the therapeutic potential of SFPS in IS through multimodal mechanistic validation. SFPS orchestrates dual neurovascular protection via synergistic activation of the HIF-1α/VEGF axis. The dual functionality of SFPS—as both a nutraceutical supplement and pharmacological agent—positions it as a compelling candidate for adjunctive IS therapy, bridging acute neuroprotective interventions with chronic vascular rehabilitation strategies. This work not only deciphers the molecular basis of marine herbal medicine’s “blood-activating” properties but also establishes a methodological framework for evaluating natural products in cross-species disease models. By integrating in vitro and in vivo paradigms, we provide a blueprint for translating traditional medicinal concepts into mechanistically grounded therapeutic innovations.
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