Using the broth microdilution method in Müller-Hinton (MH) broth, we evaluated the minimum inhibitory concentration (MIC) of Scymicrosin_7–26. Briefly, Scymicrosin_7–26 was serially two-fold diluted in MH broth. Bacterial suspensions with a density of 1 × 10⁶ colony-forming units per milliliter (CFU/mL) were prepared using mid-logarithmic phase cultures. Each well of 96-well plates received 50 μL aliquots of both drug dilutions and bacterial suspensions. Wells containing MH broth with bacteria but no antimicrobial peptide served as the positive control, while wells containing only sterile MH broth were assigned as negative control. Following overnight incubation (16–18 h, 37 °C), the MIC was designated as the lowest concentration achieving complete inhibition of visual growth. MBC assessment involved subculturing from clear wells onto agar plates, with MBC defined as the minimum concentration demonstrating bactericidal activity (≥99.9% reduction) against the original inoculum (Huo et al., 2020).

A single isolate from each of the five clinical multidrug-resistant pathogens was selected for subsequent experiments. The MIC₅₀ value, defined as the minimal concentration inhibiting 50% of strains, identifies isolates that balance susceptibility and resistance, thus representing a moderate resistance level within the population (Kowalska-Krochmal and Dudek-Wicher, 2021; García-Viñola et al., 2025). The screening procedure was as follows: Step 1: The minimum inhibitory concentration (MIC) of Scymicrosin_7–26 against all isolates was determined, and the MIC₅₀ for each bacterial species was calculated. Step 2: Strains exhibiting MIC values equal to the MIC₅₀ of their respective species were identified, ensuring that the selected isolates demonstrated intermediate susceptibility to the antimicrobial peptide. Their antibiotic susceptibility profiles were then characterized (Supplementary Figures S4–S8). Step 3: Based on the susceptibility profiles, strains that exhibited predominant sensitivity to all tested antibiotics were prioritized as experimental isolates. If multiple strains met this criterion within a species, one was randomly chosen for further study.

One representative strain from each of the five bacterial species—designated AB1 (CR-AB), KP1 (CR-KP), EC1 (ESBL-EC), PA1 (CR-PA), and MRSA1—was selected. Following an established protocol (Rao et al., 2021), each strain was diluted in MH broth to 1 × 10⁶ CFU/mL. Bacterial suspensions (50 μL) were exposed to 50 μL of Scymicrosin_7–26 in 96-well plates, producing final concentrations of 0 (growth control), 0.5, and 1 × MIC. The starting OD₆₀₀ was measured immediately after mixing. Plates were maintained at 37 °C, and bacterial growth was assessed through OD₆₀₀ measurements at 2-h intervals until control wells reached mid-log phase. Established antibiotics (polymyxin B and vancomycin at 1 × MIC) were employed as positive controls, with all experimental conditions replicated three times.

Bacterial strains were prepared at 1 × 10⁶ CFU/mL in fresh MH broth and treated with Scymicrosin_7–26 to reach final concentrations of 0 (untreated control), 1, and 2 × MBC. Incubation was carried out at 37 °C with orbital shaking (190 rpm). At established time intervals, 100 μL samples were collected, diluted serially in 10-fold steps, and 50 μL of each dilution was spotted onto LB agar. After 24 h at 37 °C, viable bacteria were enumerated (Zhu et al., 2021). The results were plotted as survival rate versus time.

The combination effects of Scymicrosin_7–26 with established antibiotics were evaluated via checkerboard microdilution assay (Riool et al., 2020). Bacterial suspensions (1 × 10⁶ CFU/mL) were inoculated into 96-well plates. Scymicrosin_7–26 and the test antibiotic were serially diluted in two dimensions across the plate. After overnight incubation (16–18 h, 37 °C), the MICs of single agents and drug combinations were documented for the five test strains (AB1, KP1, EC1, PA1, MRSA1).

The fractional inhibitory concentration index (FICI) was determined according to the standard formula: FICI = (MIC of drug A in combination/MIC of drug A alone) + (MIC of drug B in combination/MIC of drug B alone).

Based on FICI values, drug interactions were classified as follows: synergistic (FICI ≤ 0.5), additive (0.5 < FICI ≤ 1.0), indifferent (1.0 < FICI ≤ 2.0), and antagonistic (FICI > 2.0).

Following published procedures (Ko et al., 2020), we examined how physiological salt conditions affect Scymicrosin_7–26’s efficacy. Bacterial strains (EC1, KP1, AB1, PA1, MRSA1) were grown overnight and adjusted to 1 × 10⁶ CFU/mL in MH broth. The peptide was serially diluted in MH broth supplemented with either (1) 4 μM FeCl₃, 2.5 mM CaCl₂, and 150 mM NaCl for salt stability assessment, or (2) 5, 10, and 20% (v/v) fetal bovine serum (FBS) for serum stability analysis. MIC determinations followed standard microdilution protocols, with triplicate measurements within each experiment and three separate experimental runs.

The potential for resistance development to Scymicrosin_7–26 was investigated using a serial passage method (Yu et al., 2021). PA1 cultures were transferred to fresh MH medium supplemented with Scymicrosin_7–26 at sub-MIC levels and cultivated at 37 °C. Cultures from 0.5 × MIC wells were harvested after 24 h, diluted 1:1000 in fresh medium, and exposed to a new gradient of peptide concentrations. This daily passaging was continued for 30 days. Polymyxin B and tigecycline were used as control antibiotics. With each transfer, the fold-increase in MIC compared to the original baseline was recorded.

PA1 biofilm formation under Scymicrosin_7–26 exposure was quantified in 96-well plates with peptide concentrations (0, 0.5, 1, 2, 4 × MIC). After 24 h static incubation (37 °C), wells were aspirated, phosphate-buffered saline (PBS)-washed, and methanol-fixed (10 min). Air-dried biofilms underwent crystal violet staining (1%, 20 min), distilled water washing, and ethanol elution for OD₅₉₅ measurement. Technical triplicates and three biological repeats were performed.

Mature PA1 biofilms were established in 96-well plates (24 h, 37 °C), washed with PBS, and challenged with Scymicrosin_7–26 (0, 0.5, 1, 2, 4 × MIC) in fresh MH broth for 24 h at 37 °C. The remaining biofilm was then measured according to the crystal violet method in section 2.1.7.

The cytotoxicity of Scymicrosin_7–26 was evaluated against RAW264.7, Beas-2B, HaCaT, and HEK293T cell lines. Following 24 h culture in complete medium (high-glucose DMEM with 10% FBS) at 1 × 10⁴ cells/well in 96-well plates, cells were treated with Scymicrosin_7–26 (3–48 μM) in fresh medium. Blank controls (medium only) and negative controls (untreated cells) were established. Viability was determined after 24 h using CCK-8 assay (10 μL/well, 2 h incubation at 37 °C) with detection at 450 nm.

The cell survival rate was quantified by the formula:

The absorbance readings for the peptide-treated groups, blank control, and negative control were designated as OD_A, OD_B, and OD_C, respectively. Six replicates were used for each condition.

Hemolysis assay was performed with 4% human erythrocyte suspensions. Erythrocytes were exposed to varying concentrations of Scymicrosin_7–26, 1% Triton X-100 (positive control), and PBS (negative control) for 1 h at 37 °C. Following centrifugation at 4000 × g for 5 min (room temperature, RT), 100 μL of each supernatant was transferred to a 96-well plate. Hemoglobin release was determined by measuring absorbance at 540 nm.

The hemolysis rate was determined as follows:

In this formula, A, A₀, and A_T refer to the absorbance readings of the experimental groups, PBS control, and Triton X-100 control, respectively. All assays were performed in triplicate.

N-phenyl-1-naphthylamine (NPN), a environment-sensitive fluorescent probe, was employed to examine the outer membrane disruption ability of Scymicrosin_7–26. This probe shows minimal fluorescence in aqueous solution but markedly increased emission when incorporated into membrane hydrophobic compartments. In brief, PA1 suspensions (1 × 10⁶ CFU/mL) were loaded with 10 μM NPN for 10 min under dark conditions. After treatment with Scymicrosin_7–26 at 1×, 2×, and 4 × MIC concentrations, along with polymyxin B control (4 μg/mL), fluorescence kinetics were monitored at 1-min intervals (excitation 350 nm, emission 420 nm) using a BioTek plate reader until signal equilibrium.

The effect of Scymicrosin_7–26 on bacterial membrane integrity was tested with the LIVE/DEAD® BacLight™ viability kit. PA1 and MRSA1 (1 × 10⁶ CFU/mL) treated with 1 × MIC peptide for 1 h were stained with SYTO 9/PI combination (20 μM and 20 μg/mL) for 15 min at 37 °C in darkness. Microscopic observation was conducted immediately using a fluorescence microscope.

The morphological effects of Scymicrosin_7–26 on bacterial cells were investigated through SEM imaging, following a previously described method with slight modifications (Kalsy et al., 2020). PA1 and MRSA1 cultures in logarithmic growth phase were normalized to 1 × 10⁸ CFU/mL and exposed to Scymicrosin_7–26 (1 × and 2 × MIC) for 1 h at 37 °C. After centrifugation (3,000 × g, 10 min) and triple PBS washing, cells were fixed in 2.5% glutaraldehyde (4 °C, overnight). Dehydration through graded ethanol series, critical point drying, and gold coating preceded SEM observation.

For TEM observation, samples were processed based on a previously described protocol with customized changes³⁵. Bacterial pellets from peptide-treated cultures (prepared as in 2.2.3) underwent the following processing: primary fixation with 2.5% glutaraldehyde at 4 °C overnight; post-fixation with 1% osmium tetroxide for 2 h at 4 °C; rinsing with PBS; dehydration through a graded acetone series (50, 70, 90, and 100%); embedding in Epon 812 resin and thermal polymerization (70 °C, 48 h), 65–70 nm sections were prepared using a UC6 ultramicrotome. The sections were sequentially stained with 3% uranyl acetate and 1% lead citrate prior to TEM observation.

The DNA-binding capability of Scymicrosin_7–26 was analyzed by an agarose gel retardation assay, as previously reported (Xie et al., 2015). Genomic DNA isolation from PA1 and MRSA1 strains was performed with a commercial bacterial DNA extraction kit. Aliquots of DNA (approximately 400 ng in 10 μL TE buffer) were incubated with increasing concentrations of Scymicrosin_7–26 (0, 3, 6, 12, 24, 48, and 96 μM) at room temperature for 30 min. After incubation, the reaction mixtures were analyzed by 1% agarose gel electrophoresis. A Bio-Rad gel imaging system was used to visualize DNA migration patterns.

Bacterial intracellular ROS production was monitored with the fluorescent indicator 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA), following a published method (Jayathilaka et al., 2021). Bacterial suspensions (OD₆₀₀ ≈ 0.8) of PA1 and MRSA1 in PBS were loaded with 10 μg/mL DCFH-DA and continuously shaken at 37 °C for 1 h. Excess fluorescent probe was removed through three successive PBS washes. DCFH-DA-loaded bacteria were incubated with Scymicrosin_7–26 gradient concentrations (0, 0.5, 1, 2, 4 × MIC) for 60 min. Species-appropriate positive controls (polymyxin B for PA1; lysostaphin for MRSA1) were included. Fluorescence intensity was measured with excitation at 485 nm and emission at 528 nm.

RAW264.7 cells (1 × 10⁶ cells/mL) were distributed into 6-well plates (2 mL/well) containing high-glucose DMEM and incubated for 24 h at 37 °C with 5% CO₂. The study included five experimental conditions: (1) untreated control; (2) LPS-stimulated model (100 ng/mL); and (3–5) treatment groups receiving 2-h pretreatment with Scymicrosin_7–26 (3, 6, or 12 μM) prior to 22-h LPS co-incubation. After treatment, culture media were harvested for subsequent analysis while adherent cells were processed for RNA and protein extraction.

Cellular total RNA was isolated with Trizol reagent. Complementary DNA (cDNA) synthesis was carried out with the PrimeScript® Reverse Transcription Kit following the supplier’s instructions. Using PerfectStart® Green qPCR SuperMix, RT-qPCR analyses were carried out on a Thermo Fisher Applied Biosystems real-time PCR platform. All primer sequences utilized in this work are detailed in Supplementary Table S4.

Commercial ELISA kits were employed to evaluate the levels of IL-1β, IL-6, and TNF-α in cell culture supernatants, strictly adhering to the manufacturer’s protocols.

According to the Griess reaction protocol in the Nitric Oxide Assay Kit, we measured nitrite accumulation in culture media as an indicator of nitric oxide production.

Following the respective treatment, cells underwent three PBS washes before being incubated with the fluorescent probe DCFH-DA (10 μM) for 30 min at 37 °C in darkness. After incubation with DCFH-DA, the cells were washed thoroughly with PBS to eliminate residual extracellular fluorophore. Intracellular ROS levels, indicated by fluorescence, were observed and imaged using a fluorescence microscope.

Treated cells were subjected to cold PBS washes and RIPA buffer lysis on ice. Proteins were denatured (99 °C, 10 min) in loading buffer and electrophoresed on 10% SDS-polyacrylamide gels. Proteins were transferred to PVDF membranes, blocked with 5% BSA (1 h, RT). Primary antibody incubation (4 °C, overnight) preceded TBST washes and HRP-secondary antibody treatment (1 h, RT). Visualization used a chemiluminescent detector, with three biological replicates.

The LPS neutralization activity of Scymicrosin_7–26 was evaluated according to a previously described method (Nell et al., 2006). Briefly, the peptide at concentrations ranging from 1.5 to 48 μM was co-incubated with 100 ng/mL LPS at 37 °C for 30 min. Following incubation, residual LPS levels were measured using a commercial LPS detection kit according to the manufacturer’s instructions. The neutralization percentage was calculated based on the reduction in LPS activity relative to the control (without peptide).

The membrane penetration capability of Scymicrosin_7–26 was evaluated in RAW264.7 macrophages using inverted fluorescence microscopy. Cells were plated in 12-well plates at 5 × 10⁵ cells per well and adhered for 12 h. FITC-labeled peptide was administered in high-glucose DMEM at concentrations ranging from 0 to 12 μM for 1 h. Following treatment, cells were rinsed with PBS, fixed with 4% paraformaldehyde (20 min), and blocked with 10% goat serum. Immunostaining was performed using an anti-F4/80 primary antibody (1:500, overnight) followed by a Cy3-conjugated secondary antibody (1 h, room temperature). Nuclei were counterstained with DAPI after thorough washing. Fluorescence images were acquired using an inverted fluorescence microscope.

After overnight culture in 24-well plates (4 × 10⁵ cells/mL, 500 μL/well), RAW264.7 cells were treated, PBST-washed, and fixed with 4% PFA (15 min). Blocking with 5% BSA (1 h, RT) preceded anti-P65 primary antibody incubation (overnight, 4 °C). Cy3-conjugated secondary antibody was applied (1 h, RT, dark), followed by Hoechst 33342 nuclear staining (10 min) and fluorescence imaging.

The antibacterial efficacy of Scymicrosin_7–26 was initially assessed against a panel of five clinically prevalent multidrug-resistant bacteria isolated from respiratory specimens. The peptide demonstrated antibacterial effects across all tested strains. Analysis of MIC₅₀ and MIC₉₀ values revealed that Scymicrosin_7–26 was most active against Escherichia coli, Acinetobacter baumannii, and methicillin-resistant Staphylococcus aureus (MRSA), followed by Klebsiella pneumoniae, with Pseudomonas aeruginosa exhibiting the highest MIC values (Table 1). The susceptibility of the five strains selected according to the aforementioned criteria to Scymicrosin_7–26 is summarized in Table 2.

Figures 1A–E illustrates the growth kinetics of the tested bacterial strains. Sub-inhibitory concentrations (0.5 × MIC) of Scymicrosin_7–26 significantly retarded the growth of all five strains. At the 1 × MIC concentration, bacterial growth was completely suppressed. Similar inhibitory profiles were documented in positive control groups administered 1 × MIC of either polymyxin B or vancomycin. These findings indicate that Scymicrosin_7–26 exerts concentration-dependent suppression of bacterial growth in the tested multidrug-resistant pathogens.

Time-kill assays were performed to dynamically monitor the bactericidal activity of Scymicrosin_7–26. As shown in Figures 1F–J, exposure to 1 × MBC of the peptide resulted in complete eradication of strains KP1 and AB1 within 60 min, EC1 and PA1 within 20 min, and MRSA1 within 5 min. When the concentration was increased to 2 × MBC, the killing kinetics were accelerated: KP1 and AB1 were eliminated within 40 min, EC1 within 20 min, PA1 within 10 min, and MRSA1 within 3 min. Notably, the killing rate for EC1 was more rapid at 2 × MBC during the initial 20-min period compared to 1 × MBC.

Combination therapy was assessed using the checkerboard microdilution method. As summarized in Supplementary Table S5, the combination of Scymicrosin_7–26 with polymyxin B, tigecycline, imipenem, amikacin, vancomycin, or lincomycin against strains AB1, KP1, EC1, PA1, and MR1 yielded fractional inhibitory concentration index (FICI) values all below 2.00, indicating no antagonistic interactions were observed for any of the tested combinations.

The stability of Scymicrosin_7–26’s antibacterial activity under a range of physiological ion and fetal bovine serum (FBS) concentrations was evaluated. In the presence of a physiological Na⁺ concentration, only a marginal increase in MIC was noted for strains EC1, KP1, and MRSA1. Exposure to a physiological concentration of Ca²⁺ resulted in a modest increase in the MIC for all five bacterial strains, with the effect being most pronounced for PA1. Conversely, physiological Fe³⁺ concentration did not alter the MIC against any of the strains. These results demonstrate that Scymicrosin_7–26 retains robust antibacterial activity in environments mimicking physiological salt conditions. The MIC values remained unchanged in the presence of 5% fetal bovine serum (FBS). However, they increased at higher FBS concentrations (10 and 20%). This effect was most pronounced in strain PA1, while the other tested strains showed only moderate changes (Table 3).

A serial passage experiment was conducted to assess the potential for resistance development. After 30 days of continuous exposure, the MIC of tigecycline and polymyxin B against PA1 increased by 8-fold compared to the baseline, whereas the MIC of Scymicrosin_7–26 remained unchanged (Figure 2A). For MRSA1, the MIC of tigecycline and vancomycin increased by 16-fold and 4-fold, respectively, while the MIC of Scymicrosin_7–26 again showed no increase (Figure 2B).

Scymicrosin_7–26 effectively inhibited biofilm formation at concentrations as low as 0.5 × MIC for both PA1 and MRSA1, with complete inhibition achieved at higher concentrations. Against pre-formed mature biofilms, the peptide also displayed eradication activity. At 0.5 × MIC, a partial removal effect was observed. For PA1, the biofilm biomass was reduced to 26.48% of the control at 4 × MIC. For MRSA1, the biomass decreased to 52.83% at 1 × MIC and was nearly completely eradicated at concentrations ≥2 × MIC (Figures 2C–G).

The cytotoxicity profile of Scymicrosin_7–26 was assessed in Beas-2B, HEK293T, and RAW264.7 cell lines. As shown in Figures 3C–E, the peptide exhibited low to negligible cytotoxicity across a range of concentrations. Cell morphology remained normal at non-cytotoxic concentrations, whereas characteristic shrinkage and fragmentation were observed at cytotoxic doses (Figure 3A).

The hemolysis rate was calculated based on the OD₅₄₀ value of the supernatant. As illustrated in Figures 3B,F, the peptide induced no significant hemolysis at concentrations up to 12 μM. Even at 24 μM and 48 μM, the hemolysis rates remained very low, at 1.76 and 7.63%, respectively, indicating a high hemocompatibility within its effective antibacterial concentration range.

As shown in Figure 4E, fluorescence intensity in all groups reached its peak within 2 min. Notably, all Scymicrosin_7–26 treatment groups exhibited higher fluorescence intensities than the polymyxin B control group, with a clear concentration-dependent increase. These results indicate that Scymicrosin_7–26 can quickly and effectively permeabilizes the outer membrane of P. aeruginosa PA1.

As demonstrated in Figure 4A, treatment with 1 × MIC Scymicrosin_7–26 resulted in nearly complete red fluorescence (indicating dead cells) in PA1 cultures, confirming severe membrane damage. In contrast, untreated control cells exhibited predominantly green fluorescence (viable cells), indicating intact membranes. A similar pattern was observed for MRSA1 (Figure 4B), where exposure to 1 × MIC Scymicrosin_7–26 also induced extensive membrane disruption, as evidenced by the dominance of red fluorescence.

Figures 4C,D illustrates the morphological changes in the representative Gram-negative strain PA1 and Gram-positive strain MRSA1. Untreated control cells displayed smooth, intact surfaces. Following treatment with 1 × MIC Scymicrosin_7–26, PA1 cells exhibited widespread surface wrinkling, while MRSA1 cells showed visible deformation and damage. These morphological disruptions were more severe at 2 × MIC. Interestingly, the surface damage pattern induced by Scymicrosin_7–26 differed from that caused by polymyxin B (which induced vesicle formation on PA1), suggesting a distinct mechanism of action for the antimicrobial peptide.

Ultrastructural changes were further investigated by TEM (Figures 4C,D). Control cells of both strains exhibited intact membranes, dense cytoplasm, and no content leakage. After 1-h exposure to 1 × MIC Scymicrosin_7–26, PA1 bacterial cells displayed visible dissociation of the inner membrane from the cell wall and partial cytoplasmic leakage. At 2 × MIC, cell boundaries became blurred, surface structures were severely compromised, and content leakage was exacerbated. Polymyxin B treatment resulted in cytoplasmic loosening in PA1. For MRSA1, 1 × MIC Scymicrosin_7–26 induced substantial cell lysis and content release, which intensified at 2 × MIC, resembling the effects observed with lysostaphin treatment. These TEM observations corroborate the SEM findings, confirming the membrane-disruptive action of Scymicrosin_7–26.

To investigate possible intracellular mechanisms, we examined the DNA-binding affinity of Scymicrosin_7–26. As shown in Figure 4F, Scymicrosin_7–26 began to retard the migration of PA1 genomic DNA at 24 μM, while MRSA1 DNA showed retardation at 12 μM. The retardation effect intensified with increasing peptide concentrations, suggesting that Scymicrosin_7–26 may contribute to bacterial cell death by binding to genomic DNA.

Reactive oxygen species (ROS) are oxidative molecules produced under cellular stress, which are implicated in cellular damage and can ultimately induce cell death. As shown in Figures 4G,H, treatment with a sub-inhibitory concentration (0.5 × MIC) of Scymicrosin_7–26 already elevated intracellular ROS levels in both PA1 and MRSA1. Dose-responsive ROS generation was detected in both strains at higher peptide concentrations, with ROS levels surpassing those induced by polymyxin B or lysostaphin treatments.

RAW264.7 cell viability was remarkably enhanced by LPS stimulation, reaching 249% of control values (Figure 5A). Scymicrosin_7–26 administration at 3–24 μM concentrations produced a dose-responsive reduction in cellular viability, normalizing it to baseline levels. This observed reduction in cell viability is not a result of cytotoxicity, but rather stems from the inhibition of LPS-induced proliferative signaling, an effect potentially associated with activation of the Akt pathway (Gao et al., 2022). Exposure to 48 μM of the peptide drastically suppressed cell survival to 5.8%. Consequently, 3, 6, and 12 μM doses were chosen for follow-up anti-inflammatory experiments.

Transcript expression of pivotal inflammatory mediators—IL-1β, IL-6, TNF-α, and inducible nitric oxide (iNOS) was measured by RT-qPCR. According to Figure 5B-E, LPS challenge significantly enhanced the transcriptional activity of all four investigated genes. However, treatment with Scymicrosin_7–26 resulted in a concentration-dependent suppression of their mRNA expression. These data suggest that the peptide effectively inhibits the transcription of inflammatory mediators in the established RAW 264.7 inflammation model.

The Griess assay and ELISA were employed to determine the levels of NO and cytokine concentrations (IL-1β, IL-6, TNF-α) in culture supernatants, respectively. Under LPS stimulation, all four inflammatory markers were significantly elevated (Figures 5F–I). Treatment with Scymicrosin_7–26 led to a notable reduction in the production of IL-1β, IL-6, TNF-α and NO, indicating that Scymicrosin_7–26 can effectively attenuate the release of key inflammatory factors in this cellular model.

The anti-inflammatory effects of Scymicrosin_7–26 were further substantiated through examination of iNOS and COX-2 protein expression. As depicted in Figure 5J–L, LPS challenge markedly upregulated both iNOS and COX-2 protein levels, whereas Scymicrosin_7–26 treatment produced a concentration-dependent suppression of their expression. These results demonstrate that the peptide also inhibits the expression of intracellular inflammatory enzymes in macrophages, further supporting its role in modulating inflammatory signaling.

In the LPS neutralization assay, incubation of LPS with Scymicrosin_7–26 across a concentration range of 1.5–48 μM demonstrated no significant difference in neutralization rate compared to the 0 μM control group (Figure 6D). This indicates that Scymicrosin_7–26, at these concentrations, does not neutralize LPS under the applied in vitro conditions.

RAW264.7 cells were stained with the macrophage surface marker F4/80 for the plasma membrane, DAPI for nuclei, and FITC-labeled Scymicrosin_7–26 for the peptide localization. As shown in Figure 6B, no FITC green fluorescence was observed in the control group without Scymicrosin_7–26. At 1.5 μM, faint fluorescent signals began to appear on the plasma membrane and within the cytoplasm of a small number of RAW264.7 cells. With increasing peptide concentrations, both the intensity and distribution of green fluorescence intensified in a dose-dependent manner, showing clear localization to cellular membranes and cytoplasmic regions. These results demonstrate that Scymicrosin_7–26 effectively enters RAW264.7 cells in a concentration-dependent manner within the tested range of 1.5–12 μM.

When stimulated, immune cells produce diverse ROS that not only cause tissue damage but also perpetuate inflammatory cascades (Chen et al., 2020). As shown in Figure 6A, Scymicrosin_7–26 treatment effectively reduced ROS generation in LPS-activated RAW 264.7 cells (Figure 6E).

The MAPK pathway represents a central regulator of inflammation, with its core components—P38, ERK, and JNK—playing critical roles (Haftcheshmeh et al., 2022). The results revealed that LPS treatment significantly upregulated the phosphorylation levels of P38, ERK, and JNK (Figure 6C). Quantitative analysis further confirmed these observations, showing significant elevations in the p-P38/P38, p-ERK/ERK, and p-JNK/JNK ratios (Figures 6H–J). Treatment with Scymicrosin_7–26 concentration-dependently reversed these phosphorylation events, indicating that the peptide exerts its anti-inflammatory effects through suppression of MAPK pathway activation.

The NF-κB signaling pathway serves as another critical regulator of inflammation. Upon LPS stimulation, activation of this pathway promotes the onset of inflammatory responses (Haftcheshmeh et al., 2022). To investigate whether Scymicrosin_7–26 modulates the NF-κB pathway, we examined key phosphorylation events in this signaling cascade. LPS stimulation significantly enhanced the phosphorylation of both p65 and IκBα (Figures 6C,F,G). While total p65 levels remained consistent across groups, Scymicrosin_7–26 treatment progressively reduced phospho-p65 and phospho-IκBα levels in a dose-dependent manner. Additionally, the LPS-induced degradation of IκBα was effectively counteracted by peptide intervention. These collective findings demonstrate that Scymicrosin_7–26 inhibits NF-κB pathway activation in macrophages.

Under stimulation by LPS or other pro-inflammatory factors, the cytoplasmic nuclear factor kappa B subunit p65 (p65) undergoes phosphorylation, transitioning from its non-phosphorylated state. The phosphorylated p65 (p-p65) subsequently translocates into the nucleus, where it binds to specific target genes and regulates their transcriptional expression, thereby modulating the expression of inflammatory mediators and other physiological responses (Florio et al., 2022). Immunofluorescence analysis (Figure 7) revealed minimal nuclear p65 signal in control cells. LPS stimulation induced pronounced p65 nuclear accumulation, whereas Scymicrosin_7–26 treatment significantly reduced p65 nuclear translocation. The nuclear translocation of p65 is a central event in NF-κB pathway activation. The results establish that Scymicrosin_7–26 mediates its anti-inflammatory activity by attenuating NF-κB signal transduction.