The SPC herbal decoction was prepared by the Pharmacy Department of the Affiliated Hospital of Qingdao University. Herbs were purchased from Linqu Pharmaceutical LTD, Ltd., Co.; ShangPharma Holdings Qingdao LTD, Ltd., Co. and Qingdao Shanda Tianyuan Chinese Medicine slices LTD, Ltd., Co. The prescribed dosages of SPC are shown in Supplementary Table S1 . Quality control of SPC was carried out by UPLC-QE- Orbitrap-MS ( Supplementary Figure S1 ).

Main reagents: Ferric chloride hexahydrate (FeCl₃·6H₂O, 98%) and 2-methylimidazole were acquired from Aladdin (Shanghai, China). Benzene-1,4-dicarboxylic acid (H₂BDC, 99%), trimesic acid (H3BTC,98%), ferric nitrate nonahydrate (Fe(NO₃)₃·9H₂O, 98.5%), zinc nitrate hexahydrate (Zn(NO₃)₂·6H₂O, AR) were purchased from Heowns. Acetonitrile and methanol were chromatographically pure, Merck, USA; formic acid was chromatographic alcohol, Thermo Fisher, USA. All solvents and reactants obtained were of analytical grade and used directly without special treatment. Deionized water was used in preparation of various solutions throughout the experiment.

Main instruments: Thermo Scientific Q-Exactive Orbitrap system, Waters Acquity H class type high performance liquid chromatograph, Waters Corporation, USA; Thermo Q Exactive Orbitrap type mass spectrometer with Nano ion source, Xcalibur mass spectrometry workstation, Waters Corporation, USA Thermo Scientific; MS105DU 1-in-100,000 balance, Mettler-Toledo Ltd., Switzerland; Powder X-ray diffraction (PXRD) measurements were recorded on a D/Max-2500 X-ray diffractometer by depositing powder on the glass substrate, 2θ from 2° to 40° with 0.02° increment. The scanning electron microscopy (SEM, HITACHI SU3500) and transmission electron microscopy (TEM, FEI Talos F200X G2) were used to analyze the morphologies.

5 g of each herb was weighed, giving a total of 50 g. The herbs were crushed into granules using a high-speed Chinese medicine crusher and passed through a 40 mesh Chinese medicine sieve. Each batch of fresh herbs was extracted three times by continuous dynamic countercurrent extraction. The procedure was as follows: fresh herbs and solvent (5 times 50% ethanol) were placed in the extraction tank for the first extraction, and the extracts was collected after 2.5 hours. Then the solvent (5 times 75% ethanol) was added to the first extraction and the second extraction was carried out, and the extracts was collected after 2 hours. Finally, the second extract was mixed with solvent (5 times 90% ethanol) and the third extracts was collected after 1 hour of extraction. The extracts are filtered, concentrated and dried, and the paste percentage of the drug is determined. No separation was carried out to avoid loss of active ingredients.

The above three extracts were combined and kept in a refrigerator at 4 °C for 24 h (total 750mL). The supernatant is obtained by centrifugation at 10000 r/min for 20 min at room temperature and filtered through 0.45 μm hollow fiber membrane. The filtrate was collected and divided equally into 4 portions. Each portion contained 12.5 g of raw herbs. Take appropriate amount of chitosan powder, add 1% glacial acetic acid to prepare 1% chitosan solution, ready for use. The alcohol extracts are concentrated under vacuum until the ratio of the amount of raw herbs (g) to the volume of drug solution (mL) is 1:8. Then it is placed on a 50 °C constant temperature magnetic stirrer and stirred for 10 min. Chitosan solution (1.6 mL for every 1 g of raw herbs) is added to the alcoholic extracts and continue stirring for 1 min. Finally the solution was taken out and left at room temperature for 12 h. Observe and record the clarity of the solution and the shape of the precipitate.

MIL-101(Fe) (Peng et al., 2022), MIL-100(Fe) (Fang et al., 2018), and ZIF-8(Zn) (Wang et al., 2023) were prepared according to the reported procedures in the literatures.

MIL-101(Fe) was synthesized from a mixture of 0.68 g (2.5 mmol) FeCl₃·6H₂O and 0.21 g (1.25 mmol) H₂BDC in 15 mL DMF. The suspension was sonicated to complete dissolution and then was sealed into a Teflon-lined stainless steel autoclave and heated at 110°C for 24 h. The orange solid was obtained by centrifugation and washed by DMF and EtOH for three times to remove the unreacted raw materials. Finally, the product was dried overnight at 70 °C.

MIL-100(Fe) was synthesized from a mixture of 2.4 g (6 mmol) Fe(NO₃)₃·9H₂O and 0.84 g (4 mmol) H₃BDC in 6 mL deionized water. The mixture was stirred at room temperature for 1 h and then was heated at 160°C for 12 h. The solid product was centrifuged and washed with deionized water and EtOH for three times for purification. Finally, the resultant MIL-100(Fe) was dried overnight at 90 °C.

ZIF-8 Zn(NO₃)₂·6H₂O (2.0 g) was added into 40 mL methanol and stirred until it was completely dissolved. Next, 2-methylimidazole (2.0 g) was added to the above solutions and stirred. Then a white solid was obtained after stirring for two hours and leaving overnight at 60 °C. The final ZIF-8 was made by drying and washing it three times with methanol.

Gram-negative E. coli ATCC 11229 and gram-positive S. aureus ATCC 6538 were selected as model microorganisms to evaluate the antimicrobial activities. Bacteria were incubated overnight at 37 °C 220 rpm. The bacterial concentration was then adjusted to ~10^6 CFU/mL using LB medium for subsequent experiments. Bacterial concentration is usually determined by the optical density of the bacteria suspensions at 600 nm (OD600 nm).

S. aureus in growth phase (OD600 = 0.6) was co-incubated with MIC under SPC for 6 h. The untreated group served as a control. Bacteria were harvested at 10,000 rpm and used as samples for metabolomics analysis. First, 10 mg solid sample were accurately weighed, and the metabolites were extracted using 80 µL methanol: water (4:1, v/v) solution. The mixture was allowed to settle at -10 °C and treated by high throughput tissue crusher at 50 Hz and followed by ultrasound at 40 kHz. After centrifugation at 13000 g, the supernatant was transferred to sample vials for analysis. Ultra-performance liquid chromatography - mass spectrometry (UPLC-MS) analysis was performed by Majorbio Bio-Pharm Technology (Shanghai, China). Chromatographic column: Infinity Lab Poroshell 120EC-C18 (150×2.1mm, 2.7μm) column, mobile phase: 0.1% formic acid water (A) - acetonitrile (B); Flow rate: 0.2 mL-min^-1; Injection volume: 2 µL; Column temperature: 35 °C; Elution method: 0~5 min, 10% B; 5~10 min, 10%-21% B; 10~20 min, 21% B; 20~30 min, 21%-70% B; 30~35 min, 70% B. Mass spectrometry conditions: positive and negative ion mode detection, sheath gas flow rate 40 psi (1 psi = 6.895 kPa), auxiliary gas flow rate 10 psi; spray voltage 3.8 kV; capillary temperature 320°C; auxiliary gas temperature 350°C; mass scan range m/z 100–1500, detection resolution 70,000 FWHM. Ionization energies 20, 30, 40 eV, resolution 17500. The samples of the control, SPC-treated, and QC groups were subjected to UPLC for component separation; a single component entered the ion source of the high vacuum mass spectrometer for ionization and was separated according to the mass-to-charge ratio (m/z) to obtain a mass spectrum. Finally, the mass spectra of the samples were analyzed. The qualitative and quantitative results of the samples were obtained. The LC-MS data from sample pellets were processed using the Majorbio Cloud Platform (Shanghai, China), and metabolites were identified. Normalized data were visualized by partial least squares-discriminant analysis (PLS-DA) using the ropls package in R. The ellipses in the PLS-DA plots were employed to characterize metabolic perturbations among groups in a Hotelling T2 region with a 95% confidence interval threshold. The variable importance in projection (VIP) was calculated based on the PLS-DA model to identify significant metabolites with a VIP >1.0 and P-value <0.05. The KEGG (http://www.kegg.com/) database was used to explore related metabolic pathways. The detailed methods can be seen in the Supplementary Information .

The SPC and MOFs were subjected to an antimicrobial experiment using the diffusion method described by Fiebelkorn et al (Fiebelkorn et al., 2003). The K–B test, commonly known as the agar disc diffusion method, is a classical method to evaluate antibacterial activities of samples through the average inhibition diameter, which is obtained by placing antibacterial agents in an agar plate containing a certain concentration of bacterial suspension and then incubating. Typically, powders of MOFs were pressed into discs. The discs were placed firmly on the agar plates which were uniformly covered with E. coli. All the plates were incubated at 37°C for 24 h, following which the zone of inhibition was measured. These data represent the average of at least three different independent experiments.

The MICs of SPC were determined by a microtiter broth dilution method (Ruparelia et al., 2008). The SPC solutions were diluted in a 96-well plate with LB medium (62.5, 31.25, 15.6, 7.8, 3.9, 1.95, 0.98 mg/mL), bacteria of the same concentration (~10^6 CFU/mL, 100 μL) were added to each well and cultured in a thermostatic incubator (37 °C) for 16–18 h. The final volume of each well in the 96-well plate is 200 μL. Ampicillin was employed as positive control, bacterial grown without treatment as negative control and LB medium as blank control. MIC was measured by the optical density values at 600 nm and minimum bactericidal concentration (MBC) was calculated by standard plate count method. These data represent the average of at least three different independent experiments.

S. aureus suspension was cultured in liquid medium until OD600 = 0.5, and 2 mL of each group was centrifuged to remove the supernatant. To the experimental group, 2 mL of SPC (MIC) solution was added and incubated for 6 h at 37 °C on a shaker. LB medium was used to replace SPC extracts in the control group, and other operations were the same as the experimental group. The bacterial solution of each group was centrifuged (5500 rpm) for 12 min, and washed 3 times with PBS. Then the S. aureus obtained by centrifugation in each group was fixed with 2.5% glutaraldehyde at 4°C for 5 h and was dehydrated with gradient ethanol (50%, 70%, 80%, 90%, 100%) for 15min. Subsequently, the morphological changes of bacteria were observed by SEM and TEM (Garcia Maset et al., 2022).

The growth curve of E. coli after MOFs treatment was determined using a method similar to that described in (Rasool et al., 2016). The E. coli suspension was resuspended to 1.0×10^6 CFU/mL, and different concentrations of MOFs were added to each group and cultured on a shaker at 37 °C. The absorbance at 600 nm was measured every 3 hours using a microplate reader. Finally, the appropriate concentration of MOF was obtained. The combined antibacterial activity of SPC and MOF can also be expressed by the bacterial growth curve. In the same way as above, SPC (final concentration of MIC) and MOF (final concentration based on screening) were added to E. coli solution (1.0×10^6 CFU/mL) to determine the absorbance every 3 h. PBS was added as a positive control group and culture medium group as a negative control group. These data represent the average of at least three different independent experiments.

All experiments were repeated at least three times. Differences between groups were assessed using Student’s t-test, one-way analysis of variance (ANOVA) and Tukey’s multiple comparison test. All analyses were performed on GraphPad Prism. Data were expressed as means ± standard deviations (Mean ± SD). P< 0.05 was considered as statistical significance. ***indicates p<0.001; **indicates p<0.01; *indicates p<0.05.

According to previous research methods, the antibacterial activity of Gram-negative strains of E. coli and Gram-positive strains of S. aureus was determined using the disk diffusion method. As shown in Figure 1A , the SPC extracts produced an inhibition zone (clear areas with no bacterial growth) against S. aureus but had little effect on inhibiting the growth of E. coli. This suggests that SPC primarily inhibits the proliferation of S. aureus. To gain deeper insights into the biological effects of SPC, we conducted antibacterial experiments using the broth microdilution method and standard plate counting method. Figure 1B demonstrates that SPC exhibited dose-dependent inhibition on the proliferation of S. aureus. The minimum inhibitory concentration (MIC) value of SPC was determined to be 7.8 mg/mL (crude drug concentration), and the minimum bactericidal concentration (MBC) value was 15.6 mg/mL (crude drug concentration). The chemical composition of SPC was characterized using UPLC-MS/MS ( Supplementary Figure S1 ). Six compounds were identified in SPC by comparing the ion information and retention times between the multistage mass spectrometry fragments with the relevant information in the literature and databases ( Supplementary Table S2 ). At a SPC concentration of 7.8 mg/mL, the concentrations of the six components were Chlorogenic acid: 7 μg/mL; Catechin: 45.4 μg/mL; Columbianetin: 1.2 μg/mL; Nodakenin: 5.2–156 μg/mL, Imperatorin: 15.5 μg/mL; Osthol: 37.4 μg/mL. Since the antimicrobial activity of SPC extracts may be the result of a combination of several components, the final choice in this study was to use the crude concentration of SPC as the MIC, i.e. 7.8 mg/mL. Several antibiotics are effective against S. aureus. So we chose ampicillin (AM) as a positive control drug and determined the MIC and MBC of AM against S. aureus using the same test method ( Supplementary Figure S2 ), with values of 1 μg/mL (MIC) and 2 μg/mL (MBC), respectively.

Changes in bacterial morphology after SPC treatment can be observed by SEM and TEM. SEM images revealed that untreated S. aureus displayed a smooth surface with 1 μm in diameter ( Supplementary Figure S3 ). However, after treatment with SPC, the bacteria exhibited irregular morphology and damaged cell walls. TEM images further confirmed these observations, showing that untreated S. aureus had uniform morphology, while S. aureus co-cultured with SPC exhibited transparent cells with blurred boundaries and cytoplasmic leakage ( Figure 1C ). SYTO 9 green fluorescent stain and PI red fluorescent stain are two common nucleic acid stains with different ability to penetrate bacteria. SYTO 9 stains all bacteria in a population regardless of cell membrane integrity. PI is a membrane-impermeable nucleic acid stain that can only penetrate bacteria with damaged cell membranes and is used to assess bacterial cell membrane integrity. As shown in Figure 1D , little red fluorescence was observed in the fluorescence images of the untreated group, in contrast, the intensity of red fluorescence was increased in S. aureus treated with SPC. This proved that most of the S. aureus in the control group was alive and the PI was blocked outside the cell membrane, whereas SPC was able to disrupt the cell membrane of S. aureus, leading to the production of red fluorescence.

S. aureus is a pathogenic bacterium known for its remarkable ability to adapt to various environmental conditions, including high temperatures and high salt concentrations. In nature, S. aureus often forms biofilms, which are structured communities of bacterial cells enclosed in a self-produced matrix of extracellular polymeric substances (EPS). Considering that biofilm formation may increase the ability of S. aureus to resist external disturbances, we conducted an investigation to evaluate the inhibitory effect of SPC on biofilm formation. The formation ability of S. aureus biofilm was assessed using the Crystal violet staining method. As the culture time was extended, the absorbance steadily increased and reached its peak at 72 hours, indicating that the biofilm had fully matured after three days ( Figure 2A ). The control group displayed a deep purple color due to the interaction between crystal violet and negatively charged surface molecules in the extracellular matrix, implying the formation of a relatively complete biofilm ( Supplementary Figure S4 ). However, the color of the SPC-treated group was noticeably lighter, revealing that SPC had the ability to inhibit biofilm formation. To quantify the biomass of the biofilms treated with various concentrations of SPC (ranging from 0 to MIC), crystal violet colorimetry was performed. The results, as shown in Figure 2B , demonstrated that SPC significantly reduced bacterial attachment in a dose-dependent manner. At 1/2 MIC, SPC exhibited a strong inhibition of biofilm formation, with an inhibition rate exceeding 80%. For mature biofilms, SPC demonstrated the ability to remove approximately 10.6% of the biofilm at MIC. As the concentration increased (up to 3 MIC), the biofilm clearance reached 50.2% ( Figure 2C ). Simultaneously, the same experiment was performed for AM and it was found that biofilm formation was almost completely inhibited at MIC concentrations, with 70.4% inhibition rate at 1/2MIC. In the biofilm removal ability, the performance of AM was greatly weakened. When the concentration of AM was increased to 3MIC, the removal rate of biofilm was only 20.5%, which may be due to the fact that AM was not able to penetrate into the extracellular matrix of the bacteria and could not play an effective role ( Figures 2B, C ). These findings hold promise for the eradication of S. aureus in diverse scenarios. To better visualize its effects, microscopic images were captured to support the biofilm inhibition and clearance of SPC. The results shown in Figure 2D align with the aforementioned conclusions. Supplementary Figure S5 displayed microscopic images of the AM treatment group. To further analyze the impact of SPC on biofilm eradication, the average thickness of the biofilm formed on Petri dish surface was analyzed using 3D CLSM. The results, depicted in Figure 2E , showed that while S. aureus can easily form a certain thickness of biofilm on the surface of Petri dish, the biofilm in the SPC-treated group was much thinner. Simultaneously, the results obtained from fluorescent staining demonstrated that SPC exerted a concentration-dependent bactericidal effect on the bacteria within the biofilm. Additionally, we also examined the inhibition effect of SPC on biofilm formation by 3D CLSM and investigated the impact on the secretion of protein, which is the main component of S. aureus biofilms. Supplementary Figure S6 displayed that the group treated with SPC could hardly form biofilms and Supplementary Figure S7 revealed a significant reduction in the levels of protein after treatment with SPC, which was consistent with the inhibitory effect on biofilm formation.

The metabolite profiles of S. aureus samples were analyzed using LC-MS. Through this analysis, we identified a total of 267 differential metabolites in the SPC-treated group compared to the control group. The significance of these metabolites was determined using VIP based on the PCAs model ( Figures 3A, B ). Among the differential metabolites, the content of L-Proline, L-Asparagine, Prolylhydroxyproline, and Pyroglutamic acid in S. aureus metabolites was found to be higher in the control group than in the treated group (P-value<0.001, Figure 3B ). The identified metabolites were further classified into different tiers based on the number of metabolites, as displayed in descending order (superclass shown in Figure 3C1 , class shown in Figure 3C2 ). To gain a deeper understanding of the metabolic pathways affected by SPC, we utilized the KEGG pathway database, which describes molecular interactions, physiological and biochemical reactions, and relationships between gene and products. Three metabolic pathways were identified: Metabolism, Genetic Information Processing, and Environmental Information Processing. Among these, amino acid metabolism and lipid metabolism were the most abundant metabolites annotated in the classification of metabolism pathway ( Figure 3E ). Additionally, the identified metabolites were categorized into various groups based on their biological functions, such as organic acids, lipids, carbohydrates, nucleic acids, peptides, vitamins and cofactors, steroids, hormones and cofactors, and antibiotics ( Figure 3F ). KEGG pathway enrichment analysis revealed significant differences in the enrichment rates of Linoleic acid metabolism and Alanine, aspartate, and glutamate metabolism between the two groups (P-value<0.001, Figure 3G ).

The above experiments confirmed the weak inhibitory effect of SPC on E. coli ( Figure 1A ). Therefore, to overcome these limitations, we introduced MOFs with antibacterial effects. Three types of MOFs (ZIF-8, MIL-100(Fe) and MIL-101(Fe)) were selected as collaborative materials for SPC. The successful synthesis of ZIF-8, MIL-100(Fe) and MIL-101(Fe) was confirmed through PXRD, TEM, and SEM, which was in full agreement with previously reported results ( Figures 4A–C ; Supplementary Figure S8 ).

These MOFs’ effects on E. coli and S. aureus were tested using the agar disk diffusion method. The MOFs were sterilized, pressed into circular sheets, and placed in LB medium containing bacteria for culturing. The results, shown in Figure 5A , indicated that all three MOFs exhibited inhibitory effects on both E. coli and S. aureus. Additionally, the relative bactericidal activity was evaluated through bacterial growth kinetics, and the concentration suitable for the combination of SPC and MOFs was determined ( Supplementary Figure S9 ). Furthermore, the effect of the combined system on E. coli, which was not sensitive to SPC, was tested. The combined system was prepared by simply mixing the appropriate concentration of SPC and MOFs. As depicted in Figure 5B , under the same treatment conditions, the mixed system had a significant effect on the growth kinetics of bacteria compared to the SPC group, indicating that the introduction of MOFs in SPC was helpful in expanding the antibacterial spectrum of SPC.