Fresh healthy leaves of Reseda arabica were collected from Al-Ahsa Oasis in Saudi Arabia’s Eastern region (25°20002.900 N 49°34039.900 E). The samples were washed with running tap water within 24 h of collection, then washed with double-distilled deionized water to remove surface debris. The cleaned leaves were allocated into 1 cm × 1 cm-size segments. Surface sterilization was carried out under aseptic conditions with succeeding washes in 70% ethanol, 4% sodium hypochlorite solution, sterilized distilled water, and finally allowed to air dry in the laminar airflow chamber (Dobranic et al., 2011). The sterilized leaf segments were added to potato dextrose agar (PDA) supplemented with 100 μg/mL streptomycin to prevent bacterial growth and incubated for 5–7 days at 28°C until endophytic fungi start to develop hyphal filaments. The growth was observed daily, and individual hypha tips were picked out on time and transferred to a new PDA medium until pure fungal colonies were obtained. Morphological identification of the isolated endophytic fungi was performed at the Regional Center for Mycology and Biotechnology, Al-Azhar University, Cairo, Egypt. All isolated endophytic fungi were screened for their antibacterial activities in addition to the ability of their extracts to reduce AgNO₃ into AgNPs (data not shown). A. parasiticus EA-1 (RCMB 150) displayed significant activity against MRSA and powerful potential to reduce AgNO₃ into AP-AgNPs. Therefore, it was further selected for the mycosynthesis of AP-AgNPs.

Sequence analysis of the ITS1-5.8S-ITS2 and ITS regions of the ribosomal ribonucleic acid (RNA) gene was performed to confirm the morphological identification of the A. parasiticus EA-1 (RCMB 150) isolate. Briefly, the total fungal genomic DNA was directly obtained from fungal mycelia. The isolated DNA was subjected to the polymerase chain reaction (PCR, Applied Biosystems, USA) using primers ITS1: TCCGTAGGTGAACCTGCGG and ITS4: TCCTCCGCTTGATATGC (White Bruns et al., 1990). Then, the BigDye Deoxy Terminator Cycle Sequencing Kit (Applied Biosystems, Darmstadt, Germany) was used to purify the amplified product, and the sequence was obtained by using an automated DNA sequencer (ABI PRISM 3700). The developed fungal sequence was compared with previous species sequences in NCBI GenBank. The identification of endophytic fungus was performed depending on the homology of amplified sequences to those in the GenBank database. The relevant accession number of isolate was obtained using MEGA version 7.0, and the phylogenetic analysis of sequences was also created.

Fungal isolate was cultured on potato dextrose broth (PDB) medium and kept at 25°C for 20 days in a shaking (150 rpm) incubator and extracted by using ethyl acetate. Filtrate was mixed with an equal volume of ethyl acetate, placed on a vortex shaker for 15 min, and kept for 4 min until the two clear separate layers. The ethyl acetate layer was separated from the aqueous layer using a separating funnel. The collected phase was evaporated using the oven at 65°C. Finally, DMSO (1 mg/mL) was used to dissolve the fungal crude extract and then stored at −20°C until further experiments.

Qualitative screening of fungal secondary metabolites (alkaloids, flavonoids, glycosides, steroids, terpenoids, tannins, saponins, and reducing sugars) was carried out following the methods of Gul et al. (2017).

The metabolites existing in the ethyl acetate extract of A. parasiticus were examined, counted, and identified using GC–MS. Briefly, the fungal extract was dissolved in spectroscopy-grade methanol. GC–MS analysis was done using Trace GC1310ISQ mass spectrometer (Thermo Scientific, Austin, TX, USA). The column oven temperature was 50°C at the start and increased at a rate of 5°C/min to 230°C and then held for 3 min, and then raised to the final temperature of 290°C and kept for 3 min. The sample (1 μL) was injected at 250°C utilizing helium as a carrier gas, split at the ratio of 1:30. Mass spectrometer functioned in the electron ionization mode at 200°C at 70 eV with a scan range of 40–1,000 m/z. The spectrum of the identified compounds was compared with the spectrum of the known compounds stored in the WILEY 09 (Wiley, New York, NY, USA) and NIST 11 (National Institute of Standards and Technology, Gaithersburg, MD, USA) library.

Ethyl acetate extract of A. parasiticus was obtained as described previously, and 200 μL of 0.5 M AgNO3 solution was added to 100 mL of fungal extract to obtain a final concentration of 1 mM AgNO3. The mixture was incubated at 25°C; 150 rpm for 2 days in the dark. Then, the mixture was checked for brown color, which indicated the formation of AP-AgNPs. The particles were harvested by ultracentrifugation at 5°C, 15,000 rpm for 10 min, and freeze-dried using benchtop lyophilizer. A schematic diagram of the mycosynthesis mechanism of AP-AgNPs is displayed in Supplementary Figure S1B.

The biosynthesis of AP-AgNPs was confirmed by recording the absorption spectra using UV–Vis spectroscopy at a wavelength of 300–700 nm (Elico UV–visible double-beam spectrophotometer). X-ray diffraction (XRD) analysis was performed by XPERT-PRO using monochromatic Cu-Ka radiation (k = 1.5406 A°) functioned at 40 kV and 30 mA at a 2 h angle design. The scanning was done in the region of 208–808. The images obtained were compared with the Joint Committee on Powder Diffraction Standards (JCPDS) library to explain the crystalline structure. Fourier transform infrared spectroscopy (FTIR) is used to identify the functional groups of biological molecules accountable for the biosynthesis and capping of AP-AgNPs. Potassium bromide was added to the dried AP-AgNP powder (100:1), and the FTIR spectrum was recorded using an FTIR Spectrophotometer (PerkinElmer) in the range of 450–4,000 cm⁻¹. TEM (TEM HF-3300—Hitachi High-Tech Canada, Inc.) is used to describe the shape and size of AP-AgNPs. The size distributions from the TEM images for AP-AgNPs were subsequently examined using ImageJ Freeware Version 1.53d downloaded from the NIH website.¹ The zeta potential of the biosynthesized AP-AgNPs was determined by a nanoparticle analyzer (Zetasizer Ver. 7.13, Malvern Instruments Ltd., UK). Energy-dispersive X-ray spectroscopy (EDX) at 20 keV was used to confirm the occurrence of nano-silver elements.

The bacteria used in this study were methicillin-resistant staphylococcus aureus (MRSA) ACLT 32571 clinical isolate obtained from Kasr Al-Aini Hospital, Cairo University, Cairo, Egypt.

MRSA suspension was fixed by 1% glutaraldehyde and then dehydrated in alcohol solutions (30, 50, 70, 80, 90, and 100%) and allowed to air dry and finally coated with Au/Pd (Quorum Q150T ES vacuum coater). Pictures were acquired by the SEM Merlin (Carl Zeiss, Jena, Germany), working at an accelerating voltage of 15 kV, SE-detector.

The effect of AP-AgNPs on membrane permeability was assessed as described (Marri et al., 1996). 10 μL of 1x AP-AgNPs, 10 μL of ortho-nitrophenyl-galactoside (ONPG), and 30 μL of the bacterial suspension were mixed in triplicate in the wells of a 96-well microtiter plate. Triton X-100 was used as the positive control. Samples were incubated at 37°C for 3 h, and the OD was measured every 30 min at 405 nm.

The red blood cells were counted and diluted to a concentration of 10⁷–10⁸ cells/ml and incubated at room temperature for 1 h with serially diluted AP-AgNPs at a ratio of 4:1 in a final volume of 100 μL. The optical density at 570 nm was measured with a microplate reader. HC₅₀ values were obtained following the method of Cao et al. (2012). Saline solution at a concentration of 0.85% was used as the negative control, and 0.1% Triton X-100 was used as the positive control.

The leakage of intracellular proteins and nucleic acids was assessed as described (Moghimi et al., 2016). Briefly, the bacterial suspension was centrifuged at 7000 rpm for 4 min, filtrated, and examined at the absorption of 260 nm and 280 nm using an ultraviolet spectrophotometer (Agilent, USA).

MRSA cells were cultured in Brain Heart Infusion (BHI) culture medium for 24 h at 37°C and were centrifuged at 3000 rpm. Subsequently, cells were washed with PBS buffer and fixed overnight in 2.5% glutaraldehyde and 4% paraformaldehyde in 0.1 M cacodylate buffer, with pH 7.2. Then, the cells were fixed for 30 min in 1% OsO4, with dehydration in acetone, and embedded in a solution of Polybed 812. Finally, ultra-thin sections were selected on 300-mesh copper grids and stained with uranyl acetate (5%) and lead citrate (1%). The images were photographed with an FEI Tecnai G2 Spirit transmission electron microscope, working at 120 kV.

The bacterial cell survival was assessed after 24 h of treatment with AP-AgNPs and observed by CLMS on an inverted Carl Zeiss LSM 780 confocal laser scanning microscope (Carl Zeiss, Jena, Germany). The samples were stained for 20 min with acridine orange (green fluorescent) and propidium iodide (red fluorescent) to distinguish between undamaged and cell surface damaged bacteria.

PI staining using flow cytometry was used to determine the cell membrane integrity following the method of Ning et al. (2017). The MRSA suspensions were washed with PBS, and the bacterial precipitate was re-suspended by adding 100 μL 1× binding buffer. Subsequently, 10 μL PI staining solution was added to the suspensions and mixed gently. After 10–15 min in darkness at room temperature, samples were measured by flow cytometry (FACS CantoI1, USA) in 1 h.

MRSA cells (1.5 × 10⁸ CFU/mL) were mixed with AP-AgNPs (25 to 200 μg/mL) in LB broth in 96-well plates, and cells were incubated for 24 h at 37°C. After incubation, each well was rinsed three times with sterile 0.85% saline by discarding the culture supernatant, and then, the adherent biofilm was fixed with methanol and stained with 0.2% crystal violet (7 min). Cell growth was determined by measuring the OD at 595 nm (Shi et al., 2023). AP-AgNPs mixed in polystyrene 24-well plates with bacterial suspensions (1.0 × 10⁶ CFU/mL) at different concentrations in LB broth. After 2 days of incubation, wells were washed with a sterile 0.85% saline, and a 0.01% final concentration of resazurin was added, followed by an incubation in the dark for 1 h. A microplate reader (Thermo Fisher Scientific, Waltham, MA, USA) was used to measure the optical density (OD 570 and OD 600) values (Paduszynska et al., 2019). Furthermore, we performed additional experiments on the inhibitory potential of AP-AgNPs in the establishment of MRSA biofilms following the method proposed by Hu et al. (2024). Briefly, pictures of biofilms were taken for control cells and for cells treated with AP-AgNPs at concentrations of 25 μg/mL (MIC), 50 μg/mL, or 100 μg/mL; 2 mL of MRSA-diluted overnight culture was used to grow biofilms on coverslips for 48 h. The cover slides were then washed with PBS, transferred to a new plate, and exposed to AP-AgNPs for 48 h. The cover slides were washed again with PBS and stained with a LIVE/DEAD BacLight Bacterial Viability Kit (Invitrogen Molecular Probes, Eugene, OR, USA). CLSM pictures were taken using an Olympus FV1000 confocal laser scanning microscope (Olympus, Tokyo, Japan) with a 60× objective lens. For the examination of SYTO 9 (green channel), we used 488 nm excitation and 520 nm emission filter settings. For PI detection (red channel), we used 543 nm excitation and 572 nm emission filter settings. Picture investigations were carried out using Fluoview version 1.7.3.0 software.

The in vivo Tape-Stripping Mouse Model was approved and licensed by the Research Ethics Committee, Deanship of Scientific Research at King Faisal University (KFU-REC-2022-OCT-ETHICS218). Male BALB/c mice (n = 24, 2 months old) were housed under conventional conditions of 25 ± 2°C and 12 h dark/light cycle, with a standard diet and water ad libitum.

The mice were anesthetized by a combination of 100 mg/mL ketamine and 20 mg/mL xylazine, and the back hair was shaved. Small pieces of elastic bandage tape were applied and immediately removed from a 2 × 2 cm² dorsal area for 10 to 15 times until the skin became enlarged and ruddy without bleeding. Subsequently, a 10 μL-MRSA inoculum (1 × 108 CFU/mL) was applied to wounds (Tatiya-Aphiradee et al., 2016). Mice were divided into four groups (n = 6 per group) as follows: Group 1, non-infected and non-treated (negative control group). Group 2, MRSA-infected group with saline. Group 3, MRSA-infected group treated topically to the wound area with a 100-μL aliquot of 10% ethanol in propylene glycol (vehicle) and 160 μg/mL of vancomycin daily for 9 successive days (as a positive control group). Group 4, the MRSA-infected group treated topically with a 100-μL aliquot of each sample (10% ethanol in propylene glycol) (vehicle), 100 μg/mL of AP-AgNPs daily for 9 successive days. Wounds photographed on days 2, 4, 6, and 8.

C-reactive protein (CRP) was evaluated with enzyme-linked immunosorbent test (ELISA) kits (R&D Systems, Kamiya Biomedical Company), following the manufacturer’s instructions.

All values expressed as mean ± SD (standard deviation) of at least three independent experiments. Power calculation was performed for using unpaired two-samples Student’s t-test (two-tailed) assuming equal variation in the two groups. Differences were considered statistically significant at *p < 0.05 and **p < 0.005.

Genotypic methods and morphological characterization were used to verify the identification of the most promising endophytic fungal isolate, EA-1 (RCMB 150), A. parasiticus. Depending on the partial sequence of the 18S ribosomal RNA gene; internal transcribed spacer 1, 5.8S ribosomal RNA gene, and internal transcribed spacer 2; the complete sequence and the partial sequence of 28S ribosomal RNA gene, the fungal sequence was submitted to NCBI GenBank database with accession number PQ460365. Higher similarity sequences were selected, MEGA 7.0 was used to construct a phylogenetic tree depending on the maximum probability method (Supplementary Figure S1A), and the isolate was closely related to a cluster of A. parasiticus according to BLAST analyses.

To identify the fungal secondary metabolites with antibacterial activity in A. parasiticus extract, we performed GC–MS analysis. The results of the screening are given in Table 1, which revealed some secondary metabolites that are present in the ethyl acetate extract of A. parasiticus. GC–MS analysis showed a demonstrative spectral output of each one of the compounds present in the examined samples. The results from the analysis of ethyl acetate extract of A. parasiticus determined that the average yield of the extract is approximately 16 major compounds as shown in Figure 1 and Table 2. 1,2-Benzenedicarboxylic acid, di-iso-octyl ester; 9-octadecenoic acid, E; and hexadecanoic acid were the major compounds with ratios 25.88, 16.93, and 8.31%, respectively.

Aspergillus parasiticus-based AgNPs (AP-AgNPs) were mycosynthesized using ethyl acetate extract of the endophytic fungus A. parasiticus isolated from the leaves of Reseda arabica plant. The color of the reaction mixture (mycelial extract and AgNO3) was detected at regular intervals, which changed from white to yellowish brown after 48 h (Supplementary Figure S1B). This color change is attributed to the process of excitation of surface Plasmon resonance within the biologically produced AP-AgNPs. The biosynthesis of AP-AgNPs in the aqueous solution was identified by recording the absorption spectra using UV–Vis spectroscopy. The maximum peak of fungus-mediated AP-AgNPs biosynthesis was observed at 434 nm (Figure 2A). The XRD pattern obviously displays the major peaks at (2-theta) 38.19, 44.37, 64.56, and 77.47 corresponding to the (111), (200), (220), and (311) planes, respectively (Figure 2B). In addition, two unassigned peaks were found at 32.25° and 46.21°. Unpredicted crystalline structures (32.25° and 46.21°) are also present.

The FTIR examination of AP-AgNPs showed strong peaks at 3421.1, 2924.41, 1633.96, 1384.56, 1073.63, and 617.36 cm⁻¹ (Figure 2C). These peaks were equivalent to N-H stretching of primary amine of the protein, alkane C-H stretching, the stretching of conjugated alkane C=C, methylene tails of the protein, C-N of aliphatic amines of polyphenols, and O-H stretching, respectively. TEM examination of the size of AP-AgNPs revealed that the shape of AP-AgNPs is spherical, and the average particle size is 27.7 nm. The diameter range is 5.34–72.12 nm, and the geometric mean equal value is 24.93 ± 1.60 nm (Figures 3A,B). The stability of AP-AgNPs was assessed by zeta potential (Figure 3C). The zeta potential value was negative (− 25.4 ± 3.13 Mv), suggesting the good colloidal nature of biosynthesized nanoparticles. Energy-dispersive spectroscopy (EDS) was applied to obtain qualitatively evaluations of chemical compositions in nanoparticle samples; however, it is able to provide semi-quantitative results. The EXD (or EDS) results in Figure 3D showed variable intensities of signals at the X-axis of the several binding energies (keV), confirming the presence of diverse elements in the sample. A strong signal at 3 keV showed the occurrence of metallic silver in AP-AgNPs. Other weak-to-moderate signals were found in addition to the above-mentioned signals. These comprised the occurrence of oxygen and carbon because of the preparation of the sample using a copper grid coated with carbon in addition to the occurrence of carbon due to capping biomolecules. Figure 3E of the EDX spectrum displayed the elemental composition present except AP-AgNPs in the sample. The spectrum displayed that silver, constituting 53.29% of the total, was the greatest important element, followed by carbon (33.5%) and oxygen (13.21%), representing the establishment of AP-AgNPs.

We examined the anti-MRSA activity of AP-AgNPs using the agar well diffusion method. The AP-AgNPs showed higher anti-MRSA action as compared to vancomycin where the diameter of the inhibition zone (IZD) was 48 and 42 mm, respectively (Table 3; Figure 4A). The MIC values of AP-AgNPs and vancomycin were 25 and 50 μg/mL, respectively (Table 3). The experiments of killing kinetics established there was a significant reduction in cell quantity after adding AP-AgNPs for several hours. Precisely, the curve of killing kinetics in Figure 4B revealed that it would take 2 h at least to recognize 1 log decrease in live cells, confirming the higher efficacy of AP-AgNPs as a potent anti-MRSA agent.

The morphological alterations of cellular membranes of MRSA treated with AP-AgNPs for 24 h were examined by scanning electron microscopy (SEM). The untreated cells had intact, smooth, and spherical morphology (Figure 5A). The MRSA cells treated with 12.5 μg/mL (1/2 MIC) of AP-AgNPs displayed a moderate rise in the number of lysed cells. The treatment with higher concentrations of AP-AgNPs (25 and 50 μg/mL) resulted in a roughened bacterial surface with boss-like protuberances and unusual protrusions that were not detected in the control cells. In addition, these cells showed severely changed surfaces and noticeable cleavage sites (Figure 5A). Several lysed cells complemented by cellular debris and discharge of intracellular components could be perceived.

As shown in Figure 5B, non-treated MRSA displayed homogeneous cell walls without any obvious modifications and with complete cytoplasm as assessed using transmission electronic microscopy (TEM). In addition, non-treated MRSA showed the distinctive structures of Staphylococci morphology: rounded cells with a completely thick cell wall envelope and distinct membranes (Figures 5B,a–d). The cytosol showed a consistent electron density and proliferating cells with a central septum. The cell wall presented a tripartite structure consisting of an outer that displays an extremely stained fibrous surface, an intermediate transparent region, and an electrodense inner thin zone. The plasma membrane was detected immediately below this electrodense layer of the wall (Figures 5B,a–d). On the other hand, despite MRSA treated with AP-AgNPs showing rounded morphology and consistent electron density in the cytosol (Figures 5B,e–h), some cells presented modifications in the cell wall: without a tripartite-layers structure and no noticeable electrodense inner thin zone. In these cells, the observation of the plasma membrane was easier, rather than those under control conditions. Alterations in the outer extremely stained fibrous surface and intermediate transparent area of the cell wall were also detected.

To investigate the bacterial cell membrane damage by AP-AgNPs, fluorescent microscopy was conducted in the presence of propidium iodide as a marker of damaged membrane and acridine orange to stain all cells. Figure 6A displays the fluorescence images of MRSA after 24-h cultivation with AP-AgNPs at different concentrations. In the absence of AP-AgNPs, all cells had a green fluorescence representing an undamaged cytoplasmic membrane. In the presence of AP-AgNPs at ½ and 1× MIC red fluorescence was observed in approximately 30 and 60% of bacteria, respectively (Figures 6A,B). At 2× MIC, AP-AgNPs caused the loss of the membrane integrity by 92% of cells, respectively (Figures 6A,B).

Furthermore, PI staining using flow cytometry estimated cell membrane integrity. Generally, PI has no access to stain cellular DNA with an integral membrane. Fluorescence intensity signifies the number of impaired cells (Han et al., 2020). As shown in Figure 6C, MRSA treated with AP-AgNPs displayed a high percentage of stained cells by PI as compared to either control or vancomycin-treated cells. As shown in Figure 6D, after 30 min of treatment, the optical density values in the AP-AgNP group and the Triton X-100 group were higher as compared to the negative control group, demonstrating that AP-AgNPs increased the membrane permeability of MRSA. The results of intracellular protein and nucleic acids leakage are displayed in Figures 6E,F. AP-AgNPs exhibited a significant effect on MRSA cellular damage as revealed by releasing a high amount of protein and nucleic acids contents of MRSA cells as compared to control non-treated cells.

We examined the effect of AP-AgNPs on the suppression of MRSA biofilm formation by crystal violet staining to measure the mass of biofilm. As shown in Figure 7A, AP-AgNPs displayed a dose-dependent inhibitory effect on the biofilm medium, the formation of MRSA biofilm. Resazurin viability test was applied to measure the occurrence of active biofilm bacteria (Figure 7B). AP-AgNPs excreted an inhibitory effect on the metabolic activity of MRSA in a dose-dependent manner (Figure 7B). Furthermore, we explored the effects of AP-AgNPs on the adhesion of the MRSA biofilm on the glass surface using SEM. As shown in Figures 7C,a,c, control non-treated MRSA showed a significant bacterial biofilm growth, which led to clusters with complex morphology observed on the surface of the glass. In contrast, AP-AgNPs significantly inhibited the bacterial biofilm formation (Figures 7C,b,d). Furthermore, the effect of AP-AgNPs on established biofilms was investigated using confocal laser scanning microscopy (CLSM) (Figure 7D). After treatment for 48 h, the control group showed living bacterial cells (Figures 7D,a) while, treatment with 25 μg/mL (MIC) of AP-AgNPs reduced the bacterial population and decreased the number of bacteria in the biofilm (Figures 7D,b), and removed the biofilms. Furthermore, biofilm bacteria are killed by AP-AgNPs at concentrations of 50 μg/mL and 100 μg/mL, and these concentrations of AP-AgNPs were also able to separate biofilms (Figures 7D,c,d).

As a prerequisite step before applying AP-AgNPs in animals, the cytotoxicity of AP-AgNPs against mammalian cells was examined by a hemolysis test. The results confirmed that AP-AgNPs had lower hemolytic activity at a high concentration, and the HC₅₀ value of AP-AgNPs was 340 μg/mL. At 200 μg/mL, the hemolytic activity of AP-AgNPs in human erythrocytes was lower than 30% (Figure 8A). At this concentration, AP-AgNPs efficiently suppressed the growth of MRSA cells.

To examine the potential wound healing potential of AP-AgNPs against MRSA-induced skin infection, we used a tape-stripping mouse model. In this model, infected mice with MRSA were treated topically with either saline, vancomycin, or AP-AgNPs. Topical treatment of MRSA-induced skin infection in mice with AP-AgNPs reduced the size of wounds significantly at days 2 and 8 as compared to control infected non-treated mice (Figures 8B,C). Precisely, on day 6, mice in the AP-AgNP and vancomycin groups showed fast control of the infection, as demonstrated by the absence of pus under the scab and greater wound healing; however, in the control group, pus persisted under the scab (Figures 8B,C). Conversely, scabs still strongly adhere to the non-treated tissues and approximately 30% of unhealed wounds continued noticeable.

To examine the in vivo anti-MRSA activity of AP-AgNPs, the bacterial burden in skin lesions was evaluated during the timeline of wound healing as described in the material and methods. Interestingly, treatment of MRSA-infected wounds with AP-AgNPs showed a significant reduction in the percentage of MRSA colonies that reached 0% after 6 days of treatment compared to vancomycin treatment mice (Figure 8D).

As shown in Figure 9A, compared to normal skin, abundant inflammatory cell infiltration (mostly neutrophil), great quantities of fragments of pus cells, necrotic tissues, and MRSA clusters were detected in the skin tissues after 2 days of infection by MRSA. On day 8, the healing effect of AP-AgNPs was equivalent to that of vancomycin, with totally healed wound tissues. No MRSA clusters or pus cell fragments were detected, and inflammatory cells were fewer disseminated or not detected (Figure 9B). Furthermore, a greater number of fibroblasts, substantial neovascularization, and even newborn hair follicle tissues were detected in the subcutaneous tissue just underneath the wound surface of AP-AgNP-treated groups, demonstrating good healing (Figure 9B). After 8 days, immunohistochemical (IHC) examination was performed to measure the expression of interleukin (IL-6), which is a significant inflammatory marker, in the hypodermis. The tissue areas displayed a substantial decrease in the infection-related inflammation of the wounds after treatment with AP-AgNPs on day 8 (Figure 9C). More thicker new shaped collagen fibers at the center of the wound area, were detected through Masson straining (Figure 9D) and serum C-reactive protein (CRP) levels were expressively decreased in the AP-AgNP group (Figure 9E). Our results proposed that AP-AgNPs could penetrate through the skin into the infected areas to eliminate MRSA.