The clinical drug-resistant strains (P. aeruginosa 01~10) were obtained from a tertiary care hospital located in Kaili City, Guizhou Province, with all isolates originating from hospitalized patients. The P. aeruginosa CMCC 10104 was obtained from the Key and Characteristic Laboratory of Modern Pathogen Biology at Guizhou Medical University. All strains were preserved in 15% glycerol at -80°C and cultured in LB medium at 37°C with shaking at 220 rpm until reaching the logarithmic growth phase.

The preparation of AMP-17 was carried out according to the method described by (Yang et al., 2022). The protein was purified using a Ni-NTA column, and the concentration of AMP-17 was determined using a BCA protein assay kit.

Referring to the judgment criteria in the American Committee for Clinical Laboratory Standardization (CLSI), the subculturing of bacterial broth in MHB medium was repeated twice. One monoclonal colony was selected, and inoculated into 5 mL of LB liquid medium until the logarithmic growth phase. The bacterial suspension was adjusted to a McFarland turbidity standard using a densitometer, followed by conversion to colony-forming units (CFU). The bacterial concentration was then standardized to 1×10⁶ CFU/mL using 1× PBS for subsequent experiments. The concentration of AMP-17 and 1.0×10⁶ CFU/mL bacterial suspension were incubated in 96-well plates at 37°C for 16 h. The MIC value was determined as the concentration of the drug in the wells with no visible bacterial growth. The concentration of the antimicrobial peptide corresponding to no bacterial growth on the LB plate was used as the MBC. 100 μL of bacterial suspension was diluted to the appropriate concentration from the wells judged as MIC and the wells higher than the concentration of the drug in the wells, and then evenly coated on LB plates and incubated at 37°C for 24 h. MHB medium was used for the blank control, MHB medium with bacterial suspension for the negative control, and 4 μg/mL of polymyxin B for the positive control.

The growth curve analysis of bacteria was based on previous methods with modifications (Boulaamane et al., 2024). The bacterial suspension (1.0×10⁶ CFU/mL) was added into 96-well plates and incubated with different concentrations (1/2~2 ×MIC) of AMP-17 at 37°C for 24 h. The absorbance values at 600 nm were measured hourly, and the results were recorded and the growth curves were plotted. The peptide antibiotic Polymyxin B (2×MIC) was used as the positive control, MHB as the blank control, and MHB medium with bacterial suspension was used as the negative control.

The time-kill kinetics experiment was performed as described in previous study (Lasko et al., 2022). AMP-17 (1/2~2 ×MIC) was incubated with 1×10⁶ CFU/mL of P. aeruginosa CMCC 10104 and 04 suspension at 37°C for 24 h. When incubated for 0, 1, 2, 4, 6, 8, 10, 12, 24 h, 10 μL of the AMP-17-activated suspension was serially diluted in MHB medium at ten-fold dilution to the appropriate concentration. 5 μL of the dilution was pipetted onto LB plates, and the colonies were counted and the time-kill curve was plotted after the colonies had grown.

To investigate the activity of AMP-17 under salt conditions and in the presence of serum, we performed MIC assays using the microbroth dilution method. P. aeruginosa CMCC10104 and 04 were inoculated in Mueller-Hinton broth (MHB) containing gradient concentrations of AMP-17. The culture media were supplemented with either 150 mM NaCl, 4.5 mM KCl, 2.5 mM CaCl₂, 10% murine serum, or fetal bovine serum (FBS). The inoculated 96-well plates were incubated at 37°C for 16 h, followed by MIC determination to evaluate potential alterations in the antibacterial activity of AMP-17.

The motility assay refers to the theory of (Fang et al., 2022). P. aeruginosa CMCC 10104 and 04 in logarithmic growth phase was prepared into 1.0 × 10⁶ CFU/mL bacterial suspension and set aside. When the medium temperature was lowered to 50°C, AMP-17 at final concentrations of 1/4 ×MIC and 1/2 ×MIC was added to the medium, and 2 μL of the suspension was pipetted onto 0.3% (w/v) agar medium containing tryptic peptone (10 g/L), NaCl (10 g/L) and yeast extract (5 g/L). The plates were allowed to dry naturally and then incubated orthotropically at 37°C for 24 h. The bacterial motility area was measured and the effect of AMP-17 on the motility of P. aeruginosa was assessed.

Biofilm formation is a prevalent phenomenon in wound environments and constitutes a significant barrier to the efficacy of antimicrobial agents. In order to evaluate the potential of AMP-17 against biofilms formation in vitro, an antibiofilm assay was performed as follows (Rajan et al., 2023). A bacterial suspension of P. aeruginosa CMCC10104 and 04 (1.0×10⁶ CFU/mL) was prepared in Tryptic Soy Broth (TSB) and co-cultured with serial dilutions of AMP-17 (1/2~2 ×MIC) in 96-well microtiter plates at 37°C for 24 hours. Sterile TSB served as a blank control, while MHB with bacteria acted as a negative control. After incubation, non-adherent cells were removed by aspirating the supernatant and washing twice with sterile phosphate-buffered saline (PBS). The remaining biofilms were methanol-fixed (15 min), air-dried, and stained with 0.1% (w/v) crystal violet for 30 min at room temperature. Excess stain was washed off with PBS, and biofilm-associated dye was solubilized in anhydrous ethanol (200 µL/well) at 37°C for 30 min. The absorbance at 595 nm was measured using a microplate reader to quantify biofilm biomass.

A bacterial suspension of P. aeruginosa CMCC 10104 and 04 at a concentration of 1.0×10⁶ CFU/mL was prepared and incubated at 37°C for 24 h. Following the removal of nonadherent bacteria by washing with sterile phosphate-buffered saline, 200 μL of AMP-17 at varying concentrations were added to the wells and incubated at 37°C for 2 hours. Subsequently, the biofilm biomass was quantified by measuring the absorbance at 595 nm after staining with crystalline violet. TSB served as the blank control, whereas MHB containing the bacterial suspension functioned as the negative control.

To visualize biofilm eradication more comprehensively, the experiment refers to the theory of (Tchatchiashvili et al., 2024). The biofilms were dual-stained with 10 μM propidium iodide (PI) and 5 μM SYTO 9. PI selectively labels dead or membrane-compromised bacterial cells, while SYTO 9 stains both live and dead cells, enabling the differentiation of cell viability within the biofilm. The stained biofilms were then subjected to imaging using a CLSM with excitation wavelengths of 488 nm and 561 nm for SYTO 9 and PI, respectively.

The quantification of pyocyanin was performed by the method of (Gasu et al., 2019). A bacterial suspension of P. aeruginosa CMCC 10104 was prepared at a concentration of 1.0×10⁶ CFU/mL. AMP-17 was then added to achieve final concentrations of 1/2 to 2 ×MIC, and the mixture was incubated at 37°C for 24 hours. Following incubation, the culture was centrifuged at 5000 rpm for 10 minutes to obtain the supernatant. Prior to pyocyanin measurement, bacterial concentrations in all samples were adjusted to 1 × 10⁶ CFU/mL through turbidimetric assessment and plate counting to ensure standardized comparisons. 5 mL of chloroform was added to the supernatant, and after thorough vortexing, the mixture was allowed to stand for 8 minutes. The sample was then centrifuged at 4500 rpm for 10 minutes, and the lower, blue-colored chloroform phase was collected. Next, 1 mL of 0.2 mol/L hydrochloric acid was added to the collected phase, and the mixture was allowed to stand for 30 minutes with intermittent vortexing until extraction was complete. After a final centrifugation at 4500 rpm for 8 minutes, the resulting pink supernatant was collected. The pyocyanin concentration (μg/mL) was calculated using the following formula, where 17.072 represents the molar extinction coefficient of pyocyanin at 520 nm:

Bacterial suspension of P. aeruginosa CMCC 10104 and 04 were prepared at 1.0×10⁶ CFU/mL and treated with AMP-17 at a final concentration of 2 ×MIC. The sample was incubated at 37°C for 8 hours, then centrifuged at 3000 g for 10 minutes to pellet the cells. The pellet was washed once with sterile PBS, discarding the supernatant thereafter. The cells were resuspended in 2.5% glutaraldehyde and fixed overnight at 4°C in the dark. Following fixation, the cells underwent sequential dehydration in graded ethanol solutions (30%, 50%, 70%, 80%, 90%, 95%, and 100%). The dehydrated samples were dried under reduced pressure and subsequently sputter-coated with gold using an ion sputterer. SEM was then employed to observe the cellular morphology, and representative micrographs were collected. A PBS-treated group served as the negative control.

For TEM observation, the samples were prepared in a manner analogous to that used for scanning electron microscopy. Briefly, the samples were centrifuged at 3000 g for 10 minutes, and the supernatant was discarded. The pellet was then fixed in 2.5% glutaraldehyde at 4°C overnight in the dark. Following this primary fixation, the samples were post-fixed in 1% ephorbic acid at 4°C for 2 hours. Subsequently, the specimens underwent graded dehydration in ethanol for 20 minutes per concentration step, followed by two washes in 100% acetone, each lasting 15 minutes. The dehydrated samples were then embedded in Epon 812 by immersion. Ultrathin sections were prepared, double-stained with uranyl acetate and lead nitrate, and examined using TEM, with micrographs collected.

To assess membrane fluidity (Verstraeten et al., 2021), the bacterial suspension of P. aeruginosa CMCC 10104 and 04 were adjusted to a concentration of 1.0×10⁶ CFU/mL. Laurdan dye was introduced to achieve a final concentration of 10 μM, and the mixture was incubated at 37°C for 1 hour. Following incubation, the cells were washed once with sterile PBS. Subsequently, 100 μL aliquots of the bacterial suspension were exposed to AMP-17 at concentrations of 1/2 to 2 ×MIC for 1 hour. Fluorescence intensity was measured using a multifunctional microplate reader, with an excitation wavelength of 350 nm and emission wavelengths of 435 nm and 490 nm. The Laurdan Generalized Polarization was calculated using the following equation:

The transmembrane potential refers to the theory of (Song et al., 2021). Various concentrations of AMP-17 were introduced to the 1.0×10⁶ CFU/mL bacterial suspension, followed by the addition of DiSC₃ (5) dye to a final concentration of 0.5 μM. The mixture was incubated in the dark at 37°C for 15 minutes. Subsequently, 100 μL of the treated suspension was transferred to a black 96-well microplate. Fluorescence intensity was measured using a multifunctional fluorometer, with excitation and emission wavelengths set at 622 nm and 670 nm, respectively. Fluorescence changes were continuously monitored and recorded over a 1-hour period.

To detect changes in ΔpH in P. aeruginosa, the 1.0×10⁶ CFU/mL bacterial cells were then incubated with 1 μM of the pH-sensitive fluorescent probe BCECF-AM at 37°C for 30 minutes under low-light conditions (Kim, 2024). Following this incubation, the cells were treated with varying concentrations of AMP-17 at 37°C for 1 hour, again under dark conditions. Fluorescence intensity was measured using a multifunctional fluorometric plate reader, with excitation and emission wavelengths set at 488 nm and 535 nm, respectively.

P. aeruginosa CMCC 10104 was cultured to a concentration of 1.0×10⁶ CFU/mL and incubated with AMP-17 at final concentrations of 1/2 to 2 ×MIC at 37°C for 8 hours. A control group was treated with an equal volume of sterile 1×PBS under the same conditions. Following incubation, 5 μM SYTO 9 and 10 μM PI were added to each sample, and the mixtures were incubated at 37°C for 15 minutes in the dark. Subsequently, bacterial suspension was placed on microscope slides. Fluorescence images were captured using a CLSM.

To assess outer membrane permeability, we referred to the method of (Yang et al., 2023). The 1.0×10⁶ CFU/mL bacterial suspension was incubated with AMP-17 (at final concentrations of 1/2~2 ×MIC) at 37°C in a 96-well black microplate. Following this, N-phenyl-1-naphthylamine (NPN) dye was added to each well to a final concentration of 10 μM, and the mixtures were incubated in the dark. Fluorescence intensity was measured using a multifunctional microplate reader, with excitation and emission wavelengths set at 350 nm and 420 nm, respectively.

The inner membrane permeability experiment was performed as described in previous study (Mikkola et al., 2019). The 1.0×10⁶ CFU/mL bacterial suspension was incubated with PI at a final concentration of 10 μM for 15 minutes in the dark. Subsequently, AMP-17 was added to achieve final concentrations of 1/2 to 2 ×MIC, and the mixture was incubated at 37°C for 1 hour in 96-well black microplates. Fluorescence intensity was measured using a multifunctional microplate reader, with excitation and emission wavelengths set at 622 nm and 670 nm, respectively.

To evaluate ROS production, the experiment was performed as described in previous study (Baker et al., 2021). The 1.0×10⁶ CFU/mL bacterial suspension was incubated with AMP-17 (the final concentrations of 1/2~2 ×MIC). Subsequently, 2’,7’-dichlorodihydrofluorescein diacetate (DCFH-DA) was added to a final concentration of 10 μM, and fluorescence intensity was continuously monitored at 37°C for 1 hour. Real-time fluorescence measurements were conducted using a multifunctional microplate reader, with excitation and emission wavelengths set at 485 nm and 528 nm, respectively.

To establish a mouse wound model, female Balb/c mice (6 weeks old, 18 ~ 20 g) were purchased from SPF Biotechnology Co., Ltd (Beijing, China). The animals were housed under a standard 12-hour light/12-hour dark cycle. All animal experiments were conducted in accordance with the Ethical Principles in Animal Research adopted by Guizhou Medical University and were approved by the Institutional Animal Care and Use Committee (The animal ethics review form number: No. 2302013).

After anesthetizing the mice, full-thickness oval wounds with a long axis of 10 mm were created on their backs using a puncher. Subsequently, 100 μL of P. aeruginosa solution (1.0 × 10⁸ CFU/mL) was applied to induce infection. The wound infection model was established by maintaining continuous infection for 2 days. The mice were then randomly divided into three groups (n = 6): control, vehicle and AMP-17. Wounds not infected with P. aeruginosa and treated with 100 μL volume of normal saline were designated as the control group, while wounds infected with P. aeruginosa and treated with the same volume of normal saline were designated as the vehicle group. AMP-17 was administered at a concentration of 2 ×MIC. A 100 μL volume of the respective sample was applied dropwise to the wound surface using a 1 mL syringe. Infected wounds received daily treatments for 7 days (100 μL once a day), with daily monitoring of wound changes. After 11 days, the mice were sacrificed, and the healing effects of each group were evaluated through hematoxylin-eosin (H&E) staining and Masson staining.

Wound tissue was collected in a sterile environment, weighed, and homogenized in 1 mL of normal saline. Serial dilutions were prepared, and the homogenate was inoculated onto LB solid culture plates, which were incubated for 24 hours at 37 °C. The bacterial load on the wound surface was calculated using the formula:

Additionally, the wound tissue was homogenized in RIPA buffer for 5 minutes at 4°C. The levels of vascular endothelial growth factor (VEGF), interleukin-6 (IL-6), and tumor necrosis factor-alpha (TNF-α) in the supernatant were determined using ELISA kits.

The experiment was performed with three biological replicates per group. All data were processed by Graph Pad Prism 8.0.1 and expressed as mean ± standard deviation. T-test was used to analyze the differences between the two groups, and P< 0.05 was considered significant. The wound area was measured and calculated using ImageJ software.

Ten multidrug-resistant P. aeruginosa strains were collected from clinical samples and exhibited varying degrees of resistance to five commonly used antibiotics: cephalosporins, carbapenems, aminoglycosides, quinolones, and peptides ( Table 1 ). AMP-17 showed 1 ×MIC values of 25 µg/mL and MBC of 50 µg/mL against both the standard strain P. aeruginosa CMCC 10104 and the clinical isolates. Subsequent experiments were conducted using P. aeruginosa 04 as the representative clinical strain, selected based on its extensive multidrug resistance profile among the tested isolates.

The growth curve of P. aeruginosa CMCC 10104 and 04 without AMP-17 treatment was “S” shaped, and the bacterial growth reached the stationary phase after 16 h. Treatment with 1 ×MIC of AMP-17 inhibited bacterial growth within 10 hours, with only marginal OD₆₀₀ increase observed by 24 hours. In contrast, 2×MIC treatment completely suppressed growth throughout the 24-hour incubation period. ( Figure 1A ). Time-kill kinetics experiments confirmed that AMP-17 exhibits bactericidal effects in a concentration- and time-dependent manner. At a concentration corresponding to 2 ×MIC, complete eradication of the bacterial population was achieved within 8 hours, and this inhibitory effect persisted for an additional 16 hours without any evidence of regrowth ( Figure 1B ). Notably, at the 6-hour time point, AMP-17 (2 ×MIC) demonstrated significantly enhanced bactericidal activity against the drug-resistant P. aeruginosa strain 04 compared to the reference strain CMCC10104, with the bacterial load being reduced by 3.091 log₁₀ CFU/mL.

Notably, systematic evaluation of the salt and serum stability of AMP-17 revealed the following findings ( Table 2 ). The MIC of AMP-17 demonstrated remarkable stability, showing no significant alterations in the presence of physiological concentrations (150 mM NaCl, 4.5 mM KCl, or 2.5 mM CaCl₂) compared to the control group. Importantly, under simulated physiological conditions using culture media supplemented with 10% FBS or murine serum, AMP-17 maintained antimicrobial efficacy, with no change in MIC values.

To investigate the effect of AMP-17 on P. aeruginosa motility, we utilized semisolid agar plates ( Figure 2A ). Bacterial migration was significantly reduced in the presence of AMP-17 after 24 hours of incubation. Notably, compared to the untreated control, exposure to AMP-17 at 1/4 ×MIC concentration induced reductions in swarming motility areas of P. aeruginosa CMCC10104 and strain 04 by 47.63% and 48.91%, respectively. Upon escalating the AMP-17 concentration to 1/2 ×MIC, both strains exhibited remarkably diminished motility zones with more pronounced inhibition rates of 88.04% and 90.26% ( Figure 2B ). These findings collectively suggest that AMP-17 may exert its antimicrobial effects through suppression of bacterial motility mechanisms. These findings suggest that AMP-17 impairs bacterial motility, thereby enhancing its bactericidal activity.

Pyocyanin, a virulence factor produced by P. aeruginosa, plays a significant role in disrupting the host defense system and contributes to the pathogenesis of infections. As shown in Figure 2C , increasing concentrations of AMP-17 led to a significant reduction in pyocyanin production across all groups compared to the untreated strains. These findings suggest that AMP-17 effectively inhibits pyocyanin production, thereby potentially reducing the virulence of P. aeruginosa and aiding in infection control.

Biofilms serve as protective barriers for bacteria, shielding them from antibiotic-induced eradication and contributing to the development of bacterial resistance. As illustrated in Figure 2D , the formation of P. aeruginosa biofilm was effectively inhibited by 1/2 ×MIC of AMP-17. Significantly, no statistically discernible eradication of mature biofilms was observed compared to the untreated control at 1/2×MIC ( Figure 2E ). However, AMP-17 at concentrations (1 and 2 ×MIC) elicited significant structural disintegration of biofilms.

To assess the bactericidal activity against biofilm-resident viable cells, we conducted SYTO9/PI dual fluorescence staining with confocal laser scanning microscopy and evaluated the proportion of live to dead bacteria within the biofilm ( Figures 2F, G ). Results revealed a dose-dependent intensification of red fluorescence (indicative of membrane-compromised cells) concomitant with diminished green fluorescence (viable cells), demonstrating that AMP-17 not only disrupts biofilm architecture but also induces substantial bacterial mortality within the biofilm.

SEM revealed that untreated P. aeruginosa cells exhibited intact morphology with smooth, undamaged surfaces ( Figure 3A ). Following 24-hour treatment with 2×MIC AMP-17, both P. aeruginosa CMCC 10104 and 04 strains exhibited distinct ultrastructural alterations. SEM analysis revealed localized membrane rupture accompanied by cytoplasmic efflux in CMCC 10104 cells, whereas 04 cells displayed characteristic depression-like pores distributed across the bacterial surface. These morphological modifications suggest that AMP-17 may compromise cellular integrity through disruption of membrane architecture in P. aeruginosa.

TEM analysis further predicted the membrane-targeting effects of AMP-17. Untreated controls maintained characteristic membrane architecture with preserved cytoplasmic density and organelle organization ( Figure 3B ). In contrast, AMP-17-treated cells (2 ×MIC) exhibited discontinuous membrane continuity, accompanied by periplasmic space dilation. Notably, cytoplasmic content redistribution and nucleoid material dispersion were observed in treated specimens. These ultrastructural perturbations collectively indicate simultaneous disruption of membrane architecture and intracellular organization.

Fluorescence analysis using the NPN probe revealed modulatory effects of AMP-17 on the outer membrane permeability of P. aeruginosa. The control group exhibited minimal fluorescence signal fluctuations during the initial 5-minute monitoring period ( Figure 4A ), whereas AMP-17-treated samples demonstrated a progressive increase in fluorescence intensity, suggesting outer membrane permeability alterations. Parallel experiments with PI staining showed time-dependent enhancement of fluorescence signals ( Figure 4B ), correlating with modifications in inner membrane permeability. Live/dead cell viability assays ( Figure 4C ) revealed concentration-dependent responses to AMP-17 treatment. Bacterial samples treated with 1/2×MIC AMP-17 showed limited red fluorescence emission, whereas 1×MIC exposure elevated red fluorescence intensity relative to untreated controls, suggesting compromised membrane integrity. At 2×MIC concentration, the entire bacterial population exhibited complete red fluorescence conversion. These findings imply that AMP-17’s antibacterial mechanism likely involves coordinated modulation of both outer and inner membrane permeability.

Laurdan GP analysis revealed concentration-dependent elevation of generalized polarization values in AMP-17-treated groups (1 ×MIC and 2 ×MIC) relative to untreated controls, implying potential compound-mediated modulations in membrane fluidity ( Figure 5A ). This phenomenon appears consistent with previously documented alterations in membrane permeability.

Given the well-established correlation between membrane permeability changes and PMF dynamics, a critical bioenergetic parameter that integrates membrane potential (Δψ) and transmembrane proton gradient (ΔpH) to support bacterial survival, we systematically investigated these components. Membrane potential perturbations were evaluated using a DiSC₃(5) fluorescent probe. Following co-incubation with P. aeruginosa, fluorescence baselines remained stable throughout the 5-minute equilibration period. Subsequent administration of AMP-17 at varying concentrations induced dose-responsive fluorescence intensity increases ( Figure 5B ), indicative of possible membrane depolarization events. For transmembrane proton gradient assessment, the BCECF-AM probe was implemented. This non-fluorescent compound undergoes intracellular esterase-mediated conversion to BCECF, a pH-sensitive fluorophore whose emission intensity demonstrated a positive correlation with AMP-17 concentrations ( Figure 5C ), suggesting potential disruption of proton gradient homeostasis.

In this study, intracellular ROS accumulation in P. aeruginosa cells was quantitatively assessed using the non-fluorescent probe DCFH-DA. Upon oxidation by ROS, DCFH-DA is converted to the highly fluorescent compound DCF, the fluorescence intensity of which was measured using a fluorescence microplate reader to determine ROS levels. Treatment with 2 ×MIC of AMP-17 resulted in a higher concentration of ROS compared to the 1 ×MIC group, with ROS levels increasing over time ( Figure 5D ). These findings suggest that AMP-17 induces ROS accumulation in P. aeruginosa cells and induces bacterial death.

To evaluate the in vivo efficacy of AMP-17, a murine wound infection model was established. One hour after creating full-thickness skin wounds on the dorsum of BALB/c mice, the wounds were topically inoculated with 1×10⁸ CFU/mL of P. aeruginosa 04 ( Figure 6A ). At 48 hours post-wounding, the AMP-17 treatment group received a topical application of 50 μg/mL AMP-17, while the vehicle group was treated with an equal volume of PBS as a control. Wound tissues were harvested on days 7, 9, and 11 for analysis. In the absence of P. aeruginosa infection, the wound healing rate in the control group reached 74.18% by day 11, whereas the vehicle and AMP-17 group exhibited healing rates of 39.25% and 86.35%, respectively ( Figures 6B, C ). Bacterial load analysis revealed that AMP-17 treatment significantly reduced the wound bacterial burden by 1.22 Log₁₀ CUF/g, 2.29 Log₁₀ CUF/g, and 3.73 Log₁₀ CUF/g on days 7, 9, and 11, respectively, compared to the model group ( Figure 6D ). Histopathological examination demonstrated that the AMP-17 group exhibited markedly reduced inflammatory cell infiltration and fibroblast proliferation, along with enhanced collagen deposition, compared to both the control and model groups ( Figure 6E ), and the expression of VEGF was increased, suggesting that AMP-17 promotes wound healing ( Figure 6F ). Furthermore, while the expression levels of pro-inflammatory cytokines IL-6 and TNF-α were significantly upregulated in the model group, AMP-17 treatment markedly downregulated their expression. Collectively, these results indicate that AMP-17 exhibits potent antimicrobial activity against P. aeruginosa 04 infection in vivo and promotes wound healing by suppressing infection-induced inflammatory responses.