The peptides used in the study were synthesized by GL Biochem (Shanghai) Ltd. with an amidated C-terminus, and the purity of the peptides was more than 95%. P. aeruginosa PAO1, P. aeruginosa ATCC27853, P. aeruginosa ATCC9027, and P. aeruginosa CCTCC93066 used in study were cultured using Luria-Bertani (LB) broth medium at 37°C.

The minimum inhibitory concentration (MIC) of the peptides against P. aeruginosa was measured using a broth microdilution assay recommended by the Clinical and Laboratory Standards Institute guidelines (CLSI, 2019). Briefly, exponential-phase P. aeruginosa cells were diluted in LB medium to 10⁵–10⁶ cfu/mL, and the peptides were dissolved and serially diluted in 0.9% saline. Then, the bacterial dilution (80 μL) and peptide dilution (20 μL) were added into sterile 96-well cell culture plates. The final concentration of each peptide was 3.13 μg/mL, 6.25 μg/mL, 12.5 μg/mL, 25 μg/mL, 50 μg/mL, and 100 μg/mL, respectively. Each concentration was conducted in triplicate. Thereafter, the plates were incubated with continuous shaking at 200 rpm for 18–24 h at 37°C. The lowest peptide concentration with no bacterial growth was determined as the MIC.

Hemolytic activity was used to evaluate the in vitro toxicity of the peptides, using the method described previously (Yuan et al., 2022). Briefly, fresh mouse red blood cells (mRBCs) were washed with and then resuspended in 0.9% saline to 2% (v/v), and the peptides were dissolved and serially diluted in 0.9% saline. Then, the mRBCs suspension (100 μL) and peptide dilution (100 μL) were added into sterile 96-well cell culture plates. The final concentration of each peptide was 25 μg/mL, 50 μg/mL, and 100 μg/mL, respectively. Each concentration was conducted in triplicate, and 1% Triton X-100 and no peptide (0.9% saline) treatment were set as the positive and negative controls, respectively. Thereafter, the plates were incubated with continuous shaking at 200 rpm for 1 h at 37°C. After incubation, the plates were centrifuged at 1,000 g for 10 min, and 100 μL of the supernatant was transferred to a new 96-well plate, and the absorbance was measured at 490 nm. Hemolytic activity was evaluated using the following equation: Hemolysis% = (H_sample − H_negative)/(H_positive − H_negative) × 100%. H: absorbance at 490 nm.

The stability of the peptide was determined using the method described previously (Mirzaei et al., 2022; Yuan et al., 2022). For the thermal stability assay, the peptide was incubated at 60°C for 24 h. Then, the MIC of the peptide against P. aeruginosa PAO1 was measured. For salt stability assay, the exponential phase P. aeruginosa PAO1 cells were washed with and diluted in NaCl-free LB medium to 10⁵–10⁶ cfu/mL with specific amounts of NaCl, CaCl₂, MgCl₂, or KCl, respectively. The peptide was dissolved and serially diluted in sterile water. Then, the bacterial dilution (80 μL) and peptide dilution (20 μL) were added into sterile 96-well cell culture plates. The final concentration of each peptide was 3.13 μg/mL, 6.25 μg/mL, 12.5 μg/mL, 25 μg/mL, 50 μg/mL, and 100 μg/mL, respectively. Each concentration was conducted in triplicate. The final concentration of NaCl, CaCl₂, MgCl₂, and KCl was 150 mM/300 mM, 1 mM/2 mM, 1 mM/2 mM, and 2 mM/4 mM, respectively. Thereafter, the MIC of the peptide against P. aeruginosa PAO1 was measured.

The time-killing kinetics assay of the peptide against P. aeruginosa PAO1 was performed using the method described previously (Li et al., 2020). Briefly, exponential-phase P. aeruginosa PAO1 cells were diluted in LB medium to 10⁵–10⁶ cfu/mL, and the peptide was dissolved and serially diluted in 0.9% saline. Then, the bacterial dilution (400 μL) was incubated with peptide dilution (100 μL) at 37°C with continuous shaking at 200 rpm. The final concentration of the peptide was 12.5 μg/mL, 25 μg/mL, and 50 μg/mL, respectively. No peptide (0.9% saline) treatment served as a negative control. Aliquots were collected at 0 min, 15 min, and 30 min, serially diluted in 0.9% saline, and plated on LB agar plates. The plates were incubated at 37°C for 18–24 h, and the CFU was counted.

The action site of the peptide on P. aeruginosa was determined using the method described previously (Li et al., 2014). Briefly, exponential-phase P. aeruginosa PAO1 cells were diluted in LB medium to 10⁶–10⁷ cfu/mL, and the FITC-labeled peptide was dissolved in 0.9% saline. Then, the bacterial dilution was incubated with the FITC-labeled peptide dilution at a volume ratio of 4:1. The final concentration of the FITC-labeled peptide was 10 μg/mL. After incubation for 20 min at 37°C, the cells were washed with and resuspended in PBS, and then immobilized on a glass slide. The cells were observed using a confocal laser-scanning microscope.

The interaction of the peptide with LPS was evaluated using the method described previously (Cao et al., 2012). Briefly, exponential-phase P. aeruginosa PAO1 cells were diluted in LB medium to 10⁵–10⁶ cfu/mL, and the peptides were dissolved and serially diluted in 0.9% saline containing LPS. Then, the bacterial dilution (80 μL) and peptide dilution (20 μL) were added into sterile 96-well cell culture plates. The final concentration of each peptide was 3.13 μg/mL, 6.25 μg/mL, 12.5 μg/mL, 25 μg/mL, 50 μg/mL, and 100 μg/mL, respectively. Each concentration was conducted in triplicate. The final concentration of LPS was 100 μg/mL, 200 μg/mL, or 500 μg/mL, respectively. Thereafter, the plates were incubated with continuous shaking at 200 rpm for 18–24 h at 37°C. The MIC of the peptide in the presence of LPS was determined.

The influence of the peptide on the membrane integrity of P. aeruginosa cells was evaluated using the method described previously (Li et al., 2016). Briefly, exponential-phase P. aeruginosa PAO1 cells were washed with and diluted in 0.9% saline to 10⁶–10⁷ cfu/mL, and then incubated with SYTOX green (final concentration: 5 μM) in the dark for 10 min at 37°C. The peptide was dissolved and serially diluted in 0.9% saline. Then, the bacterial dilution (50 μL) and peptide dilution (50 μL) were added into a Costar 96-well flat-bottom black plate. The final concentration of the peptide was 6.25 μg/mL, 12.5 μg/mL, and 25 μg/mL, respectively. Each concentration was conducted in triplicate, and no peptide (0.9% saline) treatment was served as a negative control. Thereafter, the fluorescence was measured every 2 min at the excitation and emission wavelengths of 488 and 525 nm, respectively.

The influence of the peptide on the membrane potential of P. aeruginosa cells was evaluated using the method described previously (Lee and Lee, 2014). Briefly, exponential-phase P. aeruginosa PAO1 cells were washed with and diluted in 0.9% saline to 10⁶–10⁷ cfu/mL, and then incubated with DiBAC4(3) (final concentration: 10 μM) in the dark for 10 min at 37°C. The peptide was dissolved and serially diluted in 0.9% saline. Then, the bacterial dilution (50 μL) and peptide dilution (50 μL) were added into a Costar 96-well flat-bottom black plate. The final concentration of the peptide was 6.25 μg/mL, 12.5 μg/mL, and 25 μg/mL, respectively. Each concentration was conducted in triplicate, and no peptide (0.9% saline) treatment was served as a negative control. Thereafter, the fluorescence was measured every 2 min at the excitation and emission wavelengths of 488 and 525 nm, respectively.

The influence of the peptide on ROS generation of P. aeruginosa cells was evaluated using the method described previously (Shi et al., 2021). Briefly, exponential-phase P. aeruginosa PAO1 cells were washed with and diluted in 0.9% saline to 10⁶–10⁷ cfu/mL, and then incubated with DCFH-DA (final concentration: 10 μM) in the dark for 20 min at 37°C. The peptide was dissolved and serially diluted in 0.9% saline. Then, the bacterial dilution (50 μL) and peptide dilution (50 μL) were added into a Costar 96-well flat-bottom black plate. The final concentration of the peptide was 6.25 μg/mL, 12.5 μg/mL, and 25 μg/mL, respectively. Each concentration was conducted in triplicate, and no peptide (0.9% saline) treatment was served as a negative control. Thereafter, the fluorescence was measured every 10 min at the excitation and emission wavelengths of 488 and 525 nm, respectively.

The interaction of the peptide with nucleic acids was evaluated using the method described previously (Li Z. et al., 2022). Briefly, approximately 300 ng of DNA (plasmid pET-28a) or RNA (P. aeruginosa PAO1 RNA) was incubated with varying concentrations of the peptide for 10 min at room temperature. Then, the mixtures were electrophoresed in a 1% agarose gel. The migration of nucleic acids in the gel was visualized using a Bio-Rad Gel Documentation system.

The influence of the peptide on biofilm formation of P. aeruginosa was evaluated using the method described previously (Artini et al., 2022). Briefly, exponential-phase P. aeruginosa PAO1 cells were diluted in LB medium to 10⁵–10⁶ cfu/mL, and 200 μL of the bacterial dilution was added into sterile 96-well cell culture plates. After incubation for 4 h at 37°C, the wells were washed with PBS to remove the non-adherent cells. Then, 200 μL of LB medium containing varying concentrations of the peptide was added to the wells. The final concentration of the peptide was 6.25 μg/mL, 12.5 μg/mL, and 25 μg/mL, respectively. Each concentration was conducted in triplicate, and no peptide (0.9% saline) treatment was served as a negative control. After static cultivation for another 24 h at 37°C, biomasses were quantified using a crystal violet staining assay.

The influence of the peptide on the motility of P. aeruginosa was evaluated using the method described previously (She et al., 2018; Chen et al., 2020). Briefly, exponential-phase P. aeruginosa PAO1 cells were diluted in LB medium to 10⁶–10⁷ cfu/mL. To investigate swimming motility, 0.3% LB agar medium (1 mL) supplemented with varying concentrations of the peptide was poured into sterile 24-well cell culture plates. Then, 2 μL of the bacterial dilution was spotted in the center of the agar plate. After incubation for 18–24 h at 37°C, the swimming zone was observed and pictured with a camera. To investigate swarming motility, 0.5% LB agar medium (1 mL) supplemented with varying concentrations of the peptide was poured into sterile 24-well cell culture plates. Then, 2 μL of the bacterial dilution was spotted in the center of the agar plate. After incubation for 18–24 h at 37°C, the swarming zone was observed and pictured with a camera.

The influence of the peptide on the pyocyanin production of P. aeruginosa was evaluated using the method described previously (Pan et al., 2023). Briefly, exponential-phase P. aeruginosa PAO1 cells were diluted in LB medium to 10⁶–10⁷ cfu/mL, and the peptides were dissolved and serially diluted in 0.9% saline. Then, the bacterial dilution was incubated with the peptide dilution at a volume ratio of 4:1. The final concentration of each peptide was 6.25 μg/mL and 12.5 μg/mL, respectively. Each concentration was conducted in triplicate, and no peptide (0.9% saline) treatment was served as a negative control. After incubation for 16 h at 37°C with continuous shaking at 200 rpm, pyocyanin was extracted using 0.6 mL of chloroform from 1 mL supernatant of the bacterial culture. Then, the organic layer was extracted using 0.2 mL of HCl (0.2 N), and the absorbance of the upper layer was measured at 520 nm.

The influence of the peptide on the elastase of P. aeruginosa was evaluated using the method described previously (Musthafa et al., 2012; Xu et al., 2015). Briefly, exponential-phase P. aeruginosa PAO1 cells were diluted in LB medium to 10⁶–10⁷ cfu/mL, and the peptides were dissolved and serially diluted in 0.9% saline. Then, the bacterial dilution was incubated with the peptide dilution at a volume ratio of 4:1. The final concentration of each peptide was 6.25 μg/mL and 12.5 μg/mL, respectively. Each concentration was conducted in triplicate, and no peptide (0.9% saline) treatment was served as a negative control. After incubation for 6 h at 37°C with continuous shaking at 200 rpm, the cultures were centrifuged at 12,000 r/min for 10 min at 4°C. Then, the supernatant was filtered using a 0.22 μm syringe filter. Thereafter, the filtered supernatant (100 μL) was incubated with ECR solution (900 μL: ECR 20 mg/mL, 0.1 M Tris–HCl, 1 mM CaCl₂, pH 7.2) for 4 h at 37°C with continuous shaking at 200 rpm. After incubation, the reaction mixture was incubated on ice for 10 min, and then centrifuged at 12,000 r/min for 10 min at 4°C. The absorbance of the supernatant was measured at 520 nm.

Male BALB/c mice (20–30 g), obtained from Henan SKBEX Biology Co., Ltd., were maintained under a standard condition of humidity (50 ± 5%), temperature (25 ± 2°C), and dark–light cycles (12 h each) with free access to food and water. All the animal-associated experiments were conducted following the Animal Care and Ethics guidelines with protocols approved by the Animal Care and Use Committee of Henan University of Technology and Science. At the end of the experiments, all the mice were humanely euthanized by intraperitoneal injection of excessive pentobarbital.

A mouse subcutaneous infection model was used to evaluate the in vivo anti-P. aeruginosa activity of the peptide according to the method previously described (Li S. et al., 2022). Briefly, exponential-phase P. aeruginosa PAO1 cells were washed with and resuspended in 0.9% saline to approximately 10⁹ CFU/mL. Each mouse was subcutaneously injected with 50 μL of the bacterial suspension at the back near the tail. Then, the mice were randomly divided into three groups (six mice per group). One hour after infection, 50 μL of 0.9% saline (negative control), peptide solution (500 μg/mL in 0.9% saline), or ciprofloxacin solution (positive control, 500 μg/mL in 0.9% saline) was subcutaneously injected into the infected area of each mouse in the corresponding group. All the treatments were administered once a day continuously for 3 days. On the 4^th day, the mice were humanely euthanized, and the infected area of each mouse was sterilized using 10% povidone/iodine solution and 70% ethyl alcohol after shaving the fur. Thereafter, half of the skin abscesses were excised and homogenized in 0.9% saline. After being serially diluted in 0.9% saline, the homogeneous samples were cultured on LB agar at 37°C for 18–24 h. The number of colony-forming units (CFU) per gram of the skin abscess was subsequently calculated. Another half of the samples were used for hematoxylin and eosin staining, and the infiltration of inflammatory cells was observed.

The data were expressed as mean ± standard error of the mean (SEM) and analyzed using the GraphPad Prism 6 software. Differences between groups were analyzed using one-way ANOVA.

Based on the peptide Helepsin, three peptide derivatives (HTP, HTP1, and HTP2) were designed (Figure 1A). HTP was a truncated peptide from Helepsin; HTP1 and HTP2 were designed based on HTP by amino acid substitution with lysine. As shown in Figure 1B, the MICs of Helepsin against the tested P. aeruginosa strains were all 17.3 μM, while HTP had no inhibitory effects against the tested P. aeruginosa strains at 66.9 μM. The MICs of HTP1 against the tested P. aeruginosa strains were close to that of Helepsin, which were 16–32 μM. HTP2 displayed an improved activity against the tested P. aeruginosa strains compared to that of Helepsin with the MICs all 7.7 μM.

Hemolytic activity was used to evaluate the in vitro toxicity of the peptides. As shown in Figure 1C, Helepsin showed about 91% hemolysis at the concentration of 25 μg/mL (8.6 μM), while HTP showed only about 22% hemolysis at the concentration of 100 μg/mL (66.9 μM), and HTP2 showed only about 18% hemolysis at the concentration of 100 μg/mL (61.7 μM). Although HTP1 only showed about 13% hemolysis at the concentration of 25 μg/mL (16 μM), it showed about 72% hemolysis at the concentration of 100 μg/mL (63.9 μM).

According to the antibacterial activity and hemolytic activity, HTP2 was selected, and its stability was evaluated. As shown in Figure 1D, there are no changes in the MICs for HTP2 against P. aeruginosa after incubation at 60°C for 24 h. Thus, HTP2 had good thermal stability. When treated with HTP2 with different salts at varying concentrations, MICs showed varying degrees of changes. Low concentration monovalent ions (2 mM KCl, 4 mM KCl) had little effect on the antibacterial activity of HTP2, but high concentration monovalent ions (300 mM NaCl) had a greater impact on the antibacterial activity of HTP2. And low concentrations of divalent cations (1 mM/2 mM CaCl₂, 2 mM MgCl₂) have a significant impact on activity. Therefore, the impact of different ions, especially cations, on the activity of HTP2 varies.

Electrostatic interaction mediates the activity of AMPs, and LPS is the main anionic component on the surface of P. aeruginosa cells. Thus, the interaction between HTP2 and LPS was determined. As shown in Figure 1D, the MICs of HTP2 increased in the presence of additional LPS, indicating a decrease in the activity of HTP2 against P. aeruginosa. These results indicated that HTP2 could interact with LPS.

To gain insights into the killing mode of HTP2 against P. aeruginosa, the time-killing kinetics were performed. As shown in Figure 2A, the amount of P. aeruginosa PAO1 cells did not show any changes after being treated with HTP2 at a concentration of 12.5 μg/mL for 30 min. When treated with HTP2 for 15 min at the concentrations of 25 μg/mL and 50 μg/mL, the amount of P. aeruginosa PAO1 cells decreased by about 1-log and about 3-log, respectively. Thus, HTP2 killed P. aeruginosa cells in a concentration-dependent manner.

To determine the action sites of HTP2, P. aeruginosa PAO1 cells were treated with FITC-HTP2. As shown in Figure 2B, a strong fluorescence distribution appeared on the cell surface, while there was a weak fluorescence distribution inside the cell. These results indicated that FITC-HTP2 mainly bound to the surface of P. aeruginosa PAO1 cells, and it could also act inside the cell.

To evaluate the influence of HTP2 on the membrane integrity of P. aeruginosa cells, the membrane permeability assay was performed using the fluorescence dye SYTOX green. As shown in Figure 2C, there is a significant increase in fluorescence intensity within a few minutes after treating P. aeruginosa PAO1 cells with HTP2 at the concentration of 6.25 μg/mL, 12.5 μg/mL, or 25 μg/mL, respectively. These results indicated that the membrane integrity of P. aeruginosa cells was disrupted by HTP2.

To evaluate the influence of HTP2 on the membrane potential of P. aeruginosa cells, the membrane potential assay was performed using the fluorescence dye DiBAC4(3). As shown in Figure 2D, there is a rapid decrease in fluorescence intensity after treating P. aeruginosa PAO1 cells with HTP2 at the concentration of 6.25 μg/mL, 12.5 μg/mL, or 25 μg/mL, respectively. These results indicated that the membrane potential of P. aeruginosa cells was disrupted by HTP2.

To evaluate the influence of HTP2 on the production of ROS in P. aeruginosa cells, the amount of ROS was detected using the fluorescence dye DCFH-DA. As shown in Figure 2E, there is a significant increase in fluorescence intensity after treating P. aeruginosa PAO1 cells with HTP2 at the concentration of 6.25 μg/mL, 12.5 μg/mL, or 25 μg/mL for 10 min or 20 min, respectively. These results indicated that HTP2 induced the production of ROS in P. aeruginosa cells.

To evaluate whether HTP2 could interact with nucleic acids, the migration of nucleic acid in gel in the presence of HTP2 was tested. As shown in Figure 2F, the migration of RNA (Figure 2Fa) and plasmid DNA (Figure 2Fb) is retarded by HTP2. These results indicated that HTP2 could interact with different types of nucleic acids.

Biofilm is an important pathogenic factor for P. aeruginosa, therefore, the influence of HTP2 on biofilm formation of P. aeruginosa was evaluated. As shown in Figure 3A, after treatment with HTP2 at the concentration of 6.25 μg/mL, 12.5 μg/mL, and 25 μg/mL, the biomass of P. aeruginosa biofilms was reduced by about 24.1% (p < 0.05), 41% (p < 0.05), and 71.7% (p < 0.05), respectively. Thus, HTP2 could significantly inhibit the biofilm formation of P. aeruginosa.

Motility ability plays a crucial role in the adhesion, colonization, and invasion processes of bacteria with flagella, therefore, the influence of HTP2 on the motility ability of P. aeruginosa was evaluated. As shown in Figure 3B, after treatment with HTP2 at the concentrations of 6.25 μg/mL and 12.5 μg/mL, the size of the bacterial lawn area was significantly smaller than that of the negative control (non-HTP2) treatment, respectively. Thus, HTP2 could significantly inhibit the motility ability of P. aeruginosa.

Pyocyanin can enhance the pathogenicity of P. aeruginosa through multiple modes, therefore, the influence of HTP2 on pyocyanin production of P. aeruginosa was evaluated. As shown in Figure 3C, after treatment with HTP2 at the concentrations of 6.25 μg/mL and 12.5 μg/mL, the production of pyocyanin decreased by about 9.6% (p < 0.05) and 19.6% (p < 0.05), respectively. Thus, HTP2 could significantly inhibit the pyocyanin production of P. aeruginosa.

Elastase plays an important role in the invasiveness of P. aeruginosa, therefore, the influence of HTP2 on the elastase production/activity of P. aeruginosa was evaluated. As shown in Figure 3D, after treatment with HTP2 at the concentrations of 6.25 μg/mL and 12.5 μg/mL, the elastase production/activity decreased by about 7.7% (p > 0.05) and 18.1% (p < 0.05), respectively. Thus, HTP2 could inhibit the production/activity of the elastase of P. aeruginosa.

A mouse subcutaneous infection model was used to evaluate the activity of HTP2 against P. aeruginosa under physiological conditions. As shown in Figure 4D, the bacterial load of P. aeruginosa from the infection area of HTP2-treated mice and Ciprofloxacin-treated mice was significantly decreased (p < 0.05) compared to that from the negative control group. And there was no significant difference between the HTP2-treated group and the Ciprofloxacin-treated group. Moreover, HTP2 significantly inhibited inflammatory infiltration (Figure 4B) in the infection area compared to that of the negative control group (Figure 4A). Thus, HTP2 had good anti-P. aeruginosa activity under physiological conditions.