The bacteria strains Escherichia coli ATCC 25922, Escherichia coli UB1005, Salmonella typhimurium ATCC 14028, Salmonella typhimurium C7731, Pseudomonas aeruginosa ATCC 27853, Pseudomonas aeruginosa PAO1, Staphylococcus aureus ATCC 29213, Staphylococcus aureus ATCC 25923, Methicillin-resistant Staphylococcus aureus (MRSA) 43300, Enterococcus faecalis ATCC 29212, and Staphylococcus epidermidis ATCC 12228 were obtained from the College of Veterinary Medicine, Northeast Agricultural University (Harbin, China).

All the amphiphiles designed in this study were synthesized by GL Biochem (Shanghai) Ltd. (Shanghai, China) using solid-phase peptide synthesis.

The secondary structure of the peptides (150 μm) was measured at wavelengths ranging from 195 to 250 nm using a J820 spectropolarimeter (Jasco, Tokyo, Japan). The mean residue ellipticity was converted using the following equation:

Where θ_obs is the observed ellipticity corrected for the buffer, c is the concentration of the peptides, l is the path length, and n is the number of amino acids in the peptides (Crusca et al., 2018).

The bacteria were cultured overnight in the MHB medium and transferred to the new MHB the next day until the bacteria reached the exponential growth stage. The bacteria were diluted with MHB to 2 × 10⁵ CFU/ml. About 50 μl of polypeptide and equal volume of bacterial suspension was added to a 96-well plate to give the final concentration of polypeptide between 1 and 128 μm. The last two columns are negative (MHB only) and positive (bacterial and MHB) controls and incubated at 37°C for 18–24 h. Optical density was measured at an absorbance wavelength of 492 nm using a microplate reader (Tecan GENios F129004, Tecan, Austria). At least three independent measurements were taken. Each trial was conducted two times in parallel. A total of three independent assays were carried out.

The hemolytic activity of the amphiphiles was assessed by determining the hemolytic rate of the peptide to hRBCs. The cytotoxicity of each peptide was assessed with IPEC-J12 and HEK 293T cells. The specific experimental method is the same as the previous study (Xu et al., 2020; Xue et al., 2021). Each trial was conducted two times in parallel. A total of three independent assays were carried out.

The salt and serum stability of polypeptides were determined by improved MIC method (Lv et al., 2014). The salt and serum stability of polypeptides were determined by improved MIC method. Peptides were incubated with BSA containing different final concentrations of salt ions (150 mM NaCl, 4.5 mM KCl, 6 μm NH₄Cl, 8 μm ZnCl₂, 1 mM MgCl₂, 2 mM CaCl₂, and 4 μm FeCl₃) (Wang et al., 2018). To determine the stability of the peptide in serum, peptide was incubated with different proportions of serum (50%, 100%) at 37°C for 4 h. The following steps are the same as those for antimicrobial activity. Each trial was conducted two times in parallel. A total of three independent assays were carried out.

Continuous culture method was used to evaluate the drug resistance of peptides. Colistin was used as model antibiotic to test the resistance of E. coli 25922. First, minimum inhibitory concentration (MIC) was determined for peptide and Colistin Next, the bacteria with each sub-MIC concentration were diluted to 2 × 10⁵ CFU/ml with MHB for the MIC determination of the next passage. The process was repeated 30 times.

The bacteria cultured to logarithmic growth phase were diluted to OD600 nm = 0.4 (3 × 10⁸-4 × 10⁸ CFU/ml) and diluted 1,000 times in phosphate buffer saline (PBS). The peptides with 1, 2, 4, 8, and 16 times the minimum inhibitory concentration (MIC) were incubated with the bacterial solution in a polypropylene test tube with a total volume of 1 ml at different time intervals (3, 5, 15, 30, 60, and 120 min). About 50 μl of the mixed solution was absorbed and diluted to an appropriate multiple (100 or 1,000). About 20 μl was extracted and evenly coated on MHA solid medium plate. After overnight incubation at 37°C, the colony number was calculated and the colony number curve was plotted for each time point. A total of three parallels per trial and four trials were repeated.

The protocol used in this in vivo test was approved by the Animal Care Use Committee of Northeast Agricultural University (NEAU-[2011]-9), and all the animal care and treatment protocols were in line with the standards stipulated in the Regulations on The Management of Experimental Animals of Heilongjiang Province, China (revised in 2016).

A total of 32 female C57BL/6 mice (weighing about 20 g) aged from 6 to 8 weeks were selected. The mice in each group were divided into 4 groups by blind method (n = 8): (a) control group (saline), (b) infection group (E. coli 25922) (c) treatment group (E. coli 25922 + TP(i+4) 1&2&5. (d) treatment group (E. coli 25922 + Colistin). Mice in control group were intraperitoneally injected with 100 μl normal saline, and mice in other groups were intraperitoneally injected with 100 μl (~1 × 10⁸ CFU/ml) diluted bacterial solution. The peptide treatment group was intraperitoneally injected with TP(i+4) 1&2&5 (5 mg/kg) at 0 and 1 h after infection, and the control group and antibiotic treatment group were injected with normal saline and colistin (1 mg/kg), respectively. The mice were then euthanized 12 h after infection. After weighing liver, lung, spleen, and kidney, the bacterial load of each organ was determined, and the histopathological changes in the organ sections were analyzed by H&E staining. Serum levels of inflammatory factors IL-6, IL-1β, and TNF-α were detected by ELISA kits (Shanghai Jinma).

The LPS and LTA binding affinities of the peptides was assessed with a fluorescent dye BC displacement method. In brief, LPS or LTA was mixed with BC dye at final LPS/LTA and BC concentrations of 50 and 5 μg/ml. The dye was incubated in the dark for 4 h, and the polypeptide was mixed with the dye. Fluorescence was measured using a fluorescence spectrophotometer (Hitachi, Japan) at an excitation wavelength = 580 nm and emission wavelength = 620 nm. LPS-binding affinities were calculated using the following equation:

Where F_t is the fluorescence measured after peptide treatment, F_n is the negative control (the initial fluorescence of BC with a lack of the peptides), and F_p is the positive control (the fluorescence measured after polymyxin B (10 μg/ml) treatment). Because of its strong binding affinities, polymyxin B was used as the positive control (Kumar and Shin, 2020).

The OM permeability of polypeptides was evaluated by fluorescent dye NPN. E. coli 25922 in the exponential phase were centrifuged at 5,000 g for 5 min at 4°C to discard the MHB, and the remaining bacterial cells were washed three times with HEPES (5 mM, pH = 7.4, containing 5 mM glucose) and resuspended with HEPES to OD600 nm = 0.2. Next, NPN dye (10 μm) and bacterial suspension were incubated for 30 min at 37°C in the dark. Afterward, peptides of equal volume and different concentrations were mixed. Fluorescence leakage induced by polymyxin B (20 μg/ml) was used as positive control. Fluorescence was measured using a fluorescence spectrophotometer (Hitachi, Japan) at an excitation wavelength = 350 nm and emission wavelength = 420 nm. The formula is as follows:

Where F_t is the fluorescence measured after peptide treatment, F_n is the negative control (the initial fluorescence of NPN with a lack of peptides), and F_p is the positive control (the fluorescence measured after polymyxin B (10 μg/ml) treatment). Owing to its strong OM permeability, polymyxin B was used as the positive control.

Fluorescent dye DiSC3(5) method was used to determine the CM depolarization ability of polypeptides. E. coli 25922 and S. aureus 29213 in the exponential phase were centrifuged at 5,000 g for 5 min at 4°C to discard the MHB, and the remaining bacterial cells were washed three times with HEPES (5 mM, pH = 7.4, containing 20 mM glucose) and resuspended with HEPES to OD600 nm = 0.05. Then, DiSC3(5) dye (0.4 μm) and bacterial suspension were incubated for 1 h at 37°C in the dark. Next, 0.1 M KCl was added and then incubated with the bacteria for 30 min, and 2 ml of the mixture was added to each well of a 24-well plate. Afterward, fluorescence was measured using a fluorescence spectrophotometer (Hitachi, Japan) at an excitation wavelength = 622 nm and emission wavelength = 670 nm until a stable reduction in the fluorescence was achieved. Subsequently, the peptides at different concentrations were added to the 24-well plate, and the fluorescence was measured to 1,500 s.

The damage of peptide to bacterial membrane was evaluated by Becton-Dickinson, USA. The bacteria were cultured to logarithmic stage, washed with PBS, and modulated to OD600 nm = 0.2. The peptides with different concentrations were incubated with bacteria at 37°C for 1 h. Then 1 h later, it was incubated with PI (10 μg/ml) at 4°C for 30 min. Samples without peptide treatment were used as controls.

Fluorescence microscopy was used to further evaluate the ability of peptides to destroy bacterial membranes. The peptides with different concentrations were incubated with bacteria at 37°C for 1 h. Then, SYTO 9 green fluorescent nucleic acid staining and PI red fluorescent nucleic acid staining were performed for 30 min. No peptide-treated bacteria were used as controls. Next, images are obtained using a fluorescence microscope.

The bacteria were cultured to logarithmic stage, washed with PBS, and suspended again to OD600 nm = 0.2. About 4 μm of peptide with bacteria was incubated for 60 min, with no peptide as control. The method refers to the previous method in our laboratory (Lai et al., 2019; Yu et al., 2021).

For statistical analyses, ANOVA was performed using SPSS software, one-way ANOVA was used across multiple groups (one-way ANOVA), and p < 0.05 was considered to indicate an extremely significant difference.

The specific substitution positions and naming are shown in Figure 1, 1–4 are the substitution positions of polar surface positively charged amino acids, and paired Lys substitutions N5 G9, G9 G13, S2 K6, and K6 G10 are named as 1, 2, 3, and 4, respectively. Because it is substituted based on the hydrogen bond formation position (i i+4) on the original peptide TP, so it is uniformly named as TP(i+4). Therefore, the peptides designed by substituting Lys for the N5 G9, G9 G13, S2 K6, and K6 G10 positions of the TP peptide were named as TP(i+4) 1, TP(i+4) 2, TP(i+4) 3, and TP(i+4) 4. Subsequently, considering whether the substitution has a cumulative effect, two and three pairs of different substitution combinations were subsequently designed (TP(i+4) 1&2, TP(i+4) 1&3, TP(i+4) 1&4, TP(i+4) 2&3, TP(i+4) 2&4, TP(i+4) 3&4, TP(i+4) 1&2&3, TP(i+4) 1&2&4, and TP(i+4) 2&3&4). In the second step, the most active TP(i+4) 1&2 was selected as the basis for the second transformation. The amino acids where Trp replaces T8 A12, L7 V11, and K6 G10 are named as TP(i+4) 1&2&5, TP(i+4) 1&2&6, and TP(i+4) 1&2&7. Similarly, their two and three different pairs of combinatorial substitutions were also designed and named as TP(i+4) 1&2&5&6; TP(i+4) 1&2&5&7, TP(i+4) 1&2&6&7, and TP(i+4) 1&2&5&6&7. The key physical and chemical parameters of the designed and synthesized peptides and the projection of the spiral wheel are shown in Table 1 and Supplementary Figure S2.

The synthesized amphiphiles were designed to measure their molecular weights by mass spectrometry (MS) and reversed-phase high-performance liquid chromatography (RP-HPLC). As shown in Table 1, the measured molecular weight is basically consistent with the theoretical molecular weight.

The secondary structure of these peptides in different solutions was determined by circular dichroism (CD), and the results are shown in Figure 1B and Supplementary Figure S1. CD results show that there is a strong negative peak at 208 nm and a weak negative peak at 222 nm. This indicates that these peptides exhibit α-helical structure in membrane simulation solution (Lee et al., 2013). Meanwhile, we also analyze the α-helical content (Supplementary Figure S3) in different solutions. Lys replacing the hydrogen bond formation position improves the α-helical tendency of the peptide compared to TP. Compared with TP(i+4) 1&2, the α-helical tendency of the derivatives obtained from the hydrophobic surface replaced by Trp decreased to a certain extent. Through the determination of secondary structure, we found that the introduction of Trp reduced the helix content in different amplitude. The interaction between amino acid side groups has a great influence on the formation of helix structure (Louis-Jeune et al., 2012).

The MICs of the designed peptides against gram-negative bacteria and gram-positive bacteria are shown in Table 2. In order to investigate the effect of substitution of positive charge at ith and i+4th positions on the antibacterial activity, 14 peptides were designed. Based on the experimental research, we found that compared with the original peptide TP, all of the peptides except TP(i+4) 3 showed inhibitory activity against negative bacteria. The TP(i+4) 1&2 of the first and second pairs replaced at the same time had the best activity. Based on the TP(i+4) 1&2, the effect of Trp replacing the hydrophobic active center and expanding the hydrophobic surface at both ends on the activity of the peptide was investigated. Compared with the TP(i+4) 1&2, other peptides showed compromised antimicrobial activity against both gram-negative and gram-positive bacteria. At the same time, TP(i+4) 1&2&5 was screened out by analysis and comparison.

The amount of charge has a direct effect on the activity of the amphiphiles, but with the development of research, the factors that affect the activity are not limited to the amount of positive charge carried by the peptide, but also depend on the degree of charge aggregation. As shown in Figure 1A, we simulated the structural diagrams of these amphiphiles. The substitution of different sites results in distinct charge aggregation of peptides. We know that from our preliminary activity results, the four peptides that replace a pair of amino acids at the ith and i+4th positions on the polarity plane with Lys are in the order of activity, which was TP(i+4) 2 > TP(i+4) 1 ≈ TP(i+4) 4 > TP(i+4) 3. As expected, TP(i+4) 2-positive charges closely distributed on one side had better activity and TP(i+4)1 and TP(i+4)4 with similar degree of charge aggregation and as we predicted, they had similar antibacterial activity. Compared with the other three peptides, TP(i+4) 3 has a lower charge density. At the same time, we also found through the analysis of its secondary structure TP(i+4) 3 which has a lower helical content than the TP. Many previous reports have suggested that the tendency to form α-helical structures is correlated with the antibacterial activity of α-helical AMPs (Zhao et al., 2013). According to the helical content data in Supplementary Figure S3, the contents of TP(i+4) 3 in TFE and SDS are 54.85 and 61.94%. Thus, we conclude that secondary structure is another important reason for its low activity. Based on the investigation of the initial activity of these amphiphiles substituted by Lys, we find that the stack effect of replacing two pairs is better than a pair effect and the three pairs effect. The antibacterial activity did not increase as the replacement logarithm increased. Excessive positive charge residue is easy to produce electrostatic obstruction which is not conducive to the activity of the peptide. Among them, the relationship between charge and biological activity is not linear and seems to show a threshold. The results showed that the antimicrobial activity of the peptides did not increase when the positive charge was above the threshold. The peptide in this study had the best antibacterial activity at a net charge of +5. This is consistent with the previous research (Lyu et al., 2019). TP(i+4) 1&2 with the best activity was selected to explore the effects of Trp replacement at the H-H positions on the hydrophobic center and the sides of the hydrophobic surface on the peptide activity and cell selectivity. We found that the design and synthesis of TP(i+4) 1&2&6 by replacing the position of the hydrophobic center did not increase the activity as expected, but decreased the activity compared with TP(i+4) 1&2. We discussed this result and concluded that Trp as a common amino acid in natural AMPs, which carries a large indole side chain, so the interaction between the side chains is more obvious (Lopes et al., 2013; Feng et al., 2020). As shown in Figure 1A, the Trp residues in the hydrophobic center were in contact with each other and wound together. Due to the interaction of side chains, the ability of the tryptophan side chain to insert into the bacterial plasma membrane was weakened, so the hydrophobic surface of TP(i+4) 1&2&6 was reduced as a whole, affecting its activity. In the experimental process, we also found that the H-H position of Trp at both ends of the hydrophobic surface significantly improved the activity of the amphiphiles, and the TP(i+4) 1&2&5 showed best compromised antimicrobial activity against both gram-negative and gram-positive bacteria. However, TP(i+4) 1&2&5&7 was significantly lower than TP(i+4) 1&2&6&7 and TP(i+4) 1&2&5&6 by analyzing the activity of multiple pairs of substituted peptides. By analyzing its hydrophobicity, we found that the value of TP(i+4) 1&2&5&7 was 0.879, much higher than that of TP(i+4) 1&2&6&7 and TP(i+4) 1&2&5&6. We infer that the relationship between hydrophobicity and bioactivity is not linear. When the hydrophobicity exceeded the threshold, the antibacterial activity of the antimicrobial peptides decreased rather than increase.

The increasing number of experiments has prompted us to continue to investigate the obstacles to the practical application of these amphiphiles. In addition to their antimicrobial activity, the toxicity of these peptides in practical use is another important consideration (Zhu et al., 2020). To understand the effect of different substitution sites on the cytotoxicity of the peptides, we measured the cytotoxicity and hemolysis activity of these designed peptides. It was observed that Lys substituted peptides had weak cytotoxic effects on HEK 293T and IPEC-J2 cells (Figure 2B and Supplementary Figure S4). Similarly, these peptides rarely cause hemolysis even at maximum concentrations of 64 μm (Figure 2A). Through the significance analysis of these peptides, it was found that there was no significant difference between these 14 peptides. However, toxicity measurements of the amphiphiles based on the substitution of Trp on the hydrophobic surface showed that the introduction of hydrophobic amino acids enhanced the toxicity of the peptides. Except for TP(i+4) 1&2&6 and TP(i+4) 1&2&5, the other peptides showed high cytotoxicity and hemolysis. TP(i+4) 1&2&6 and TP(i+4) 1&2&5 maintained ~50% cell viability even at 128 μm. TP(i+4) 1&2&5 showed lower hemolysis than other peptides, and the hemolysis rate was <50% below 64 μm.

By comparing the toxicity of polypeptides formed by Lys substitution, it was found that paired Lys substitution at H-H formation sites increased activity without increasing toxicity. Trp is a hydrophobic amino acid with aromatic side chains. It is a special amino acid in peptide design. Previous studies have pointed out that introducing amino acids will also bring toxicity while improving activity. Therefore, at the beginning of the design, we predicted that selecting Trp as hydrophobic amino acid to substitute H-H position would bring certain toxicity to the peptide. Thus, as expected, the introduction of Trp increases the activity and toxicity of these peptides. However, by analyzing the toxicity of the peptides, we unexpectedly found that the substitution of tryptophan for TP(i+4) 1&2&6 in the hydrophobic center did not increase its toxicity. Meanwhile, the virulence of TP(i+4) 1&2&5&6 and TP(i+4) 1&2&6&7 was lower than that of TP(i+4) 1&2&5&7. We infer that the location of hydrophobic centers can effectively reduce peptide toxicity by Trp substitution. For further evaluating the cell selectivity of the peptides, we also calculated the SI which defined as the ratio of MHC to the geometric mean (GM) (Dufourc, 2008). As shown in Table 3, compared with other peptides TP(i+4) 1&2&5 has a good therapeutic index (SI = 64), indicating that TP(i+4) 1&2&5 has a greater application potential in the body.

The complex physiological environment is an important reason to limit the application of AMP in vivo (Shao et al., 2020). According to the above analysis results, TP(i+4) 1&2 and TP(i+4) 1&2&5 were selected for stability study. The stability difference between the selected optimal peptide for positive charge replacement and the optimal peptide for hydrophobic replacement was discussed. Therefore, we used an improved MIC method to evaluate the effects of different salt ion environments on peptide activity. Table 4 shows that different salt ion environments have no or only partial influence on MIC values of TP(i+4) 1&2&5. However, for peptide TP(i+4) 1&2, salt ions have a great influence on its activity. As indicated in Table 4, the stability of TP(i+4) 1&2&5 and TP(i+4) 1&2 in 50 and 100% serum was also determined. TP(i+4) 1&2&5 active serum had less effect compared to TP(i+4) 1&2.

Although the peptides exhibit good antibacterial activity, its antibacterial activity will be reduced or even lost under physiological conditions of salt and serum. Poor stability is one of the main factors that limit the clinical application potential of AMPs. In many studies on natural peptides, it is not difficult to find that cations such as Na⁺ and Mg²⁺ are more likely to affect the activity of AMPs (Yu et al., 2011; Chu et al., 2013). Since the electrostatic force between the negative charge on the outer membrane of bacteria and the cationic AMPs attracts each other, therefore, that any factor that reduces the electrostatic interaction between the two, such as high ionic strength, will affect the antibacterial activity of AMPs (Han et al., 2016). However, in this study, TP(i+4) 1&2&5 was shown to have a strong inhibitory effect on gram-negative and gram-positive bacteria in the presence of salt. The TP(i+4) 1&2 of the positive charge replacement design is greatly affected by salt ions. They have the same positive charge and are close to amphiphilicity. However, TP(i+4) 1&2&5 was formed by replacing ith and i+4th with Trp, expanded the hydrophobic surface, and had higher hydrophobicity. It can be inferred that the strategic addition of hydrophobic residues to peptide sequences enhances the affinity of AMPs to bacterial membranes and thus the ability to overcome salt sensitivity. The serum environment had different degrees of inhibition on TP(i+4) 1&2&5 and TP(i+4) 1&2. This phenomenon can be explained using a balance between the free peptide molecule and the protein binding peptide molecule. As more free molecules combine their microbial targets, balance will be gradually released to more peptide molecules, which facilitates maintaining their antibacterial activity (Deslouches et al., 2005). In addition, the antibacterial activity decreased with the increase of serum concentration. This may be due to high concentrations of albumin binding to antimicrobial peptide molecules, which makes the peptide less active (Svenson et al., 2007).

We evaluated the toxicity and activity of these peptides. Finally, TP, TP(i+4) 1&2, and TP(i+4) 1&2&5 were selected to study the mechanism of action. To make the mechanism description more reasonable and credible, the peptides are implemented in a comprehensive membrane operating system for typical bacteria. The membrane manipulation of E. coli 25922 and S. aureus 29213 from outside to inside was shown in the main body, from qualitative to quantitative, and from molecular components to morphology. The membrane permeation of gram-negative bacteria and gram-positive bacteria was studied with fluorescent probes, and the direct interaction between peptides and bacteria was observed under various microscopes.

The likelihood of antimicrobial peptides causing bacterial resistance is considered low but bacterial resistance is still an important research parameter (Andersson et al., 2016; Marston et al., 2016; Cw et al., 2021). Figure 4A shows the potential of TP(i+4) 1&2, TP(i+4) 1&2&5, and colistin to induce bacterial resistance. Colistin resistance appeared as early as the sixth generation, with a 256-fold increase in MIC values after 30 generations and the antibacterial efficacy of TP(i+4) 1&2&5 and TP(i+4) 1&2 did not change during the trial. The results of the mechanism of action study in the previous step showed that the designed peptides exert their antibacterial action mainly by penetrating and disrupting bacterial cell membranes. Bacteria can only resist the physical disruption of the peptide by altering their membrane structure, and therefore, the possibility of AMPs inducing resistance in bacteria is low. Polymyxin, as a control, showed a 256-fold increase in MIC values after 30 generations of polymyxin at the end of the assay period. It has been shown that resistance to polymyxin is associated with plasmid-mediated delivery of the gene MCR1 between strains and the addition of cationic moieties (L-Ara4N and pEtN) to the LPS. In summary, TP(i+4) 1&2&5 and TP(i+4) 1&2 act in a similar manner to most AMPs, where the positive charge on the surface of AMPs is electrostatically attracted to the negative charge of the bacterial membrane, and when the concentration of AMPs reaches a certain threshold, their hydrophobic ends insert into the bacterial membrane disrupting it and causing it to leak until death. This mode of action greatly reduces the likelihood of drug resistance in microorganisms and gives AMPs an unprecedented advantage in preventing the development of bacterial drug resistance.

Thus, the resistance of E. coil 25922 to TP(i+4) 1&2 and TP(i+4) 1&2&5 and the bactericidal dynamic curve of the target amphiphilic peptide against the infected strain were evaluated before preclinical testing. The time killing curve of E. coli 25922 was determined, and it was found that the target peptide caused the decrease of bacteria within 0–15 min, and then, the bactericidal rate tended to slow down. However, neither peptide at 1 × MIC concentration could provide sufficient power to completely kill the pathogen within 2 h (Figure 4B). With the increase of peptide concentration, TP(i+4) 1&2&5 at 4 × MIC can completely kill E. coli 25922 within 2 h. 8 × MIC and 16 × MIC can kill pathogens within 3 min. TP(i+4) 1&2 do not completely kill E. coli 25,922 even at 16 × MIC concentration. The hydrophobicity is an important factor that affects the antibacterial efficiency. TP(i+4) 1&2 has poor bactericidal effect because TP(i+4) 1&2 and TP have low hydrophobicity.

Using FCM and the nuclear fluorescent dye PI, which can penetrate the damaged membrane, the damaging activity of peptide on the bacterial membrane was evaluated. Meanwhile, fluorescence microscopy was used to observe the ability of the peptide to destroy bacterial membranes (Shi et al., 2006). The degree of membrane rupture (PI fluorescence signal) increased with the increase of peptide concentration (Figure 4C). TP(i+4) 1&2&5 caused more than 80% of E coli 25922 rupture and more than 70% of S aureus 29213 rupture. Compared with TP(i+4) 1&2 and TP, TP(i+4) 1&2&5 has stronger ability of sterilization by membrane breaking. At the same time, fluorescence imaging analysis was performed using the non-membrane permeable nucleic acid dye propidium iodide (PI) and the membrane permeable nucleic acid dye SYTO 9. The fluorescence images showed a similar trend to the FCM results (Figure 4D). These results further confirm that TP(i+4) 1&2 and TP(i+4) 1&2&5 kill bacteria by destroying the integrality of the bacterial membranes.

For a more intuitive observation of the activity mechanism of the peptide, SEM and TEM further examined the effects of TP, TP(i+4) 1&2, and TP(i+4) 1&2&5 on the cell morphology of E. coli 25922 and S. aureus 29213 (Figure 5). The control group was E. coli 25,922 and S. aureus 29,213, which were not treated with peptide. The test results could be observed that their bacterial cell membrane surface was full and smooth with no signs of rupture and the bacterial membrane was in good condition. The TP-treated bacterial cell membranes showed slight wrinkles on the surface, but no obvious pores or fragmentation was observed. The TP(i+4) 1&2- and TP(i+4) 1&2&5-treated E. coli 25922 showed visible folds and large pores and breaks on the surface of the bacterial cell membrane, but TP(i+4) 1&2 was less effective than TP(i+4) 1&2&5. Since there is no activity against TP(i+4) 1&2-positive bacteria, the cell membrane state of S. aureus 29213 treated with TP(i+4) 1&2 in the figure is not significantly different from the control. The effects of TP, TP(i+4) 1&2, and TP(i+4) 1&2&5 on the membrane ultrastructure of E. coli 25922 and S. aureus 29213 were further visualized using TEM. It was clearly observed that the membranes of the control group without peptide treatment were clear and intact, with no structural changes and dense internal cell structure. The outer and plasma membranes of E. coli 25922 and S. aureus 29213 treated with TP(i+4) 1&2&5 were clearly separated, the cell membranes were ruptured, and the cytoplasm was leaking. However, since TP(i+4) 1&2 had no activity against the positive bacteria, there was no significant difference between the treated S. aureus 29213 and the control.

Preliminary in vitro antibacterial activity and toxicity determination and analysis provided the basis for AMPs in vivo. Through the comprehensive study and screening of the activity and toxicity of these synthetic peptides as well as the mechanism of action, the potential application of the ultimate target amphiphiles in vivo was studied. We evaluated the in vivo therapeutic effects of TP(i+4) 1&2&5 in the E. coli 25922-mediated C57BL/6 mouse peritonitis model and selected colistin as a control. TP(i+4) 1&2&5 and colistin treatment significantly reduced bacterial load in multiple organs (Figure 6A). Histopathological examination showed that TP(i+4) 1&2&5 therapy significantly reduced multi-organ inflammatory response in varying degrees compared with the infected group. The infiltration of lung and hepatitis cells decreased, and the bleeding of kidney and spleen decreased. No significant lesions were observed in these organs after the peptide treatment (Figure 6C). In addition, to evaluate the therapeutic effect of the target peptide in vivo, serum levels of interleukin-1β (IL-1β), interleukin-6 (IL-6) and tumor necrosis factor-α (TNF-α) were measured. The test results showed a significant reduction in serum levels of IL-1β, IL-6, and TNF-α compared to the E. coli 25922-infected group. IL-1β is considered to be a pro-inflammatory cytokine that plays an important role in the host's defense response to acute infection (Schmidt and Lenz, 2012; Li et al., 2017). IL-6 is a major mediator of sepsis and can cause septic shock through hemorrhage and severe oedema. TNF-α is secreted by immune cells (activated macrophages, monocytes, and T cells). The overexpression of TNF-α is associated with body damage such as apoptosis and tissue inflammation and also induces the recruitment of other inflammatory cells (Jiang et al., 2016; Gao et al., 2021). As the results shown in Figure 6B, the levels of inflammatory factors were significantly lower in the peptide-treated group than in the infected group. The reduction in these proinflammatory cytokines suggests that TP(i+4) 1&2&5 also has potential anti-inflammatory effects in vivo.