Polyaspartamide-based antimicrobials for combatting bacterial infections

Abstract

The increasing challenge of antimicrobial resistance has stimulated research on antibacterial polymers, which offer enhanced therapeutic potential due to their facile synthesis, low immunogenicity, and superior stability compared to natural antimicrobial peptides. Herein, a series of cationic polyaspartamides (PASP_nDA_m) with different side chain lengths (m) and degrees of polymerization (n) were synthesized through ammonolysis reaction of poly(β-benzyl-L-aspartate) (PBLA) to systematically investigate their antibacterial activity and biocompatibility. All PASP_nDA_m exhibited antibacterial activity. Among the PASP_nDA_m, PASP₁₀DA₆ (degree of polymerization of 10, bearing side chains modified with 1,6-hexamethylenediamine) displayed potent activities against bacteria (S. aureus and E. coli, MIC = 7.8 μg mL⁻¹) and had the highest selectivity index of 96. Further mechanistic study revealed that PASP₁₀DA₆ killed bacteria by disrupting the integrity of the bacterial membrane, thereby leading to bacterial death. Based on this unique bactericidal mechanism, PASP₁₀DA₆ showed fast bacterial killing kinetics and was less likely to induce bacterial resistance for up to 15 generations. In a mouse skin bacterial infection model, PASP₁₀DA₆ showed effective in vivo antibacterial activity and negligible toxicity. Hence, this study provides a promising strategy for treating clinical bacterial infections.

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1. Introduction

The discovery of penicillin by Alexander Fleming in 1928 marked the beginning of the antibiotic era, revolutionizing the treatment of previously fatal diseases.^1,2 However, the widespread use of antibiotics has created a significant challenge in the form of antimicrobial resistance (AMR). The emergence and evolution of AMR represent a grave challenge to public health, fueled by the selective pressure exerted by antibiotic exposure.³ Over time, bacteria have evolved diverse mechanisms to resist the effects of antibiotics, including enzymatic degradation, activation of efflux pumps, and genetic mutations that confer resistance.^4,5 The ongoing arms race between microbes and antibiotics has led to the emergence of multidrug-resistant pathogens, rendering conventional antibiotics ineffective and threatening the efficacy of contemporary medical therapies.^6,7 In this post-antibiotic era, characterized by a paucity of treatment alternatives and the resurgence of infections previously under control, the development of novel antimicrobial approaches has become an urgent necessity.

Natural antimicrobial peptides (AMPs) are small cationic molecules produced by various organisms as part of their innate immune system, playing a crucial role in defending against a wide range of pathogens.⁸ AMPs have garnered significant attention for their potent antimicrobial activity and unique mechanisms of action.⁹ However, their practical application is hindered by high production costs, physiological instability and potential cytotoxicity.^10,11 Antibacterial poly(amino acids) offers a cost-effective and scalable solution that mimics the membrane-disruptive mechanisms of AMPs^12–15 while overcoming AMPs’ limitations through tunable molecular structures that enhance stability and improve efficacy.¹⁶ Polyaspartamides, a class of biocompatible and biodegradable polymers derived from aspartic acid, have found widespread applications in fields including drug delivery and tissue engineering owing to their versatile properties.^17,18 Research on antimicrobial polyaspartamides began in 1966, when “α,β-poly-DL-aspartic acid 3-aminopropylamide” was prepared from polysuccinimide (PSI) through aminolysis with 1,3-propanediamine. These polyaspartamides showed notable antimicrobial activity against Cryptococcus neoformans, Histoplasma capsulatum, Staphylococcus aureus, and Mycobacterium smegmatis.^19,20 However, to date, research on antibacterial polyaspartamides remains relatively scarce and is primarily confined to the effects of specific degrees of polymerization (DPs)^21–26 and side chain structures²⁷ on their antibacterial performance, lacking effective screening of different DPs and side chain structures as well as systematic studies on the structure–activity relationships between these structural parameters²⁸ and antibacterial activity. Such screening and in-depth exploration of structure–activity relationships are essential for guiding the rational design and performance optimization of antibacterial polyaspartamides.

In this study, we conducted a thorough investigation into the influence of the DP and side chain length on the antimicrobial activity and biocompatibility of PASP_nDA_m obtained from the aminolysis of PBLA with different diamines (n denotes the DP of polymer, and m denotes the number of carbon atoms located between two amino groups in the diamines). Based on the screening tests, PASP₁₀DA₆ had better antibacterial efficacy and biocompatibility compared to the other tested polyaspartamides. PASP₁₀DA₆ exerted its bactericidal effect through membrane disruption. Furthermore, in a bacterial infection model on mouse epidermis, PASP₁₀DA₆ demonstrated superior therapeutic performance compared to conventional antibiotics, highlighting its potential as an effective antimicrobial agent (Scheme 1).

Scheme 1 Schematic of the synthesis of PASP₁₀DA₆ for the treatment of S. aureus infection in mouse epidermis.

2. Experimental section

2.1. Synthesis of PASP_nDA_m

The aminolysis reaction was performed following a modified literature procedure.²⁷ PBLA₁₀ (200 mg, 0.095 mmol) was dissolved in anhydrous DMSO (10 mL) in a flame-dried flask under a nitrogen atmosphere. Ethylene diamine (50 eq. excess of benzyl ester units) was added, and the reaction mixture was stirred at room temperature for 1 h. After 1 h, the aminolyzed product was precipitated into an excess of cold diethyl ether (100 mL). The precipitate was collected by centrifugation (10 000 rpm, 10 min) and washed with cold ether (20 mL) to remove residual ethylene diamine and solvent. The aminolyzed polymer was dissolved in deionized water (5 mL) and transferred to a pre-treated dialysis tube (MWCO 200 Da). Dialysis was conducted against HCl solution (0.01 M, pH = 1.0) at room temperature with six changes of the dialysis medium over 48 h. The dialyzed polymer solution was lyophilized to yield the final product, PASP₁₀DA₂, as an off-white fluffy solid (85 mg). Additionally, aminolysis of PBLA samples with different DPs (20, 30, and 40) and different diamines (1,3-propanediamine, 1,4-butanediamine, 1,5-pentanediamine, and 1,6-hexanediamine) was conducted using the same method.

2.2. Minimum inhibitory concentration (MIC) assay

The MICs of PASP_nDA_m against S. aureus and E. coli were evaluated using the broth microdilution method according to Clinical and Laboratory Standards Institute (CLSI) guidelines. Bacterial strains were grown overnight on Luria–Bertani (LB) broth at 37 °C with shaking at 120 rpm. Then, bacteria were suspended in Mueller-Hinton (MH) broth. The bacterial suspensions were further diluted in MH broth to obtain a final inoculum of approximately 1 × 10⁶ CFU mL⁻¹. The PASP_nDA_m compound was dissolved in sterile deionized water to prepare a stock solution of 1 mg mL⁻¹. Two-fold serial dilutions were prepared in deionized water in a 96-well microtiter plate (100 μL per well). Each well was inoculated with another 100 μL of the diluted bacterial suspensions (1 × 10⁶ CFU mL⁻¹). Positive (bacteria suspended in equal volumes of MH broth and sterile deionized water) and negative controls (equal volumes of MH broth and sterile deionized water) were included. Plates were incubated at 37 °C for 24 h, and bacterial growth was assessed by measuring absorbance at 600 nm (OD₆₀₀). The MIC was defined as the lowest concentration of PASP_nDA_m that completely inhibited visible growth of the test organism. The MICs of the PASP_nDA_m against Methicillin-resistant Staphylococcus aureus (MRSA) were evaluated according to the same method.

2.3. In vitro cytotoxicity assay

L929 mouse fibroblast cells were seeded in 96-well plates at a density of 1 × 10⁴ cells per well in DMEM supplemented with 10% fetal bovine serum and 1% penicillin–streptomycin. The plates were incubated for 24 h at 37 °C in a humidified incubator containing 5% CO₂ to allow cell attachment. PASP_nDA_m was dissolved in sterile medium at a concentration of 1 mg mL⁻¹. Serial two-fold dilutions were prepared in cell culture medium to the desired final concentrations. After culturing for 24 h, the medium was removed and replaced with 100 μL of the PASP_nDA_m solutions (A_sample). Cells treated with medium only served as the non-toxic negative control (A_control). One hundred microliters of each dilution was added to the wells, with untreated cells serving as the negative control and wells containing only medium serving as the blank control. Each concentration was tested in triplicate. The plates were incubated for an additional 24 h. After 24 h, 10 μL of MTT solution in PBS (5 mg mL⁻¹) was added to each well. The plates were returned to the incubator for 4 h to allow metabolically active cells to form formazan crystals. The MTT solution was then discarded, and the insoluble formazan crystals were dissolved in 150 μL of DMSO per well with gentle shaking for 10 min. The absorbance was measured at 492 nm using a microplate reader. Cell viability was calculated as

Cell viability (%) = OD₄₂₀(A_sample)/OD₄₂₀(A_control) × 100%. (1)

The polymer concentration that inhibited 50% of cell growth (IC₅₀) was determined by non-linear regression analysis.

2.4. In vivo S. aureus-infected mouse wound model

Female Kunming mice (6–8 weeks old, weighing 18–20 g) were purchased from Liaoning Changsheng Biotechnology Ltd (Liaoning, China). All animal procedures were performed in accordance with the Guidelines for Care and Use of Laboratory Animals of Jilin University and approved by the Animal Ethics Committee of Jilin University.

An in vivo model of S. aureus-infected mouse wounds was established by inducing standardized wounds (d = 10 mm) on the epidermis of mice, followed by inoculation with S. aureus. After 24 h of wound infection, mice were treated with a topical solution containing PASP₁₀DA₆ on days 3, 5, and 7. In the PBS group (PG), only S. aureus infection was applied. In the vancomycin treatment group (VG), 20 μg mL⁻¹ vancomycin was used for the treatment. In the low-concentration PASP₁₀DA₆ treatment group (LG), 50 μg mL⁻¹ PASP₁₀DA₆ prepared in sterile water was administered after bacterial infection, while in the high-concentration PASP₁₀DA₆ treatment group (HG), 100 μg mL⁻¹ PASP₁₀DA₆ prepared in sterile water was used for treatment after bacterial infection. The non-infected group (NG) served as a model where wounds were created without bacterial infection. Skin samples were collected from the infected sites 24 h after the initial treatment (days 4) to assess bacterial burden. Samples were then homogenized and plated for bacterial enumeration. Additionally, photographs of the wound site were taken every two days to monitor the progression of infection and wound healing. On day 14, mice were euthanized, and wound skin samples were collected for histological examination. Hematoxylin and eosin (H&E) staining and Masson's trichrome staining were performed to evaluate tissue morphology, inflammation, and wound healing. Furthermore, vital organs, including the heart, liver, spleen, lungs, and kidneys, were harvested and subjected to histological examination using H&E staining to assess potential systemic effects of PASP₁₀DA₆ treatment. Blood samples were collected for routine blood analysis to evaluate systemic toxicity after treatment with PASP₁₀DA₆.

3. Results and discussion

3.1. Synthesis and characterization of PASP_nDA_m

The synthetic pathway of PASP_nDA_m is illustrated in Fig. 1a. The ¹H NMR spectrum confirmed the successful synthesis of β-benzyl L-aspartic acid N-carboxyanhydride (BLA-NCA), with a singlet peak at 6.31 ppm assigned to the NH proton of the NCA ring, a methine peak d (–CO–CH–NH–) at approximately 4.59 ppm, and a methylene peak e (–CH–CH₂–CO–) at 3.15–2.77 ppm, consistent with the expected structure (Fig. 1b).²⁹ Different DPs of PBLA_n were synthesized using the ring-opening polymerization of BLA-NCA, where n denotes the feeding molar ratio of BLA-NCA (monomer) to n-hexylamine (initiator). The molecular weight (M_n and M_w) and polydispersity indices (Đ) of PBLA_n were determined by gel permeation chromatography (GPC) (Fig. S1). The M_n values of PBLA₁₀, PBLA₂₀, PBLA₃₀, and PBLA₄₀ obtained by GPC were 2100, 4100, 5800, and 8000 g mol⁻¹, respectively, corresponding to DPs of 10, 20, 28, and 39, which closely match the feeding ratios of BLA-NCA to n-hexylamine, and these polymers showed low Đ values, ranging from 1.20 to 1.06 (Fig. 1c). The DP of PBLA was also decided from the proton nuclear magnetic resonance (¹H NMR) spectrum (Fig. S2–S5) by comparing the integral ratio of the peak area at 7.0–7.2 ppm (the benzene ring) with that at 0.7 ppm (the methyl group of the n-hexylamine end group). The DPs of PBLA₁₀, PBLA₂₀, PBLA₃₀, and PBLA₄₀ calculated from the ¹H NMR spectra were 10, 15, 25, and 42, respectively, which are also close to the feeding ratios of BLA-NCA to n-hexylamine.^27,28 PASP_nDA_m polymers were successfully synthesized through the aminolysis of PBLA using linear diamines. The ¹H NMR spectrum of PASP₁₀DA_m revealed the disappearance of peaks corresponding to the benzyl ring (7.0–7.2 ppm) and the benzylic methylene (4.9–5.0 ppm). Peaks corresponding to the initiator (n-hexylamine, CH₃CH₂–) remained unchanged, while new methylene peaks (1.0–2.0 ppm) from the linear diamines were observed (Fig. 1d and e). The DP of these polyaspartamides showed no significant change. These observations suggested that the linear diamines were successfully grafted onto the polypeptide backbone. PASP₁₀DA₂, PASP₁₀DA₃, PASP₁₀DA₄, PASP₁₀DA₅ and PASP₁₀DA₆ were successfully synthesized. Similarly, PBLA polymers with other DPs (20, 30 and 40) were successfully aminolyzed using linear diamines of varying side chain lengths (Fig. S6–S20).

Fig. 1 Synthesis of PASP_nDA_m. (a) Synthesis scheme for PASP_nDA_mvia ring-opening and aminolysis reactions. Here, n stands for the DP (n = 10, 20, 30, and 40), and m stands for the number of carbon atoms in the linear diamines (m = 2, 3, 4, 5, and 6). (b) ¹H NMR spectra of BLA in deuterated chloroform, with a few drops of deuterated trifluoroacetic acid, and BLA-NCA in deuterated chloroform. (c) Molecular weight, DP, and polydispersity of PBLA_n determined by GPC and ¹H NMR spectroscopy, where α is calculated from the number-average molecular weight obtained by GPC; β is determined from the ¹H NMR spectrum of PBLA. (d) ¹H NMR spectra of PASP₁₀DA_mvia aminolysis (m = 2, 3, 4, 5, and 6). (e) Chemical structure of PASP₁₀DA_m.

3.2. Antibacterial activity and biocompatibility of PASP_nDA_m

We conducted a systematic evaluation of the antimicrobial efficacy of these PASP_nDA_m against both Gram-positive (S. aureus) and Gram-negative (E. coli) bacteria by assessing their ability to inhibit bacterial growth (Fig. S21). All PASP_nDA_m exhibited inhibitory activity against both S. aureus and E. coli. As displayed in Fig. 2a and Table S1, PASP₁₀DA₅ exhibited the weakest inhibitory effect against S. aureus, with an MIC of 125 μg mL⁻¹, while PASP₄₀DA₄ showed the strongest inhibition, with the lowest MIC of 3.9 μg mL⁻¹. Most PASP₄₀DA_m samples showed lower MIC values compared to PASP₁₀DA_m, showing that increasing the DP could contribute to the improved antibacterial effectiveness. Moreover, in most cases, PASP_nDA₆ (n = 10, 20, 30, and 40) consistently presented lower MIC than polypeptides with other diamine side chains. For example, the MIC of PASP₁₀DA₆ against S. aureus was 7.8 μg mL⁻¹, which was significantly lower than those of PASP₁₀DA_m (m = 2, 3, 4 and 5). Like the inhibitory effect against S. aureus, PASP_nDA_m also proved an inhibitory effect against E. coli. The MICs of PASP₁₀DA₂ and PASP₁₀DA₅ were 62.5 μg mL⁻¹, while PASP₁₀DA₆ and PASP₄₀DA₃ exhibited significantly lower MICs of 7.8 μg mL⁻¹ (Fig. 2b and Table S2). Interestingly, PASP₁₀DA₆ maintained its effectiveness in killing E. coli, which suggested that PASP₁₀DA₆ could effectively kill both Gram-negative and Gram-positive bacteria. Generally, the observed variations in MIC across different PASP_nDA_m suggest that both the DP and side chain length play critical roles in the antimicrobial activity of PASP_nDA_m. Higher DP appeared to enhance the interaction of the polymer with bacteria, leading to enhanced membrane disruption activity.^30,31 At DPs of 10 and 20, the antibacterial activity of PASP_nDA_m increased with increasing side chain length. However, at DPs of 30 and 40, antibacterial activity initially increased and then decreased as the side chain length increased. The length of the side chain had a notable influence on the antimicrobial activity of PASP₁₀DA_m; as the side-chain carbon number increased from 2 to 6, the MIC of PASP₁₀DA_m decreased from 125 to 7.8 μg mL⁻¹. For side chain containing 2, 3, 4 and 5 carbon atoms, the antibacterial activity of PASP_nDA_m increased with increasing DP. In contrast, for a side chain with 6 carbon atoms, a slightly opposite trend was observed. In addition, we verified the antibacterial properties of the polymers against clinically drug-resistant bacteria. As shown in Table S3, PASP_nDA_m showed certain antibacterial performance against MRSA. Considering the complex environment of infected tissues, we examined whether the antibacterial activity of PASP₁₀DA₆ would be affected under variable conditions. After different treatments, the MIC value of PASP₁₀DA₆ remained unchanged (Table S4), demonstrating the good stability of the polymer.

Fig. 2 Antibacterial effect and toxicity of PASP_nDA_m. (a) MIC of PASP_nDA_m against S. aureus (μg mL⁻¹). (b) MIC of PASP_nDA_m against E. coli (μg mL⁻¹). (c) IC₅₀ values of PASP_nDA_m against L929 cells (μg mL⁻¹). (d) Selectivity index of PASP_nDA_m (calculated as the ratio of IC₅₀ for L929 cells to MIC against S. aureus).

Further experiments were conducted to evaluate the cytotoxicity and hemolytic effects of these polypeptides. The cytotoxicity and hemolytic activities of PASP_nDA_m were measured against the mouse fibroblast cell line L929 (Fig. S22) and rabbit red blood cells (Fig. S23), respectively. PASP_nDA₂ and PASP₁₀DA_m exhibited good biocompatibility, with IC₅₀ values around 1000 μg mL⁻¹; however, most of these had low antimicrobial activity. Notably, PASP_nDA₆ showed a trend of increased cytotoxicity with increasing DP. The IC₅₀ value was 750 μg mL⁻¹ for PASP₁₀DA₆, which decreased to 80 μg mL⁻¹ for PASP₄₀DA₆ (Fig. 2c and Table S5). Based on these observations, the cytotoxicity risk of PASP_nDA_m against L929 cells increases with both the DP and side chain length. Regarding hemolytic activity, most PASP_nDA_m samples showed HC₅ values greater than 2000 μg mL⁻¹, showing that these polymers were well-tolerated by red blood cells (Table S6). This suggests a low possibility of causing hemolysis by PASP_nDA_m at a concentration around their MIC values. Based on the antibacterial and cytotoxicity assays, the selectivity index (SI) was then calculated as the ratio of IC₅₀ for L929 cells to the MIC against S. aureus to evaluate the balance between antimicrobial activity and cytotoxicity (Table S7). Due to its low MIC values against bacteria and high IC₅₀ values for L929 cells, PASP₁₀DA₆ exhibited the highest SI of 96 among all tested PASP_nDA_m (Fig. 2d), showing that PASP₁₀DA₆ could be an effective and safe therapeutic agent.

3.3. Bactericidal mechanisms of PASP₁₀DA₆

To evaluate the antibacterial mechanism of PASP₁₀DA₆, scanning electron microscopy (SEM) was used to reveal surface alterations in S. aureus and E. coli following treatment with the PASP₁₀DA₆. After treatment with PASP₁₀DA₆, obvious morphological changes were observed on the surfaces of both S. aureus and E. coli, characterized by surface depressions and blurred boundaries on the bacteria (Fig. 3a). By contrast, negligible alterations were observed in the morphology of bacteria treated with norfloxacin and PBS. Norfloxacin exerts its bactericidal effect by inhibiting bacterial DNA gyrase and topoisomerase IV, essential enzymes involved in bacterial DNA replication and transcription.³² This mechanism effectively halts bacterial proliferation without causing significant changes to bacterial surface morphology. Next, a live/dead assay was conducted to further investigate bacterial membrane damage after treatment with PASP₁₀DA₆. The simultaneous use of SYTO 9 and PI dyes allowed differentiation between live and damaged bacteria, i.e., viable bacteria with intact membranes exhibit strong green fluorescence and minimal PI uptake, while bacteria with damaged membranes show increased red fluorescence from PI and potentially reduced SYTO 9 signals.³³ Following a 3-hour exposure period, PASP₁₀DA₆-treated samples displayed a higher proportion of red-stained bacteria under fluorescence microscopy compared to norfloxacin-treated samples. By contrast, PBS and norfloxacin treatment groups showed no significant red fluorescence, suggesting that these treatments do not disrupt bacterial membranes. Therefore, the presence of red fluorescence in the PASP₁₀DA₆-treated group confirmed its ability to induce membrane damage (Fig. 3b and S24).

Fig. 3 Bactericidal mechanism of PASP₁₀DA₆ against S. aureus and E. coli. (a) SEM images of S. aureus and E. coli after treatment with 1 × MIC of norfloxacin (0.04 μg mL⁻¹) and PASP₁₀DA₆. (b) Fluorescence images of S. aureus post-treatment with 1 × MIC norfloxacin (0.04 μg mL⁻¹) and PASP₁₀DA₆. (c) CD spectra of PASP₁₀DA₆ before and after interaction with POPG or POPG + POPE. (d) Surface potential alterations in S. aureus and E. coli. exposed to PASP₁₀DA₆. (e) Membrane depolarization effects of PASP₁₀DA₆ on S. aureus. (f) Membrane depolarization effects of PASP₁₀DA₆ on E. coli. (g) Permeability of the E. coli membrane in response to PASP₁₀DA₆ exposure. (h) Leakage of nucleic acids from S. aureus and E. coli treated with PASP₁₀DA₆.

To gain further insights into the antibacterial mechanism of PASP₁₀DA₆, the interaction between PASP₁₀DA₆ and the bacterial membrane was investigated. The relationship between the antimicrobial efficacy of polypeptides and their secondary structures is a crucial aspect of understanding their mode of action.³⁴ Our observations revealed that PASP₁₀DA₆ remained in a random coil conformation in both pure and POPG-containing aqueous solution (Fig. 3c). PASP₁₀DA₆ did not undergo secondary structural changes when transitioning from pure aqueous solution to membrane-simulating environments (POPG-containing solutions), suggesting that its antibacterial activity does not rely on a transition of secondary structure. The absence of a secondary structure transition upon membrane interaction may contribute to the low cytotoxicity of PASP₁₀DA₆. This was because keeping a random coil conformation could reduce its tendency to insert into mammalian cell membranes, thereby minimizing membrane disruption and cytotoxic effects.³⁵ Adsorption of cationic polypeptides onto the negatively charged surfaces of bacteria can result in alterations in the bacterial ζ-potential. As shown in Fig. 3d, when the concentration of PASP₁₀DA₆ was 50 μg mL⁻¹, the surface potential of the bacteria increased by 10 mV. The shift in the bacterial ζ-potential suggests that the cationic PASP₁₀DA₆ successfully binds to the bacterial membrane through electrostatic interactions. The action of PASP₁₀DA₆ on the bacterial membrane was further investigated through membrane depolarization experiments. As displayed in Fig. 3e and f, PASP₁₀DA₆ demonstrated its potential in disrupting bacterial homeostasis through significant membrane depolarization in both S. aureus and E. coli. The degree of membrane depolarization increased with increasing concentrations of PASP₁₀DA₆. This result provided evidence of the polypeptide's ability to destabilize bacterial membranes, leading to impaired bacterial function and contributing to its antimicrobial effect.³⁶

The ortho-nitrophenyl-β-galactoside (ONPG) cleavage method is particularly useful in the study of the effects of polypeptides on the integrity of bacterial membranes. ONPG cannot traverse the bacterial membrane to be cleaved by cytoplasmic β-galactosidase into the yellow compound o-nitrophenol (ONP) unless permeabilization of the bacterial membrane is increased.³⁷ Enhanced permeability changes were observed in E. coli and S. aureus exposed to PASP₁₀DA₆, as indicated by increased ONPG cleavage (Fig. 3g and S25). The observed increase in membrane permeability further confirms the results of the bacterial membrane depolarization after treatment with polypeptides.^38–40 Furthermore, nucleic acid leakage from bacteria after the treatment with PASP₁₀DA₆ was investigated by UV-Vis absorption spectroscopy (Fig. 3h). Peaks at 260 nm were clearly observed after treatment with PASP₁₀DA₆, indicating a large amount of nucleic acid leakage.^41,42 Taken together, all the abovementioned results indicate that the antimicrobial mechanism of PASP₁₀DA₆ involves the binding of PASP₁₀DA₆ to bacterial membranes via electrostatic interactions, triggering membrane depolarization and disruption.

3.4. Killing kinetics, resistance development, and biofilm disruption of PASP₁₀DA₆

It has been reported that AMPs exhibit more rapid bactericidal kinetics than antibiotics.⁴³ Accordingly, we investigated the bactericidal rate of PASP₁₀DA₆. Antimicrobial kinetics analysis revealed the rapid bactericidal action of PASP₁₀DA₆ (Fig. 4a and b). Specifically, PASP₁₀DA₆ eliminated 100% of S. aureus within 20 min at 1 × MIC, while 2 × MIC was required to achieve similar effects against E. coli. The killing rate of Gram-negative bacteria was slightly lower compared to Gram-positive bacteria, possibly due to the presence of the outer membrane in Gram-negative bacteria.⁴⁴ At 4 × MIC, PASP₁₀DA₆ effectively eradicated all bacteria within approximately 1 min. By contrast, vancomycin and norfloxacin did not show significant bactericidal activity within 20 min, similar to the control group (Fig. 4c and d). These results highlight that the bactericidal efficacy of PASP₁₀DA₆ is both time- and concentration-dependent. Moreover, the membrane-disrupting bactericidal mechanism of PASP₁₀DA₆ leads to rapid bacterial death, which differs from the mechanisms of vancomycin and norfloxacin.

Fig. 4 Antimicrobial activity and resistance induction by PASP₁₀DA₆. (a) Time-dependent plating results for S. aureus treated with PASP₁₀DA₆; “van.” means vancomycin (2 μg mL⁻¹). (b) Time-dependent plating results for E. coli treated with PASP₁₀DA₆; “nor.” means norfloxacin (0.04 μg mL⁻¹). (c) Survival of S. aureus at different time points following incubation with antimicrobial agents, based on data given in (a). (d) Survival of E. coli at different time points following incubation with antimicrobial agents, based on data given in (b). (e) Resistance induction in S. aureus by vancomycin and PASP₁₀DA₆. (f) Resistance induction in E. coli by norfloxacin and PASP₁₀DA₆. (g) Inhibition of S. aureus biofilm formation by PASP₁₀DA₆, using vancomycin (MIC = 2 μg mL⁻¹) as a control. (h) Biofilm dispersion in mature S. aureus cultures. (i) Inhibition of E. coli biofilm formation by PASP₁₀DA₆, with norfloxacin (MIC = 0.04 μg mL⁻¹) as a control. (j) Dispersion of mature E. coli biofilms. *p < 0.05, **p < 0.01, and ***p < 0.001.

Bacterial resistance to antibiotics is a significant global health threat.^45,46 This resistance not only complicates treatment strategies but also increases healthcare costs. Unlike traditional antibiotics, most AMPs act on bacterial membranes, a mechanism that reduces the likelihood of resistance development.^47–49 As expected, over a 15-day period, PASP₁₀DA₆ did not induce bacterial resistance in both S. aureus and E. coli, while the MIC values of vancomycin and norfloxacin increased by 256-fold and 32-fold, respectively (Fig. 4e and f). The absence of resistance development in bacteria treated with PASP₁₀DA₆ emphasizes its potential as a sustainable antimicrobial strategy.

Biofilms, formed by microbial communities, pose significant hazards, as they provide protection for pathogens, promote antibiotic resistance, and cause surface contamination to increase the risk of chronic infections.⁵⁰ Preventing and controlling biofilm formation are crucial for keeping environmental hygiene, reducing healthcare-associated infections, and safeguarding human health.^51–53 Therefore, we evaluated the inhibition effect of PASP₁₀DA₆ on the formation of biofilm. As shown in Fig. 4g, PASP₁₀DA₆ markedly inhibited the formation of S. aureus biofilms compared to vancomycin, reducing biofilm formation rate by 65% and 31% for PASP₁₀DA₆ and vancomycin, respectively. For mature biofilms, PASP₁₀DA₆ at 8-fold MIC dispersed approximately 55% of the biofilm mass, significantly outperforming vancomycin, which only achieved 24% dispersion (Fig. 4h). For E. coli, PASP₁₀DA₆ showed a 58% reduction in biofilm formation at 1-fold MIC, surpassing the 30% reduction by norfloxacin at 1-fold MIC (Fig. 4i). PASP₁₀DA₆ also proved a strong dispersion effect on mature E. coli biofilms, disrupting up to 46% at 8-fold MIC compared to 27% by norfloxacin (Fig. 4j). The inhibitory and eradication effects of PASP₁₀DA₆ on biofilms were superior to those of antibiotics at the same MIC.

3.5. In vivo antibacterial efficacy

Inspired by the high antibacterial activity and good biocompatibility of PASP₁₀DA₆, we evaluated the therapeutic efficacy of PASP₁₀DA₆ against S. aureus-infected epidermal wounds. Bacterial infections were induced in epidermal wounds, followed by treatment with PASP₁₀DA₆ and vancomycin. The progression of infection and wound healing was monitored to assess the effectiveness of PASP₁₀DA₆ in reducing bacterial burden and promoting recovery under physiological conditions. The mice were divided into five groups: the PBS group (PG, infected but non-treated), vancomycin treatment group (VG, 20 μg mL⁻¹), low-concentration PASP₁₀DA₆ treatment group (LG, 50 μg mL⁻¹), high-concentration PASP₁₀DA₆ treatment group (HG, 100 μg mL⁻¹), and the non-infected group (NG). The timeline and sampling points were carefully planned to capture both immediate and extended impacts of the treatments (Fig. 5a). Representative images illustrate the progression of infected skin lesions during treatment (Fig. 5b), with the diameter of the infected area visibly decreasing in the PASP₁₀DA₆-treated groups. On the fourth day, PASP₁₀DA₆ was able to eliminate more than 99% of the bacteria at the wound site, while vancomycin treatment achieved a reduction of only 90% (Fig. 5c). Due to its significant reduction of bacterial burden at the wound site, the PASP₁₀DA₆-treated group showed a notably smaller wound area on day seven compared to the PG and the VG (Fig. 5d). Throughout the wound healing process, the VG and PASP₁₀DA₆-treated groups showed comparable effects in reducing wound area (Fig. 5e). These findings indicate that PASP₁₀DA₆ could effectively kill bacteria and then accelerate the wound healing process.

Fig. 5 Application of PASP₁₀DA₆ in the treatment of murine skin infection. (a) Timeline of PASP₁₀DA₆ administration and sample collection points. (b) Representative images of infected skin wounds after treatment (ring diameter: 10 mm). (c) Bacterial count results from the tissue coating plates after fragmentation of infected tissues on day 4. (d) Bar graph comparing wound area percentages across different groups on the seventh day. (e) Quantitative analysis of lesion size over the treatment period. *p < 0.05, **p < 0.01, and ***p < 0.001. Error bars indicate standard deviation (n = 5).

IL-1β (Interleukin-1β), a marker of pro-inflammatory M1 macrophages, indicated an active inflammatory response associated with bacterial infection in the mouse epidermal wound model. Conversely, ARG-1 (Arginase-1), a marker of anti-inflammatory M2 macrophages, suggested a transition toward tissue repair and resolution of infection, reflecting controlled inflammation and progression toward wound healing. Fluorescence staining with DAPI/IL-1β, as well as DAPI/ARG-1, was performed to analyze the healing of the mouse wound (Fig. 6a and b). In the later stages of wound healing, red fluorescence was predominantly localized in the wound area, and there were no significant differences in IL-1β fluorescence among all groups, which might reflect that inflammation in the wound had reached a steady state by the end of treatment (Fig. 6c). However, differences were observed in ARG-1 fluorescence levels across the groups. The HG exhibited the lowest green fluorescence intensity for ARG-1, lower than that of the NG group. Conversely, the PG demonstrated the highest fluorescence intensity, while the VG and the LG showed no significant differences from each other (Fig. 6d). Low ARG-1 expression in the HG indicated that, by the end of the treatment period, wounds in the HG had restored near-normal tissue structure, potentially surpassing the non-infected NG. By contrast, high ARG-1 expression in the PG suggested ongoing rapid tissue repair. These results aligned with H&E staining observations. These results indicate that PASP₁₀DA₆ could promote wound healing and tissue repair.

Fig. 6 Immunofluorescence analysis of mouse wound sections. (a) Immunofluorescence staining for IL-1β (red) and cell nuclei with DAPI (blue). (b) Immunofluorescence staining for ARG-1 (green) and cell nuclei with DAPI (blue). (c) Quantification of relative areas of red fluorescence (IL-1β). (d) Quantification of relative areas of green fluorescence (ARG-1). Statistical significance is denoted as *p < 0.05, **p < 0.01, and ***p < 0.001.

Following the animal experiments, physiological toxicity assessments were conducted to ensure the antimicrobial efficacy without causing significant harm to normal tissues. Histological examination of heart, liver, spleen, lung, and kidney tissues revealed no significant changes (Fig. S26), indicating minimal biotoxicity of PASP₁₀DA₆. Hematological analysis further supported the biocompatibility of PASP₁₀DA₆. Hematological results showed no significant changes in white blood cells (WBC), lymphocytes (LYM), neutrophils (NEU), platelets (PLT), red blood cells (RBC), or hemoglobin (HGB) levels (Fig. S27), indicating that PASP₁₀DA₆ did not cause immune activation, inflammation, or hematological abnormalities. These results indicate that PASP₁₀DA₆ is a safe and effective antibacterial poly(amino acid) suitable for treating bacterial infections.

4. Conclusion

In summary, this study systematically investigated the impact of side chain lengths and DP on the antibacterial properties of polyaspartamides. In vitro antibacterial experiments showed that antimicrobial activity increased with increasing length of side chains and DP. However, when the DP of PASP_nDA_m exceeded 20, further increases in DP did not improve antimicrobial efficacy of PASP_nDA_m but instead led to increased hemolytic and cytotoxic effects. Among all the tested PASP_nDA_m, PASP₁₀DA₆ exhibited a relatively low MIC and the highest selectivity against both S. aureus and E. coli. The antimicrobial mechanism of PASP₁₀DA₆ involved adsorption onto negatively charged bacterial surfaces via electrostatic interactions and disruption of membrane integrity. Unlike traditional antibiotics, PASP₁₀DA₆ did not induce bacterial resistance and was more effective in the inhibition of biofilm formation and dispersion of mature biofilms. In vivo antibacterial studies demonstrated that PASP₁₀DA₆ not only effectively eradicates bacterial pathogens but also promoted wound healing. These findings underscore the potential of PASP₁₀DA₆ as a valuable antimicrobial agent with promising applications for treating infections that require biofilm prevention and resistance management.

Author contributions

Wenlong Zhang: writing—original draft, investigation, formal analysis, and methodology; Guowenlie Gao: formal analysis and methodology; Yuqian Ji: formal analysis and methodology; Mingming Zhao: formal analysis; Pengqi Wan: writing—review and editing, supervision, and funding acquisition; Chunsheng Xiao: writing—review and editing, conceptualization, supervision, investigation, formal analysis, resources, and funding acquisition; Xuesi Chen: writing—review and editing, conceptualization, supervision, investigation, formal analysis, and resources. All the authors approved the final version of the manuscript.

Conflicts of interest

There are no conflicts to declare.

Data availability

The data supporting this article have been included as part of the SI.

The Supplementary Information provides additional experimental section, NMR spectra of intermediates and polymers, and supporting figures that complement the main findings of the study. See DOI: https://doi.org/10.1039/d5bm00994d.

Acknowledgements

The work was financially supported by the National Natural Science Foundation of China (52222307, U21A2089) and the Youth Growth Technology Project of the Jilin Provincial Department of Science and Technology (20230508153RC).

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