Light-mediated activation of nitric oxide and antibacterial polymers for anti-biofilm applications

Abstract

Bacterial biofilms remain a major challenge in treating persistent infections due to their dense extracellular matrix and inherent antibiotic resistance. Herein, we propose a light-responsive nanoparticle system (PNO@Ir) that integrates a nitric oxide (NO) donor polymer (PNO) with the photosensitizer fac-Ir(ppy)₃. Upon green light irradiation, NO release and activation of primary amine-containing antibacterial polymers are triggered via a dual mechanism involving triplet–triplet energy transfer (TTET) and photoinduced electron transfer (PeT). Under mildly acidic and hypoxic conditions, protonation of the exposed amines induces nanoparticle reorganization, leading to surface charge reversal and enhanced bacterial affinity. Both in vitro and in vivo studies, including a murine wound infection model, demonstrate that this cascade-activation strategy disrupts methicillin-resistant Staphylococcus aureus (MRSA) biofilms. This work presents a synergistic and spatiotemporally controllable platform for NO delivery and antibacterial polymer activation, offering significant potential for combating antibiotic-resistant bacterial infections.

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Zhiqiang Shen

Dr Zhiqiang Shen received his PhD degree in Science from the School of Chemistry and Materials Science at the University of Science and Technology of China (USTC) in 2022. He subsequently conducted postdoctoral research at USTC and is currently an Associate Research Fellow at the Suzhou Institute for Advanced Research, USTC. His research focuses on the design and synthesis of biomedical polymers and gas-delivery polymeric systems for biomedical applications.

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1. Introduction

Antibiotic resistance within bacterial biofilms represents a central challenge in the treatment of infectious diseases, arising from the biofilm's unique structure and dynamic microenvironment.¹ Biofilms evade antimicrobial agents through multifaceted defense strategies, in which pH gradients and hypoxic environments play critical roles in the development of resistance.^2–5 The dense extracellular matrix not only acts as a physical barrier impeding drug penetration but also fosters an acidic and oxygen-depleted environment through localized metabolic processes, such as anaerobic respiration, which generate acidic byproducts (e.g., CO₂ and organic acids).² This self-constructed microenvironment drives phenotypic adaptations in bacteria: the low pH and hypoxia suppress metabolic activity, promoting the formation of slow-growing or dormant subpopulations that exhibit intrinsic tolerance to antibiotics.^6–8 Concurrently, acid stress triggers transcriptional reprogramming, such as the upregulation of multidrug efflux pumps (e.g., NorB)⁹ and stress response regulators (e.g., RpoS),¹⁰ further enhancing resistance mechanisms.^11,12 Notably, pH and hypoxia play dual regulatory roles, and they are both the consequences of biofilm metabolic activity and key modulators that exacerbate resistance by impairing drug efficacy and altering bacterial physiology.^5,13 As a result, biofilm-induced resistance presents a complex and multifaceted obstacle in clinical treatment.^5,14

To address this issue, various strategies have been developed, including nanoplatforms incorporating gaseous signaling molecules,^4,15–19 antimicrobial peptides,^20–22 and amine-containing cationic polymers.^23–25 Among these, pioneering work by Meyerhoff,²⁶ Handa,²⁷ Schoenfisch,²⁸ and others²⁹ has demonstrated the potential of nitric oxide (NO) as an effective anti-biofilm agent, which primarily involves the inhibition of extracellular matrix production,³⁰ disruption of metabolic pathways,^31–33 and modulation of signaling molecules such as cyclic-di-GMP and quorum sensing (QS) regulators.³⁴ However, conventional small-molecule NO donors, such as S-nitrosothiols (RSNOs) and N-diazeniumdiolates (NONOates), suffer from limitations including spontaneous NO release and short half-lives,³⁵ which hinder their ability to achieve spatiotemporal control over NO delivery and thereby reduce their therapeutic efficacy. Light activation has emerged as a promising non-invasive approach for achieving controllable NO release.³⁶ Nevertheless, most light-responsive NO donors are triggered by ultraviolet (UV) or near-UV light, which is associated with phototoxicity and limited tissue penetration, rendering them unsuitable for biological applications.³⁷ Therefore, the development of NO donors that can be activated by longer-wavelength light is of significant importance for advancing biofilm-targeted therapies.¹⁵

In addition, the anti-biofilm activity of antimicrobial peptides (AMPs) arises from their unique amphiphilic structures, characterized by the synergistic presence of cationic and hydrophobic domains.^38–40 The positively charged amine groups electrostatically bind to negatively charged phospholipids on the bacterial membrane surface (e.g., phosphatidylglycerol and cardiolipin), facilitating membrane targeting and translocation. Simultaneously, the hydrophobic domains insert into the lipid bilayer, disrupting membrane integrity and causing pore formation or membrane lysis, ultimately leading to leakage of intracellular contents and bacterial death. Inspired by this mechanism, many currently developed amine-containing cationic polymers are designed to achieve anti-biofilm effects: under mildly acidic conditions, the amine groups become protonated, generating cationic charges that, when combined with hydrophobic segments and controlled polymer chain length,^23,41–43 enable effective biofilm disruption.

In contemporary anti-biofilm strategies, combination therapies using two or more agents with distinct mechanisms are often employed to achieve synergistic effects, allowing for reduced individual drug dosages and minimized systemic toxicity.^24,44–47 Here, we developed a photoresponsive polymeric nanoparticle system that integrates light-triggered NO release with the in situ generation of antibacterial polymers. Specifically, we designed an N-nitrosamine-based NO donor featuring a coumarin scaffold and a cleavable carbamate group, which was polymerized via reversible addition–fragmentation chain transfer (RAFT) to yield a NO-releasing random copolymer (PNO, Scheme 1a). Upon co-assembly with the photosensitizer fac-Ir(ppy)₃, the resulting nanoparticles (PNO@Ir) exhibited dual NO activation through triplet–triplet energy transfer (TTET) and photoinduced electron transfer (PeT) under 500 nm light irradiation under hypoxic conditions. Concurrently, light-induced carbamate cleavage exposed primary amines, enabling the formation of antibacterial polymers. Under mildly acidic environments, protonation of these amines triggered a surface charge reversal, enhancing the nanoparticles’ interaction with bacterial membranes and biofilms (Scheme 1b). This spatiotemporally controlled cascade activation strategy demonstrated promising anti-biofilm efficacy against methicillin-resistant Staphylococcus aureus (MRSA) both in vitro and in a murine wound infection model (Scheme 1c).

Scheme 1 Schematic illustration of (a) light-triggered NO release and antimicrobial polymer generation from the random copolymer PNO via carbamate cleavage under 500 nm light in the presence of fac-Ir(ppy)₃. (b) Self-assembly of PNO and fac-Ir(ppy)₃ into nanoparticles (PNO@Ir), which undergo NO release and surface charge switching under hypoxia and mild acidity conditions. (c) Light-activated NO release and antimicrobial polymer formation from PNO@Ir micelles in biofilm microenvironments, enabling disruption of MRSA biofilms.

2. Experimental section

2.1. Materials

Iodomethane, dibutyltin dilaurate (DBTL), tetrabutylammonium perchlorate, sodium nitrite (NaNO₂), sodium ascorbate (SA), potassium carbonate (K₂CO₃), butyl methacrylate (BMA), poly(ethylene glycol)methyl ether methacrylate (M_n = 500) and 2-phenyl-4,4,5,5-tetramethylimidazoline-1-oxyl 3-oxide (PTIO) were purchased from Sinopharm Chemical Reagent Co., Ltd. 2,2'-Azobis(4-methoxy-2,4-dimethylvaleronitrile) (V70), 2,2′-azobis(2-methylpropionitrile) (AIBN), 2-isocyanatoethyl methacrylate and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) were purchased from Sigma Aldrich. LIVE/DEAD® BacLight™ bacterial viability kit reagents (L13152, Molecular Probes, Thermo Fisher) were used as received. Water was deionized with a Milli-Q SP reagent water system (Millipore) to a specific resistivity of 18.4 MΩ cm. Benzyl 4-cyano-4(((ethylthio)carbonothioyl)thio)pentanoate (Ph-CTA), CouNO, CouNH, CouNMe, CouNO monomer and CouNMe monomer were synthesized according to the previously reported literature.^15,48 All other reagents were purchased from Sinopharm Chemical Reagent Co., Ltd and used as received unless otherwise stated.

2.1.1. Sample preparation. Synthetic schemes employed for the preparation of the CouNO monomer, CouNH monomer and CouNMe monomer; PNO and PNMe are shown in Schemes S1 and S2 (ESI†).

2.2. Synthesis of the CouNH monomer (Scheme S1, ESI†)

CouNH (0.8 g, 2.57 mmol) was dissolved in THF (20 mL), and DBTL (150 μL) was added. After this, 2-isocyanatoethyl methacrylate (0.48 g, 3.08 mmol) was added into the mixture. The mixture was stirred at room temperature overnight. After evaporation of the solvent under vacuum, the crude product was further purified by flash chromatography to obtain the CouNH monomer as a yellowish solid (0.99 g, yield: 83%).

2.3. Synthesis of PNO through RAFT polymerization (Scheme S2, ESI†)

The CouNO monomer, V70, Ph-CTA, OEGMA, and BMA were added into a reaction tube and dissolved in 0.3 mL of DMSO with a magnetic stirring bar. The reaction tube was carefully degassed by three freeze–pump–thaw cycles and was sealed under vacuum. After stirring at 37 °C for 20 h, the polymerization reaction was quenched by liquid nitrogen. The tube was then opened, and the reaction mixture was precipitated into an excess of diethyl ether (50 mL). The above dissolution–precipitation cycle was repeated three times. The precipitate was then collected by filtration and was dried in a vacuum oven overnight at room temperature, affording a yellowish solid. The degree of polymerization, DP, of the resulting random polymer was analyzed by ¹H NMR analysis and was denoted as P(OEGMA_0.38-co-BM_0.25-co-CouNO_0.37)₆₀ (abbreviated as PNO, Scheme S2, ESI†). GPC analysis revealed an M_n of 21.5 kDa and an M_w/M_n of 1.25 (Table S2, ESI†).

2.4. Synthesis of PNMe through RAFT polymerization (Scheme S2, ESI†)

The CouNMe monomer, AIBN, Ph-CTA, OEGMA, and BMA were added into a reaction tube and dissolved in 0.3 mL of DMSO with a magnetic stirring bar. The reaction tube was carefully degassed by three freeze–pump–thaw cycles and was sealed under vacuum. After stirring at 70 °C for 8 h, the polymerization reaction was quenched by liquid nitrogen. The tube was then opened, and the reaction mixture was precipitated into an excess of diethyl ether (50 mL). The above dissolution–precipitation cycle was repeated three times. The precipitate was then collected by filtration and was dried in a vacuum oven overnight at room temperature, affording a yellowish solid. The degree of polymerization, DP, of the resulting random polymer was analyzed by ¹H NMR analysis and was denoted as P(OEGMA_0.39-co-BM_0.25-co-CouNMe_0.36)₆₁ (abbreviated as PNMe, Scheme S2, ESI†). GPC analysis revealed a M_n of 25.8 kDa and a M_w/M_n of 1.27 (Table S2, ESI†).

2.5 500 nm Light-mediated NO release and quantification of NO release

To study the NO release behavior of the CouNO monomer, the fac-Ir(ppy)₃ photosensitizer and the CouNO monomer were dissolved in DMSO and the mixture was irradiated with an LED lamp (500 nm; Zhongshan Hongye Luminescence Co. Ltd). The evolution of UV/Vis absorbance spectroscopy was then recorded and the NO-releasing amounts were calculated following the reported protocol using the following equations.

A₁ = c₀l(ε^A₁x^A+ ε^B₁x^B)

A₂ = c₀l(ε^A₂x^A+ ε^B₂x^B)

where A₁ and A₂ are the absorbances of the CouNO monomer and CouNH monomer at the wavelengths of λ₁ and λ₂, respectively; ε^A₁, ε^A₂, ε^B₁, and ε^B₂ are the molar extinction coefficients of the CouNO monomer and CouNH monomer at the wavelengths of λ₁ and λ₂ (Fig. S4, ESI†), respectively; C₀ is the initial concentration of the CouNO monomer and l is the light path length (=1 cm); x^A and x^B are the molar contents of the CouNO monomer and CouNH monomer at any irradiation time points. Note that the quantitative conversion of the CouNO monomer and CouNH monomer led to the release of an equivalent amount of NO. Therefore, the NO-releasing amounts could be indirectly calculated by UV/Vis absorbance spectroscopy. For micellar nanoparticles (0.2 g L⁻¹) of PNO, the 500 nm light-triggered NO release was conducted in the presence of SA (10 mM). Following a similar protocol, the UV/Vis absorbance spectra under red light irradiation were monitored and the NO-releasing contents were calculated.

2.6 Photolysis monitoring by high performance liquid chromatography (HPLC).

To monitor the photodegradation process of the CouNO monomer in the presence of fac-Ir(ppy)₃, the mixture of the CouNO monomer (1 mM) and fac-Ir(ppy)₃ (200 μM) was dissolved in DMSO. Subsequently, the mixture was exposed to irradiation using a 500 nm LED lamp. In this process, a certain DMSO solution of these compounds was diluted with a mixture of acetonitrile and water to test HPLC. The peak areas of the CouNO monomer and products were monitored and recorded during the process.

2.7 Electrochemical measurements

All electrochemical measurements were implemented using a CHI (model 600E) electrochemical analyzer/workstation equipped with a three-electrode setup, consisting of a glassy carbon working electrode, a Pt wire counter electrode, and an Ag/AgCl reference electrode. Data were acquired in anhydrous DMF, by cyclic voltammetry (CV), with a scan rate of 0.1 V s⁻¹, in the presence of a tetrabutylammonium tetrafluoroborate electrolyte (0.1 M). The electrolyte was degassed with argon at 25 °C for 30 min prior to measurement and the argon atmosphere was maintained during the measurement process.

2.8 Fabrication of fac-Ir(ppy)₃-loaded micelles (PNO@Ir)

fac-Ir(ppy)₃ (0.2 mg) and PNO (2 mg) were dissolved in 1 mL of DMSO and the DMSO solution was quickly injected into 8 mL of DI water. The DMF was removed by dialysis (MWCO = 14 kDa). The resultant micellar nanoparticles were used for further experiments.

2.9 In vitro antibacterial assay

For methicillin-resistant Staphylococcus aureus (MRSA, USA300 LAC) bacterial strain, 3–5 individual colonies were inoculated into fresh Luria–Bertani (LB) media and incubated at 37 °C for 16–18 h. Then, 40 μL of culture medium was diluted with fresh LB (100-fold) and regrown at 37 °C to a mid-log phase (OD₆₀₀ = 0.5–0.7). Bacterial cells were then harvested and washed twice with sterile PBS via centrifugation (10 000 rpm for 5 min, 4 °C), adjusted with sterile PBS to 1.5 × 10⁶ cfu per mL, and inoculated into zero-dilution wells of a preset 96-well microplate. Varying concentrations of micellar aqueous dispersions (100 μL) of PNMe@Ir, PNO@Ir and NP@Ir micelles were added into each zero-dilution well in a 96-well microplate. 50 μL of the adjusted bacterial suspension was inoculated into each zero-dilution well of a preset microplate, to achieve 5 × 10⁵ cfu per mL in each well (150 μL). The microplate was then incubated at 37 °C for 10 min, followed by illumination for 30 min under 500 nm (30 mW cm⁻²). After this, the treated bacteria were further incubated at 37 °C for 1 h and diluted with sterile PBS buffer (100×), followed by plating the dilutions (20 μL) onto LB agar plates for overnight incubation at 37 °C to form visible colonies.

2.10 N-Phenylnapthylamine (NPN) assay

In the NPN uptake assay, mid-log-phase MRSA bacteria were diluted to a working suspension (OD₆₀₀ = 0.6) with HEPES buffer (5 mM HEPES, pH 7.4, containing 5 mM glucose) and then treated with PBS, 1% Triton X-100, PNO@Ir and PNMe@Ir micelles (0.2 g L⁻¹) without or with 500 nm irradiation for 30 min (30 mW cm⁻²). After incubation for 30 min, 200 μL of the cell suspension and 10 μL of the NPN probe (0.4 mM, dissolved in acetone) were added into a polystyrene 96-well plate. After this, the fluorescence emission at 420 nm of the bacterial suspension was read using a microplate reader (Thermo Fisher) with an excitation wavelength of 350 nm. The NPN uptake was calculated using the following formula:

NPN uptake % = (F − F₀)/(F_c − F₀) × 100%

where F is the fluorescence intensity in the presence of micellar nanoparticles, F₀ is the fluorescence intensity in the absence of micellar particles with MRSA, and F_c is the fluorescence intensity of MRSA in the presence of the NPN probe and 1% Triton X-100.

2.11 Protein leakage assay

For the protein leakage assays of bacteria, the mixtures of bacterial suspension (OD₆₀₀ = 0.6) with PBS, 1% Triton X-100, PNO@Ir and PNMe@Ir micelles (0.2 g L⁻¹) without or with 500 nm irradiation for 30 min (30 mW cm⁻²), PBS and 1% Triton X-100 were used as the negative and positive controls, respectively. The bacterial suspension was incubated at 37 °C for 30 min and then centrifuged (10 min, 10 000 rpm), and the supernatant was used to measure protein contents by an enhanced BCA protein assay kit. The absorbance intensity at 562 nm was recorded and the protein concentrations were calculated against a standard calibration curve using bovine serum albumin (BSA) as a model protein.

2.12 General procedures for bacterial biofilm formation and harvesting

MRSA and Pseudomonas aeruginosa (PAO1, ATCC 27853) were used for different experiments. LB medium (300 μL) and MRSA/PAO1 suspension (10⁸ cfu per mL, 100 μL) were added into each well in a 24-well plate. The 24-well plate was placed in an incubator and cultured at 37 °C. After 24 h incubation, the timeworn medium was replaced with fresh LB medium, and the biofilms were cultured for another 24 h, which were used for further antibiofilm experiments.

2.13 Quantification of biofilm biomass by crystal violet staining

The cultured MRSA/PAO1 biofilms were treated with PBS, PNO@Ir and PNMe@Ir micelles (0.2 g L⁻¹) under the hypoxia environment, respectively. The biofilms were incubated at 37 °C for 4 h in an AnaeroPack™ (Mitsubishi Gas Chemical Company, Inc.). After this, the treated biofilms were irradiated or not irradiated with 500 nm (30 mW cm⁻²) for 30 min. The biofilms were incubated at 37 °C for 30 min and were washed three times with PBS, followed by the addition of 0.5 mL of the crystal violet staining agent (0.1 wt% in PBS). Afterward, the plate was incubated for 20 min before the wells were washed with PBS two times. The crystal violet was then dissolved in pure ethanol (500 μL) and was quantified by measuring the OD₅₅₀ of the homogenized suspension using a microtiter plate reader (Thermo Fisher). All the reported results were repeated in at least three independent experiments.

2.14 Evaluation of the bacterial viability by the colony forming unit (CFU) assay

The bacterial viability in biofilms after various treatments was also evaluated by the standard CFU assay. The MRSA/PAO1 biofilms were first cultured according to the above method. Briefly, after receiving various treatments, the biofilms were washed three times with PBS, and the viable MRSA/PAO1 cells in biofilms were detached into 0.5 mL of sterile PBS via ultrasound (200 W, 40 kHz) for 30 min. The harvested cell suspension was serially diluted, plated onto LB agar, and incubated at 37 °C for 18 h. The bacterial colonies were then counted. All assays included three replicates and were repeated in three independent experiments.

2.15 Live/dead staining of biofilms observed by confocal laser scanning microscopy (CLSM)

For CLSM analysis, MRSA biofilms were grown in a glass-bottom 24-well plate as described above. After various treatments, bacterial biofilms were washed twice with PBS before staining with LIVE/DEAD® BacLight™ bacterial viability kit reagents (L-13152, Molecular Probes) according to the manufacturer's instructions. Bacterial biofilms were incubated at room temperature for 20 min under dark conditions, followed by observation by CLSM. The green channel of SYTO 9 was excited at 488 nm and was collected between 500 and 545 nm; the red channel of PI was excited at 514 nm and collected between 600 and 650 nm.

2.16 Variations of bacterial morphology observed by scanning electron microscopy (SEM)

To investigate the antibiofilm mechanism, MRSA biofilms were grown on sterile cover glass slides in 24-well plates as described above. Briefly, the treated biofilms were washed three times with PBS and fixed with paraformaldehyde (2.5 wt%) at 4 °C overnight. The fixed biofilm was serially dehydrated with graded ethanol solution (30%, 50%, 70%, 80%, 90%, 95%, and 100%) and was dried for SEM observation.

2.17 MRSA-infected mouse wound model

All the animal studies were approved by the Committee on the Ethics of Animal Experiments of the University of Science and Technology of China (USTC) and were performed in strict accordance with the Animal Care and Use Committee of USTC (Approval no.: USTCACUC25120124095). First, dorsal hair of 6–8 weeks-old Balb/c male mice (purchased from the Experimental Animal Center of Anhui Medical University) were shaved. The mouse wound model was established by first excising a wound and then applying 20 μL of MRSA suspension (2 × 10⁷ cfu per mL) to the wound site. The images on −2 day were immediately taken just before bacterial inoculation. After 48 h, the biofilm infected wound was formed.

2.18 In vivo antibacterial evaluation in the mouse wound infection model

The wound infection was created by applying MRSA suspension. After 48 h, PBS (25 μL), free vancomycin (Van, 25 μL, 19 μg mL⁻¹), and aqueous dispersions (25 μL, 0.2 g L⁻¹) of PNO@Ir and PNMe@Ir micelles were applied to the biofilm infected sites, respectively. After 30 min, the infected sites of mice were treated with or without 500 nm irradiation (30 mW cm⁻²). On days 2 and 6, the infected tissues were excised and homogenized for 30 min in PBS (1 mL). The homogenate was serially diluted, plated on LB agar (60 μL), and cultured at 37 °C for 14 h. After this, the bacterial colonies on the plates were imaged and counted, and the CFU assay results are shown in Fig. 4d. On day 6, the infected tissues were excised from the mice and were fixed in 4% formaldehyde at 4 °C overnight, embedded in paraffin, and sectioned for further histological evaluation. Moreover, the changes of the infected areas and the body weights were continuously monitored and recorded through photographs.

2.19 Histology analysis by hematoxylin and eosin (H&E), Masson staining and TNF-α staining

Neutral buffered formalin (10%) was used to fix the infected sections and surrounding tissues for 24 h. The fixed tissues were embedded in paraffin and then sectioned at a 4 μm thickness using a Leica microtome (RM2016). Infected sections were stained with hematoxylin and eosin (H&E), Masson and TNF-α staining during treatment. All histological analyses were performed on at least three independent groups, and the images displayed are representatives of all replicates. Images of entire sections were recorded using an IX71 microscope (Olympus).

2.20 Cytotoxicity assay

HUVECs were cultured in DMEM supplemented with 10% fetal bovine serum (FBS) and antibiotics (100 units per mL penicillin and 0.1 g L⁻¹ streptomycin). The cell culture was maintained at 37 °C in a humidified atmosphere containing 5% CO₂. To determine cell viability, HUVECs were seeded onto a 96-well plate at a density of 10 000 cells per well, with each well containing 100 μL of DMEM. After 24 h of incubation, the DMEM was replaced with 100 μL of fresh medium containing the desired micellar nanoparticles. Following an additional 24 h incubation period, 10 μL of MTT reagent in PBS buffer (5 g L⁻¹) was added to each well. The cells were then incubated for another 4 h. Subsequently, the culture medium in each well was removed and replaced by 100 μL of DMSO. The absorbance values were recorded at a wavelength of 490 nm using a microplate reader (Thermo Fisher Scientific). The cell viability was calculated using the following equation:
where A_{490nm,treated} and A_49nm,control are the absorbance values in the presence and absence of micelles. A_490nm,blank is the absorbance value of the plate with an identical volume of MTT solution without cells, respectively.

2.21 Statistical analysis

Data are presented as mean ± standard deviations and were analyzed using Prism 8.0 software (GraphPad, San Diego, California, USA) and student's t-test. p values lower than 0.05 were considered to be statistically significant.

3. Results and discussion

3.1 500 nm-Light-triggered NO release from the CouNO monomer in the presence of fac-Ir(ppy)₃

N-Nitrosamine, a type of NO donor, has shown considerable potential as a photoresponsive NO-releasing agent. Based on our previous studies, it can be activated by light to trigger controlled NO release.^4,15,16,49 Given that coumarin-based NO donors possess self-reporting capabilities, and with the aim of synthesizing polymers suitable for biomedical applications, we have designed and synthesized a coumarin-based NO-releasing monomer, referred to as the CouNO monomer. The CouNO monomer exhibited a maximal absorbance at 330 nm, with negligible absorption above 400 nm, indicating its lack of direct responsiveness to green light irradiation (e.g., 500 nm) (Fig. 1a). Building upon our previous findings, we found that a mixture of the CouNO monomer (50 μM) and fac-Ir(ppy)₃ (10 μM) displayed a simultaneous decrease in absorbance at 330 nm and an increase at 370 nm upon exposure to 500 nm light (30 mW cm⁻²), as shown in Fig. 1b. This irradiation condition was consistently applied throughout the study. By contrast, absorbance measurements revealed no significant changes in control groups (non-irradiated) and irradiated by the photosensitizer alone (Fig. S3, ESI†). Notably, the absorbance profile of the CouNO monomer/fac-Ir(ppy)₃ mixture after 500 nm irradiation exhibited close alignment with that of CouNH monomer derivatives, providing spectroscopic evidence for light-triggered NO release (Fig. S2 and S4, ESI†). Additionally, electron paramagnetic resonance (EPR) analysis with PTIO as the spin trap confirmed light-dependent NO radical generation, with no detectable spontaneous release under dark conditions (Fig. 1c), corroborating UV/Vis absorbance spectroscopy observations. Real-time monitoring of NO release remains technically challenging owing to its inherent high reactivity and short half-life. However, spectrophotometric tracking of the stoichiometric conversion from the CouNO monomer to CouNH monomer derivatives enabled quantitative NO release quantification via absorbance changes. Calculated using the equation, the NO release reached equilibrium at approximately 50 min, with a cumulative concentration of ∼30 μM (Fig. 1d). Moreover, the fac-Ir(ppy)₃-mediated NO release exhibited concentration-dependent kinetics, and an increased fac-Ir(ppy)₃ concentration led to a faster NO-releasing rate (Fig. S2, ESI†).

Fig. 1 (a) UV/Vis absorbance spectra of fac-Ir(ppy)₃(10 μM) and the CouNO monomer (50 μM) in DMSO. (b) Evolution of UV/Vis absorbance spectra of DMSO solution of the CouNO monomer (50 μM) and fac-Ir(ppy)₃ (10 μM) under 500 nm light irradiation (30 mW cm⁻²). (c) EPR spectra of the CouNO monomer in the presence of fac-Ir(ppy)₃ (10 μM) without (blue curve) and with (red curve) 500 nm light irradiation for 10 min. In all cases, the concentrations of PTIO and CouNO monomer were 20 and 50 μM, respectively. (d) NO release profiles of DMSO solution of the CouNO monomer (50 μM) and fac-Ir(ppy)₃ (10 μM) mixture with or without 500 nm irradiation (30 mW cm⁻²).

3.2 Photolytic reaction of the CouNO monomer in the presence of fac-Ir(ppy)₃

Based on our previous work, we hypothesized that the CouNO monomer undergoes photolysis in the presence of fac-Ir(ppy)₃ under light irradiation, enabling controlled NO release. Moreover, building upon the excellent mechanistic insights from Han et al.,^50,51 with additional evidence from Wang et al.^52,53 on drug release induced by long-wavelength activation of short-wavelength-prodrugs, we proposed that the photolabile linkage within the CouNO monomer undergoes different cleavages under identical irradiation conditions. To our delight, the photodegradation process of the CouNO monomer was characterized by HPLC analysis (Fig. 2a and b). Within 20 min, the CouNO monomer was completely degraded, largely converted into the CouNH monomer, with small amounts of CouNO and CouNH also generated. From 20 to 60 min, the CouNH monomer gradually degraded under light irradiation to form CouNH, consistent with the results from direct irradiation of the CouNH monomer (Fig. S5, ESI†). Based on the changes in the peak area, the CouNO monomer was found to undergo nearly complete degradation within 30 min (Fig. 2c). To investigate the photodegradation mechanism of the CouNO monomer in the presence of fac-Ir(ppy)₃, we examined the photoluminescence of fac-Ir(ppy)₃ with different concentrations of the CouNO monomer. The results clearly show that the phosphorescence intensity of fac-Ir(ppy)₃ decreases significantly as the CouNO monomer concentration increases, indicating that the excited state of fac-Ir(ppy)₃ is effectively quenched by the CouNO monomer, likely through either an energy or an electron transfer mechanism. The Stern–Volmer constant (k_sv) and the quenching rate constant (k_q) were determined to be 2.01 × 10³ M⁻¹ and 1.58 × 10⁹ M⁻¹ s⁻¹, respectively (Fig. 2d and Fig. S6, ESI†). Similarly, the k_sv and k_q values of CouNO were also determined to be 1.91 × 10³ M⁻¹ and 1.50 × 10⁹ M⁻¹ s⁻¹ (Fig. S7, ESI†). Meanwhile, the CV results indicated that the excited-state reduction potential of fac-Ir(ppy)₃ was −1.39 V. The reduction potentials of the CouNO monomer and CouNO were determined to be −1.18 V (Fig. S8, ESI†) and −1.21 V,¹⁵ respectively. These values suggest that fac-Ir(ppy)₃ in its excited state is thermodynamically capable of reducing both the CouNO monomer and CouNO.

Fig. 2 (a) The photolytic reaction of fac-Ir(ppy)₃ and the CouNO monomer through two mechanisms (TTET/PeT). (b) HPLC trace recorded for sequential degradation of the compound CouNO monomer under the light irradiation (500 nm, 30 mW cm⁻²) with fac-Ir(ppy)₃. (c) The peak area change of the CouNO monomer was analyzed by HPLC in Fig. 2b. (d) Stern–Volmer plot of phosphorescence intensity quenching of fac-Ir(ppy)₃ (10 μM) by the CouNO monomer in Ar-saturated DMF. (e) The relative energetic dispositions for the frontier orbitals (HOMO and LUMO) of fac-Ir(ppy)₃, CouNO monomer and CouNO. (f) The singlet (S₁) and triplet (T₁) energy levels of fac-Ir(ppy)₃ and T₁ energy levels of the CouNO monomer, which reveal the rationale for the occurrence of TTET.

Additionally, we also attempted to explain the photodegradation of the CouNO monomer from a theoretical perspective. Through time-dependent density functional theory (TDDFT) at the level of B3LYP/6-31+G(d, p), we found that fac-Ir(ppy)₃ possesses a LUMO energy level of −1.70 eV, which is higher than those of the CouNO monomer (−2.47 eV) and CouNO (−2.44 eV) (Fig. 2e). This energy difference provides a thermodynamic driving force for electron transfer (ET) from PdTPTBP to CouNO upon excitation. These findings support the hypothesis that NO release from the CouNO monomer and CouNO is triggered via a PeT process. Interestingly, the triplet energy (T₁) of fac-Ir(ppy)₃ was determined to be 2.4 eV based on its phosphorescence emission spectrum. Meanwhile, the T₁ energies of the CouNO monomer and CouNO were found to be 2.24 eV and 2.23 eV, respectively (Fig. 2f). These results suggest that NO release from both molecules could also proceed via a TTET mechanism, indicating that the system operates through a dual-mechanism pathway for NO release. Furthermore, based on the work of Han et al.,⁵⁰ in combination with our computational and experimental results, we propose that the CouNO monomer may undergo carbamate bond cleavage via a TTET mechanism, thereby exposing the amine group. However, our calculations revealed that the T₁ energy level of the CouNH monomer is slightly higher than that of fac-Ir(ppy)₃, though the values are close (Fig. S9 and Table S1, ESI†). This energy mismatch makes TTET a less favourable mechanism in this case. Nonetheless, HPLC analysis showed that the CouNH monomer alone can undergo carbamate bond cleavage upon 500 nm light irradiation in the presence of fac-Ir(ppy)₃. Therefore, we tentatively attribute this process to a TTET-mediated pathway. In summary, we propose a possible degradation mechanism of the CouNO monomer in the presence of fac-Ir(ppy)₃, primarily involving TTET- and PeT-mediated NO release and carbamate bond cleavage (Fig. 2a). Further studies will be conducted to elucidate the detailed reaction pathways.

3.3 500 nm-Light-triggered NO release and linkage cleavage from micellar nanoparticles

After confirming that the CouNO monomer can release NO and achieve carbamate linkage cleavage under 500 nm light irradiation, we decided to enhance its water solubility by using RAFT polymerization to copolymerize the NO-releasing moiety with a hydrophilic monomer (poly(ethylene glycol)methyl ether methacrylate, OEGMA) and a hydrophobic monomer (butyl methacrylate, BMA) to form the copolymer PNO (Scheme S2, ESI†), which was characterized by ¹H NMR spectroscopy (Fig S10a, ESI†) and gel permeation chromatography (GPC) (Fig S11, ESI†). As a control, a polymer designated as PNMe was synthesized via RAFT polymerization (Scheme S2 and Fig. S10, ESI†). PNO and PNMe were subsequently used to form micelles, into which the photosensitizer was encapsulated. The resulting micelles were referred to as PNO@Ir (Scheme 1b) and PNMe@Ir.

Given that the biofilm microenvironment is typically weakly acidic and hypoxic,⁵ we evaluated the behavior of PNO@Ir micelles under weakly acidic conditions and in the presence of the SA. Transmission electron microscopy (TEM) and dynamic light scattering (DLS) measurements revealed that the micelle size decreased after light irradiation (Fig. 3a–c and Fig. S12, ESI†), regardless of whether the environment was neutral or weakly acidic. This phenomenon is presumed to result from the photogeneration of NO by PNO@Ir micelles under hypoxic conditions, which subsequently triggers the cleavage of carbamate bonds. This bond cleavage exposes amine groups, converting previously hydrophobic segments in the micelle core into hydrophilic ones, thereby leading to micelle dissociation. Under weakly acidic conditions, the dissociated polymer chains may form protonated amine groups, resulting in the generation of smaller nanoparticles in aqueous solution. Additionally, zeta potential measurements showed a shift toward positive values after light irradiation under acidic conditions (Fig. 3d), which supports the hypothesis of amine protonation. Importantly, UV/Vis absorbance spectroscopy confirmed that PNO@Ir micelles can release approximately 17 μM of NO under the same conditions (Fig. 3e and f). In addition, fluorescence measurements revealed that the fluorescence intensity of the coumarin moiety within the nanoparticle gradually increased with prolonged light irradiation, corresponding to the release of NO. This indicates that the nanoparticle is capable of self-reporting the reaction progress without the need for external fluorescent probes (Fig. S13, ESI†). Furthermore, under light irradiation in the presence of SA, the dissolved oxygen concentration dropped from approximately 8 mg L⁻¹ to 0.02 mg L⁻¹, while little to no change was observed without irradiation (Fig. S14, ESI†). These findings indicate that the micelles are capable of releasing NO in a hypoxic environment upon light activation.

Fig. 3 (a) and (b) TEM images of PNO@Ir micelles with or without 500 nm irradiation for 30 min in the presence of SA (10 mM) at pH 5.5, respectively. (c) Intensity-average hydrodynamic diameter distributions of PNO@Ir micelles with or without 500 nm irradiation for 30 min at pH 5.5, respectively. In all cases, the irradiation intensity was 30 mW cm⁻² under hypoxic conditions. (d) ζ potentials of aqueous dispersions (0.2 g L⁻¹) of PNO@Ir micellar nanoparticles at pH 7.4 or pH 5.5 with or without 500 nm light irradiation for 30 min (30 mW cm⁻²). (e) Evolution of UV/Vis absorbance spectra of PNO@Ir micelles (0.2 g L⁻¹) in the presence of SA (10 mM) under 500 nm irradiation. (f) NO release profiles from PNO@Ir micelles in the presence of SA (10 mM) with or without 500 nm irradiation for 30 min. In all cases, the irradiation intensity was 30 mW cm⁻².

3.4 500 nm-Light-triggered NO release and amino cation generation synergistically combat biofilms

Since we confirmed that the micelles PNO@Ir could release NO under green light irradiation and generate amino cations under mildly acidic and hypoxia conditions, we sought to evaluate their antibacterial and antibiofilm activity. We first investigated the antibacterial efficacy of PNO@Ir micelles against MRSA across varying concentrations. The results revealed that PNO@Ir micelles eradicated 80% of the bacteria at 0.2 g L⁻¹, while a significant enhancement in bactericidal activity was observed at 0.3 g L⁻¹, achieving 99% elimination (Fig. S15, ESI†). These findings underscore the promising antimicrobial potential of PNO@Ir micelles, whose unique mechanism of synergistic NO release and amino cations generation establishes a theoretical groundwork for subsequent anti-biofilm applications. It is well established that approximately 80% of chronic and recurrent microbial infections originate from bacterial biofilms.⁵⁴ Given that MRSA and PAO1 serve as paradigm representatives of Gram-positive and Gram-negative bacteria, respectively, both extensively validated for standardized antibiofilm assessments. Therefore, we selected the MRSA/PAO1 biofilm as our experimental model system.

Antibiofilm assays revealed that under 500 nm light irradiation (30 min), only PNO@Ir micelles significantly reduced biomass in both MRSA and PAO1 biofilms by crystal violet staining (Fig. 4a and Fig. S16a, ESI†). Additionally, CFU assays further confirmed the PNO@Ir's antibiofilm activity, achieving 70% and 28% reductions in viable bacteria for MRSA and PAO1 biofilms, respectively (Fig. 4b and Fig. S16b, ESI†). These findings demonstrate the PNO@Ir's better antibiofilm efficacy against MRSA compared to PAO1. Meanwhile, MRSA is a widespread pathogen responsible for a broad range of infections, from superficial skin conditions to severe diseases such as osteoarticular infections and endocarditis, leading to high morbidity and mortality;⁵⁵ we therefore focused on its antibiofilm activity. More importantly, the 500 nm-light-triggered release of NO and the generation of amino cations not only facilitated biofilm dispersion but also killed bacterial pathogens, as demonstrated by live/dead BacLight bacterial viability staining (Fig. 4c). Among all samples, PNO@Ir micelles exhibited antibiofilm activity, as indicated by a marked decrease in green fluorescence and an increase in yellow and red fluorescence (dead bacteria). Furthermore, the synergistic antibiofilm effect of NO and amino cations was further validated by SEM, which revealed substantial biofilm disruption and bacterial membrane damage upon 500 nm light irradiation (Fig. 4d). To elucidate the synergistic antibiofilm mechanism between NO and amino cations, we measured membrane permeability and protein leakage in MRSA after treatment under different conditions (Fig. S17, ESI†). PNMe@Ir micelles caused a modest increase in membrane permeability and protein release, whereas PNO@Ir micelles produced significantly greater effects, confirming the enhanced membrane disruption resulting from the combined NO release and amino-cationic activity. These findings align with previous reports demonstrating that amino-cationic polymers damage bacterial membranes and thereby kill bacteria.^56–58 Building on the aforementioned experimental evidence and the supporting literature, we propose the following mechanism for NO and cation synergy against biofilms: NO triggers biofilm dispersal via phosphodiesterase-mediated c-di-GMP depletion, converting sessile bacteria to planktonic states.^59,60 This cascade upregulates motility genes while suppressing adhesins, weakening biofilm matrices and sensitizing embedded bacteria.⁶⁰ Concurrently, light-activated cationic moieties electrostatically disrupt bacterial membranes, causing permeabilization and lysis. The synergy of NO-mediated dispersal and cationic membrane disruption eliminates biofilms significantly more effectively than either monotherapy.^24,58,61 Consistent with this dual mechanism, NO-releasing cationic micelles (PNO@Ir) achieve deeper biofilm eradication than NO-inert controls (PNMe@Ir) (Fig. 4c and d).

Fig. 4 (a) Bacterial biomass of MRSA biofilms by crystal violet staining and (b) corresponding bacterial viability after treatment with PBS, PNO@Ir and PNMe@Ir micelles without or with 500 nm light irradiation for 30 min, respectively. Data are shown as the mean ± s.d. (n = 3); **p < 0.01, ***p < 0.001, n.s., not significant. (c) 3D CLSM images of MRSA biofilms stained with the LIVE/DEAD BacLight bacterial viability kit, which were treated with PBS, PNO@Ir, PNMe@Ir micelles and Van without or with 500 nm light irradiation for 30 min (30 mW cm⁻²), respectively. The green and red channels were excited with a 488 nm (SYTO 9) and 514 nm (PI) and were collected at 500–545 nm (green) and 600–650 nm (red), respectively. (d) SEM images of MRSA biofilms treated with PBS, PNO@Ir, PNMe@Ir micelles and Van without or with 500 nm light irradiation for 30 min (30 mW cm⁻²), respectively. The arrows indicate dead bacteria with disrupted membrane integrity. In all cases, the Van and micelle concentrations were 19 μg mL⁻¹ and 0.2 g L⁻¹, respectively.

3.5 Treatment of MRSA infection in a wound infection model

After confirming the antibacterial and antibiofilm activities of PNO@Ir micelles in vitro, we further explored their potential for treating biofilm-associated infections in vivo. Given the prevalence of wound infections, this model holds significant promise for evaluating novel therapeutic agents. Notably, several NO-releasing nanoparticles have already demonstrated promising antibiofilm efficacy in similar infection models. To establish a wound infection model, an incision was made on the dorsal surface of mice, followed by the topical application of the MRSA suspension onto the wound. After 48 h of infection, visible skin necrosis developed at the wound site. The mice were then randomly divided into five groups, receiving different treatments: PNMe@Ir and PNO@Ir micelles with 500 nm light irradiation for 30 min, and PBS, PNO@Ir micelles and Van without light irradiation. Throughout the treatment period, various assessments were conducted at designated time points, including wound photography, body weight monitoring, histological analysis, and bacterial counting (CFU assays), to evaluate therapeutic efficacy (Fig. 5a). Wound photography and measurements of the relative wound size demonstrated that PNO@Ir micelles under 500 nm light irradiation significantly improved the healing of biofilm-infected wounds compared to the other four groups, indicating enhanced therapeutic efficacy and accelerated wound closure (Fig. 5b and c), which show that NO promotes wound healing.^59,62 In parallel, bacterial burden at the wound site was quantitatively assessed through CFU assays, which confirmed that PNO@Ir micelles under light exposure exhibited bactericidal activity (Fig. 5d and Fig. S18, ESI†), attributed to the synergistic effects of NO release and amino cation generation. Histological analyses further demonstrated that 500 nm-light-triggered NO release from PNO@Ir micelles reduced inflammatory cell infiltration, promoted collagen deposition, and facilitated re-epithelialization at the wound sites (Fig. 5e). Moreover, histological analysis revealed a marked decrease in the expression of pro-inflammatory cytokines, such as TNF-α, underscoring the anti-inflammatory effects of the treatment (Fig. 5e). These results align well with the established physiological roles of NO in regulating inflammation and promoting tissue regeneration during the wound healing process. Moreover, histopathological evaluation of major organs (heart, liver, spleen, lungs, and kidneys) and body weight monitoring revealed no adverse effects, confirming the biocompatibility of the NO-releasing micelles in infected mice (Fig. 5f and Fig. S19, ESI†). These experimental data represented a proof-of-concept experiment conducted under deliberately conservative conditions (low polymer dose). To obtain a more comprehensive and accurate assessment of antibiofilm performance, we tested PNO@Ir micelles at a higher concentration (0.3 g L⁻¹) in an in vivo infected-wound model. At this dose, the system produced a pronounced improvement in bactericidal activity (Fig. S20, ESI†). Because the overall activity is likely governed by the concentrations of the NO-releasing moiety and hydrophobic segment within the polymer, as well as the loading concentration of fac-Ir(ppy)₃ within the micelles, the present manuscript should be regarded as a proof-of-concept study. Importantly, cytotoxicity assays demonstrated that PNO@Ir micelles exhibited negligible toxicity toward normal cells, such as human umbilical vein endothelial cells (HUVECs), underscoring their biosafety (Fig. S21, ESI†). Collectively, these results demonstrate that the micellar system, capable of simultaneously releasing NO and generating amino cations, exhibits significant therapeutic efficacy against biofilm-associated wound infections through a dual antibacterial mechanism.

Fig. 5 (a) Experimental timeline of the in vivo treatment of bacterial infection in a wound infection model. (b) Representative images of the wound during the treatment process and (c) quantitative analysis of the infected areas receiving different treatments. Data are shown as mean ± s.d. (n = 3); **p < 0.01 compared with the PBS group. (d) Bacterial colony-forming unit separated from wound tissues with varying treatments. Data are shown as mean ± s.d. (n = 3); **p < 0.01, compared with the group receiving PNO@Ir micelles (+hv) treatment on day 2 and day 6. (e) Histological analysis on day 6 of the mice infected with MRSA receiving varying treatments. The scale bar is 50 μm. (f) Changes of body weights of MRSA biofilm-infected mice after different treatments. In all cases, the Van and micelle concentrations were 19 μg mL⁻¹ and 0.2 g L⁻¹, respectively.

4. Conclusions

In summary, we have developed a green light-responsive polymeric nanoparticle system capable of releasing NO and activating antibacterial polymers simultaneously via TTET and PeT mechanisms. This dual-activation process is specifically triggered by the hypoxic and mildly acidic microenvironment characteristic of bacterial biofilms, enabling a biofilm-adaptive, synergistic antibacterial strategy. The resulting NO and in situ generated cationic polymer collaboratively disrupt biofilm integrity and eradicate MRSA with higher efficiency. Both in vitro and in vivo studies demonstrated anti-biofilm activity and accelerated wound healing, with minimal cytotoxicity toward normal cells. This work presents a promising approach for designing next-generation anti-biofilm therapeutics and underscores the broader potential of light-activated nanomedicine.

Author contributions

Siyuan Luo: conceptualization, formal analysis, methodology, writing – original draft, and writing – review and editing. Zuotao Zhou: methodology and investigation. Yu Jin: formal analysis and writing review and editing. Haochuan Ding: investigation and methodology. Faxing Jiang: investigation, formal analysis, and conceptualization. Zhiqiang Shen: conceptualization, formal analysis, writing – review and editing, and project administration.

Conflicts of interest

The authors declare no competing financial interest.

Data availability

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

Acknowledgements

The authors gratefully acknowledge the National Key R&D Program (2024YFA1509203) and the National Natural Science Foundation of China (52303213). The image is adapted from biorender.com, with permission.

References

1. H. Koo, R. N. Allan, R. P. Howlin, P. Stoodley and L. Hall-Stoodley, Nat. Rev. Microbiol., 2017, 15, 740–755.
2. P. S. Stewart, Antimicrob. Agents Chemother., 1996, 40, 2517–2522.
3. Y. Liu, H. J. Busscher, B. R. Zhao, Y. F. Li, Z. K. Zhang, H. C. van der Mei, Y. J. Ren and L. Q. Shi, ACS Nano, 2016, 10, 4779–4789.
4. J. Cheng, G. H. Gan, S. Q. Zheng, G. Y. Zhang, C. Zhu, S. Y. Liu and J. M. Hu, Nat. Commun., 2023, 14, 7510.
5. S. B. Behbahani, S. D. Kiridena, U. N. Wijayaratna, C. Taylor, J. N. Anker and T. R. J. Tzeng, Front. Microbiol., 2022, 13, 1028560.
6. C. G. Kumar and S. K. Anand, Int. J. Food Microbiol., 1998, 42, 9–27.
7. R. M. Donlan and J. W. Costerton, Clin. Microbiol. Rev., 2002, 15, 167–193.
8. Y. F. Gao, J. Wang, M. Y. Chai, X. Li, Y. Y. Deng, Q. Jin and J. Ji, ACS Nano, 2020, 14, 5686–5699.
9. Q. C. Truong-Bolduc, G. R. Bolduc, R. Okumura, B. Celino, J. Bevis, C. H. Liao and D. C. Hooper, Antimicrob. Agents Chemother., 2011, 55, 3214–3219.
10. R. Hengge-Aronis, Microbiol. Mol. Biol. Rev., 2002, 66, 373–395.
11. J. D. Chambless, S. M. Hunt and P. S. Stewart, Appl. Environ. Microbiol., 2006, 72, 2005–2013.
12. K. Poole, J. Antimicrob. Chemother., 2012, 67, 2069–2089.
13. B. L. Miao, D. H. Wang, L. Yu, X. F. Meng, S. Y. Liu, M. Q. Gao, J. T. Han, Z. L. Chen, P. Li and S. W. Liu, Microbiol. Res., 2025, 292, 127987.
14. Y. D. Pang, H. Y. Wang, Y. Yao, D. Y. Chen, R. Yang, Z. Y. Wang, J. H. Yang, Y. M. Li and W. G. Liu, Adv. Funct. Mater., 2023, 33, 2303095.
15. Z. Q. Shen, S. Q. Zheng, S. Y. Xiao, R. Shen, S. Y. Liu and J. M. Hu, Angew. Chem., Int. Ed., 2021, 60, 20452–20460.
16. Z. H. Chen, S. Q. Zheng, Z. Q. Shen, J. Cheng, S. Y. Xiao, G. Y. Zhang, S. Y. Liu and J. M. Hu, Angew. Chem., Int. Ed., 2022, 61, e202204526.
17. Q. Zhou, T. J. Wang, K. P. Li, S. Y. Zhang, K. Wang, W. L. Hong, R. J. Liu and P. Li, J. Controlled Release, 2023, 357, 561–571.
18. G. F. Li, Z. Y. Feng, Z. S. Hou, R. Z. Chen, H. N. Cui, T. J. Wang and P. Li, Adv. Funct. Mater., 2025, 35, 2418229.
19. Z. Q. Shen, S. Q. Zheng, Y. M. Fang, G. Y. Zhang, C. Zhu, S. Y. Liu and J. M. Hu, Angew. Chem., Int. Ed., 2023, 62, e202219153.
20. J. Y. Xie, M. Zhou, Y. X. Qian, Z. H. Cong, S. Chen, W. J. Zhang, W. N. Jiang, C. Z. Dai, N. Shao, Z. M. Ji, J. C. Zou, X. M. Xiao, L. Q. Liu, M. Z. Chen, J. Li and R. H. Liu, Nat. Commun., 2021, 12, 5898.
21. Y. X. Qian, S. Deng, Z. H. Cong, H. D. Zhang, Z. Y. Lu, N. Shao, S. A. Bhatti, C. Zhou, J. G. Cheng, S. H. Gellman and R. H. Liu, J. Am. Chem. Soc., 2022, 144, 1690–1699.
22. S. M. Andrabi, S. M. S. Shahriar, Y. R. Tian, F. Hayati, Y. J. Su, F. Zhang, N. S. Sharma, G. S. Wang, S. L. Wong, Z. Yan, G. L. Huang and J. W. Xie, Adv. Funct. Mater., 2025, 2422150.
23. T. K. Nguyen, S. J. Lam, K. K. K. Ho, N. Kumar, G. G. Qiao, S. Egan, C. Boyer and E. H. H. Wong, ACS Infect. Dis., 2017, 3, 237–248.
24. R. Namivandi-Zangeneh, Z. Sadrearhami, A. Bagheri, M. Sauvage-Nguyen, K. K. K. Ho, N. Kumar, E. H. H. Wong and C. Boyer, ACS Macro Lett., 2018, 7, 592–597.
25. M. H. Chen, J. J. Wei, S. Z. Xie, X. Y. Tao, Z. L. Zhang, P. Ran and X. H. Li, Nanoscale, 2019, 11, 1410–1422.
26. W. Y. Cai, J. F. Wu, C. W. Xi and M. E. Meyerhoff, Biomaterials, 2012, 33, 7933–7944.
27. P. Singha, J. Locklin and H. Handa, Acta Biomater., 2017, 50, 20–40.
28. K. R. Rouillard, D. B. Hill and M. H. Schoenfisch, J. Cystic Fibrosis, 2020, 19, 1004–1010.
29. Y. A. Tian, X. J. Tian, T. Li and W. H. Wang, Crit. Rev. Food Sci. Nutr., 2025, 65, 647–666.
30. L. Gao, X. H. Qiao, L. X. Zhang, Y. Wang, Y. Wan and C. Chen, Nitric Oxide-Biol. Chem., 2022, 128, 50–58.
31. B. J. Privett, A. D. Broadnax, S. J. Bauman, D. A. Riccio and M. H. Schoenfisch, Nitric Oxide-Biol. Chem., 2012, 26, 169–173.
32. C. X. Gong, W. Guan, X. M. Liu, Y. F. Zheng, Z. Y. Li, Y. Zhang, S. L. Zhu, H. Jiang, Z. D. Cui and S. L. Wu, Adv. Mater., 2022, 34, 2206134.
33. H. Palza, B. Barraza and F. Olate-Moya, Annu. Rev. Mater. Res., 2022, 52, 1–24.
34. C. M. Thompson, A. H. Tischler, D. A. Tarnowski, M. J. Mandel and K. L. Visick, Mol. Microbiol., 2019, 111, 187–203.
35. E. M. Hetrick and M. H. Schoenfisch, Annu. Rev. Anal. Chem., 2009, 2, 409–433.
36. S. Y. Tao, Z. Q. Shen, J. Chen, Z. Shan, B. Huang, X. Y. Zhang, L. Zheng, J. H. Liu, T. You, F. D. Zhao and J. M. Hu, ACS Nano, 2022, 16, 20376–20388.
37. N. Ieda, Y. Hotta, N. Miyata, K. Kimura and H. Nakagawa, J. Am. Chem. Soc., 2014, 136, 7085–7091.
38. A. A. Polyansky, R. Ramaswamy, P. E. Volynsky, I. F. Sbalzarini, S. J. Marrink and R. G. Efremov, J. Phys. Chem. Lett., 2010, 1, 3108–3111.
39. L. Yang, T. M. Weiss, R. I. Lehrer and H. W. Huang, Biophys. J., 2000, 79, 2002–2009.
40. P. P. Kalelkar, M. Riddick and A. J. García, Nat. Rev. Mater., 2022, 7, 39–54.
41. C. Ergene, K. Yasuhara and E. F. Palermo, Polym. Chem., 2018, 9, 2407–2427.
42. R. Namivandi-Zangeneh, R. J. Kwan, T. K. Nguyen, J. Yeow, F. L. Byrne, S. H. Oehlers, E. H. H. Wong and C. Boyer, Polym. Chem., 2018, 9, 1735–1744.
43. Y. J. Jiang, X. Yang, R. Zhu, K. Hu, W. W. Lan, F. Wu and L. H. Yang, Macromolecules, 2013, 46, 3959–3964.
44. T. K. Nguyen, R. Selvanayagam, K. K. K. Ho, R. X. Chen, S. K. Kutty, S. A. Rice, N. Kumar, N. Barraud, H. T. T. Duong and C. Boyer, Chem. Sci., 2016, 7, 1016–1027.
45. V. W. L. Ng, X. Y. Ke, A. L. Z. Lee, J. L. Hedrick and Y. Y. Yang, Adv. Mater., 2013, 25, 6730–6736.
46. M. A. Fischbach, Curr. Opin. Microbiol., 2011, 14, 519–523.
47. G. Cottarel and J. Wierzbowski, Trends Biotechnol., 2007, 25, 547–555.
48. W. Jiang, Z. Q. Shen, Z. X. Guo, Q. Wang, Q. Li, J. M. Hu and Y. C. Wang, Nano Today, 2023, 48, 101696.
49. Z. Q. Shen, W. Jiang, S. Q. Zheng, S. Y. Luo, Z. X. Guo, Q. Wang, Y. C. Wang and J. M. Hu, Angew. Chem., Int. Ed., 2025, 64, e202419939.
50. L. Huang, L. Zeng, Y. Z. Chen, N. Yu, L. Wang, K. Huang, Y. Zhao and G. Han, Nat. Commun., 2021, 12, 122.
51. L. Zeng, L. H. Jiang, J. Y. Li, L. Huang, Y. Chen, N. Yu, L. Wang, K. Huang, J. Peng and G. Han, Angew. Chem., Int. Ed., 2023, 62, e202218341.
52. K. Q. Long, W. Lv, Z. H. Wang, Y. M. Zhang, K. Chen, N. Fan, F. Y. Li, Y. C. Zhang and W. P. Wang, Nat. Commun., 2023, 14, 8112.
53. W. Lv, Y. F. Li, F. Y. Li, X. Lan, Y. M. Zhang, L. L. Du, Q. Zhao, D. L. Phillips and W. P. Wang, J. Am. Chem. Soc., 2019, 141, 17482–17486.
54. Q. E. Grayton, H. K. Nguyen, C. A. Broberg, J. Ocampo, S. G. Nagy and M. H. Schoenfisch, ACS Infect. Dis., 2023, 9, 1730–1741.
55. A. Kaushik, H. Kest, M. Sood, B. W. Steussy, C. Thieman and S. Gupta, Pathogens, 2024, 13, 76.
56. L. D. Blackman, Y. Qu, P. Cass and K. E. S. Locock, Chem. Soc. Rev., 2021, 50, 1587–1616.
57. D. F. Hu, Y. Y. Deng, F. Jia, Q. Jin and J. Ji, ACS Nano, 2020, 14, 347–359.
58. M. S. Ganewatta, Z. K. Wang and C. B. Tang, Nat. Rev. Chem., 2021, 5, 753–772.
59. S. M. Andrabi, N. S. Sharma, A. Karan, S. M. S. Shahriar, B. Cordon, B. Ma and J. W. Xie, Adv. Sci., 2023, 10, e2303259.
60. A. W. Carpenter and M. H. Schoenfisch, Chem. Soc. Rev., 2012, 41, 3742–3752.
61. K. Z. Wang, H. H. Li, Y. Hu, J. C. Lin, G. Shu, W. Zhang, H. L. Fu, X. X. Liang, A. C. Cheng, S. B. Zhou and F. E. Xu, Adv. Funct. Mater., 2025, 35, 202418928.
62. M. Wu, Z. H. Lu, K. K. Wu, C. W. Nam, L. Zhang and J. S. Guo, J. Mater. Chem. B, 2021, 9, 7063–7075.

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Footnote

† Electronic supplementary information (ESI) available. See DOI: https://doi.org/10.1039/d5tb01132a

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