Multifunctional ROS-responsive hydrogels alleviate subconjunctival inflammation

Abstract

Persistent subconjunctival inflammation poses a significant risk for progression to vision-threatening complications such as fibrosis. Existing pharmacological interventions are limited by their administration constraints and potential bio-toxicity issues, failing to adequately address clinical requirements. There is an urgent need for effective methods to inhibit subconjunctival inflammation. Here, we prepared a multi-network hydrogel (CPES) with reactive oxygen species (ROS) scavenging properties via a one-step mixing process to suppress early subconjunctival inflammation. Due to its unique crosslinked structure, primarily comprising dynamic borate esters and thioether, CPES exhibits controllable biodegradability and ROS scavenging capacity. Experimental results reveal that CPES hydrogels possess ideal gelation stability, adequate swelling rates, robust self-adaptive and self-repair capabilities, free radical scavenging capacity, good cell compatibility, and inflammation restrain capabilities. In a rat model of subconjunctival injury inflammation, CPES effectively controlled early inflammation. Collectively, these findings establish CPES as a promising therapeutic platform for controlling subconjunctival inflammation.

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1. Introduction

Traumatic subconjunctival injury frequently triggers severe inflammatory reactions, which can progress to subconjunctival fibrosis that further impairs vision. This fibrotic transformation markedly elevates the risk of severe complications, including eyelid-scleral adhesion (adhesion between the eyelid and eyeball), recurrent pterygium, and compromised outcomes following glaucoma filtration surgery.^1–3 Current treatment strategies primarily involving pharmacological intervention and surgical repair are limited by inadequate drug delivery to the injury site, high surgical complexity, and associated infection risks.^4,5 During the inflammatory process, reactive oxygen species (ROS) serve as key pathological mediators. Excessive ROS production induces oxidative stress, leading to macromolecular damage and cellular dysfunction. This oxidative microenvironment continuously activates inflammatory signaling pathways, impedes tissue repair mechanisms, and ultimately drives fibrotic transformation.^6–8

Traditional treatment methods such as eye drops have limitations, including short ocular surface retention times, poor penetration, low bioavailability, and long-term drug toxicity.^9–11 Therefore, there is an urgent need for innovative biomaterial-based strategies to address the critical challenge of alleviating subconjunctival inflammation and fibrosis through non-pharmacological approaches. ROS-scavenging hydrogels are a class of hydrogels characterized by a three-dimensional, crosslinked structure and inherent antioxidant functionality, with excellent biocompatibility and tunable physical properties, creating a conducive environment for subconjunctival implantation to ameliorate subconjunctival inflammation.^12–15 While polymeric materials such as electrospun nanofibrous membranes have been reported for repairing subconjunctival damage, they often face challenges including acute foreign body responses and limited degradability.^16–18 Borate ester bonds are frequently employed in the fabrication of responsive hydrogels that can be degraded by specific stimuli such as pH and ROS and have been exploited in drug delivery platforms.^18–22 For example, borate ester-based hydrogels have been employed as a drug delivery platform for the administration of rapamycin to treat subconjunctival fibrosis.^23–25 However, the relatively rapid degradation kinetics of borate ester-based networks often fall short of meeting the long-term requirements for subocular implantable materials.^26–29 Sulfides can be oxidized by ROS into sulfones or sulfoxides without cleavage. When incorporated into the crosslinked network of hydrogels, they can also enhance the long-term stability and antioxidant properties of the hydrogel.^30–32

Here, we designed a dual-crosslinked and ROS-scavenging hydrogel containing multiple ROS-scavenging crosslinking structures through thiol–ene click chemistry and dynamic covalent bonding (CSMA-PBA/EPL-SH, CPES). The crosslinking structures are mainly sulfide bonds and borate ester bonds (Fig. 1). The CPES hydrogel was prepared using a one-step blending method by mixing thiolate polylysine, acrylamidated and boronated bifunctional sulfated chondroitin sulfate, and polyvinyl alcohol. Covalent thioether bonds support stable mechanical properties, while borate ester bonds confer ROS responsiveness and dynamic performance.^27,28,33,34 Experimental results indicate that the hydrogel exhibits tunable degradability, strong radical scavenging capacity, adaptability, and dynamic properties. Additionally, it effectively downregulates ROS levels within conjunctival epithelial cells, protecting them from oxidative stress-induced damage. Immunofluorescence analysis further demonstrated the enzyme-like activity of CPES hydrogel in scavenging ROS. By establishing a rat subconjunctival injury model, we validated the hydrogel's ability to restrain inflammation in vivo. Therefore, this dual-crosslinked, ROS-scavenging CPES hydrogel offers a promising therapeutic strategy for addressing subconjunctival inflammation.

Fig. 1 Schematic diagram of the preparation of CPES hydrogels, the mechanisms of reactive oxygen species scavenging and its application of subconjunctival treatment.

2. Materials and methods

2.1. Materials

1-(3-Dimethylaminopropyl)-3-ethylcarbodiimide (EDC), N-hydroxysuccinimide (NHS), and ε-poly-L-lysine (EPL) were procured from Aladdin Bio-Chem Technology Co., Ltd (Shanghai, China). DL-N-Acetylhomocysteine thionolactone, glycidyl methacrylate (GMA), chondroitin sulfate (CS), and polyvinyl alcohol (PVA) were procured from Macklin Biochemical Technology Co., Ltd (Shanghai, China). Human corneal epithelial cells (HCE-2) and rat conjunctival epithelial cells (rPCEC) were acquired from the Henan Provincial Eye Hospital. Macrophages (RAW264.7) and their culture medium were purchased from Zhongqiao Xinzhou Biotechnology Co., Ltd. The Cell Counting Kit-8 (CCK-8) was purchased from Sigma-Aldrich Trading Co., Ltd. (Shanghai, China). 2′,7′-Dichlorodihydrofluorescein diacetate (DCFH-DA) fluorescent probe and DMEM/F12 medium were purchased from Solarbio Technology Co., Ltd (Beijing, China).

2.2. Preparation of EPL-SH

Firstly, 1 g of polylysine hydrochloride was dissolved in 20 mL of deionized water, and the pH was adjusted to 9. DL-N-acetyl homocysteine sulfonyl lactate hydrochloride (0, 100, or 200 mg) was added and the reaction proceeded for 4 hours under a nitrogen atmosphere. The product was collected and dialyzed (molecular weight cut-off: 3500 Da) against deionized water with a pH of 3–5 for 3 days, replacing the dialysate 3 times daily. Depending on the different ratios of DL-N-acetyl homocysteine sulfonyl lactate hydrochloride added, the resulting sulfhydryl modified polylysine (EPL-SH) was named EPL-SH₀, EPL-SH₁₀, and EPL-SH₂₀, respectively.³⁵

2.3. Preparation of CSMA and CSMA-PBA

Methyl acrylate-sulfated chondroitin sulfate (CSMA) was prepared through the ring-opening reaction between the hydroxyl group of chondroitin sulfate and epoxypropyl methyl acrylate. Firstly, 5 g of CS was dissolved in 50 mL of deionized water, followed by the addition of 15 mL of GMA. The reaction mixture was stirred at 60 °C for 2 days. Subsequently, the product was precipitated by adding 200 mL of ethanol (repeat 3 times). The precipitate was collected, vacuum-dried, and stored at −20 °C.

Phenylboronic acid-modified methyl acrylate chondroitin sulfate (CSMA-PBA) was prepared via EDC/NHS-mediated amidation. Firstly, 2 g of CSMA was dissolved in 10 mL of deionized water. After complete dissolution, 400 mg of EDC and 200 mg of NHS were added and stirred for 2 hours to activate carboxyl groups. Then, 100 mg of 3-aminobenzoic acid was added, and the reaction was continued for 10 hours. The sample solution was collected, and dialyzed against deionized water for 3 days (changing the dialysis solution 3 times daily), with a molecular weight cutoff of 3500 Da. Finally, the sample was freeze-dried and stored at −20 °C.³⁶

2.4. Preparation of CPES hydrogel

The PVA solution was obtained by dissolving 1 g of PVA in 50 mL of deionized water at 85 °C with stirring until complete dissolution. The CPES hydrogel was prepared by simple one-step mixing of aqueous solutions: CSMA-PBA solution (200 mg mL⁻¹), EPL-SH solution (50 mg mL⁻¹), and PVA solution (20 mg mL⁻¹) in a volume ratio of 2 : 1 : 1. Hydrogels prepared with EPL-SH₀, EPL-SH₁₀, and EPL-SH₂₀ were designated CPES-1, CPES-2, and CPES-3, respectively.

2.5. Physicochemical characterizations

The lyophilized sample (20 mg) was subsequently dissolved in a suitable amount of deuterated water (D₂O). 5 mg of lyophilized sample was dissolved in 1 mL of deuterium oxide and measured using a Bruker AV400 spectrometer operating at 400 MHz to get nuclear magnetic resonance hydrogen spectrum (¹H NMR) data. Fourier transform infrared (FT-IR) spectroscopy was performed on lyophilized samples using the KBr pellet method with a scanning wavenumber range of 4000–500 cm⁻¹ at a resolution of 2 cm⁻¹. The acquisition of X-ray photoelectron spectroscopy (XPS) data was carried out by grinding the freeze-dried samples with a ball mill and then through AXIS SUPRA spectrometer. Furthermore, the crosslinked structure of the hydrogel was characterized using XPS and FT-IR.

2.6. Equilibrium swelling rate

Hydrogel samples were placed in a freeze dryer and completely dried, and the dry weight of the sample was recorded as W₀. The sample was completely immersed in phosphate-buffered saline (PBS) at pH 7.4 until equilibrium swelling was achieved. The residual liquid was gently removed with filter paper, and the swollen weight was recorded as W_x. All samples were independently repeated 3 times using the above method.³⁷ The equilibrium swelling rate is calculated as follows:

Equilibrium swelling rate (%) = (W_x − W₀)/W₀ × 100%

2.7. Degradation experiments

The degradation profiles of the hydrogels under physiological and oxidative stress conditions were evaluated hydrogels were weighed (M₀) and incubated in either PBS (pH 7.4) or PBS containing H₂O₂. At predetermined time points, samples were removed, rinsed with water, lyophilized, and weighed (M_x). The experiment was repeated 3 times.³⁸ The formula for calculating the degradation rate of hydrogel is as follows:

Degradation rate (%) = (M₀ − M_x)/M₀ × 100%

2.8. Self-healing properties and adaptive performance

To better demonstrate self-healing effects, two hydrogel pieces were stained with methylene blue and alizarin red, respectively. The stained surfaces were gently brought into contact. After a defined healing period, manual stretching was performed to assess the fusion and recovery of mechanical integrity at the interface.³⁹ To characterize the hydrogel's adaptive ability, the hydrogel was placed in a glass bottle containing glass beads to observe its ability to adapt to confined spaces.⁴⁰

2.9. Rheological tests

Rheological properties were analyzed using a rotational rheometer equipped with parallel plate geometry. Firstly, by varying the frequency (0.01–10 Hz) while keeping the strain constant at 0.1%, the storage modulus (G′) and loss modulus (G″) of the hydrogel were tested. Subsequently, strain scanning tests (0.01–1000%) were conducted with a fixed frequency of 1 Hz to assess the strain stability of the hydrogel. The intersection points of G′ and G″ was recorded, which corresponds to the hydrogel's fracture strain. Finally, the self-healing performance of the hydrogel was characterized using an alternating strain test. Alternating low (10%) and high (500%) strain amplitudes were applied at 1 Hz to evaluate the self-recovery capability over multiple cycles.⁴¹

2.10. Antioxidant performance test

The free radical scavenging activity was evaluated using 2,2-diphenyl-1-picrylhydrazyl (DPPH) and 2,2′-azinobis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) assays.^42,43 100 mg of freeze-dried hydrogel samples were incubated with pre-prepared DPPH solution or ABTS solution. After 30 minutes, the color changes of the solutions were observed and recorded. The absorbance of the solutions was measured at 517 nm (DPPH) or 734 nm (ABTS) using a microplate reader, respectively. The same method was used to mix the test solutions with blank media (DPPH: ethanol, ABTS: water), and the absorbance of the solution was measured at 517 nm or 734 nm and recorded as A₀. Absorbance of sample solutions was recorded as A_x. The calculation methods for DPPH or ABTS are as follows:

DPPH/ABTS scavenging rate (%) = (A₀ – A_x)/A₀ × 100%

2.11. Antimicrobial properties test

Staphylococcus aureus (S. aureus) and Escherichia coli (E. coli) suspension were incubated with CPES hydrogel for 6 h. Subsequently, the bacterial suspension was suspended with sterile saline. Aliquots (100 µL) of appropriate dilutions were spread onto nutrient agar. Plates were incubated overnight at 37 °C, and colony-forming units were counted and photographed.

2.12. Cytocompatibility and intracellular ROS regulation

2.12.1. Cell viability. The biocompatibility of the CPES hydrogel was evaluated using the CCK-8 cell counting method. First, rPCEC and HCE-2 were seeded into 96-well plates. Then, the cells were treated with hydrogel extracts at different concentrations (25, 50, 75, 100, 200 mg mL⁻¹) for 24 hours. Finally, CCK-8 assay solution was added, and the mixture was incubated for 3 hours. The absorbance was measured at 450 nm to calculate the cell survival rate.

2.12.2. Intracellular ROS detection. Intracellular ROS levels were assessed by DCFH-DA staining. First, rPCEC cells were uniformly seeded into 6-well plates. After removing the culture medium, the control group was treated with culture medium solution, the lipopolysaccharide (LPS) group was treated with culture medium solution containing 2 µg mL⁻¹ LPS, and the CPES group was treated with culture medium solution containing 2 µg mL⁻¹ LPS and 100 mg mL⁻¹ hydrogel extract. After incubation for 24 hours, cells were treated with culture medium solution containing 10 µM DCFH-DA for 30 minutes, washed three times with PBS, and photographed using an inverted fluorescence microscope.⁴⁴

2.13. Immunofluorescence

RAW264.7 macrophages were divided into three groups: Control, LPS, and CPES groups to assess the enzyme-like activity of the hydrogel. The CPES group was incubated in medium containing CPES hydrogel extract for 24 hours, followed by stimulation with 1 µg mL⁻¹ LPS for 16 hours. After treatment, the cells were fixed with 4% paraformaldehyde for 10 minutes, lysed with 0.5% Triton for 20 minutes, and blocked with goat serum for 30 minutes. Primary antibodies against superoxide dismutase (SOD), catalase (CAT), and glutathione peroxidase (GPx) were added and incubated at room temperature for 1 hour. FITC-labeled goat anti-rabbit IgG H&L secondary antibodies were added and incubated at room temperature for 1 hour. Cell nuclei were stained with DAPI. Finally, fluorescence images were captured and recorded under a fluorescence microscope.⁴⁵

2.14. Animal experiments

All animal procedures were approved by the Institutional Animal Care and Use Committee of the Experimental Animal Ethics Committee of Henan Eye Hospital (permit number: HNEECA-2022-21). SD rats (female, 8 weeks old) were randomly divided into control group and CPES group, with 6 rats in each group. Anesthesia was induced by intraperitoneal injection of sodium pentobarbital 2% (2 mL kg⁻¹). The eyes and surrounding skin were disinfected with povidone–iodine. A conjunctival incision was made along the temporal corneal limbus (approximately one-third of the corneal circumference) using ophthalmic scissors, blunt dissection was performed to create a conjunctival pocket. In the CPES group, a hydrogel was placed in the temporal conjunctival sac. Postoperative care included applying gatifloxacin eye gel to the conjunctival sac. Conjunctival edema and hyperemia were assessed at 3, 5, 7, and 14 postoperative days, respectively, and observation and photographs were taken using a slit lamp.

Rat ocular tissue was collected, fixed with FAS ocular fixative, paraffin-embedded, and sectioned. Samples were stained with hematoxylin and eosin (H&E) on days 3, 5, 7, and 14 postoperatively to assess the inflammatory control capacity of the hydrogel. Tissue sections were stained for interleukin-1 beta (IL-1β) and 8-hydroxy-2′-deoxyguanosine (8-OHdG) to evaluate inflammation and oxidative DNA damage, respectively. Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay was performed to assess apoptosis.

2.15. Statistical analysis

Statistical analyses were conducted using GraphPad Prism 8.0. Results are presented as the mean ± standard deviation, with each experiment repeated at least three times. Statistical significance between groups was determined by one-way analysis of variance (ANOVA) followed by Bonferroni post-test. Statistical significance was set at P < 0.05.

3. Results and discussion

3.1. Preparation and characterization of CSMA-PBA and EPL-SH

Methyl acrylate-functionalized and phenylboronic acid-modified CSMA-PBA was synthesized via a two-step approach. Additionally, thiol-functionalized EPL-SH was prepared through an amide reaction. The specific synthesis routes are detailed in Fig. S1. The chemical modification of CS to CSMA was confirmed by FT-IR and ¹H NMR spectroscopy. The FT-IR spectrum of CSMA exhibited distinct characteristic absorption peaks at 1714 cm⁻¹ and 1490 cm⁻¹, which are attributed to the C=O and C=C, respectively, confirming the successful conjugation of methyl acrylate groups onto the CS backbone (Fig. 2(A)). Further evidence for successful methacrylation was obtained from ¹H NMR. Both CSMA and CSMA-PBA spectra displayed characteristic resonance peaks at 6.2 and 5.8 ppm, respectively, corresponding to the methacrylation modification. The triple peak output of 7.7 ppm is attributed to the aromatic protons, indicating that the phenylboronic acid was successfully modified (Fig. 2(B)).

Fig. 2 Characterization of CSMA-PBA and EPL-SH. The FT-IR spectrum of (A) CSMA-PBA and (C) EPL-SH. The ¹H NMR spectrum of (B) CSMA-PBA and (D) EPL-SH.

EPL-SH was synthesized by reacting an alkaline ring-opening DL-N-acetyl homocysteine thioester with EPL. The FT-IR spectrum of EPL-SH exhibited prominent thiol absorption band (780 cm⁻¹) and a thioester carbonyl absorption band (1585 cm⁻¹), indicating the successful preparation of EPL-SH (Fig. 2(C)). Comparative ¹H NMR analysis of EPL-SH and EPL revealed characteristic peaks of methyl groups at 2.3 ppm and 1.8 ppm in the EPL-SH spectrum (Fig. 2(D)), providing conclusive evidence for the successful preparation of EPL-SH.

3.2. Preparation and characterization ofthe CPES hydrogel

CPES hydrogels were formed through the spontaneous crosslinking of CSMA-PBA and EPL-SH at room temperature. The gelation process involves the formation of borate ester dynamic bonds through the reaction of boronic acid groups with the hydroxyl groups of PVA, as well as the addition reaction between the thiol groups of EPL-SH and the vinyl groups. The formation of the crosslinked structures including of borate ester bonds, sulfide bonds, and disulfide bonds was confirmed using FT-IR and XPS. The FT-IR spectrum of the CPES hydrogel exhibited a characteristic absorption peak at 1381 cm⁻¹, corresponding to the B–O bond stretching vibration, confirming borate ester bond formation. While the absorption peak at 1190 cm⁻¹ was assigned to C–S stretching vibrations, signifying the presence of sulfide/thioether bonds in the CPES hydrogel (Fig. 3(A)). The elemental composition of the crosslinked structures was further confirmed by XPS analysis. The peaks observed at 285.4 eV, 398.4 eV, 531.6 eV, and 180.0 eV were assigned to C, N, O, and S, respectively (Fig. 3(B)). Moreover, the high-resolution C1s, N1s, and O1s spectra provide further insight into the chemical bonding environment within the hydrogel (Fig. 3(C)–(E)). In addition, the S2p spectrum displayed characteristic peaks corresponding to C–S and S–S bonds (Fig. 3(F)), supporting the formation of disulfide crosslinks.

Fig. 3 Characterization of CPES hydrogel. (A) The FT-IR spectrum of CPES hydrogel. (B) The XPS spectrum of CPES hydrogel. XPS high-resolution spectrum of (C) C1s, (D) N1s, (E) O1s and (F) S2p.

3.3. Dynamic performance of CPES hydrogel

For applications on irregular trauma surfaces and in deep tissue compartments, the shape-adaptive and autonomous self-healing capabilities of hydrogels are required. First, we used a simple irregular mold made with beads and test tubes to test the adaptability of CPES hydrogels (Fig. S2A). Upon placement of an alizarin red/methylene blue-stained CPES hydrogel atop the bead bed and static incubation for 30 min, the hydrogel spontaneously flowed and completely filled the interstitial spaces between the beads (Fig. S2A), indicating exceptional shape adaptability and moldability. The intrinsic self-repair capability was investigated by bisecting a hydrogel sample and gently bringing the freshly cut surfaces into contact. The results showed that after 1 hour of contact, the interface disappeared. The healed hydrogel could withstand manual handling and stretching without rupture. Additionally, to investigate the effect of crosslink density on the elongation at break of the hydrogel, tensile tests on hydrogels from different groups were performed. The results showed that CPES-1 exhibited the highest tensile strain at break, with the tensile properties decreasing proportionally with increasing sulfide bond density within the hydrogels (CPES-1 > CPES-2 > CPES-3) (Fig. S2B). This trend is attributed to the progressively increasing density of covalent sulfide bonds, forming a more rigid network.

3.4. Swelling, degradation and mechanical properties of CPES hydrogel

The swelling behavior and degradation kinetics of hydrogels are critical determinants of their performance in biomedical applications. As shown in Fig. 4(A), the equilibrium swelling ratio of CPES hydrogels decreased gradually with increasing thiol functionalization, which was attributed to differences in crosslinking density and amounts of hydrophobic sulfide bonds. To assess the ROS-responsive degradation behavior of the hydrogels, degradation tests were conducted under physiological (non-oxidative) and oxidative (1 mM H₂O₂) conditions. Under physiological conditions, CPES-1 was completely degraded by day 3, while CPES-2 and CPES-3 maintained structural integrity by day 7 (Fig. 4(B)), benefiting from their higher sulfide bond density. The oxidative environment accelerated the degradation kinetics of all hydrogel formulations. This accelerated degradation stems from the oxidative cleavage of dynamic borate ester bonds and potentially disulfide linkages within the network. Notably, although CPES-3 exhibited the best degradation performance, degrading completely within 4 days (Fig. 5(C)), this pronounced ROS-responsiveness highlights the therapeutic potential for targeted degradation at inflamed sites rich in ROS.

Fig. 4 Swelling behavior, degradation kinetics and mechanical performance of CPES hydrogel. (A) Swelling of CPES hydrogels. Degradation of CPES hydrogels under (B) non-oxidizing and (C) oxidizing conditions. (D) The frequency sweep, (E) strain amplitude sweep and (F) self-healing rheological test of CPES hydrogel. The data are presented as the means ± SDs.

Fig. 5 Antioxidant properties of CPES hydrogels. Optical photograph of CPES hydrogel scavenging (A) DPPH and (B) ABTS. Statistical results of CPES hydrogels scavenging (C) DPPH and (D) ABTS. The data are presented as the means ± SDs.

Rheological characterization revealed frequency-dependent viscoelastic properties (Fig. 4(D)) and Fig. S3A, S4A). CPES-1 exhibited fluid-like behavior at low frequencies (0.01–2 Hz), where the loss modulus (G″) exceeded the storage modulus (G'), potentially attributed to dynamic borate ester dissociation. CPES-2 exhibited similar viscoelastic instability at higher frequencies (2–10 Hz) due to insufficient sulfide crosslinking density. In contrast, CPES-3 maintained stable gel behavior across the entire frequency range (0.01–10 Hz), confirming excellent network stability. Strain scanning results indicated that mechanical stability significantly improved with increasing crosslinking degree (Fig. 4(E) and Fig. S3B, S4B). Alternating strain testing revealed structural collapse in CPES-3 under high strain conditions. Notably, the self-healing capability of CPES-2 is comparable to that of CPES-1, indicating that an appropriate sulfide content can maintain dynamic performance without sacrificing toughness (Fig. 4(F) and Fig. S3C, S4C). Self-healing rheological testing revealed structural collapse in CPES-3 during the second cycle, suggesting increased brittleness from excessive crosslinking. Notably, CPES-2 maintained self-healing capability equivalent to CPES-1, demonstrating that optimal thioether incorporation preserves dynamic performance without compromising resilience.

3.5. Antioxidant properties and antimicrobial properties of CPES hydrogel

Subconjunctival defects induce the accumulation of a large amount of ROS, significantly delaying subconjunctival healing. To evaluate the free radical scavenging capacity of hydrogels, we performed quantitative analyses using DPPH and ABTS as targets. The DPPH and ABTS solutions treated with different hydrogels exhibited different colors, indicating that the hydrogels had radical scavenging capacities (Fig. 5(A) and (B)). As shown in Fig. 5(C), the CPES-1 hydrogel, which has only borate ester dynamic bonds as its crosslinking structure, exhibited limited scavenging capacity for DPPH (approximately 20%). In contrast, the CPES-2 and CPES-3 hydrogels with multiple crosslinking structures exhibited excellent DPPH scavenging capabilities (57% and 66%, respectively). This suggests that the free radical scavenging capability of hydrogels is related to the content of ROS-sensitive chemical bonds. Additionally, the ABTS scavenging test yielded similar results (Fig. 5(D)). The antibacterial properties of three hydrogels were evaluated. As shown in Fig. S5, all hydrogels inhibited the growth of S. aureus and E. coli, conforming the antibacterial properties of polylysine molecules. The amino sites of polylysine were modified without affecting the antibacterial ability.

3.6. Cytocompatibility and intracellular ROS regulation

HCE-2 and rPCEC cells were incubated with different concentrations of CPES hydrogel extract for 24 hours, and the cell survival rate was detected by CCK-8. Even under the influence of a 200 mg mL⁻¹ hydrogel extract, the cell survival rate remained above 80%, indicating that the CPES hydrogel exhibited excellent cell compatibility (Fig. 6(C) and (D)). Following the validation of antioxidant properties at the cellular level, we characterized the hydrogel's ability to scavenge ROS within conjunctival epithelial cells using the DCFH-DA fluorescence assay. High ROS levels were induced in cells using LPS, resulting in higher fluorescence intensity observed by microscopy (Fig. 6(A)). After treatment with the CPES hydrogel extract, fluorescence intensity significantly decreased, consistent with the statistical results of the mean fluorescence intensity (MFI), indicating that the CPES hydrogel exhibited a certain ability to scavenge intracellular ROS (Fig. 6(A) and (B)).

Fig. 6 Intracellular ROS regulation and cytocompatibility of CPES hydrogel. Representative fluorescence microscopy images (A) and fluorescence intensity statistics (B) of rPCEC stained with DCFH-DA. (C) Metabolic activity of rPCEC after 24 hours exposure to hydrogel extracts. (D) Metabolic activity of HCE-2 after 24 hours exposure to hydrogel extracts. Scale bar: 20 µm. The data are presented as the means ± SDs (*P < 0.05, **P < 0.01, and ***P < 0.001).

3.7. Immunofluorescence staining

Antioxidant enzymes are crucial for maintaining redox homeostasis. We further confirmed antioxidant properties by assessing changes in enzyme activity before and after CPES treatment. Beyond direct ROS scavenging, we evaluated the hydrogel's capacity to augment endogenous antioxidant defenses by modulating key redox-regulatory enzymes including of SOD, CAT, and GPx in macrophages. Immunofluorescence and statistical analysis of average fluorescence intensity revealed that LPS significantly inhibited the activity of SOD, CAT, and GPx, while the enzyme activity in the CPES hydrogel group improved (Fig. 7), indicating that CPES hydrogels possess notable antioxidant capacity.

Fig. 7 CPES hydrogel enhancing macrophage enzyme activity after oxidative stress. Images of (A) CAT, (B) SOD, (C) GPx enzyme activity and (D) statistics after different treatments. Scale bar: 10 µm. The data are presented as the means ± SDs (**P < 0.01, ***P < 0.001).

3.8. In vivo therapeutic effects on subconjunctival injury

To evaluate the potential of CPES hydrogel in treating subconjunctival injury, we established an in vivo animal model of subconjunctival damage: the control group received no additional intervention, while the experimental group was treated with CPES hydrogel. On days 3, 5, 7, and 14, the stage of tissue repair was documented using a slit lamp (Fig. 8(A)) and analyzed via H&E staining (Fig. 8(B)). The results demonstrated that the CPES-treated group exhibited significantly better tissue repair outcomes compared to the control group, with a notable reduction in inflammatory cell infiltration in the tissue.

Fig. 8 CPES hydrogel inhibiting subconjunctival inflammation in vivo. (A) Typical images of conjunctival injury (B) H&E staining images of each group at 3, 5, 7 and 14 days postoperatively. Scale bar: 100 µm.

Representative inflammatory factors IL-1β (Fig. 9A and C) and oxidative stress indicator 8-OHdG (Fig. 9B and D) were selected for immunofluorescence staining analysis on conjunctival tissue to verify the anti-inflammatory effect of CPES hydrogel in vivo. The results showed that the expression levels of IL-1β factors after CPES treatment were significantly lower than those in the control group, demonstrating that CPES hydrogel has good anti-inflammatory properties in vivo, similar to the H&E staining results. Immunofluorescence quantification showed a 25% decrease in the oxidative DNA damage marker 8-OHdG after CPES treatment at day 7 (**P < 0.001). TUNEL assays confirmed no significant apoptosis in the corneal and retinal tissues of both groups (Fig. S6), indicating that the hydrogel system had good biocompatibility.

Fig. 9 CPES hydrogel decreases the expression of inflammatory cytokines in vivo. (A) IL-1β expression and (C) statistical results of the control group and the operation group at different times. The expression of (B) 8-OHdG and (D) statistics of the control group and the operation group at different times. Scale bar: 50 µm. The data are presented as the means ± SDs (**P < 0.01, and ***P < 0.001).

4. Conclusions

In this work, we have engineered CPES hydrogels through a one-step mixing method, integrating dual ROS-scavenging networks through dynamic borate esters and thioether/disulfide bonds. CPES hydrogels exhibit excellent antioxidant activity, tunable mechanical properties, outstanding biocompatibility, and anti-inflammation capabilities. In a subconjunctival injury model, CPES significantly accelerated tissue repair while concurrently mitigating oxidative damage, inflammatory cascades, and microbial infection. This work establishes a new paradigm for ocular regenerative medicine by demonstrating that rationally designed multi-network hydrogels can simultaneously resolve the pathological ROS accumulation, chronic inflammation, and infection risk. The CPES hydrogel represents a transformative platform for preventing vision-threatening subconjunctival fibrosis.

Conflicts of interest

The authors declare that there is no potential conflicts of interest and no commercial or financial relationships in this study.

Data availability

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

Supplementary information (SI) is available. See DOI: https://doi.org/10.1039/d5tb01859e.

Acknowledgements

This research was supported by the National Natural Science Foundation of China (52173143 and 82371108), Henan Province Science and Technology Research and Development Plan Joint Fund Project (242301420010), Natural Science Foundation of Henan Province of China (242300421018 and 252300421024), Henan Province Youth Health Science and Technology Innovation Talent Training Project (LJRC2024003), 2024 Young and Middle-aged Academic Leaders of Health Care in Henan Province and Henan Province Clinical Medical Scientist Training Program (HNCMS202402).

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Footnote

† These authors contributed equally to this work.

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