A redox-responsive NIR fluorescent nanoprobe for tumor microenvironment-activated surgical navigation with submillimeter precision

Abstract

Surgical precision in tumor resection critically relies on real-time intraoperative imaging, yet conventional probes face limitations in specificity and spatiotemporal control. Here, we present a tumor microenvironment (TME)-activated near-infrared (NIR) fluorescent nanoprobe (DNS–DYE/PEG–NI) that integrates dual responsiveness to hypoxia and glutathione (GSH) for submillimeter-level surgical navigation. The system comprises a GSH-activatable NIR fluorophore (λ_ex/em = 679/730 nm) quenched by 2,4-dinitrobenzenesulfonyl (DNS) moieties and hypoxia-sensitive amphiphilic PEG–NI micelles. Upon tumor accumulation via the enhanced permeability and retention (EPR) effect, a hypoxic TME triggers micelle disassembly through nitroimidazole (NI) reduction, releasing DNS–DYE. Subsequent GSH cleavage restores fluorescence via intramolecular charge transfer (ICT) recovery, achieving a 12.3-fold tumor-to-normal tissue signal ratio and >90% reduction in off-target activation compared to non-responsive controls. Systematic validation demonstrates: (1) dose-dependent fluorescence recovery (35-fold intensity increase at 10 mM GSH); (2) hypoxia-driven micelle destabilization (800% hydrodynamic diameter expansion); (3) sustained colloidal stability (12.9% size variation over 15 days); and (4) low cytotoxicity (cell viability >90% at 125 μg mL⁻¹). In vivo studies reveal precise tumor delineation within 12 h post-injection, enabling real-time resection of submillimeter lesions. By coupling TME-specific activation with prolonged tumor retention, this dual-responsive nanoprobe advances fluorescence-guided surgery toward precision oncology, reducing positive margin rates from 70% to <5% in preclinical models.

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1 Introduction

Cancer remains a major global public health challenge and the leading cause of mortality worldwide. Surgical resection, currently the most effective clinical intervention for solid tumors, relies critically on precise intraoperative identification of tumor margins and micrometastatic lesions.^1,2 However, conventional preoperative imaging techniques—such as X-ray, computed tomography (CT), and magnetic resonance imaging (MRI)—have notable intraoperative limitations, including poor visual discrimination of tumor tissue under white-light illumination, spatial misregistration due to tissue deformation during surgical manipulation, and insufficient sensitivity for detecting submillimeter lesions.^3,4 Intraoperative MRI and ultrasound, while providing real-time guidance, are hindered by high costs, bulky equipment, and prolonged procedural interruptions. These technical constraints contribute to residual tumor burden (positive surgical margins) in approximately 70–80% of cases,⁵ underscoring the imperative to develop real-time intraoperative imaging guidance systems for optimizing tumor resection completeness.

Compared with intraoperative MRI, ultrasound, and other techniques involving ionizing radiation or complex equipment, fluorescence probe-based optical imaging demonstrates unique advantages such as high spatial resolution (<1 mm), operational convenience, and radiation-free characteristics, which have established it as a cutting-edge approach for real-time navigation.⁶ The concept of fluorescence-guided surgery traces its origins to the demonstration of utilizing activatable probes for in vivo imaging. Subsequent advancements in nanotechnology, particularly the discovery of the enhanced permeability and retention (EPR) effect, laid the foundation for tumor-targeted nanocarriers for delivering probes. Near-infrared (NIR) fluorescence imaging, in particular, offers superior tissue penetration (up to 5–10 mm), minimal autofluorescence interference, and compatibility with real-time surgical workflows.^7–9 Notably, FDA-approved indocyanine green (ICG) has been widely adopted for sentinel lymph node mapping and angiography.^10,11 However, ICG suffers from critical shortcomings, including rapid systemic clearance (<10 min plasma half-life), non-specific vascular leakage, and aggregation-induced fluorescence quenching in aqueous media, which limit its utility for tumor-specific delineation.¹² Other clinically explored probes, such as 5-aminolevulinic acid (5-ALA), face challenges in penetration depth and tumor-to-background ratios. The development of intelligent probe systems integrating tumor-targeted accumulation with microenvironment-responsive activation has become imperative.

The pathological characteristics of the tumor microenvironment (TME) provide distinctive targets for probe design. Malignant proliferation in solid tumors induces abnormal metabolism and vascular dysfunction, creating hypoxic microenvironments in over 60% of malignancies,^13,14 with hypoxia severity positively correlating with tumor aggressiveness. Concurrently, tumor cells maintain redox homeostasis through upregulated expression of reductase enzymes (nitroreductase and azoreductase), nicotinamide adenine dinucleotide phosphate (NADPH), and elevated production of reduced glutathione (GSH)—reaching concentrations up to 1000-fold higher than normal tissues.^15–17 This dual microenvironmental signature, combining hypoxia and hyper-reduction, presents an ideal molecular switch for constructing tumor-specific responsive probes.

Building upon this rationale, we engineered a GSH-activatable NIR probe, DNS–DYE (Scheme 1a), through structural modification strategies to enhance tumor specificity. The probe was synthesized by conjugating DYE (λ_ex/em = 679/730 nm) with 2,4-dinitrobenzenesulfonyl chloride (DNS).¹⁸ This design capitalizes on DNS's strong electron-withdrawing effect and nitro-based fluorescence quenching properties, maintaining the probe in a fluorescence-quenched (“off”) state in normal tissues.¹⁹ To optimize pharmacokinetic performance, hydrophobic DNS–DYE was encapsulated within amphiphilic block copolymer micelles (Scheme 1c), enabling passive tumor targeting via the enhanced permeability and retention (EPR) effect.^20,21 Furthermore, hypoxia-responsive nitroimidazole (NI) moieties were incorporated into the polymer's hydrophobic segment, resulting in the construction of dual-responsive PEG–NI nanocarriers (Scheme 1b).

Scheme 1 Schematic diagram of the project design. (a) The structure of the “Off” state probe DNS–DYE; (b) the structure of the hypoxia-responsive polymer PEG–NI; (c) amphiphilic polymers and hydrophobic probes self-assemble to form micelles; and (d) intramicellar behavior of DNS–DYE/PEG–NI micelles.

The mechanistic cascade involves hypoxia-triggered micelle disassembly. Upon EPR-mediated tumor accumulation, hypoxic microenvironments induce NI reduction to hydrophilic aminoimidazole, destabilizing micellar structures and releasing DNS–DYE;²² GSH-mediated fluorescence activation can be achieved by the released probes undergoing DNS cleavage via tumor-overexpressed GSH, restoring DYE fluorescence through intramolecular charge transfer (ICT) recovery, and achieving tumor-specific “off-to-on” switching (Scheme 1d). This dual-responsive system employs a spatiotemporally coupled release-activation mechanism, demonstrating potential as an innovative solution for intraoperative real-time tumor margin delineation and complete resection rate improvement.

2 Experimental procedures

2.1 Materials

All chemicals and reagents were obtained from commercial suppliers and used without further purification unless otherwise specified. IR-780, DNS and 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) were purchased from Sigma-Aldrich (Shanghai, China), stored at −20 °C. N^ε-Carbobenzyloxy-L-lysine (Z-Lys(Z)-OH), N,N-dimethylformamide (DMF), dichloromethane (DCM), triphosgene and trifluoroacetic acid (TFA) were sourced from Aladdin (Shanghai, China). Hydrogen bromide (HBr) and acetic acid (AcOH) were obtained from Energy Chemical (Shanghai, China). p-Nitrophenyl chloroformate (PNP), 4-dimethylaminopyridine (DMAP) and 4-chlororesorcinol (4-CR) were supplied by Sun Chemical Technology (Shanghai) Co., Ltd. Triethylamine (TEA), methanol (MeOH) and anhydrous sodium dithionite (Na₂S₂O₄) were acquired from Sinopharm Chemical Reagent (Shanghai, China). Methoxy–poly(ethylene glycol)–amine (CH₃O–PEG_5k–NH₂) was acquired from J&K Scientific Ltd (Beijing, China), M_n = 5000 Da, PDI < 1.1, stored at −20 °C. Acetonitrile (ACN) was purchased from CINC High Purity Solvents (Shanghai) Co. Ltd. GSH was provided by Dalian Meilun Biotechnology Co. Ltd. For cell culture experiments, Dulbecco's modified Eagle medium (DMEM) and fetal bovine serum (FBS) were obtained from Gibco BRL (Carlsbad, CA, USA). Trypsin–EDTA (0.25%) was sourced from Invitrogen Co. (Waltham, Massachusetts, USA). All anhydrous solvents are stored over molecular sieves (3 Å) and degassed with N₂ before use. Triphosgene handling was performed in a fume hood with strict temperature control (<50 °C) to avoid decomposition. Regenerated cellulose membranes (MWCO 3.5 kDa) pre-rinsed with EDTA were used to remove preservatives.

The cell lines used in this study were purchased from the Cell Bank/Stem Cell Bank affiliated with the Shanghai Institute of Biochemistry and Cell Biology (SIBCB). Orthotopic tumors were established by injecting 1 × 10⁶ cells suspended in 50 μL of PBS/Matrigel (1 : 1 v/v) into the 1st mammary fat pad of 6-week-old female BALB/c nude mice (Slac, Shanghai). Tumors were allowed to grow for 21 days to reach ∼100 mm³. All animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Shanghai Polytechnic University and conducted in compliance with the NIH Guide for the Care and Use of Laboratory Animals.

2.2 Equipment

¹H NMR spectra were recorded using a Varian Oxford NMR spectrometer (400 MHz, Palo Alto, California, USA) at room temperature. Chemical structures were obtained using ChemBio 3D Ultra (14.0 version, Cambridge Software, Massachusetts, USA). Transmission electron microscopy (TEM) images were acquired using an FEI Tecnai G2 SpiritBIOTWIN electron microscope (ThermoFisher, USA). Dynamic light scattering (DLS) and z-potential results were measured using a Malvern Instruments Zetasizer Nano at 25 °C. The cells were observed using a fluorescence microscope (Leica, Wetzlar, Germany). Small animal in vivo fluorescence imaging was obtained using a Molecular Biology Workstation (Image Visualization and Infrared Spectroscopy, IVIS, Caliper, Newton, MA, USA).

2.3 Synthesis of the “turn-off” type probe DNS–DYE

The synthesis of compound DNS–DYE was performed according to the modified literature procedure,⁵ as illustrated in Fig. 1a. Key challenges included maintaining anhydrous conditions to prevent hydrolysis of DNBS-Cl and optimizing the reaction stoichiometry to minimize byproducts. To address these, all reagents were dried under vacuum prior to use. In a typical experiment, compound DYE (103.8 mg, 0.2 mmol, 1.0 equiv.) was precisely weighed and transferred to a 50 mL two-necked round-bottom flask under anhydrous conditions. The compound was dissolved in anhydrous DCM (10 mL) with stirring. The reaction flask was then immersed in an ice bath to maintain the temperature at 0 °C, followed by dropwise addition of TEA (11.6 μL, 24.28 mg, 0.24 mmol, 1.2 equiv.) in three equal portions. A separate solution of 2,4-DNBS-Cl (63.84 mg, 0.244 mmol, 1.2 equiv.) in anhydrous DCM (10 mL) was prepared and slowly added to the reaction mixture via a syringe. The resulting mixture was protected from light and stirred at room temperature for 12 h. After completion, the reaction solvent was removed under reduced pressure using a rotary evaporator to afford the crude product, which was further purified by column chromatography on silica gel (eluent: DCM/MeOH = 20 : 1, v/v) with R_f = 0.35 to yield DNS–DYE as a purple solid (68.7 mg, 42.78%). High resolution MS (HR-MS) calc. for C₃₄H₃₁ClN₃O₈S⁺ [S-DYE-I]⁺ 676.1515, found, 676.1517 (Fig. S3, ESI†). The product was subsequently dried under vacuum for 1 h and stored at −20 °C in light-protected containers to ensure stability.

Fig. 1 Synthetic route for (a) the “turn-off” probe DNS–DYE and (b) the hypoxia-responsive polymer PEG–NI.

2.4 Synthesis of the hypoxia-responsive polymer PEG–NI

The synthesis of PEG–NI was accomplished through four sequential steps as outlined in Fig. 1b: (1) preparation of Lys–NCA, (2) synthesis of PEG–pLys, (3) synthesis of PNP–NI, and (4) conjugation to yield PEG–NI.

2.4.1 Preparation of Lys–NCA. Under a nitrogen atmosphere, Z-Lys(Z)-OH (3.0 g, 10.7 mmol) and triphosgene (1.27 g, 4.28 mmol) were dissolved in anhydrous tetrahydrofuran (30 mL) in a 100 mL two-necked flask. Triphosgene was added in three portions to avoid exothermic decomposition. The reaction mixture was stirred at 50 °C for 5 h. After cooling to room temperature, the product was precipitated with ice-cold anhydrous hexane, filtered, and dried under vacuum overnight to obtain Lys–NCA as a white solid.

2.4.2 Synthesis of PEG–pLys. CH₃O–PEG_5k–NH₂ (1.0 g, 0.2 mmol) and Lys–NCA (735 mg, 2.4 mmol) were dissolved in anhydrous DMF (20 mL) under nitrogen protection. To prevent premature termination, the reaction was shielded from moisture. After stirring at 50 °C for 48 h, the solution was dialyzed (MWCO 3.5 kDa) against deionized water for 24 h and lyophilized to yield the intermediate product as a white solid.

The obtained white solid (200 mg) was then treated with TFA (10 mL) and HBr/AcOH solution (0.5 mL, 33 wt%) at room temperature for 2.5 h. The reaction was quenched with DCM, concentrated by rotary evaporation, re-dissolved in DMF, and dialyzed (MWCO 3.5 kDa) against water. Lyophilization afforded PEG–pLys as a white solid.

2.4.3 Synthesis of PNP–NI. A solution containing 1-methyl-2-hydroxymethyl-5-nitroimidazole (NI-OH, 300 mg, 1.0 equiv.), DMAP (0.1 equiv.), and TEA (1.0 equiv.) in THF was prepared. Separately, PNP (1.5 equiv.) was dissolved in THF, which was added dropwise to the above solution under a nitrogen atmosphere and stirred overnight. Excess PNP ensured complete esterification. The crude product was purified by silica gel column chromatography to obtain PNP–NI.

2.4.4 Final conjugation (PEG–NI). PEG–pLys (1.0 equiv.), PNP–NI (20 equiv.), DMAP (0.1 equiv.), and TEA (1.0 equiv.) were combined in anhydrous DMF (20 mL) under nitrogen protection. Steric hindrance required prolonged reaction time. After 8 h, the mixture was sequentially dialyzed (MWCO 3.5 kDa) against DMF (8 h) and deionized water overnight, followed by lyophilization to obtain PEG–NI as a white solid.

2.5 Fluorescence spectral validation of GSH-responsive activation of DNS–DYE

The “off” state probe DNS–DYE can be converted into a “turn-on” fluorescent probe (DYE) upon reaction with GSH, which can be evidenced by the increase in fluorescence intensity. For experimental control establishment, two sets of samples were prepared. A 20 μM DNS–DYE solution in ACN alone was used as the negative control. The positive control was constructed by incubating 20 μM DNS–DYE in ACN with varying GSH concentrations (5, 10, 15, 20, 30, 40 and 60 μM) to demonstrate concentration-dependent activation. All solutions were incubated at 37 °C with continuous shaking (30 rpm) for 2 h. Fluorescence measurements were performed using a fluorescence spectrometer with an excitation wavelength of 679 nm and emission detection between 700 and 900 nm. Each concentration was tested in triplicate to ensure reproducibility.

2.6 Verification of intracellular GSH-responsive activation of DNS–DYE

MDA-MB-231/luci cells were cultured in DMEM containing 10% FBS and 1% antibiotics at 37 °C with 4% CO₂. The cells were seeded at 5 × 10⁴ cells per well in 24-well plates and incubated for 24 h to achieve 80–90% confluence. DNS–DYE was dissolved in DMSO and added to the DMEM medium to final concentrations of 0.25% DMSO and 2 × 10⁻⁴ mol L⁻¹ DNS–DYE. The experiment included two groups: group 1 was incubated with DNS–DYE, and group 2 served as a control with only DMSO in the medium, with three replicates per group. After 1 h, the medium was replaced with complete DMEM, and cells were cultured for an additional 0, 1, 2, 4, 8 or 12 h. Fluorescence was measured using a microplate reader at excitation and emission wavelengths of 650 nm and 700 nm, respectively.

2.7 Verification of hypoxia conversion for PEG–NI

PEG–NI (5 mg) was dissolved in 1 mL of DMF and dried into a uniform film using a rotary evaporator. The film was further dried overnight in a vacuum desiccator to remove residual solvent. Subsequently, 5 mL of distilled water was added, and the mixture was sonicated to obtain an empty micelle solution (1 mg mL⁻¹). To simulate hypoxic conditions, Na₂S₂O₄ (100 mM) was introduced into 1 mL of the micelle solution, followed by a 5 h reaction.²³ The UV absorption spectral changes were monitored before and after the reaction using a UV spectrophotometer.

2.8 Preparation of DNS–DYE/PEG–NI

Firstly, the critical micelle concentration (CMC) value of the PEG–NI polymer was determined via pyrene fluorescence spectroscopy. Briefly, a 100 μM pyrene solution (in acetone) was prepared, and 10 μL aliquots were dispensed into 10 EP tubes, followed by solvent evaporation. A series of PEG–NI solutions (250, 100, 50, 25, 10, 5, 2.5, 1.0, 0.5, and 0.25 μg mL⁻¹ in pure water) were added to the pyrene-containing tubes. The mixtures were incubated overnight at 37 °C under constant shaking. The fluorescence intensities at excitation 337 nm and emissions 373 nm/384 nm were measured using a microplate reader. A scatter plot was generated with the logarithm of concentration as the x-axis and the fluorescence intensity ratio as the y-axis. Linear regression yielded two straight lines, and the CMC was determined as the micelle concentration corresponding to their intersection point.

Secondly, DNS–DYE/PEG–NI micelles were prepared using the thin-film hydration method. Briefly, DNS–DYE (1 mg mL⁻¹ in DMF) and PEG–NI (10 mg mL⁻¹ in DMF) solutions were first prepared. Then, 500 μL of each solution was combined in a 25 mL round-bottom flask and mixed ultrasonically. The mixture was subsequently evaporated into a uniform purple film using a rotary evaporator (40 °C, 30 rpm, 15 min), followed by vacuum drying overnight to remove the residual solvent. The film was hydrated with 5 mL of pure water and sonicated for 2.5 h, yielding a blue-purple micellar solution (1 mg mL⁻¹). The final product was stored protected from light at 4 °C.

2.9 Characterization of DNS–DYE/PEG–NI

2.9.1 Particle size and morphological analysis. For the aforementioned 1 mg mL⁻¹ micelle solution, the hydrodynamic diameter was determined by DLS using a Malvern Nano ZS particle analyzer at 25 °C. For TEM, the micelle solution was concentrated to 5 mg mL⁻¹via ultrafiltration and centrifugation. A droplet was placed at the center of an ultrathin carbon support film to form a thin liquid layer, followed by negative staining with uranyl acetate solution. After standing at room temperature for 30 min, the sample was dried under an infrared lamp (30 min, 39.7 °C). The micelle morphology was then observed.

2.9.2 Determination of drug loading capacity (DL) and encapsulation efficiency (EE). DL and EE were analyzed by high performance liquid chromatography (HPLC). A methanol solution of DNS–DYE was prepared at a concentration of 0.01 mg mL⁻¹, respectively. The ultraviolet absorption spectra were recorded within the wavelength range of 200–800 nm using a UV spectrophotometer. The maximum absorption wavelength of DNS–DYE was selected as the detection wavelength for the HPLC UV detector.

The analysis was performed on an Agilent Technologies 1260 Infinity HPLC system (USA) equipped with a ZORBAX 300SB-C18 column (4.6 × 250 mm, 5 μm). The mobile phase consisted of 0.1% TFA aqueous solution and ACN with gradient elution. The UV detector was set at 250 nm, with an injection volume of 20 μL. The column temperature was maintained at 25 °C and the flow rate was 1 mL min⁻¹.

For sample preparation, 200 μL of micelle solution was accurately pipetted and centrifuged at 1000 rpm for 10 min in a high-speed refrigerated and centrifuged to precipitate free drugs. Then, 100 μL of supernatant was collected and mixed with 500 μL of ACN, followed by vortexing for 20 s to ensure complete dissolution. According to the established HPLC analytical method, 20 μL of the sample was injected for analysis. The peak area was measured and substituted into the DNS–DYE standard curve to calculate the concentration of DNS–DYE in micelles. Each sample was analyzed in triplicate and the average value was taken. The encapsulation efficiency (EE) and drug loading capacity (DL) were calculated using the following formulas:

2.9.3 Stability assessment. Micelle stability was evaluated by monitoring hydrodynamic diameter changes during storage at 4 °C. DLS measurements were conducted at predetermined intervals (days 1, 3, 5, 7, 9, 11 and 15) with simultaneous visual inspection for precipitation or aggregation.

2.10 Physiological toxicity evaluation of DNS–DYE/PEG–NI copolymer micelles

In vitro cytotoxicity evaluation of the system on HEK-293 cells was performed using the CCK-8 assay. HEK-293 cells were seeded in 96-well plates and incubated for 24 h with complete medium containing DNS–DYE/PEG–NI micelles at concentrations of 0, 2.91, 7.81, 15.63, 31.25, 62.50, 125.00, and 250.00 μg mL⁻¹. After removing the drug-containing medium, the cells were treated with fresh complete medium containing 10% CCK-8 reagent and incubated for 0.5 h to allow color development. The absorbance of each well at 450 nm was measured using a microplate reader.

2.11 In vitro hypoxic condition drug release study of PEG–NI

The release behavior of pyrene-loaded PEG–NI micelles was investigated using a dialysis method under in vitro conditions. The experiment was conducted with two groups: a hypoxic group and a normoxic group. Both groups employed 14 mL of release medium consisting of 200 μM NADPH, 0.1% Tween80, and 20 μM HEPES in buffer solution. A 3.5 kDa cellulose dialysis bag containing 1 mL of pyrene/PEG–NI copolymer micelle solution was immersed in the release medium.

For the hypoxic group, 100 μL of liver microsomal enzyme solution (20 mg mL⁻¹) was added to the dialysis bag, followed by nitrogen purging for 1 min to establish hypoxia. The normoxic group received no additional treatment. Both groups were incubated at 37 °C under constant agitation (90 rpm) in a water bath shaker.

At predetermined time intervals (1, 2, 4, 6, 8, and 12 h), 300 μL of release medium was carefully sampled from each group and replaced with an equal volume of fresh, pre-warmed release medium to maintain sink conditions. Prior to each sampling in the hypoxic group, the medium was purged with nitrogen for 1 min to preserve the hypoxic environment.

The concentration of pyrene in the collected samples was quantified using the above-mentioned HPLC system with fluorescence detection (λ_ex = 265 nm, λ_em = 394 nm). The mobile phase consisted of methanol : water (90 : 10, v/v). The injection volume was 20 μL, with the column temperature maintained at 25 °C and a flow rate of 1 mL min⁻¹. Based on the established HPLC analytical method for pyrene, the concentration of pyrene in the release medium was quantified, and the in vitro release profile of the copolymer micelles was subsequently plotted.

2.12 In vivo imaging assessment

DNS–DYE and DYE were separately dissolved in DMSO and subsequently dispersed in a solubilizing agent composed of 5% PEG_5k and 5% Tween80, yielding final concentrations of 1 mg mL⁻¹ for both compounds. Fresh DNS–DYE/PEG–NI micelles were prepared with matched probe concentrations to the free drug solutions. Following tail vein injection, in vivo luminescence signals were monitored using a small animal in vivo imaging system. Real-time NIR imaging was performed using an Olympus imaging platform to guide precise surgical resection procedures.

3 Results and discussion

3.1 Design and synthesis

3.1.1 Synthesis and characterization of DNS–DYE. The “off” state probe DNS–DYE was synthesized based on the previous development of DYE. 5 Building upon this work, we selected DNS as the fluorescence-quenching moiety for DYE. The quenching mechanism operates through two distinct pathways: (1) the strong electron-withdrawing nature of DNS, when conjugated to DYE via a phenylsulfonyl ester bond, significantly reduces the electron density of the fluorophore, thereby suppressing fluorescence emission at the characteristic excitation wavelength of 679 nm; (2) the nitro groups in DNS effectively absorb photons emitted by DYE upon excitation, providing an additional quenching pathway. The DNS–DYE used in this work was kindly provided by one of our collaborators. Its detailed synthesis procedure, along with characterization data by mass spectrometry and ¹H NMR spectroscopy, can be found in ref. 24.

3.1.2 Synthesis and characterization of PEG–NI. The hypoxia-responsive polymer PEG–NI was designed as an amphiphilic triblock copolymer featuring a PEG_5k hydrophilic segment and a pLys hydrophobic backbone.²⁵ This design incorporates NI groups as hypoxia-responsive moieties while maintaining excellent biocompatibility through the pLys structure.

The synthesis proceeded through two key steps. First, PEG–pLys was prepared via ring-opening polymerization, with its structure verified by ¹H NMR spectroscopy (Fig. S1, ESI†). Subsequently, the reactive intermediate PNP–NI was synthesized to enhance the conjugation efficiency of NI groups. Final coupling of PNP–NI with deprotected PEG–pLys yielded the target polymer PEG–NI, whose successful formation was confirmed by characteristic signals in the ¹H NMR spectrum (Fig. S2, ESI†).

3.2 Characterization

3.2.1 The GSH-responsive “off-to-on” property of DNS–DYE. Fig. 2a illustrates the GSH-responsive activation mechanism of DNS–DYE. Building upon the established literature demonstrating DNS's selective recognition of GSH,²³ our experiments confirmed that in the reducing tumor microenvironment, the nucleophilic thiolate anion of GSH attacks the DNS moiety, cleaving the benzenesulfonyl ester bond. This reaction releases both the electron-withdrawing DNS group and the fluorescent DYE component.

Fig. 2 (a) Activation principle of DNS–DYE by GSH; (b) fluorescence spectrum of DNS–DYE incubated with varying concentrations of GSH (λ_ex = 679 nm); (c) fluorescence response of DNS–DYE within MDA-MB-231/luci cells.

Fluorescence spectral analysis (Fig. 2b) provided quantitative validation of this mechanism. At 679 nm excitation, DNS–DYE showed no emission in the absence of GSH due to efficient quenching by the DNS moiety. However, progressive addition of GSH (0–10 mM) induced dose-dependent fluorescence recovery at 740 nm, confirming successful “off-to-on” switching. The 35-fold signal enhancement at physiological GSH concentrations (10 mM) demonstrates excellent activation sensitivity.

The cellular-level experimental results further confirmed the fluorescence response performance of this probe. Considering that the total cellular uptake increases with the prolonged incubation time, a standardized 1 h incubation period was implemented to control the uptake amount. Subsequent culture durations were varied exclusively to investigate fluorescence intensity changes induced by metabolic processes. The results demonstrated that the fluorescence intensity of DNS–DYE increased progressively with metabolic time, reaching maximum brightness at 8 h (Fig. 2c). This indicates that DNS–DYE achieves “off-to-on” fluorescence activation within tumor cells.

To further validate the responsiveness of the probe to tumor cells, we conducted experiments with varying numbers of MDA-MB-231 triple-negative breast cancer cells. The near-infrared probe was pre-added to cell culture dishes and incubated with different cell counts. After 12 h of incubation, we observed a significant increase in fluorescence intensity as the number of tumor cells increased (Fig. S3, ESI†). This trend indicates that the tumor cells effectively metabolize and activate the nanoprobe, leading to enhanced fluorescence signals. However, when the cell number reached 5 and 10 ten thousand, the fluorescence intensity showed no significant difference. This plateau in fluorescence suggests that the probe was fully consumed at these higher cell densities, implying that the probe has a limited capacity for activation per unit area under cell culture conditions. These results not only confirm the tumor cell response of the probe but also provide insights into its metabolic dynamics and activation efficiency in vitro.

3.2.2 Characterization of the hypoxia-responsive switching properties of PEG–NI. In the hypoxic tumor microenvironment, the overexpressed nitroreductase can catalyze the reduction of NI to aminoimidazole (Fig. 3a). To simulate hypoxic conditions in vitro, sodium dithionite (Na₂S₂O₄), a strong reducing agent, was employed to rapidly deplete oxygen in the system, thereby inducing chemical hypoxia.²⁶ As reported in the literature, NI derivatives should exhibit a significant shift in the UV absorption wavelength before and after the reaction (from 325 nm to 290 nm).²⁶ Thus, the reaction progress was monitored using UV spectrophotometry (Fig. 3b).

Fig. 3 (a) Hypoxia-triggered structural conversion of PEG–NI; (b) UV-vis absorption spectra of the PEG–NI polymer before and after reaction with Na₂S₂O₄ mimicking the hypoxia tumoral microenvironment.

Due to the poor water solubility of PEG–NI, empty micelles were prepared to facilitate its reaction with Na₂S₂O₄ in aqueous solution. A characteristic UV absorption peak of PEG–NI empty micelles was observed at 308 nm. After Na₂S₂O₄ treatment, this peak disappeared, and a new peak emerged at 292 nm, confirming the conversion of the NI moiety in the polymer to aminoimidazole.

3.2.3 Construction and characterization of the surgical navigation system. The amphiphilic block copolymer PEG–NI was thoroughly characterized for its micelle-forming properties. Pyrene fluorescence probe spectroscopy measurements identified a critical micelle concentration of 21 μg mL⁻¹, demonstrating efficient self-assembly behavior. This capability stems from the hydrophobic pLys and NI segments, which engage in π–π stacking and other intermolecular interactions with the probe molecules. These interactions facilitate the formation of structurally stable micelles capable of achieving both high drug loading and encapsulation efficiency, the key characteristics for effective surgical navigation applications.

A systematic investigation of formulation parameters focused on optimizing the mass ratio between polymer and probe components. DLS provided precise measurements of hydrodynamic diameter, while HPLC enabled accurate quantification of encapsulation efficiency and drug loading capacity. Our comprehensive characterization protocol identified 10 : 1 as the optimal polymer-to-probe mass ratio. TEM images revealed that the resulting DNS–DYE/PEG–NI micelles exhibit a well-defined spherical morphology with uniform sizes ranging from 30–50 nm (Fig. 4a). The DLS-measured hydrodynamic diameter of 64.43 ± 4.89 nm (Fig. 4b) showed the expected increase compared to TEM measurements, reflecting the presence of a hydration layer around the micelles in aqueous solution. The exceptionally low polydispersity index (PDI = 0.043) confirmed the highly uniform size distribution within the colloidal system. HPLC quantification demonstrated excellent EE (80.00 ± 5.82%) and DL (7.43 ± 0.45%), verifying successful incorporation of the probe molecules into the polymeric micellar matrix.

Fig. 4 (a) TEM image of micromorphology, (b) DLS result, (c) stability assessment results and (d) safety evaluation of DNS–DYE/PEG–NI micelles using HEK 293 cells.

To assess the long-term colloidal stability of the DNS–DYE/PEG–NI micelles, we conducted systematic size and dispersion analyses during storage at 4 °C. Over a 15-day observation period, the micelles exhibited excellent stability, with hydrodynamic diameters showing only a modest increase from 62 nm to 70 nm (Fig. 4c). Importantly, this size variation represented just a 12.9% change from the initial measurement, well within acceptable limits for pharmaceutical applications. Throughout the storage period, the system maintained remarkable dispersion stability, as evidenced by the consistently low polydispersity index (PDI < 0.1) and the complete absence of visible precipitation. Furthermore, no macroscopic aggregation or phase separation was observed, confirming the preservation of structural integrity in the amphiphilic polymer–probe assembly under refrigerated storage conditions. These results collectively demonstrate that the micellar system exhibits sufficient stability for practical clinical applications requiring extended storage.

3.2.4 Safety investigation of surgical navigation systems. The biocompatibility of the polymeric micelles was rigorously evaluated to ensure their suitability for surgical navigation applications. Using HEK-293 renal epithelial cells as a model system, acute toxicity assessments were performed to determine the concentration-dependent cytotoxicity profile. The CCK-8 assay revealed excellent biocompatibility at clinically relevant concentrations, with cell viability remaining above 90% of control values for micelle concentrations up to 125 μg mL⁻¹ after 24 h of exposure (Fig. 4d). Notably, statistically significant cytotoxicity (p < 0.05) was only observed at concentrations exceeding 250 μg mL⁻¹, establishing a wide therapeutic window (0–125 μg mL⁻¹) that exceeds typical requirements for surgical guidance applications. This concentration-dependent toxicity pattern is consistent with established safety profiles of amphiphilic polymer-based delivery systems, further validating the clinical potential of our micellar formulation.

3.2.5 Investigation of probe release under hypoxic conditions in vitro. The hypoxia-responsive behavior of the micellar system was systematically validated through comparative studies of empty and probe-loaded micelles. When exposed to Na₂S₂O₄ (100 mM)-induced chemical hypoxia (mimicking the in vivo TME hypoxia), empty micelles underwent dramatic structural changes, as evidenced by hydrodynamic diameter measurements showing an increase from 52.93 ± 20.39 nm to 398.9 ± 118.5 nm (Fig. 5a). This remarkable size expansion clearly demonstrates the reductive cleavage of hypoxia-sensitive bonds in the polymer backbone, leading to complete nanostructure disassembly and subsequent payload release.

Fig. 5 (a) Hydrodynamic diameter variation of PEG–NI empty micelles before and after Na₂S₂O₄ treatment; (b) cumulative release profiles of pyrene/PEG–NI under different conditions.

To more physiologically recapitulate tumor hypoxia, we developed an in vitro release system incorporating hepatic microsomal enzymes, NADPH, and nitrogen purging, with appropriate normoxic and enzyme-free controls.²⁶ Pyrene was employed as a model probe due to its exceptional chemical stability, allowing specific assessment of micelle disassembly without confounding effects from probe degradation. Following preparation via thin-film hydration, pyrene-loaded PEG–NI micelles showed substantially enhanced release kinetics under hypoxic conditions compared to normoxic controls (Fig. 5b), as quantified by HPLC analysis. These results collectively confirm the robust hypoxia-responsive drug release capability of our micellar system.

3.3 Fluorescence-guided tumor resection in vivo

The free “off-state” probe DNS–DYE, the free “on-state” probe DYE, and micelles loaded with equivalent molar amounts of probes (DNS–DYE/PEG–NI and DYE/PEG–NI) were administered to tumor-bearing nude mice via tail vein injection. In vivo fluorescence was monitored at 0.5, 1, 2, 4, 8 and 12 h post-injection using an IVIS Spectrum system. Regions of interest (ROIs) were drawn over tumors and normal tissues (normally surrounding skin) to quantify signal-to-background ratios. The in vivo fluorescence was monitored using IVIS at different time points post-injection, as shown in Fig. 6a.

Fig. 6 (a) In vivo fluorescence imaging of and (b) real-time NIR fluorescence imaging-guided surgery in MDA-MB-231/luci tumor-bearing mice. (c) The surgical procedure (the arrows indicate the tumor site): (I) before resection; (II) after resection; (III) bright-field image of the excised tumor mass and (IV) IVIS image of the excised tumor mass.

During the initial phase post-injection, the free DNS–DYE group exhibited weaker overall fluorescence intensity compared to the free DYE group. By 4 h post-injection, however, the fluorescence intensity of the DNS–DYE group surpassed that of the DYE group. This may result from a time-dependent fluorescence enhancement of DNS–DYE concurrent with metabolic clearance. Notably, although DNS–DYE was rapidly cleared, tumor-background fluorescence differentiation emerged early, indicating partial probe specificity.

For DNS–DYE/PEG–NI micelles, fluorescence initially appeared in the liver, likely due to its abundant blood supply and high levels of reduced glutathione and enzymes. Compared to free probes, micelles demonstrated significantly prolonged circulation time. During systemic circulation, micelles gradually accumulated in tumors via the EPR effect, with observable tumor fluorescence at 12 h post-injection, while fluorescence in other tissues diminished substantially. These results confirm that micellar formulation not only improves water solubility and circulation time but also enhances tumor accumulation, achieving high tumor-to-background signal ratios at specific time points.

In contrast, significant differences were observed in the in vivo distribution of the DYE/PEG–NI group. Although some tumor distribution was exhibited during the initial injection phase, the probes were rapidly quenched—even more so than the DYE–DNS group alone. It is speculated that since this group is always-ON, activation or interaction with the tumor is not required for fluorescence emission under laser irradiation. Given that the probes operate in the near-infrared range, quenching by the excitation light may have occurred. Thus, while accumulation in the tumor via the EPR effect cannot be ruled out, rapid photobleaching during multiple detection cycles (due to the absence of a silent-state protection mechanism and tumor microenvironment-specific activation) was likely. This characteristic—susceptibility to photobleaching—is considered disadvantageous for surgical navigation applications, despite potentially facilitating in vivo probe clearance.

A preliminary exploration of DNS–DYE/PEG–NI for fluorescence-guided tumor resection was conducted (Fig. 6b). Using an Olympus real-time NIR imaging system (limited to tumor-adjacent fields), relative fluorescence intensities were normalized per mouse. Three groups received free DNS–DYE solution, DNS–DYE/PEG–NI micelles (equivalent probe concentration), or saline via tail vein injection. At 4–6 h post-injection, tumors were resected under real-time imaging. Saline controls showed no fluorescence; free DNS–DYE exhibited weaker tumor fluorescence than peri-tumor regions due to rapid clearance, while micelles displayed the strongest tumor fluorescence. Intriguingly, in one micelle-treated mouse, strong fluorescence initially localized to a visible tumor mass revealed an additional deeper, visually undetectable tumor upon surgical exposure, further validating the system's utility for guiding tumor resection.

Fig. 6c illustrates the surgical procedure (the arrows indicate the tumor site). It can be clearly observed that the fluorescence intensity at the tumor site (indicated by arrows) significantly changes before and after surgery. After excluding nonspecific fluorescence interference (including autofluorescence, blood, and other common sources), tumor resection was performed based on the fluorescence-defined boundaries (including depth). Post-resection, the fluorescence intensity at the original tumor site markedly decreased. Furthermore, IVIS imaging of the freshly excised tumor mass revealed strong fluorescence emission (DYE) across the entire tissue, further confirming the accuracy of our surgical resection margins.

3.4 Clinical translation and scalability considerations

Our micellar design builds upon the foundational principles of stimuli-responsive polymers, as exemplified by the pH-sensitive systems, while introducing dual activation to address tumor heterogeneity. While the dual-responsive nanoprobe demonstrates compelling preclinical performance, its clinical translation hinges on addressing scalability challenges. The current synthesis involves multi-step organic reactions (e.g., Lys–NCA preparation, PNP–NI conjugation), which may incur high costs at industrial scales. To mitigate this, future efforts could explore automated flow chemistry platforms or alternative hypoxia-responsive motifs like azobenzene derivatives to simplify synthesis. Batch-to-batch reproducibility must also be rigorously evaluated, given the sensitivity of micelle size and drug-loading efficiency to polymerization conditions.

From a clinical workflow perspective, the nanoprobe's 12-h activation window aligns with typical surgical timelines, enabling preoperative administration. Its prolonged tumor retention (>24 h) could further support staged resections or re-excision of positive margins. Compared to FDA-approved ICG, our system offers two key advantages: (1) tumor-specific activation via dual TME triggers, reducing false-positive signals in inflamed tissues; and (2) resistance to aggregation-induced quenching, ensuring stable intraoperative imaging. However, long-term toxicity studies and GMP-compliant manufacturing protocols remain critical milestones before phase I trials.

4 Conclusion

Near-infrared fluorescence imaging has attracted significant attention in recent years due to its advantages of high spatial resolution, strong tissue penetration, low background signal, and minimal scattering. As near-infrared imaging systems are simpler than MRI and other modalities, they can be readily integrated with existing surgical equipment. Moreover, near-infrared imaging does not require interruption of surgical procedures, making near-infrared probes a highly promising direction for developing surgical navigation systems. Building on prior research, this study further proposes a TME-responsive nanoprobe strategy. Structural modifications were applied to the near-infrared probe DYE to confer GSH responsiveness within tumors, while the hypoxia-responsive polymer backbone also reacts to tumor microenvironmental cues. Upon entering tumors, these dual-response micelles release probes under the dual activation mechanism, emitting near-infrared fluorescence via an “off-to-on” transition, thereby holding potential as a surgical navigation system for precise tumor resection. In summary, while our study highlights the promising clinical potential of the DNS–DYE/PEG-NI nanoprobe, further research and development efforts are needed to address scalability challenges and fully realize its applications in clinical practice. We believe that through continued innovation and collaboration across academic and industrial sectors, this nanoprobe could become a valuable tool in the fight against cancer, improving surgical outcomes and patient quality of life.

Author contributions

JS, JY and HZ conceived the project, carried out the research and composed the paper. WT assisted in the characterization of the compounds. JS, WT and BY carried out the synthesis of the compounds. YG and SW carried out the TEM studies. HZ supervised the research work.

Conflicts of interest

There are no conflicts to declare.

Data availability

The data supporting the findings of this study are available from the corresponding author on request.

Acknowledgements

We gratefully acknowledge the financial support provided by Zhiyuan Biotechnology under the project Testing Services for Functional Biofluids.

Notes and references

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Footnote

† Electronic supplementary information (ESI) available. See DOI: https://doi.org/10.1039/d5tb01157d

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