NIR-IIb fluorescence ratiometric imaging of tumor-associated neutrophils for immunotherapy efficacy monitoring and strategy optimization

Li, Yang; Lin, Jinyan; Zou, Jianhua; Zeng, Yongyi; Liu, Xiaolong

doi:10.1038/s41467-025-65927-1

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Published: 05 December 2025

NIR-IIb fluorescence ratiometric imaging of tumor-associated neutrophils for immunotherapy efficacy monitoring and strategy optimization

Nature Communications volume 16, Article number: 10880 (2025) Cite this article

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Abstract

Imaging the immune activation status of tumor-associated neutrophils (TANs) provides insights into cancer immunotherapeutic strategy improvement. Here we show a ratiometric nanoprobe responding to tumor microenvironment and enabling NIR-IIb quantitative fluorescence imaging of TANs with overexpression of neutrophil elastase (NE). The ratiometric nanoprobe integrates erbium-based nanoparticles (ErNPs) with NE-cleavable peptide-caged near-infrared-II (NIR-II) fluorescence dye. Upon systemic delivery, the nanoprobe actively targets tumor microenvironment, where NE cleaves the caging peptide to trigger energy transfer within ErNPs under 808 nm excitation, yielding enhanced NIR-IIb fluorescence emission at 1532 nm. In contrast, the NE-independent fluorescence of ErNPs at 1532 nm remains stable when excited at 980 nm, providing a self-calibrated signal. Across Hepa1-6 and H22 tumor models during immunotherapy, the nanoprobe translates TANs’ phenotype shift into precise ratiometric signals, enabling quantitative dynamic imaging of TANs’ immune status changes. Subsequently, the responders to immunotherapy among mice could be effectively identified by fluorescence ratiometric imaging of TANs. This strategy offers a non-invasive way to track immune status in real time and could guide timely adjustments to immunotherapy strategies, potentially improving outcomes for patients.

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Introduction

Immunotherapy represents one of the most promising approach in contemporary oncology by activating the host immune system to elicit robust and sustained anti-tumor responses^1,2. However, its clinical application is hampered by the prolonged treatment duration and considerable interpatient variability in response to identical regimens^3,4,5,6. More critically, there is currently a lack of effective methods to assess immunotherapy efficacy in the early stage. Therefore, there is an urgent need to develop tools capable of evaluating early treatment effects, with the aim of predicting therapeutic outcomes and facilitating timely, dynamic adjustments to treatment strategies⁷.

Imaging the immune activation status of tumor-associated neutrophils (TANs) provides significant insights into cancer immunotherapy strategy optimization^8,9. As essential components of tumor immune microenvironment, TANs are broadly categorized into anti-tumor “N1” and pro-tumor “N2” phenotypes, which play critical roles in tumor progression and immunotherapy response^10,11. N2-TANs can secrete neutrophil elastase (NE), a serine protease that can decompose elastin, collagen, and fibronectin, resulting in the promotion of tumor angiogenesis, invasion and metastasis¹². Consequently, NE is regarded a pivotal biomarker of N2-TANs, serving as a molecular target for visualizing its immune status changes^13,14. Theoretically, NE expression will be significantly down-regulated after effective immunotherapy, otherwise, it may remain unchanged or be up-regulated^13,14. Therefore, dynamic monitoring of TANs’ immune activity based on the expression of NE can provide a critical basis for assessing immunotherapy efficacy, predicting outcomes, and optimizing therapeutic strategies.

Tissue biopsy and peripheral blood analysis are traditional techniques for detecting TANs in clinical cancer diagnosis. Both methods, however, are invasive and provide only limited, static, and slow information, thereby limiting their utilization in precise TANs monitoring^15,16. Consequently, there is an urgent need for the development of non-invasive and dynamic tools for in vivo TANs analysis. Traditional imaging modalities, such as computed tomography (CT), positron emission tomography (PET), and magnetic resonance imaging (MRI) could offer non-invasive and dynamic diagnostic abilities for tumor tissues, but they have limited ability to detect signals at the cellular level due to poor molecular sensitivity and specificity^17,18. Molecular fluorescence imaging could enable the dynamic visualization and quantification of immune cells with high sensitivity and specificity^{18,19,20,21,22,23,24,25}. Fluorescence probes have been recently utilized for TANs imaging^20,26,27. For example, Lin and co-workers developed a series of NE-activatable fluorescence probes by conjugating diverse fluorinated amide chains onto hemicyanine dye, which enables NIR-I fluorescence imaging of TANs for early tumor detection²⁰. Pu and co-workers also developed an NE-activatable fluorescence nanoprobe based on “semiconductor polymer-hemicyanine dye-peptide” conjugate, achieving quantitative fluorescence/photoacoustic imaging of TANs²⁷. However, these probes are limited to first near-infrared window (NIR-I, 700-900 nm) fluorescence with shallow penetration depth and substantial autofluorescence interference, restricting their in vivo imaging applications. The second near-infrared window (NIR-II, 1000-1700 nm) fluorescence imaging offers advantages such as longer emitting wavelengths, lower photon absorption and scattering in biological tissues, deeper tissue penetration, lower background autofluorescence, and higher temporal and spatial resolution, thus being a more promising alternative^28,29,30. Additionally, fluorescence probes relying on single-wavelength fluorescence intensity variations are inherently susceptible to various factors, including probe distribution heterogeneity, tissue depth-dependent signal attenuation, instrumentation, and environmental parameters, which hinder quantitative imaging under complex physiological conditions. Therefore, there remains a pressing need for NIR-II fluorescence probes and multi-wavelength imaging technologies that enable interference-resistant, precise, and quantitative TANs imaging. Recently, fluorescence ratiometric imaging probes have demonstrated improved accuracy, reliability, and reproducibility in quantitative imaging through the introduction of self-calibrating capability with a reference signal. This strategy eliminates dependence on single-wavelength fluorescence signal variations and alleviates interference from confounding variables, achieving more accurate quantitative imaging^31,32,33. Therefore, our goal is to develop activatable NIR-II fluorescence ratiometric probes by integrating NIR-II fluorescence imaging modality with ratiometric imaging technology, enabling high-precision in vivo quantitative imaging of TANs for subsequent applications in monitoring immunotherapy efficacy and optimizing immunotherapy strategies.

Inspiringly, rare-earth nanoprobes possess characteristics such as tunable NIR-II wavelengths, multichannel emission, sharp emission bands, excellent photostability, and low toxicity⁷. Several ratiometric rare-earth fluorescent probes based on aggregation-caused emission enhancement (ACIE) processes or Förster resonance energy transfer (FRET) pathway have been developed for quantitative imaging of ROS, pH, and other biomarkers in the tumor microenvironment^34,35. Consequently, the NIR-II fluorescence ratiometric imaging technology with rare-earth nanoprobes is expected to enable ultrasensitive, precise, and quantitative imaging of TANs’ function information within tumor microenvironment.

In this work, we show a NIR-IIb fluorescence ratiometric nanoprobe for quantitative visualization of TANs in vivo, as explicitly depicted in Fig. 1. The nanoprobe is prepared by introducing a NE-cleavable peptide-caged NIR-II fluorescence dye (custom-synthesized benzo[cd]indole hemicyanine, Bi-HCy-OH) and a cRGD-targeting polymer network on the surface of erbium-based nanoparticle (ErNPs). The NE-cleavable peptide-caged Bi-HCy-OH (Bi-HCy-valine-proline-alanine-alanine-N-acetyl, Bi-HCy-VPAA-Ac) absorbed on the surface of ErNPs exhibit no fluorescence due to intramolecular charge transfer (ICT) effect. In the presence of NE, Bi-HCy-VPAA-Ac molecule reacts with NE to form Bi-HCy-OH, which results in the fluorescence recovery. Under excitation at 808 nm, Bi-HCy-OH absorbs the incident laser and transferred the excitation energy to ytterbium ion (Yb³⁺) of ErNPs via FRET pathway, subsequently achieving the dye sensitization effect of ErNPs, and finally emitting amplified NIR-II fluorescence signal at 1532 nm (defined as F_{1532em, 808ex}). In addition, 980 nm laser cannot excite Bi-HCy-OH molecules, but can excite ErNPs directly. Therefore, 1532 nm emission excited at 980 nm can be used as the reference signal (defined as F_{1532em, 980ex}). The fluorescence intensity at 1532 nm excited at 808 nm is responsive to NE concentration, and fluorescence intensity ratio (F_{1532em, 808ex})/(F_{1532em, 980ex}) significantly change before and after NE response. The Bi-HCy-ErNPs nanoprobes can possess a ratiometric response for NE with good linear correlation. Therefore, our NIR-IIb fluorescence ratiometric nanoprobes can precisely and quantitatively visualize TANs in vivo, thus reflecting the immune status change, monitoring immunotherapy efficacy, and distinguishing between the immunotherapy-irresponsive and immunotherapy-responsive individuals, thus providing the opportunity to optimize immunotherapy strategy for personalized cancer immunotherapy.

**Fig. 1: Schematic illustration of the study design.**

Results

Synthesis and characterization of NE-responsive NIR-II fluorescence probes

The custom-synthesized NIR-II fluorescence dye, Bi-HCy-OH (benzo[cd]indole hemicyanine), was designed and synthesized through a series of reactions, utilizing commercial indole as the starting fluorophore (Supplementary Fig. 1). Then the Bi-HCy-VPAA-Ac was designed and synthesized between Bi-HCy-OH as a fluorophore and p-aminobenzyl alcohol-VPAA-Ac (pABA-VPAA-Ac) as a recognizer towards NE biomarker (Supplementary Fig. 1a). The chemical structures of Bi-HCy-OH, pABA-VPAA-Ac, and Bi-HCy-VPAA-Ac were verified by proton nuclear magnetic resonance (¹H NMR) spectra, ¹³C NMR spectra, and mass spectrometry (MS) analysis (Supplementary Figs. 2–13 and Supplementary Data 1–25).

To explore the NE responsiveness of Bi-HCy-VPAA-Ac, we tested the absorption and fluorescence emission spectra of Bi-HCy-OH, Bi-HCy-VPAA-Ac, and Bi-HCy-VPAA-Ac plus NE. Upon NE addition, the absorption peak of Bi-HCy-VPAA-Ac red-shifted from ~665 nm to ~794 nm, mirroring the spectrum of Bi-HCy-OH (Fig. 2a). Compared to Bi-HCy-VPAA-Ac alone, the emission peak of Bi-HCy-VPAA-Ac plus NE significantly increased by 15.0-fold at ~962 nm, reaching levels comparable to Bi-HCy-OH (Fig. 2b). Furthermore, the conversion of Bi-HCy-VPAA-Ac into Bi-HCy-OH triggered by NE was proved by mass spectra (Fig. 2c–e, Supplementary Figs. 12–13, Supplementary Data 1-4, and Supplementary Data 21-25).

**Fig. 2: NE responsiveness of Bi-HCy-VPAA-Ac and characterization of Bi-HCy-ErNPs.**

To investigate the responsive mechanism of Bi-HCy-VPAA-Ac towards NE, we calculated the highest occupied molecular orbital (HOMO) and lowest unoccupied molecular orbital (LUMO) energy for Bi-HCy-OH and Bi-HCy-VPAA-Ac (Fig. 2f). The bandgap between the LUMO and HOMO of Bi-HCy-OH ( ~ 1.34 eV) is narrower compared to Bi-HCy-VPAA-Ac (~2.05 eV), suggesting that the electron donor (4-vinyl-2,3-dihydro-1H-xanthen-6-ol) in Bi-HCy-OH possesses a stronger “push-pull” electron system and electron-supplying capacity than that (1-(2-(2-acetamidopropanamido)propanoyl)-N-(3-methyl-1-oxo-1-((4-(((4-vinyl-2, 3-dihydro-1H-xanthen-6-yl)oxy)methyl)phenyl)amino)butan-2-yl)pyrrolidine-2-carboxamide) in Bi-HCy-VPAA-Ac. This could result in a stronger ICT effect to red-shift the absorption and emission peaks for Bi-HCy-OH compared to those for Bi-HCy-VPAA-Ac.

Synthesis and characterization of NIR-IIb fluorescnece ratiometric nanoprobes

To establish the FRET pathway, we initially adopted core-shell ErNPs, NaYbF₄: Er, Ce@NaYF₄: Yb, Lu, capable of energy transfer from dye to Yb³⁺ and subsequently to Er³⁺. Initially, NaYbF₄: Er, Ce core nanocrystals were synthesized via a typical co-precipitation method, then the NaYF₄: Yb, Lu shell was warped on the core surface by seed-mediated shell growth approach. Subsequently, the Bi-HCy-VPAA-Ac dye, with hydrophobic aromatic rings, was absorbed on the surface of ErNPs within a hydrophobic cavity created by oleic acid/1-octadecene ligands. Lastly, the cRGD-targeting and interpenetrating polymer network was further functionalized on the surface of dye-sensitized NaYbF₄: Er, Ce@NaYF₄: Yb, Lu through hydrophobic interactions and successive 1-(3-dimethylaminopropyl)−3-ethylcarbodiimide hydrochloride (EDC) chemistry between PMH/8Arm-PEG-NH₂, 8Arm-PEG-NH₂/PAA, and PAA/“DSPE-PEG-NH₂/8Arm-PEG-NH₂”, forming the final Bi-HCy-ErNPs nanoprobes. This surface functionalization could be expected to improve the nanoprobes’ stability and biocompatibility, enhance tumor accmulation, and protect the nanoprobes from degradation and aggregation, thus optimizing imaging performance. The synthesis of both ErNPs and Bi-HCy-ErNPs nanoparticles were verified by transmission electron microscope (TEM) images, high-resolution TEM (HRTEM) images, fast Fourier Transform (FFT) analysis, and selected area electron diffraction (SAED) patterns (Fig. 2g and Supplementary Fig. 14). Specifically, HRTEM images revealed distinct lattice stripes, with a lattice spacing of 2.18 Å for the core and 0.87 Å for the shell, proving the formation of core-shell structure of ErNPs. The high-angle annular dark field-scanning transmission electron microscopy (HAADF-STEM) and STEM-energy dispersive X-ray (STEM-EDX) elemental mapping images revealed the formation of ErNPs, with homogeneous distribution of Yb, Er, Ce, Lu, Y, Na, and F elements within the spatially defined regions, (Fig. 2h). Moreover, the introduction of both O and P within structure indicated the formation of Bi-HCy-ErNPs. The average hydrodynamic particle sizes of both ErNPs and Bi-HCy-ErNPs, determined by dynamic light scattering (DLS), were ~21.68 nm and ~29.69 nm, respectively (Fig. 2i). The powder X-ray diffraction (PXRD) patterns of both ErNPs and Bi-HCy-ErNPs presented their α-phase structure of NaYF₄/NaYbF₄ (Fig. 2j), which was conducive to efficient fluorescence emission at a low crystallographic symmetry. The fluorescence lifetime of Bi-HCy-OH and Bi-HCy-ErNPs was determined as ~1.43 and ~0.72 ns, respectively (Fig. 2k). This indicated the energy transfer from Bi-HCy-OH to ErNPs with an estimated energy transfer efficiency of ~49.65%. The X-ray photoelectron spectroscopy (XPS) verified the composition of Yb, Er, Ce, Lu, Y, Na, F, O, and P elements in the Bi-HCy-ErNPs nanoprobes (Supplementary Fig. 15).

To verify the feasibility of sensitization effect of Bi-HCy-OH towards ErNPs, we investigated their absorption and emission spectra. The absorption of ErNPs at 808 nm was appreciably weak, leading poor excitation efficiency at this wavelength (Fig. 3a). Consequently, we selected Bi-HCy-OH as a fluorophore for synthesizing NE-responsive Bi-HCy-VPAA-Ac, with the aim of sensitizing ErNPs. This selection was based on Bi-HCy-OH’s robust emission peak at ~962 nm (Fig. 3b), which aligned well with absorption peak of Yb³⁺ in ErNPs (Fig. 3c). This could help energy transfer from Bi-HCy-OH to Er³⁺ through Yb³⁺ underlying the FRET mechanism (Fig. 3d-f). With Lu³⁺-assisted dye-triplet-sensitization, ErNPs could emit intensified fluorescence at 1532 nm upon 808 nm excitation (Fig. 3f and Supplementary Fig. 16). Notably, the heavy atom Lu played a pivotal role in the formation of Bi-HCy-ErNPs, and its introduction into Bi-HCy-ErNPs was critical for intensifying fluorescence (Supplementary Fig. 16). After Bi-HCy-OH was absorbed onto the surface of ErNPs, the ErNPs exhibited a remarkable increase in fluorescence intensity when excited at 808 nm, but the fluorescence intensity remained constant when excited at 980 nm (Fig. 3g, h). This further proved the successful sensitization effect of Bi-HCy-OH towards ErNPs at 808 nm excitation, establishing a foundation for designing of NE-responsive NIR-IIb fluorescence ratiometric nanoprobes. Upon NE addition to Bi-HCy-VPAA-Ac, a newly red-shifted absorption peak emerged at ~794 nm, would be intensified, while the initial absorption peak at ~665 nm would be decreased (Fig. 3i). The emission peak at ~948 nm underwent a slight shift to ~962 nm, accompanied by a significant intensification (Fig. 3j). Consequently, upon NE triggering, the emission peak of Bi-HCy-ErNPs at 1532 nm increased significantly when excited at 808 nm, while keeping stable at 980 nm excitation (Fig. 3k-l). Moreover, this phenomenon exhibited dynamics dependent on both incubation time (Supplementary Fig. 17) and NE concentration (Fig. 3m, n). The NIR-II fluorescence ratiometric signals was linearly proportional to the NE concentration ranging from 0 to 1.6 μM (Fig. 3o, p), indicating that the Bi-HCy-ErNPs nanoprobes possessed high sensitivity for quantitative NE imaging, with a limit of detection (LOD, 3σ/slope) of ~0.37 ng/mL. Additionally, the nanoprobes demonstrated exceptional particle size stability, dye-loading stability, and optical stability in physiological environment including 10% FBS over time (Supplementary Figs. 18-22).

**Fig. 3: NE responsiveness of Bi-HCy-ErNPs.**

Selective response of NIR-IIb fluorescence ratiometric nanoprobes towards NE

To assess the selective response of Bi-HCy-ErNPs nanoprobes towards NE, we employed the NE inhibitor (sivelestat), which molecular structure could compete with substrates to bind to the catalytic site of NE, to block their interactions³⁶, thus effectively inhibiting NE activity. Meanwhile, the cathespin S, cathespin C, cathespin G, caspase-1, caspase-3, granzyme B, proteinase 3, cathepsin B, carboxylesterase, aminopeptidase N, leucine aminopeptidase, common ions (Na⁺, K⁺, Fe²⁺, and Ca²⁺), and some small molecules (glutathione and hydrogen peroxide) were taken as the controls. Then we conducted the NIR-IIb fluorescence spectroscopy and imaging studies (Fig. 4a). Upon exposure to the controls, minimal changes were observed in both F_{1532em, 808ex} and F_{1532em, 980ex} signals (Fig. 4b–g and Supplementary Figs. 23, 24). Consequently, no significant variation was observed in the ratio of F_{1532em, 808ex}/F_{1532em, 980ex}. In contrast, upon NE triggering, a significant increase was observed in the F_{1532em, 808ex} signal, while the F_{1532em, 980ex} signal remained stable, resulting in a significant increase in the ratio of F_{1532em, 808ex}/F_{1532em, 980ex}, and such ratio signal increase can be significantly abolished by adding the NE inhibitor. Furthermore, the increase in the F_{1532em, 808ex} signal exhibited a concentration-dependent relationship with NE, as demonstrated by NIR-IIb fluorescence spectra (Fig. 4h and Supplementary Fig. 25). These findings suggested that Bi-HCy-ErNPs nanoprobes exhibited a highly selective response towards NE, enabling their effective recognition in complex biosystems.

**Fig. 4: Sensitivity and specificity of NIR-IIb fluorescence ratiometric imaging of Bi-HCy-ErNPs towards NE.**

In vitro TANs imaging using NIR-IIb fluorescence ratiometric nanoprobes

Motivated by in vitro imaging ability of Bi-HCy-ErNPs towards NE, we investigated the efficacy of Bi-HCy-ErNPs nanoprobes in detecting endogenous NE in different types of cells, including N1-neutrophils, N2-neutrophils, CD8⁺ T cells (CD8⁺ Ts), M1-bone marrow derived macrophages (M1-BMDM), NIH/3T3, and Hepa1-6 cell. Prior to in vitro imaging, we initially assessed the biocompatibility of Bi-HCy-ErNPs nanoprobes by standard 3-(4, 5-dimethyl-2-thiazolyl)-2, 5-diphenyl-2-tetrazolium bromide method. After incubating various cell types with nanoprobes for 24 h, we observed over 80% cell viability, even at the concentration of up to 1000 μg/mL, indicating the excellent biocompatibility of Bi-HCy-ErNPs (Supplementary Figs. 26, 27). Next, we utilized the nanoprobe to detect intracellular NE levels. Various cell lines were incubated in a 96-well plate and further incubated with Bi-HCy-ErNPs nanoprobes for 8 h. Subsequently, the NIR-IIb fluorescence images and signals at 1532 nm were recorded upon 808 and 980 nm excitation. The results showed that the N2-neutrophils emitted strong F_{1532em, 808ex} signals as well as robust F_{1532em, 980ex} signals, concurrently accompanied by a high ratiometric signal (Fig. 5a–c). In contrast, other cell types, when incubated with Bi-HCy-ErNPs over identical time intervals, manifested weak F_{1532em, 808ex} signals, yet still presented strong F_{1532em, 980ex} signals, along with a relatively low ratiometric signal. Furthermore, pre-exposure to the NE inhibitor sivelestat (100 μM) significantly decreased the NIR-IIb fluorescence of N2-neutrophils incubated with Bi-HCy-ErNPs at 808 nm excitation, whereas the fluorescence remained stable at 980 nm excitation (Fig. 5a–c). Consequently, the ratiometric signal was significantly decreased. These results indicated high NE-responsive selectivity of Bi-HCy-ErNPs nanoprobes towards N2-neutrophils^17,27,37. Additionally, the bio-TEM imaging revealed that N2-neutrophils incubated with Bi-HCy-ErNPs effectively internalized the probes and then distributed them within intracellular spaces (Fig. 5d).

**Fig. 5: In vitro NIR-IIb fluorescence ratiometric imaging of Bi-HCy-ErNPs.**

The effectiveness of Bi-HCy-ErNPs nanoprobes was further investigated by using NIR-IIb fluorescence microscopy. The F_{1532em, 980ex} signals exhibited similarity across all cells, whereas the F_{1532em, 808ex} signals and ratiometric signals were significantly elevated in the N2-neutrophils compared to the N1-neutrophils, NE inhibitor-treated N2-neutrophils, and other cells (Fig. 5e, f). The result was likely attributable to the interaction between Bi-HCy-ErNPs and intracellular NE, which sensitized ErNPs through the time-dependent formation of Bi-HCy-OH molecules (Fig. 5g, h and Supplementary Fig. 28). After assessment using a simple linear regression model, a remarkably strong positive correlation was found between NIR-IIb fluorescence ratiometric signal of Bi-HCy-ErNPs and specific dye-labeled antibodies towards cellular biomarkers with a correlation coefficient (R) 0.84-0.98 and a Pearson’s r value (ρ) of 0.91-0.96 (Fig. 5i). The data further proved the fluorescence activation of Bi-HCy-ErNPs required NE biomarkers from N2-neutrophils. Collectively, these results demonstrated that Bi-HCy-ErNPs nanoprobes had the potential to serve as reporters for distinguishing cells with varying NE levels, which might enable quantitative imaging of TANs with high specificity.

In vivo TANs visualization using NIR-IIb fluorescence ratiometric nanoprobes

Encouraged by the nanoprobe’s in vitro exceptional performance for precise NIR-IIb fluorescence imaging towards NE, we conducted further investigations in Hepa1-6 tumor-bearing C57BL/6 mice and H22 tumor-bearing BALB/c mice^38,39,40. Prior to in vivo imaging, we initially assessed the blood compatibility of Bi-HCy-ErNPs nanoprobes. The appreciably low hemolysis rate indicated that Bi-HCy-ErNPs nanoprobes were applicable to intravenous administration (Supplementary Fig. 29). When the tumor grew to a certain volume, the tumor-bearing mice received immunotherapy on day 2, 4, and 6 followed by intravenous injection of nanoprobe through the tail vein on day 7 (Fig. 6a). Whole-body NIR-IIb fluorescence imaging at 1532 nm was recorded at various time points upon excitation at 808 and 980 nm. Upon excitation at 980 nm, a strong and persistent NIR-IIb fluorescence signal was observed in the tumors of both non-treatment and the treatment groups (Fig. 6b), which was attributed to the integrated passive and active tumor-targeting of nanoprobes⁴¹ with their prolonged circulation lifetime (Fig. 6b, and Supplementary Figs. 30–33). Conversely, the tumors of the treatment group emitted significantly weaker NIR-IIb fluorescence signals compared to non-treatment group upon excitation at 808 nm. Moreover, upon intratumor injection of NE inhibitor into non-treatment groups, the NIR-IIb fluorescence of Bi-HCy-ErNPs nanoprobes at the tumor sites was significantly declined when excited at 808 nm, but remained relatively stable at 980 nm excitation, indicating the selectivity of Bi-HCy-ErNPs nanoprobes towards NE in vivo. Quantitative analysis revealed that the ratiometric signals of tumor regions in vivo in the treatment group were ~8.0-fold higher compared to those in non-treatment group (Fig. 6c). NIR-IIb fluorescence imaging of tumor lysate also showed stronger ratiometric signals in the treatment group as opposed to non-treatment group (Fig. 6d, e), aligning with the in vivo NIR-IIb fluorescence imaging results. These results were further confirmed by the gold standard hematoxylin and eosin (H&E) staining (Supplementary Figs. 34–35). In addition, body weight assessment, H&E staining, blood biochemical assay, and hematological indexes exhibited that intravenous administration of Bi-HCy-ErNPs did not lead to apparent histological abnormalities/lesions and had no pronounced inflammation/infection, nephrotoxicity, hepatotoxicity, or hematological toxicity (Supplementary Figs. 36–38). We also investigated biodistribution and clearance profiles of Bi-HCy-ErNPs by inductively coupled plasma optical emission spectrometry (ICP-OES). It was observed that the nanoprobes from urine and feces reached their maximum at the 4th day and gradually decreased to normal levels over 14 days (Supplementary Fig. 39). Moreover, ~85% of injected nanoprobes were excreted from the body within 14 days (Supplementary Fig. 40). This indicated that the nanoprobes could be cleared from the body through both hepatobiliary and renal excretion pathways over time. Regarding long-term potential immune responses, the key cytokines like pro-inflammatory cytokines (TNF-α, IL-12, IL-6, and IFN-γ) and anti-inflammatory cytokine (IL-10) showed no significant changes between nanoprobe-injected and PBS-injected groups (Supplementary Figs. 41–43), indicating Bi-HCy-ErNPs caused no abnormal cytokine secretion, reducing adverse immune responses. Additionally, common immune cells revealed no significant population changes after nanoprobe use (Supplementary Fig. 44). This implied that Bi-HCy-ErNPs maintained immune cell homeostasis during long-term use. Overall, these results demonstrated the excellent in vivo biosafety profile of Bi-HCy-ErNPs nanoprobes, supporting their potential for clinical translation. To investigate the correlation between activatable signals of nanoprobes and intratumoral neutrophils, we performed immunofluorescence staining at the end of imaging process. The NIR-IIb fluorescence signal of Bi-HCy-ErNPs in tumor sections from non-treatment group excited at 808 nm was 10.2-fold higher stronger than in those from the treatment group, and showed a high co-localization of ~81.4% with NE-antibody (YA5765, MedChemExpress, USA)-labeled TANs (Fig. 6f, g). Clinical data have demonstrated that elevated levels of M1-tumor-associated macrophages (M1-TAMs) and CD8⁺ Ts were closely correlated with a favorable therapeutic prognosis^42,43. In contrast, an increase in N2-TANs was strongly indicative of lower survival rates among cancer patients^43,44. Thus, the trend of immune status changes reflected by the level variations of M1-TAMs/CD8⁺ Ts was opposite to that of N2-TANs. As depicted in Fig. 6f–l, the change in immunofluorescence signals of N2-TANs following immunotherapy treatment was diametrically contrary to that of M1-TAMs and CD8⁺ Ts, which further validated the high accuracy of our N2-TANs imaging.

**Fig. 6: In vivo NIR-IIb fluorescence ratiometric imaging of Bi-HCy-ErNPs.**

In vivo TANs visualization using NIR-IIb fluorescence ratiometric nanoprobes for immunotherapy efficacy monitoring

To explore the potential of Bi-HCy-ErNPs nanoprobe in immunotherapy efficacy monitoring, we conducted the prolonged NIR-IIb fluorescence ratiometric imaging (Fig. 7a). As illustrated in Fig. 7b, c, on day 8, the NIR-IIb fluorescence ratiometric signals in the treatment group surged abruptly, indicating that the immunotherapy started to produce effects, leading to a change in the immune status of TANs and a significant decrease in NE expression. The results were corroborated by the H&E staining of tumor slices (Supplementary Fig. 45). We further carried out long-term NIR-IIb fluorescence ratiometric imaging and immunotherapy efficacy monitoring (Fig. 7d). As depicted in Fig. 7e, f, the treatment group led to markedly intensified NIR-IIb fluorescence ratiometric signals, while the NIR-IIb fluorescence ratiometric signals remained unaltered in non-treatment group. In addition, we noticed a negative correlation between the NIR-IIb fluorescence ratiometric signal and tumor volumes/weights in Hepa1-6 tumor-bearing mice (Fig. 7g, h), which was consistent with the H&E staining of tumor slices (Supplementary Fig. 46). Moreover, the population of TANs, TAMs, CD8⁺ Ts, mature dendritic cells (mDCs), tumor-infiltrating effector memory T cells (Tems), and tumor-infiltrating regulatory T cells (Tregs) was further quantified by flow cytometry at the end of imaging (Fig. 7i, j, and Supplementary Figs. 47–51). The proportion of M1-TAMs, CD8⁺ Ts, mDCs, and Tems in the treatment group was significantly increased compared to that in the non-treatment mice. Additionally, there was a significant decrease in the proportion of M2-TAMs and Tregs in the treatment group, in contrast to the non-treatment group. Notably, compared to the non-treatment group, the number of TANs in the treatment group was markedly reduced to ~5.54%, nearly 4.24-fold lower than the non-treatment group (Fig. 7i). The results indicated a robust correlation between the NIR-IIb fluorescence ratiometric signals of Bi-HCy-ErNPs and TANs’ immune status changes, suggesting its promising potential as a tool for assessing immunotherapeutic responses.

**Fig. 7: In vivo NIR-IIb fluorescence ratiometric imaging and immunotherapy efficacy monitoring.**

In vivo TANs visualization using NIR-IIb fluorescnece ratiometric nanoprobes for immunotherapy efficacy monitoring, individual stratification, and immunotherapy strategy optimization

To assess the potential of Bi-HCy-ErNPs nanoprobe in individual stratification, we established Hepa1-6 tumor-bearing C57BL/6 and H22 tumor-bearing BALB/c mouse models to distinguish between responder and non-responder to immunotherapy via NIR-IIb fluorescence ratiometric imaging (Fig. 8a). Initially, nine C57BL/6 mice were inoculated with Hepa1-6 cells to establish Hepa1-6 tumor-bearing C57BL/6 mice. After tumor reached a certain range of volume, all nine mice underwent immunotherapy on days 0, 2, and 4. On day 20, all nine mice received an intravenous injection of Bi-HCy-ErNPs. Subsequently, we assessed NE responsiveness at 24 h post-injection using NIR-IIb fluorescence ratiometric imaging. As depicted in Fig. 8b-d, six mice (mice 4-9) were categorized into “responder group” based on robust NIR-IIb fluorescence ratiometric signals due to high NE levels within tumor microenvironment. Conversely, the remaining three mice were assigned to “non-responder group” owing to feeble NIR-IIb fluorescence ratiometric signals, which could be ascribed to relatively low NE levels within tumor microenvironment. In addition, we observed a negative correlation between the NIR-IIb fluorescence ratiometric signal and tumor volumes/weights/NE expression levels in Hepa1-6 tumor-bearing mice as well (Fig. 8e, f and Supplementary Fig. 52). Furthermore, the results obtained by H&E (hematoxylin and eosin)/TUNEL(terminal deoxynucleotidyl transferase dUTP nick end labeling)/Ki-67 staining as well as TANs/CD8⁺ Ts staining of tumor slices from Hepa1-6 tumor-bearing mice, proved the distinctions in cellular damage between NE-responsive and NE-irresponsive groups upon immunotherapy (Fig. 8g). These findings revealed significant differences in TAN immune status between NE-responsive and NE-nonresponsive groups, which could be visualized in real-time using Bi-HCy-ErNPs nanoprobes. In addition, NE is primarily secreted by neutrophils, and its expression level is closely associated with tumor invasion and metastasis. No systematic classification was established between NE expression levels and specific tumor types^{9,45,46,47,48,49}. Thus, the similar results were also observed in the H22 and 4T1 tumor-bearing BALB/c mice (Fig. 8b-e, f, g, and Supplementary Figs. 53-55).

**Fig. 8: In vivo NIR-IIb fluorescence ratiometric imaging for immunotherapy efficacy monitoring and individual stratification.**

Immune checkpoint inhibitors played major roles in immune escape by inhibiting immune responses and promoting immune tolerance toward tumor cells. Among these, PD-L1 and CTLA-4 have been identified as promising targets. The combined blockade of Immune checkpoint inhibitors might provide potentially synergistic effects due to the differences in the timing and location of their reactions. Therefore, in cases where monotherapy with anti-PD-L1 antibody (HY-P99145, MedChemExpress, USA) showed suboptimal efficacy, combination therapy with anti-CTLA-4 antibody (HY-P99132, MedChemExpress, USA) might achieve enhanced immunotherapeutic effects. Additionally, lenvatinib, a VEGFR inhibitor, has been shown to provide potential synergistic effects when used in conjunction with immune checkpoint inhibitors by inhibiting angiogenesis and reversing immunosuppressive tumor microenvironment. To further assess the potential of Bi-HCy-ErNPs nanoprobe in personalized immunotherapy, we delved deeper into immunotherapy efficacy monitoring, individual stratification, and immunotherapy strategy optimization (Fig. 9a). First, fifteen C57BL/6 mice were inoculated with Hepa1-6 cancer cells. Once tumor grew to a certain range of volume, all fifteen mice underwent immunotherapy via intraperitoneal injection of anti-PD-L1 antibody on days 0, 2, and 4. On day 13, an intravenous injection of Bi-HCy-ErNPs was administered to all the mice. Subsequently, individual stratification was carried out at 24 h post-injeciton, employing NIR-IIb fluorescence ratiometric imaging. As presented in Fig. 9b–e, nine mice (mice 7−15) were classified as responder group, whereas the remaining six mice (mice 1−6) were designated to non-responder group. Moreover, a negative correlation between the NIR-IIb fluorescence ratiometric signal and tumor volumes/weights was detected across all groups (Fig. 9e, f). Then, the non-responder group (mice 1-6) were further randomly divided into two groups including adjustor group (mice 1-3) and non-adjustor group (mice 4-6). Among the adjustor group, 3 mice further received immunotherapy by receiving intraperitoneal injection of anti-PD-L1 antibody plus anti-CTLA-4 antibody⁵⁰ at the day 14, 16, and 18. At the day 20 and 27, the mice further received intravenous injection with Bi-HCy-ErNPs for imaging on days 21 and 28 to monitor immunotherapy efficacy. As illustrated in Figs. 9b-d, g, h and Supplementary Fig. 56, the in vivo NIR-IIb fluorescence ratiometric signals showed a gradual increase from day 14, via day 21, to day 28, suggesting an effective therapeutic response after adjusting the immunotherapy regimen. Furthermore, the comprehensive results, including ex vivo NIR-IIb fluorescence ratiometric imaging, TANs/CD8⁺ Ts staining, tumor evolution monitoring, flow cytometric assay of TANs/TAMs/CD8⁺ Ts, as well as H&E/TUNEL/Ki-67 staining of tumor slices from Hepa1-6 tumor-bearing mice (Fig. 9i–o), further validated the enhanced therapeutic efficiency after optimizing immunotherapeutic interventions. It also clearly demonstrated that the NIR-IIb fluorescence ratiometric imaging based on Bi-HCy-ErNPs probe could reliably identify the responders to immunotherapy. Then, the effectiveness of immunotherapy could be assessed in advance, which would in turn provide the possibility for timely adjustment of the immunotherapy strategy. This supported its use in guiding drug selection for precision medicine. Overall, our designed probe offered a promising visualization tool to monitor in vivo immune status change of TANs with high molecular imaging sensitivity. More importantly, our work held the potential to further expand into individual stratification so as to customize personalized treatment plan, with the aim of improving therapeutic efficacy in cancer immunotherapy.

**Fig. 9: In vivo NIR-IIb fluorescence ratiometric imaging for immunotherapy efficacy monitoring, individual stratification, and immunotherapy strategy optimization.**

Method

Software Information

Schematic illustrations, 3D models, and graphical elements were designed and rendered from scratch using the software including 3ds Max 2022.2, Cinema 4D 2023.2, and Adobe Photoshop 2022.3.2, and Adobe Illustrator CS6.

Synthesis of Bi-HCy-VPAA-Ac probe

The probe was synthesized utilizing hemicyanine structure. Subsequently, the indole structure was introduced to enlarge the conjugated system of fluorescence probe, leading to a substantial redshift in both excitation and emission wavelengths. The following details elaborated on the synthesis process:

Synthesis of Compound 1 (1-ethylbenzo[cd]indol-2(1H)-one)

1.0 mmol of benz[cd]indol-2(1H)-one was dissolved in dimethylformamide. At 0 °C, 1.5 mmol of sodium hydride and 1.2 mmol of iodoethane were added to the solution and stirred for 2 h. After returning to 25 °C, the mixture was poured into deionized water and then extracted using ethyl acetate. The organic phase collected from this process was dried using anhydrous sodium sulfate. Subsequently, the solvent was removed through evaporation, and the product was purified by column chromatography with a yield of ~75.8%. ¹H NMR spectrum of compound 1 is included in Supplementary Fig. 2.

Synthesis of Compound 2 (1-ethyl-2-methylbenzo[cd]indol-1-ium iodide)

1.0 mmol of 1-ethylbenzo[cd]indol-2(1H)-one was dissolved in anhydrous tetrahydrofuran and stirred at 25 °C under nitrogen atmosphere. Then 2.0 mmol of CH₃MgCl dissolved in anhydrous tetrahydrofuran was slowly added dropwise to the solution. The mixture was stirred at 60 °C for 2 h, then cooled to 25 °C, and slowly added dropwise into ice-deionized water containing 37% HCl. Next, a saturated KI solution was added, and the mixture was stirred for 30 min. The resulting precipitate was collected by filtration, washed with deionized water and ethyl acetate, and dried under vacuum to yield a red solid with a yield of ~62.9%. ¹H NMR spectrum of compound 2 is included in Supplementary Fig. 3.

Synthesis of Compound 3 (2-bromo-1-cyclohexene-1-carboxaldehyde)

1.0 mmol of PBr₃ was slowly added dropwise to dichloromethane solution containing 1.5 mmol of dimethylformamide at 0 °C. Upon reaching 25 °C, the mixture was stirred for ~30 min until a suspended solid formed. Subsequently, 0.3 mmol of cyclohexanone dissolved in dichloromethane was slowly added, and the mixture was stirred at 25 °C for 8 h. After the reaction was complete, the mixture was quenched by pouring it into ice deionized water. Subsequently, the product was extracted with dichloromethane, and the organic phase was dried with anhydrous sodium sulfate. The solvent was removed through evaporation, and the product was purified by column chromatography with a yield of ~62.5%. ¹H NMR spectrum of compound 3 is included in Supplementary Fig. 4.

Synthesis of Compound 4 (6-methoxy-2, 3-dihydro-1H-xanthene-4-carbaldehyde)

1.0 mmol of 2-bromo-1-cyclohexene-1-carboxaldehyde was dissolved in dimethylformamide. 0.5 mmol of Cs₂CO₃, and 2.0 mmol of 2-hydroxy-4-methoxybenzaldehyde was added in the solution. The solution was stirred overnight under nitrogen atmosphere. After that, the solution was poured into deionized water. Subsequently, the organic phase was extracted with dichloromethane, washed three times with deionized water, and further washed three times with saturated saline solution, subsequently dried by anhydrous sodium sulfate. Ultimately, the solvent was evaporated, and the crude product was isolated by column chromatography with a yield of ~70.3%. ¹H NMR spectrum of compound 4 is included in Supplementary Fig. 5.

Synthesis of Compound 5 (6-hydroxy-2, 3-dihydro-1H-xanthene-4-carbaldehyde)

1.0 mmol of 6-methoxy-2, 3-dihydro-1H-xanthene-4-carbaldehyde was dissolved in dried dichloromethane at 0 °C. Subsequently, 1.5 mmol of BBr3 dissolved in dichloromethane was added slowly dropwise into the solution. The reaction mixture was stirred vigorously for 3 h and subsequently quenched with deionized water. The collected organic phase was dried over anhydrous sodium sulfate and purified by column chromatography with a yield of ~51.6%. ¹H NMR spectrum of compound 5 is included in Supplementary Fig. 6.

Synthesis of Compound 6 (e-1-Ethyl-2-(2-(6-hydroxy-2, 3-dihydro-1H-xanthen-4-yl)vinyl)benzo[cd]indol-1-ium iodide, Bi-HCy-OH)

In nitrogen atmosphere, 1.0 mmol of 1-ethyl-2-methylbenzo[cd]indol-1-ium iodide and 1 mmol of 6-hydroxy-2, 3-dihydro-1H-xanthene-4-carbaldehyde were added in ethanol and refluxed overnight with stirring. After the reaction was complete, the mixture was cooled to 25°C, and the resulting precipitate was collected via filtration. The product was subsequently purified by column chromatography with a yield of ~30.5%. ¹H and ¹³C NMR NMR of compound 6 are included in Supplementary Figs. 7 and 8.

Synthesis of Bi-HCy-VPAA-Ac

Firstly, HOOC-VPAA-Ac was synthesized by solid-phase peptide synthesis. Secondly, 1.0 mmol of HOOC-VPAA-Ac, 2.0 mmol of ethoxycarbonyl-2-ethoxy-1, 2-dihydroquinoline (EEDQ), and 2.0 mmol of p-aminobenzyl alcohol (pABA) were dissolved in dichloromethane and stirred at 25 °C for 6 h. After that, the mixture was purified by high performance liquid chromatography to obtain pABA-VPAA-Ac with a yield of ~89.6%. ¹H and ¹³C NMR spectra of pABA-VPAA-Ac are included in Supplementary Fig. 9. Thirdly, 1.0 mmol of pABA-VPAA-Ac was dissolved in anhydrous tetrahydrofuran, and subsequently, 3.0 mmol of PBr₃ was added. The mixture was stirred at 0 °C for 2 h. Then, tetrahydrofuran was removed via rotary evaporation, and the residue was dissolved in an excess of ethyl acetate. The solution was washed three times with NaHCO₃ solution and subsequently with saturated saline solution. After concentration and drying, the residue was dissolved in anhydrous acetonitrile, and then 0.2 mmol of Bi-HCy-OH and 100 µL of N, N-diisopropylethylamine (DIPEA) were added. The mixture was stirred at 55 °C for 4 h. Subsequently, acetonitrile was removed via rotary evaporation, and the residue was purified by high-performance liquid chromatography. The product, Bi-HCy-VPAA-Ac, was then freeze-dried with a yield of ~9.2%. ¹H and ¹³C NMR spectra of Bi-HCy-VPAA-Ac are included in Supplementary Fig. 10.

Preparation of precursors

1 mmol of rare-earth oxide (Yb₂O₃, Er₂O₃, Ce₂O₃, Y₂O₃, and Lu₂O₃) was respectively dissolved in 10 mL of 50% aqueous trifluoroacetic acid, refluxed at 90^oC for 4 h, and evaporated under N₂ protection. Afterward, the white powder precursors of Yb(CF₃COO)₃, Er(CF₃COO)₃, Ce(CF₃COO)₃, Y(CF₃COO)₃, and Lu(CF₃COO)₃, were obtained for subsequent uses.

Synthesis of NaYbF₄: Ce, Er core nanoprobes

NaYbF₄: Ce, Er nanoprobes were synthesized via co-precipitation of trifluoroacetate precursors with oleylamine (OM), oleic acid (OA), and 1-octadecene (ODE). A solution containing 0.96 mmol of Yb(CF₃COO)₃, 0.02 mmol of Er(CF₃COO)₃, 0.02 mmol of Ce(CF₃COO)₃, and 1 mmol of CF₃COONa was mixed with 6 mL of OM, 6 mL of OA, and 15 mL of ODE in a three-necked flask. The mixture was stirred vigorously at 25 °C for 1 h ad then heated to 120 °C under vacuum for another 1 h to remove deionized water and oxygen impurities. Subsequently, the solution was slowly heated to 300 °C under argon atmosphere for 30 min. After cooling to 25 °C, an excess of ethanol was added, and the nanocrystals were collected by centrifugation at 9000 x g for 10 min. The nanocrystals were washed with ethanol three times and dispersed in 10 mL of cyclohexane for further use.

Synthesis of NaYbF₄: Ce, Er@NaYF₄: Yb, Lu core-shell nanoprobes

NaYbF₄: Ce, Er@NaYF₄: Yb, Lu core-shell nanoprobes were synthesized via a seed-mediated shell growth approach. 0.5 mmol of Y(CF₃COO)₃, 0.2 mmol of Yb(CF₃COO)₃, 0.3 mmol of Lu(CF₃COO)₃, 1 mmol of CF₃COONa, and the as-prepared core nanoprobes were added to a three-necked flask containing a mixture of 10 mL of OA and 10 mL of ODE while stirring vigorously. The mixture was gradually heated under vacuum to 120 °C for 1 h to effectively remove deionized water and oxygen impurities. Subsequently, the solution was slowly heated under an argon atmosphere to 300 °C and held for 1 h. Following cooling to 25 °C, an excess of ethanol was added to the solution. The obtained nanoprobes were centrifuged at 12, 000 x g for 10 min, thoroughly washed with ethanol, and finally dispersed in 10 mL of cyclohexane.

Introduction of Bi-HCy-VPAA-Ac onto NaYbF₄: Ce, Er@NaYF₄: Yb, Lu core-shell nanoprobes

The NaYbF₄: Ce, Er@NaYF₄: Yb, Lu dispersion was evaporated to remove the cyclohexane by rotary evaporator and ultrasonically dispersed in 5 mL of chloroform. Subsequently, 2 mg of Bi-HCy-VPAA-Ac dissolved in ethanol were added and the mixture was continuously stirred overnight. After that, the dispersions were washed using ethanol by centrifugation and redispersed in cyclohexane. The obtained Bi-HCy-ErNPs nanoprobes were stored at 4 °C for further use.

Synthesis of Bi-HCy-ErNPs nanoprobes by surface functionalization of cRGD-targeting, interpenetrating polymer network

To keep the ultra-bright fluorescence imaging and zero fluorescence photo-bleaching ability within aqueous and biological media, the Bi-HCy-VPAA-Ac dye-sensitized NaYbF₄: Ce, Er@NaYF₄: Yb, Lu core-shell nanoprobes were surface functionalized with cRGD-targeting, interpenetrating polymer network. The network consists of four distinct polymer layers, featuring an innermost layer composed of poly(maleic anhydride-alt-1-octadecene) (PMH), succeeded by an eight-arm branched poly(ethylene glycol) amine (8Arm-PEG-NH₂) layer, a poly(acrylic acid) (PAA) layer, and an outermost layer composed of 1, 2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino (poly(ethylene glycol)] (DSPE-PEG-NH₂)/8Arm-PEG-NH₂/DSPE-PEG-cyclic arginine-glycine-aspartic acid (DSPE-PEG-cRGD). Regarding the surface functionalization of PMH, 80 mg of PMH (average molecular weight: 30,000–50,000) was dissolved in 5 mL of chloroform. Subsequently, 32 mg of Bi-HCy-VPAA-Ac dye-introduced NaYbF₄: Ce, Er@NaYF₄: Yb, Lu dispersed in cyclohexane was added to the solution. Next, the mixture was agitated for 1 h and then subjected to rotary evaporation to remove the cyclohexane. Following this, 80 mg of 4-(dimethylamino)pyridine (DMAP) dissolved in 6 mL of aqueous solution was added, and the mixture was ultrasonicated until the ErNPs were uniformly dispersed. The dispersions were centrifuged twice at 18, 000 x g for 30 min to remove the excess PMH/DMAP. The precipitates were redispersed in 3 mL of morpholineethanesulfonic acid (MES) solution (pH 8.5). Regarding the surface functionalization of 8Arm-PEG-NH₂, 12 mg of 8Arm-PEG-NH₂ (average molecular weight: 40, 000) dissolved in 3 mL of MES solution was added. Then, 8 mg of EDC was added, and the mixture was agitated for 2 h. Next, 2-amino-2-(hydroxymethyl)-1, 3-propanediol (Tris-base, 40 mg) and EDC (20 mg), both dissolved in MES solution, were added to block the excess carboxyl group from PMH. Subsequently, the mixture was agitated for another 2 h, washed by centrifugal filter (300 kDa) to remove excess PEG/EDC, and subsequently dispersed in 3 mL of MES solution. Regarding the surface functionalization of PAA, 4 mg of PAA (average molecular weight: 1, 800) dissolved in 3 mL of MES solution was added to the above dispersion. Then, 8 mg of EDC dissolved in MES solution was added, and the mixture was agitated for 2 h. Next, the solution was washed by centrifugal filter (300 kDa) to remove excess PAA/EDC and dispersed in 3 mL of MES solution. Regarding the surface functionalization of DSPE-PEG-NH₂, 8Arm-PEG-NH₂, and DSPE-PEG-cRGD, 4 mg of DSPE-PEG-NH₂ (average molecular weight: 5000), 1 mg of 8Arm-PEG-NH₂ (average molecular weight: 40, 000), and 0.8 mg of DSPE-PEG-cRGD, dissolved in 3 mL of MES solution, were added to the above dispersion. Then, 8 mg of EDC dissolved in MES solution was added, and the mixture was agitated for 2 h. Subsequently, 20 mg of Tris-base and 10 mg of EDC, both dissolved in MES solution, were added into the above solution to block the excess carboxyl group from PAA. Finally, the mixture was agitated for another 2 h and washed by centrifugal filter (300 kDa) to remove excess PEG/EDC. The obtained Bi-HCy-ErNPs nanoprobes were stored at 4 °C for further use.

In vitro NIR-II fluorescence imaging of nanoprobes

The tested nanoprobes were introduced into either PCR tube or centrifuge tube cap and placed on an imaging system for in vitro NIR-II fluorescence imaging. They were excited with either an 808 nm or 980 nm laser at a power density of 15 mW/cm² and their fluorescence images were acquired by an NIR-II camera using a 1500-1600 nm band-pass emission filter.

In vitro NIR-II fluorescence imaging of cells

Naive neutrophils were isolated from the blood of healthy mice using the Biolegend MojoSort Mouse Neutrophil Isolation Kit. Then, neutrophils were polarized towards “N1” or “N2” phenotype to investigate the specificity of Bi-HCy-ErNPs nanoprobes⁵¹. For polarizing neutrophils towards “N1” phenotype, the process was carried out in the complete medium supplemented with an N1-polarization cocktail containing LPS (100 ng/mL), IFN-γ (50 ng/mL), and IFN-β (10, 000 U/mL) at 37 °C in a humidified air atmosphere containing 5% CO₂ for 48 h¹⁷. For polarizing neutrophils towards “N2” phenotype, the process was carried out in the complete medium (adjusted to 6.7) supplemented with an N2 polarization cocktail containing L-lactate (25 mM), adenosine (10 μM), TGF-β (20 ng/mL), IL-10 (10 ng/mL), prostaglandin E2 (20 ng/mL), and granulocyte colony stimulating factor (100 ng/mL) at 37 °C in a humidified hypoxic chamber containing 2% O₂ and 5% CO₂ for 48 h¹⁷. After the incubation of tested nanoprobes for different incubation periods, neutrophils were rinsed three times with PBS, and subsequently imaged using a NIR-II fluorescence imaging system.

Hepa1-6, 4T1, H22 and NIH/3T3 cells were employed as controls, which were authenticated by short tandem repeat (STR) before usage as follows: Genomic DNA was extracted using Axygen genomic DNA extraction kit and amplified with PowerPlex®21 system using 21-short tandem repeat (STR) amplification protocol. STR loci and sex-determining gene Amelogenin were detected on an ABI 3730XL Genetic Analyzer. Data were processed using Gene Mapper ID 3.2 software (Applied Biosystems). Appropriate positive and negative controls were run and confirmed for each cell line. The resulting STR profiles of the tested cell lines matched with the STR data of corresponding cell lines in Cellosaurus database (Supplementary Table 1). In addition, these cell lines were routinely tested for mycoplasma contamination using a commercial Mycoplasma PCR Detection Kit (Beyotime, C0301S) and confirmed to be mycoplasma-free before all experiments.

Animal model

Female C57BL/6 and BALB/c mice (6−9 weeks) were purchased from Guangzhou Yaokang Biotechnology Co., Ltd. Before imaging, the mice were shaved and anesthetized with 4% isoflurane. To establish the tumor-bearing mice model, the experimental mice received the subcutaneous inoculation of cells at the sides.

In vivo NIR-II fluorescence imaging

Twenty microliters (20 µL) of Bi-HCy-ErNPs nanoprobes, suspended in PBS at a concentration of 1 mg/mL, were intravenously administered to tumor-bearing mice at varying time points after tumor grew to a certain volume. NIR-II fluorescence imaging was conducted employing an imaging system, utilizing either an 808 nm or 980 nm laser as the excitation source and equipped with a 1500−1600 nm band-pass emission filter. The mice were anesthetized via inhalation of 1-2% isoflurane in air throughout the duration of experiment. The acquired images were processed using Image J software, where false coloring was applied and the Region of Interest (ROI) values were extracted.

In vitro/In vivo quantitative detection

For in vitro calibration and controls, a series of solutions with known concentrations of NE were prepared. The nanoprobe was incubated with these solutions, and the fluorescence intensity ratio (F_{1532em, 808ex}/F_{1532em, 980ex}) was measured. A calibration curve was then constructed by plotting the fluorescence intensity ratio against the NE concentrations. This calibration curve was used to quantify NE levels. For in vivo calibration and controls, after the administration of the nanoprobe, the fluorescence signals from the tumor region were monitored at various time points. The fluorescence intensity ratio (F_{1532em, 980ex}/F_{1532em, 808ex}) was measured, and correlated with the tumor volume, the NE levels determined by enzyme-linked immunosorbent assay (ELISA), and other biochemical methods to establish a quantitative relationship.

Discussion

Cancer immunotherapy has revolutionized the treatment paradigm in oncology⁵². However, due to the influence of tumor type, staging, intratumoral heterogeneity, and individual variations, significant heterogeneity persists in patients’ immune microenvironments, leading to marked disparities in treatment responses^52,53. Therefore, implementing precision personalized immune status evaluation and dynamic monitoring of immunotherapeutic efficacy is critical for patients undergoing immunotherapy. Tumor-infiltrating leukocytes, encompassing T cells, TAMs, and TANs, serve as pivotal biomarkers for immune monitoring⁴³. Their functional states, subtype abundance, and spatial architecture critically govern immunotherapy outcomes⁴³.

Within the tumor microenvironment, a high density of infiltrating neutrophils has emerged as a critical prognostic indicator for poor prognosis in most cancer types, such as liver cancer, lung cancer, and oral cancer. As mentioned above, TANs at the tumor infiltrating edge often presented as the N1 phenotype, while those in the tumor center tended to be the N2 phenotype^10,11. This might be related to the fact that the combined parameters of low intratumoral TANs, high stromal TANs, and high necrosis have been shown to indicate significantly greater 3 year disease-free survival in cancer patients^12,13,14. Due to the complexity of TANs’ subpopulations, there was still a lack of sufficient research on how to accurately describe TANs’ subpopulations and quantify TANs infiltration using reliable biomarkers to predict tumor prognosis based on TANs density. Our work leveraged the advantages of NIR-IIb fluorescence ratiometric imaging, enabling real-time quantitative visualization and monitoring of TANs, thereby providing a powerful tool for predicting tumor immunotherapy responses. Therefore, quantitative dynamic visualization of NE expression to assess immune status changes in TANs within the tumor immune microenvironment is of great significance for monitoring immunotherapy efficacy, predicting immunotherapy outcomes, identifying potential immunotherapy targets, and screening and optimizing immunotherapy strategies.

Tumor biopsy and peripheral blood analysis are currently the standard clinical methods for obtaining tumor-associated neutrophil (TAN) information. However, biopsy is invasive and delayed in providing results, and only reflects local information, while blood analysis lacks spatial resolution and is confounded by nonspecific interference¹⁵. Molecular imaging technologies offer a novel approach to noninvasively monitor the tumor immune microenvironment dynamically and visualize the immune status of TANs. Nevertheless, conventional imaging modalities (e.g., CT, PET, and MRI) suffer from limitations such as strong background signal, poor sensitivity, single-target detection capability, and compromised tissue specificity^17,18. In contrast, activatable fluorescence probes utilize an “Off-On” signal-switching mechanism to specifically respond to target enzyme activity, enabling molecular-level precision in visualization imaging^18,54. Despite the simplicity of imaging techniques relying on single-channel fluorescence intensity changes^17,55, their signal accuracy is inherently compromised by probe biodistribution, environmental interference, and instrument variability, limiting applications to qualitative assessments.

Fluorescence ratiometric imaging technology overcomes these limitations through a dual-signal self-calibration mechanism, which quantifies targets via reference-to-response signal ratio changes, thereby significantly enhancing detection accuracy^31,32,33. Current optical probes are primarily restricted to visible light or NIR-I fluorescence²⁷. However, this system remains limited by shallow tissue penetration depth and substantial autofluorescence interference. In contrast, NIR-II fluorescence imaging, particularly within the NIR-IIb sub-window, could minimize tissue photon scattering as well as absorption coefficients^28,29,30, enabling deep-tissue imaging while achieving significant improvement in signal-to-noise ratio (Supplementary Figs. 57–59). By synergizing ratiometric self-calibration with NIR-II optical advantages, we propose that activatable NIR-II fluorescence ratiometric probes could dynamically resolve TANs’ immune status with spatiotemporal precision, ultimately advancing personalized immunotherapy through real-time quantitative efficacy assessment. Although we acknowledge that the design strategy such as cascaded FRET pathway between NIR-II fluorescence dye and ErNPs have been reported by Zhang and co-workers³⁵, the existing nanoprobes are insufficient for in vivo quantitative dynamic imaging of TANs. Nevertheless, our tailored probe meets this need via the energy-transfer cascade, providing a potential tool for monitoring immunotherapy efficacy and guiding immunotherapy decisions. In our future work, the innovation from the bottom layer of probe design will also be an important research direction.

In summary, given the pivotal role of TANs in immunotherapy, this study addressed the critical challenge of achieving high-precision, high-specificity quantitative dynamic imaging of TANs within the tumor microenvironment for real-time immune status monitoring. Leveraging the synergistic advantages of fluorescence ratiometric imaging technology and NIR-IIb fluorescence imaging, we developed a NIR-IIb fluorescence ratiometric nanoprobe with highly selective responsiveness towards NE to quantitatively visualize TANs for immunotherapy efficacy monitoring and strategy optimization. Upon active targeting towards the tumor and subsequent interaction with NE overexpressed in N2-TANs, our probe exhibited a fluorescence ratiometric response: the 808 nm-excited NIR-IIb signal at 1532 nm showed significant amplification, whereas the 980 nm-excited reference signal at 1532 nm remained stable. This dual-channel self-calibration mechanism allowed NIR-IIb fluorescence ratiometric imaging to distinguish immunotherapy-responsive individuals with low N2-TANs levels, from immunotherapy-irresponsive individuals with high N2-TANs levels. Furthermore, this approach enabled dynamic correlation visualization between N2-TANs’ immune status changes and therapeutic outcomes. The elaborate design provided a potential strategy for quantitative dynamic imaging of N2-TANs, enabling both the monitoring of immunotherapy efficacy and the optimization of immunotherapy strategy. Consequently, a personalized immunotherapy plan could be implemented under the imaging guidance, bridging the gap between immune status quantification and clinical decision-making.

Statistics and reproducibility

All data are presented as mean ± standard deviation with individual data points indicated. All experiment was repeated three times independently with similar results. Group differences were evaluated by one-way ANOVA (multiple groups) or two-tailed unpaired Student’s t-test (two groups) in GraphPad Prism 8.0.2. Exact p-values are reported in the figures and legends. Statistical significance was defined as *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001. Correlations were quantified with Pearson’s correlation coefficient.

Ethics Statement

All animal experiments were performed in compliance with the protocols approved by the Animal Ethics Committee of Mengchao Hepatobiliary Hospital of Fujian Medical University (MCHH-AEC-2023-17). In accordance with the requirements, the tumor size of mice must not exceed 2000 mm³.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The experimental data generated in this study are provided in the Supplementary Information file, Supplementary Data 1-25 file, and Source Data file. NMR spectrometry data are available in Supplementary Figs. 2-7, Supplementary Fig. 8a, b, Supplementary Figs. 9-10, and Supplementary Data 5–16 (raw figure exported from NMR spectrometry software for NMR spectrometry data), and mass spectrometry data are available in Fig. 2c, d, Supplementary Fig. 8c, Supplementary Figs. 11-13, Supplementary Data 1-4, and Supplementary Data 17-25 (raw figure exported from mass spectrometry software for mass spectrometry data). No datasets require deposition in public repositories, and no access restrictions apply. Source data are provided with this paper.

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Acknowledgements

Y.L. and J.L. contributed equally to this work. This work was partially supported by the National Natural Science Foundation of China (Grant No. 62475264, Y.L.; Grant No. 62175031, X.L.; Grant No. 62275050, Y.Z.), the Fujian Provincial Natural Science Foundation of China (Grant No. 2025Y0056, Y.L.). We thank Wenlong Li, Kaiyuan Wang, Jie Hu, Yingping Huang, Shasha He, MengLi Ma, Yongwei Guo, Qinglai Wang, Yun Zhang, Haomiao Zhu, Fulin Lin, Peiyuan Wang, Xiaolong Zhang, Zhixiong Cai, Ming Wu, Qiang Luo, Yueyang He, Yanling Lin, Xiaohang Liu, Fukai Zhu, Shuangyan Bao, Zhankun Hao, Zhiyang Fan, Chen Yang, Li Li, and Haotian Ding for their invaluable help and advice. We also thank Shiyanjia Lab (www.shiyanjia.com) for XPS analysis.

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These authors contributed equally: Yang Li, Jinyan Lin.

Authors and Affiliations

State Key Laboratory of Structural Chemistry, Fujian Institute of Research on the Structure of Matter, Chinese Academy of Sciences, Fuzhou, P. R. China
Yang Li, Jinyan Lin & Xiaolong Liu
The United Innovation of Mengchao Hepatobiliary Technology Key Laboratory of Fujian Province, Mengchao Hepatobiliary Hospital of Fujian Medical University, Fuzhou, P. R. China
Yang Li, Jinyan Lin, Yongyi Zeng & Xiaolong Liu
Xiamen Key Laboratory of Rare Earth Photoelectric Functional Materials, Xiamen Institute of Rare-Earth Materials, Chinese Academy of Sciences, Xiamen, P. R. China
Yang Li, Jinyan Lin & Xiaolong Liu
Department of Occupational and Environmental Health, School of Public Health, Xi’an Jiaotong University, Xi’an, P. R. China
Jinyan Lin
Departments of Diagnostic Radiology, Surgery, Chemical and Biomolecular Engineering, and Biomedical Engineering, Yong Loo Lin School of Medicine and Faculty of Engineering, National University of Singapore, Singapore, Singapore
Jianhua Zou

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Yang Li
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Jinyan Lin
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Jianhua Zou
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Contributions

Y.L., J.L., J.Z., Y.Z., and X.L. conceived and designed the research. Y.L. and J.L. performed the experiments. Y.L. and J.L. analysed the data and wrote the manuscript. J.Z., Y.Z., and X.L. supervised the project. All authors discussed the results throughout the project and approved the final version of the manuscript.

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Correspondence to Jianhua Zou, Yongyi Zeng or Xiaolong Liu.

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Nature Communications thanks Hak Soo Choi, Bengang XING and Haijun Yu, and the other anonymous reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

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Li, Y., Lin, J., Zou, J. et al. NIR-IIb fluorescence ratiometric imaging of tumor-associated neutrophils for immunotherapy efficacy monitoring and strategy optimization. Nat Commun 16, 10880 (2025). https://doi.org/10.1038/s41467-025-65927-1

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Received: 05 February 2025
Accepted: 27 October 2025
Published: 05 December 2025
Version of record: 05 December 2025
DOI: https://doi.org/10.1038/s41467-025-65927-1