Intracellular Monitoring of Luminescent Ag2S Nanoparticles in Fibroblasts

Mahshid Hashemkhani, Thomas Naillon, Lise Abiven, Stéphane Roux, Bruno

Viana, Corinne Chanéac, Florence Gazeau

► To cite this version:

Mahshid Hashemkhani, Thomas Naillon, Lise Abiven, Stéphane Roux, Bruno Viana, et al. Intra-cellular Monitoring of Luminescent Ag2S Nanoparticles in Fibroblasts. Advanced Optical Materials, 2025, 27 (11), pp.279. 10.1002/adom.202501425. hal-05329093

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# Intracellular Monitoring of Luminescent Ag₂S Nanoparticles in Fibroblasts

Mahshid Hashemkhani, Thomas Naillon, Lise Abiven, Stéphane Roux, Bruno Viana,*
Corinne Chanéac,* and Florence Gazeau*

Interaction between nanoparticles (NPs) and cells is a key concern in nanotechnology-based biomedical applications. This study investigates cytocompatible silver sulfide (Ag2S) quantum dots (QDs), which emit strongly in the second near-infrared (NIR II) region, for their potential in image-guided therapies. Since cellular microenvironments contain acidic pH, hydrogen peroxide (H2O2), and reactive oxygen species (ROS), which can affect luminescence, Ag2S NPs (AS NPs) coated with three ligands: DTDTPA (dithiolated diethylene triamine pentaacetic acid), 11MUA (11 mercaptoundecanoic acid), and PEG (polyethylene glycol) were studied in fibroblast cells over 9 days. Long term photoluminescence (PL) tracking showed distinct behavior. AS DTDTPA and AS 11MUA exhibited increasing uptake but decreasing PL, while AS PEG showed both enhanced PL and uptake over time. No significant changes in NP size or structure explained the PL differences. Exposure to H2O2 revealed that AS DTDTPA and AS 11MUA lost PL due to oxidation or shell detachment, reducing surface passivation. In contrast, AS PEG showed intensified PL, possibly from removal of metallic silver and increased PL lifetime. These results highlight time dependent PL changes in cellular and oxidative environments. DTDTPA and PEG coatings may act as ROS scavenger and promoter, respectively, while enabling NIR II imaging and offering insights into NP design for stable optical tracking and therapeutic use.

## 1. Introduction

Advancements in the synthesis and modification of nanoparticles (NPs) to finely control their physical, chemical, and biological properties opened the way for new generation multifunctional nanoprobes for theranostic applications and personalized

medicine.[1-4] In particular, fluorescent semiconductor quantum dots (QDs) have attracted great attention due to their size-dependent photoluminescence (PL) and broad excitation spectra. QD can be designed to generate strong luminescence in the near-infrared-II (NIR-II) biological window, which is well suited for medical imaging due to deeper tissue penetration and a higher signal-to-background ratio as a result of lower tissue autofluorescence in the NIR-II range.[5,6] Heavy metal-free binary QDs such as CuS and Ag₂S (AS) due to their biocompatibility and colloidal stability have been utilized as imaging agents, photothermal and/or photodynamic therapy (PTT/PDT) agents, etc.[7,8]

Despite the rapid advancement in developing innovative NPs for nanomedicine, only a limited number have successfully reached the clinic so far.[9] This concern arises from their distinct properties compared to their bulk material counterparts, which could result in posing unknown risks to patients.[10] In addition, due to the transformations NPs undergo within the cells, their behavior post-internalization may differ significantly. To determine NPs performance in cells, many aspects of their behavior must be investigated and understood.[11,12] Several studies have shown that the physical and chemical properties of NPs such as their size, shape, homogeneity, surface chemistry, etc. could alter in physiological environment and have an impact on their cellular behavior.[3,13] For

M. Hashemkhani, L. Abiven, F. Gazeau

NABI

Université Paris Cité

CNRS UMR7057, 45 Rue Saint des Peres, Paris 75006, France

E-mail: florence.gazeau@u-paris.fr

The ORCID identification number(s) for the author(s) of this article can be found under https://doi.org/10.1002/adom.202501425

© 2025 The Author(s). Advanced Optical Materials published by Wiley-VCH GmbH. This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited.

DOI: 10.1002/adom.202501425

T. Naillon, L. Abiven, C. Chanéac

LCMCP

Sorbonne Université

CNRS, 4 Place Jussieu, Paris 75005, France

E-mail: corinne.chaneac@sorbonne-universite.fr

T. Naillon, B. Viana

IRCP

CNRS

PSL Chimie Paris Tech

11 Rue P. & M. Curie, Paris 75005, France

E-mail: bruno.viana@chimieparistech.psl.eu

S. Roux

Laboratoire Chrono-environnement UMR6249 CNRS-UFC

University of Franche-Comté

Besançon 25030, France
<figure><img src="image_3.png" /><figcaption>**Scheme 1.** Surface ligands used for the synthesis of AS NPs and their binding modes to the nanoparticle surface for the three different ligand types.</figcaption></figure>

instance, reduced particle size results in having larger surface areas that is associated with higher reactive sites on the surface enhancing their cellular uptake.[14,15] Smaller particles tend to show higher structural defects altering their electronic properties on the surface of NPs, thereby creating reactive groups resulting in both increase and/or decrease in NPs luminescence. It was seen in the study of Li et al. that the aggregation of La₀.06Ce₀.26Tb₀.14PO₄ NPs enhanced their luminescence by a factor of 4.[16]

$H_2O_2$ plays multifunctional role in cell biology mechanisms involving oxidative stress and intracellular signal transduction.[17] It is the most common representative and stable form of non-radical ROS in living organisms.[18] Hydrogen peroxide ($H_2O_2$) and ROS such as superoxide ($O_2^•$), hydroxyl radical ($\cdot OH$), and singlet oxygen ($^1O_2$) are produced endogeneously through the mitochondrial respiration process, or exogenously by exposure to deleterious conditions including infectious agents and UV light.[19] The ROS molecules are highly reactive molecules capable of interacting with various organic substrates, potentially triggering extensive alterations or damage to ells and/or NPs through chain reactions.[20] Therefore, there have been various studies to investigate the NPs performance influenced by $H_2O_2$ and ROS molecules or the use of reactive NPs as ROS sensors inside cells. Of the many analytical techniques that have been reported, optical (absorption, luminescence, and fluorescence) and electrochemical (amperometry) approaches are the most popular to study the changes in NPs in the presence of ROS and $H_2O_2$.[21] It has been reported that $H_2O_2$ increased the luminescent intensity of $CaS:Ln^{3+}$ NPs ($Ln = Ce, Nd, and Er$) due to the $4f \to 5d$ transition of $Ln^{3+}$ enhancing the NIR-II luminescent of NPs. Another case on increasing the luminescent of NPs is the plasmonic electron transfer in Ag/AS Janus NPs as demonstrated by Zhang et al.[6] On the other hand, it has also been reported a decrease in the luminescent of particles by $H_2O_2$, such as Au-11MUA nanorods, $NaYF_4:Yb^{3+}/Er^{3+}$ upconversion

NPs, $Ce^{3+}$ doped vanadate, $Eu^{3+}$-doped gadolinium.[13,17,19] However, in most cases, $H_2O_2$ has no influence on the luminescence of particles.[18]

Herein, we propose three types of AS NPs that differ in their ligand corona, to monitor their optical and cellular behavior in vitro (**Scheme 1**). All three contain a thiol (—SH) group, which binds strongly to silver atoms, ensuring nanoparticle stabilization and preventing aggregation. However, they differ in their organic chains, which influence their specific properties. HS-PEG provides steric stabilization, enhances hydrophilicity, and is bio-compatible, making it highly suitable for pharmaceutical applications. 11MUA, with its carboxyl (—COOH) group, improves aqueous solubility and enables the attachment of biomolecules, making it particularly effective for imaging, biomolecular detection, and therapy. DTDTPA, distinguished by its two thiol (—SH) groups, forms even stronger bonds with silver atoms and contains multiple carboxylate (—COO⁻) groups, facilitating bio-conjugation with proteins, fluorophores, or therapeutic agents while maintaining nanoparticle dispersion.[22,23] The effect of these capping agents (different size, shape, and surface charge) namely DTDTPA (dithiolated diethylene triamine pentaacetic acid), 11MUA (11-mercaptoundecanoic acid), and PEG (polyethylene glycol) was studied in terms of variations in the size, shape and surface charge of the NPs formed. Fibroblasts are the key component of tumor microenvironment and the building blocks of connective tissues with high NP uptake. Therefore, the fate of AS NPs and changes in their colloidal and optical properties in fibroblasts were investigated for long incubation time up to 9 d. Based on the changes in the luminescence of NPs reported in the literature, the effect and mechanisms of $H_2O_2$ on the luminescent of AS NPs were investigated in this paper.[24,25] Various change in the luminescent of AS NPs in this study offers a simple and selective optical sensing system to detect $H_2O_2$ and subsequently ROS molecules in cells.
# 2. Experimental Section

## 2.1. Materials

All chemicals were of analytical grade or highest purity. Silver nitrate (AgNO₃, 99.0%), sodium sulfide nonahydrate (Na₂S·9H₂O, 99.99%), 11-mercaptoundecanoic acid (95%), poly(ethylene glycol) methyl ether thiol (6 kDa), lipopolysaccharide (LPS), and N-acetylcysteine (NAC) were purchased from Sigma Aldrich Chemistry. DTDTPA is a non-commercial product and was prepared as previously reported.[26] Roswell Park Memorial Institute (RPMI) 1640 medium (with L-glutamine and 25 mM HEPES) and Dulbecco's modified Eagle medium (DMEM) was obtained from Multicell, Wisent Inc. (Canada). Fetal bovine serum was provided from Capricorn Scientific GmbH (Germany). Trypsin-EDTA and Penicillin/Streptomycin solutions were purchased from Wisent Inc. (Canada). Alamar blue and phosphate buffered saline (PBS) tablets were purchased from Biomatik Corp.

## 2.2. Synthesis of AS QDs

AS-11MUA QD was synthesized in aqueous media using microwave. Briefly, 20 mM AgNO₃, and 11MUA in the molar ratio of 11MUA/Ag: 3 was dissolved in 10 mL deionized (DI) water and stirred for 24 h. The pH of the solution was adjusted to 8 using NH₃ (30%w) and HCL (37%w). Na₂S. 9H₂O (Ag/S: 2) was first dissolved in 4 mL DI water, then added to the reaction mixture before putting the reaction solution in the microwave. The reaction solution was heated to 100 °C for 15 min in a monomodal microwave (Anton Paar Monowave 300, 1200 tr min⁻¹). At the end of the reaction, as-prepared QDs were separated from solution by centrifugation after destabilization of the suspension with ethanol and then washed with DI water using dialysis cassette (Thermo Scientific Slide-A-Lazer G2, 10 kDa) for 48h.

To synthesize AS-PEG (PEG: 6 kDa), the procedure was the same as explained for AS-11MUA with the molar ratio of PEG/Ag/S: 2/2/1 and without pH adjustment. The same synthesis method and parameters were used in the case of AS-DTDTPA with the molar ratio of DTDTPA/Ag/S: 2/2/1 and the microwave temperature was set to 150 °C for 15 min.

### 2.2.1. Characterizations

The absorption spectra of all NPs were recorded using Shimadzu UV-3600 UV-Vis spectrophotometer. Since the bandgap energy of the QDs is size dependent, the relationship between the size and the band gap energy of the QDs was stated by Brus equation:[27,28]

$$ \Delta E = \frac{\hbar^2 \pi^2}{8R^2} \left[ \frac{1}{m_e} + \frac{1}{m_h} \right] - 1.8 \frac{e^2}{\epsilon_{Ag_2S} 4\pi \epsilon_0 R} \quad (1) $$

where E<sub>g</sub>(Bulk) is the band gap energy of the semiconductor, h is the Planck's constant, m<sub>e</sub> and m<sub>h</sub> are the mass of the excited electron and hole, e is the charge of an electron, ε is the dielectric constant of the material and R is the radius of the QDs.[27] This equation offers a convenient method to estimate the theoretical particle size from the band gap energy instead of particle size measured by transmission electron microscopy (TEM),

scanning electron microscopy (SEM), X-ray diffraction (XRD), or other characterization tools.

Lifetime and quantum yield (QY) of NPs were recorded on an Edinburgh FLS100 Spectrometer. Hydrodynamic size and surface charge of NPs were determined by dynamic light scattering (DLS) and zeta potential measurements using Malvern Zetasizer. TEM was carried out by a TECNAI 120 Spirit Twin at an acceleration voltage of 120.0 kV, with a Gatan Orius 1000 camera model.

## 2.3. Cell Culture

Fibroblast cells were cultured in complete DMEM media supplemented with 2% fetal bovine serum (FBS) and 1% penicillin-streptomycin antibiotic solution according to ATCC recommendations. 2% FBS limits the proliferation of fibroblasts allowing to track them for longer time (9 d).[29] Trypsin-EDTA was used for the detachment of the cells from the surface.

## 2.4. Determination of Cellular Viability

Cell viability was evaluated by performing the Alamar blue assay on fibroblasts measuring the mitochondrial metabolic activity of cells. Cells were seeded at a density of 3500 cells per well into 96-well culture plates in culture medium and incubated at 37 °C and 5% CO₂ atmosphere for 24 h. Following incubation, the medium was replenished; cells were treated with AS NPs (0–4 µg mL⁻¹ [Ag]) and incubated for further 48 h. Then, the medium was removed, and cells were washed with PBS. To evaluate the cytotoxicity over 9 d, the cells were prepared as mentioned before and NPs were incubated for 1, 3, 5, and 9 d with cells. Alamar blue reaction solution was added to the cells and incubated for 2 more hours. Absorbance was measured at 540 and 580 nm in each well with a iD3 SpectraMax microplate reader (Molecular Devices, CA).

## 2.5. Cellular Uptake

### 2.5.1. Qualitative Analysis

The in vitro cell imaging of NPs was performed with a confocal microscope (Leica dmi8/SP8) to show the cell uptake qualitatively. Fibroblast cells were grown on a 3 mL glass-bottom petri dish at a density of 1.5 × 10³ cells per well in complete medium, cultured at 37 °C under 5% CO₂ for 24 h. NPs at 1 µg mL⁻¹ [Ag] were introduced to the cells in complete medium, and 1 and 3 d after incubation, the cells were washed three times with PBS to remove the un-internalized NPs. Then, the cells were fixed with paraformaldehyde solution (4% in PBS) and incubated for 20 min. To stain the cell nuclei, cells were incubated for an additional 15 min with 2 µg mL⁻¹ of DAPI. After rewashing the cells with PBS three times to remove the unbound DAPI dye, 1 mL PBS was left in each well to protect the cells against drying. The fixed cell samples were imaged using a confocal microscope with two different filters for NIR region (λ<sub>ex</sub>: 510–550 nm and λ<sub>em</sub>: 710 nm long pass with a HC PL APO CS2 63 × /1.40 oil immersion objective, exposure time: 14.6 s) and DAPI (λ<sub>ex</sub>: 325–375 nm and λ<sub>em</sub>: 435–485 nm). The same experimental procedure was also performed for control cells, which were not treated
with any NPs. Images were processed and merged using ImageJ software.

### 2.5.2. Quantitative Analysis

To quantify the NPs' uptake, inductively coupled plasma mass spectrometry (ICP-MS) analysis was performed to determine the internalized Ag content of treated cells. Fibroblasts were seeded on a 250 mL flask at a density of $3 \times 10^6$ cells per well in a complete medium and cultured for 24 h at 37 °C and 5% CO$_2$. Next day, the cells were incubated with AS NPs at $1 \mu g mL^{-1}$ [Ag] for 1, 3, 5, and 9 d. Then, the cells were washed with PBS to remove un-internalized particles and were collected after being detached from the well surface with trypsin. Cells were digested with HNO$_3$ for 1 week at RT to dissolve NPs, and Ag content was determined with ICP-MS ($n = 3$).

### 2.5.3. TEM Analysis

Fibroblasts were prepared as mentioned earlier with $1 \mu g mL^{-1}$ [Ag] for 1, 3, 5, and 9 d. Cells were detached by trypsinization, washed twice with PBS, and fixed with 2% glutaraldehyde in 0.1 M Na cacodylate buffer pH 7.2, for 1 h at room temperature. Thin sections (70 nm) were collected onto 200 mesh copper grids, and counterstained with lead citrate. Images were acquired with a charge-coupled device camera (AMT). TEM images of cells were acquired using a Hitachi HT7700 operating at 80 kV (MIMA2 platform, INRAE, Jouy-en-Josas, France).

## 2.6. PL Measurement of AS NPs in Fibroblasts

Fibroblast cells were incubated and AS NPs were added as explained in quantitative analysis in Section 2.5. Then, the cell pellet ($3 \times 10^6$ cells per well) was washed using PBS after NP incubation after 1, 3, 5, and 9 d. 200 µL of cell pellet with NPs were added to 96-well plate and their luminescence were measured using the setup shown in Scheme S1 (Supporting Information). To evaluate the cell-induced changes in the shape of the emission band, PL emission spectra of AS NPs dispersed in water or internalized in cells were recorded using an infrared hyperspectral camera thermoelectric cooled InGaAs camera (PyLoN-IR, Princeton Instruments). 730 nm LED excitation beam was collimated with a 5 cm focal collimator. The data was normalized to the emission of equal number of cells without any particles.

## 2.7. Determination of ROS (ROS Generation)

### 2.7.1. In Solution

$1 \mu g mL^{-1}$ of AS-DTDTPA, AS-11MUA, and $10 \mu g mL^{-1}$ AS-PEG were incubated with $10 mM H_2O_2$ 30% v/v for 1, 3, 5, and 9 d. To compensate for the significantly higher ligand proportion in the AS-PEG sample (96.2% compared to 49.8% and 45.3% for

the AS-11MUA and AS-DTDTPA samples, respectively), its mass concentration in the suspension was increased tenfold (Figure S2, Supporting Information). Then, the PL of NPs were measured as explained in Section 2.6.

### 2.7.2. In Vitro

To measure the ROS level quantitatively in cells, ROS assay utilized using 2'-7'dichlorofluorescin diacetate (DCHF-DA). After diffusion of DCHF-DA into the cells and its deacetylation by cellular esterase, the resulting product is then oxidized by ROS into fluorescent 2'-7'dichlorofluorescin (DCF). So, fibroblast cells were seeded on 96-well plates for 24 h. Then, cells were washed with PBS and DCHF-DA ($0.045 \mu g mL^{-1}$ in media) were added to the cells and were kept in dark for 40 min in incubator. After removing the DCHF-DA and washing the cells with PBS, cells were treated with $10 mM NAC$ (ROS scavenger) or $50 ng mL^{-1}$ LPS (ROS producer) 30 min prior to the NPs' addition and then, NPs were added at $1 \mu g mL^{-1}$ Ag concentration. The ROS levels in the cells were determined every 2 h using ROS generation kits (Abcam cellular ROS Detection Kit (orange)) at $\lambda_{ex}/\lambda_{em} = 480/530$ nm following the manufacturer's instructions using an iD3 SpectraMax microplate reader (Molecular Devices, CA).

## 2.8. Statistical Analysis

Two-way ANOVA with Tukey's multiple comparison test of GraphPad Prism 8 software package was employed for statistical analysis (GraphPad Software, Inc., USA). All data were expressed as mean ± standard deviation (SD), and $p < 0.05$ was considered statistically significant.

# 3. Results and Discussion

## 3.1. Synthesis, Chemical, and Photophysical Characterization and Cytotoxicity of AS NPs

Aqueous AS NPs were synthesized from AgNO$_3$ and Na$_2$S using a simple and fast microwave method with three different capping agents: PEG, DTDTPA, and 11MUA. The structure and length of the ligands significantly impact the optical properties of the final NPs. Therefore, PEG, DTDTPA, and 11MUA were selected to investigate their effects on the optical properties of NPs with and without cells. The growth of NPs depends on the complexation strength of the ligands, which varies according to their functional groups and solvation, resulting in different particle sizes. Consequently, these AS NPs emit in the NIR-II region, with emission maxima between 980 and 1250 nm, depending on their particle size relative to the QD structure (Figure 1a).

Since the particles were synthesized with different coatings, direct comparison is challenging. However, the molar ratio of coating/Ag/S was 2/2/1 in AS-DTDTPA and AS-PEG, while it was 6/2/1 in AS-11MUA. In general, organic coatings passivate the surface of the growing crystal, playing a crucial role in controlling crystal size, surface quality, and colloidal stability. A higher coating concentration leads to increased particle size due to greater
<figure><img src="image_4.png" /><figcaption>**Figure 1.** a) Absorption calibrated (PL) and b) UV-vis spectra of AS-DTDTPA, AS-11MUA, and AS-PEG NPs. c) TEM images and the corresponding histogram of (i, ii) AS-DTDTPA, (iii, iv) AS-11MUA, and (v, vi) AS-PEG NPs, respectively. d) Ag 3d, e) S 2p XPS spectra of AS-DTDTPA, AS-11MUA, and AS-PEG NPs. *The 1000 nm long-pass filter was replaced by an 830 nm long-pass. f) Binding modes of thiol ligands on Ag₂S NPs.</figcaption></figure>

organic adsorption on the NP surface, resulting in a red shift in the luminescence spectra. The higher coating/Ag ratio in AS-11MUA resulted in a red shift due to its larger crystal size (3.82 nm) and TEM-measured size (5.8 nm).

As mentioned earlier, the crystal size of QDs obtained by the Brus equation is calculated based on the optical shift in UV-Vis or PL spectra. The calculated size is usually smaller than the actual physical size, as it represents the Bohr radius, which is influenced by electron-hole interactions.[28] Additionally, this method assumes a perfectly spherical particle. In contrast, TEM provides the actual physical size of individual NPs, accounting for shape, aggregation, and crystallinity. Therefore, surface defects, ligand effects, and NP shape lead to the optical size determined by the Brus Equation (based on the electronic band gap) being relatively smaller than the actual TEM size.

Furthermore, the synthesis temperature of AS-DTDTPA was higher (150 °C) than that of the other NPs (100 °C), enhancing Ostwald ripening. This resulted in an increased crystal size (3.67 nm) and TEM-measured size (4.4 nm) of AS-DTDTPA compared to AS-PEG (crystal size: 3.42 nm, TEM: 3.2 nm), even though both had the same coating/Ag/S molar ratio, leading to a red shift in the PL spectra (Figure 1a and Table S1, Supporting Information). Consequently, AS-11MUA and AS-DTDTPA exhibited higher PL peak positions than AS-PEG NPs. The absorption onset at 850-890 nm enables their application in NIR-II imaging (Figure 1b).

Crystal sizes determined by the Brus Equation can be used to evaluate the influence of reaction variables on particle size and, consequently, luminescence wavelength. However, TEM and Brus equation sizes do not always correlate well in the literature, as the reported electron and hole effective masses for AS differ from those obtained by TEM. The hydrodynamic sizes (number-based) of AS-DTDTPA, AS-11MUA, and AS-PEG were

6.2, 10.9, and 9.3 nm, respectively. As expected, TEM-measured sizes were slightly smaller than DLS-measured sizes due to the presence of coating materials. NP agglomeration based on intensity-weighted hydrodynamic size can be seen in Table S1 (Supporting Information), though it was limited by the organic shell, providing excellent colloidal stability.

Negative charges recorded by zeta potential measurements at pH ≈ 7.5 (Table S1, Supporting Information) were attributed to anionic groups on the NP surfaces. AS-DTDTPA had a zeta potential of -1.5 mV due to -SH and -COOH groups, while AS-11MUA had a zeta potential of -24.7 mV due to -COOH groups. Surprisingly, AS-DTDTPA exhibited a low charge (-1.5 mV) despite containing carboxyl groups, which are typically deprotonated at this pH. This suggests that the carboxyl groups remain coordinated to silver. Compared to 11MUA, which also has carboxylate groups, this result can be explained by the chelate effect, where DTDTPA's structure increases its affinity for metal ions compared to monodentate ligands. Non-polar groups like -CH₃ on the surface of AS-PEG do not directly contribute to charge but may still result in a weak negative charge, consistent with the observed value (-1.3 mV).

The hydrodynamic size and charge of NPs can vary significantly depending on the solution medium due to factors such as ionic strength, pH, and the presence of proteins or macromolecules. Therefore, the size and charge of AS NPs were measured in DMEM medium with 2% FBS, as used in cell culture. The number-based hydrodynamic sizes increased to 10.1 nm (AS-DTDTPA), 13.5 nm (AS-11MUA), and 18.7 nm (AS-PEG) due to the adsorption of serum proteins and electrolytes. The charge also changed, with zeta potentials increasing to -5.5 mV (AS-DTDTPA), -10.4 mV (AS-11MUA), and -1.1 mV (AS-PEG) in culture medium, likely due to protein adsorption reducing electrostatic interactions and shielding the NP surfaces.
To further characterize the surface state and chemical composition of AS NPs, XPS analysis was performed. Ag 3d peaks at 367.8 eV (3d5/2) and 373.8 eV (3d3/2) correspond to Ag⁺ in the AS core and were observed in all three AS-DTDTPA, AS-11MUA, and AS-PEG NPs (Figure 1d).[4,6] Additional Ag peaks in AS-PEG at lower intensity (368.3 eV for 3d5/2 and 374.3 eV for 3d3/2) suggest the presence of a metallic Ag core.[30] A broad S 2p signal (Figure 1e,f) was fitted to two doublets at 161.2 eV (2p3/2) and 162.3 eV (2p1/2), corresponding to Ag–S covalent bonds in AS-DTDTPA, AS-11MUA, and AS-PEG' core.[5] The S 2p signal at a higher BE of 162.2 eV (2p3/2) and 163.4 eV (2p1/2) is assigned to the S atoms of ligands that are bound to the core atoms (Ag–S–R).[31,32] DTDTPA contains two thiol groups—one bound to the AS core, with the other possibly remaining free on the NP surface. In AS-11MUA, the presence of unbound thiolates suggests coordination to the AS surface via carboxyl groups. This indicates that while most thiol s contribute to covalent bonding with the core, some remain unbound, potentially affecting NP optical properties over time in solution and cells. Since AS-PEG lacks additional functional groups, only Ag–S doublets were detected. Another doublet at 163.4 eV (2p3/2) and 164.6 eV (2p1/2) is assigned to unbound thiol (–SH) groups in AS-DTDTPA and AS-11MUA.

Additionally, doublets at 168.1 eV (2p3/2) and 169.2 eV (2p1/2) in AS-11MUA were attributed to oxidation products formed during synthesis. Both –COOH and –SH groups contribute to particle formation, and –COOH binding to the core may lead to thiol oxidation into sulfenic (RSOH), sulfinic (RSO₂H), and, most commonly, sulfonic (RSO₃H) acids.[33] Notably, all three synthesized AS NPs did not require an oxygen-free atmosphere, unlike most colloidal semiconductor NP syntheses reported in the literature.[4,7] As a result, these air-insensitive AS NPs remained stable at ambient conditions for months.

The viability of fibroblasts treated with 0–4 µg Ag mL⁻¹ was determined by Alamar blue assay evaluating mitochondria metabolic activity, in comparison to the non-exposed cells, after 48 h of incubation with the different AS NPs in DMEM media containing 2% FBS. According to ISO-10993-5, viability value higher than 80% is considered as non-toxic.[8] Therefore, none of the particles were toxic in this range of concentration after 48 h of incubation as represented in Figure S3a (Supporting Information). Besides, we studied the fate of fibroblasts exposed to the NPs over 9 d of incubation at 1 µg Ag mL⁻¹ since this concentration is non-toxic and high enough to study their change in cells. AS-PEG and AS-11MUA NPs were highly cytocompatible over 9 d without any noticeable cytotoxicity. However, AS-DTDTPA reduced the viability of fibroblasts to ≈60% and 50% at days 5 and 9 of incubation, respectively (Figure S3b, Supporting Information). Such toxicity was already reported for aqueous Au@DTDTPA particles at concentration of 100 µg NPs mL⁻¹ produced by Butterworth et al. that reduced the viability of PC3, DU145, and PNT2-C2 cells after 24 h of incubation.[34]

## 3.2. Internalization of AS NPs into the Cells

The internalization of the NPs by fibroblast was qualitatively determined by TEM and fluorescence microscopy and quantified by intracellular [Ag] detected by ICP-MS to elucidate the intracellular

distribution and fate of NPs after 1, 3, 5, and 9 d of incubation with AS at 1 µg mL⁻¹ [Ag]. In the case of AS-11MUA and AS-DTDTPA, the uptake of NPs was increased over time from 4% to 10% wt (AS-DTDTPA) and 5% to 25% wt (AS-11MUA) from day 1 to day 9 of incubation as seen in Figure S3c (Supporting Information). On the other hand, the NPs' uptake in AS-PEG increased from 4% to 53% wt from day 1 to day 5 and then decreased to 42% wt at day 9. If these particles exhibit the smaller core size, their hydrodynamic diameter is larger than AS-DTDTPA. In fact, it has been reported that the size, shape, surface charge, heterogeneity, hydrophobicity/hydrophilicity, surface functional groups and properties govern the uptake of NPs into cells as well as the adsorption of proteins onto their surface.[35]

The NPs' distribution and their fate in fibroblast cells were followed by TEM at different times after NPs' exposure to the cells. A wide range of intracellular responses could be observed when NPs were internalized by cells. Endocytosis pathways including pinocytosis, the formation of caveolae and clathrin, and caveolae/clathrin-independent uptake may take place.[11] As reported in the literature, particle sizes less than 50–120 nm are preferred for endocytic uptake which is the case for all three AS NPs.[11,35] As shown in **Figures 2** and S4 (Supporting Information), NPs interacted with the cell plasma membrane at day 1 and then quickly internalized into the membrane-closed endosomes. Although polydisperse AS-DTDTPA created aggregates in the solution and increased their number-based hydrodynamic size to 10 nm in DMEM media with 2% FBS, the NPs formed pretty large and diffuse aggregates (>50 nm) after cell uptake making therefore impossible the measurement of size for every single particle (Figure 2a). Adsorption of water molecules and proteins on the surface of AS-DTDTPA NPs were supposed to form a steric layer on the surface making steric repulsion between pairs that results in higher colloidal stability. However, low charge of NPs (–5.5 eV in 2% DMEM) resulting from the formation of dithiol bridges by the reaction of –SH groups lead to the formation of aggregates in AS-DTDTPA in cells. On the contrary, AS-11MUA and AS-PEG were taken up by the cells in endosomes.

Endosomes are vesicles formed to engulf NPs from extracellular environment through endocytosis. On the other hand, lysosomes are specialized organelles with acidic pH (= 4.5–5) containing hydrolytic enzymes for breaking down various biomolecules. Upon degradation of NPs by cells, the endosome fuses with a lysosome allowing the contents to be degraded by lysosomal enzymes. AS-11MUA are promptly incorporated and dispersed in the membrane-closed endosomes after one day. Then, they tend to create aggregates due to the protein adsorption on their surface with high NPs concentrations surrounded by an electron dense lysosomal matrix at days 5 and 7 to degrade NPs. The aggregates seen in the lysosomes after day 3 suggested the implication of endogenous lysosomal proteins in the transformation of NPs into cells. The mean size of AS-11MUA and AS-PEG NPs increased slightly over 3 d however it is equal to the initial sizes at day 9 indicating neither degradation nor dissolution of NPs. Such difference in size could be attributed to the difficulty in NPs size measurement and to uncertainty (counting smaller number of NPs, segregation of particle without the same polydispersity than in suspension, lower magnitude of TEM images, and difficulty identifying particle contours in the cell).
<figure><img src="image_5.png" /><figcaption>**Figure 2.** TEM images of fibroblasts treated with 1 µg Ag mL⁻¹ of a) AS-DTDTPA, b) AS-11MUA, and c) AS-PEG NPs at days 1, 3, 5, 9. Scale bars are 500, 200, and 100 nm.</figcaption></figure>

Upon internalization of NPs, they were transferred to the acidic microenvironment of endosome/lysosome, which normally lead to degradation of the coating materials and/or the NPs over time and eventual excretion/metabolization of NPs and their degradation products.[36,37] Exocytosis depends on the size of NPs; bigger NPs are slower exocytosed and consequently lower NPs uptake may be seen after some time.[37] Therefore, agglomeration of NPs after entering the cells reduced the exocytosis kinetics. The reason for the reduction of intracellular NPs' in cells at day 9 compared to day 5 could be attributed to the mentioned fact.

One of the great advantages of luminescence AS QDs is their application as imaging agents in NIR-II which is well suited for medical application due to high signal/noise and deeper penetration.[7] The fluorescence microscopy images of fibroblasts treated with and without AS NPs on day 1 and 3 of incubation

in **Figure 3** revealed numerous punctuated intracellular spots of luminescent NPs that is in good agreement with above TEM evidence of endosomal/lysosomal confinement of NPs and previous results in the literature for AS NPs with different coating (AS coated with branched polyethyleneimine-25 kDa and L-cysteine, AS coated with N-acetyl-L-cysteine-coated).[38,39] After comparing the NIR signals at each day to the non-exposed control in Figure 3, AS-11MUA showed the lowest signal compared to the other NPs at day 1. However, the signal from AS-11MUA started to increase at day 3. The data showed that the AS-11MUA became brighter from day 1 to day 3 while AS-DTDTPA exhibited lower signal after day 3 and there was not significant change in the brightness of AS-PEG. The data obtained in Figure 3 demonstrated the highest intracellular fluorescent intensity for AS-PEG NPs compared to the other synthesized NPs after 3 d.
<figure><img src="image_6.png" /><figcaption>**Figure 3.** a) Fluorescence microscopy images of fibroblasts treated with 1 µg Ag mL⁻¹ of AS-DTDTPA, AS-11MUA, AS-PEG NPs, and untreated control cells for 1 and 3 d. The exposure time for the samples at each day is constant compared to each other's and the fluorescent signals were compared to the control at each day. Red: luminescence from AS (λ<sub>ex</sub>/λ<sub>em</sub> = 510–550/710 nm long-pass filter), blue: nuclear stain (λ<sub>ex</sub>/λ<sub>em</sub> = 325–375/435–485 nm). The scale bar is 15 µm. b) Quantitative analysis of normalized fluorescent intensity of confocal microscopy images by the intracellular silver ion content showing intracellular AS NPs in fibroblast at 1 µg mL⁻¹ [Ag] after 1 and 3 d incubation (*p* < 0.05).</figcaption></figure>

### 3.3. Evolution of Optical Properties of AS NPs in Cells

Our experimental plan, maintaining ultralow proliferation of cells over time due to low serum supplementation, allows us to investigate the fate of AS NPs in fibroblasts on a period of 9 d. TEM observations revealed that the different AS NPs, albeit they show different sizes, levels of cellular uptake and intralysosomal distributions, globally maintain their size, shape, and arrangement over time with no evident sign of structural degradation. However, we observed different evolutions of the intracellular luminescence after particle uptake as illustrated in Figure 3. Therefore, we questioned how the optical properties of NPs can be modified by cellular uptake and how they evolve along the cellular processing during the 9 d of incubation. In general, it has been reported that fluorescence of NPs can be quenched by close proximity interaction in aggregates, can be diminished by NPs degradation involving core dissolution, or modified by chemical modifications resulting in surface reactions with their environment.[4] Such mechanism, if relevant in our systems, would be all the more pronounced during long-term incubation with cells and in the acidic enzyme rich environment of lysosomes.[4] Besides, the acidic endosome/lysosome boosts the degradation of surface coating and thus the degradation of the shell, this would result in loss of colloidal stability and florescence quenching.[4] The PL intensity of AS-DTDTPA, AS-11MUA, and AS-PEG was studied in the cells for different incubation times, from 1 to 9 d, as presented on Figure 4a–c and compared to that of NPs suspension used for cell exposure (red curve) and to the PL spectrum of control non-exposed cells. The presence of an emission peak on PL spectra at around 1015 nm (black curve) of cells without NPs is attributed to water in cells.[40,41]

For AS-DTDTPA and AS-11MUA, PL was very low after 1 d incubation and thus slightly enhanced over time (Figure 4a,b), possibly due to the increased NPs internalization over 9 d as determined by ICP-MS (Figure S3c, Supporting Information). Since

the amount of the internalized NPs were low after 1 d of incubation (around 5%), the emission peaks related to the cells at ≈1015 nm were more dominant. For a better representation of this evolution over time, the integral of the PL spectra was normalized to the emission of NPs without the cells (area under the emission peak of initial NPs suspension = 100%, Figure 4d,e). While the slight increase in luminescence of AS-DTDTPA with increasing exposure time seems to correlate with the increase in the number of internalized particles, this trend was almost the same for AS-11MUA sample. Luminescence by prolonged presence of AS-PEG NPs in the acidic microenvironment evolved in a completely different way. As illustrated in Figure 4c, the luminescence of AS-PEG after internalization in cells at day 1 was similar to that of initial suspension of NPs although the quantity of NPs is much lower (less than 10%). Then, luminescence enhanced significantly over 9 d and reaches almost 250%, while only around 50% of AS-PEG NPs are internalized for longer time of exposure (days 5 and 9). Clearly, such changes cannot be explained solely by the difference in internalized NPs number after incubation. In fact, it was reported that the presence of ROS and H₂O₂ in the cells also had an impact on the luminescence of NPs.[6,18]

To find out the role of luminescence dependent ROS for such changes, NPs' emission was monitored upon H₂O₂ addition in suspension (without cells) over 9 d. Thus, each NPs' suspension was incubated with 10 mM H₂O₂, and PL and lifetimes were monitored over a period of 9 d to investigate the kinetic of luminescence evolution.

### 3.4. Evolution of Optical Properties of AS NPs Suspension in Response to H₂O₂

Acidity and overexpression of H₂O₂ are typical features of tumor microenvironment that could be triggered by ROS production.
<figure><img src="image_7.png" /><figcaption>**Figure 4.** a–c) PL spectra of cells at different incubation time with NPs in comparison to the PL spectrum of pristine NPs in suspension, normalized to the emission of equal number of cells. d,–f) Total area under the PL spectrum of cells normalized with that of pristine NPs suspension of a,d) AS-DTDTPA, b,e) AS-11MUA, and c,f) AS-PEG NPs in fibroblast over 9 d.</figcaption></figure>

ROS are also key signaling molecules that play an important role in the progression of inflammatory disorders. Therefore, it is assumed that metabolism is affected by ROS in the cellular environment. Conversely, ROS are able to alter the radiative and/or non-radiative recombination as previously mentioned by Zhang et al.[6] NPs coating, surface structure, and defects have a great impact on the NPs' relaxation pathway. They may participate in non-radiative recombination and energy losses between vibrational modes of the ligand and/or surface trap states.

### 3.4.1. Effect of $H_2O_2$ on AS-DTDTPA

It was reported that thiolate groups in cysteine residues or thiol-terminated capping agents such as dodecanthiol (DDT) and 11MUA could act as a hole trap and ROS-scavenger, reducing consequently the NPs' emission.[21,42,43] DTDTPA molecules consist of two thiol groups where one of them is coordinated to AS core through formation of Ag–S bond. The other thiol group could be: i) oxidized to organic disulfide product (RS-SR) in the presence of $H_2O_2$ upon interaction with another AS molecules or another –SH groups on the same AS molecule (Figure 5a-i).[21,24,44] ii) The thiol group on the surface could be oxidized and detach from the core, reducing the surface passivation and luminescence (Figure 5a-ii).[21,24,44] iii) Another mechanism for luminescence quenching is illustrated in Figure 5a-iii; the electron-transfer reaction that occurred on the surface where $H_2O_2$ was reduced to $O_2$, which in turn lie in electron-hole traps on the NPs.[24] iv) Redox reaction in the presence of untreated Ag

and $H_2O_2$ could interrupt the electron's communication and reconstructed electron-defective AS to emit photons.[6] In the presence of Ag in the structure of AS NPs, the electron transfer from plasmonic Ag to AS could quench the fluorescence. However, elimination of untreated Ag resulted in an increase in the luminescence of NPs (Figure 5a-iv). It is worth mentioning that there are no noticeable changes in the PL peak position and its shape upon treating with $H_2O_2$ showing no change in the size of NPs and their bandgap in $H_2O_2$. The emission intensity of AS-DTDTPA in solution with $H_2O_2$ molecules as demonstrated in Figure 5b reduced over time (emission intensity: D1 > D3 > D5 > D9) due to the mentioned mechanisms of i-iii.

### 3.4.2. Effect of $H_2O_2$ on AS-11MUA

The impact of $H_2O_2$ on the emission spectra of AS-11MUA (no cells) was similar to that of AS-DTDTPA (mechanisms of i-iii), but with a weaker effect in terms of reducing luminescence (Figure 5c). These results were consistent with the study of Shiang et al. upon incubation of $H_2O_2$ and Au-11MUA nanorods.[21] It was reported a luminescence quench due to the oxidation of Au–S bonding in the presence of $H_2O_2$; the same mechanism that could happen in AS-11MUA NPs (Figure 5a-i,ii). The increase in the hydrodynamic size of AS-11MUA after $H_2O_2$ incubation over time leads to slight aggregation suggesting a destabilization of the particles following degradation of the ligand shell as mentioned earlier (Table S2, Supporting Information).
<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /><figcaption>**Figure 5.** a) Schematic illustration of the different fate of AS NPs upon incubation with H₂O₂. PL spectra of b) AS-DTDTPA, c) AS-11MUA, and d) AS-PEG NPs treated with 10 mM H₂O₂ for 9 d. The inset shows the luminescent of NPs without and with H₂O₂ for 9 d using 808 nm laser. S 2p XPS spectra of e) AS-DTDTPA, f) AS-11MUA, and g) AS-PEG NPs with and without incubation with 10 mM H₂O₂ for 9 d.</figcaption></figure>
<figure><img src="image_10.png" /><figcaption>**Figure 6.** a–c) PL spectra of cells alone or of cells at different incubation time with only NPs, NPs with 10 mm NAC (ROS scavenger) and NPs with 50 ng mL⁻¹ LPS (ROS producer) in comparison to the PL spectrum of pristine NPs in suspension, normalized to the emission of equal number of cells. d–f) Total area under the PL spectrum of cells normalized with that of pristine NPs suspension of a–c) AS-DTDTPA, d–f) AS-11MUA, and g–i) AS-PEG NPs in Fibroblast over 9 d.</figcaption></figure>

### 3.4.3. Effect of H₂O₂ on AS-PEG

In the case of AS-PEG, the PL intensity as seen in Figure 5d increased after incubation with H₂O₂ from day 1 to day 5 and decreased slightly afterward at day 9. Recently, Zhang et al. have shown sixfold enhancement in the fluorescent intensity of Ag/AS NPs 24 h after the addition of H₂O₂. [6] They have reported that fluorescence quenching of Ag/AS NPs is due to the electron transfers from plasmonic Ag to semiconductor AS NPs.[6,24] Herein, the unreacted Ag in the structure of AS-PEG (Figure 1d) that could create the hole formation and electron transfer (PL quenching) would be removed by redox reaction with H₂O₂ triggering the radiative recombination of AS-PEG NPs (mechanism iv, Figure 5a-iv). The inset in Figure 5b–d shows the luminescence of all three NPs using 808 nm camera without and with H₂O₂ for 9 d. Moreover, the hydrodynamic size and the PL peak position did not change significantly with H₂O₂ (Figure 5d, Table S2, Supporting Information).

To supports these hypothesis, XPS analysis was performed after H₂O₂ addition. Ag and S doublets corresponding to the formation of AS were seen in Figure S5 (Supporting Information) and Figure 5e–g, respectively. The Ag doublets corresponding to the AS core showed a slight shift (0.3–0.5 eV) after H₂O₂ incubation (Figure S5, Supporting Information). Moreover, the intensity of the metallic silver doublets at 368.3 and 673.3 eV decreased after H₂O₂ proving the elimination of unreacted Ag as mentioned earlier. A slight shift in the S 2p doublets at 161.2 eV (2p3/2) and 162.3 eV (2p1/2) can be seen in Figure 5e,f after reacting with H₂O₂. Interestingly, the ratio of Ag–S peaks compared to the –SH peaks reduced after reaction with H₂O₂ in both AS-DTDTPA and AS-11MUA (Figure 5e,f). In addition, the intensity of S 2p peaks attributed to the bounded thiolates at 162.4 eV (2p3/2) and 163.5 eV (2p1/2) were lower compared to the unbounded thiolate peaks at 163.5 eV (2p3/2) and 164.6 eV (2p1/2) confirming the detachment of DTDTPA and 11MUA from AS core that could subsequently reduce the NPs
emission after H₂O₂ exposure. The formation of doublets at 168.1 eV (2p3/2), 169.2 eV (2p1/2) and 169.3 eV (2p3/2), 170.4 eV (2p1/2) in AS-DTDTPA and AS-11MUA NPs were attributed to sulfinic (RSO₃⁻) and sulfonic (RSO₃⁻) groups, which is possibly formed during the oxidation of dangling bonds at the surface of AS NPs.[33] In the case of AS-PEG, the presence of doublets at 168.1 eV (2p3/2) and 169.2 eV (2p1/2) also explains the oxidation of NPs in the presence of H₂O₂. Lin et al. have previously shown the formation of CdSO₄ and Ag₂SO₄ upon the addition of H₂O₂ to CdS-Ag₂S QDs.[24] This leads to the reduction of PL intensity as a result of surface oxidation.[24] Taken together, this change in the structure of AS-DTDTPA and the oxides formation explains the luminescence quench in the presence of H₂O₂ in solution.

To further study the mechanism of radiative and/or non-radiative recombination, the PL decay curves were studied according to the biexponential function of namely $[I(t): A_1\exp(-t/\tau_1) + A_2\exp(-t/\tau_2)]$, where $I(t)$ is the PL intensity at delay time (t), $\tau_1$ and $\tau_2$ are the decay times, and $A_1$ and $A_2$ are the fractional contributions of the two components of the two reported processes (Figure S6, Supporting Information).[5] The average lifetimes calculated from $\langle r \rangle \approx (A_1 \tau_1^2 + A_2 \tau_2^2) / (A_1 \tau_1 + A_2 \tau_2)$ where $\tau_1$ (fast component) arises from the surface trap states while $\tau_2$ (slow component) comes from deeper donor-acceptor recombination of trapped electrons and holes relevant to the intrinsic defects.[45] Within this approach, radiative decay is related to deep donor-acceptor states surface passivation, while non-radiative decay is controlled by surface states.[45]

The average lifetime of AS-DTDTPA with H₂O₂ increased slightly up to day 3 (day 3: 13.1 µs) and reduced afterwards (day 9: 12.4 µs; Figure S7 and Table S2, Supporting Information). However, fast decay component remained unchanged for the first 3 d and reduced at day 5 and day 9. In addition, the contribution of surface states (fast decay-A₁) increased over time and the effect of deep donor-accepter recombination (slow decay-A₂) decreased with H₂O₂ incubation time. The reason attributes to the fact that the interaction of AS-DTDTPA and H₂O₂ lead to the formation of non-radiative RS-SR bounds and/or the detachment of coating from AS. It was also reported that the lifetime depends on the exact location of hole trapping by Ag⁺, as the overlap with the delocalized electron wave function varies.[46,47] These results were in agreement with Wang et al. that has shown a significant PL quenching in MoS₂ QDs coated with glutathione (have thiol groups) under H₂O₂ addition due to the oxidation of dangling bonds and MoS₂ core.[48] However, most of the studies show no influence of H₂O₂ on luminescent and lifetime of particles; the same happened in AS-11MUA (lifetime with H₂O₂: 12.6–12.9 µs).[18] In AS-11MUA, $\tau_1$, $\tau_2$, A₁, A₂, and the average lifetime values remained almost constant indicating that the PL decay time was related to the non-radiative traps upon H₂O₂ incubation. Moreover, AS-11MUA in Figure 5c showed almost the same PL intensity before and after H₂O₂ addition (at D > 1), confirming most of the studies in the literature that H₂O₂ has no effect on luminescence.[18]

It was believed that the fast decay component ($\tau_1$) was contributed to the structural defects and a certain amount of them still remained in the AS-PEG after H₂O₂ incubation. As shown in Table S2 and Figure S7a (Supporting Information), the fast decay component was increased with H₂O₂ over 5 d and its con-

tribution to the emission profile (A₁) decreased from 36% for AS-PEG NPs to 5.5%. Furthermore, the contribution of radiative related recombination (A₂) increased from 64% to 92% in AS-PEG (no H₂O₂) and AS-PEG with H₂O₂ for 9 d (AS-PEG-H₂O₂-D9), respectively, confirming the predominance of radiative states with longer the decay times. Overall, fast decay component became more and more suppressed under the effect of H₂O₂, suggesting that the amount of structural defects was considerably reduced while PL emission was mainly correlated to intrinsic defects.

**3.5. Modulating ROS in Cells Govern AS PL Depending on the Capping**

To compare the data obtained from NPs interaction in solution with H₂O₂ and the NPs in fibroblasts, AS-DTDPA NPs underwent fluorescent quenching under the effect of H₂O₂ due to mechanisms explained in Figure 5ai,iii, a shift in Ag doublets (Figure S5a, Supporting Information), a decrease in the ratio of Ag–S peaks, an increase in –SH related peaks (Figure 5b), and increase in the contribution of surface state recombination (A₁). These data demonstrated the loose of organic content protecting the AS core resulting in luminescent quench that is similar to the reduction in luminescence of NPs in Fibroblasts.[49] The decrease in AS-DTDTPA’s luminescence was not only related to the acidic microenvironment of lysosomes but also it was related to the reactions with ROS and H₂O₂ in cells.

In the case of AS-11MUA, H₂O₂ lead to a slight reduction of luminescence due to the mechanisms shown in Figure 5ai–iii. The shift in Ag doublets in Figure S5b (Supporting Information), and formation of R-SH peaks proved the oxidation and detachment of organic compound that could result in the PL reduction. Knowing the fact that the H₂O₂ had no noticeable effect on the lifetime parameters of AS-11MUA and comparing their luminescence in cells and in solution with H₂O₂ showed that ROS and H₂O₂ exhibited low effect on their luminescence reduction. Since the acidic microenvironment of endosome/lysosome also had an impact on particles’ luminescence, that might be the reason for the PL reduction observed Figure 4b. The large aggregate formation of AS-11MUA in cells that has shown by TEM (Figure 2b) is another reason for their luminescent quench.

Lastly, H₂O₂ and ROS related mechanism increased AS-PEG luminescence in solution due to the effect shown in Figure 5a-iv, reduction in metallic silver doublets (Figure S5c, Supporting Information), and significant increase in the contribution of radiative related recombination (A₂) and their lifetime (Figure S7d,e, Supporting Information). In comparing the PL enhancement in fibroblasts, it was clearly demonstrated that the enhancement was directly linked to the reaction with H₂O₂ and ROS within the cells.

To further study the impact of ROS and H₂O₂ on the luminescence of NPs in cells, N-acetyl-L-cysteine (NAC) and lipopolysaccharide (LPS) were incubated with cells. NAC is a common antioxidant and a natural scavenger of free radicals and ROS due to the presence of sulfhydryl group.[6,50] Thereby, it has been commonly utilized as a pharmaceutical drug for the diseases induced by free radicals and ROS. Therefore, the cells were treated
<figure><img src="image_11.png" /><figcaption>**Figure 7.** a) ROS production and b) cell viability of fibroblasts upon incubation with AS NPs at 1 µg mL⁻¹ [Ag] after 24 h incubation with and without NAC or LPS. The data are expressed as mean ± S.D. (n = 4), (*p* < 0.05 compared to the control). Untreated cells were used as control.</figcaption></figure>

with NAC, 30 min prior to NPs addition over 9 d, to investigate their effect on luminescence. On the other hand, to induce ROS generation in the reaction LPS has been utilized.[51-53] LPS can trigger inflammation leading to the activation of immune signaling pathways and ROS generation.[54]

The PL spectra of AS-DTDTPA and the area under its curve showed a reduction in the luminescence of NPs in the presence of both LPS and NAC (Figure 6a-i). Since luminescence of these NPs were quenched in the presence of H₂O₂ (Figure 5a,b), the intensity reduction in the presence of LPS that produced more ROS are in agreement with luminescent reduction in Figure 6a. The consumption of ROS by NAC leads to the decrease in the luminescence of AS-DTDTPA with NAC compared to only NPs (Figure 6b). Presence of ROS in the cell microenvironment decreased their luminescence (NPs+cells) compared to NPs without cells (NPs suspension; Figure 6c). The presence of ROS scavenger, NAC, in cell media reduced their emission (even if the NPs are not inside cells) leading to the dramatic reduction in PL intensity compared to AS-DTDTPA in cells (Figure 6b,c).

The results obtained from AS-11MUA with LPS reveal that the emission intensity and its spectral integral were not affected dramatically by the ROS in the first 5 d while a dramatic enhancement is obtained at day 9 that could be due to a delayed effect of ROS or activation of the fibroblasts (Figure 6d-f). The effect of NAC, on the other hand, exhibited nearly the same emission intensity as the ones with cells over 9 d. This clearly shows that ROS molecules do not have any influence on the luminescence of AS-11MUA NPs.

Lastly, incubation of AS-PEG with cells not only improved the PL intensity and the area under each curve from days 1 to 9, but also the intensity was significantly higher when NPs were incubated with LPS due to the elevation of ROS quantity in the cells and formation of radiative recombination (Figure 6g-i). When AS-PEG NPs were utilized with NAC, due to the very low H₂O₂ level in the environment, weak fluorescence was recorded. These data evidently showed that ROS is the main reason for the change in the luminescence of AS-PEG NPs in cells.

To quantify the intracellular ROS, DCFH-DA was used as a fluorescent probe with/without LPS and NAC. The intracellular esterases hydrolyzes DCFH-DA upon enzymatic reaction forming non-fluorescent DCFH, which in the presence of ROS, DCFH is oxidized to highly fluorescent DCF.[5,55] Since the mechanism of ROS affecting each NPs behavior vary and ROS generation by cells is indeed a slow reaction, the level of ROS as a function of time was determined up to 24 h with and without NAC/LPS (Figure 7a). The ROS level was improved almost twofold in contact with LPS compared to NPs themselves proving the generation of higher ROS using LPS on cells with all three NPs. As expected, NAC scavenged the production of ROS in all NPs which agrees with the results obtained in Figures 4 and 6. It is worth mentioning that even though the LPS and NAC's concentrations were not toxic in the range of study, their incubation with NPs enhanced the cell cytotoxicity that could affect the level of ROS (Figure 7b). The highest ROS level was observed after 24 h in AS-PEG. This is in line with the results obtained in Figures 4c and 6g-i.

One should point out that the change in the luminescent of AS NPs depends on the coating, surface functional groups, NPs size, and the effect of ROS and H₂O₂.[4] However, it should be noted that we achieved strong fluorescence intensity in AS-PEG and AS-11MUA over time. These AS NPs can be used for long-term studies especially in cancer treatment since the production of ROS is higher in cancer cells and the NPs can be track in NIR-II.

## 4. Conclusions

A one-step microwave-assisted synthesis produced aqueous AS NPs with a TEM size under 6 nm and NIR-II emission (950-1200 nm). Studies on fibroblast uptake showed that AS NPs with a PEG coating achieved the highest cellular uptake due to their small hydrodynamic size, followed by AS-11MUA and AS-DTDTPA. Long-term PL tracking over 9 d revealed distinct luminescence behavior: AS-DTDTPA and AS-11MUA showed initial PL decreases post-internalization but later increased, whereas
AS-PEG exhibited sustained PL enhancement. However, the pattern did not correspond with the observed luminescence. In order to understand the reason behind that, the effect of H₂O₂ in solution on the PL spectra of AS NPs were studied thoroughly.

To investigate this, the effects of H₂O₂ on AS NP PL were examined. H₂O₂ exposure reduced AS-DTDTPA and AS-11MUA luminescence due to (i) oxidation/detachment of unbound—SH groups, observed as Ag–S doublet reduction and increased thiol peaks in XPS, and (ii) formation of electron-hole traps reducing lifetime and increasing its non-radiative contribution (A₁). By contrast, AS-PEG luminescence increased, likely due to (i) H₂O₂ elimination of metallic Ag (evident from reduced Ag doublets in XPS) and (ii) increased lifetime with enhanced radiative recombination (A₂). Overall, PL enhancement in fibroblasts was directly linked to H₂O₂ and ROS interactions within cells. It is worth mentioning that no substantial changes in particle organization, size, or structure were observed in cells over 9 d; however, we did observe significant variation in the PL spectra. AS-DTDTPA and AS-PEG coatings demonstrate potential as ROS scavengers and ROS promoters, respectively.

## Supporting Information

Supporting Information is available from the Wiley Online Library or from the author.

## Acknowledgements

The authors acknowledge the Plateforme MIMA2 (INRA, Jouy en Josas, France) for electron microscopy preparation and observation, Samia Hidalgo (IPGP: Institut de physique du globe de Paris) for inductively coupled plasma mass spectrometry (ICP-MS). The authors acknowledge the following grants for financial support: the European Commission for providing funding for this project under the Horizon 2020 research and innovation program for Marie Skłodowska-Curie Individual Fellowship, with the acronym "SANTE" and project number: 101110756 and French Research Agency: Agence Nationale de la Recherche (ANR-21-CE09-0030 NANOTHERMOMETRY), the IdEx Université Paris Cité, ANR-18-IDEX-0001 (IVETH plateform), by the Region Ile de France under the convention SESAME 2019—IVETH (EX047011) (IVETH plateform).

## Conflict of Interest

The authors declare no conflict of interest.

## Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

## Keywords

Ag₂S, H₂O₂, photoluminescence, quantum dots, reactive oxygen species

Received: May 7, 2025

Revised: July 9, 2025

Published online:

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