Mesoporous polydopamine@CeO₂ nanoparticles for photoacoustic-guided photothermal therapy in suppressing arthritis

Summary

Rheumatoid arthritis (RA) is characterized by poor oxygen supply and overproduction of reactive oxygen species (ROS), thereby exacerbating synovial inflammation. We synthesized MPDA@CeO₂ nanoparticles (NPs) by incorporating mesoporous polydopamine (MPDA) NPs and CeO₂ for photoacoustic imaging (PAI)-guided synergistic light-controlled PTT/oxygen-boosting/ROS-eliminating therapy. Upon irradiation with a near-infrared (NIR) laser, MPDA@CeO₂ NPs trigger photothermal conversion to eradicate inflammatory cells and release CeO₂, which have a catalytic activity to eliminate ROS and release oxygen. The oxygen-release and ROS-depleting efficiency was simultaneously augmented by heat to further combat the harmful environment of inflammation. We validated the therapeutic effect of MPDA@CeO₂in vitro and in vivo. We further applied multiwavelength PAI to observe the distribution of NPs and the change in oxygen saturation during the treatment process in vivo. The study highlights the therapeutic value of MPDA@CeO₂ in inhibiting arthritis in RA through imaging-guided synergistic therapy.

Graphical abstract

Highlights

• •MPDA@CeO₂ NPs offer a potential therapy for RA via PTT/ROS/oxygen synergy
• •Photoacoustic imaging tracks NP distribution and oxygenation dynamics in inflamed joints
• •Heat-triggered CeO₂ release alleviates oxidative stress and hypoxia in RA microenvironment

Health sciences; Biophysics; Materials science

Introduction

Rheumatoid arthritis (RA) is an autoimmune, systemic, chronic inflammatory disease characterized by persistent arthritis and bone erosion.¹ Prompt and effective treatment is of paramount importance for patients with RA to avoid detrimental consequences such as constant pain, irreversible joint dysfunction, and impaired work ability.² However, due to the unknown etiology and heterogeneous clinical manifestations of the disease, the management of the disease remains a challenge. Disease-modifying antirheumatic drugs (DMARDs) are the first-line therapeutic strategy to achieve the goals of RA treatment, including remission and low disease activity.³ However, failure of long-term inflammation treatment and drug discontinuation because of accompanying side effects of DMARDs have become major clinical challenges in the treatment of RA.⁴ There is still a lack of effective methods to control severe systemic and local inflammation in patients with RA. In this regard, it is necessary to develop new treatment strategies for arthritis based on the pathophysiological characteristics.

Tissue hypoxia, the decline in oxygen saturation, is a hallmark of inflammatory arthritis in RA. In the thickened synovium of RA, the proliferation of inflammatory cells increases the consumption of oxygen. The synovial microvasculature is highly malformed and dysfunctional, which jeopardizes oxygen supply. Hence, tissue hypoxia in inflamed synovium is the consequence of an imbalance between increased oxygen demand (due to increased metabolism) and reduced oxygen supply (due to vascular dysfunction). Researchers have performed direct measurements of oxygenation of the synovial tissues in vivo both in animal models and RA patients, which validated the decrease in oxygen pressure in inflamed synovium in RA.^5,6,7,8 The hypoxic microenvironment of synovial tissues further aggravates the abnormal synovial proliferation and cellular dysfunction, thereby exacerbating inflammation activity. Previous studies have also identified the overexpression of hypoxia-inducible factor (HIF-1) in RA synovitis, the primary regulator in response to altered cellular oxygen pressure, which is associated with the production of inflammation cytokines and bone and cartilage erosions.⁹ In light of its role in RA, hypoxia has served as a therapeutic target for RA.¹⁰

It has also been proven that the hypoxic environment of synovial tissues is involved in the generation of reactive oxygen species (ROS), a category of detrimental metabolites that aggravate synovitis and structural destruction.¹¹ The overproduction of ROS can cause oxidative damage to cellular components and molecules, such as DNA damage, protein inactivity, and mitochondria damage.^12,13 In addition, ROS can provoke the production of inflammatory cytokines and the differentiation of immune cells, thereby further intensifying inflammation.^14,15,16 Previous studies have also shown that ROS can be a sensitive indicator of disease activity and treatment response.^17,18,19 Accordingly, hypoxia alleviation and ROS elimination may be regarded as possible targets for the therapy of RA.

Another efficient therapeutic target for RA is the elimination of inflammatory cells. The activation of innate inflammatory cells, such as T and B lymphocytes, dendritic cells, macrophages, and neutrophils—which aggregate in the infected joints in response to autoimmune factors—mainly contributes to the chronic inflammatory process and subsequent joint and bone destruction.^20,21 These inflammatory cells secrete proinflammatory cytokines and other chemical compounds, resulting in a cascade of antagonistic pathophysiological events such as angiogenesis, synovial thickening, and bone erosion.²² Thus, it is critical in RA treatment to inhibit inflammatory cells and inhibit the release of cytokines.

Versatile nanomaterials have been extensively studied in the context of RA treatment. Not only can they serve as vehicles for anti-inflammatory agents but they can also offer therapeutic advantages, such as photodynamic therapy (PDT), photothermal therapy (PTT), immunotherapy, and their combinations. Nanomaterials are also distinguished by their imaging characteristics, which facilitate imaging-guided theranostic treatment strategies for RA.²³ PTT is a noninvasive treatment method based on the photothermal effect of nano-photosensitizers, which has shown promise in treating RA. PTT is generated when photosensitizers convert light to heat in a dosage- and time-dependent manner. External laser irradiation with adjustable range and dose enables accurate targeting of tumors to minimize harm to adjacent tissues. The photosensitizers for PTT commonly have photoacoustic (PA) characteristics that can be visualized by PA imaging (PAI), a novel imaging technique for clinical use.^24,25,26 Polydopamine (PDA) is a broadly utilized nanopolymer that possesses high photothermal conversion efficiency, which makes it a perfect material to use for PTT, but also to produce thermal reaction shift of phase transformation materials for light-controlled drug release.²⁷ The reductive functional groups of PDA also enable it to scavenge ROS, making it an excellent choice for designing anti-inflammatory nanomaterials.²⁸

Apart from PTT, catalase-like nanomaterials that are designed to improve H₂O₂-rich hypoxic milieu and scavenge ROS have also been utilized to suppress inflammation. Ceria (CeO₂) is one of these nanomaterials with regenerative antioxidant properties for anti-inflammation.^29,30,31 Of note, CeO₂-induced oxygen release can be further enhanced through increased temperature that is produced by the PTT effect.

In this study, we combined mesoporous PDA (MPDA) NPs, a kind of biocompatible nano-photosensitizers with high drug-loading efficiency, and CeO₂ to produce MPDA@CeO₂ NPs for PAI-guided synergistic light-controlled PTT/oxygen-boosting/ROS-eliminating therapy. MPDA NPs constituted the core of MPDA@CeO₂, and CeO₂ NPs were deposited within the pores of the mesoporous structures. Upon irradiation with a near-infrared (NIR) laser, MPDA@CeO₂ NPs exhibited excellent photothermal conversion to eradicate inflammatory cells and release CeO₂. The oxygen-release and ROS-depleting efficiency was simultaneously augmented by heat to further combat the harmful environment of inflammation. We validated the therapeutic effect of MPDA@CeO₂ in vitro and in vivo. In addition, making use of the PA characteristics of the NPs and the function of PAI in monitoring oxygenation, we applied multiwavelength PAI to observe the NPs and their oxygen-release efficiency during the treatment process. After injecting the NPs into inflamed tissues, the localization of the NPs was visualized by PAI. Tissue oxygenation was enhanced subsequently, which could also be monitored in vivo by PAI. Therefore, MPDA@CeO₂ can be deemed as a theranostic nanomaterial that facilitates both imaging guidance and synergistic therapy for treating RA.

Results and discussion

Preparation and characterization of MPDA@CeO₂ NPs

Uniformly sized MPDA@CeO₂ NPs were first synthesized using the aqueous phase method. The loading of hydrophobic CeO₂ into MPDA was achieved through physical adsorption. This mechanism allows the hydrophobic CeO₂ nanoparticles to adhere to the MPDA matrix without the need for additional chemical reactions or stabilizers. When MPDA@CeO₂ NPs are subjected to external laser irradiation, the nanoparticles absorb the laser energy and convert it into heat, resulting in a temperature increase within the nanoparticle structure. This heat generation disrupts the stability of the nanoparticles, causing structural changes and partial disintegration, thus releasing CeO₂. The transmission electron microscopy (TEM) imaging revealed a high degree of dispersion of MPDA@CeO₂ NPs (Figures 1A and 1B). We employed X-ray diffraction (XRD) to investigate the crystal phases of MPDA@CeO₂ nanoparticles (Figure 1C). According to the XRD patterns showing the crystal phases of NPs, the synthesized MPDA@CeO₂ displayed diffraction peaks corresponding to the cubic CeO₂ phase’s (111) plane (Figure 1C).

Figure 1.

Characterization of MPDA@CeO₂ NPs

(A) Transmission electron microscopy (TEM) imaging of MPDA, CeO₂, and MPDA@CeO₂ NPs, scale bar: 200 &100 nm; (B) X-ray diffraction (XRD) of MPDA@CeO₂ NPs; (C) Diameter distribution of MPDA, CeO₂, and MPDA@CeO₂ nanoparticles measured by dynamic light scattering (DLS). Data are represented as mean ± SEM.

Dynamic light scattering (DLS) was used to determine the diameter of MPDA@CeO₂ particles (Figure 1D). The hydrodynamic particle size of MPDA was 235.1 ± 2.7 nm, the size of CeO₂ was 10.3 ± 1.7 nm, and the size of MPDA@CeO₂ was 256.3 ± 10.3 nm, indicating that CeO₂ NPs were successfully loaded onto the mesoporous structure of MPDA. The polydispersity index (PDI) of MPDA, CeO₂, and MPDA@CeO₂ was 0.233 ± 0.006, 0.141 ± 0.003, and 0.167 ± 0.06, respectively, indicating the high dispersibility of the nanoparticles. The zeta potential of MPDA@CeO₂ was −17.2 ± 1.073 mV. And the zeta potential chart of MPDA@CeO₂ is illustrated in Figure 1. The element mapping of MPDA@CeO₂ NPs is shown in Figure S1, highlighting the distribution of elements, including cerium (Ce), carbon (C), hydrogen (H), and oxygen (O). The result verified the composition of MPDA and CeO₂ of MPDA@CeO₂ NPs.

Photothermal and photoacoustic properties of MPDA@CeO₂

As shown in Figure 2A, MPDA@CeO₂ NPs exhibited a high extinction peak at 810 nm in ultraviolet-visible (UV-vis) spectroscopy. The absorbance of MPDA@CeO₂ in UV-vis spectroscopy increased with the NPs’ concentration, indicating the surface plasmon resonance absorption of MPDA. We further investigated the plasmonic photothermal properties of MPDA@CeO₂ under the NIR laser with a wavelength of 808 nm, laser energy of 1 W/cm⁻², and duration time of 5 min. When irradiated by an 808nm NIR laser, the temperature of the MPDA@CeO₂ solution increased rapidly and reached a peak at about 45°C in 5 min. The increase in temperature exhibited a time-dependent, power density–dependent, and dosage-dependent trend, which also suggested a good photothermal effect of MPDA@CeO₂ (Figures 2B–2D and 2G). The photothermal conversion efficiency of MPDA@CeO₂ was about 37.87%. Five cycles of heating and cooling tests were conducted to determine the photothermal reversibility of MPDA@CeO₂ (Figure 2C). The photothermal performance of MPDA@CeO₂ remained unchanged after repeated NIR laser irradiation, indicating excellent photothermal stability of MPDA@CeO₂ (Figure 2E).

Figure 2.

Plasmonic photothermal properties of MPDA@CeO₂ NPs

(A) UV-vis spectroscopy of MPDA@CeO₂ at different concentrations.

(B) Heat-generating ability of MPDA@CeO₂ NPs with different times and dosages.

(C) Five cycles of heating and cooling tests of MPDA@CeO₂.

(D) The photothermal performance of MPDA@CeO₂ with different laser energies.

(E–G) (E) Temperature change of MPDA@CeO₂ NPs after laser irradiation for 15 min (F and G) Heatmap of the photothermal performance of MPDA@CeO₂ NPs. (E) The linear fitting plot of time versus -Ln(θ) according to the cooling period in (F). (G) Infrared thermal images of MPDA-CeO₂ NP at different power densities and concentrations.

We further assessed the PA property of MPDA@CeO₂ at two wavelengths, namely 750 and 830 nm using a multimodal PA/US system (Mindray Biomedical Technology Corporation, Shenzhen, China). The PA signals at the two wavelengths were also quantified by calculating the pseudo-color pixels. MPDA@CeO₂ NPs showed significant PA signals at both wavelengths, which also had a concentration-dependent tendency, as shown in Figure S3.

H₂O₂-decomposing and oxygen-producing capacity of MPDA@CeO₂ NPs

Methylene blue is a dye that has a strong absorbance peak at 664 nm. When it interacts with reactive species, such as those produced during the decomposition of H₂O₂, its concentration decreases, leading to a reduction in absorbance. MPDA@CeO₂ NPs are designed to have catalytic properties. Cerium oxide can catalyze the decomposition of H₂O₂ into water and oxygen. When H₂O₂ is decomposed, it generates ROS which can interact with methylene blue. Therefore, we can evaluate the H₂O₂-decomposing capacity of MPDA@CeO₂ NPs by measuring the absorbance of methylene blue at 664 nm. We performed the H₂O₂ assay and tested oxygen bubble production of MPDA@CeO₂, MPDA, and CeO₂ with or without laser irradiation at a pH of 7.4. The solution became more colorless with the increase in reaction time, and the height of the absorption peak of methylene blue at 664 nm gradually decreased. Figure 3 illustrates that the H₂O₂-decomposing capacity of MPDA@CeO₂ NPs increased with concentration and time and could also be augmented by heating. We found a gradual increase in the dissolved-oxygen concentration in the solution after the reaction of MPDA@CeO₂ NPs and H₂O₂, which also exhibited a positive correlation with the concentration of H₂O₂.

Figure 3.

Detection of H₂O₂-decomposing and oxygen-producing capacity of MPDA@CeO₂ NPs

(A) Absorption spectra of methylene blue for different concentrations of H₂O₂ solution reacting with MPDA@CeO₂ NPs for 1 h. With the increase in H₂O₂ concentration, the peak of methylene blue gradually decreased. The decomposing capacity of MPDA@CeO₂ NPs increased after heating.

(B) Absorption spectra of methylene blue for 160 mM H₂O₂ solution with MPDA@CeO₂ NPs for 1, 2, and 4 h. With the increase in reacting time, the peak absorbance gradually decreased. The absorbance after heating also decreased.

(C) Photographs of the mixture of H₂O₂ and MPDA@CeO₂ NPs of different concentrations exposed to methylene blue. The color faded more with the increase in concentration and heating.

(D) Detection of dissolved-oxygen concentration in the solution of H₂O₂ and MPDA@CeO₂ NPs of different concentrations. The dissolved-oxygen concentration increased with the reaction time, and solutions with a higher concentration of H₂O₂ presented with a higher oxygen concentration.

Cytotoxicity and PTT of MPDA@CeO₂ NPs in vitro

For the in vitro experiment, we utilized the macrophage cell line RAW 264.7, which can release inflammatory cytokines when activated and has been regarded as the representative cell line in studies about RA. We induced cell inflammation by adding lipopolysaccharide (LPS) to RAW 264.7 cells and examined the PTT effect of MPDA@CeO₂ NPs with laser irradiation on the LPS-induced cells. The cytotoxicity of MPDA@CeO₂, MPDA, and CeO₂ was enhanced with laser exposure (IC₅₀ values: 42.05 μg/mL of MPDA@CeO₂ + laser, 583.2 μg/mL of MPDA + laser, 64.8 μg/mL of CeO₂ + laser). MPDA@CeO₂ with laser irradiation presented the highest cytotoxicity among these groups. We also observed cell death using calcein AM staining for live cells and propidium iodide staining for dead cells under confocal laser scanning microscopy (CLSM). For the cells treated with phosphate-buffered saline (PBS), PBS + laser, and MPDA, only green fluorescence was observed. Mainly green fluorescence and sparse red fluorescence were found in the cells treated with MPDA@CeO₂ and CeO₂, suggesting the good biocompatibility of MPDA@CeO₂ NPs without laser exposure. Red fluorescence increased in all NPs after laser irradiation, and the cells treated with MPDA@CeO₂ and laser irradiation presented the strongest red fluorescence. The quantified results of the ratio of red fluorescence also verified the greatest cellular growth inhibition of MPDA@CeO₂ with laser (Figure 4).

Figure 4.

Cell viability of MPDA@CeO₂ NPs

(A) Cell viability of CeO₂, MPDA, and MPDA@CeO₂ NPs with laser in different concentrations.

(B) Calcein AM staining and propidium iodide staining of RAW 267.4 cells treated with PBS, CeO2, MPDA, and MPDA@CeO2 NPs with or without laser. Scale bar: 20 μm.

(C) Quantification of cell viability of AM-PI staining. Data are represented as mean ± SEM. ∗: p < 0.05; ∗∗: p < 0.01; ∗∗∗: p < 0.001; ∗∗∗∗: p < 0.0001 (Unpaired two-tailed Student’s t test).

H₂O₂-decomposing and proinflammatory-inhibition effect of MPDA@CeO₂in vitro

The ROS-scavenging capacity of MPDA@CeO₂ NPs was determined by measuring the level of H₂O₂ in the cells. We used an H₂O₂ assay and quantified the fluorescence intensity, which represented intracellular H₂O₂ concentrations, to evaluate the H₂O₂-decomposing rate of different NPs. As demonstrated in Figure 5, showing the H₂O₂ fluorescence and the quantitative results of fluorescence intensity, the cells treated with MPDA and CeO₂ showed a slight decrease in green fluorescence intensity compared with the cells treated with PBS and PBS with laser. After laser irradiation, the fluorescence intensity of the cells treated with MPDA, CeO₂, and MPDA@CeO₂ decreased. The cells treated with MPDA@CeO₂ and laser had the lowest H₂O₂ concentration.

Figure 5.

H₂O₂-decomposing and proinflammatory-inhibition effects of MPDA@CeO₂in vitro

(A and B) H₂O₂ assay of RAW 267.4 cells treated with PBS, CeO₂, MPDA, and MPDA@CeO₂ NPs with or without laser; Scale bar: 20 μm. (C–F) ELISA of HIF-1 α, IL-6, and TNF-α; (G–I) western blot of HIF-1 α, IL-6, and TNF-α in vitro. Data are represented as mean ± SEM. ∗: p < 0.05; ∗∗: p < 0.01; ∗∗∗: p < 0.001; ∗∗∗∗: p < 0.0001 (Unpaired two-tailed Student’s t test).

Then, HIF-1α and proinflammatory cytokines (TNF-α and IL-6) were assessed using western blot (WB) and enzyme-linked immunosorbent assay (ELISA), as shown in Figure 5. The two measurements showed a similar trend in the cytokine levels. With laser exposure, the cytokine levels decreased in the cells treated with MPDA, CeO₂, and MPDA@CeO₂. Specifically, the cells treated with MPDA@CeO₂ with laser irradiation had the lowest HIF-1α, TNF-α, and IL-6 expression. HIF-1α is closely related to the hypoxic environment of the inflammatory tissues. Thus, the decrease in HIF-1α indicated that MPDA@CeO₂ helped improve the hypoxic status of inflammatory tissues. The decrease in TNF-α and IL-6, two crucial inflammatory cytokines that are related to clinical manifestations, also revealed the role of MPDA@CeO₂ in regulating cytokines and suppressing inflammation. These in vitro experiments validated the ability of MPDA@CeO₂ to scavenge ROS and inhibit proinflammatory cytokine expression when exposed to laser irradiation.

PTT and oxygen generation effects of MPDA@CeO₂in vivo

The PTT effect and oxygen production capability of MPDA@CeO₂ were tested in vivo on an adjuvant-induced arthritis (AIA) rat model. The rats presented arthritis on day 21 after injection of 0.2 mg complete Freund’s adjuvant. To explore the in vivo PTT effect of the NPs, the AIA rats received an intra-articular injection of MPDA, CeO₂, and MPDA@CeO₂ solutions, under the 808-nm laser irradiation with the same conditions (laser energy of 1 W/cm⁻², time duration of 5 min). During the laser irradiation, we recorded the thermal images of the inflamed joints. As shown in Figures 5A and 5B, after 5 min of 808-nm laser irradiation at the target region, the temperature of the inflamed joint climbed rapidly to approximately 131°F in the rats treated with MPDA@CeO₂, to about 107°F in the rats treated with MPDA, and to 106°F in the rats treated with CeO₂. The temperature of the control group did not change significantly. The results suggested that MPDA@CeO₂ also presented a good photothermal effect in vivo, which was in line with the in vitro results of the photothermal property.

We performed dual-wavelength PAI on the AIA rats before and after the laser irradiation. Oxygenated hemoglobin and deoxygenated hemoglobin have similar light absorption in the infrared range, which varies markedly at the wavelength of 600–700 nm. Hence, we chose 750 and 830 nm wavelengths of PAI to measure the tissue oxygenation. In the PA system applied in the study, the ratio of PA signal pixels at the two wavelengths was calculated to obtain the relative oxygen saturation (SO₂) value. The SO₂ values were represented as pseudo-colors in the SO₂ mapping section of the PA system, including red for relatively high oxygenation and blue for relatively low oxygenation. Due to the PA property of the NPs, the injected NPs could also be visualized at 750 and 830 nm. MPDA@CeO₂ and MPDA turned out to be bright blue signals on the SO₂ mapping section, while CeO₂ was less obvious on SO₂ mapping.

After 6 min of laser irradiation, the red signals within the inflamed tissues of the AIA rats treated with MPDA@CeO₂ remarkably increased and subsided after 30 min. The rats injected with CeO₂ had a modest increase in red signals inside the inflamed tissues, which diminished after 30 min (Figure 6). We also quantified the area of blue. Upon laser irradiation, the local temperature of the inflamed tissue rose, thereby boosting blood flow, and oxygen was released during the process, increasing blood oxygen and thus the photoacoustic red signal in the area. The distribution of NPs and oxygen release was successfully monitored by PAI.

Figure 6.

Thermal changes and photoacoustic imaging of AIA rats treated with PBS, CeO₂, MPDA, and MPDA@CeO₂ NPs with or without laser

(A and B) Thermal changes in AIA rats treated with NPs with or without laser in 6 min.

(C and D) Photoacoustic imaging of the AIA rats after injection of nanoparticles in 0, 5, 10, 20, and 30 min.

We observed the metabolism of MPDA@CeO₂ NPs in vivo through PAI. As shown in Figure S4, a prominent red signal was observed in the subcutaneous soft tissue layer of the rats’ left paw according to the PA images. Subsequently, the signal gradually diminished, and by day 14 the PA signal disappeared from the soft tissue, concentrating only on the skin layer. After day 21, the signal completely disappeared from the tissues of the AIA rats.

The therapeutic effect of MPDA@CeO₂ NPs in vivo

The ROS-scavenging effect of the NPs was evaluated by examining the ROS production in the treated tissues. The inhibition of hypoxia was verified by measuring the expression level of HIF-1α, the most important hypoxia-related factor. Proinflammatory cytokines, including TNF-α and IL-6, are also hallmarks of RA and cause clinical manifestations and detrimental outcomes. The reduction in the levels of these molecules could validate the anti-inflammatory effect of MPDA@CeO₂ NPs with laser irradiation. The ROS production was examined by quantifying the fluorescence intensity of 7-dichlorofluorescein diacetate (DCFH-DA) of the collected tissues. The expression levels of HIF-1α, TNF-α, and IL-6 were examined through ELISA and immunohistochemistry (IHC).

As shown in Figure 7, the inflammatory tissues treated with CeO₂, MPDA, and MPDA@CeO₂ with laser irradiation showed the lowest score of histological inflammation. They had a reduction in ROS production compared with the groups that did not receive laser irradiation, indicating that both MPDA and CeO₂ NPs could scavenge ROS. The tissues treated with MPDA@CeO₂ and laser irradiation had the lowest ROS production. The results suggested that the capacity of ROS scavenging improved after combining the two NPs.

Figure 7.

In vivo assessment of the therapeutic effect of MPDA@CeO₂ with laser irradiation

(A and B) HE staining and SO staining results of the articular tissues.

(C–E) IHC results of TNF-α, IL6, and HIF-1α levels in the inflammatory tissue of the AIA rats. The rats treated with MPDA@CeO₂ NPs and laser irradiation showed the lowest HIF-1α and TNF-α expression levels. Scale bar: 50 μm.

(F) Histological scores of the AIA rats.

(G) ROS production within the inflammatory tissues of the AIA rats that received NPs with or without laser irradiation.

(H and I) Quantitative IHC and ELISA results of TNF-α, IL6, and HIF-1α levels. Data are represented as mean ± SEM. ∗: p < 0.05; ∗∗: p < 0.01; ∗∗∗: p < 0.001; ∗∗∗∗: p < 0.0001 (Unpaired two-tailed Student’s t test).

HIF-1α, TNF-α, and IL-6 levels decreased with the addition of NIR laser irradiation of the rats injected with CeO₂, MPDA, and MPDA@CeO₂ according to the ELISA and IHC analysis. The MPDA@CeO₂ with laser irradiation group also had the lowest expression in the analyses. The NPs also exhibit good biocompatibility, according to Figure S5, which shows no significant changes in liver and kidney function indicators in AIA rats before and after the injection of the NPs. The hematoxylin-eosin (HE) staining of the major organs of the rats showed that no damage was caused to these organs, as shown in Figure S6. Overall, MPDA@CeO₂ NPs displayed good efficacy in suppressing inflammation, improving hypoxia, and scavenging ROS in inflammatory joints under the guidance of PAI, indicating its utilization potential for treating RA.

Limitations of the study

In this study, we injected the nanoparticles intra-articularly to treat monoarthritis. Materials with active targeting ability for treating systemic inflammation should be developed in further studies. Also, the therapeutic effect was validated by in vivo animal study. Further biological safety assessment is needed for clinical application.

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Meng Yang (yangmeng_pumch@126.com, yangmeng@pumch.cn).

Materials availability

This study did not generate new unique reagents.

Data and code availability

The original datasets generated and analyzed in this study will be made available from the lead contact upon reasonable request. No code was generated in this study. Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

Acknowledgments

This work was supported by National Natural Science Foundation of China (62325112, U22A2023, and 82302207); Ministry of Science and Technology of the People's Republic of China; National Key Research and Development Program of China (2023YFC2411700 and 2023YFC2411705); Non-profit Central Research Institute Fund of Chinese Academy of Medical Sciences (2024-RC320-02); National High-Level Hospital Clinical Research Funding (2022-PUMCH-C-009, 2022-PUMCH-B-064, and 2022-PUMCH-D-002); China Postdoctoral Science Foundation (2022TQ0044 and 2024M763745).

We would like to express our sincere gratitude for the research support provided by Theranostics and Translational Research Facility, the National Infrastructures for Translational Medicine, Peking Union Medical College Hospital, Chinese Academy of Medical Sciences and Peking Union Medical College. We thank LetPub (www.letpub.com) for its linguistic assistance during the preparation of this manuscript.

Author contributions

Conceptualization, L.H.Z., W.J.J., Z.C.Y., Y.M., and H.Y.L.; methodology, L.H.Z., W.J.J., Z.C.Y., Z.R., and W.M.; investigation, L.H.Z., W.J.J., and Z.C.Y.; writing – original draft, L.H.Z., W.J.J., and Z.C.Y.; writing–review and editing, Y.M., J.Y.X., S.D.S., and H.Y.L.; funding acquisition, Y.M., H.Y.L., L.H.Z., and Z.C.Y.; resources, J.Y.X., Y.M., S.D.S., and H.Y.L.; supervision, Y.M., J.Y.X., S.D.S., and H.Y.L.

Declaration of interests

The authors have declared that no competing interest exists.

STAR★Methods

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
Anti-HIF-1α	Abcam	ab247533; RRID: AB_3696415
Anti-TNFα	Abcam	ab220210; RRID: AB_2892586
Anti-IL-6	Abcam	ab9324; RRID: AB_307175
GAPDH	Abcam	ab9485; RRID: AB_307275
β-actin	Abcam	ab8226; RRID: AB_306371
Biological samples
Synovial tissues in mice	This paper	N/A
Chemicals, peptides, and recombinant proteins
PBS	Gibco	70011044
PVDF membrane	Thermo Fisher	LC2007
BSA	MERCK	V900933-100G
WST	beyotime	S0103
MTT solution	beyotime	C0009S
calcium AM and PI	beyotime	C2012
SDS buffer	Sigma-Aldrich	S3401
Critical commercial assays
SOD assay kit	Sigma-Aldrich	19160-1KT-F
H2O2 assay kit	Sigma-Aldrich	ab138874
HIF-1α ELISA kit	Abcam	ab275103
TNFα ELISA kit	Abcam	ab46070
IL-6 ELISA kit	Abcam	ab234570

Experimental model and study participant details

Mouse models

The AIA rat model is a classical animal model mimicking the autoimmune of RA. We used female Wistar rats aged 6–8 weeks (Beijing Vital River Laboratory Animal Technologies Co., Ltd., Beijing, China) for modeling. The animals were kept in an environment maintained at a temperature of 22 ± 1°C, with a relative humidity of 50 ± 1% and a light/dark cycle of 12 h each. The animals were provided with sterilized standard rodent chow and autoclaved water ad libitum by trained technicians. Female rats were selected for arthritis model to mimic the higher prevalence of RA in human females. The study protocol was approved by the Institutional Animal Care and Use Committee of the animal experiment center our institution (BJLongan-L-L-0002). The in vivo study was conducted in accordance with the guidelines and regulations for animal experiments at our institution.

On the first day of modeling, the left hindlimb paws of the rats were injected with 0.2 mg complete Freund’s adjuvant (CFA, Sigma-Aldrich, St. Louis, MO, USA) intradermally. On Day 21 after the injection, it was determined whether arthritis had been successfully established in the rats by identifying redness and swelling of the left paws of rats.

Cell line culture

The in vitro validation of cellular uptake of siRNA was tested on the murine macrophage cell line RAW 264.7 (National Infrastructure of Cell Line Resource, Shanghai, China). The cell line was authenticated via STR profiling (16 loci) by the supplier using Cellosaurus database. Cells were cultured in DMEM (Gibco, USA) supplemented with 10% fetal bovine serum (FBS, Gibco, USA) and 1% double-antibody (Gibco, USA). Cells were incubated at 37°C in a 5%CO2 humidified atmosphere.

Method details

Synergy of MPDA@CeO₂

First, MPDA nanoparticles were synthesized using a one-pot method. First, 0.36 g of surfactant F127 and 0.36 g of 1,3,5-Trimethylbenzene (TMB) were dissolved in a mixture of 60 mL of ethanol and 65 mL of water and stirred for 30 min. Then, 90 mg of trimethylolaminomethane (TRIS) and 60 mg of dopamine hydrochloride were added and stirred at room temperature for 24 h. Subsequently, the solution was centrifuged, and the precipitate was washed with ethanol and acetone. After washing, the sample was placed in a mixed solution of ethanol and acetone (2:1 v/v) and subjected to ultrasonic treatment three times. Next, 1 mmol of cerium (III) acetate and 12 mmol of oleylamine were dissolved in 15 mL of xylene, and the mixture was stirred at room temperature for 2 h and then heated to 90°C. After adding 1 mL of deionized water, the aging solution was kept at 90°C for 3 h and then cooled to room temperature. The obtained CeO₂ nanoparticles were washed with ethanol. MPDA@CeO₂ nanoparticles were synthesized using the aqueous phase method. Different amounts of CeO2 nanoparticles were mixed with 20 mg of MPDA nanoparticles in 40 mL of deionized water. The mixture was stirred overnight at room temperature and filtered to obtain the precipitate. The precipitate was washed with ethanol and water, and dried under vacuum conditions at 70°C. MPDA@CeO₂ nanoparticles with an MPDA to CeO₂ ratio of 1:0.8 were selected for further experiments, because it achieves a saturation loading of the CeO2 NPs under this ratio. Increasing the ratio beyond 1:0.8 would lead to an oversaturation of CeO₂, while decreasing the ratio would result in a lower loading capacity.

Characterization of the NPs

A high-resolution transmission electron microscope (HRTEM, JEM-1400, HITACHI, Japan)) operated at 200 kV was used to examine the size, morphology, and element mapping of the nanoparticles. A dynamic light scattering instrument (Zetasizer Nano-ZS, Malvern Instruments) was used to measure the hydrodynamic diameter of MPDA, CeO₂, and MPDA@CeO₂ NPs. Powder XRD patterns of MPDA@CeO₂ NPs were obtained on an AXS D8 advance (Bruker, Germany). The loading efficiency of MPDA@CeO₂ NPs was measured by thermogravimetric analysis (Thermogravimetric Analyzer TGA55, Discovery).

Evaluation of photothermal and photoacoustic properties of MPDA@CeO₂ NPs

A spectrophotometer (UV-1750) (Shimadzu, Kyoto, Japan) was used to measure the UV-vis-NIR absorption spectra of the NPs. Then the laser irradiation for photothermal effect was conducted using the following settings: NIR wavelength of 808 nm, laser energy of 1 W/cm⁻², and time of 5 min. We examined six different concentrations of MPDA@CeO₂ NPs under laser irradiation (0, 12.5, 25, 50, 100 and 200 μg/mL). During the evaluation of the photothermal effect, the temperature changes of the NP solution were recorded by a thermal camera, and the specific temperature value was presented on the screen.

SOD-catalytic activity, H₂O₂ decomposing, and oxygen production

We measured the SOD-catalytic activity of the NPs using a SOD assay kit (Sigma-Aldrich) according to the manufacturer’s protocol. We diluted 1 mL of water-soluble tetrazolium salt (WST) solution into 19 mL of buffer solution to prepare a water-soluble tetrazolium salt (WST) working solution. We also prepare an enzyme working solution of 15 μL of enzyme solution and 2.5 mL of dilution buffer. SOD activity was determined via the creation of water-soluble formazan dye when reducing WST with a superoxide anion. WST working solution and enzyme working solution were added at a concentration of 300 μg/mL to MPDA, CeO₂, and MPDA@CeO₂ NPs, and were incubated at room temperature for 20 min. Using a microplate reader to measure the absorbance at 450 nm, the slope of the inhibition rate was compared to the slope of the blank to determine the SOD activity.

At room temperature, 200 μg/mL of MPDA, CeO₂, and MPDA@CeO₂ NPs were mixed with 100 μM H₂O₂, respectively. The H₂O₂ concentration was determined using an H₂O₂ assay kit (Sigma-Aldrich) by measuring the solution’s absorbance at 405 nm with or without 808 nm laser irradiation.

We also dissolved 200 g/mL of MPDA, CeO₂, and MPDA@CeO₂ NPs with 100 μM H₂O₂ solution. Oxygen production levels were monitored every 10 min in the grey-scale modality of an ultrasound system (Mindray Corporation, Shenzhen, China) with or without 808 nm laser irradiation.

Cytotoxicity and PTT of MPDA@CeO₂ nanoparticles in vitro

Cytotoxicity assay

MTT test was used to determine cytotoxicity in vitro. We cultivated RAW 264.7 cells for 24 h. Then, varied concentrations of MPDA@CeO₂ solutions (0, 12.5, 25, 50, 100, and 200 μg/mL) were added to the cells and incubated for 24 h at 37°C. Subsequently, a 10% MTT solution was applied and incubated for 4 h at 37°C. Then the MTT solution was discarded and 200 L of DMSO was added into each well for 20 min to dissolve the action products. The IC₅₀ values were determined using a microplate reader to determine the absorbance at 490 nm.

The LPS-induced RAW264.7 macrophages were treated with 200 g/mL PBS, MPDA, CeO₂, and MPDA@CeO₂, with or without 808 nm laser irradiation, respectively. After washed by PBS, the cells received 808 nm laser irradiation for 5 min, using the same conditions as described above. The MTT test was used to examine the cytotoxicity of the NPs. Next, live and dead cells were stained with calcium fluorescein (AM) and PI, and cell pictures were captured using a laser confocal fluorescence microscope (Olympus, Tokyo, Japan), to further validate the photothermal effect of MPDA@CeO₂ in killing inflammatory cells.

Intracellular H₂O₂ decomposing

We used a hydrogen peroxide assay kit (ab138874, Abcam, Sigma-Aldrich) to measure the H₂O₂ decomposing efficiency of MPDA@CeO₂ NPs. Firstly, we added 100 μM of H₂O₂ to the cells. Then the cells were incubated with 200 μg/mL PBS, MPDA, CeO₂, and MPDA@CeO₂ NPs for 24 h, with or without 808 nm laser irradiation, respectively. The 808 nm laser was implemented on the cells for 5 min. After incubating the cells for 3 h, the cellular H₂O₂ concentrations were assessed by the H₂O₂ assay. We used confocal fluorescence microscopy to identify the H₂O₂ concentrations with Ex/Em of 490/520 nm.

Intracellular inflammatory cytokines reducing

The cellular levels of HIF-1α, TNF-α, and IL-6 were tested by qPRC, western blot, and ELISA, based on the manufacturer’s protocols.

Western bolts were conducted according to the method of Hnasko (1). Sodium dodecyl sulfate (SDS) buffer was mixed with the samples, boiled for 10 min, and loaded onto gel wells. Then polyacrylamide gel electrophoresis (PAGE) (Power Supplies Basic, Bio-Rad, USA was used for separating the proteins, which were transferred on a polyvinylidene fluoride (PVDF) membrane. The membrane was then treated with antibodies (ab247533, ab220210, ab9324, Abcam, Aobsen Biochem Co., Ltd., Shanghai, China). The bands were analyzed using a blotting detection system (SYNGENE G: BOXChemiXR5, Eppendorf, Germany) and a gel documentation system (Gel-Pro32, Bio-Rad, Hercules, CA, USA).

ELISA kits (RTA00, R&D Systems, MN, USA) were used to measure the procytokines’ concentrations according to the manufacturer’s instructions. The cells were added to the microplates that were coated with antibodies, and incubated for 90 min at 37°C. Then detection antibodies were added to wells and incubated for 2 h at room temperature. Horseradish peroxidase (HRP) was added after washing with PBS to detect binding protein indirectly. Absorbance was measured at 450 nm. Dilution folds were used to adjust the results to final concentrations.

In vivo synergistic photothermal/oxygen releasing/ROS scavenging therapeutic effect

AIA rat model and treatment grouping

The rats were randomly allocated to six groups, which were treated with 200 μL of (1) PBS, (2) PBS + laser, (3) MPDA, (4) MPDA + laser, (5) CeO₂, (6) CeO₂ + laser, (7) MPDA@CeO₂, (8) MPDA@CeO₂, + laser intra-articularly. The settings of laser irradiation include 808 nm of laser, 1 W/cm² of energy, and a treatment duration of 5 min. There were totally six rats in each group. During the 5 min of laser irradiation, the thermal images of the rats every 30 s were acquired using a thermal camera. The AIA rats received a total of three times of laser irradiation on days 21, 23, and 25.

PAI of NPs and tissue oxygenation

We used a multimodal photoacoustic/ultrasound system that incorporated a commercial ultrasound machine (Resona 7, Shenzhen Mindray Biomedical Electronics Co., Ltd., Shenzhen, China) and an optical parametric oscillator laser (Spotlight 600-OPO and Innolas laser GmbH). The rats received PAI before injection of NPs, immediately after injection, and 0, 5, 10, 20, and 30 min after laser irradiation. We selected 750 nm and 830 nm similar scattering coefficients to calculate SO₂. The PA-SO₂ image was pseudocolored and superimposed on the US image on the PA/US system. Red and blue pixels represented high and low SO₂ values, respectively. An onboard algorithm was designed to quantify the SO₂ values and PA signal areas automatically after tracing the region of interest (ROI) of the synovium.

SO₂ is the percentage of the hemoglobin bound to oxygen to the total hemoglobin, of which the formula is:

$$S{O}_2=Hb{O}_2/\left(Hb{O}_2+Hb\right)\times 100\%$$

The light absorption of oxyhemoglobin (HbO2) and deoxyhemoglobin (Hb) varies markedly at the wavelength of 600∼700 nm and becomes identical in the infrared range. In this system, the SO₂ values were calculated using the signals of 750 nm (λ₁) and 830 nm (λ₂), expressed as the following equation:

$$S{O}_2=\frac{C_{HbO2}}{C_{Hb}+C_{HbO2}}=\frac{{\epsilon }_{Hb}^{{\lambda }_1}{A}^{{\lambda }_2}-{\epsilon }_{Hb}^{{\lambda }_2}{A}^{{\lambda }_1}}{A^{{\lambda }_1}\left({\epsilon }_{HbO2}^{{\lambda }_2}-{\epsilon }_{Hb}^{{\lambda }_2}\right)+A^{{\lambda }_2}\left({\epsilon }_{Hb}^{{\lambda }_1}-{\epsilon }_{HbO2}^{{\lambda }_1}\right)}\times 100\%$$

(CHb and CHbO₂: Hb and HbO₂ content; ${\epsilon }_{Hb}^{\mathrm{\lambda }1}$, ${\epsilon }_{Hb}^{\mathrm{\lambda }2}$, ${\epsilon }_{HbO2}^{\mathrm{\lambda }1}$, ${\epsilon }_{HbO2}^{\mathrm{\lambda }2}$: extinction coefficients of Hb and HbO₂ at λ1 and λ2; A^λ1 A^λ2: photoacoustic intensity at wavelengths λ₁ and λ₂).

Pathological examinations of inflammatory cytokines

Detection of ROS production in the collected tissues

First, we applied the enzymatic digestion method to prepare single-cell suspension. The collected tissues were washed with PBS to remove the necrotic components, fibers, fat, and blood vessels of the paws. Then the tissues were cut into small pieces with a diameter of about 1 mm. The pieces were washed with PBS and added to an enzymatic digestion solution at the temperature of 37°C for 20–30 min. The content was shaken intermittently during the digestion. After digestion, the tissues were filtered with a 00-mesh nylon net to collect the cells. The cells were centrifuged at 500 g for 10 min, and the precipitate was kept and washed 1–2 times with PBS. The cells were then diluted with DCFH-DA to a density of 1×10⁶ to 2×10⁷/mL. Then the cells were incubated at 37°C for 30 min, and the mixture was inverted every 5 min to ensure full contact between the probe and cells. The collected suspension was centrifuged at 1000 g for 10 min, and the precipitate was collected and washed 1–2 times with PBS. The cells were resuspended in PBS for fluorescence detection using a Microplate System (Multiskan go, Thermo, USA). The optimal excitation wavelength is 488 nm, and the emission wavelength is 525 nm.

ELISA

The concentrations of cytokines in the harvested tissues were also measured by ELISA using ELISA kits (RTA00, R&D Systems, MN, USA) according to the manufacturer’s instructions.

IHC evaluation

The harvested samples were fixed in formalin, decalcified for 6 h, and embedded in paraffin. The slides were deparaffinized and blocked in methanol. HE staining was performed first. After being treated with 5% bovine serum albumin (BSA, Aobsen Biochem, Shanghai, China), the samples were incubated with antibodies (ab228649, ab220210, ab9324, Abcam, Aobsen Biochem Co., Ltd., Shanghai, China) for 2 h at 37°C. Next, the secondary antibodies were added to the samples for 30 min at room temperature and incubated with High Sensitivity Streptavidin-HRP conjugate (HSS-HRP) for 30 min. The tissues were then covered with diaminobenzidine/aminoethylcarbazole (DAB/AEC) substrate. The staining was observed using a microscope (Olympus, Tokyo, Japan), and positive cells were counted.

Quantification and statistical analysis

IBM SPSS Statistics for Windows (version 20.0; IBM Corp, Armonk, NY, USA) was used to perform the statistical analysis. The quantitative data was expressed as continuous variables (mean ± standard error of the mean [SEM]) and analyzed using the Mann–Whitney U test. When comparing more than two independent samples, the Kruskal-Wallis test was used. A p value less than 0.05 was considered as statistical significance.

Published: May 3, 2025