Abstract

   Rationale: Inflammatory osteolysis, characterized by abundant immune
   cell infiltration and osteoclast (OC) formation, is a common
   complication induced by bacterial products and/or wear particles at the
   bone-prosthesis interface that severely reduces long-term stability
   after implantation. Molecular nanoclusters are ultrasmall particles
   with unique physicochemical and biological properties that have great
   potential as theranostic agents for treating inflammatory diseases.

   Methods: In this study, heterometallic PtAu[2] nanoclusters with
   sensitive nitric oxide-responsive phosphorescence turn-on
   characteristics and strong binding interactions with cysteine were
   designed, making them desirable candidates for the treatment of
   inflammatory osteolysis.

   Results: PtAu[2] clusters exhibited satisfactory biocompatibility and
   cellular uptake behavior, with potent anti-inflammatory and anti-OC
   activities in vitro. In addition, PtAu[2] clusters alleviated
   lipopolysaccharide-induced calvarial osteolysis in vivo and activated
   nuclear factor erythroid 2-related factor 2 (Nrf2) expression by
   disrupting its association with Kelch-like ECH-associated protein 1
   (Keap1), thereby upregulating the expression of endogenous
   anti-inflammatory and anti-oxidative products.

   Conclusion: Through the rational design of novel heterometallic
   nanoclusters that activate the endogenous anti-inflammatory system,
   this study provides new insights into the development of
   multifunctional molecular therapeutic agents for inflammatory
   osteolysis and other inflammatory diseases.

   Keywords: Metal cluster, Alloy cluster, Osteolysis, Osteoclast, Nrf2

Introduction

   Total joint replacement (TJR) is an effective surgical modality for
   treating end-stage joint pathologies such as osteoarthritis, rheumatoid
   arthritis, and traumatic arthritis [45]^1^, [46]^2. Although TJR can
   improve articular morphology, function, pain relief, and quality of
   life, approximately 3-5% of patients experience postoperative aseptic
   loosening and periprosthetic infection [47]^3. These complications can
   be induced by bacterial products and/or wear particles generated by
   prosthesis components, leading to inflammatory osteolysis [48]^4.
   Acute/chronic inflammatory reactions and bone erosion at the
   bone-prosthesis interface caused by bacterial products such as
   lipopolysaccharides (LPS) can severely reduce the long-term stability
   of prostheses. Indeed, patients with inflammatory osteolysis may
   require multiple revision surgeries and even experience implant failure
   [49]^5. In addition to surgical intervention, medications such as
   bisphosphonates and non-steroidal anti-inflammatory drugs (NSAIDs) are
   often required to prevent the progression of inflammation and bone
   erosion. However, over-administration can induce adverse side effects,
   making it highly important to investigate new curative strategies for
   periprosthetic inflammatory osteolysis [50]^6.

   Osteolysis commonly occurs around areas of inflammation and is
   characterized by the infiltration of abundant osteoclasts (OCs) and
   immune cells [51]^7. As a principal source of inflammatory cytokines,
   macrophages play a central role in the pathogenesis of osteolysis and
   can differentiate into OCs, the main effector cells of osteolysis
   [52]^8. Accumulating evidence suggests that inflammatory cytokines such
   as tumor necrosis factor-α (TNF-α), interleukin (IL)-1β, IL-6, and
   inducible nitric oxide synthase (iNOS) may stimulate OC formation, even
   in the absence of the receptor activator of NF-κB ligand (RANKL)
   [53]^9^-[54]^12. It has also been reported that RANKL and TNF-α exert
   synergistic effects on osteoclastogenesis, with low RANKL levels
   directly stimulating OC precursors to become mature OCs [55]^13. IL-1β
   is another cytokine that is abundant in inflamed tissues and can
   enhance inflammatory osteolysis by cooperating with TNF-α to stimulate
   the expression of RANKL and its receptor, RANK [56]^14. This clear
   interdependency between inflammatory cytokines and RANKL suggests the
   importance of blocking both the inflammatory reactions and
   osteoclastogenesis in an osteolytic environment.

   In recent years, nanotherapies have attracted increasing attention
   because of their beneficial characteristics, including accurate
   targeting, responsive release, and flexible functionalization [57]^15.
   Molecular nanoclusters are ultrasmall particles with core-shells of
   less than 3 nm [58]^16. Unlike large nanoparticles, nanoclusters have
   precise molecular structures, uniform molecular sizes, good
   biocompatibility, facile synthesis, and promising biomedical
   applications [59]^17^-[60]^19. Nanocluster cores can be homometallic
   (e.g., Au, Ag, Cu, and Pt) or heterometallic with discrete energy
   levels, which can endow nanoclusters with unique chemical, biological,
   optical, and electronic properties [61]^16. For example, Au clusters
   have been widely studied for the diagnosis and therapy of inflammation,
   cancer, implant-associated infections, and COVID-19 [62]^20^-[63]^28,
   whereas heterometallic or alloy nanoclusters, such as Pt-Au clusters,
   have been reported to improve the physicochemical and biological
   performance of cancer therapies compared to homometallic nanoclusters
   [64]^29^, [65]^30. The enhanced biomedical activity of heterometallic
   or alloy clusters is probably due to their improved stability and
   structural diversity resulting from the complementary advantages of
   different metal ions, which can favorably bind to proteins and enzymes
   and exert biological effects. Consequently, alloy nanoclusters have
   great potential as theranostic agents for treating inflammatory
   diseases.

   In this study, we designed a platinum(II)-gold(I) (PtAu[2]) nanocluster
   protected by bis(diphenylphosphinomethyl)phosphine as an effective
   anti-inflammatory and anti-osteolytic agent (Figure [66]1A). Owing to
   its d^8-d^10 heterometallic nature, the ultrasmall PtAu[2] cluster
   (PtAu[2]-1) with electron-rich 4-ethynylbenzene-1,2-diamine acts as a
   highly-sensitive luminescence sensor in response to nitric oxide (NO),
   producing distinct phosphorescence turn-on characteristics that allow
   the detection of inflammation, as well as showing strong binding
   interactions with cysteine. In addition, the PtAu[2] clusters exhibited
   satisfactory biocompatibility and cellular uptake behavior as well as
   potent anti-inflammatory and anti-OC effects in vitro and alleviated
   LPS-induced calvarial osteolysis in vivo. More importantly, PtAu[2]
   clusters activated nuclear factor erythroid 2-related factor 2 (Nrf2)
   expression by disrupting its association with Kelch-like ECH-associated
   protein 1 (Keap1), thereby increasing the release and nuclear
   translocation of Nrf2, which upregulates the expression of endogenous
   anti-inflammatory and anti-oxidative products (Figure [67]1B). Unlike
   other reported nanoscale anti-inflammatory systems with complicated
   synthetic procedures, PtAu[2] clusters can activate endogenous
   anti-oxidative systems without excess drug loading and modification.
   Taken together, our findings provide new insights into the development
   of multifunctional molecular therapeutic agents for inflammatory
   osteolysis and other inflammatory diseases.

Figure 1.

   [68]Figure 1
   [69]Open in a new tab

   Schematic representation of the design, synthesis, and potential
   application of PtAu[2] clusters to treat inflammatory osteolysis. (A)
   Synthesis of PtAu[2] clusters. (B) Application of PtAu[2] clusters to
   manage inflammatory osteolysis by disrupting the Keap1-Nrf2
   association, increasing the release and nuclear translocation of Nrf2,
   and upregulating the expression of endogenous anti-inflammatory and
   anti-oxidative products (created with BioRender.com).

Materials and methods

Reagents and antibodies

   Escherichia coli-derived lipopolysaccharide (LPS) was obtained from
   InvivoGen (San Diego, CA, USA). Recombinant mouse RANKL and macrophage
   colony-stimulating factor (M-CSF) were purchased from R&D (Minneapolis,
   MN, USA). Penicillin, streptomycin, and minimal essential medium alpha
   (α-MEM) were purchased from Gibco BRL (Gaithersburg, MD, USA). Fetal
   bovine serum (FBS) was purchased from Avantor (Radnor, PA, USA). The
   Cell Counting Kit-8 (CCK-8) was obtained from Beyotime Biotechnology
   (Shanghai, China). Methylisothiourea sulfate (SMT) was purchased from
   Selleck (Houston, TX, USA). Dihydroethidium (DHE) was purchased from
   Med Chem Express (Shanghai, China). The Prime Script RT reagent Kit and
   TB Green^TM Premix Ex Taq^TM II were obtained from Takara Biomedical
   Technology (Beijing, China). The primary antibody against iNOS
   (Thermofisher #PA1-036) was purchased from Thermofisher Scientific
   (Waltham, MA, USA). Primary antibodies against phospho-p65 (CST #3033),
   p65 (CST #8242), Keap1 (CST #4678), Nrf2 (CST #12721), HO-1 (CST
   #26416), and NQO1 (CST #62262) were purchased from Cell Signaling
   Technology (CST, Danvers, MA, USA). Primary antibodies against β-Actin
   (Affinity #AF7018), GAPDH (Affinity #AF7021), NFATc1 (Affinity
   #DF6446), CTSK (Affinity #DF6614), and TNF-α (Affinity #AF7014) were
   purchased from Affinity Biosciences (Cincinnati, OH, USA).

General procedures

   All syntheses were conducted under dry argon using Schlenk technique
   and vacuum-line system. All solvents were dried and distilled from
   appropriate drying agents prior to use. 1,2-Diamine-4-iodobenzene,
   di-tert-butyl dicarbonate ((BOC)[2]O) and other starting materials were
   commercially available without further purification.
   Bis(di-o-tolylphosphino-methyl)phenylphosphine (dTolmp) was prepared by
   the synthetic procedure described in the literature [70]^31.

Physical measurements

   The ^1H and ^31P NMR spectra were performed on a Bruker Avance III (400
   MHz) and ECZ600R (600 MHz) spectrometer with SiMe[4] as the internal
   reference and H[3]PO[4] as the external reference, respectively.
   Splitting patterns are designated as singlet (s), doublet (d), and
   triplet (t). Splitting patterns that could not be interpreted or easily
   visualized are designated as multiplet (m) and broad (br). The UV-Vis
   absorption spectra were measured on a Perkin-Elmer Lambda 35 UV-Vis
   spectrophotometer using a 10 mm path quartz cell. The high-resolution
   mass spectrometry (HRMS) was conducted on an Impact II mass
   spectrometer using dichloromethane and methanol mixtures as mobile
   phases. The emission and excitation spectra together with the emissive
   lifetimes were measured on Edinburgh FLS-920 fluorescence spectrometer.
   The emission spectra and lifetimes were measured upon excitation at 397
   nm. The morphology of the material was observed by a field emission
   transmission electron microscope (FETEM, FEI Tecnai F20).

Crystal structural determination

   The X-ray single-crystal diffraction data of PtAu[2]-1a were collected
   on a Bruker D8 Venture diffractometer using IμS 3.0 microfocus source
   Mo-Kα radiation (λ = 0.71073 Å) and PHOTON II CPAD detector. Frames
   were integrated with the Bruker SAINT software package (V8.38A) using a
   SAINT algorithm. Data were corrected for absorption effects using the
   multi-scan method (SADABS) [71]^32. The structure was solved and
   refined using the Bruker SHELXTL Software Package, a computer program
   for automatic solution of crystal structures, and refined by the
   full-matrix least-squares method with ShelXle Version 4.8.6, a Qt
   graphical user interface for the SHELXL [72]^33. All non-hydrogen atoms
   were refined anisotropically, whereas the hydrogen atoms were generated
   geometrically and refined using isotropic thermal parameters. CCDC
   2211858 contains the supplementary crystallographic data for this
   paper. These data can be obtained free of charge from the Cambridge
   Crystallographic Data Centre via www.ccdc.cam.ac.uk/data_request/cif.

Titration of PtAu[2]-1 cluster with NO

   Sodium nitroprusside (Na[2][Fe(CN)[5]NO]), a widely used exogenous NO
   donor [73]^34^, [74]^35, was dissolved in distilled water as a 10 mM
   stock solution immediately before the experiments to avoid direct light
   exposure. The titration experiment was performed by addition of one- to
   eleven-fold aqueous solution of Na[2][Fe(CN)[5]NO] to dimethyl
   sulfoxide (DMSO)-phosphate-buffered saline (PBS) (v/v = 1:1) solution
   of PtAu[2]-1 cluster (the concentration of 1.0 × 10^-5 M). The
   phosphorescent sensing behavior of PtAu[2]-1 cluster to nitric oxide
   was monitored by the measurement of the emission spectrum change at a
   concentration of 1.0 × 10^-5 M in DMSO-PBS (v/v = 1:1) solution upon
   excitation at 398 nm.

Titration of PtAu[2]-1 cluster with L-cysteine

   To investigate the possible binding interaction of PtAu[2] cluster with
   cysteine, the titration experiment was performed by addition of one- to
   sixteen-fold aqueous solution of L-cysteine to a DMSO-PBS (v/v = 1:1)
   solution of PtAu[2]-1 cluster (the concentration of 1.0 × 10^-5 M). The
   binding interaction was monitored through the measurement of UV-Vis
   absorption and emission spectroscopy.

Cellular uptake assay

   RAW 264.7 murine macrophages were seeded in confocal dishes, then
   incubated with 100 ng/mL LPS plus 20 μg/mL PtAu[2] clusters for 12 h.
   Then, cells were incubated with Hoechst 33342 (C1027; Beyotime
   Biotechnology, Shanghai, China) for nuclear staining. Meanwhile, a
   commercial kit for NO detection (DAF-FM DA) was used for comparison,
   according to the manufacturer's instructions. After several times of
   rinsing with warm PBS, the cells were observed in confocal laser
   scanning microscopy (CLSM).

   For evaluation of lysosome colocalization, the cells were stained with
   Lyso-Tracker Red for another 30 min at 37 °C. After rinsing several
   times, the colocalization of PtAu[2] clusters and endo/lysosomes was
   observed via CLSM.

Quantitative real‑time PCR analysis

   To analyze the mRNA level of pro-inflammatory genes, RAW 264.7 murine
   macrophages were cultured in 6-well plates at a density of 5 × 10^5
   cells/well, then stimulated with LPS (100 ng/mL) plus various
   concentrations of PtAu[2] clusters (5, 10, and 20 μg/mL) for 24 h. To
   evaluate the mRNA level of OCs-related genes, bone marrow macrophages
   (BMMs) were cultured in 6-well plates at a density of 2 × 10^5
   cells/well supplemented with M-CSF (30 ng/mL) and RANKL (100 ng/mL), in
   the presence of various concentrations of PtAu[2] clusters (5, 10, and
   20 μg/mL) for 4 days. The total RNA of each group was extracted by
   using an Axygen RNA Miniprep Kit (Axygen, Union City, CA, USA),
   according to the manufacturer's instructions. Reverse transcription was
   performed based on RNA templates. TB Green^TM Premix Ex Taq^TM II was
   used for a real-time PCR (RT-qPCR) assay on an ABI 7500 Sequencing
   Detection System (Applied Biosystems, Foster City, CA). 5 μL of TB
   Green, 3 μL of ddH[2]O, 1 μL of cDNA, 0.4 μL of each primer and 0.2 μL
   ROX Dye2 were mixed to establish a 10 μL reaction system. Cycling
   conditions were set as 40 cycles (95 °C for 5 s and 60 °C for 30 s).
   The melting curves were checked to verify the amplification
   specificity. The relative gene expression was calculated using the
   comparative 2^-ΔΔCT method, as described previously [75]^36. The
   housekeeping gene was set as Gapdh. The primer sequences are listed in
   [76]Table S1.

Immunofluorescence staining

   RAW 264.7 murine macrophages were cultured with LPS (100 ng/mL) in the
   presence of various concentrations of PtAu[2] clusters (5 and 20 μg/mL)
   for 24 h. Then, the cells were fixed, permeabilized, and blocked. After
   that, the cells were incubated with primary antibody against iNOS
   (1:100; PA1-036; Thermofisher, Waltham, MA, USA), p-p65 (1:100; CST
   #3033; CST, Danvers, MA, USA) and Nrf2 (1:100; CST #12721; CST,
   Danvers, MA, USA). The second antibodies for immunofluorescence were
   incubated for another 1 h. DAPI was used for nuclei staining. Then, the
   cells were detected via CLSM. The relative fluorescence intensity was
   analyzed by ImageJ software (NIH, Bethesda, MD, USA).

Enzyme-linked immunosorbent assay (ELISA)

   RAW 264.7 murine macrophages were cultured with LPS (100 ng/mL) in the
   presence of various concentrations of PtAu[2] clusters (5, 10, and 20
   μg/mL) for 24 h. Then the supernatants were collected, and the TNF-α,
   IL-1β, IL-6, and IL-10 contents of the supernatants were detected
   according to the ELISA kit instructions (Ruixin Biotech, Fujian,
   China).

Detection of reactive oxygen species (ROS)

   RAW 264.7 murine macrophages were seeded in confocal dishes, and
   cultured with 100 ng/mL LPS plus 20 μg/mL PtAu[2] clusters for 24 h.
   After that, the cells were incubated with 5 μM DHE probe (HY-D0079; Med
   Chem Express, China) for 30 min at 37 °C. Hoechst 33342 was used for
   nuclear staining. After 3 times rinsing with warm PBS to remove the
   excess dye, the cells were detected via CLSM. ImageJ software (NIH,
   Bethesda, MD, USA) was used for semi-quantitative analysis of relative
   ROS fluorescence.

GSH/GSSG measurement

   GSH and GSSG Assay Kit (S0053; Beyotime Biotechnology, Shanghai, China)
   was used for the measurement of intracellular GSH/GSSG level, referred
   to the manufacturer's instructions. Briefly, RAW 264.7 murine
   macrophages were cultured in 6-well plates at a density of 5 × 10^5
   cells/well, then stimulated with LPS (100 ng/mL) plus various
   concentrations of PtAu[2] clusters (5, 10, and 20 μg/mL) for 24 h. The
   cells were collected and lysed, then frozen and thawed in liquid
   nitrogen and 37 °C repetitively. After centrifugation, the supernatant
   of each sample was collected to measure total glutathione. The sample
   absorbance was measured at a wavelength of 412 nm, with the standard
   curve used for further calculation. The formula was shown as follow:
   GSH = Total Glutathione - GSSG × 2.

Malonaldehyde (MDA) measurement

   Cellular Lipid Peroxidation MDA Assay Kit (A003-4-1; Nanjing Jiancheng
   Bioengineering Institute, Jiangsu, China) was used for the measurement
   of intracellular MDA level, referred to the manufacturer's
   instructions. Briefly, RAW 264.7 murine macrophages were cultured in
   6-well plates at a density of 5 × 10^5 cells/well, then stimulated with
   LPS (100 ng/mL) plus various concentrations of PtAu[2] clusters (5, 10,
   and 20 μg/mL) for 24 h. The cells were collected, lysed, and
   centrifuged at 12,000 g for 15 min. After that, the supernatant of each
   sample was collected for MDA measurement. The sample absorbance was
   measured at a wavelength of 532 nm.

Superoxide Dismutase (SOD) measurement

   Total Superoxide Dismutase Assay Kit with WST-8 (S0053; Beyotime
   Biotechnology, Shanghai, China) was used for the measurement of SOD
   activity. Briefly, RAW 264.7 murine macrophages were cultured in 6-well
   plates at a density of 5 × 10^5 cells/well, then stimulated with LPS
   (100 ng/mL) plus various concentrations of PtAu[2] clusters (5, 10, and
   20 μg/mL) for 24 h. The cells were collected by 0.25% trypsin for
   subsequent experiments. The SOD activity was measured strictly
   according to the manufacturer's instructions. The results were
   presented as SOD activity relative to the control group, which was set
   at 100%.

Tartrate-resistant acid phosphatase (TRAP) staining assay

   BMMs were seeded into 96-well plates at a density of 8 × 10^3
   cells/well, then cultured with M-CSF (30 ng/mL) and RANKL (100 ng/mL),
   in the presence of various concentrations of PtAu[2] clusters (2.5, 5,
   and 10 μg/mL). The culture medium was replaced every 2 days until the
   formation of OCs was observed on day 5. After a 20 min fixation with 4%
   paraformaldehyde (PFA), the cells were stained with TRAP staining
   solution at 37 °C for 1 h. OCs were defined as TRAP-positive cells with
   more than three nuclei. An optical microscope (Olympus, Tokyo, Japan)
   was used for image capturing. The OCs number and area were quantified
   by ImageJ software (NIH, Bethesda, MD, USA).

Detection of podosome actin belt

   BMMs were cultured with M-CSF (30 ng/mL) and RANKL (100 ng/mL), in the
   presence of various concentrations of PtAu[2] clusters (2.5, 5, and 10
   μg/mL). After mature OC formation, the cells were fixed and
   permeabilized, then incubated with TRITC Phalloidin (1:100; CA1610;
   Solarbio, Beijing, China) for 40 min. After that, the nuclei were
   stained with DAPI for 5 min. The cells were washed with PBS three
   times, then observed via CLSM. The number of podosome actin belts was
   quantified by ImageJ software (NIH, Bethesda, MD, USA).

Bone resorption assay

   At a density of 1 × 10^4 cells/well, BMMs were seeded into 96-well
   Corning Osteo Assay Surface plates (Corning, NY, USA), then cultured
   with M-CSF (30 ng/mL) and RANKL (100 ng/mL), in the presence of various
   concentrations of PtAu[2] clusters. The culture medium was replaced
   every 2 days until the formation of mature OCs. After culture for
   another 1 day, the mature OCs were eliminated by 5% sodium
   hypochlorite. The resorption pits were imaged using an optical
   microscope. The bone resorption area was analyzed by ImageJ software
   (NIH, Bethesda, MD, USA).

RNA sequencing

   RNA sequencing was performed to explore the potential mechanisms for
   the anti-inflammatory effect of PtAu[2] clusters. Briefly, RAW 264.7
   murine macrophages were seeded in 6-well plates at a density of 5 ×
   10^5 cells/well, then cultured with 100 ng/mL LPS plus 20 μg/mL PtAu[2]
   clusters for 24 h. TRIzol reagent was used to collect the total RNA.
   The gene expression level was measured by Biomarker Technologies
   (Beijing, China). Gene ontology (GO) enrichment analysis and Kyoto
   Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis
   were performed using BMKCloud (www.biocloud.net).

Immunoblotting

   For RAW 264.7 murine macrophages, the cells were seeded in 6-well
   plates at a density of 5 × 10^5 cells/well, then stimulated with 100
   ng/mL LPS plus various concentrations of PtAu[2] clusters (5, 10, and
   20 μg/mL) for 24 h. For BMMs, the cells were seeded in 6-well plates at
   a density of 3 × 10^5 cells/well with M-CSF (30 ng/mL) and RANKL (100
   ng/mL), then treated in the presence or absence of 10 μg/mL PtAu[2]
   clusters for 0, 1, 3 and 5 days. Then, the total protein of each sample
   was extracted by using whole cell lysis buffer with a protease
   inhibitor cocktail (P8340; Sigma-Aldrich, Shanghai, China). After the
   lysates were centrifuged at 12,000 g for 15 min, the supernatant of
   each sample was collected. Nuclear proteins were extracted via the
   nuclear extraction kit (Thermofisher, Waltham, MA, USA) by high-speed
   centrifugation, according to the manufacturer's instructions. Protein
   concentrations were measured using Bicinchoninic acid (BCA) assay
   (P0012; Beyotime Biotechnology, Shanghai, China). Then, the proteins
   were dissolved in the loading buffer, separated by 10% SDS-PAGE, and
   transferred to 0.22 µm polyvinylidene fluoride (PVDF) membranes. After
   1 h blocking in 5% BSA in 1 × TBST (Tris-buffered saline with Tween 20)
   at room temperature, the membranes were incubated with the primary
   antibodies (β-Actin, 1:1000; GAPDH, 1:1000; iNOS, 1:1000; p-p65,
   1:1000; p65, 1:1000; Keap1, 1:1000; Nrf2, 1:500; HO-1, 1:1000; NQO1,
   1:1000) overnight at 4 ºC. After that, the secondary antibodies were
   incubated with membranes for 1 h at room temperature. Odyssey V3.0
   image scanning (Li-COR. Inc., Lincoln, NE, USA) was used for blot
   detection. The blot results were analyzed and quantified by ImageJ
   software (NIH, Bethesda, MD, USA).

Immunoprecipitation

   For each sample, a total of 600 μg protein lysates were pre-incubated
   with IgA/G beads (Sigma, Shanghai, China). Anti-Keap1 antibody (Keap1,
   1:50) and protein IgA/G beads were used to precipitate with endogenous
   Keap1. Then, the Keap1-Nrf2 immuno-complex was subjected to Western
   blotting.

LPS-induced calvarial osteolysis model

   The Animal Care and Experiment Committee of Ninth People's Hospital
   Affiliated to Shanghai Jiao Tong University School of Medicine
   (SH9H-2022-A874-1) approved the animal protocol. All experiments were
   performed according to the guidelines for Ethical Conduct in the Care
   and Use of Nonhuman Animals in Research by the American Psychological
   Association. The murine calvarial osteolysis model was used to evaluate
   the in vivo therapeutic effect of PtAu[2] clusters on inflammatory
   response and bone resorption, based on the previous reports [77]^37^,
   [78]^38. Briefly, fifteen 8-week-old C57/BL6 male mice (approximate
   weight 25 ± 2 g) were randomly divided into three groups with five
   animals per group: (1) Sham group (with PBS injection); (2) LPS group
   (10 mg/kg body weight of LPS and PBS injection); (3) PtAu[2] group (10
   mg/kg body weight of LPS and 20 μg/mL PtAu[2] clusters injection).
   Collagen sponges (4 mm × 4 mm × 2 mm) soaked with PBS or LPS were
   implanted in the middle of the calvaria under general anesthesia. 3
   days after implantation, all the mice were subject to subcutaneous
   injections of calvarias every other day over another 11-day period.
   After experiments, the mice were euthanized. The whole calvaria was
   harvested and washed with PBS, then fixed in 4% PFA for 48 h for
   radiographic and histological analysis.

Micro-computed tomography

   Micro-computed tomography (CT) scanning was performed using a
   high-resolution micro-CT (μCT-100, SCANCO Medical AG, Switzerland). The
   resolution of the scanning was 10 μm; the X-ray energy was set at 70
   kv, 200 μA; and a fixed exposure time was 300 ms. The microstructure
   indicators of bone volume/tissue volume (BV/TV), trabecular number
   (Tb.N), trabecular separation (Tb.Sp), and trabecular number (Tb.Th)
   were measured in a three-dimensional region of interest (ROI) using
   evaluation analysis software (Version: 6.5-3, SCANCO Medical AG,
   Switzerland).

Histological and immunohistochemical analysis

   After micro-CT scanning, the samples were decalcified in 10% EDTA (pH =
   7.4) for 4 weeks, then embedded in paraffin. Histological sections were
   prepared for hematoxylin and eosin (H&E) and TRAP staining.
   Immunofluorescence staining was performed with antibodies against TNF-α
   (1:100; AF7014; Affinity, Cincinnati, OH, USA), Nrf2 (1:100; AF0639;
   Affinity, Cincinnati, OH, USA), and HO-1 (1:100; AF5393; Affinity,
   Cincinnati, OH, USA). The images of stained slices were captured under
   a high-quality microscope (Leica DM4000B). The relative fluorescence of
   stained slices was quantified by ImageJ software (NIH, Bethesda, MD,
   USA). The number of OCs and TRAP-positive multinucleated OCs per field
   (Oc.S/BS) were calculated.

In vivo biosafety evaluation

   14 days after model establishment, the major organs of mice were
   collected for H&E staining. The systemic toxicity of PtAu[2] clusters
   was evaluated based on the H&E staining of the heart, liver, spleen,
   lung, and kidney. Meanwhile, blood routine examinations and serum
   biochemical analyses were performed by collecting the blood samples.

In vivo biodistribution of PtAu[2] clusters

   To detect the biodistribution and metabolism of PtAu[2] clusters in
   major organs, 8-week-old C57/BL6 male mice (n = 3) were subcutaneously
   injected with PtAu[2] clusters on calvarias at a dose of 100 μg/mL (100
   μL). Two days after injection, mice were sacrificed and the major
   organs were harvested. Then, the organ tissues were weighed,
   homogenized, and dissolved in aqua regia. Inductively coupled plasma
   mass spectrometry (ICP-MS) was used to measure the amounts of Au and Pt
   in each sample, and the percentage of injected dose per gram of tissue
   (%ID g^-1) was calculated.

Statistical Analysis

   The results were analyzed by using the Prism 8.0 statistical software
   package (GraphPad Software Inc, San Diego, CA, USA). The data were
   uniformly presented in the form of mean ± standard deviation (SD).
   Differences between the two groups were analyzed using the two-tailed
   unpaired Student's t-test, after the homogeneity test of variance.
   One-way analysis of variance (ANOVA) with Tukey's post hoc tests was
   used for multiple group comparisons. Significant differences were
   determined to be at *p < 0.05 and **p < 0.01.

Results and Discussion

Synthesis and characterization of PtAu[2] clusters

   Because 4-ethynylbenzene-1,2-diamine is highly sensitive to air,
   PtAu[2]clusters were first synthesized using t-butyloxy carbonyl (BOC)
   protected ethynyl ligands. As shown in Figure [79]1A and [80]Scheme S1,
   the BOC-protected PtAu[2]-1a cluster was prepared by reacting the
   mononuclear Pt(II) precursors,
   bis(di-o-tolylphosphino-methyl)phenylphosphine (dTolmp),
   chloro(tetrahydrothiophene)gold(I) (Au(tht)Cl), and NaSO[3]CF[3] in a
   1:2:2:2 molar ratio at room temperature [81]^39^, [82]^40. The
   structure of the BOC-protected PtAu[2]-1a cluster ([83]Table S2) was
   determined using single crystal X-ray diffraction, which confirmed that
   the PtAu[2] framework was supported by double dTolmp ligands. The
   PtAu[2] cluster ([84]Figure S1) is highly stabilized by four
   five-membered chelating rings, as well as significant d^8-d^10
   heterometallic interactions as the Pt-Au distance (2.9245(2) Å) is much
   shorter than the sum of the van der Waals radii for Pt and Au (ca. 3.3
   Å). Removal of the BOC protective groups gave the target PtAu[2]-1
   cluster in 45% yield (Figure [85]2A). Both the PtAu[2]-1 and
   BOC-protected PtAu[2]-1a clusters were fully characterized using ^1H
   and ^31P NMR spectrometry ([86]Figures S2, S3, S5, and S6) and
   high-resolution mass spectrometry ([87]Figures S4 and S7). Transmission
   electron microscopy (TEM) further indicated that the PtAu[2]-1 clusters
   were ultrasmall and widely dispersed, with a uniform nanomorphology and
   a diameter of approximately 2 nm (Figure [88]2B).

Figure 2.

   [89]Figure 2
   [90]Open in a new tab

   Characterization of PtAu[2] clusters. (A) Coordination structure of the
   PtAu[2]-1 cluster. (B) TEM image of the PtAu[2]-1 clusters. (C)
   Excitation and emission spectra of the BOC-protected PtAu[2]-1a and the
   BOC-removed PtAu[2]-1 clusters at a concentration of 1.0 × 10^-5 M in
   DMSO-PBS (v/v = 1:1). (D) Schematic diagram showing the phosphorescent
   turn-on effect of the PtAu[2]-1 cluster in response to NO. (E)
   NO-responsive emission spectra change of the PtAu[2]-1 cluster at a
   concentration of 1.0 × 10^-5 M in DMSO-PBS (v/v = 1:1) solution,
   showing a gradual increase in emission intensity with the addition of 1
   to 11 equivalent Na[2][Fe(CN)[5]NO]. (F) Dependence of phosphorescence
   intensity on the molar ratio of Na[2][Fe(CN)[5]NO] vs PtAu[2]-1
   cluster. (G) Emission spectral change of the PtAu[2]-1 cluster (1.0 ×
   10^-5 M) in DMSO-PBS (v/v = 1:1) solution in response to an aqueous
   solution of L-cysteine, showing a gradual decrease in the peak at 542
   nm. (H) Dependence of emission intensity at 542 nm on the molar ratio
   of L-cysteine vs PtAu[2]-1 cluster.

   Upon excitation at λ[ex] = 398 nm, the BOC-protected PtAu[2]-1a cluster
   displayed intense green luminescence, peaking at 534 nm with an
   emissive lifetime of 2.0 μs and 26% quantum yield in DMSO-PBS (v/v =
   1:1) at room temperature (Figure [91]2C). The long-lived emission with
   a micro-second range lifetime and a large Stokes shift suggested that
   phosphorescent characteristics may arise from the mixed triplet excited
   states of ^3[acetylide → dTolmp] ^3LLCT (ligand-to-ligand charge
   transfer) and ^3[acetylide → PtAu[2]] ^3LMCT (ligand-to-metal charge
   transfer) transitions [92]^39^, [93]^40. In contrast, the PtAu[2]-1
   cluster exhibited relatively weak luminescence centered at 542 nm with
   an emissive lifetime of 1.7 μs and a quantum yield of 1.9%. The much
   weaker phosphorescence of the PtAu[2]-1 cluster compared to the
   BOC-protected PtAu[2]-1a cluster is likely due to the strong
   electron-donating character of the diamino groups, which facilitate
   intramolecular electron transfer that deactivates triplet excited
   states through non-radiative decay.

   The long-term stability of the PtAu[2]-1 cluster was further
   characterized using UV-Vis absorption and phosphorescence spectra. The
   inappreciable changes in the bands centered at 395 and 495 nm due to
   [acetylide → dTolmp] LLCT (ligand-to-ligand charge transfer) and
   [acetylide → PtAu[2]] ligand-to-metal charge transfer (LMCT)
   transitions, imply that the PtAu[2]-1 cluster maintains sufficient
   stability in the DMSO-PBS (v/v = 1:1) solution after it has been put
   aside for three days ([94]Figure S8). Likewise, the inappreciable
   changes in the phosphorescent bands centered at 534 nm, ascribed to
   ^3[acetylide → dTolmp] LLCT (ligand-to-ligand charge transfer) and
   ^3[acetylide → PtAu[2]] LMCT triplet excited states, suggest that the
   PtAu[2]-1 cluster is sufficiently stable in DMSO-PBS (v/v = 1:1)
   solution after it has been put aside for three days ([95]Figure S9).

   The binding of weakly phosphorescent PtAu[2]-1 cluster to molecules,
   such as NO, through diamino groups to produce a triazole ring
   effectively inhibited intramolecular electron transfer [96]^41,
   resulting in dramatically enhanced phosphorescence (Figure [97]2D).
   Therefore, we performed phosphorescent sensing experiments by titrating
   the PtAu[2]-1 cluster in DMSO-PBS (v/v = 1:1) with an aqueous solution
   of sodium nitroprusside (Na[2][Fe(CN)[5]NO]), a widely used exogenous
   NO donor [98]^42^, [99]^43. As depicted in Figures [100]2E and 2F, the
   addition of sodium nitroprusside progressively intensified the
   phosphorescent emission peak at 542 nm, resulting in a four-fold
   phosphorescent enhancement with 10 equivalents of sodium nitroprusside.
   In addition, gradual titration with sodium nitroprusside caused a
   progressive blue-shift of the emission peak for the PtAu[2]-1 cluster
   to 533 nm, due to the formation of PtAu[2]-1b cluster (Figure [101]2E)
   with the benzotriazole-acetylide ligand. Thus, the PtAu[2]-1 cluster
   displays excellent phosphorescent turn-on effects in response to NO and
   is a promising biosensor for detecting NO in inflammatory environments.

   As an effective binding agent for cysteine residues, Au^+ has been
   reported to react with cysteine thiol groups in specific peptides and
   proteins, thereby exerting biological effects [102]^44^, [103]^45. For
   instance, Au^+ can exert anti-inflammatory effects by inhibiting
   inflammatory ligand-receptor recognition and increasing the release of
   protective proteins. To further investigate the possible binding
   interactions between the heterometallic PtAu[2] clusters and cysteine,
   titration experiments were performed by adding an aqueous L-cysteine
   solution to the PtAu[2]-1 cluster in DMSO-PBS (v/v = 1:1). As shown in
   [104]Figures S10 and S11, the 390 nm absorbance peak of the PtAu[2]-1
   cluster was dose-dependently reduced by the addition of cysteine, which
   also progressively reduced its emission intensity at 542 nm (Figures
   [105]2G and 2H). Taken together, these results suggest a potential
   binding interaction between cysteine and the PtAu[2]-1 cluster, which
   may provide this heterometallic cluster with anti-inflammatory
   activity.

Biocompatibility and cellular uptake of PtAu[2] clusters

   Next, we investigated the biocompatibility and cellular uptake behavior
   of PtAu[2] clusters. Live/dead staining was performed to evaluate the
   cytotoxicity of PtAu[2] clusters at various concentrations. As shown in
   Figures [106]3A and 3B, PtAu[2] clusters exhibited neglectable
   cytotoxicity to RAW 264.7 macrophages at a dose of 20 μg/mL. However,
   an increased number of PI-positive cells and a reduced number of
   Calcein-AM-positive cells were observed when the concentration reached
   40 μg/mL. To further evaluate the effect of PtAu[2] clusters on cell
   viability, CCK-8 assays were performed. As shown in Figure [107]3C,
   PtAu[2] clusters hardly affected the viability of RAW 264.7
   macrophages, even at a dose of 40 μg/mL for 24 h. However, clear
   cytotoxicity was observed at a concentration of 40 μg/mL after
   prolonged treatment for 48 h. Similarly, PtAu[2] clusters displayed
   negligible cytotoxicity in BMMs at a dose of 40 μg/mL after 48 h
   (Figure [108]3D), but dose-dependently reduced cell viability was
   observed at concentrations of 20 μg/mL or higher when treated for 96 h.
   Consistently, flow cytometry revealed that PtAu[2] clusters only
   marginally affected cell cycle phases at doses of up to 20 μg/mL
   ([109]Figure S12). To further explore whether PtAu[2] clusters
   adversely affected blood cell homeostasis, hemolysis tests were
   performed by incubating red blood cells (RBCs) with various
   concentrations of PtAu[2] clusters. As shown in Figure [110]3E, PtAu[2]
   clusters slightly reduced RBC viability at doses up to 20 μg/mL, with a
   hemolysis rate lower than 3%. Collectively, these results demonstrate
   that PtAu[2] clusters exhibit satisfactory biocompatibility at
   concentrations up to 20 μg/mL in RAW 264.7 macrophages and could be
   used at concentrations of 10 μg/mL or below in BMMs without
   compromising cell viability.

Figure 3.

   [111]Figure 3
   [112]Open in a new tab

   Biocompatibility and cellular uptake of PtAu[2] clusters. (A) Live/dead
   staining of RAW 264.7 macrophages treated with various concentrations
   of PtAu[2] clusters for 24 h. (B) Quantitative analysis of Calcein-AM
   positive cells and PI positive cells. (C) Viability of RAW 264.7
   macrophages treated with various concentrations of PtAu[2] clusters for
   24 and 48 h. (D) Viability of murine BMMs treated with various
   concentrations of PtAu[2] clusters for 48 and 96 h. (E) Hemolysis assay
   of PtAu[2] clusters. (F) Representative images of LPS-activated RAW
   264.7 macrophages stained with DAF-FM DA and detected by CLSM. (G)
   Representative images of LPS-activated RAW 264.7 macrophages incubated
   with PtAu[2] clusters detected by CLSM. Data represent the mean ± SD of
   three independent experiments (two-tailed unpaired Student's t-test).
   ** and ^## p < 0.01 compared to the 0 μg/mL group.

   Having evaluated their biocompatibility, we examined the cellular
   uptake behavior of PtAu[2] clusters and the feasibility of
   NO-responsive phosphorescence in inflammatory environments. Since NO is
   an inflammatory molecule produced by inflammatory macrophages, we
   detected NO-responsive phosphorescence in RAW 264.7 macrophages
   stimulated by LPS for 12 h and treated with PtAu[2] clusters at a
   concentration of 20 μg/mL, and used a commercial NO detection (DAF-FM
   DA) kit for comparison. As shown in Figure [113]3F, the fluorescence
   intensity of DAF-FM DA was markedly enhanced after LPS stimulation,
   indicating an increase in NO production. Similarly, the phosphorescence
   intensity of PtAu[2] clusters was significantly enhanced in LPS
   activated-macrophages but not in unstimulated macrophages (Figures
   [114]3G and S13). Weak phosphorescence was observed in LPS-activated
   macrophages following the addition of SMT, an iNOS inhibitor,
   suggesting that phosphorescence enhancement was mainly triggered by NO
   generation ([115]Figure S14). Furthermore, we observed that most
   PtAu[2]clusters escaped lysosomes after 12 h, with little
   colocalization observed due to the cationic property of the PtAu[2]
   clusters ([116]Figure S15). Taken together, these results demonstrate
   that PtAu[2] clusters exhibit satisfactory cellular uptake and
   NO-responsive phosphorescence enhancement in inflammatory macrophages.

Anti-inflammatory effects of PtAu[2] clusters in vitro

   Encouraged by the satisfactory biocompatibility and cellular uptake
   behavior of PtAu[2] clusters, we next explored whether these
   nanoclusters exhibited anti-inflammatory effects in inflammatory
   macrophages. RAW 264.7 macrophages were challenged with LPS in the
   presence of various concentrations of PtAu[2] clusters (5, 10, and 20
   μg/mL) for 24 h. As shown in Figure [117]4A, RT-qPCR assays revealed
   that Tnfα, Il1β, Il6, and Nos2 mRNA expression was upregulated
   following LPS stimulation but was dose-dependently inhibited after
   treatment with PtAu[2] clusters. Next, we investigated the effect of
   PtAu[2] clusters precursors on the inflammatory reaction by RT-qPCR.
   The results demonstrated that, unlike the complete PtAu[2] clusters,
   the precursors of PtAu[2] clusters (dTolmp, Pt(PPh[3])[2] and
   Au(tht)Cl) exhibited a marginal inhibitory effect on pro-inflammatory
   gene expression ([118]Figure S16), indicating that the structural
   integrity is important for the anti-inflammatory effect of PtAu[2]
   clusters. Consistently, immunofluorescence staining revealed that iNOS
   expression was significantly elevated following LPS stimulation, while
   it was suppressed by various concentrations of PtAu[2] clusters in
   LPS-activated RAW 264.7 macrophages (Figure [119]4B). LPS stimulation
   also markedly upregulated the phosphorylation and nuclear translocation
   of NF-κB p65, which were dose-dependently inhibited by PtAu[2] clusters
   (Figure [120]4C). Furthermore, we investigated the expression of iNOS,
   p-p65, and p65 using western blotting. As shown in Figure [121]4D, the
   expression of iNOS and the phosphorylation of p65 were markedly
   upgraded following LPS stimulation, whereas they significantly
   decreased by treatment with PtAu[2] clusters in a dose-dependent
   manner. In addition, to determine the effect of PtAu[2] clusters on
   inflammatory cytokines at the protein level, we performed ELISA to
   measure the production of inflammatory cytokines. PtAu[2] clusters
   dose-dependently reduced the production of the pro-inflammatory
   cytokines TNF-α, IL-1β, and IL-6 but hardly affected the production of
   the anti-inflammatory cytokine IL-10 (Figure [122]4E). Taken together,
   these results suggest that PtAu[2] clusters suppress inflammatory
   reactions in LPS-stimulated RAW 264.7 macrophages.

Figure 4.

   [123]Figure 4
   [124]Open in a new tab

   PtAu[2] clusters suppress inflammatory reactions in LPS-stimulated RAW
   264.7 macrophages. (A) RT-qPCR analysis of pro-inflammatory gene
   expression in LPS-activated RAW 264.7 macrophages treated with various
   concentrations of PtAu[2] clusters. (B) iNOS immunofluorescence in
   LPS-activated RAW 264.7 macrophages treated with various concentrations
   of PtAu[2] clusters. (C) p-p65 immunofluorescence in LPS-activated RAW
   264.7 macrophages treated with various concentrations of PtAu[2]
   clusters. (D) Expression of iNOS, p-p65, and p65 in total cellular
   lysates after treatment with various concentrations of PtAu2 clusters.
   (E) ELISA results for inflammatory cytokines production by
   LPS-activated RAW 264.7 macrophages treated with various concentrations
   of PtAu[2] clusters. (F) Confocal images of LPS-activated RAW 264.7
   macrophages treated with various concentrations of PtAu[2] clusters and
   stained with DHE. Data represent the mean ± SD of three independent
   experiments (one-way ANOVA with Tukey's post hoc test). *p < 0.05; **p
   < 0.01.

   When activated by various inflammatory stimuli, macrophages exhibit
   respiratory bursts and produce excessive ROS, which leads to oxidative
   stress. Oxidative stress may further aggravate inflammatory reactions,
   triggering a vicious cycle [125]^46^, [126]^47. Therefore, we
   investigated the effect of PtAu[2] clusters on ROS overproduction in
   inflammatory macrophages. As shown in Figure [127]4F, PtAu[2] clusters
   effectively reduced intracellular ROS levels, with weaker DHE
   fluorescence than that of the LPS-activated group. Moreover, the level
   of MDA, a well-established marker of oxidative stress, was efficiently
   reduced by PtAu[2] clusters treatment ([128]Figure S17). Furthermore,
   both the intracellular GSH level and SOD activity were restored by
   treatment with PtAu[2] clusters in a dose-dependent manner ([129]Figure
   S18 and S19), thereby increasing the cellular resistance to oxidative
   stress. Collectively, these data suggest that PtAu[2] clusters suppress
   oxidative stress and increase the anti-oxidative capacity of
   LPS-stimulated RAW 264.7 macrophages.

Effect of PtAu[2] clusters on OC formation, fusion, and function

   Having demonstrated the potent anti-inflammatory effects of PtAu[2]
   clusters, we next investigated whether PtAu[2] clusters exert
   inhibitory effects against OCs, the principal effector cells of
   inflammatory osteolysis. A maximum concentration of 10 μg/mL of PtAu[2]
   clusters was used based on their cytotoxicity in BMMs. As shown in
   Figure [130]5A, BMMs differentiated into mature OCs with multiple
   nuclei in the presence of RANKL for five days. However, the formation
   of TRAP-positive OCs was reduced in a dose-dependent manner by various
   concentrations of PtAu[2] clusters (2.5, 5, and 10 μg/mL), which
   reduced the number and area of the OCs (Figure [131]5D and 5E).
   Notably, the nuclear number of mature OCs also significantly decreased
   in a dose-dependent manner following treatment with PtAu[2] clusters,
   indicating the inhibitory effect of PtAu[2] clusters on OC fusion
   ([132]Figure S20). To further evaluate the effect of PtAu[2] clusters
   on OC fusion, cytoskeletal podosome actin belts, which indicate OC
   precursor cell fusion ability, were visualized by immunofluorescence.
   As shown in Figures [133]5B and 5F, PtAu[2] clusters dose-dependently
   suppressed the formation of podosome actin belts, further indicating
   that PtAu[2] clusters reduced OC fusion. In addition, we used Osteo
   Assay Surface plates with hydroxyapatite coating to evaluate the effect
   of PtAu[2] clusters on bone resorption, an important function of mature
   OCs. Extensive bone resorption was observed when BMMs were stimulated
   with RANKL for 6 days; however, the bone resorption area was reduced in
   a dose-dependent manner by various concentrations of PtAu[2] clusters
   (Figures [134]5C and 5G). To further investigate the effect of PtAu[2]
   clusters on transcript levels in OCs, RT-qPCR was performed to analyze
   the expression of OC-related genes, including Trap (osteoclast
   differentiation), Ctr (osteoclast differentiation), Dcstamp (osteoclast
   fusion), and Ctsk (bone resorption). As expected, PtAu[2] clusters
   markedly downregulated the expression of OC-related genes in a
   dose-dependent manner (Figure [135]5H). Moreover, western blotting
   results demonstrated that the expression of NFATc1 (a key
   transcriptional factor in osteoclastogenesis) and CTSK (a key protein
   in OCs-mediated bone resorption) gradually increased over time in the
   control group but was reduced after treatment with 10 μg/mL PtAu[2]
   clusters ([136]Figure S21). Taken together, these results indicate that
   PtAu[2] clusters effectively reduce RANKL-induced OC formation, fusion,
   and function in BMMs.

Figure 5.

   [137]Figure 5
   [138]Open in a new tab

   PtAu[2] clusters reduce RANKL-induced osteoclastogenesis, podosome
   actin belt formation, and OC-mediated bone resorption. (A)
   Representative tartrate-resistant acid phosphatase (TRAP) staining
   images of BMMs stimulated with RANKL for 5 days and treated with
   various concentrations of PtAu[2] clusters. (B) Representative
   immunofluorescence images of podosome actin belts in BMMs stimulated
   with RANKL for 5 days and treated with various concentrations of
   PtAu[2]clusters. (C) Representative images of bone resorption pits on
   Osteo Assay Surface plates coated with hydroxyapatite produced by OCs
   treated with various concentrations of PtAu[2] clusters. (D) Number of
   OCs per well. (E) Area of OCs per well. (F) The number of podosome
   actin belts per well. (G) Bone resorption area per well. (H) RT-qPCR
   results of OCs-related gene expression in BMMs cultured with RANKL for
   4 days in presence of various concentrations of PtAu[2] clusters. (D,
   E, and G) Data represent the mean ± SD of four independent experiments
   (one-way ANOVA with Tukey's post hoc test). (F and H) Data represent
   the mean ± SD of three independent experiments (one-way ANOVA with
   Tukey's post hoc test). *p < 0.05; **p < 0.01.

Anti-inflammatory mechanisms of PtAu[2] clusters

   To further elucidate the potential mechanisms underlying the
   anti-inflammatory and anti-OC properties of PtAu[2] clusters, we
   performed RNA sequencing. Principal components analysis (PCA) revealed
   that the Control, LPS, and LPS + PtAu[2] groups had different
   transcriptomic profiles (Figure [139]6A). The Venn diagram illustrates
   that the three groups shared 9341 genes, with 495, 56, and 85 genes
   exclusively expressed in the Control, LPS, and LPS + PtAu[2] groups,
   respectively (Figure [140]6B). The Volcano plots displayed that 2672
   genes were significantly upregulated with 2551 genes downregulated
   after LPS stimulation, while 128 genes were significantly upregulated
   and 109 genes were downregulated after treatment with PtAu[2] clusters
   (fold change ≥ 2.0, p < 0.05) ([141]Figure S22). As shown in Figure
   [142]6C, the clustered heat map shows that the expression of various
   pro-inflammatory genes, such as Tnf, Nlrp3, Ptgs2, Mmp13, Il1b, Il12b,
   Il6, and Il23a, was markedly elevated following LPS stimulation,
   whereas it was significantly reduced after treatment with PtAu[2]
   clusters. Meanwhile, it was noted that the expression of a series of
   anti-inflammatory and anti-oxidative genes, including Gstp1, Cat,
   Hmox1, Mgst2, Gclm, and Prdx1, was upregulated after PtAu[2] clusters
   treatment compared to the LPS group. GO enrichment analysis of the
   differentially expressed genes between the LPS and LPS + PtAu[2] groups
   demonstrated significant changes in immune response, positive
   regulation of NF-κB transcription factor activity, and positive
   regulation of OC differentiation (Figure [143]6D). The GO analysis
   chord plot also revealed that treatment with PtAu[2] clusters
   downregulated the expression of genes related to the positive
   regulation of OC differentiation (Tnfα, Il12b, Il23a), positive
   regulation of NO biosynthetic processes (Klf2, Tnfα, Il1β), and other
   inflammatory biological processes, but upregulated the expression of
   genes related to angiogenesis (Hmox1, Angptl2, Plxdc1), glutathione
   biosynthetic processes (Gclm and Mgst2), and other anti-inflammatory
   biological processes (Figure [144]6E). We also performed KEGG analysis
   of the differentially expressed genes between the LPS and LPS + PtAu[2]
   groups to identify the top 20 enriched pathways, which included
   cytokine-cytokine receptor interactions, the TNF, NF-κB, and JAK-STAT
   signaling pathways (Figure [145]6F). In addition, gene set enrichment
   analysis (GSEA) revealed that genes related to cytokine-cytokine
   receptor interactions and OC differentiation were significantly
   enriched and downregulated after treatment with PtAu[2] clusters,
   compared with the LPS group (Figure [146]6G). Taken together, these
   results suggest that PtAu[2] clusters may exert anti-inflammatory
   effects by disrupting cytokine-cytokine receptor interactions, thereby
   inhibiting downstream signaling pathways such as the NF-κB and JAK-STAT
   signaling pathways. These findings are consistent with previous reports
   stating that gold clusters can inhibit the activation of cytokine
   receptors by cysteine residue-containing cytokine peptides through
   direct interactions between Au^+ and thiol groups in the stimulating
   peptides [147]^20^, [148]^45.

Figure 6.

   [149]Figure 6
   [150]Open in a new tab

   Transcriptomic analysis of the anti-inflammatory mechanisms of PtAu[2]
   clusters. (A) PCA of the transcriptomic profiles in the Control, LPS,
   and LPS + PtAu[2] groups. (B) Venn diagram. (C) Clustered heat map of
   the representative inflammation-related genes (fold change ≥ 2.0 and p
   < 0.05). (D) Dot plot of biological processes related to the
   differentially expressed genes between the LPS and LPS + PtAu[2]
   groups. (E) Chord plot of GO analysis of the differentially expressed
   genes between the LPS and LPS + PtAu[2] groups. (F) KEGG pathway
   enrichment analysis of the differentially expressed genes between the
   LPS and LPS + PtAu[2] groups. (G) GSEA of the differentially expressed
   genes between the LPS and LPS + PtAu[2] groups.

PtAu[2] clusters promote Nrf2 release from the Keap1/Nrf2 complex

   Since RNA sequencing analysis suggested that PtAu[2] cluster treatment
   upregulated the expression of Nrf2 downstream genes with
   anti-inflammatory and anti-oxidative properties, including Hmox1, Gclm,
   Cat, Prdx1, Mgst2, and Gstp1 (Figure [151]6C), we performed RT-qPCR
   analysis to verify these findings. As shown in Figure [152]7A, PtAu[2]
   clusters markedly increased the mRNA expression of Hmox1 and Nqo1 in a
   dose-dependent manner without affecting that of Nrf2. Thus, PtAu[2]
   clusters may upregulate the transcriptional expression of antioxidants
   downstream of Nrf2, to increase endogenous antioxidant production and
   reduce intracellular ROS. Nrf2 has been reported to bind to Keap1,
   which is anchored to the actin cytoskeleton and promotes Nrf2
   degradation in unstimulated cells [153]^48. The conformational changes
   in Keap1 induced by oxidative stress or chemicals with electrophilic
   properties allow Nrf2 to dissociate from Keap1 and translocate into the
   nucleus, where it increases the production of downstream antioxidants
   [154]^49. To further explore the effect of PtAu[2] clusters on Nrf2
   activation, immunofluorescence was used to detect Nrf2 expression in
   LPS-challenged macrophages treated with PtAu[2] clusters. As shown in
   Figure [155]7B, increased Nrf2 expression and nuclear translocation
   were observed in LPS-challenged macrophages treated with PtAu[2]
   clusters compared to LPS-challenged macrophages. Considering the strong
   interaction reported between Au^+ and thiol groups, we speculated that
   PtAu[2] clusters might activate Nrf2 expression by modifying cysteine
   residues in Keap1, leading to a conformational change that promotes the
   release and nuclear translocation of Nrf2 and the upregulation of
   downstream anti-inflammatory and anti-oxidative genes (Figure [156]7C).

Figure 7.

   [157]Figure 7
   [158]Open in a new tab

   PtAu[2] clusters promote the release of Nrf2 from the Keap1/Nrf2
   complex. (A) RT-qPCR analysis of Nrf2 and downstream gene expression in
   LPS-activated RAW 264.7 macrophages treated with various concentrations
   of PtAu[2] clusters. (B) Immunofluorescence analysis of Nrf2 in
   LPS-activated RAW 264.7 macrophages treated with various concentrations
   of PtAu[2] clusters. (C) Schematic diagram of the modulation of Nrf2
   and Keap1 conformation by PtAu[2] clusters (created with
   BioRender.com). (D) Expression of Keap1/Nrf2 signaling members in total
   cellular lysates after treatment with various concentrations of PtAu[2]
   clusters. (E) Quantitative analysis of western blotting results. (F)
   Nrf2 expression in cytoplasmic lysates. (G) Nrf2 expression in nuclear
   lysates. (H) Co-immunoprecipitation assays of Keap1-Nrf2. Data
   represent the mean ± SD of three independent experiments (one-way ANOVA
   with Tukey's post hoc test). *p < 0.05; **p < 0.01.

   To verify this hypothesis, we evaluated the protein expression of
   Keap1, Nrf2, and its downstream antioxidants by western blotting of
   total cellular lysates of LPS-challenged macrophages treated with
   PtAu[2] clusters. As expected, PtAu[2] clusters downregulated Keap1
   expression but upregulated the expression of Nrf2 and its downstream
   antioxidants, including HO-1 and NQO1, in a dose-dependent manner
   (Figures [159]7D and 7E). Similar results of increased expression of
   Nrf2 and its downstream HO-1 and NQO1 could also be observed in
   RANKL-stimulated BMMs with PtAu[2] clusters treatment ([160]Figure
   S23). Furthermore, cytoplasmic Nrf2 expression was upregulated by
   PtAu[2] clusters, indicating increased Nrf2 protein stabilization and
   accumulation (Figure [161]7F). Consistent with the immunofluorescence
   results, PtAu[2] clusters increased nuclear translocation of Nrf2,
   suggesting increased Nrf2 activation (Figures [162]7G and S24). To
   elucidate the underlying mechanism, we examined whether PtAu[2]
   clusters affect the association between Keap1 and Nrf2 in LPS-activated
   macrophages. Co-immunoprecipitation assays confirmed the low levels of
   immunoprecipitation between Keap1 and Nrf2 following treatment with
   PtAu[2] clusters (Figure [163]7H). In addition, we also explored
   whether PtAu[2] clusters affect other endogenous anti-inflammatory and
   anti-oxidative pathways by RT-qPCR ([164]Figure S25). The results
   demonstrated that PtAu[2] clusters hardly affected gene expression of
   Foxo1, Sirt1, and Ppargc1a in LPS-stimulated macrophages, indicating
   that these signaling molecules may not be involved in the
   anti-inflammatory and anti-oxidative efficacy of PtAu[2] clusters.
   Taken together, these findings suggest that PtAu[2] clusters disrupt
   the association between Keap1 and Nrf2, thereby promoting the release
   and nuclear translocation of Nrf2 as well as the activation of
   endogenous anti-inflammatory and anti-oxidative products.

Inhibitory effect of PtAu[2] clusters on LPS-induced calvarial osteolysis in
vivo

   Having determined the biocompatibility, cellular uptake,
   anti-inflammatory phenotypes, and anti-inflammatory mechanisms of
   PtAu[2] clusters in vitro, we investigated their therapeutic efficiency
   in vivo by establishing a murine model of LPS-induced calvarial
   osteolysis. As shown in Figure [165]8A, LPS-soaked collagen sponge
   blocks were implanted into mouse calvaria to cause inflammation-induced
   bone loss, and local PBS or PtAu[2] injections were administered every
   other day. After 14 days, the mouse calvaria were harvested for
   radiographic and histological assessment. Micro-CT revealed extensive
   bone erosion on the calvarial bone surface in the LPS group, with
   numerous large and deep resorption pits (Figure [166]8B). In contrast,
   PtAu[2] clusters significantly reduced the scope and extent of bone
   destruction, as evidenced by a reduction in the number of resorption
   pits. Quantitative analysis also illustrated that key morphometric
   parameters, including bone volume/tissue volume (BV/TV), trabecular
   bone number (Tb.N), and trabecular bone separation (Tb.Sp), were
   markedly improved in the PtAu[2] group compared to the LPS group
   (Figure [167]8C).

Figure 8.

   [168]Figure 8
   [169]Open in a new tab

   Inhibitory effect of PtAu[2] clusters on LPS-induced calvarial
   osteolysis in vivo. (A) Schematic diagram of in vivo experiments. (B)
   Representative micro-CT scanning images of mice calvaria. (C)
   Quantitative analysis of micro-CT scanning images. (D) Representative
   H&E, TRAP, and immunostaining images of TNF-α, Nrf2, and HO-1. (E)
   Quantification of the number of TRAP-positive OCs. (F) Quantification
   of OcS/BS%. (G) Quantitative analysis of relative TNF-α fluorescence.
   (H) Quantitative analysis of relative Nrf2 fluorescence. (I)
   Quantitative analysis of relative HO-1 fluorescence. (C) Data represent
   the mean ± SD of five independent animals (one-way ANOVA with Tukey's
   post hoc test). (E, F, G, H, and I) Data represent the mean ± SD of
   four independent animals (one-way ANOVA with Tukey's post hoc test). *p
   < 0.05; **p < 0.01.

   Histological evaluation confirmed the protective effect of PtAu[2]
   clusters on bone resorption and inflammatory reactions in vivo.
   Consistent with the micro-CT findings, H&E staining revealed that the
   local LPS injection induced extensive osteolysis, whereas PtAu[2]
   clusters alleviated this effect (Figure [170]8D). In addition, TRAP
   staining demonstrated that the number of TRAP-positive OCs was
   increased by LPS injection but reduced by PtAu[2] clusters (Figures
   [171]8E and 8F). Immunofluorescence staining further suggested that LPS
   significantly increased TNF-α expression, whereas PtAu[2] clusters
   markedly reduced TNF-α expression (Figure [172]8G). Consistent with the
   in vitro results, both in vivo western blotting and immunofluorescence
   staining illustrated that PtAu[2] clusters effectively activated Nrf2
   expression ([173]Figure S26 and 8H). Consistent with these results,
   immunofluorescence staining revealed that PtAu[2] clusters elevated
   HO-1 expression, an important downstream anti-oxidative product of Nrf2
   (Figure [174]8I). Collectively, these results indicate that PtAu[2]
   clusters can attenuate LPS-induced osteolysis by inhibiting bone
   resorption and inflammatory activity, and activating endogenous
   anti-inflammatory systems.

   Moreover, we examined the biosafety of PtAu[2] clusters. H&E staining
   revealed no obvious unfavorable effects on the histological features of
   the major organs, including the heart, liver, spleen, lungs, and
   kidneys ([175]Figure S27). Meanwhile, complete blood panel analysis and
   serum biochemistry analysis also demonstrated that PtAu[2] clusters did
   not exhibit toxicity 11 days after PtAu[2] clusters treatment
   ([176]Figure S28). Furthermore, ICP-MS was performed to evaluate the
   biodistribution and metabolism of PtAu[2] clusters 48 h after
   administration. The results showed that the liver, spleen, and
   intestine exhibited the highest dosage distribution ([177]Figure S29),
   which is consistent with a previous report on the biodistribution of
   bimetallic clusters [178]^30, indicating possible hepatobiliary or
   fecal elimination of PtAu[2] clusters. Together, these results suggest
   a satisfactory level of biosafety as well as an excretion pathway of
   PtAu[2] clusters.

Conclusions

   In this study, we demonstrated that PtAu[2] nanoclusters with
   electron-rich 4-ethynyl benzene-1,2-diamine show specifical
   NO-responsive phosphorescent properties and exhibit strong binding
   interactions with cysteine, thereby qualifying them as sensitive
   inflammation detectors, as well as an ideal anti-inflammatory agent.
   The potent anti-inflammatory effects of PtAu[2] clusters can be
   attributed to the following mechanisms: i) interrupting
   cytokine-cytokine receptor interactions to inhibit downstream
   inflammatory signaling pathways; and ii) activating Nrf2 expression by
   disrupting the Keap1-Nrf2 association, causing increased Nrf2 release
   and nuclear translocation to upregulate the expression of endogenous
   anti-inflammatory and anti-oxidative products. Furthermore, in vivo
   experiments confirmed that PtAu[2] clusters alleviated inflammatory
   osteolysis by inhibiting osteoclastogenesis and increasing Nrf2
   expression, despite the molecular structural basis of PtAu[2] clusters
   binding to their targeted proteins remains to be investigated by
   experimental crystal structure studies. From the perspective of
   clinical translation, PtAu[2] clusters could be directly used as
   therapeutic agents or coatings for implants, or in combination
   therapies with other drugs to reduce systemic toxicity, while these
   nanoclusters may possess certain limitations, including inadequate
   ROS-scavenging properties and a lack of responsiveness to various
   stimuli. Nevertheless, by developing novel heterometallic nanoclusters
   as activators of endogenous anti-inflammatory systems, this study
   provides new insights into the use of multifunctional molecular
   therapeutic agents for the treatment of inflammatory osteolysis and
   other inflammatory diseases.

Supplementary Material

   Supplementary materials and methods, figures and tables.
   [179]Click here for additional data file.^ (9.2MB, pdf)

Acknowledgments