Microenvironment-responsive NIR-IIb multifunctional nanozyme platform for bacterial imaging and specialized anti-anaerobic bacteria periodontal therapy

Abstract

Periodontitis is a chronic inflammatory disease caused by plaque. In order to remove pathogens and promote tissue repair, the following steps need to be taken simultaneously: localizing the diseased area, improving the anaerobic microenvironment, as well as addressing the anti-inflammatory and osteogenic needs. This study aims to address these issues by developing a responsive near-infrared-IIb nanozyme system (DMUP), assembled from lanthanide-doped down-converted nanoparticles and multi-enzymatically active nanozyme. DMUP binds to bacterial membranes via the bacterial targeting peptide ubiquicidin_29-41 (UBI_29-41). Upon responding to the inflammatory microenvironment, it releases manganese (Mn) nanozyme and paeonol (Pae), and localized infected areas by fluorescent bacterial imaging in the near-infrared IIb (NIR-IIb) region. In particular, the released Mn nanozyme reacts with hydrogen peroxide in the inflammatory microenvironment to generate oxygen (O₂) in situ, thereby improving the anoxic environment to inhibit anaerobic bacteria. On the other hand, as a metal oxide nanozyme, Mn nanozyme scavenges reactive oxygen species (ROS) by mimicking the cascade process of superoxide dismutase and catalase. The phenolic antioxidant Pae shifts macrophages from pro-inflammatory (M1-type) to anti-inflammatory (M2-type) through the Akt/mTOR pathway. It can synergize with Mn nanozyme to regulate the inflammatory microenvironment, thereby reducing inflammation, promoting osteogenic genes expression, and accelerating periodontal tissues regeneration.

Graphical abstract

Introduction

Periodontitis, a chronic inflammatory disease caused by plaque [1], has become the sixth most common chronic disease, affecting 734 million people in the world [2]. Typically, periodontitis treatment involves subgingival scaling and root planning (SPR) and antibiotics [3]. However, the periodontal pockets have a complex structure and are covered with soft tissue, which makes it challenging to accurately define the infected area during curettage. As a result, the removal effect of SPR is seriously affected. Porphyromonas gingivalis (P. gingivalis) is an anaerobic bacterium that causes periodontitis [4]. Moreover, it is also difficult to inhibit such strains in the hypoxic periodontal pocket area by broad-spectrum antibiotics (e.g., minocycline, MC, clinical drug commonly used in periodontitis). Periodontitis progression also relates to the host immune response, which is overactivated with the increase of pathogenic bacteria and their metabolites [5] This immune stress triggers a series of antibody responses that modify the periodontal microenvironment by exacerbating hypoxia [6,7,8], as well as increasing reactive oxygen species (ROS) and inflammatory factors [9,10,11]. The imbalanced periodontitis microenvironment further leads to tissue acidification, which in turn affects immune cell function and accelerates tissue damage [12]. These complex interactions further reduce the effectiveness of conventional treatments. Therefore, it is crucial to develop a treatment strategy for periodontitis that includes bacterial visualization (localization of the diseased area), inhibition of anaerobic bacteria, and improvement of the inflammatory microenvironment.

With the advantages of safety, real-time monitoring, non-invasiveness, and deep tissue penetration, the near-infrared-II (NIR-II, wavelength 1000–1700 nm) region fluorescence imaging is an emerging biomedical imaging method [13]. In particular, due to their high chemical stability, excellent photostability, and narrow emission band performance, lanthanum-doped NIR-IIb (wavelength 1500–1700 nm) down-conversion nanoparticles (DCNPs) have attracted significant attention in tumor imaging [14,15,16]. Thus, when combined with suitable bacterial-targeting ligands, this material could be potentially utilized for the visual localization of infected regions [17,18,19]. For example, by using the bacteria-targeting peptide ubiquicidin (UBI_{29 − 41}), the electrostatic adsorption with phosphatidylglycerol (PG) on the bacterial cytosol membrane can effectively increase the recognition of bacteria by the imaging group [20, 21]. The oral cavity is an open environment with diverse dynamics. Therefore, the microenvironmental characteristics of bacteria or diseases can also be used to construct an auto-responsive fluorescence system. This strategy can improve the imaging specificity of pathogenic bacteria in the lesion area and localize the bacterial infection area [22, 23]. Notably, few studies have used bacterial fluorescent probes to image bacteria in periodontal pockets. The high hydrogen peroxide (H₂O₂) content, as well as the acidic microenvironment in the inflammatory tissues, could be utilized to develop responsive fluorescent probes. Thus, the efficiency of bacterial removal during scaling procedures can be potentially improved by constructing a responsive fluorescence imaging platform based on DCNPs to visualize bacteria in periodontal pockets.

After achieving precise imaging, another focus of the therapeutic system is to improve the microenvironment (hypoxia and inflammation) in the infected area. With catalase (CAT)-like activity [24], manganese (Mn) nanozyme catalyzes the disproportionation of H₂O₂ to produce oxygen (O₂). This reaction leads to a significant improvement in the hypoxic microenvironment, which restricts the anaerobic bacterial electron transport chain and inhibits the anaerobic bacteria [25, 26]. Mn nanozyme also possesses superoxide dismutase (SOD)-like activity, which scavenges another ROS (superoxide anion, ·O₂⁻). Thus, it is able to act as an exogenous ROS scavenger, thereby modulating immune-inflammatory products, protecting the endogenous antioxidant system to alleviate the inflammatory response of tissues [27, 28]. By forming an antioxidant system (Mn nanozyme-Pae) with natural antioxidants such as paeonol (Pae) [29], this approach increases cellular autophagic flux and alters the functional morphology of immune cells by targeting intracellular lysosomes. This ameliorates the immune microenvironment to promote an inflammatory environment conducive to osteogenesis, thus achieving the desired therapeutic effect of anti-inflammation and osteogenesis.

This study developed a responsive NIR-IIb multifunctional nanozyme system (β-NaErF₄:Yb³⁺@NaLuF₄@mSiO₂-Mn-UBI_{29 − 41}-Pae, DMUP), which is capable of responding to multiple stimuli to realize bacterial imaging and inflammatory microenvironment modulation in the periodontitis microenvironment. With Er³⁺ and Yb³⁺ doped NaF nanocrystals as the core, the system was coated with mesoporous silica (mSiO₂) to enhance hydrophilicity. Furthermore, the zeolite imidazole skeleton (ZIF-8) metal frame was utilized to precipitate Mn nanozyme in the pores of mSiO₂. Subsequently, the system was grafted with bacterial-targeting ligand UBI_{29 − 41} on its surface and loaded with Pae (Fig. 1). The resultant system demonstrated the following advantages: (1) Bacterial imaging. Aggregation of DCNPs and Mn nanozyme produced a fluorescence resonance energy transfer (FRET) effect leading to NIR-IIb fluorescence quenching [30]. Thus, under normal conditions, the DMUP nanoparticles exhibited a fluorescence-quenching state. However, under acidic inflammatory conditions, the hydrogen ions (H⁺) protonated the ligands in ZIF-8, thereby disrupting the metal-organic framework and releasing Mn nanozyme and Pae from the pores [31, 32]. The released Mn nanozyme and Pae rapidly restored and turned on the NIR-IIb fluorescence of DMUP. Furthermore, by utilizing the targeting property of UBI_{29 − 41}, DMUP bound to the bacterial surface with specificity to achieve fluorescence localization in the infected area (Fig. 1 I). (2) Anti-anaerobic bacteria. The released Mn nanozyme reacted with the abundant H₂O₂ and H⁺ in the acidic periodontitis environment to generate O₂ [33, 34], which effectively inhibited the growth of pathogenic anaerobic bacteria (Fig. 1 II). (3) ROS scavenging and anti-inflammation. Mn nanozyme is known to provide CAT-like and SOD-like enzyme activities [27, 28]. It exhibited anti-inflammatory effects by scavenging ROS and reducing tissue inflammation through inhibiting the expression of inflammatory factors (Fig. 1 III). (4) Promoting bone regeneration. By using Pae to target lysosomes, the system increased the cellular autophagic flux [29, 35], thereby promoting the transition of macrophages from M1-type to M2-type. This adjustment of the immune microenvironment reduced inflammation and created an environment favorable for tissue repair and regeneration. Consequently, bone matrix proteins were protected, thereby maintaining osteoblast function and activity, which promoted bone regeneration (Fig. 1 IV) [36, 37]. In summary, the multifunctional nano-system DMUP integrated bacterial imaging with anti-anaerobic, anti-inflammatory, and bone regeneration-promoting properties, demonstrating its potential applications in periodontitis treatment.

Fig. 1

Schematic structure and working principle of DMUP. (a) Down-conversion nanoparticles containing rare earth Er were coated with mSiO₂ and surface-grafted with the bacterial targeting ligand UBI_{29 − 41} to form DCMS-UBI. Through the scaffolding of a metal-organic framework, there was intra-pore precipitation of Mn nanozyme (DMU). The remaining pores were loaded with Pae, a traditional Chinese medicine monomer with anti-inflammatory and immune-modulatory properties to form DMUP. (b) DMUP targeted bacterial membranes while the metal-organic framework disintegrated in a low-pH inflammatory microenvironment, thereby releasing Mn nanozyme and restoring NIR-IIb fluorescence properties. This was utilized for bacterial imaging. The released Mn nanozyme catalyzed endogenous H₂O₂ to produce O₂, which inhibited anaerobic bacteria. (c) The released Mn nanozyme and Pae acted as an antioxidant system to inhibit the macrophage Akt/mTOR pathway, increased autophagic activity, and promoted macrophage transformation to the M2 anti-inflammatory type. Furthermore, Mn nanozyme scavenged ROS and produced O₂, synergistically improving the periodontitis microenvironment and promoting the regeneration of periodontal bone tissue. DCNPs: NaErF₄:Yb@NaLuF₄; DCMS: DCNPs@mSiO₂; DCMS-COSR: DCNPs@mSiO₂-COSR; DM-COSR: DCMS-MnO₂-COSR; DMU: DCMS-MnO₂-UBI_{29 − 41}; DMUP: DCMS-MnO₂-UBI_{29 − 41}-Pae

Methods

Synthesis of the core structure of DCNPs

The composition followed a reported scheme with slight modifications [38]. First, lanthanum acetate [1.0 mmol, Re(CH₃CO₂)₃ (Re = Er/Yb = 8/2)] was added to a flask containing oleic acid (OA, 6 mL) and 1-octadecene (ODE, 15 mL), heated to 150 ℃ to remove water. After cooling to room temperature (RT), a methanol solution of ammonium fluoride (NH₄F, 4.0 mmol) and sodium hydroxide (NaOH, 2.5 mmol) was added, followed by stirring for 30 min at 50 ℃. Subsequently, the temperature was raised to 120 ℃ to remove the methanol, and then rapidly heated to 300 ℃ in an argon atmosphere and kept for 50 min. After cooling to RT, ethanol was added to precipitate the DCNPs, which were centrifuged (11000 rpm, 10 min). The product (DCNPs) was dispersed in 20 mL of cyclohexane for further use.

Synthesis of core-shell (DCNPs) structures

The core-shell DCNPs were synthesized by a seed-induced growth method, and prepared by the same co-precipitation procedure. Typically, 1 mmol lutetium (III) acetate hydrate (Lu(CH₃CO₂)₃·xH₂O) was added to a solution containing ODE (15 mL) and OA (6 mL) in a 100 mL three-necked flask. The mixture was then heated to 150 ℃ for 30 min to form a clear solution. Subsequently, 4 mL of β-NaErF₄:Yb cyclohexane solution was added to the mixed solution, stirred at 70 ℃ for 20 min, and cooled to room temperature. Next, 10 mL of a methanol solution containing 4 mmol NH₄F and 2.5 mmol NaOH was added and stirred at 50 ℃ for 30 min. Then, the temperature was raised to 120 ℃ for 30 min for methanol removal. Subsequently, the system was heated to 300 ℃ for 50 min in an argon atmosphere, and after cooling to RT, the core-shell nanocrystals were precipitated with ethanol and dispersed in 20 mL of cyclohexane.

Preparation of DCNPs@mSiO₂-COSR (DCMS-COSR)

DCNPs@mSiO₂ were prepared using the inverse micellar method and modified with surface carboxylation. A 0.2 g cetyltrimethylammonium bromide (CTAB) was prepared in 30 mL deionized water solution, then 2 mL DCNPs solution was added, and stirred in a water bath at 80 ℃ until clarified, followed by ultrasound for 24 h. A 50 µL mixed solution of TEOS and Carboxyethylsilanetriol sodium salt (CHNaOSi, molar ratio of 1:1) in ethanol (10 mL) was configured. The sonicated mixed solution was placed in a 70 ℃ water bath, and the TEOS/CHNaOSi mixed solution was added dropwise, and the reaction was maintained for 6 h to coat the DCNPs surface with carboxyl-functionalized mSiO₂ (DCNPs@mSiO₂-COOH). Subsequently, the products were washed using anhydrous ethanol, 1 wt% NaCl methanol solution was used to remove the remaining CTAB from the products, and finally anhydrous ethanol was used to wash the products again before they were lyophilized and set aside.

Take 100 mg DCNPs@mSiO₂-COOH, 40 mg HOBT, and 42 mg EDC, respectively, and dissolve in 5 mL of deionized water. After magnetic stirring for 0.5 h, centrifugation to precipitate, re-dissolved in 10 mL of 50% ethanol solution (anhydrous ethanol: deionized water = 1:1). Then, add 64 µL of Benzyl Mercaptan, magnetic stirring for 1 h. Next, perform centrifugal precipitation, lyophilization, and storage. Finally, UBI_{29 − 41} was grafted onto the surface of the DCNPs@mSiO₂-COSR (DCMS-COSR).

Synthesis of DCNPs@mSiO₂-Mn-COSR (DM-COSR)

Metal-organic frameworks of ZIF-8 were synthesized in DCMS-COSR pores. First, 2-methylimidazole was loaded inside the DCMS-COSR pores by taking 200 mg of 2-methylimidazole completely dissolved in 3 mL of deionized water. And 100 mg of DCMS-COSR was taken and dissolved in 2 mL of deionized water. The above solutions were mixed and stirred magnetically for 2 h. The precipitate was collected by centrifugation and re-dissolved in 3 mL of deionized water. Then, 25 mg of Zinc nitrate hexahydrate (Zn(NO₃)₂·6H₂O) was dissolved in 2 mL of deionized water, mixed with the above solution, centrifuged with magnetic stirring for 1 h to collect the precipitate, and re-dissolved in 3 mL of deionized water. Subsequently, Mn nanozyme was attached to the ZIF-8 metal frame in the wells. 0.25 mg of potassium permanganate (KMnO₄) was dissolved in 1 mL of deionized water, and mixed with the above solution. The precipitate was collected by centrifugation with magnetic stirring for 30 min, and the product was lyophilized to obtain the DM-COSR.

Synthesis of DCNPs@mSiO₂-Mn-UBI_{29 − 41} (DMU)

A total of DM-COSR (50 mg) and UBI_{29 − 41} (0.1 mg) were used for the reaction, stirred at 4 ℃ for 24 h, centrifuged to precipitate, lyophilized to collect the product (DMU), and stored at 4 ℃.

Synthesis of Pae-loaded DMUP

A mixture of DMU (1.0 mg mL^− 1) with Pae/ethanol solution (4.0 mg mL^− 1) was stirred in a light-blocking vial at RT for 24 h. The mixture was centrifuged to obtain the supernatant and the absorbance of the supernatant was measured. A calibration curve for Pae was plotted based on the absorbance at 203.6 nm of a UV-2600 spectrophotometer under the same conditions. The loading ratio was calculated according to the following equation. Subsequently, it was washed, and vacuum freeze-dried to obtain DMUP powder.

$$\begin{aligned}\eqalign{& {\rm{ Loading\ ratio }}\ (\% ) = \\&\quad ({\rm{ weight\ of\ Pae\ in\ DMUP }} / {\rm{\ weight\ of\ DMUP }})\times 100\% \cr}\end{aligned}$$

(1)

Effect on macrophage polarization

Mouse monocyte macrophage leukemia (RAW264.7) cells were inoculated on cell slides in six-well plates at a density of 1 × 10⁶ cells per well and warmed at 37 ℃ for 24 h. Cells were pretreated with 200 µg mL^− 1 of different samples for 24 h, treated with 10 µg mL^− 1 lipopolysaccharide (LPS) for 12 h, and then washed three times with phosphate-buffered saline (PBS). Subsequently, cells were fixed with 4% paraformaldehyde for 15 min, washed several times with PBS, and closed with 3% bovine serum albumin (BSA) for 30 min. CD86 (1:100 dilution) and CD206 (1:200 dilution) primary antibodies were co-cultured with cells at 4 ℃ overnight, and washed with PBS. Next, samples were incubated with Cy3-labeled goat anti-rabbit IgG antibody (1:300 dilution) for 1 h. After three times of washing, the nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI). Finally, cells were observed using a fluorescence microscope (ECLIPSE C1, Nikon, Japan).

Hemolysis test

To obtain erythrocytes, fresh blood from rats was centrifuged at 1500 rpm for 15 min, washed gently, and diluted with saline. Then, 100 µL of erythrocyte solution was mixed with 1.1 mL of different concentrations of DMUP solution (25, 50, 100, 200, 400 µg mL^− 1). Meanwhile, 100 µL of erythrocyte solution was mixed with 1.1 mL of physiological saline and deionized water, which were used as negative and positive controls, respectively. After incubation at 37 ℃ for 3 h, the supernatants of all groups were collected by centrifugation at 1500 rpm for 15 min. The absorbance at 540 nm of each sample was determined by a microplate reader. The hemolysis rate was calculated according to the following formula:

$$\text { Hemolysis ratio } (\%) =\left(O D_e - O D_p\right) /\left(O D_w - O D_p\right) \times 100 \%$$

(2)

Intracellular ROS scavenging activity

Bacterial-derived LPS was selected for inducing endogenous oxidative stress in RAW264.7 cells. To induce endogenous oxidative stress, RAW264.7 cells were inoculated in 24-well plates at a density of 5 × 10⁵ cells per well and incubated for 24 h. Then, 100 µL of 10 µg mL^− 1 LPS containing Dulbecco’s modified eagle culture medium (DMEM) and different concentrations of samples were co-cultured for 12 h. The medium was then replaced with 2,7-dichlorodihydrofluorescein diacetate (DCFH-DA), which could be oxidized to 2,7-dichlorofluorescein (DCF) by ROS, and then co-cultured with the cells for 30 min. The cells were observed with an inverted fluorescence microscope, and the fluorescence quantification of DCF was measured by fluorescence analysis (Ex: 488 nm, Em: 525 nm).

In vivo imaging experiments

All animal experimental protocols were approved by the Animal Ethics Committee of Nanchang University, China (NCULAE-20221228041). Eighteen male Sprague Dawley (SD) rats (8 weeks old, 150–200 g) were randomly divided into 7 groups (n = 6). 500µL suspension of physiological saline with various concentrations (0, 10⁴, 10⁵, 10⁶, 10⁷, 10⁸, 10⁹CFU mL^− 1) of mixed P. gingivalis, Escherichia coli (E. coli), and Staphylococcus aureus (S. aureus) was injected into the bilateral hind leg muscles of the rats. After three days, infected rats with muscle infections were obtained for the experimental groups. Moreover, the infected muscles of the right side of the rats were sampled. H&E staining was conducted to observe the cell morphology of the muscle tissues and to verify the successful establishment of the infection model. Subsequently, NIR-II small animal imaging was performed 10 min after the injection of 500 µL DMUP at the synchronized site (groups of 0, 10⁶, 10⁷, 10⁸, and 10⁹CFU mL^− 1) to observe the DMUP-labeled fluorescence imaging area images.

Modeling and treatment of experimental periodontitis

To evaluate the combined therapeutic efficacy of DMUP in periodontitis, we established a typical bacterial-induced periodontitis model in rats. After 1 week of acclimatization, 24 male SD rats (8 weeks old, 150–200 g) were randomly divided into 4 groups (n = 6): no treatment group (Sham), periodontitis + PBS (Con), periodontitis + MC, and periodontitis + DMUP. 3–0 silk threads were used to ligate the left maxillary second molars of rats, while daily injections of a mixed bacterial solution of P. gingivalis, S. aureus, and E. coli mixture (10⁹ CFU mL^− 1, 500 µL), and high-sugar drinking water (100 g L^− 1 glucose). After a total of 14 days, 100 µL of PBS, MC, DMU, and DMUP (200 µg mL^− 1) was injected into periodontal pockets every alternate day. Four weeks later, maxillae with teeth harvested from the executed rats were collected for micro-computed tomography (Micro-CT) analysis, H&E staining, immunohistochemical staining, and Masson staining. Samples were scanned by Micro-CT (NEMO NMC-200, PINGSeng Healthcare Inc.). The vertical distance between the ABC and CEJ as well as the bone volume per tissue volume (BV/TV), and trabecular thickness (Tb. Th) were measured by three-dimensional reconstruction (AVATAR 1.7.0, PINGSeng Healthcare Inc.).

Statistical analysis

All quantitative data are expressed as mean ± s.d. Data processing and statistical analysis were performed using GraphPad Prism 9.5.0. All experiments were repeated at least three times. Data were analyzed using a t-test and one-way analysis of variance (ANOVA). Data are expressed as means ± s.d. (n ≥ 3), with *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001 denoting significance.

Results and discussion

Synthesis and characterization of DMUP

Figure 2a illustrates the synthesis process of the NIR-IIb fluorescence imaging multifunctional nanozyme platform (DMUP). First, a thermal decomposition method was utilized to synthesize 980 nm NIR-responsive DCNPs with a core-shell structure (Fig. 2b and c, and S1) [38]. The resulting DCNPs were homogeneous monodispersed hexagonal core-shell nanoparticles with a particle size of about 35 nm. To increase the hydrophilicity and facilitate the subsequent functional steps, a layer of mSiO₂ approximately 30 nm thick was grown on the surface of the DCNPs, resulting in DCMS (Fig. 2d and S2). Subsequently, the DCMS surface was functionalized to form DCMS-COSR. Then, Mn nanozyme was precipitated into the pores of DCMS-COSR, followed by the grafting of UBI_{29 − 41} onto its surface and the loading of Pae into remaining pores, ultimately forming the resultant complex called DMUP. As shown in Fig. 1, the transmission electron microscopy (TEM) image revealed that the Mn nanozyme nanoparticles appeared on the surface of the DCMS in a characteristic ribbon-like structure [27, 30]. Moreover, the scanning electron microscopy (SEM) results showed a raised shape at the pores (Fig. S3). The successful preparation of DMUP was also proved by the elemental mapping results (Fig. 2f).

Fig. 2

Synthesis and characterization of DMUP. (a) A schematic diagram showing the synthesis of DMUP. The TEM images of (b) DCNPs core, (c) DCNPs, (d) DCMS, and (e) DMUP. (f) Elemental mapping analysis of DMUP. (g) Zeta potential changes of the DM-COSR before and after being surface grafted of UBI_{29 − 41}. (h) XRD patterns of DCNPs, DCMS, and DMUP. (i) XPS energy spectrum for DMUP. (j) The fitted peak spectrum of the Mn element after deconvolution. (k) FTIR spectra of Core, DCNPs, DCMS, DCMS-COOH, DCMS-COSR, and DMU. (l) NIR-IIb fluorescence broad spectra of the DCNPs core, DCNPs and DCMS. (m) NIR-IIb fluorescence spectrum of DCNPs under 980 nm excitation and NIR-IIb absorption spectrum of DMUP. (n) Fluorescence spectra of DMUP synthesized with different concentrations of Mn doping in the NIR-IIb band. Data are means ± s.d. (n ≥ 3)

The grafting on the mSiO₂ surface began with modifying the N-terminus of UBI_{29 − 41} with cysteine (Fig. S4). Then, the DCMS-Mn (denoted as DM) surface reacted via thioesterification to form DM-COSR (Fig. S5) [39]. Finally, as shown in Figure S6, the peptide was covalently attached via native chemical ligation (NCL) to form DCNP@mSiO₂-Mn-UBI_{29 − 41} (denoted as DMU) [35]. We also complemented the covalent docking simulations, with Fig. S7 showing that the thiol and amine groups of cysteine in UBI_{29 − 41} could form covalent bonds with the carbonyl group of DM-COSR, resulting in a stable amide bond with a docking score of − 9.226 kcal/mol (a docking score below − 5 kcal/mol indicates bond stability). The successful synthesis of DMU was confirmed by comparing the changes in zeta potential and pH during the grafting process (Fig. 2g and S8). Fig. 2h shows the X-ray diffraction (XRD) pattern of the system. The diffraction peaks could be attributed to the hexagonal phase crystalline form of β-NaYF₄ (JCPDS No. 00-016-0334). Additionally, as for the mSiO₂ of DCMS, a broad absorption band of amorphous SiO₂ could be observed at 2θ = 22°. In addition, the results of Fig. 2h showed characteristic peaks consistent with the ZIF-8 crystal phase (JCPDS No. 00-062-1030), and matched with diffraction peaks of MnO₂ (JCPDS No. 00-072-1982). This supported the formation of metal-organic frameworks in DMUP pores and the hypothesis that MnO₂ served as the nanozyme’s active component. And ZIF-8 metal-organic framework contributed to acid-responsive functionality. The ultraviolet-visible spectroscopy (UV-vis) spectrum of DM-COSR revealed the characteristic broad absorption peak after Mn nanozyme precipitation (Fig. S5). Moreover, the X-ray photoelectron spectroscopy (XPS) fitting results revealed the Mn(IV) oxidation state in the DMUP (Fig. 2i and j). It was shown that the successful precipitation of Mn nanozyme nanoparticles demonstrated the potential performance of tetravalent manganese antioxidant nanozyme [40]. Meanwhile, fourier transform infrared (FTIR) was utilized to detect the functional groups (Fig. 2k). The FTIR spectra revealed C − C and C − H peaks at 2900 cm^− 1, which were characteristic of the DCNPs [38]. The C = O stretching vibration peaks between 1700 and 1725 cm^− 1, and the enhancement of the O − H bonds between 2500 and 3300 cm^− 1, demonstrated the successful modification of the − COOH (DCMS-COOH). Moreover, the characteristic peaks of the aryl ring at 750–800 cm^− 1 indicated the success of thioesterification (DCMS-COSR) [39].

The fluorescence properties during the synthesis of DMUP were characterized as shown in Fig. 2l. DCMS exhibited a strong emission peak in the 1500–1600 nm NIR-IIb fluorescence band under 980 nm excitation. The broad absorption of Mn nanozyme in the NIR band (Fig. 2m) led to FRET action along with the precipitation of Mn nanozyme in the pore. As shown in Fig. 2n, the NIR-IIb fluorescence of DMUP was gradually quenched. The final effectively quenched KMnO₄ synthesized concentration was estimated to be 62.5 µg mL^− 1. The successful loading of Pae was illustrated in Fig. S9 and S10. The XPS spectrum and inductively coupled plasma-mass spectrometry (ICP-MS) results of DMUP revealed the presence of Er, Yb, Lu, and Si elements in DMUP, further confirming the successful loading of each component in DMUP (Fig. S11–S15).

Assessment of physicochemical properties and biosafety of DMUP

ROS are byproducts of normal metabolism, typically neutralized by the antioxidant system of body under normal physiological conditions. During the progression of periodontitis, the immune system becomes hyperactive due to the increase in anaerobic pathogenic bacteria. This process increases the amount of ROS in the periodontal tissues, which causes an imbalance between ROS and the antioxidant system, resulting in a toxic response, thereby exacerbating periodontitis [41]. Meanwhile, with the accumulation of plaque, the periodontal microenvironment shifts from a normoxic to a hypoxic phase. The resultant deep hypoxic environment further exacerbates the colonization of pathogenic anaerobic bacteria [42, 43]. Therefore, in order to modulate the changes in the periodontitis microenvironment, this study verified the acid response, antioxidant system release, CAT-like, SOD-like, and catalytic antioxidant nano-enzymatic properties of DMUP.

Figure 2a shows the involvement of reactive oxygen species in cellular respiration. First, O₂ gained an electron to form ·O₂⁻, which was then converted to H₂O₂ by SOD. Next, CAT decomposed H₂O₂ to H₂O and O₂ [28]. During the anti-inflammation process, the excess ROS is scavenged to produce O₂, which alleviates the hypoxic environment and inhibits the anaerobic bacteria [26]. The ICP-MS and UV-vis spectroscopy (Fig. S16 and S17) results revealed that Mn oxides and Pae release increased under acidic conditions, providing the basis for a cascade catalytic reaction to excess ROS. Then, Fig. 2b demonstrated that DMUP could recover NIR-II fluorescence in acidic conditions due to the removal of Mn oxides from the pores, which eliminated FRET effects [30]. Electron spin resonance (ESR) was utilized to confirm that DMUP exhibited CAT-like and SOD-like activities by scavenging ·OH, ·O₂⁻ and H₂O₂. Figure 2c–e showed that DMUP scavenges significant amounts of ·OH, ·O₂⁻ and H₂O₂. Additionally, this study also addressed the issue of hypoxia in periodontal pockets. The H₂O₂ produced by the hyperactive immune response in inflamed tissues and the periodontal pocket bacterial biofilm during metabolism aggravates tissue damage [44]. Mn nanozyme incorporation provided DMUP with a strong ability to decompose H₂O₂ into O₂ (Fig. S18), which was extremely significant in the treatment of periodontal diseases. Adequate O₂ production effectively alleviated periodontal pocket hypoxia, improved the hypoxic environment [26], and inhibited the proliferation of anaerobic pathogenic bacteria. As shown in Fig. 2f, DMUP could catalyze O₂ production in different concentrations of H₂O₂ solution.

Encouraged by the excellent ROS scavenging and catalytic H₂O₂ O₂-producing ability of DMUP in vitro, we next tested these properties in cells. First, the effect of DMUP on the proliferation of the macrophage RAW264.7 cells, an important immune line of defense, was evaluated. As shown in Fig. S19-S22, the results of cell counting kit-8 (CCK-8), cell live/dead staining, and hemolysis test indicated that DMUP at a concentration of 200 µg mL^− 1 was considered safe and used in subsequent cell experiments. Subsequently, bacterial LPS was selected to induce an inflammatory model of endogenous oxidative stress in RAW264.7 cells [34]. The ROS probe DCFH-DA was utilized to determine the antioxidant effect of DMUP. Compared with the control (Con) group, the LPS group significantly promoted ROS production with strong fluorescence (Fig. 2g and h). In contrast, there was a fluorescence quenching in the DMUP group, indicating a reduction in the intracellular ROS level. This clearly suggested that DMUP reduced endogenous oxidative stress. Next, the intracellular O₂ production capacity of DMUP in RAW264.7 was investigated by tris (4,7-diphenyl-1,10-phenanthroline) ruthenium (II) dichloride ([Ru(dpp)₃]Cl₂), a luminescent probe commonly used for O₂ identification and quantification (Fig. 2i and j). In contrast to the Con group, the amplification of red luminescence indicated a decrease in the O₂ content of the LPS-induced inflammatory cells. After incubating with DMUP for 2 h, the attenuation of red luminescence confirmed O₂ production (Fig. S23). This result demonstrated that DMUP had a good intracellular ability to catalyze O₂ production. Therefore, the catalytic decomposition of ROS could reduce inflammation by regulating the redox imbalance and hypoxic state at the inflammatory site.

Fig. 3

Physicochemical properties and biosafety of DMUP. (a) Antioxidant and O₂-producing antimicrobial principle of DMUP. (b) NIR-II fluorescence spectra of DMU, DMUP, and DMUP (pH = 6.5). (c − e) ESR spectra of scavenging of ·O₂^–, ·OH, and H₂O₂ by DMUP. (f) Analysis of the O₂ yield as a result of the interaction of DMUP with different concentrations of H₂O₂. (g) Representative fluorescence microscopy images of DMUP on intracellular ROS. Blue: nucleus of cell, Green: intracellular ROS. (h) Relative fluorescence quantification results of intracellular ROS by DMUP. (i) Images of fluorescence microscopy detection intracellular O₂. Blue: nucleus of cell, Red: hypoxia in cells. (j) Results of O₂ content occupancy analysis. Data are means ± s.d. (n ≥ 3), ns: no significant difference, **p < 0.01, ***p < 0.001, ****p < 0.0001

Validation of DMUP targeted bacteria and imaging capabilities

UBI_{29 − 41} targets bacterial surfaces by electrostatically adsorbing to phosphatidylglycerol and attracting hydrophobic bacterial membranes. Therefore, UBI_{29 − 41} could be specifically targeted to the bacterial surface by DMUP through surface grafting, which was also the basis for accurate bacterial imaging (Fig. 4a). SEM was used to study the interaction between DMUP and P. gingivalis, E. coli, and S. aureus after 30 min of incubation at pH 6.5 (Fig. 4b − d). The corresponding images revealed that DMUP was able to target and adhere to the surface of all three bacteria. The targeting efficacy of DMUP was evaluated by loading fluorescein isothiocyanate (FITC, denoted as DMUP-F) into DMUP. DMUP-F was co-cultured with P. gingivalis for 5, 10, 20, and 30 min and observed under a fluorescence microscope (Fig. S24a). Combined with the results of the quantitative fluorescence analysis (Fig. S24b), it was confirmed that DMUP effectively targeted bacteria after co-cultivation for 10 min.

The DMUP (200 µg mL^− 1) was evaluated for imaging P. gingivalis, E. coli, and S. aureus. As shown in Fig. 4e, all three bacteria were mixed in equal proportions, grouped in different concentrations, and incubated with DMUP for 10 min. A NIR-II small animal imager was used to detect the fluorescence intensity of each group. The bacterial concentration positively correlated with the fluorescence intensity in the 10⁴−10⁹ CFU mL^− 1 bacterial concentration group (Fig. 4f). In contrast, the fluorescence intensity of the 10¹–10⁴ CFU mL^− 1 bacterial concentration group was too weak to be recorded. Moreover, there was no significant difference in the fluorescence intensity in the 10⁹–10¹¹ CFU mL^− 1 bacterial concentration group. This lack of difference was likely due to the limited acidity of the environment around the bacteria, which did not decrease indefinitely with the increased bacterial concentration [45]. Therefore, the findings of the experiments indicated that the effective bacterial detection concentration was 10⁴ CFU mL^− 1 or more when using a DMUP concentration of 200 µg mL^− 1.

The classical model of bacteria-induced periodontitis in rats was used to evaluate the effect of DMUP for imaging bacteria in periodontal pockets (Fig. 4g and h). The results showed that DMUP still had the ability to image bacteria in the rugged structure of the periodontal pockets. The Con group had the highest fluorescence intensity, while the DMUP group had the lowest. Then, comparative bacterial plating experiments were conducted on the periodontal pockets of different groups to evaluate bacterial load differences. Figure S25 showed that the DMUP group exhibited the strongest antibacterial effect, consistent with the intensity of fluorescence in the corresponding group. These results supported DMUP’s targeting and imaging capabilities in periodontal infection sites. However, the periodontal pockets of rats were too small, making it difficult to observe bacterial concentration-dependent changes in fluorescence intensity. To better simulate larger-scale in vivo imaging, we selected an SD rat leg muscle infection model. This model exhibits strong similarities to periodontal tissue infections, such as excess ROS, hypoxic conditions, and the coexistence of anaerobic and aerobic bacteria [46]. The bacteria displayed prominent proliferation when injected at 10⁶ CFU mL^− 1 in the rat (Fig. S26). Furthermore, the muscle fibers were infiltrated with many inflammatory cells (Fig. S27), indicating that injecting 1 mL bacteria (10⁶ CFU mL^− 1) or more could successfully create a model leg muscle infection. To further verify the performance of in vivo bacterial imaging, as illustrated in Fig. 4i, different concentrations of mixed bacteria were imaged in the leg muscle after injecting 500 µL DMUP (200 µg mL^− 1). The imaging results of small animals, analyzed by the fluorescence intensity ratio, showed a positive correlation with the injected bacterial concentrations, indicating that DMUP could effectively image the bacteria within the inflammation (Fig. 4j).

Fig. 4

Imaging capabilities of DMUP-targeted bacteria. (a) Schematic of DMUP-targeted bacteria and acid-responsive fluorescence imaging. Representative SEM images of DMUP targeting (b) P. gingivalis, (c) E. coli, and (d) S. aureus. (e) NIR-II fluorescence images of DMUP on different concentrations of mixed bacteria, and (f) quantification results of fluorescence intensity. (g) NIR-II images of infected areas of periodontitis in rats, and (h) the results of fluorescence relative intensity. (i) NIR-II images of the bacterial infection model in the leg, and (j) the results of fluorescence relative intensity. Data are means ± s.d. (n ≥ 3), ns: no significant difference, *p < 0.05, **p < 0.01

Study of antimicrobial and anti-biofilm activity of DMUP

Bacteria and plaque as initiators of periodontitis. The antimicrobial activity of DMUP was examined on the following bacteria: P. gingivalis (typical pathogenic anaerobic bacteria), S. aureus (typical aerobic bacteria), and E. coli (typical facultative anaerobic bacteria). First, exogenous H₂O₂ (1 mM) was added to the medium and the pH was adjusted to 6.5. A titrimetric method was utilized to examine the antimicrobial activity of different concentrations of DMUP against 10⁵ CFU mL^− 1 of P. gingivalis, E. coli, and S. aureus. The results revealed that the minimal inhibitory concentrations of DMUP against P. gingivalis and E. coli were 200 µg mL^− 1 and 1600 µg mL^− 1, respectively (Fig. S28). In contrast, the antimicrobial efficiency against the aerobic S. aureus was lower, with no significant bacterial inhibition at 1600 µg mL^− 1, which was in keeping with the antimicrobial needs of periodontitis. Since the main causative organisms of periodontitis are anaerobes, treatment focuses on anti-anaerobes. Therefore, in conjunction with the biosafety concentration, we chose the lowest antimicrobial concentration of DMUP (200 µg mL^− 1) against the anaerobic bacterium P. gingivalis for subsequent experiments.

SEM was used to observe the different degrees of cell membrane rupture and contraction of P. gingivalis, E. coli, and S. aureus by 200 µg mL^− 1 of DMUP (Fig. 5a). As shown in Fig. 5b, the spread plate method revealed antimicrobial rates of about 99.9%, 95.7%, and 63.5%. This indicated that DMUP exhibited varying antimicrobial activities against the three bacteria. Particularly, the inhibition of the anaerobic bacteria by DMUP was superior to that shown by a conventional drug and the MC group. Moreover, since plaque biofilm was considered to be the initiating factor of periodontitis, the efficiency of DMUP for removing P. gingivalis biofilm was examined by live/dead staining (Fig. S29 and S30) and crystal violet (CV) staining experiments. As shown in Fig. 5c and d, the results revealed that with a removal rate of about 96%, DMUP exhibited the best removal efficiency against P. gingivalis biofilm. Against E. coli and S. aureus, the removal rates were about 81%, and 27%, respectively. Thus, this indicated that DMUP exhibited superior performance in removing anaerobic bacterial biofilm.

This study also elucidated the antibacterial mechanism of DMUP. The destruction of bacterial cell membranes in a biofilm was investigated with a protein leakage assay. The results revealed that protein leakage was consistent with those of the anti-biofilm experiments (Fig. S31–S33). Subsequently, the expression changes in genes related to P. gingivalis, including fimbrial protein A (fimA), keratinsulfate glycoprotein (kgp), arginine-specific gingipain A (rgpA), and arginine-specific gingipain B (rgpB), were detected after treatment. The analysis revealed that DMUP effectively reduced the expression levels of the above genes (Fig. 5e–h). Thus, the findings suggested that DMUP destroyed the cell membranes of anaerobic bacteria by inhibiting the expression of the virulent genes of P. gingivalis. With an antibacterial efficiency higher than that of conventional drug MC, DMUP exhibited an excellent anti-biofilm effect against anaerobic bacteria. However, its antibacterial performance against aerobic bacteria was limited. As periodontitis progresses, anaerobic bacteria primarily contribute to tissue destruction [47]. Thus, the superior performance of DMUP in inhibiting anaerobic bacteria was in line with the requirements for periodontitis treatment.

Fig. 5

Antibacterial and anti-biofilm activity of DMUP. (a) Representative plate images and SEM images after co-incubation of DMUP with P. gingivalis, E. coli, and S. aureus, respectively. (b) Results of antimicrobial properties after quantification. (c) Representative light microscopic pictures of CV staining of three biofilms. (d) Anti-biofilm rates of different groups for the three biofilms. (e–h) Changes in the expression of P. gingivalis virulence genes after treatment in different groups. Data are means ± s.d. (n ≥ 3), ns: no significant difference, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

Experimental studies of anti-inflammatory in vitro

Several signaling pathways, such as NF-κB and MAPK, are activated by excess ROS generated by an inflammatory response, which promotes the expression of inflammatory factors [48, 49]. ROS clearance can block intracellular signaling functions and damage cellular structures, alleviating symptoms and retarding disease progression [50, 51]. Thus, an important component of the treatment of inflammatory and immune diseases involves controlling ROS levels and reducing the expression of inflammatory factors. In this study, the activated inflammation was modeled with the mouse macrophage cell line RAW264.7 cells. The anti-inflammatory capacity of DMUP in vitro was determined by examining the gene expression of key pro-inflammatory and anti-inflammatory mediators in the inflammatory process after inducing inflammation in the RAW264.7 cells (Fig. 6a). In the quantitative real-time polymerase chain reaction (qRT-PCR) results, the gene expression of pro-inflammatory factors tumor necrosis factor-α (TNF-α), transforming growth factor-β (IL-1β), and interleukin-6 (IL-6) increased significantly in LPS-treated RAW264.7 cells, while the gene expression of anti-inflammatory factor transforming growth factor-β (TGF-β) decreased relatively (Fig. 6b and c and S34). After DMUP (200 µg mL^− 1) treatment, there was a significant decrease in the expression of TNF-α, IL-1β, and IL-6. In contrast, expression of the anti-inflammatory factor TGF-β was elevated. Furthermore, the expressions of TNF-α and IL-6 were significantly reduced in the DMU group. In contrast, the expression of IL-1β was not statistically different from that of the LPS group. The gene expression of TNF-α and IL-1β was significantly more reduced in the Pae group than that in the DMU group. However, the reduction in the expression of IL-6 was lower in the Pae group than that in the DMU group. The changes in the expression of the three pro-inflammatory factors revealed that the DMUP group displayed the strongest ability to reduce them. Although there were differences between the Pae and DMU groups in the expression of the pro-inflammatory factor TGF-β, DMUP had the most pronounced effect on its elevation. This might be attributed to the synergistic anti-inflammatory effect of Mn nanozyme and polyphenolic antioxidants in the DMUP system.

The effect of DMUP on the immune function of RAW264.7 cells was examined by performing qRT-PCR assays to detect the M1-type and M2-type markers inducible nitric oxide synthase (iNOS) and arginase 1 (Arg-1) (Fig. 6d and e). Additionally, immunofluorescence (IF) assays were performed to detect cluster of differentiation 86 (CD86) and cluster of differentiation 206 (CD206) (Fig. 6f). The results exhibited an increase in the intensity of the M1-type macrophage markers iNOS and CD86. In contrast, the intensity of the M2-type macrophage markers, Arg-1 and CD206, decreased. This led to an increase in the M1/M2 ratio, which was indicative of a pro-inflammatory profile. The expression of anti-inflammatory factors in Fig. 6b–g, showing that the Pae group (the natural antioxidant) reduced the expression of genes of M1-type inflammatory cells and promoted the production of M2-type anti-inflammatory cells, compared to the LPS group. Mn nanoenzyme mimicked CAT and SOD enzyme activities [24], effectively scavenging excess ROS in the inflammatory environment. This property of Mn nanoenzyme regulated redox balance, reduced M1 macrophage-mediated inflammation, and promoted M2 macrophage differentiation, which played an anti-inflammatory and immunomodulatory role. The DMU group (Mn nanoenzyme loaded system) showed lower expression of TNF-α, iNOS, and CD86 (M1 markers) while increasing TGF-β, Arg-1, and CD206 (M2 markers). These results indicated that Mn nanozymes also contributed to macrophage phenotype switching. The results of the DMUP group were the combined effect of Mn nanozyme and Pae. Then, the high M1/M2 ratio of LPS group was reversed when the macrophages were cultured with DMUP (Fig. 6g). The above results suggested that DMUP had anti-inflammatory properties that favored anti-inflammation and tissue repair.

Fig. 6

In vitro anti-inflammatory efficacy evaluation of DMUP. (a) Schematic representation of induced inflammation RAW264.7 cells, as well as anti-inflammatory and induced autophagy. Created with BioRender. com. Gene expression of (b) pro-inflammatory factor TNF-α and (c) anti-inflammatory factor TGF-β after group treatment. (d) Expression of Arg-1, a representative gene of cellular anti-inflammatory phenotype M2, and (e) expression of iNOS, a representative gene of pro-inflammatory phenotype M1. (f) Representative fluorescence images of different treatment groups. (g) Quantitative fluorescence images of M1 (CD86), and M2 (CD206) phenotypes of RAW264.7 cells. Data are means ± s.d. (n ≥ 3), ns: no significant difference, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

Molecular mechanisms of anti-inflammation and induction of autophagy in vitro

To determine the molecular mechanisms of DMUP anti-inflammatory and autophagy, RAW264.7 cells were divided into RAW, RAW + LPS, and RAW + LPS + DMUP groups (abbreviated as Con, LPS, and DMUP) for seq-RNA and bioinformatics analysis induction in vitro. The results of the principal component (PC) analysis (Fig. S35) showed reasonable homogeneity within individual groups and differences between groups. All three sample groups met the quality control requirements. Based on the results of the volcano plot (Fig. S36), trend plot (Fig. S36), and trend heat map (Fig. 7a and S35), 89 genes were found to be significantly upregulated, and 319 genes were significantly downregulated in the DMUP group compared with the LPS group. Additionally, the results of the gene ontology (GO) analysis of these differentially expressed genes (DEGs, Fig. 7b) showed an enrichment of genes related to immune, inflammatory, and metabolic responses. The results of the kyoto encyclopedia of genes and genomes (KEGG) analysis revealed that these genes were significantly related to the PI3k-Akt, mTOR, NF-κB, and TNF signaling pathways (Fig. 7c), which further supported that DMUP played a role in immune cell autophagy and inflammation regulation.

Autophagy is an intracellular degradation process that reduces inflammatory injuries via the removal of potential pro-inflammatory molecules and maintains intracellular stability as well as immune homeostasis. The activation of autophagy has the potential to alleviate the expression of inflammatory factors, such as TNF-α and IL-1β. In contrast, autophagy is increased by the expression of anti-inflammatory cytokines, such as TGF-β, limiting the over-activation of inflammatory signals [52]. One of the important pathways in the regulation of autophagy is the Akt/mTOR pathway [53], which significantly affects the transition of macrophages from M1-type to M2-type via the regulation of autophagic activity [54]. Of these, the mechanistic target of rapamycin complex 1 (mTORC1) is a major negative regulator of autophagy, which inhibits autophagy via the suppression of key molecules for autophagy initiation (e.g., ULK1). Additionally, mTORC1-mediated inhibition could initiate the autophagy process, affecting the activity and functional status of macrophages. This could result in reduced production of pro-inflammatory factors, suppressing M1-type macrophage pro-inflammatory potential, promoting their transition to M2-type macrophages, facilitating the M2-type characteristics, and enhancing their tissue repair ability and anti-inflammatory cytokine production [55]. Here, the impact of DMUP on autophagic activity was confirmed via a number of autophagosomes and autophagic protein markers. The Monodansylcadaverine (MDC) staining results revealed that M1-type macrophages treated by DMUP possessed higher autophagosome content than untreated M1-type macrophages (Fig. S39 and S40). Next, Western blot (WB) assay was used to determine the expression of autophagic activity protein markers, autophagy initiation markers (ULK1), autophagic vesicle markers (ATG16, ATG5), and autophagy flux markers (LC3II/LC3I). The relative band intensities suggested that the levels of ULK1, ATG16, ATG5, and LC3II/LC3I were elevated in M1-type macrophages treated with DMUP (Fig. 7d and e). This implied that DMUP-activated autophagy and enhanced the autophagic flux in M1-type macrophages. Next, the signaling changes of Akt and mTOR in DMUP-treated M1-type macrophages were investigated. Initially, the Akt and mTOR signaling was active in M1-type macrophages; However, after incubation with DMUP, p-Akt and p-mTOR signaling was inactivated (Fig. S41 and S42). The above results suggested that DMUP promoted autophagic activity via the inactivation of the Akt/mTOR signaling pathway to further prevent macrophage conversion to pro-inflammatory M1-type subtype, facilitating tissue anti-inflammation and repair.

Thus, the multi-enzyme system of DMUP reduced the expression of inflammatory cytokines, enhanced the autophagic flux of immune cells, and regulated both immune cell function and inflammation reduction. This, in turn, improved the tissue repair potential and anti-inflammatory cytokine production, preserving the intracellular stability and immune homeostasis and finally reducing the inflammatory injury.

Fig. 7

To investigate the molecular mechanisms of DMUP anti-inflammation and autophagy induction at the cellular level. (a) Clustering heatmap of RNA-seq with gene expression results (LPS vs DMUP), blue: low expression; red: high expression. (b) GO enrichment analysis (LPS vs DMUP). (c) KEGG enrichment analysis (LPS vs DMUP), and related pathways pooled by differential genes. (d) WB assay determination of autophagy-related protein expression levels and (e) quantitative analysis. Data are means ± s.d. (n ≥ 3), *p < 0.05, **p < 0.01, ***p < 0.001

Regulating the effect of the inflammatory environment on in vitro osteogenesis

During inflammation, regulation of the mechanisms involved in the alteration of macrophage polarization status, autophagy induction, decreasing ROS and pro-inflammatory factor levels, and increasing anti-inflammatory factor expression can create a microenvironment favoring osteogenesis [55]. The study also explored the role of DMUP-regulated inflammatory state macrophages in the regulation of osteogenesis in vitro.

As shown in Fig. 8a, the conditioned medium was configured. First, the induction of RAW264.7 cells by LPS to the inflammatory state. Next, DMUP (200 µg mL^− 1) was added and cultured for 24 h. The supernatant was used to culture the mouse embryonic osteoblast precursor (MC3T3-E1) cell [56]. The results showed that the conditioned medium, post-LPS induction (Con group), reduced the alkaline phosphatase (ALP) activity of MC3T3-E1 cells. Also, the DMUP-treated conditioned medium enhanced ALP activity of MC3T3-E1 cells (Fig. 8b). The alizarin red staining (ARS) showed similar results (Fig. 8c). Their micrographs were also significantly different (Fig. 8d). The DMUP-conditioned medium group stimulated the formation of mineralized nodules. Subsequently, the results of the qRT-PCR assay showed that DMUP upregulated the expression of osteogenesis-related genes collagen type I (COL-1), osteocalcin OCN, and runt-related transcription factor 2 (Runx2) (Fig. 8e–g).

The osteopontin (OPN) and Runx2 are two vital proteins involved in osteogenesis, which include various osteogenesis-related pathways (wnt/β-catenin, TGF-β/Smad, PI3K/Akt) and impact osteoblast differentiation, bone matrix formation, and mineralization [57]. Here, a WB assay was performed to evaluate the expression of OPN and Runx2. The DMUP-conditioned medium group exhibited elevated expression of OPN and Runx2 in MC3T3-E1 cells (Fig. 8h–j). Thus, the DMUP-conditioned inflammatory microenvironment affected osteogenic metabolism at both the gene and protein levels, enhancing mineralized nodule formation and promoting cellular osteogenic potential.

Fig. 8

Effect of DMUP on osteoblast mineralization after adjusting the inflammatory microenvironment. (a) Schematic of conditioned medium preparation. Created with BioRender. (b) Semi-quantitative results of ALP staining. (c) Semi-quantitative results of ARS staining of mineralized nodules, and (d) microscopic staining diagrams. Changes in the expression of osteogenesis-related genes (e) COL-1, (f) OCN, and (g) Runx2. (h) WB assay determination of osteogenesis-related protein expression levels, and quantification of (i) OPN and (j) Runx2. Data are means ± s.d. (n ≥ 3), ns: no significant difference, *p < 0.05, **p < 0.01, ***p < 0.001

In vivo assessment of the anti-inflammatory effect of DMUP

The classic bacterial-induced periodontitis model in rats, using silk-knot and mixed bacterial fluids (P. gingivalis, E. coli, and S. aureus) injections, was used to evaluate the in vivo anti-inflammatory effect of DMUP (Fig. 9a) [55]. Based on the bleeding analysis by intraoral probing in rats 28 days post-treatment in different groups, the DMUP group exhibited the least amount of bleeding (Fig. S43). After injecting the bromothymol blue indicator into the periodontal pocket, the inflamed tissue appeared yellow-green (Fig. S44a and S44b). The pH testing of the supernatant from homogenized infected periodontal tissue also showed a yellow-green color (Fig. S44c and S44d). Compared to the color standard of the indicator (Fig. S44e), the pH value was approximately 6.5, confirming an acidic environment. Blood tests after DMUP treatment showed no significant changes in white blood cells (WBC), mean corpuscular hemoglobin (MCH), and platelets (PLT) levels (Fig. S45). H&E staining of main organs (Fig. S46) showed no inflammation or tissue damage in the DMUP-treated group. These results confirmed that DMUP had biosafety in vivo. Subsequently, Masson and hematoxylin-eosin (H&E) staining of periodontal tissues revealed significant inflammatory cellular infiltration and collagen loss in the Con group (Fig. 9b and S47), whereas DMUP ameliorated these abnormalities and enhanced collagen deposition. The results of the semi-quantitative analyses also showed a better effect than that of the MC group (Fig. 9c and d). Next, immunohistochemical staining was used to determine the expression of pro-inflammatory factors (IL-6, IL-1β, and TNF-α) and anti-inflammatory factors (IL-10) to further assess the in vivo anti-inflammatory effects of DMUP (Fig. 9e and S48–50). The DMUP group showed reduced expression of IL-6, IL-1β, and TNF-α and enhanced expression of IL-10, further confirming that DMUP exhibited a satisfactory anti-inflammatory effect and was stronger than that of the MC group (Fig. 9f and g and S51, S52). The results suggested that DMUP reduced the inflammatory response and promoted periodontal tissue regeneration, thus playing an effective therapeutic role in the bacterial periodontitis model.

In order to evaluate the in vivo osteogenic effect of DMUP, the therapeutic effect of DMUP was evaluated using Micro-CT and histological staining after establishing a classical bacterial-induced periodontitis model in rats. The vertical distance (attachment loss, AL) between the alveolar bone crest (ABC) and the cementoenamel junction (CEJ) was considered the therapeutic index by 3D reconstruction [58]. The Con group had the longest distance between ABC and CEJ compared with the Con group, showing significant loss of alveolar bone and successful periodontitis modeling (Fig. S53). DMUP increased the values of BV/TV, Tb. Th and bone mineral density (BMD) (Fig. 9h–j and S54) and reduced the distance between ABC and CEJ compared to the Con group. The results of H&E staining of alveolar bone volume at the heel bifurcation (Fig. S55 and S56) showed that DMUP group had the highest bone volume among the groups. Next, IF was used to study the expression of OCN and Runx2 by different groups (Fig. 9k). The red fluorescence represented the expression of OCN and Runx2, respectively, and the blue color represented the DAPI stained cell nuclei. Based on the results of the quantitative fluorescence, the DMUP group exhibited elevated expression of OCN and Runx2, which was significantly higher than that of the MC group (Fig. 9l and m). Thus, DMUP significantly regulated bone formation and bone differentiation-related factors, promoting bone formation in periodontitis. The MC group also exhibited some bone formation effects. However, those were significantly different from the DMUP group, indicating that the osteogenic effect of DMUP was better than treatment with MC.

Fig. 9

In vivo assessment of anti-inflammatory osteogenic effect in animals. (a) Schematic diagram of periodontitis modeling on the maxillary left side of SD rats. Created with BioRender. (b) H&E and Masson staining of maxillary left side tissues of different groups after treatment, and (c) quantification of the number of inflammatory cell infiltration and (d) collagen content of periodontal tissues. (e) Representative images of tissue immunohistochemistry for positive expression of IL-6 and IL-10, and (f, g) quantitative results of positive expression. (h) Representative images of Micro-CT of maxillary left jaw after treatment in different groups, (i) Tb.Th parameters and (j) BMD values. (k) Immunofluorescence staining images of osteogenesis-related proteins OCN and Runx2 in tissue and (l, m) quantification results. Data are means ± s.d. (n = 6), ns: no significant difference, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

Conclusion

In summary, we have developed a responsive NIR-IIb multifunctional nanozyme platform for treating periodontitis. This platform has bacterial imaging, anti-anaerobic, anti-inflammatory, and bone-forming functions. It accurately captures the diseased area flora in complex environments by responding to the inflammatory microenvironment. Additionally, the platform releases Mn nanozyme in response to acid, which catalyzes the production of O₂ from endogenous H₂O₂. This process works with Pae to regulate macrophage function and reduces the inflammatory factor content, resulting in improved periodontitis treatment outcomes. To further demonstrate the effectiveness of imaging infected areas in larger tissues, we verified the performance of DMUP, not only in an animal periodontitis model, but also in imaging bacterially infected areas in the legs of rats. This paves the way for potential application in treating larger tissue volumes. However, the system has limitations. First, the complex synthesis methods of materials pose a challenge for future large-scale production and in vivo metabolism. Second, in clinical applications, larger infected areas require treatment with scaling. Additionally, considering the complex flora in the oral environment, the effectiveness of combined treatment still needs further verification. Nevertheless, the combination of bacterial imaging and nanozyme for periodontitis is extremely significant and can be extended to the localization and treatment of other bacterial infectious diseases, such as various types of implants, bone plates, artificial joints, and vascular scaffolds.