Dexamethasone Long-Term Controlled Release from Injectable Dual-Network Hydrogels with Porous Microspheres Immunomodulation Promotes Bone Regeneration

Abstract

Long-lasting, controlled-release, and minimally invasive injectable platforms that provide a stable blood concentration to promote bone regeneration are less well developed. Using hexagonal mesoporous silica (HMS) loaded with dexamethasone (DEX) and poly(lactic-co-glycolic acid) (PLGA), we prepared porous DEX/HMS/PLGA microspheres (PDHP). In contrast to HMS/PLGA microspheres (HP), porous HMS/PLGA microspheres (PHP), DEX/PLGA microspheres (DP), and DEX/HMS/PLGA microspheres (DHP), PDHP showed notable immuno-coordinated osteogenic capabilities and were best at promoting bone mesenchymal stem cell proliferation and osteogenic differentiation. PDHP were combined with methacrylated silk (SilMA) and sodium alginate (SA) to form an injectable photocurable dual-network hydrogel platform that could continuously release the drug for more than 4 months. By adjusting the content of the microspheres in the hydrogel, a zero-order release hydrogel platform was obtained in vitro for 48 days. When the microsphere content was 1%, the hydrogel platform exhibited the best biocompatibility and osteogenic effects. The expression levels of the osteogenic gene alkaline phosphatases, BMP-2 and OPN were 10 to 15 times higher in the 1% group than in the 0% group, respectively. In addition, the 1% microsphere hydrogel strongly stimulated macrophage polarization to the M2 phenotype, establishing an immunological milieu that supports bone regrowth. The aforementioned outcomes were also observed in vivo. The most successful method for correcting cranial bone abnormalities in SD rats was to use a hydrogel called SilMA/SA containing 1% drug-loaded porous microspheres (PDHP/SS). The angiogenic and osteogenic effects of this treatment were also noticeably greater in the PDHP/SS group than in the control and blank groups. In addition, PDHP/SS polarized M2 macrophages and suppressed M1 macrophages in vivo, which reduced the local immune–inflammatory response, promoted angiogenesis, and cooperatively aided in situ bone healing. This work highlights the potential application of an advanced hydrogel platform for long-term, on-demand, controlled release for bone tissue engineering.

1. Introduction

With more than 10 million patients suffering from bone defects and bone injuries worldwide each year, a large demand exists for bone repair products. Autologous and allogeneic bone grafts are the gold standard for repairing bone defects exceeding critical dimensions. However, the obvious drawbacks of these methods, such as donor site pain, bleeding, and limited sources, limit their clinical application. Artificial materials have become increasingly popular in bone repair, and bone graft materials synthesized from polymers, hydrogels, ceramics, metals, and other materials continue to be developed, contributing to advances in the field of bone repair. Unfortunately, although significant progress has been achieved, these bone tissue engineering materials still have limitations. For example, shaping materials for repairing irregular bone defects, secondary trauma associated with implantation procedures, abnormal inflammatory responses to bone regeneration, and bone resorption in defects under pathological conditions (osteoporosis, bone tumors, etc.) are all issues that need to be considered when constructing graft materials. The healing of bone defects is divided into three main phases, inflammation, repair, and remodeling, and the whole process usually takes several months or even years.¹

Interest in platforms that enable the sustained and effective controlled release of drugs that promote bone regeneration has increased; these platforms can deliver drugs stably to specific sites and then sustain a steady release to promote bone healing. Currently, the majority of drugs used in the clinic are delivered via an immediate or primary release mechanism. Zero-stage delivery (controlled release) systems release the drug at a constant rate throughout the dosing cycle and extend the drug’s circulation lifetime, maintaining the drug concentration for a longer period.² Compared with slow-release technology, controlled release (zero-stage release) technology allows the carrier material to release the drug continuously for a specific period, providing a stable blood concentration for clinical treatment. Zero-stage release of drugs can reduce adverse side effects, reduce the frequency of drug administration, and improve patient compliance, overcoming the shortcomings of traditional drug delivery methods.³ Unfortunately, fewer scaffold-based drug delivery systems have been reported that can simultaneously achieve drug release for at least 2 months and promote bone regeneration.⁴ In addition, currently available controlled release materials mainly include inorganic materials such as mesoporous silica, gold nanoparticles, graphene, and bio glass and polymeric materials such as collagen, PCL, and PLGA. These materials can be loaded with drugs and delivered to the target site to release the drug upon chemical or physical stimulation. However, most of the current carriers have a short drug release cycle, usually approximately 2 weeks to 1 month, and only a small number of carrier materials can achieve a zero-grade controlled release effect, making it difficult to achieve controlled drug release throughout the entire bone defect healing cycle.⁵⁻¹⁰

In addition, the development of a bone graft repair material that can be injected in a minimally invasive manner to accommodate irregularly shaped defects and that can demonstrate a durable controlled-release effect that promotes bone regeneration is urgently needed. In the injectable drug delivery platform, degradable polyester microspheres, which can deliver drug molecules to the target via injection, inhalation, and other routes and delay the drug action time, have good biocompatibility and degradation properties; however, the short drug release cycle of polyester microspheres alone cannot meet the long-term therapeutic demand.¹¹ Previous studies have shown that inorganic drug-carrying materials, such as hexagonal mesoporous silica (HMS), have a large burst release and a short release duration, whereas HMS-modified poly(lactic-co-glycolic acid) copolymer (PLGA) microspheres can significantly prolong the duration of drug release; however, the release process is divided into three phases—burst, delayed, and plateau—and controlled release (zero-stage release) cannot be achieved.¹² Hydrogels are hydrophilic cross-linked polymers that absorb water and swell to form a 3D network and have been widely studied for their excellent biocompatibility. The injectable and in situ cross-linking properties of hydrogels make minimally invasive surgery possible, avoiding the secondary trauma of open surgery. Injectable hydrogels can support the body’s innate ability to heal by providing a temporary matrix for internal host cell growth and new blood vessel formation. In addition, injectable delivery ensures that the hydrogel remains within the defect area and adapts to the shape of the defect, even in irregularly shaped bone defects.¹³ For example, Xiaohu Zhou et al. prepared an nHA-loaded gelatin/arginine hydrogel as an injectable hydrogel with the potential to accommodate and cover irregular maxillofacial bone defects.¹⁴ Sodium alginate (SA) is a natural polymer that is biocompatible, easily cross-links in Ca²⁺-rich solutions to form hydrogels, has a certain degree of plasticity, is simple and inexpensive to prepare, and has been widely used in research on bone fillers.¹⁵ However, SA lacks cell adhesion sites, and its high-water absorption leads to rapid swelling and degradation of the hydrogel, which can be compensated by MA-modified filipin protein (SilMA). Compared with single-network hydrogels, dual-network hydrogels have greater mechanical strength and bioactivity and thus have attracted much attention.^16,17 However, hydrogels typically have shorter drug release cycles and greater burst release. The integration of microspheres into tissue-engineered scaffolds can provide the required level of controlled release of payloads.¹⁸ In previous studies of drug-loaded degradable polyester microspheres, most of the microspheres were compounded with hydrogel systems or scaffolding materials (e.g., gelatin or methylcellulose) to enable studies of their in vivo osteogenic effects and their application in bone repair.¹⁹ Compounding drug-carrying microspheres with dual-network hydrogels can further modulate the drug release behavior of the hydrogels; in addition, modulating the pore structure of the microspheres affects the drug release behavior.^20,21

Dexamethasone (DEX), a synthetic corticosteroid widely used in clinical practice, is effective at inducing the growth, proliferation, and osteogenic differentiation of mesenchymal stem cells and has been used in studies of bone tissue engineering scaffolds.²² When the DEX concentration is 10–100 nM, a higher concentration is more favorable for promoting osteogenesis.²³ Adverse effects on bone healing occur when the DEX concentration exceeds 1000 nM.²⁴ Dose-dependent side effects and complications (e.g., adrenocortical necrosis, diarrhea, and wasting) associated with long-term dexamethasone use are unacceptable.²⁵ Therefore, unique design ideas should be introduced in the construction of dexamethasone formulations so that the drug can effectively reach the lesion site to exert anti-inflammatory and bone-enhancing effects and reduce the adverse effects caused by systemic absorption. For example, Heo et al. prepared click-cross-linked HA hydrogels encapsulating DEX/PLGA microspheres, and the composite gels showed much slower cumulative release of Dex early in vivo and ex vivo, followed by sustained release for 28 days, than microspheres alone.²⁶ Neelam et al. developed injectable pullulan–poly(ethylene glycol) hydrogels with covalently attached Dex and showed that these hydrogels could sustain the release of PEG-Dex conjugates for 28 days under pH-sensitive conditions, and 74.54 and 55.15% of the PEG-Dex conjugates were released at pH 6.5 and 7.4, respectively. The release of the PEG-Dex conjugate from the hydrogel increased cell viability and proliferation and induced osteoblast differentiation.²⁷ Chen et al. prepared biphasic calcium phosphate nanoparticle/collagen composites loaded with DEX with different microgrooves and showed that microgrooves in the composite folds promote angiogenesis and stimulate new bone formation.²⁸ Aikaterini-Rafailia Tsiapla et al. loaded DEX on polymeric cellulose acetate and detected drug release for up to 181 days.⁵ However, the drug profile of this system was close to primary release and did not monitor the drug concentration, which is crucial for immunomodulation of osteogenic effects. Less injectable bone repair materials that can release DEX for more than two months and achieve controlled release behavior over a longer period have been reported.

In addition to the long-term zero-grade release of the drug and the injectable nature of the delivery platform, the excessive immune–inflammatory response that occurs during bone defect regeneration is another therapeutic challenge. Although inflammation is a mechanism for the early initiation of healing, timely and effective elimination of inflammation is necessary to create a favorable environment for bone regeneration.²⁹ Macrophages play an important role in immunomodulation, mainly through polarization into M1 and M2 phenotypes. M1 macrophages are pro-inflammatory cells that recruit inflammatory factors to activate the immune response against pathogen invasion in the early stages of inflammation, but excessive inflammation leads to tissue fibrosis and delayed healing. In contrast, M2 macrophages are anti-inflammatory cells with potent anti-inflammatory effects and are primarily involved in the clearance of inflammatory factors, tissue healing and remodeling, and immunomodulation.³⁰ Therefore, biomaterials must be able to modulate macrophage polarization from the M1 to the M2 phenotype. Similarly, the environment surrounding a bone defect affects bone repair. In pathological bone defects (e.g., those associated with osteoporosis and tumors), osteoclasts are abnormally active and adhere to bone tissue, promoting bone resorption and affecting bone regeneration.³¹ DEX has potent anti-inflammatory effects while promoting osteoinductive osteogenesis. Li et al. reported that DEX inhibited the production of M1-type macrophages and suppressed the secretion of interleukin-6 (IL-6) and inducible nitric oxide synthases (iNOs), which resulted in the inhibition of osteoclast activity and the process of bone resorption.³²

Based on the above findings, we prepared HMS using the gel–sol method and then added DEX to the pores of HMS. Then, porous DEX/HMS/PLGA microspheres (PDHP) were prepared and compounded with SilMA and SA to construct an injectable dual-network hydrogel platform for light curing and drug delivery (PDHP/SS). We obtained a hydrogel platform with long-term controlled release that promoted a stable blood drug concentration by adjusting the content of the porous drug-carrying microspheres in the hydrogel, which induced the formation of an immune microenvironment enriched with M2-type macrophages and promoted the osteogenic differentiation of bone mesenchymal stem cells (BMSCs) for bone regeneration and repair (Figure 1A). Finally, we documented the osteogenic and angiogenic effects of the composite hydrogel as well as its ability to achieve immunomodulation through macrophage polarization through in vivo experiments (Figure 1B). The prolonged and controlled release of PDHP/SS induced macrophage M2 polarization in vivo and inhibited macrophage M1 polarization, which alleviated the localized immune-inflammatory response, promoted angiogenesis, and synergistically facilitated in situ bone regeneration. In conclusion, our study provides an injectable and in situ-cured dual-network hydrogel platform for the delivery of DEX-loaded porous microspheres; this hydrogel platform can effectively coordinate bone and angiogenesis and inhibit inflammatory responses through the long-term zero-sequence release of DEX, which provides a new idea for the development of bone graft materials.

Figure 1.

Schematic diagram of the synthesis process, in vitro cell proliferation, and osteogenic induction. (A) By preparation of HMS and DEX/HMS/PLGA microspheres (DHP), the porous DEX/HMS/PLGA microspheres (PDHP) that can best promote the proliferation and osteogenic differentiation of BMSCs were selected. SilMA was prepared and combined with SA and PDHP to obtain an injectable photocured drug-carrying hydrogel platform (PDHP/SS). The optimal concentration of porous drug-loaded microspheres in the hydrogel platform was optimized by evaluating the proliferation and osteogenic differentiation of BMSCs, drug release performance, mechanical strength, and degradation performance in vitro. (B) PDHP/SS were most effective at facilitating cranial defect repair in SD rats, and compared with those in the control and blank groups, the PDHP/SS group showed significantly increased vascularization and osteogenesis and alleviated the local immune–inflammatory response in situ. HP, PHP, DP, DHP, and PDHP are abbreviations for HMS/PLGA microspheres, porous HMS/PLGA microspheres, DEX/PLGA microspheres, DEX/HMS/PLGA microspheres, and porous DEX/HMS/PLGA microspheres, respectively.

2. Materials and Methods

2.1. Materials

Ethyl alcohol (EtOH; Chemical Reagent Factory), tetraethoxysilane (TEOS; Chemical Reagent Factory), dichloromethane (DCM; Chemical Reagent Factory), poly(vinyl alcohol) (PVA; Sigma-Aldrich), dodecyl amine (DDA; Sigma-Aldrich), poly(lactic-co-glycolic acid) (PLGA; M_w = 31,000 Da; ratio of lactic to glycolic acid: 50:50; Daigang Biomaterials), sodium alginate (SA; Sigma-Aldrich), high-glucose Dulbecco’s modified Eagle’s medium (DMEM; Gibco), phosphate-buffered saline (PBS; Gibco), fetal bovine serum (FBS; 10270-106, Gibco), Cell Counting Kit-8 (CCK-8; Dojindo Laboratories), and Alizarin Red S (ARS; Sigma-Aldrich) were used. Rat BMSCs and RAW264.7 cells (macrophage line) were purchased from American Type Culture Co. (ATCC, Manassas, VA).

2.2. Preparation of Hydrogels Containing Drug-Loaded Porous Microspheres

2.2.1. Preparation of HMS

HMS was synthesized via a modified version of a previously reported process.³³ Briefly, a templating solution was prepared by dissolving 12 g of DDA in 70 mL of EtOH and 70 mL of distilled water and stirring for 1 h. A second solution was prepared by mixing 44.6 mL of hot TEOS (70 °C) and 40 mL of EtOH and stirring for approximately 30 min. The two solutions were then mixed by stirring for an additional 18 h. The mixture was then aged for 30 min, washed with distilled water, and extracted with HCl–EtOH, followed by drying and calcination.

2.2.2. Preparation of Porous Drug-Loaded HMS/PLGA Microspheres

DEX-HMS particles were prepared by adding 0.6 g of HMS particles to 20 mL of deionized water containing 0.06 g of DEX. The solution was constantly stirred for 24 h. The sedimented phase was isolated, washed with deionized water three times, and then dried. The obtained DEX-HMS particles were sieved to obtain particles smaller than 50 μm in diameter for further use.

Porous drug-loaded HMS/PLGA microspheres (PDHP) were synthesized based on a standard protocol.³⁴ Briefly, 3 g of PLGA was dissolved in 15 mL of methylene chloride with 0.15 g of DEX-HMS particles and 1 mL of deionized water. The emulsion mixture was stirred for 3 h, poured into 1000 mL of an aqueous 1% PVA solution, and stirred at 300 rpm for 4 h. The resultant PDHP were isolated, washed with deionized water, and air-dried for 24 h. The DEX/PLGA microspheres (DP), HMS/PLGA microspheres (HP), DEX/HMS/PLGA microspheres (DHP), and porous HMS/PLGA microspheres (PHP) were prepared, as described above, but 25 mg of DEX was used instead of 50 mg of DEX-HMS, and no water was added to the DP; 50 mg of HMS was used instead of 50 mg of DEX-HMS, and no water was added to the HP; no water was added to the DHP; and 50 mg of HMS was used instead of 50 mg of DEX-HMS for the PHP. All the microspheres were sieved to a diameter less than 150 μm for further use.

2.2.3. Preparation of SilMA/SA Dual-Network Hydrogels Containing DEX-Loaded Porous Microspheres

The silkworm chrysalis was removed, weighed to obtain a 60 g sample, boiled in 6 L of a 0.5% Na₂CO₃ solution for 45 min, and then washed with deionized water. The above steps were repeated three times until the silk fibroin became white, after which the sample was dried in an incubator. The dried silk fibroin protein was dissolved in a 9 mol/L LiBr solution at a concentration of 10 g/L and then incubated for 1 h at 40 °C. After the protein was fully dissolved, it was placed in a dialysis bag for 3 days, and the water was changed twice a day, after which the mixture was lyophilized to obtain the silk fibroin protein. After the boiled silk fibroin protein fiber was dissolved in LiBr, a 424 mM glycidyl methacrylate solution was added to the mixture, and the reaction proceeded at 300 rpm and 60 °C for 3 h. After filtration, dialysis bags with a molecular weight of 12–14 kDa were used for 4 days of dialysis, and the deionized water was replaced every 4 h. The bags were then placed at −80 °C overnight, freeze-dried, and stored at 4 °C until use.

To prepare 12.5 mL of 0.25% lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP) in PBS, 0.8 g of SilMA, and 0.25 g of SA were dissolved in the above solution, and 0, 31.25, 62.5, 125, and 250 mg of PDHP were added to the solution and dispersed evenly. Blue light was applied for 30 s, and a 0.5 mol/L calcium chloride solution was added for 2 min. Microsphere-containing hydrogels with mass-to-volume ratios of 0, 0.25, 0.5, 1.0, and 2.0% were obtained and are referred to as 0, 0.25, 0.5, 1.0, and 2.0%, respectively.

2.3. Characterization of Materials

2.3.1. Characterization of DEX-HMS

The morphologies of HMS and DEX-HMS were characterized using HRTEM (JEM2011) with an accelerating voltage of 50–200 kV, a point resolution of 0.23 nm, and a lattice resolution of 0.14 nm. X-ray diffraction was performed using a Rigaku D/max-2200 (Japan, Cu Ka, g = 1.54056 nm) instrument with a voltage of 40 kV and a current of 30 mA. The diffraction pattern was collected over the 2θ range of 1–10° with a scanning speed of 1°/min and a step size of 0.02°. N₂ adsorption–desorption isotherms were obtained with a Micromeritics Tristar 3000 pore analyzer under continuous adsorption conditions. Initially, the powder sample was placed over the sample bulb, heated at a temperature of 80 °C under a 0.1 MPa vacuum, and incubated overnight to remove moisture. The specific surface area was estimated from the nitrogen sorption data over a relative pressure P/P₀ range from 0.0 to 1.0. The Brunauer–Emmett–Teller (BET) and Barrett–Joyner–Halenda (BJH) methods were used to determine the surface area, pore size distribution, and pore volume. The DEX encapsulation efficiency of the DEX-HMS particles was determined by suspending 50 mg of DEX-HMS in 5 mL of PBS (pH 7.4) and incubating the suspension at 37 °C, with a periodic analysis of the supernatant.

2.3.2. Characterization of Microspheres and Hydrogels Containing Microspheres

Infrared spectra of silk fibroin and SilMA were obtained with KBr disks in the range 4000–400 cm^–1. The ¹H NMR spectra of silk fibroin and SilMA were examined by using a Bruker DPX FT-NMR spectrometer (9.4T, Bruker Analytik GmbH, Karlsruhe, Germany) at a frequency of 400 MHz to determine the degree of methacrylation. Then, 700 μL of deuterium oxide (D₂O; Sigma-Aldrich) was used as a solvent for every 5 mg of sample. The SilMA solution was filtered through a 0.45 μm filter before analysis.

1 ml of gel was collected with a 1 mL syringe and injected into a clear glass bottle. Blue light was applied for 30 s, after which the sample bottle was immediately placed upside down on the test bench. At room temperature, the gel flow in the inverted sample bottle was observed for each group at 0 min, 0.5 h, and 72 h. The gel material was collected with a 1 mL syringe, the gel was injected with the needle removed, and the injection of the gel was observed.

The surface morphology of the microspheres and the surface and cross-sectional morphology of the SilMA/SA hydrogels and SilMA/SA hydrogels containing DEX-loaded porous microspheres were observed by using scanning electron microscopy (SEM; Q25, FEI).

The scaffold compression test was conducted with a universal testing machine (INSTRON 5967) at a speed of 1 mm/min at room temperature. Six samples were prepared for each group. The prepared samples were 10 mm in diameter and 8 mm in height. On the stress–deformation curve, the compressive strength of the hydrogel was recorded as the pressure when the compressive deformation reached 90%. The compression modulus of the hydrogel was calculated with a linear deformation range of 0.5 to 20%.

The swelling ability of the hydrogels was evaluated by weighing the freeze-dried scaffold, and the initial mass was recorded as W_a. Then, the scaffold was swelled in a 37 °C PBS solution for 24 h, the liquid on the surface of the scaffold was carefully removed with filter paper, and the weight was recorded as W_b. Each group had 6 parallel samples. The hydration degree of the hydrogels was obtained using the following formula

1

The PBS degradation method was used to evaluate the degradation of the hydrogels. The freeze-dried hydrogels were weighed, and the initial mass was recorded as W_d. The hydrogels were then immersed in a PBS solution and placed in a shaker at 37 °C and 100 rpm. The scaffolds were removed at specific time points (7, 14, 21, and 28 days), washed with deionized water and anhydrous ethanol, lyophilized, and weighed, and the mass was recorded as W_f. Each group included 4 parallel samples. The remaining mass of the support was calculated with the following formula

2

DHP and PDHP (0.2 g) were suspended in methylene chloride (30 mL) to sediment the insoluble DEX-HMS particles. The sedimented particles and all supernatants were collected. The amount of drug encapsulated in the drug-loaded particles (C_DEX-HMS) was measured. The amount of drug encapsulated in the collected supernatant, which was calculated as the amount of DEX encapsulated in the PLGA matrix (C_PLGA), was also measured. Therefore, the total drug encapsulation efficiency of the drug-loaded HMS/PLGA microspheres was calculated using the formula (C_DEX-HMS + C_PLGA)/C_TID, in which C_TID represents the total initial dosage of DEX. DP (0.2 g) were dissolved in 3 mL of methylene chloride. After dissolution, 27 mL of PBS was added. The encapsulation efficiency of DEX was determined by measuring the amount of DEX in the supernatant after two centrifugation steps by using a UV spectrophotometer.

In vitro DEX release trials were performed in a shaking incubator at 60 rpm and 37 °C. 50 mg of microspheres or hydrogels containing 50 mg of microspheres were soaked in 20 mL of PBS (pH 7.4). Sample media were collected at regular time intervals with equal volumes of PBS.

2.4. Cell Experiments

2.4.1. Cell Culture

All cell experiments were performed in 24-well plates with 1.5 mL of medium added to each well. BMSCs or RAW264.7 cells were cultured in DMEM supplemented with 10% FBS and 1% penicillin/streptomycin. All the cells were maintained at 37 °C in a humid atmosphere with 5% CO₂. Cell subculturing was performed after the cells reached 75–85% confluence. For all of the experiments in this study, only early passage BMSCs (passages 3–7) were used. Before cell culture, all of the samples were subjected to γ irradiation at a dose of 10 kGy using a 60 Co source. All of the samples were incubated in cell culture medium for 2 h before use to improve cell attachment. In the microsphere-containing experiments, each group was treated with 2 mg of microspheres per well. In the microsphere-containing hydrogel experiments, the volume of microsphere-containing hydrogel per well was the same for all groups, with the 0, 0.25, 0.5, 1.0, and 2.0% groups containing 0, 1.25, 2.5, 5, and 10 mg of PDHP, respectively.

2.4.2. Cell Proliferation

Briefly, sterilized samples were placed into 24-well plates, and 5 × 10⁴ BMSCs were seeded on the samples. The proliferation of BMSCs was measured at 1, 3, and 7 days using a Cell Counting Kit 8 (CCK8) assay. At designated times, the cells were washed twice with PBS, and CCK8 assay solution [10% (v/v)] in growth medium was added to each plate and incubated for 2 h. The solution was transferred to a 96-well plate at 100 μL per well, and the absorbance (OD value) was measured at 450 nm. The difference in the OD of the samples between groups was the difference in the cell proliferation activity of the samples.

2.4.3. Polarization of Macrophages

RAW264.7 cells were inoculated into 48-well plates containing microspheres or hydrogels at a density of 5 × 10⁴ cells per well. The medium was changed every 2–3 days (the medium for each group of samples was collected separately and mixed with osteogenic induction solution at a ratio of 1:5 to prepare macrophage-conditioned medium for spare use). On days 1 and 3, real-time fluorescence quantitative PCR was used to detect the expression of the M1 genes tumor necrosis factor-alpha (TNF-α) and interleukin-1 beta (IL-1β) and the M2 genes CD206 and arginase (ARG). Glyceraldehyde phosphate dehydrogenase (GAPDH) was used as an internal reference gene.

2.4.4. Osteogenic Differentiation of BMSCs

BMSCs were plated in hydrogels solidified in 24-well plates at a seeding density of 10 × 10⁴ cells/well. After 24 h of adhesion, the medium was replaced with osteogenesis induction medium containing 50 ng/mL of ascorbic acid (Sigma-Aldrich), 100 nM DEX, and 10 mM beta-glycerophosphate (Sigma-Aldrich). At predetermined time points, qRT–PCR was used to measure the expression of the typical osteogenesis-related genes, bone morphogenetic protein 2 (BMP2), alkaline phosphatase (ALP), osteopontin (OPN), and runt-related transcription factor 2 (Runx2).

Briefly, total RNA was isolated using an RNA isolation kit (CW0581M) according to the manufacturer’s protocol. After RNA isolation, DNase I treatment was performed, and the total RNA concentration and purity were measured at 260 nm by using a Nanodrop spectrophotometer (Thermo). Reverse transcription (RT) of RNA was performed to synthesize cDNA using an HIFI cDNA synthesis kit (CW2569M). For RT, cDNA was synthesized from 1 mg of total RNA using an HIFI cDNA synthesis kit (CW2569M). Quantitative reverse transcriptase–polymerase chain reaction (qRT–PCR) amplification was performed with an Applied Biosystems QuantStudio 6 Flex Real-Time PCR (Thermo) using an UltraSYBR Mixture (Low ROX) (CW2601M). The sequences of the forward primers and reverse primers used are listed in Table 1. Relative mRNA levels were calculated and normalized to those of GAPDH. All six qRT–PCR assays were performed.

Table 1. Sequences of the Primers Used for the Real-Time Polymerase Chain Reaction (PCR).

gene transcript	forward primer sequence (5′–3′)	reverse primer sequence (5′–3′)
glyceraldehyde phosphate dehydrogenase (GAPDH)	GCCATGAGGTCCACCACCCT	AAGGTCATCCCAGAGCTG
alkaline phosphatase (ALP)	GGAGATGGTATGGGCGTCTC	GGACCTGAGCGTTGGTGTTA
runt-related transcription factor 2 (RUNX2)	TCGGAGAGGTACCAGATGGG	AGGTGAAACTCTTGCCTCGT
osteopontin (OPN)	TGAAACTCGTGGCTCTGATG	GATGAACCAAGCGTGGAAAC
collagen type I (COLI)	TTCTCCTGGCAAAGACGGAC	CTCAAGGTCACGGTCACGAA
bone morphogenetic protein-2 (BMP-2)	ACCCGCTGTCTTCTAGTGTTG	TTCTTCGTGATGGAAGCTGAG
CD206	ATGGATGTTGATGGCTACTGG	TTCTGACTCTGGACACTTGC
arginase (ARG)	CATATCTGCCAAAGACATCG	GGTCTCTTCCATCACCTTGC
tumour necrosis factor-α (TNF-α)	GGGTGTTCATCCATTCTC	GGTCACTGTCCCAGCAT
interleukin-1 beta (IL-1β)	TACAGGCTCCGAGATGAACA	AGGCCACAGGTATTTTGTCG

A total of 5 × 10⁴ BMSCs were added to 48-well plates containing the samples and incubated for 24 h. The supernatant was aspirated and discarded, osteogenic induction medium or macrophage-conditioned medium was added, and the culture medium was changed periodically. Alizarin Red S (0.5%, pH 4.2; Sigma-Aldrich) was used to assess matrix mineralization. ALP activity was used as a measure of bone regeneration. ALP staining was performed at the specified time points using an ALP staining kit; ALP enzyme activity was quantified using the BCA method; and calcium nodules were stained using Alizarin Red S (0.5%, pH 4.2; Sigma-Aldrich).

2.5. Animal Experiments

2.5.1. In Vivo Implantation

All animal experiments were performed under a protocol approved by the Institutional Animal Care and Use Committee of the Guangdong Quality Supervision and Testing Station for Medical and Health Care Appliances. 24 eight week old male Sprague-Dawley (SD) rats (Guangdong Quality Supervision and Testing Station for Medical and Health Care Appliances, Guangzhou, China) were used in this study and randomly divided into 4 groups: (1) the blank control group (blank, n = 6); (2) the SilMA/SA hydrogel group (SS, n = 6); (3) the SilMA/SA hydrogel group containing 1% PHP (PHP/SS, n = 6); and (4) the SilMA/SA hydrogel group containing 1% PDHP porous DEX-HMS/PLGA microspheres (PDHP/SS, n = 6). SD rats were anaesthetized using a mixture of 10% chloral hydrate and 25% urethane. A rat calvarial bone defect model was established on both sides of the cranium by drilling a 5 mm diameter hole with a Zoletil50 (10 mg kg^–1). Briefly, routine skin preparation, disinfection, and towel placement were performed at the site of the defect. A lateral longitudinal incision of approximately 1.5 cm in length was made in the skull. The skin and periosteum were cut layer by layer, and a cylindrical defect (diameter = 5 mm) was drilled on both sides of the cranium herringbone line using a medium-speed grinding drill. The defect was formed by rotation with hemostatic forceps, and the deep residual bone mass was removed with a small curet. The hydrogels (50 μL) were injected into the defects, and the blank groups were left untreated. Blue light was applied for 30 s, and a 0.5 mol/L calcium chloride solution was added for 2 min. The wounds were sutured, and a prophylactic antibiotic was administered to avoid infections.

2.5.2. Micro-CT Analysis

At the end of the study period (4- and 8 weeks postsurgery), the rats were euthanized, and the calvarial bone was harvested from each rat. After fixation with 4% paraformaldehyde for 48 h, a Micro CT instrument (ZKKS-MCT-Sharp, Guangzhou Zhongke Kaisheng Medical Technology Co. Ltd.) was used to scan the cranium samples and quantitatively analyze the newly formed bone within the defects. The sample scanning conditions were set as follows: a scanning voltage of 70 kV, a power of 7 W, 4-frame superposition, an angle gain of 0.72°, an exposure time of 100 ms, and one rotation to complete the scan.

2.5.3. Histological Analysis

After fixation with 4% paraformaldehyde for 48 h, the cranium samples were decalcified in a 10% EDTA demineralizing solution for one month. Then, a 5 μm thick section was cut from the paraffin-embedded tissue for histological evaluation. Haematoxylin and eosin (H&E) staining and Masson’s trichrome staining were performed on sections from each sample to identify new bone formation. For immunofluorescence histochemical staining, the sections were immersed in 3% (w/v) H₂O₂ and blocked with a 3% (v/v) BSA solution. Following enzymatic antigen retrieval, the sections were incubated with primary antibodies against OCN (rat, 1:100 dilution; Abcam, USA), COLI (rat, 1:2000 dilution; Abcam, USA), VEGF (rat, 1:200 dilution; Abcam, USA), CD31 (rat, 1:2000 dilution; Abcam, USA), CD163 (rat, 1:100 dilution; Abcam, USA), or inducible nitric oxide synthase (rat, 1:2000 dilution; iNOS, Abcam, USA). After two rinses with PBS, the samples were incubated with the corresponding horseradish peroxidase-labeled secondary antibodies, followed by 4′,6-diamidino-2-phenylindole (DAPI) for visualization. Nuclei were counterstained with hematoxylin.

2.6. Statistical Analysis

The experiments were repeated six times, and the results are presented as the means ± standard deviations. All the statistical analyses were performed using SPSS software. A one-way analysis of variance followed by Dunnett’s multiple comparisons test was used to evaluate the significance of differences between experimental groups. p values < 0.05, 0.01, and 0.001 were considered to indicate statistical significance.

3. Results

3.1. Characterization, Cell Proliferation, and ALP Activity of PDHP Microspheres and Preparation of SilMA

We first prepared HMS with a high specific surface area by using the sol–gel method and then loaded DEX into the pores of the HMS to prepare a long-acting, controlled-release hydrogel platform that can induce bone formation. Using DEX/HMS, PLGA, and pore-forming agents, porous drug-carrying microspheres were prepared by using the emulsifying solvent volatilization method. The porous structure of the microspheres and the effect of drug loading on the proliferation and osteogenic differentiation of BMSCs were studied, and the microspheres were selected for further experiments. Moreover, we modified the extracted silk fibroin protein to determine whether SilMA was successfully obtained.

As shown in the TEM images (Figure 2A), the pore structure of the DEX/HMS system was less clear than that of the HMS system. The N₂ adsorption–desorption isotherms, pore size distributions, BET surface areas, and pore volumes of HMS and DEX-HMS are shown in Figure 2B,C. The N₂ adsorption–desorption isotherms of HMS suggest that the pores resemble type IV isotherms. After the drug loading process, nearly complete pore blockage was observed; the specific surface area of HMS decreased from 1173.47 to 627.98 m²/g, the mean pore size of HMS increased from 2.23 to 3.42 nm, the pore volume of HMS decreased from 0.11 to 0.04 m³/g, and the pore shape resembled a type IV isotherm. A pore volume of 0.04 m³/g indicated that DEX had adsorbed into the HMS pores. As shown in Figure 2D, HMS and DEX-HMS present well-defined XRD patterns, and the d (100) reflections of HMS and DEX-HMS are not consistent, although the d (100) reflections of the mesoporous materials of the typical HMS system can be identified. The peak of the d (100) reflections for DEX-HMS is lower than that of HMS prior to drug loading. Considering the BET results, this difference might have occurred because some of the mesopores of DEX-HMS collapsed. Indeed, the effective adsorption of DEX into the HMS pore system was evidenced by pore blockage, a greater mean pore size, a smaller surface area, and a smaller pore volume, consistent with the BET results. The loading ratio of DEX-HMS was 99.5%, and the above results indicated that DEX was successfully loaded into the HMS.

Figure 2.

TEM images (A), N₂ adsorption–desorption isotherms (B), pore size distributions (C), and XRD patterns (D) of HMS and DEX-HMS. SEM images showing the surface morphology of HMS/PLGA microspheres (HP), porous HMS/PLGA microspheres (PHP), porous DEX-HMS/PLGA microspheres (PDHP), DEX/PLGA microspheres (DP), and DEX-HMS/PLGA microspheres (PHP) (E). CCK-8 assay (F) indicating the viability of BMSCs after treatment for 1, 3, and 7 days. ALP activity (G) showing the osteogenic potential of BMSCs after treatment for 7 days. FTIR spectra and H NMR spectra of SF and SilMA (H, I). *p < 0.05, **p < 0.01, and ***p < 0.001.

SEM images of HP, PHP, PDHP, DP, and DHP are shown in Figure 2E. All of these microspheres were prepared using a single emulsion solvent evaporation technique and showed good sphericity with a particle size less than 150 μm. A porous structure was observed on the surface of both the PHP and PDHP. The cytocompatibility of the microspheres was analyzed using a CCK-8 assay after 1, 3, and 7 days of coculture. As shown in Figure 2F, an increase in absorbance from day 1 to day 7 was recorded, which indicated that the cells were viable in all of the samples. The viability of the BMSCs cultured on PDHP was greater than that of the cells cultured on the other four media in all the groups, which suggested that the PDHP had the greatest cytocompatibility. In the comparison of the pro-proliferative activity of BMSCs in the remaining groups, PHP enriched with a pore structure was superior to HP, while DHP, drug-loaded microspheres with added HMS, were significantly more effective than DP. Specifically, the cell proliferative activity of the PHP group was superior to that of the DP group but not significantly different from that of the DHP group. The ability of the microspheres to induce osteogenic differentiation in the BMSCs was determined primarily on the basis of the expression of the marker protein ALP. The secreted ALP content was analyzed quantitatively after the cells were cultured for 7 days, and the PDHP group exhibited the highest ALP activity (Figure 2G). The ALP-promoting properties were also significantly different between the remaining groups, with DHP > PHP > DP > HP. These results indicate that HMS/PLGA microspheres with porous structures loaded with DEX can better promote the proliferation and osteogenic differentiation of BMSCs.

As shown in Figure 2H, in the FTIR spectrum of SilMA, amide I (1640 cm^–1), amide II (1511 cm^–1), and amide III (1234 cm^–1) are the three characteristic amide peaks of SF. RR’C=CH₂ is the characteristic peak at 951 cm^–1, and the obvious increase in the size of this peak indicates an obvious increase in the amount of double bonds and successful grafting of the methyl acrylamide group. ¹H NMR spectroscopy was performed on SF and SilMA to further verify the access of the target groups (Figure 2I). The increase in the peaks associated with the methacrylate vinyl group at δ = 6.2–6 ppm and 5.8–5.5 ppm and the increase in the peak associated with the –CH₃ group at δ = 1.8 ppm are consistent with the literature.³⁵ This result indicates that SilMA was successfully prepared.

3.2. Drug-Loaded Microspheres Induce Macrophage Polarization and Immunomodulation of Osteogenic Differentiation In Vitro

The immune microenvironment surrounding the graft influences the bone regeneration process. In this section, we investigated gene expression induced by each group of drug-loaded microspheres (DP, DHP, and PDHP) to promote macrophage polarization toward the M1 or M2 phenotype and used the results to assess the osteogenic differentiation properties of BMSCs in the presence of macrophage-conditioned medium.

As shown in Figure 3A, compared with DP and DHP, PDHP most significantly upregulated the expression of M2-type genes (CD206 and ARG) and suppressed the expression of M1-type genes (TNF-α and IL-1β), whereas DP was superior to DHP. In the presence of macrophage-conditioned medium, the PDHP group exhibited significantly upregulated expression of osteogenesis-related genes (ALP, BMP, and BMP-2) compared to the other groups at all time points. Genes (ALP, BMP-2, OPN, and RUNX2) (Figure 3D). The DP group exhibited osteogenic induction significantly greater than that of the DHP group. This property was also reflected in the staining and quantification of ALP on day 7 and calcium nodules on day 14 (Figure 3B,C), with an increasing trend of DHP, DP, and PDHP, and the most significant changes in the PDHP group.

Figure 3.

Polymerase Chain Reaction (PCR) assay showing the effects of the 3 groups of drug-loaded microspheres on macrophage polarization (A), ALP staining, and activity, calcium nodule staining and quantification, and PCR assay showing the effects of drug-loaded microsphere-macrophage conditioned medium on the osteogenic differentiation of BMSCs (B–D). *p < 0.05, **p < 0.01, and ***p < 0.001.

3.3. Characterization of the Physical and Chemical Properties of the Hydrogels Containing Microspheres

In this portion of the research, we systematically evaluated the injectable properties, cured stability, surface and side morphology, compressive properties, swelling and water absorption properties, degradation properties, and drug release properties, of hydrogel platforms with different porous drug-carrying microsphere contents. The effects of the microsphere content on the main physical and chemical properties of the hydrogel platform were studied.

We conducted a syringe injection experiment and an inversion experiment on the hydrogels to determine the injectability of the composite hydrogels containing drug-loaded microspheres. As shown in Figure 4A, the hydrogels could be extruded from the syringe and quickly cured and maintained their shape after injection, which proves that the SilMA/SA composite hydrogels can be injected after compositing with the drug-loaded microspheres and can maintain their shape after injection and that the SilMA/SA composite hydrogels containing microspheres are injectable and can effectively fill defects with irregular shapes. Figure 4C shows that the composite hydrogels can still maintain their original shape after being inverted for more than 72 h under gravity, which proves their good stability.³⁶ Therefore, composite hydrogels can be injected and adapt to bone defects of different shapes and are not easily lost from bone defects after injection.

Figure 4.

Characterization of the microspheres and hydrogels containing microspheres. Injectability (A) and stability (C) of different hydrogels. (B) SEM images showing the morphology of the surface and longitudinal section of hydrogels containing the microspheres. (C) Images of the inversion test for 0, 0.5, and 72 h after the curing and injection of SilMA/SA and 0.25–2.0% SilMA/SA-M, respectively. [D(1)] and [D(2)] show the compressive stress tests of the five hydrogels. (E–G) swelling, water absorption, and degradation properties of the different hydrogels. *p < 0.05, **p < 0.01, and ***p < 0.001.

Figure 4B shows the morphologies of the surface and longitudinal sections of the hydrogel platforms. Typically, SilMA hydrogels have a macroporous structure and good pore connectivity, while SA hydrogels are denser and have many folded structures on their surface. After SA was doped into SilMA, a denser, nonporous, and heavily wrinkled structure could be observed on the surface and cross-section of the scaffolds, possibly due to the filling of the macroporous structure of SilMA by SA.³⁷ After the addition of porous microspheres to the SilMA/SA network, the porous structure on the surface of the microspheres was not visible because of the hydrogel wrapping the microspheres. As the microsphere content increased, the number of microspheres on the surface of the scaffold increased; as the amount of hydrogel wrapping the microspheres increased, the pore wall inside the scaffold became thinner, and the pore structure became more irregular.^38,39 This result indicates that the pore structure of the hydrogel platforms can be significantly affected by the addition of microspheres.

The addition of microspheres led to an increase in the compressive capacity and Young’s modulus of the hydrogel, with the hydrogel containing 0.5% microspheres showing the best performance, and the mechanical properties deteriorated with a further increase in the microsphere content [Figure 4D(1,2)]. This difference may be due to the addition of an appropriate number of microspheres to the hydrogel network to fill part of the pore structure, while a further increase in the microsphere concentration may change or destroy the cross-linked structure of the SilMA/SA hydrogels, which, in turn, affects the mechanical properties.

Figure 4E–G shows that the molecular chain structure of the hydrogel was altered by the added microspheres, resulting in decreases in the swelling and absorption rates of the hydrogel, and the swelling rate tended to correlate with the degradation rate. The degradation rate of the microsphere-containing hydrogel decreased significantly in a three week degradation experiment, and the effective material filling at the site of prolonged bone defects provided an osteogenic and vasculogenic medium that was favorable for the repair and regeneration of bone defects.

As shown in Figure 5A(1–3), during the first 48 days, the 0.25, 0.5, and 1% groups showed a stable release profile with near zero release, while the 2% group abruptly released the drug between days 16 and 24. The DP group had the greatest abrupt release, with the drug being released by day 36; the DHP group had the lowest abrupt release and the slowest release rate with a significant plateau period; and the PDHP group showed a slow release in the first 6 days, after which the drug release accelerated and approached a zero-level release, releasing nearly 65% of the total drug on day 48. The results showed that the use of dual network hydrogel-coated porous microspheres significantly modulated the release of DEX. After 48 days, the rate of DEX release increased with the degradation of the microspheres and hydrogel. After prolonged immersion in PBS, the backbone of the hydrogel was gradually eroded, and on day 96, the drug release rates of the 0.25 and 0.5% groups increased further and rapidly, whereas the drug was basically depleted from PDHP, and the release entered into a relatively smooth phase. Overall, the drug release behavior of the PDHP group was closer to zero-level release than that of the DP and DHP groups. On day 128, the total DEX release was 100, 85, 65, 60 and 100% for the four hydrogel groups (0.25, 0.5, 1 and 2%, respectively) and PDHP, respectively. The long-term controlled DEX release was beneficial for the ability of the hydrogel to produce osteogenic effects and inhibit bone resorption during bone defect repair. The encapsulation rates of DEX by DP, DHP, and PDHP were 52.5, 98.4 and 82.3%, respectively. We calculated the concentration of DEX in PBS at each time point [Figure 5C(1)] and the concentration of DEX in DMEM at each time point [Figure 5C(2)] to investigate the effect of the concentration of DEX released by the material on cell behavior. The concentration of DEX in DMEM in the DP group was significantly greater than 1000 nM at each time point on days 1–4, whereas the concentration of DEX in the DHP and PDHP groups was less than 1000 nM at each time point on days 1–16. Overall, the concentration of DEX in the PDHP group was higher than that in the DHP group. In the microsphere-containing hydrogel group, the DEX concentration in the 2% group was slightly greater than 1000 nM on days 1, 2, and 6; the DEX concentration in the 0.25% group was lower than that in the other groups (less than 400 nM) at each time point on days 1–16; the concentration in both the 0.5 and 1.0% groups was less than 1000 nM; and the concentration in the 1.0% group was consistently higher than that in the 0.5% group.

Figure 5.

Studies of the drug release performance of microspheres and hydrogels containing microspheres. Cumulative DEX release from the microspheres and hydrogels containing the microspheres in PBS is shown in [A(1–3)]. The amount of Dex released in PBS and DMEM (simulation calculations) at each time point is shown in [B,C(1–2)], respectively.

3.4. Effects of the Hydrogels on Cell Proliferation and Osteogenic Differentiation In Vitro

We evaluated the proliferation of BMSCs using a CCK8 assay and evaluated the osteogenic differentiation of BMSCs using PCR, ALP activity, ALP staining, and calcium nodule staining to systematically study the effects of hydrogel platforms containing different proportions of microspheres on the proliferation and osteogenic differentiation of BMSCs.

Good cytocompatibility is a prerequisite for an ideal biomedical material. BMSCs were cocultured with each hydrogel and assayed with CCK8 at specific time points, and the results are shown in Figure 6A. Compared with the blank hydrogels, the drug-loaded microsphere-containing hydrogels significantly promoted the proliferative activity of the BMSCs at all time points. On day 1, no significant difference in BMSCs proliferation was observed between the different drug-loaded microsphere-containing hydrogel groups; however, on days 3 and 7, the absorbance of the 1% group was significantly higher than that of the other groups. The results indicated that the 1% group was the most effective at promoting the proliferation of BMSCs compared to the other groups.

Figure 6.

Effects of different hydrogels on cell proliferation and osteogenic differentiation. (A) BMSCs proliferation on days 1, 3, and 7 after treatment with hydrogels containing different concentrations of microspheres. [B(1,2)] Osteogenesis-related gene expression on days 7 and 14 after treatment with the hydrogels. [C(1,2)] ALP and calcium nodule quantification. (D,E) ALP staining and Alizarin Red staining results. *p < 0.05, **p < 0.01, and ***p < 0.001.

We measured the expression of osteogenesis-related genes (ALP, BMP, OPN, and Runx2) via qRT–PCR on days 7 and 14 to evaluate the osteogenic differentiation of the hydrogel-treated BMSCs in each group, and the results are displayed in Figure 6B(1,2). The expression of each analyzed gene was significantly upregulated in the 1% group at the two time points, with values significantly greater than those of the control group and the other experimental groups. ALP expression did not significantly differ between the 2% group and the control group, while ALP expression was significantly higher in the 0.25 and 0.5% groups than in the control group. Regarding the expression of BMP and OPN, the groups showed a decreasing trend in the order of 1% > 0.5% > 0.25% > 0%, while the 2% group did not significantly differ from the control group. On day 7, Runx2 expression increased in the following order: 0, 0.25, 2, 0.5, and 1% groups. Unexpectedly, on day 14 of coculture, the expression of Runx2 in the 0.25 and 2% groups decreased, but the difference from that in the control group was not significant. ALP regulates the formation of calcium phosphate in the extracellular matrix and is an important biomarker for early bone formation. Consistent with the ALP quantification and staining results obtained on day 7 [Figure 6C(1),D], the addition of drug-loaded microspheres led to a significant increase in hydrogel-induced ALP expression, which was confirmed by Alizarin Red calcium nodule staining on day 14 [Figure 6C(2),E]; this change is a key indicator of the late stage of osteogenic differentiation. In conclusion, in the in vitro cell proliferation and osteogenic differentiation assays, the 1% group promoted BMSCs proliferation and osteogenic differentiation significantly more effectively than did the other groups. In the subsequent study, the 1% group was selected as the experimental group for the animal bone defect repair experiment.

3.5. Effects of Drug-Loaded Microsphere Hydrogels on Macrophage Polarization and Immunomodulation of Osteogenic Differentiation Properties In Vitro

We examined the expression of macrophage polarization genes (TNF-α, IL-1β, CD206, and ARG) and assessed osteogenic differentiation using macrophage-conditioned medium cocultured with BMSCs to systematically investigate the effects of hydrogel platforms containing different ratios of microspheres on macrophage polarization and immunomodulatory properties.

RAW264.7 cells were cocultured with each hydrogel, and macrophage-polarization-related gene expression was assayed on days 1 and 3. The results are shown in Figure 7A. Compared with the blank hydrogels, the drug-containing microsphere hydrogels significantly promoted the expression of macrophage M2 phenotype genes (CD206 and ARG) while suppressing the expression of inflammatory genes (TNF-α and IL-1β) at all time points. In terms of the ability to induce macrophage polarization toward the M2 phenotype, the effect of the 1% group was the most significant, with the remaining groups showing an increasing trend in the order of the 0, 0.25, 2, and 0.5% groups.

Figure 7.

PCR assay showing the effects of the 5 groups of hydrogels containing different concentrations of microspheres on macrophage polarization (A). ALP staining and activity, calcium nodule staining and quantification, and PCR assay showing the effects of hydrogels containing different concentrations of microspheres in macrophage conditioned medium on the osteogenic differentiation of BMSCs (B–D). *p < 0.05, **p < 0.01, and ***p < 0.001.

We examined the expression of osteogenesis-related genes (ALP, BMP, OPN, and Runx2) on days 7 and 14 to assess the effect of macrophage-CM on the osteogenic differentiation of hydrogel-treated BMSCs in each group, and the results are shown in Figure 7D. Compared with the control group, the drug-loaded microsphere hydrogel group exhibited significantly increased expression of osteogenic genes in BMSCs. At both time points, the expression of each analyzed gene was significantly upregulated in the 1% SS group, with values significantly greater than those of the control group and the other experimental groups. The 2% group showed significantly lower expression of ALP, BMP-2, and Runx-2 than the control group on day 7 and significantly higher expression of individual genes than the control group on day 14. In contrast, the immunomodulatory osteogenic properties of the 2% SS group were lower than those of the 0.5 and 0.25% groups. This finding is in general agreement with the results of the quantitative analysis and staining of ALP on day 7 and calcium nodules on day 14 (Figure 7B,C), which revealed that the addition of drug-loaded microspheres affected ALP expression and mineralized osteogenesis. In conclusion, the 1% group possessed the most significant immuno-coordinated osteogenic properties in the in vitro macrophage polarization induction and immunomodulation of BMSCs osteogenic differentiation.

3.6. In Situ Bone Reconstruction and Regeneration

We established a SD rat cranial defect model to investigate the bone repair effect of the drug-loaded hydrogel platform. The 1% SS treatment with the most significant in vitro osteogenic and immunomodulatory properties was selected as the experimental group and was labeled PDHP/SS. For gradient comparison, a sham-operated group (blank), a hydrogel group (SS), a hydrogel group containing 1% PHP (PHP/SS), and an experimental group (PDHP/hydrogel) were also established. Micro-CT, H&E, Masson’s trichrome, and immunofluorescence histochemical staining (OCN, COL1, CD163, iNOS, VEGF, and CD31) were used to systematically evaluate bone tissue, angiogenesis, and the immune response of macrophages.

As shown in Figure 8A, at 1- and 2 months postsurgery, representative microCT images of the defect site were similar and consistent with the in vitro osteogenic differentiation results. At 1 month, new bone formation at the defect site was significantly greater in the PDHP/SS group than in the other groups. At 2 months, a further increase in calcification and bone tissue formation was observed, and new bone was connected into a block in the PDHP/SS group, but a large bone defect persisted, and no signs of bone healing were observed in the remaining groups. The control and experimental hydrogel groups showed bone tissue growth from the periphery to the center of the defect site over time. The quantitative analysis of relevant microCT parameters confirmed this trend (Figure 8B–F). At 1 and 2 months, the bone volume fraction (BV/TV) and BMD data showed the most significant increase in new bone volume in the PDHP/SS group and a decrease in new bone volume in the PHP/SS, SS, and blank groups, in that order. Among the main indicators of trabecular space morphology, the trabecular thickness (TB.th) and trabecular number (TB.N) increased in the blank, SS, PHP/SS, and PDHP/SS groups, whereas the trabecular separation (TB.sp) decreased in these groups in that order, which was consistent with our expectations.

Figure 8.

In vivo bone regeneration in a rat cranial defect model. Representative micro-CT images of new bone formation at the defect site after 1 and 2 months of treatment. (b–f) Quantitative analysis of the micro-CT parameters (BV/TV, bone mineral density (BMD), TB.th, TB.N, and TB.sp) of regenerated bone at the defect site. *p < 0.05, **p < 0.01, and ***p < 0.001.

We performed histological staining to further assess the presence of new bone at the interface of the hydrogel and the bone defects (Figure 9). H&E staining showed that after 1 month, compared to the blank group, the material group showed a thicker layer of lamellar collagen fiber tissue, with a central lumen and a rich distribution of transverse blood vessels. The PDHP/SS group showed denser neoplastic connective tissue than did the PHP/SS and SS groups, which was dominated by collagen fibers. However, the hydrogel without added microspheres exhibited a faster rate of hydrolysis, uneven distribution of neoplastic tissue, and less internal vascular shadowing. The neoplastic tissues of all three groups were further ossified after 2 months, with better homogeneity of maturation of the neoplastic bone tissue in the PDHP/SS group, which tended to be tightly integrated with the host native bone. The cavity area was still significantly larger in the SS group than in the PHP/SS group. Unexpectedly, the results of Masson’s trichrome staining were consistent with the results of H&E staining, as shown in Figure 9. Dark blue indicates mature bone tissue, light blue indicates fibrous connective tissue, and red indicates myofibrous tissue. After 1 month, the bone defects were mainly filled with fibrous connective tissue, but the connective tissue in the PDHP/SS group was thick and dense, with many red-stained blood vessel walls, and the connective tissue in the remaining groups was relatively sparse. After 2 months, the fibrous connective tissue had formed into mature bone tissue at the defect site after implantation of the hydrogel. After implantation of the hydrogel, the new bone at the defect site was tightly connected to the native bone under the guidance of the material, and the osteogenic effect of the treatment on the PDHP/SS group was the most significant. Moreover, the effects on the other groups were in the order of PHP/SS > SS > blank. These results indicate that the PDHP-functionalized SilMA/SA hydrogel can induce in situ bone tissue regeneration without the addition of exogenous seed cells or growth factors.

Figure 9.

Histological analysis after H&E staining (light red represents new collagen fiber tissue, dark red represents mature collagen tissue, and the dashed box is the enlarged interface diagram of the defect area). Masson’s trichrome staining (light blue represents collagen fiber tissue, dark blue represents mature bone tissue, red represents muscle fiber tissue, and the dotted box is the enlarged interface map of the defect area) of regenerated bone at the defect site after 1 and 2 months.

Immunofluorescence staining for OCN, Col1, CD163, and iNOS was performed after 1 month, and OCN, Col1, CD163, iNOS, VEGF, and CD31 were detected after 2 months to assess osteogenesis, angiogenesis, and immunoreactivity at the defect site. The results are shown in Figure 10. Compared with the other groups, the PDHP/SS group had obvious Col1 and OCN protein deposition at the defect site, indicating better osteogenesis in the DM/hydrogel group, which was consistent with the results of microCT, H&E staining, and Masson’s trichrome staining (Figures 8 and 9). Notably, the expression of the M2 macrophage marker CD163 was significantly upregulated in the PDHP/SS group and was significantly greater than that in the other two groups, while the expression of the M1 macrophage marker iNOS was significantly downregulated, suggesting that PDHP/SS could regulate the polarization of macrophages from the M1 phenotype to the M2 phenotype in vivo. The functional polarization of macrophages plays an important role in the quality and efficiency of bone repair in organisms. The M1 phenotype appears in the early stages of inflammation and recruits many inflammatory cells to induce inflammation, which affects the healing outcome, whereas the M2 phenotype promotes angiogenesis and tissue repair to restore the tissue to its original state. Unexpectedly, the PDHP/SS combination had the greatest effect on the expression of proangiogenic factors with the highest VEGF positivity, and the remaining groups had nonsignificant VEGF fluorescence, followed by the hydrogel and blank groups. According to the results of the above animal experiments, compared with the other hydrogels, the hydrogel containing PDHP promoted bone formation and vascular regeneration, had a stronger immunomodulatory effect, induced macrophage polarization in the M2 direction, and inhibited M1 polarization.

Figure 10.

Immunofluorescence staining of regenerated bone tissue at the defect site after 1 and 2 months. The tissue from the defect area was cut into slices and incubated with primary antibodies (OCN—green, COLI—red, VEGF—green, CD31—red, CD163—green, or iNOS—red) overnight. The cell nuclei were fluorescently stained with DAPI—blue after an incubation with the peroxidase-labeled secondary antibody.

We performed H&E staining of sections of major organs (heart, liver, spleen, lungs, and kidneys) from experimental rats and normal rats at month 2 to evaluate the in vivo biocompatibility of the hydrogels. As shown in Figure S1, a significant difference in H&E-stained sections of major organs was not observed between experimental rats and normal rats. Cardiac sections showed well-arranged cardiomyocytes without bruising or edema, inflammatory cell infiltration, or coagulative necrosis. The liver tissue had a normal cellular morphology, the structure of the liver lobules was intact, and no pitting necrosis, fatty degeneration, etc. were observed. The white marrow, marginal zone, and red marrow structures were clearly visible in the splenic tissue section without abnormal changes. No interstitial thickening was observed in the alveoli, and no congestion of the capillaries and no obvious accumulation of fluid were detected in the alveolar cavities. Renal tissue exhibited an intact glomerular structure, and no granular-free material was found in the interstitial tissue. These results indicate that the hydrogel materials had good biocompatibility in all the groups.

4. Discussion

Minimally invasive, injectable, and in situ-curable drug-controlled hydrogel platforms have received widespread attention due to their ability to accommodate irregularly shaped bone defects and provide stable blood concentrations to accelerate the bone regeneration process during bone repair. However, the currently available hydrogel systems lack long-lasting, controlled release abilities throughout the bone repair cycle. Based on these findings, we prepared and optimized porous HMS/PLGA microspheres loaded with DEX and combined the porous drug-loaded microspheres with SilMA and SA to form a dual-network hydrogel platform that can be injected via in situ photocuring and can release drugs continuously for more than 4 months. Both PDHP and DP microspheres had better immuno-coordinated osteogenic properties than DHP. We chose PDHP microspheres, which are generally better able to modulate macrophage polarization and thus better regulate the osteogenic differentiation of MSCs and provide a longer and more stable release of DEX than DP and DHP microspheres to be compounded with the hydrogel. When the content of microspheres in the hydrogel ranged from 0.25 to 1%, the hydrogel platform was released in vitro for 48 days at the zero stage. When the microsphere concentration was 1%, the hydrogel platform had the best biocompatibility and immunomodulatory effect on the osteogenesis of the BMSCs. The hydrogel containing 1% drug-loaded porous microspheres had the greatest effect on the repair of skull defects in SD rats. Compared with those in the control group and the blank group, vasogenesis and osteogenesis in the PDHP/SS group were significantly increased; M2 macrophage polarization was induced in vivo, and M1 macrophage polarization was inhibited. This treatment alleviates local immune inflammation, promotes vascularization, and collaboratively promotes in situ bone regeneration.

The long-term controlled release performance of a hydrogel platform strongly guarantees that it can provide a stable blood drug concentration. PLGA is one of the most widely used biodegradable materials and has good biocompatibility, and it is favored by researchers for developing bone graft materials as carriers for drug delivery systems.^40,41 The HMS material is characterized by a large specific surface area, mesoporous structure, and facile synthesis.¹² Silicon is an essential trace element for bone mineralization, and it can induce a favorable immune response by polarizing MPs into a healing-promoting M2 phenotype.⁴² This property may also be one of the reasons why DHP microspheres have a better ability to promote the osteogenic differentiation of BMSCs by mediating MP polarization than DP microspheres by inducing the M2 polarization of MPs and inhibiting M1 polarization (Figures 2 and 3). However, HMS, an inorganic material, has a large amount of drug burst release and a short release duration. In this study, the drug loading rate of the DEX/HMS system reached 99.5%, and DEX was dispersed within the HMS mesopores, as observed using SEM and confirmed by the BET and XRD results. The effective adsorption of DEX to the HMS pore system was evidenced by blocked pores, a greater average pore size, and a lower surface area and pore volume, which are consistent with the results of a previous study (Figure 2A–D).¹² PLGA microspheres also have a general release time of approximately 1 month, and HMS-modified PLGA microspheres can significantly prolong the time of slow drug release; however, zero-level release is still not possible (Figure 5).¹² Furthermore, the porous structure endowed with HMS/PLGA microspheres maintained long-term drug release, which may be attributed to the strong drug loading and slowing ability of HMS. More importantly, the drug release behavior of the porous HMS/PLGA microspheres was closer to zero-grade release than that of the PLGA and HMS/PLGA microspheres, which provides ideas for further preparation of composites that can release drugs at zero grade for a long time. Degradable polyester-based microspheres are not convenient for bone repair alone due to their shape and size. Hydrogel-based drug delivery systems are highly biocompatible, and satisfactory osteogenic results have been obtained in previous studies. We hypothesized that the hydrogels could enhance the delayed release of the drug from these porous DEX/HMS/PLGA microspheres.^43,44 Here, SilMA/SA composite PDHP was utilized to construct a drug-loaded hydrogel system. In the 128 day drug release experiment, the porous drug-loaded microsphere-containing hydrogels in the 0.25, 0.5, and 1% groups exhibited stable sustained release behaviors during the first 48 days, with near-zero release (controlled release). Compared with the slow release of drugs, the controlled release of drugs results in the continuous release of drugs for a certain period, and during the early stage of bone defect repair, the continuous administration of a stable dose of DEX can not only inhibit the process of bone resorption but also promote the proliferation of osteoblasts and regulate their immune function to a certain extent.⁴⁵ Interestingly, the drug release rate of degradable polyester microspheres was positively correlated with the rate of erosion, with PDHP alone degrading rapidly in PBS and exhibiting nearly first-order drug release.⁴⁶ However, the drug release rate of the PDHP microspheres was significantly decreased by hydrogel encapsulation, which may be attributed to the protective effect of the hydrogel on the delay of the erosive destruction of PDHP.

The long-acting, controlled-release drug-carrying hydrogel system has a strong ability to coordinate osteogenesis, especially in bone defects with abnormally active bone resorption due to bone tumors, osteoporosis, etc. By analyzing microspheres promoting the proliferation and differentiation of BMSCs in vitro, we found that the porous structure of the surface endowed the microspheres with stronger osteogenic properties, which were significantly enhanced by the addition of DEX (Figure 2F–G). DEX released from porous microspheres may facilitate the promotion of bone marrow mesenchymal stem cell proliferation and osteogenesis.²² Continuous administration of DEX at a stable dose not only promotes the proliferation of BMSCs and osteogenic differentiation but also regulates the immune microenvironment of bone grafts, alleviates inflammatory reactions, and enhances the survival rate of implants.³² With the development of bone tissue engineering approaches, hydrogels with unique hydrophilic properties have attracted widespread attention, but they lack cell recognition sites and have a low osteogenic capacity.⁴⁷ ALP is an active marker of early osteogenesis and promotes mineralization and calcium salt deposition, and calcium nodule deposition is one of the hallmarks of bone maturation.⁴⁸ BMP2 has great osteogenic differentiation potential for BMSCs, and the transcription factor RUNX2 is a downstream factor in the BMP2 signaling pathway and a major regulator of osteoblast development and bone formation. OPN secreted by osteoblasts contributes to the binding of hydroxyapatite, which plays an important role in regulating the process of bone matrix mineralization.⁴⁹ In terms of the in vitro osteogenic performance of the drug-loaded microsphere hydrogels, we determined the expression of osteogenic markers at specific time points. The drug-loaded microsphere hydrogels were significantly more effective than the SilMA/SA hydrogels at promoting BMSCs proliferation and osteogenic differentiation in vitro, with the 1% group exhibiting the best performance (Figure 6), which may be related to the fact that the concentration of DEX released from the 1% group could best promote the proliferation and osteogenic differentiation of BMSCs. When the concentration of DEX was less than 1000 nM, the regulation of the osteogenic differentiation of MSCs by DEX was concentration dependent; i.e., the higher the DEX concentration was, the more favorable it was for promoting osteogenic differentiation.^23,50 However, DEX concentrations greater than 1000 nM adversely affect the osteogenic differentiation of the cells involved and bone healing.^24,51−53 The concentration of DEX in the DP group in DMEM on days 1–4 exceeded 1000 nM, while the DEX concentration in both the DHP and PDHP groups on days 1–16 was less than 1000 nM, and overall, the DEX concentration in the PDHP group was higher than that in the DHP group. The DEX concentration in the DP group significantly exceeded 1000 nM in the first 4 days, which was unfavorable for the induction of the osteogenic differentiation of BMSCs by DP. The ALP activity of BMSCs cocultured with the three groups of microspheres tended to decrease in the order of PDHP > DHP > DP, which was consistent with what has been reported in the literature.

Binding of DEX to its receptor inhibits the signaling and activation of inflammatory transcription factors such as nuclear factor kappa B (NF-κB) and activator protein-1 (AP-1), thereby suppressing the production of inflammatory mediators and producing anti-inflammatory and immunosuppressive effects. For the NF-κB signaling pathway involved in M1 polarization, DEX can inhibit NF-κB activation by enhancing the transcription of NF-suppressor protein α (I-κBα), which increases the level of I-κBα and inhibits NF-κB activation. Moreover, the activated DEX receptor can directly interact with activated NF-κBp65 to block the binding between NF-κB and target DNA, which ultimately inhibits M1 polarization of macrophages and attenuates the inflammatory response to DEX by inhibiting the activation of F-κBp65.⁵⁴ In addition, the anti-inflammatory effects of DEX are also related to its ability to induce M2 polarization. The ability of DEX to modulate macrophage polarization depends on its concentration. For example, Nakamura et al. reported that the expression of inflammatory genes (CXCL10, CXCl11, PTGS2, TNFα, and IL1β) in human macrophages decreased significantly when the concentration of DEX increased from 10 to 100 nM and that the expression of inflammatory genes further decreased when the concentration of DEX increased from 100 to 1000 nM.⁵⁵ In a study by Bensiamar et al., increasing the concentration of DEX from 10 to 100 nM resulted in a significant decrease in the levels of inflammatory factors, such as TNFα and IL6, secreted by BMSCs and TPA-treated THP-1 cells (dTHP-1); the levels of inflammatory genes in the 1000 nM DEX-treated group were slightly lower than those in the 100 nM group.⁵⁶ Takato et al. showed that the level of CD206, a phenotypic marker of anti-inflammatory M2 macrophages secreted by immortalized porcine nephrogenic macrophages, was significantly elevated when the concentration of DEX increased from 0 to 80 nM.⁵⁷ Asger et al. reported that the sequential treatment of porcine whole blood with lipopolysaccharide and DEX resulted in the release of TNF-α close to the same level when the DEX concentration was increased from 255 nM to 2547 nM.⁵⁸ Si et al. reported that both 10 μM and 100 μM DEX downregulated TNF-α and IL-12 and upregulated IL-10 and TFG-β in LPS-stimulated MPs.⁵⁹ After coculturing, the three groups of microspheres with macrophages, trends of DHP > PDHP ≈ DP in the expression of inflammatory genes and PDHP > DP > DHP in the expression of anti-inflammatory genes were observed, which may be attributed to the fact that increasing the concentration of DEX is beneficial for enhancing its ability to inhibit macrophage M1 polarization and promote M2 polarization in a certain concentration range; however, when the concentration of DEX exceeds a certain range, its ability to regulate macrophage fractional polarization does not change significantly. However, although PLGA is a nontoxic and harmless material that can be absorbed by the human body, it tends to be an inflammatory stimulant for MPs.⁶⁰ Considering the drug release properties of drug-carrying microspheres and their effects on the osteogenic differentiation of BMSCs, MP fractional polarization, and mediating the MP-induced osteogenic differentiation of BMSCs, we chose PDHP microspheres, which can release DEX for a long time, have a release profile close to zero-grade release, are the most capable of promoting the osteogenic differentiation of BMSCs and have good immunoregulatory effects on osteogenesis, and can be combined with a double-networked hydrogel to construct an injectable drug-carrying hydrogel platform.

In the microsphere-containing hydrogel group, the concentration of DEX in DMEM was greater than 1000 nM for the DP group at each time point on days 1–4, whereas the DEX concentration was less than 1000 nM for both the DHP and PDHP groups at each time point on days 1–16. Overall, the DEX concentration in the PDHP group was higher than that in the DHP group. In the microsphere-containing hydrogel group, the DEX concentration in the 2% group was slightly greater than 1000 nM on days 1, 2, and 6; the DEX concentration in the 0.25% group was lower than that in the other groups (less than 400 nM) at each time point from days 1–16; the concentrations in both the 0.5% and 1.0% groups were less than 1000 nM; and the concentration in the 1.0% group was consistently greater than that in the 0.5% group. Although the DEX concentration in the 2% group was higher than that in the 1% group overall, the DEX concentration in the 2% group was slightly greater than 1000 nM at individual time points, making it weaker than that in the 1.0% group in terms of its ability to induce osteogenic differentiation of BMSCs. Overall, the group treated with 1% microsphere-containing hydrogels had the strongest ability to induce osteogenic differentiation, which is also consistent with what has been reported in the literature. The 1% group had the best performance in mediating macrophage polarization and the immunomodulation of osteogenesis. Overall, the amount of DEX released from the composites tended to increase when the content of PDHP microspheres increased from 0.25 to 2.0% over 16 days. In general, higher concentrations of DEX were more conducive to inhibiting the inflammatory differentiation of MPs and promoting M2 differentiation. However, with increasing microsphere content, the amount of acidic byproducts generated during the degradation of the composites increased, leading to an increase in the M1 polarization of the MPs.⁶⁰ In this study, the 1% group exhibited the most successful immunomodulation, which may be mainly due to the combined effects of DEX release and the PLGA degradation products.

The hydrogel platform, which can be injected and cured in situ, can adapt to irregular bone defects and has been widely studied. Good biocompatibility is also necessary for the regeneration and repair of bones and other tissues. Moreover, the hydrogel platform should exhibit suitable swelling and water absorption performance, degradation performance, and mechanical strength so that it will not be excessively deformed or rapidly degraded after implantation in the body. The hydrogel system developed in this study is biocompatible and degradable. It is also injectable and curable in situ and can be adapted for irregularly shaped bone defects, avoiding the secondary trauma associated with invasive surgery (Figure 4A,C).⁶¹ PLGA is one of the most widely used biodegradable materials, and HMS is a widely used bio ceramic; all of these materials have good biocompatibility.⁶² As a natural hydrogel, SF can be easily modified and prepared into 3D networks with a high-water content, and they are considered promising biomedical materials. The methacrylation of SF by glycidyl methacrylate introduces a double bond on the SF molecule, which allows SilMA to form light-curable hydrogels due to the introduction of additional chemical groups. After the addition of SA to SilMA, the hybrid hydrogel exhibited improved biocompatibility and degradation properties.^63,64 Compounding PDHP on SilMA/SA did not weaken the stability or injectability. Hydrogels provide an excellent extracellular matrix for colonization by bone regenerative cells; however, the poor compressive properties of hydrogels limit their use in bone defects exposed to stress. We utilized SilMA and SA to construct a dual-network cross-linked structure, and the incorporation of porous HMS/PLGA composite microspheres drastically improved the mechanical properties and reduced the swelling, water absorption, and degradation properties of the hydrogels, further expanding their application potential (Figure 5D–G).^65,66

We selected PDHP/SS as the experimental group based on the above findings and created a rat cranial bone defect model for in vivo experiments to further validate the osteogenic properties of this hydrogel and compare them with those of the control group (PHP/SS, SS) and the blank group. Micro-CT results showed that as a filler material for bone defects, the hydrogel was able to guide bone mineralization from the periphery to the center. In both the first and second months, the amount of new bone and the density and thickness of the bone trabeculae were significantly greater in the experimental group than in the other groups (Figure 8). According to the results of H&E staining and Masson’s trichrome, the experimental group had thicker lamellar fibrous connective tissue and was rich in internal striated blood vessels (Figure 9). These results indicated that the hydrogel containing drug-loaded microspheres exerted significantly stronger osteogenic and angiogenic effects than those of the other treatments. The in vivo osteogenic effect of PHP/SS was superior to that of SS due to the incorporation of PHP microspheres, which have bone-enhancing effects, whereas the effect of SS on the blank group may be due to the presence of a hydrogel that provides a medium for bone regeneration. During bone regeneration, COLI is deposited in the bone matrix, induces differentiation of BMSCs, and contributes to the formation of mineralized nodules. OCN is involved in the regulation of calcium ion stabilization and bone mineralization and is a marker of late osteoblasts.⁴⁹ The immunomodulatory capacity of grafts can facilitate the induction of osteogenic and angiogenic microenvironments. iNOS and CD163, which are M1 and M2 macrophage markers, respectively, can be used to assess the polarization behavior of macrophages during bone repair, whereas the expression of angiogenesis-related genes, such as VEGF and CD31, recruits endothelial cells for vascularization at defects and promotes osteogenesis.⁶⁷ As shown in the images of immunofluorescence staining (Figure 10), the DM/hydrogel group exhibited substantial accumulation of osteogenesis-related proteins (OCN and ColI) at the defect site. In terms of the immunomodulatory ability, the expression of the macrophage M2 marker CD163 was significantly upregulated, whereas that of the macrophage M1 marker iNOS was significantly suppressed, further suggesting that PDHP/SS has a greater ability to promote the polarization of macrophages from the M1 to M2 phenotype than the other two treatments, forming an immune microenvironment enriched with M2-type macrophages, which play a positive role in the early stage of bone repair. These behaviors can alleviate inflammation during the early stage of bone repair and promote the transition of bone healing from the inflammatory phase to the healing phase.⁶⁸ Notably, the vascularization of tissue-engineered materials is an important process in bone defect repair and is a key factor affecting bone repair and remodeling.⁶⁹ Studies have shown that an immune microenvironment enriched with M2-type macrophages favors angiogenesis during bone regeneration, which was supported by the significant VEGF-positive staining in immunofluorescence assays at month 2 in the PDHP/SS group.⁷⁰

In this study, although hydrogels usually do not release drugs for a long period, compounding the porous drug-carrying microspheres obtained from the study, which are close to the zero-grade release effect, with the dual-network hydrogel can further modulate the drug release behavior of the hydrogel. Here, we prepared 1% PDHP/SS with an effective DEX release cycle of 120 days, completely covering the bone regeneration cycle. What is more, DEX was released at a near zero level for about 48 days and had a low burst release with an overall concentration between 400 and 1000 nM, which was conducive to the modulation of macrophage polarization throughout the bone regeneration cycle and the formation of an immune microenvironment that contributes to bone regeneration and better promotes bone regeneration. In summary, the hydrogel platform constructed in this study has excellent performance in coordinating osteogenesis and provides new ideas for the development of bone grafting materials. However, future studies are needed to validate the safety of these hydrogel platforms for humans, and additional studies are needed in other large animal models, such as rabbits, dogs, and pigs, to strengthen preclinical studies.

5. Conclusions

The construction of a long-term and stable drug delivery system to promote the regeneration of different types of bone defects while reducing the level of damage is still a major challenge. Our injectable and light-curable porous drug-carrying microsphere-containing hydrogel drug-controlled release system can be applied to bone defects of different shapes. PDHP encapsulated in the SilMA/SA dual-network hydrogel can be used to effectively control the release of DEX for more than 4 months, providing a stable blood concentration to promote bone regeneration for the repair of bone defects, and can be adapted for the treatment of osteoporosis. PDHP are suitable for the treatment of bone defects involving excessive bone resorption, such as defects associated with osteoporosis and tumors. The composite hydrogel has good biocompatibility and osteogenic activity and can induce macrophage M2 polarization, inhibit macrophage M1 polarization, relieve inflammation, and promote the regeneration of blood vessels at the bone defect site in vitro and in vivo, accelerating the process of in situ bone regeneration. This study provides a potential minimally invasive filler material for clinical application in the field of bone defect repair. This study provides a more long-term DEX-controlled release system for bone defect repair. This system is injectable, has in situ photocuring properties, can reduce the damage caused by invasive operations, can adapt to different bone defects requiring long-term stable drug delivery under different clinicopathological conditions, and has great clinical application potential.

Data Availability Statement

If possible and applicable, we will deposit data that support the findings of our research in a public repository.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsami.4c06661.

• H&E staining of heart, liver, spleen, lung, and kidney sections from normal rats and experimental rats (PDF)

The authors declare no competing financial interest.

Notes

Ethics statement: All animal experiments were performed under the protocol approved by the Institutional Animal Care and Use Committee of the Guangdong Quality Supervision and Testing Station for Medical and Health Care Appliances.