Development of Small Molecular Drug-Loaded Microspheres for Enhanced Cartilage Regeneration

Abstract

Articular cartilage has limited self-repair ability, making osteoarthritis a major challenge due to aging and mechanical stress. While current therapies provide partial functional improvement, their outcomes remain suboptimal, highlighting the need for regenerative strategies. Mesenchymal stem cells (MSCs) show promises due to their chondrogenic potential, and kartogenin (KGN) enhances this process. This study examines KGN-loaded gelatin microspheres (KMs) for cartilage regeneration. KMs were synthesized via gelatin-KGN dispersion in ice-cooled hexane, followed by glutaraldehyde cross-linking. Scanning electron microscopy showed porous KMs (200-800 μm) with interconnected pores (2-10 μm), while Fourier transform infrared spectroscopy confirmed gelatin and KGN presence. hMSCs were cultured with KMs, showing no cytotoxicity. Gene expression analysis revealed upregulated chondrogenic markers (SOX9, ACAN, COMP, COL2A1). Western blotting and immunofluorescence confirmed increased chondrogenic protein production. Sulfated glycosaminoglycan content increased over four weeks, indicating extracellular matrix maturation. This study demonstrates that KMs effectively deliver KGN, enhancing MSC chondrogenesis. Small molecule-based biomaterials may offer an alternative to growth factors in osteochondral tissue engineering, warranting further in vivo validation.

Highlights:

• Kartogenin-loaded microspheres support cartilage regeneration using a small molecule approach

• Microspheres exhibit porous structure enabling cell attachment and sustained KGN delivery

• No cytotoxicity observed; microspheres are biocompatible with human MSCs

• Chondrogenic gene and protein markers significantly upregulated with microsphere treatment

• Enhanced extracellular matrix production confirms functional chondrogenic differentiation

• Provides a growth factor-free strategy for osteochondral tissue engineering

Introduction

Articular (hyaline) cartilage, which is characterized by being aneural, avascular, alymphatic, and hypocellular, has long been a key subject of clinical research^[1]. Its extracellular matrix (ECM) contains a high-water content, with water making up 80% of its total weight, emphasizing its critical role in joint lubrication and wear resistance. The dry weight of the cartilage consists mainly of type II collagen and proteoglycans, which are essential for load-bearing functions^[2]. Under normal conditions, chondrogenic tissues have limited healing ability. Cartilage damage poses a significant clinical challenge due to its poor regenerative capacity. Several factors, such as aging, joint instability, excessive mechanical stress, diet, hormones, crystal deposition, bone microfractures, and immune-related mechanisms, play a role in the development of osteoarthritis (OA)^[3]. OA is often mistakenly thought to be caused by decreased metabolic activity, but it is actually an active catabolic process. In response to cartilage injury, there is an increase in matrix synthesis and chondrocyte proliferation. However, excessive metabolic activity leads to the release of lysosomal enzymes, which break down the cartilage matrix and reduce proteoglycan content, worsening the disease^[4]. Over time, cartilage degradation outpaces its synthesis, leading to erosion and lesion formation.

The synthesis and secretion of ECM components in cartilage can be influenced and enhanced by signaling molecules in the surrounding ECM^[5]. Previous research has investigated biomaterial-based approaches to replicate the natural cartilage environment^[6]. One such strategy involved creating a gelatin-chondroitin-6-sulfate-hyaluronan tri-copolymer, which effectively promoted ECM secretion, lacunae formation, and glycosaminoglycan (GAG) production^[7]. This tri-copolymer showed promise as a scaffold for cartilage tissue engineering. However, using chondrocytes as a cell source remains impractical due to donor site morbidity. Current clinical therapies for cartilage repair still have limitations, rarely achieving complete functional restoration or returning damaged tissue to its native state.

Full-thickness cartilage defects are frequently seen, but no standard medical treatment has proven to be effective for long-term repair. Techniques like marrow stimulation, including drilling and microfracture, are typically restricted to smaller defect areas and involve the injection of multipotent stem cells. In contrast, the implantation of cultured autologous cells or engineered tissue constructs offers a more promising solution for regenerating hyaline-like cartilage^[8]. Cell-based regenerative therapies have emerged as an innovative surgical approach for repairing cartilage and play a critical role in managing cartilage-related disorders^[9]. However, these treatments face various challenges, such as donor site morbidity, the formation of fibrous cartilage, poor graft integration, and the loss of cell viability during storage. Major obstacles in cartilage reconstruction include selecting appropriate cell sources, designing optimal scaffolds, ensuring biomechanically functional tissue formation, and achieving smooth integration with the native tissue after implantation^[10].

Mesenchymal stem cells (MSCs) are considered a promising cell source for regenerative medicine due to their ability to self-renew and their secretion of bioactive molecules, such as growth factors and cytokines, which help prevent rejection of allogeneic cells^[11]. These bioactive factors contribute to creating a regenerative microenvironment at defect sites, reducing tissue damage and promoting repair by interacting with resident cells. MSCs can be easily isolated from bone marrow aspirates, expanded in culture while preserving their multipotency, and have been extensively studied in preclinical trials for tissue engineering applications^[12]. Their therapeutic potential in repairing and reconstructing damaged mesenchymal tissues is widely acknowledged.

Kartogenin (KGN), a small molecule, has been demonstrated to promote chondrocyte differentiation both in vitro and in animal models^[13]. Research shows that KGN regulates the Runt-related transcription factor 1 (CBFβ-Runx1) pathway to induce chondrogenesis. It encourages mesenchymal stem cells (MSCs) to differentiate into matrix-producing chondrocytes by upregulating type II collagen (Col2a1), SRY-box 9 (Sox9), aggrecan (Acan), and tissue inhibitors of metalloproteinases (TIMPs), while simultaneously downregulating Runx2-related genes to prevent chondrocyte hypertrophy^[14]. However, the full potential of KGN in combination with biomaterial scaffolds remains under investigation.

Gelatin, a natural polymer derived from collagen, is widely utilized in pharmaceutical and medical applications due to its biodegradability and biocompatibility in physiological environments^[15]. Various gelatinbased matrices have been developed for controlled-release applications. The degradation rate of gelatin hydrogels in vivo is affected by their cross-linking density, which also influences the release profile of biomolecules^[15]. Research indicates that gelatin/biomolecule complexes are released through enzymatic degradation of the gelatin carrier in vivo^[16]. Gelatin-based hydrogels or sponges can control the release of growth factors, thus enhancing their biological function in tissue regeneration^[17]. Based on these properties, this study proposes the development of a gelatin microsphere system loaded with kartogenin for cartilage tissue regeneration.

Materials and methods

Materials

Glutaraldehyde (GA, CAS No. 111-30-8), kartogenin (KGN, CAS No. 4727-31-5), and gelatin (CAS No. 9000-70-8) were purchased from Sigma-Aldrich (St. Louis, MO, USA).

Preparation of Kartogenin Microspheres (KMs)

A solution was prepared by dissolving 10 g of gelatin and 10 M kartogenin (KGN) while stirring magnetically at 50 °C for 30 minutes. The resulting mixture was transferred to a 10 mL syringe and then dispersed into an ice-cooled hexane bath to form kartogenin microspheres (KMs). Cross-linking of the microspheres was subsequently carried out using a 1% glutaraldehyde solution.

Morphology of KMs

The microstructure of the kartogenin microspheres (KMs) was analyzed using scanning electron microscopy (SEM). The samples were first sputter-coated with gold and then mounted on adhesive copper stubs before being imaged with a Hitachi S3000/N electron microscope.

Functional Group Identification

Fourier transform infrared (FTIR) spectroscopy was employed to analyze the functional groups of organic compounds by detecting changes in dipole moments. FTIR spectra of various scaffolds were recorded using a Perkin Elmer spectrophotometer (Waltham, MA, USA) over a wavelength range of 450–4000 cm⁻¹.

Isolation and Expansion of Human MSCs (hMSCs) for In Vitro Experiments

The use of human mesenchymal stem cells (hMSCs) was approved by the National Taiwan University Hospital Institutional Review Board (NTUH IRB No. 201704005 RINA). hMSCs were sourced from bone marrow aspirates collected during total hip and knee joint replacement surgeries. Mononuclear cells were isolated using Ficoll-Paque PLUS (GE Healthcare, Amersham, UK). hMSCs from passages 3 to 4 were utilized for the subsequent experiments.

Cell Viability

The viability of L929 cells (BCRC Strain Collection, Hsinchu, Taiwan) was evaluated using a water-soluble tetrazolium (WST-1) assay in accordance with the ISO 10993-5 standard. An extract medium was prepared by dissolving 0.2 g/mL of the scaffold in high-glucose DMEM (Sigma-Aldrich) and incubating it at 37 °C for 24 hours. L929 cells were seeded in 96-well plates at a density of 5 × 10³ cells per well and cultured at 37 °C for 1 day. The culture medium was then replaced with the extract medium, and incubation continued for 1 to 3 days. Prior to measurement, 10 μL of WST-1 reagent was added to each well and incubated for 4 hours. Formazan formation was quantified using a Sunrise spectrophotometric plate reader (ELISA reader, Tecan, Männedorf, Switzerland), with absorbance recorded at 450 nm (reference filter at 600 nm). Cell viability was calculated using the equation,

$$\mathrm{Cell} \mathrm{viability} (\%)=\frac{(\mathrm{OD} \mathrm{experiment} - \mathrm{OD} \mathrm{background}) \times 100}{(\mathrm{OD} \mathrm{control} - \mathrm{OD} \mathrm{background})}$$

Cytotoxicity

Cytotoxicity was assessed using the CytoTox 96 assay kit (Promega, Madison, WI, USA) to measure extracellular lactate dehydrogenase (LDH) release. After transferring the suspension medium, the LDH substrate solution was added to fresh enzymatic assay plates. Following a 30-minute incubation, a stop solution was added to each well, and absorbance was measured at 490 nm using a Tecan Sunrise ELISA spectrophotometric plate reader. Cytotoxicity was calculated using the equation,

$$\mathrm{Cytotoxicity}(\%)\frac{(\mathrm{Experimental} \mathrm{value} - \mathrm{Negative} \mathrm{control}) \times 100}{(\mathrm{Positive} \mathrm{control}- \mathrm{Negative} \mathrm{control})}$$

Live/Dead Assay

Cytotoxicity was further evaluated using live/dead staining. Cell constructs were incubated with 4 μM calcein AM and 4 μM propidium iodide (PI) (Life Technologies, Carlsbad, CA, USA) for 30 minutes. Live cells exhibited green fluorescence (calcein AM, ex/em ~495 nm/~515 nm), while dead cells displayed red fluorescence (PI, ex/em ~540 nm/~615 nm).

Gene Expression

Total RNA was extracted from cell constructs at various time points using a Total RNA Miniprep Purification Kit (GeneMark, Zhubei, Taiwan). First-strand cDNA was synthesized with random hexamers and reverse transcriptase (Vivantis, CA, USA; Cat No: RTPL12) under the following PCR conditions: 95 °C for 3 minutes (denaturation), followed by 40 cycles of 95 °C for 20 seconds, 60 °C for 30 seconds (annealing), and 72 °C for 30 seconds (elongation). Real-time RT-PCR was conducted using TOOLS 2X SYBR qPCR Mix (Biotools, Taipei, Taiwan) on a CFX Connect Real-Time PCR Detection System (BioRad, Hercules, CA, USA). Gene expression levels were normalized to GAPDH and calculated using the 2^-ΔΔCt method. The relative expression of four cartilage-related genes (ACAN, SOX9, COL2A1, and COMP) was measured on days 7, 14, 21, and 28. Chondro-specific primers (Biotools) used for chondrogenic differentiation are listed in Table 1.

Table 1. Primers used for chondrogenic differentiation.

Gene name	Forward primer (5′–3′)	Reverse primer (5′–3′)
SOX9	GGCAAGCTCTGGAGACTTCTG	CCCGTTCTTCACCGACTTCC
ACAN	TAAAAAGGGCACAGCCACCAC	GTGAGCTCCGCTTCTGTAGTC
COMP	CAAGGCCAACAAGCAGGTTT	TATGTTGCCCGGTCTCACAC
COL2A1	ATGAGGGCGCGGTAGAGA	GCCAGCCTCCTGGACATC
GAPDH	AATGGGCAGCCGTTAGGAAA	GCCCAATACGACCAAATCAGAG

Western Blotting

After four weeks of co-culture with KMs, hMSCs from each group were washed with phosphate-buffered saline (PBS) and lysed using pre-cooled lysis buffer (Biotools). Three samples per group were selected for analysis. Total protein concentration was determined using a BCA Protein Assay (Pierce Chemicals, Rockford, IL, USA). Equal amounts of protein were separated by SDS-PAGE (8%–12%) and transferred to a PVDF membrane (Merck Millipore, Burlington, MA, USA). The membranes were blocked with 5% (w/v) non-fat milk for 1 hour at 25 °C and incubated overnight at 4 °C with primary antibodies (Abcam, Cambridge, UK) targeting ACAN (ab3778, 1:1000), COL2A1 (ab34712, 1:1000), COMP (ab300555, 1:1000), and GAPDH (ab8245, 1:1000) as an internal control. Following incubation with secondary antibodies for 2 hours at 37 °C, protein bands were visualized using enhanced chemiluminescence (ECL). Relative protein expression was normalized to the internal control and the control group using ImageJ software (NIH, Bethesda, MD, USA).

Biochemical Analysis

Sulfated glycosaminoglycan (sGAG) content was quantified using the 9-dimethyl methylene blue chloride (DMMB) method with a Blyscan Assay Kit (Biocolor, Carrickfergus, UK). Total DNA was extracted using the Genomic Geno Plus DNA Extraction Miniprep System (Viogen, Taipei, Taiwan) and quantified with the Quant-iT PicoGreen dsDNA Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA). The sGAG-to-dsDNA ratio was averaged across samples. Bone- and cartilage-specific proteins were measured using commercial ELISA kits (FineTest, Wuhan Fine Biotechnology, Wuhan, China).

Immunofluorescence Staining

Samples were incubated overnight at 4 °C with primary antibodies (Abcam) targeting ACAN (rabbit monoclonal, 1:200) and COL2A1 (rabbit polyclonal, 1:200). The following day, samples were washed and incubated with secondary antibodies (1:200) at room temperature for 30 minutes, then stained with DAPI (Thermo Fisher Scientific) to visualize the nuclei. After a final PBS wash, the samples were examined using a fluorescence microscope (Axio Imager ZI, Zeiss, Germany).

Statistical analysis

The results are expressed as mean ± standard deviation (SD). Statistical analysis was conducted using one-way ANOVA (SPSS Statistics 29, IBM, Armonk, NY, USA), with significance set at p < 0.05

Results

Morphology of KMs

Figure 1 shows the scanning electron microscopy (SEM) images of the KMs, displaying particle sizes that range from 200 μm to 800 μm. The surface of the particles also reveals pore sizes varying between 2 μm and 10 μm.

Figure 1. The scanning electron microscopy (SEM) images of KMs at different magnifications: (A) 50× and (B) 3000×, reveal an open and interconnected porous structure with uniformly distributed pores throughout the KMs.

Functional Group Identification

Figure 2 shows the distinct peaks in the infrared spectrum of the KMs. The presence of gelatin was identified by peaks at 2800–2950 cm⁻¹ (CH stretching), 1650 cm⁻¹ (C=O stretching), and 3430 cm⁻¹ (O-H stretching). Furthermore, the FTIR spectrum of KGN in the KMs displayed clear bands at 1537–1596 cm⁻¹, corresponding to the C=C bending vibration from the aromatic ring of KGN.

Figure 2. The FTIR spectrum of the KMs.

Biocompatibility of the KMs

The biocompatibility of KMs was evaluated in L929 fibroblasts using WST-1 and LDH assays on days 1, 2, and 3 of culture (Figure 3). The results showed no significant differences in cell viability between the control group and the KMs-treated groups, suggesting that KMs are non-toxic to L929 fibroblasts. Additionally, live/dead staining revealed no noticeable change in the ratio of dead (red) to live (green) cells across various KMs extract concentrations, further supporting that KMs do not induce cytotoxicity.

Figure 3. Evaluating the biocompatibility of KMs. (A) Cell viability of KMs by WST-1 assay (n = 12, *** p < 0.001 compared with control). (B) Cytotoxicity of KMs by LDH assay (n = 12, *** p < 0.001 compared with control). (C) Live/dead staining of KMs.

Gene Expression Analysis by Qpcr

qPCR experiments were conducted over four weeks to assess gene expression in hMSCs cultured with KMs (Figure 4). To evaluate chondrogenic differentiation, RNA interference was used to analyze the expression of key chondrogenic markers, including SOX9, ACAN, COMP, and COL2A1. The temporal expression patterns of SOX9, ACAN, COMP, and COL2A1 were analyzed over a 28-day period. Notably, SOX9 and ACAN exhibited peak expression at day 14, suggesting their involvement in the early to mid-stages of chondrogenesis. In contrast, COL2A1 and COMP expression gradually increased, reaching their highest levels at day 21 or 28, which indicates their role in the later stages of cartilage matrix maturation. These results demonstrate a time-dependent expression profile, highlighting the dynamic progression of chondrogenic differentiation induced by KMs. Overall, gene expression was significantly upregulated in hMSCs cultured with KMs compared to the control group, indicating successful differentiation into prechondroblasts.

Figure 4. The results present the relative chondrogenic gene expression levels in hMSCs cultured with KMs, normalized to the control group. The upregulation of key chondrogenic markers, including SOX9, ACAN, COMP, and COL2A1, indicates enhanced chondrogenic differentiation in the presence of KMs (n = 12, ** p < 0.01 compared with day7, *** p < 0.001 compared with day7).

Quantification and expression of biomarkers

An increase in the levels of chondrogenic proteins ACAN, COMP, and COL2A1 served as a positive indicator of chondrogenesis. Western blot analysis confirmed elevated expression of these biomarkers in hMSCs cultured with KMs, further validating that KMs effectively promote chondrogenesis in vitro (Figure 5).

Figure 5. Western blotting was performed to assess the expression of chondrogenic proteins in hMSCs co-cultured with KMs for four weeks. (A) The amount of western blot product in hMSCs was set to 1.0, and the relative ratio of the protein expression is indicated on the ordinate. (B) Chondrogenic protein expression levels were quantified and normalized to GAPDH as an internal control. The results demonstrate that KMs enhance the expression of chondrogenic markers, further supporting their role in promoting chondrogenesis.

Biochemical analysis

The DMMB method revealed a significant increase in the sGAG/dsDNA ratio with prolonged culture time up to 4 weeks (Figure 6), indicating progressive extracellular matrix (ECM) deposition and glycosaminoglycan (GAG) accumulation. This suggests enhanced chondrogenic differentiation and matrix maturation, highlighting the potential of KMs to support cartilage tissue development and regeneration.

Figure 6. Biochemical analysis of chondrogenic differentiation. Ratio of sGAG to dsDNA content after 7, 14, 21, and 28 d of differentiation (n = 12, ** p < 0.01 compared with control, *** p < 0.001 compared with control).

Immunofluorescence analysis

After culturing hMSCs with KMs for four weeks, immunofluorescence staining was performed. On day 28, the staining showed positive expression of chondrogenic proteins (Figure 7). These findings demonstrate that KMs enhance the attachment of hMSCs, promoting the production of cartilage tissue-engineered constructs. The distinct coloration enabled clear identification and localization of these essential components within the cartilage matrix, providing valuable insights into the chondrogenic differentiation of hMSCs and the overall effectiveness of KMs in supporting cartilage tissue formation. The presence of these biomarkers underscores the successful maturation and development of the engineered cartilage constructs.

Figure 7. Immunofluorescence staining was performed to visualize chondro-specific biomarkers. Aggrecan was stained red, while type II collagen was stained green.

Discussion

Current approaches for treating chondral defects and osteoarthritis primarily rely on conservative management methods. However, no single treatment has proven effective in achieving long-lasting, complete functional repair of cartilage and bone tissues^[18]. The challenges faced in these treatments stem from the complex biological, biochemical, and biomechanical nature of the chondral unit, which is under constant pressure, frequent movement, and has intricate interactions with the subchondral bone. These difficulties are further compounded by the limited healing potential of articular cartilage, which has low regenerative capacity^[19]. Moreover, the age-related decline in the number and proliferative ability of endogenous stem cells, along with their reduced regenerative capacity, poses additional hurdles to developing safe and effective therapies. Given the significant economic and social burden of chondral-related conditions and the lack of effective long-term solutions, there is an urgent need to create innovative treatment strategies that can promote cartilage regeneration^[20]. A successful chondral tissue engineering construct requires three critical elements: a reliable and accessible source of cells, a three-dimensional scaffold that promotes cell attachment and encourages chondrogenesis, and a delivery system for chondrogenic signals. Choosing appropriate cell sources for osteochondral tissue engineering is a critical yet challenging task, as different cell types may have distinct differentiation potentials that can impact the success of chondrogenesis^[21].

Mesenchymal stem cells (MSCs) are a preferred choice due to their numerous benefits and the relative ease with which they can be obtained through minimally invasive methods^[22]. Studies have shown that MSCs have a strong potential to differentiate into various bone lineages^[23], and beyond bone, they can differentiate into cementum, periodontal-like tissues, and capillary-like structures^[24]. Additionally, MSCs possess immunomodulatory properties, which may provide antiinflammatory benefits and enhance immune tolerance during transplantation^[25]. Recently, small molecules, in addition to traditional growth factors, have emerged as potential modulators of MSC behavior. These small molecules can selectively influence stem cell differentiation and are more cost-effective, making them promising candidates for clinical applications. In this study, we aimed to enhance the regeneration of osteochondral defects using the small molecule KGN embedded in KMs. ECM-based biomaterials derived from native tissues have been widely studied in tissue engineering. One study demonstrated that ECM can promote cell recruitment, infiltration, and differentiation without the need for additional growth factors^[26].

KGN has been widely studied for its chondrogenic potential. It promotes the differentiation of mesenchymal stem cells (MSCs) by upregulating key chondrogenic markers such as SOX9, ACAN, COL2A1, and lubricin, leading to the formation of cartilage nodules rich in proteoglycans and collagen type II, typical of hyaline cartilage^{[27, 28]}. In addition to its pro-chondrogenic effects, KGN has been reported to elevate levels of tissue inhibitors of matrix metalloproteinases (TIMPs), suggesting chondroprotective properties. Preclinical studies further indicate that KGN can prevent cartilage degeneration and subchondral bone changes in osteoarthritis models^[29]. Moreover, KGN-integrated scaffolds have demonstrated efficacy in enhancing cartilage regeneration, partly through their ability to recruit endogenous cells without the need for exogenous cell transplantation^[30].

In this study, the novel KMs showed substantial potential in promoting cartilage regeneration. In vitro, MSCs adhered to, proliferated on, and differentiated in response to KMs. While the current results confirm that KMs promote the chondrogenic differentiation of hMSCs, a more nuanced understanding emerges when examining the temporal differences among specific marker genes. SOX9 and ACAN, typically associated with early to mid-stage chondrogenesis, peaked around day 14, indicating their role in the initiation and regulation of cartilage lineage commitment. Conversely, COL2A1 and COMP reached peak expression at later time points (day 21 or 28), reflecting their involvement in extracellular matrix deposition and tissue maturation. This staggered expression pattern underscores the sequential activation of chondrogenic programs and highlights the capacity of KMs to support the full spectrum of cartilage development, from lineage specification to matrix assembly. These findings emphasize the importance of temporally coordinated gene expression in engineered cartilage constructs and reinforce the potential of KMs as a scaffold-free delivery platform for small-molecule-induced cartilage regeneration.

Notably, the levels of dsDNA, sGAG, chondrogenic biomarkers, and relative mRNA expression were significantly upregulated in cells cultured with KMs, indicating enhanced ECM secretion.

These results suggest that KMs effectively support bone marrow MSCs in their chondrogenic differentiation, facilitating the regeneration of chondral defects. Compared to traditional growth factors, small molecules such as KGN offer several practical advantages that enhance their suitability for tissue engineering applications. Small molecules are generally more cost-effective, easier to synthesize and modify, and exhibit greater chemical and thermal stability, allowing for more consistent formulation and storage. Additionally, they typically have lower immunogenicity and can penetrate tissues more efficiently due to their low molecular weight. These properties contribute to better control of dosage and release kinetics when incorporated into delivery systems like microspheres. In this study, the use of KGN-loaded microspheres (KMs) leveraged these benefits, enabling sustained and bioactive release of the small molecule to drive chondrogenic differentiation. These findings support the broader potential of smallmolecule-based strategies as a robust and clinically translatable alternative to protein-based growth factors in regenerative medicine.

Considering the observed upregulation of chondrogenic markers such as SOX9, ACAN, and COL2A1, it is reasonable to hypothesize that the CBFβ–Runx1 signaling pathway may have contributed to the differentiation process in our model. This pathway, previously identified as a key downstream mechanism of kartogenin (KGN), has been shown to regulate mesenchymal stem cell differentiation into chondrocytes by stabilizing Runx1 and enhancing its interaction with CBFβ, ultimately promoting transcription of chondrogenic genes while suppressing hypertrophic transition^{[13, 14]}. Although we did not directly assess pathway activity in this study, the gene expression trends observed are consistent with this mechanistic model. Therefore, we speculate that the KGN-loaded microspheres (KMs) may exert their effects in part through activation of the CBFβ–Runx1 axis, which warrants further investigation in future mechanistic studies.

Conclusion

The composite KMs, composed of small-molecule drug-based ECM microspheres, demonstrated outstanding performance. These KMs not only exhibited excellent biocompatibility but also promoted rapid chondrogenesis, underscoring their potential for clinical applications. Our results suggest that the use of small molecules, as opposed to traditional growth factors, could improve the feasibility of osteochondral tissue engineering for regenerating chondral defects. However, a key limitation of this study is the absence of preclinical trials. Looking forward, future in vivo studies are essential not only to confirm the efficacy of KMs in cartilage regeneration but also to evaluate several critical parameters. First, the long-term release kinetics of KGN from the gelatin microspheres must be characterized to ensure sustained therapeutic effects over the course of cartilage repair. Second, the biodegradation behavior of the gelatin matrix in vivo needs to be systematically assessed to confirm that its by-products do not interfere with tissue remodeling or cell viability. Lastly, the potential immunogenicity of both the biomaterial and the encapsulated molecule should be carefully monitored, as host immune responses could impact scaffold integration, local inflammation, or systemic tolerance. Addressing these aspects in future preclinical models will be pivotal for translating KMs into safe and effective clinical applications.

Funding

This research received grants from Shin Kong Wu Ho-Su Memorial Hospital (2022SKHADR028).

Competing Interests

None