3D-printed biomimetic scaffolds with precisely controlled and tunable structures guide cell migration and promote regeneration of osteochondral defect

GelMA and lithium phenyl-2,4,6trimethylbenzoylphosphinate (LAP) were synthesized as previously [22, 28–31]. Proton nuclear magnetic resonance (¹H-NMR) was used to verify the degree of substitution of methacrylic anhydride (MA) in GelMA.

To prepare the hydrogel ink for the bilayer scaffold, the lyophilized GelMA was dissolved in phosphate-buffered saline (PBS) at 10 and 15% concentrations at 42 °C. KGN (200 μM) (Selleck Chemicals, Houston, TX, USA), photo-initiator LAP (0.1%), and UV-absorber phenol red (0.04%) were then added to prepare the ink for the upper layer. Solutions containing GelMA and 0.1% LAP were used as the precursor for the bottom layer.

The integrated bilayer scaffold was manufactured by a custom-made DLP-based 3D printer [32, 33]. The scaffold diameter was 5 mm with a 1 mm high upper scaffold and a 3 mm high bottom scaffold. The printed scaffold was immersed in PBS and stored at 4 °C for 14 d to evaluate its stability. The non-structural GelMA scaffold was prepared for comparison. For the test group, the bottom 3 mm scaffold was printed with GelMA solution, followed by printing the upper 1 mm scaffold with ink containing GelMA and KGN small molecules. Scaffolds with 100, 200, and 400 μm longitudinal apertures, respectively, were printed for mechanical tests. Scaffolds with different structures were printed for cell infiltration and migration tests.

To evaluate the macroscopic pore structure of the hydrogel scaffolds, the samples were stained with 4',6-diamidino-2-phenylindole (DAPI) and observed using an optical microscope. Briefly, DAPI was diluted 1:1000 and used as a dye. The printed upper scaffold was soaked in DAPI for 15 min, washed with PBS three times before they were observed and imaged under an Olympus upright two-photon confocal microscope (S6D, Olympus, Germany).

To evaluate the microstructure of the hydrogel scaffold, the sample was examined using a field emission scanning electron microscope (Nova Nano 450, Thermo FEI, Czech Republic) after gold–palladium coating.

Uniaxially unconfined compression tests were performed to evaluate the effect of the pores' size and distribution on the scaffolds' mechanical properties. A universal material testing machine (Zwick/Roell with a 5 kN sensor) was used to measure compressive force. The bottom lotus scaffolds were printed with 10% and 15% GelMA. The pore diameters of the scaffolds are 0, 100, 200, and 400 μm. During all experiments, the probe was pressed down at a constant speed of 1 mm min⁻¹ until the height of the scaffold was 65% of the initial height. The compressive modulus was calculated by linear fitting the stress–strain curve in the range of 40%–60% strain. The upper- and bottom-layer scaffolds with 200 μm pore size were printed separately and tested for compressive stress–strain measurements. The experiment process is the same as above. The compression modulus was calculated by linear fitting according to the strain range of 20%–40%.

In order to further prove that the scaffold could function normally under mechanical loading, consecutive cycle compression test was performed on the scaffold with 200 μm pore size. The scaffold was compressed until the strain reached 50% at a rate of 2 mm min⁻¹, and the specimen was relaxed at the same deformation rate. The above two steps were repeated as one whole cycle for ten times with no waiting time between cycles.

The cell counting kit-8 (CCK-8) assay was done to evaluate cell proliferation. First, to obtain the extracted liquid, 1 ml of cross-linked GelMA hydrogel was soaked in 10 ml of culture medium for 24 h. L929 cells (mouse fibroblast cells, Cell Bank of the Chinese Academy of Sciences) were seeded at a density of 1000 cells per well and incubated for 24 h at 37 °C. Afterward, the cell medium was renewed, and cells were further incubated with extract liquid for 1, 3, and 5 d. At every test point, the sample solution was removed, and cells were incubated with CCK-8 agent (1:10 diluted with the culture medium) at 37 °C for 2 h. The absorbance values were read at 450 nm using SynergyMx M5 (iD5, Molecular Devices, USA). Each test group has six replicate samples.

To assess cell viability on the hydrogel, a live/dead assay was performed. Briefly, 500 μl GelMA was added to a 3.5 cm dish and photo-crosslinked. C3H cells (C3H/10T1/2, mouse MSCs, Cell Bank of the Chinese Academy of Sciences) were seeded at a density of 5 × 10⁵ cells ml⁻¹ and cultured in a humidified atmosphere of 5% CO₂ at 37 °C for 1, 3 and 5 d. At each test point, a live/dead assay kit was administrated to the cells on hydrogel. Samples were observed under an Olympus upright two-photon confocal microscope.

2.5.1. Cell migration test in vitro

2.5.2. Cell migration test in vivo

Adult male New Zealand white rabbits (2.8–3.2 kg) were used to evaluate the function of the 3D-printed integrated bilayer scaffold in vivo. The treatment was conducted under the standard guidelines approved by the Ethics Committee of Zhejiang University (ZJU20210289). The surgery was performed under general anesthesia with an abdominal cavity and ear veins injection of 3% sodium pentobarbital (40 mg kg⁻¹). The skin was treated with iodophor in a sterile environment. A 1 cm long incision was made along the inside of the patellar ligament to expose the knee joint. A cylindrical shallow osteochondral defect (4 mm diameter and 2 mm depth) was made at the femoral trochlear by a dental drill. Forty defects were generated and randomly divided into five groups: blank group (B), non-structural scaffold group (N, the 3D-printed scaffold without any pore channels), 'lotus-' scaffold group (L, the 3D-printed scaffold with only 'lotus-' pores), 3D-printed structural scaffold group (P, the 3D-printed scaffold with both 'radial' and 'lotus-' pores), and P + KGN (the 3D-printed structural scaffold combined with KGN) group. The scaffolds were implanted into the defect, and then the tissue was disinfected with iodophor and sutured immediately. For the blank group, the defect was sterilized and sutured directly. After the operation, the experimental animals were intramuscularly injected with penicillin at 400 000 U d⁻¹ for 3 d. Samples were collected on days 7 and 42 after surgery.

For the full-thickness osteochondral defect model, the operation process was the same as above, while the defect size was 5 mm in diameter and 4 mm deep. Joints were randomly divided into four groups: blank group (B), non-structural group (N), 3D-printed structural group (P), and 3D-printed bilayer scaffold with KGN in the upper layer group (P + KGN). There are 12 joints in each group. Animals were sacrificed at 8 and 16 weeks postoperatively, and joint samples from each group were collected for further analysis.

The harvested joint samples were immediately photographed and then assessed according to the general evaluation standards of the International Cartilage Repair Society (ICRS), specifically including the degree of defect repair, integration to the border zone, and macroscopic appearance [34]. The samples were fixed in 4% paraformaldehyde (PFA) for 2 days and decalcified with 10% ethylene diamine tetraacetic acid (EDTA) for 8 weeks. Then, the samples were dehydrated in gradient ethanol, hyalinized in xylene, embedded in paraffin, and sectioned to slices with a thickness of 10 μm using a microtome. Repair of the defect was assessed by H&E and safranin-O/fast-green (SO/FG) staining.

To test the compressive mechanical properties of the repaired cartilage at 16 weeks post-operation (n = 3 per group), the universal material testing machine (Instron-5543 with a 1 kN sensor) was used.

After the rabbit joint samples were harvested at 16 weeks post-operation, the joints were investigated by micro-computed tomography (micro-CT) scanning imaging system (SCANCO Medical AG, μCT 100, Bassersdorf, Switzerland) provided by Hangzhou Yuebo Biological Company. A 3D 180° rotation scan was performed at a rotating angular velocity of 0.8° s⁻¹ to reconstruct the cylindrical regeneration area with a layer thickness of 18 μm using 60 kV scanning voltage. Quantitative analysis, including bone volume/tissue volume (BV/TV), trabecular number (Tb.N), and trabecular separation/spacing (Tb.Sp) was obtained.

GraphPad Prism software was used for statistical analysis. The difference between the two groups was analyzed by t-test. One-way analysis of variance with Tukey post-test was used when there were more than two variables. All results were presented as mean ± standard deviation. Statistical significance was defined as p values < 0.05.

In this study, as shown in scheme 1, the bilayer-integrated hydrogel scaffolds were successfully fabricated via DLP 3D printing to accurately mimic the osteochondral architecture. To boost the efficacy of the cartilage repair, the growth factor KGN was added to the upper layer of the bilayer scaffolds.

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Prior to the scaffold fabrication, as first reported by Van Den Bulcke et al [35], a portion of gelatin was modified with MA to produce GelMA with a 34% degree of substitution (figure 1(A)). ¹H-NMR analysis was used to observe lysine conjugation with MA. The lysine methylene proton (2H) peaks around 2.9 ppm (figure 1(A-b)) decreased significantly in the spectrum of GelMA, exhibiting complete interlocking of the lysine moiety with MA. Further, the appearance and enhancement of the peaks corresponding to the acrylic protons (2H, around 5.3–5.7 ppm, figure 1(A-a)) and the methyl protons (3H, around 1.8–2.1 ppm, figure 1(A-c)) of grafted methacrylamide groups were observed in GelMA samples [36]. This confirmed the successful conjugation to MA.

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The Fourier transform infrared spectroscopy (FTIR) spectra (figure 1(B)) showed that the –OH stretching, N–H stretching and N–H bending, C–H stretching and C–H bending vibrations of the GelMA hydrogel are at ≈3442, ≈1637, ≈1431, ≈2930, and ≈1451 cm⁻¹, respectively, indicating successful modification of gelatin to GelMA.

Then, the bilayer scaffold was 3D-printed by DLP technology. Figure 1(D-a, b) showed the top and side views of the 3D-printed hydrogel scaffold, respectively. It exhibited rounded edges and clear boundaries. The scaffold immersed in PBS can maintain its overall shape for at least 14 d (supplementary figure 1). Meanwhile, the scaffold exhibited overlap of reloading and unloading curves across ten cycles, suggesting highly reversible compressibility and mechanical stability (supplementary figure 2). The surface morphology of the 'lotus- and radial-' scaffold (figure 1(E)) was further characterized by a two-photon confocal laser scanning microscopy, and the lotus-distribution pores with a diameter of 200 μm can be observed. Furthermore, as shown in figure 1(F), the radial-distribution pore structure of the upper scaffold and the abundant micropores inside the scaffold could be observed by scanning electron microscopy (SEM).

To explore the structural flexibility of the scaffold, we prepared bilayer scaffolds with different longitudinal pore sizes: diameters of 100 μm, 200 μm, and 400 μm. The microscopy results (figure 2(A)) and the SEM images (supplementary figure 3) showed that the size of the channels can be precisely controlled.

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Using this printing strategy, we prepared bilayer scaffolds with different concentrations of GelMA hydrogels. The SEM morphology (figures 2(B-a) and (C-a)) showed that both 15% and 10% GelMA hydrogels had high porosity and precisely defined interconnected porous structures. We analyzed the relationship between the load and compression deformation of two GelMA concentrations (10% and 15%) with different pore sizes (figures 2(B-b, c) and (C-b, c)). The compression modulus of 15% GelMA hydrogel scaffolds with pore sizes of 0, 100, 200, and 400 μm were 388.0 ± 17.52, 344.3 ± 16.5, 302.0 ± 15.00, and 203.3 ± 10.41 kPa, respectively; while the compression modulus of 10% GelMA hydrogel scaffolds with pore sizes of 0, 100, 200, and 400 μm were 178.0 ± 7.211, 157.3 ± 6.429, 139.0 ± 5.568, and 107.3 ± 3.055 kPa, respectively. 3D-printed integrated bilayer scaffolds fabricated with 15% GelMA had higher mechanical properties compared to those printed with 10% GelMA. The scaffold with a pore size of 400 μm had the highest specific surface area up to 4.338 m² m⁻³. The non-porous scaffold had a higher compressive strength (388.0 ± 17.5 kPa), but its specific surface area (1.895 m² m⁻³) was significantly lower than the other scaffolds. Considering the balance between mechanical property and surface area, a 15% GelMA hydrogel with a pore size of 200 μm (302.0 ± 15.0 kPa) was selected for further study. This hydrogel exhibited good mechanical properties and a relatively large specific surface area (2.861 m² m⁻³) (supplementary table 1), providing mechanical support and enough surface for cell attachment.

After determining the composition and pore size of the scaffold, the compressive mechanical properties of the upper and bottom layers of scaffolds were further investigated (figure 2(D)). The bottom layer of scaffold (558.4 ± 22.9 kPa) exhibited significantly higher compressive mechanical properties compared with the upper layer of scaffold (339.3 ± 9.1 kPa) (***p < 0.001). Therefore, our strategy provided a functional bilayer scaffold with layer-specific strength, contributing to mimicking the anisotropy of natural osteochondral tissue.

Next, bovine serum albumin (BSA) was chosen as the model drug to test its release profile from scaffolds. As shown in figure 2(E), BSA was released slowly and steadily, reaching 66.23 ± 3.31% on day 7. This suggested that our scaffold fabricated with GelMA hydrogel can slowly release small molecules or drugs for synergetic effect.

The CCK8 assay was performed to assess the cytotoxicity of GelMA hydrogel when cells were cultured in hydrogel extract liquid (figure 3(A)). The absorbance values of cell viability increased over 5 d in both groups. The results affirmed a significant rise in cell numbers over time, comparable to the control group. The result indicated that the GelMA hydrogel did not inhibit cell proliferation.

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To further evaluate the cell compatibility, the live/dead staining test was performed (figures 3(B) and (C)). During the 7 day culture, almost no C3H cells were stained with red fluorescence, which indicated that GelMA hydrogel had no obvious toxicity. On the 7th day, the cell survival rate reached 99.63 ± 0.09%. It proved that the GelMA hydrogel had great biocompatibility and did not affect cell proliferation. As shown in figure 3(D), the cytoskeleton of C3H cells adhering to the hydrogel was stained in green with fluorescein isothiocyanate after culturing for 1, 4, 7 d, which showed that the cells spread well on the hydrogel. Taken together, the GelMA hydrogel was suitable for cell adhesion, spreading, and proliferation.

We further evaluated the effect of the structure of 3D-printed hydrogel scaffolds on cell migration and infiltration in vitro and in vivo (figure 4(A)). In the in vitro experiment, C3H cells stained with DiI were allowed to adhere to the surface of culture dishes for 24 h before the 3D-printed structural (P) and non-structural (N) hydrogel scaffolds were separately placed in the dishes. Confocal microscopy was used to observe cell migration into the scaffold after 1, 3, and 5 d. 3D reconstructions of the confocal images (figure 4(B)) showed a significant increase of cells in the P group over time. Figure 4(C) showed cell distribution in hydrogel scaffolds after 1 d culture. Quantitative analysis (figure 4(D)) showed that the number of migrating cells in the P group (29.2 ± 6.6) was approximately four times greater than the N group (7.6 ± 0.1) (**p< 0.01).

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We further implanted the 3D-printed structural and non-structural hydrogel scaffolds into rats subcutaneously to evaluate the cell infiltration in vivo (figures 4(E)–(G)). 3D reconstructed images showed that 3 d after implantation in vivo, the number of cells in the P group (276.8 ± 12.1) was approximately 1.4 times higher than that in the N group (225.0 ± 11.1) (**p < 0.01). 5 d after implantation, the number of cells in the P group (554.3 ± 22.2) was still significantly higher than that in the N group (457.8 ± 20.1) (^*** p < 0.001). The above results confirmed that the 3D-printed structural hydrogel scaffold effectively promoted cell migration or infiltration into the scaffold.

In addition, we subcutaneously implanted different structural scaffolds in rats to compare their effects on cell infiltration. The hydrogel scaffolds were harvested 5 d after implantation. DAPI staining showed that both the number of cells and depth of migration were significantly enhanced when both 'lotus-' and 'radial-' pores were present (figures 5(A)–(C)). Quantitative analysis (figure 5(B)) showed that the maximum vertical infiltration distances of cells were 279.0 ± 56.9, 222.3 ± 63.4, 141.3 ± 68.0, and 25.6 ± 14.6 μm in the 3D-printed structural (P), only 'lotus-' (L), only 'radial-' (R), and non-structural (N, without any pores) groups, respectively. The maximum vertical migration distance of cells in the P group was significantly higher than in groups R and N. However, it was not significantly different from that in the L group, indicating the critical role of the 'lotus-' pores on the longitudinal migration of cells. Figure 5(C) shows that the number of cells in the P group (354.5 ± 12.2) was significantly higher than that in the L (223.5 ± 64.0), R (120.5 ± 43.9) and N (65.6 ± 26.9) groups. It demonstrated that the scaffold with both 'lotus- and radial-' distribution pore structures provided the best support for cell infiltration and migration into the scaffold. Next, we further observed that the cells infiltrated the scaffold mainly along the 3D-printed pore structures instead of elsewhere. Therefore, the scaffolds were sliced horizontally and vertically in two different directions (figure 5(D) and supplementary figure 4). The results of H&E staining of the P group (figure 5(D)) showed that, in both the horizontal and vertical directions, there were indeed more cells and greater migration depth in the pore structure (marked by the black dashed line) than that outside the pore area. Overall, the 3D-printed scaffold with 'lotus- and radial-' distribution pore structures facilitated cell migration to the interior of the scaffold along the established pores.

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Following the results of the in vitro experiments, we evaluated the feasibility of cartilage repair using a 'lotus- and radial-' 3D-printed structural scaffold in a rabbit cartilage defect model (diameter = 4.0 mm, depth = 2 mm). Five different groups were investigated: blank group, non-structural group (N, the 3D-printed scaffold without any pore channels), 'lotus-' scaffold group (L, the 3D-printed scaffold with only 'lotus-' pores), 3D-printed structural scaffold group (P, the 3D-printed scaffold with both 'radial' and 'lotus-' pores), and P + KGN (the 3D-printed structural scaffold combined with KGN) group. After 7 d and 6 weeks, the rabbits were euthanized to harvest the repaired knee joints for gross morphological observation and histological analysis. There was no mortality or infection in any animal after the operation.

From the representative histological evaluation of the repaired regions (figure 6(A)), we observed that none of the hydrogel scaffolds in the cartilage defect were degraded 7 d after surgery. Even so, there was better integration of the implanted hydrogel scaffolds in the P group with 'radial-' pores compared to the blank and N group, possibly because the 'radial-' pores facilitated the horizontal migration of cells.

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In order to evaluate the therapeutic effects of different scaffolds, we analyzed the gross specimens at 6 weeks after surgery according to the ICRS macroscopic assessment scoring criteria (figures 6(B) and (C)). We observed that the defect site in the P + KGN group was relatively flat and well-repaired. Although the defect was still visible, its boundary with the surrounding area was blurred. The ICRS scores of the P + KGN group (10.67 ± 0.58) were significantly higher than the blank group (1.44 ± 0.29) and the N group (2.44 ± 0.91) (**p < 0.01). The P group, with a score of 8.56 ± 1.47, still had partial defects. On the contrary, there was no significant difference in the repair effect between the N and L group (6.22 ± 1.79) compared with the blank group (p > 0.05), where a clear boundary with the surrounding healthy cartilage could be seen. Figure 6(B) also showed images of H&E and SO/FG staining of the repaired cartilage after 6 weeks. Cluttered fibrous tissue was regenerated in the blank group, and the defect was not filled. In the N and L group, most of the hydrogel remained in the defect area, preventing the growth of new tissue. In contrast, in the P group, the defective cartilage site not only formed neo-tissue but also appeared SO staining positive area, indicating that the regeneration of the P group was started yet semimature and incomplete. Notably, the P + KGN group showed the best repair performance, with a large number of SO staining positive areas and typical round articular chondrocytes. These results showed that 3D-printed scaffolds with 'lotus- and radial-' distribution pores promoted cartilage regeneration, and the addition of KGN accelerated the regeneration process.

Next, we evaluated the osteochondral regenerative capacity of the 3D-printed integrated bilayer scaffolds. Four different groups were investigated: blank group, non-structural group (N, the 3D-printed scaffold without any pore channels), 3D-printed structural group (P, the 3D-printed bilayer scaffold with both 'radial' and 'lotus-' pores), and P + KGN (the 3D-printed bilayer scaffold combined with KGN in the upper layer) group. Full-thickness osteochondral defects (diameter = 5.0 mm, depth = 4.0 mm) were made on the bilateral knee joints of rabbits (figure 7(A)). At sacrifice (8, 16 weeks post-implantation), no obvious infection or implantation response was found in the repaired tissue of the four groups of rabbits. Then, the regenerative capability of osteochondral tissues was assessed by gross morphological observations, histologic analyses, micro-CT, and mechanical tests.

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As shown in figure 7(B), the gross morphological images showed that the osteochondral defects in the blank group exhibited a heavily cavitated surface at both 8 and 16 weeks. Compared to 8 weeks, the N group showed some regeneration at the defect site at 16 weeks but still exhibited a poor repair outcome. Blood infiltration could be observed as red areas in the P group at 8 weeks. At 16 weeks, the P group showed improved regeneration and integration, but the irregular surface still differed from natural cartilage. The P + KGN group exhibited optimal osteochondral repair as early as week 8, with the defects at 16 weeks being replaced with typical cartilage-like tissues. At 8 weeks, the ICRS scores of the blank, N, P, and P + KGN groups were 2.222 ± 0.7698, 2.889 ± 0.3849, 5.556 ± 1.018 and 6.556 ± 1.018, respectively. At 16 weeks, the ICRS scores were 6.778 ± 1.836, 8.778 ± 0.7698, 9.556 ± 0.3849, and 9.778 ± 0.5092 in the blank, N, P, and P + KGN groups, respectively. Based on macroscopic observations and the ICRS scoring system, the P + KGN group showed better repair than the N and P groups, especially at 8 weeks (figures 7(B)–(D)).

Micro-CT was used to assess the subchondral bone regeneration at 16 weeks after surgery. Figure 7(E) shows representative images of each group after 3D reconstruction. The formation of new subchondral bone tissue was most pronounced in the P + KGN group, where the defect area was almost completely filled. The P group also showed some improvement, but there was a significant cavity in the defect area. Admittedly, the cavity in the N group was larger, while the blank group only had very little new subchondral bone formation along the edges of the defect area. Quantitative analysis of the newly formed subchondral bone in the defect area confirmed the micro-CT reconstruction results (supplementary figure 5). The BV/TV value verified that the bone repair effect of the P + KGN group was significantly better than that of the blank group (*p< 0.05). Compared to the other groups, the subchondral bone at the defect site was well repaired with or without KGN. These results indicated that the 3D-printed structural scaffolds had the potential to promote the regeneration of the subchondral bone.

We further evaluated the mechanical properties of the articular cartilage samples at 16 weeks postoperatively (figures 7(F) and (G)). The results showed that the reduced modulus of the cartilage treated with P + KGN (6.32 ± 0.45 GPa) was significantly greater than both the N group (1.30 ± 0.18 GPa) and the blank group (0.37 ± 0.02 GPa) (****p< 0.0001). There was no significant difference between the P + KGN group and the P group (5.59 ± 0.34 GPa) (p > 0.05).

Histological analysis of regenerated osteochondral tissue after 8 and 16 weeks was performed by H&E and SO/FG staining in figure 8. At 8 weeks, the defect area in the blank group was only filled with fibrous tissue, and a small amount of matrix staining was observed. In other groups, the implanted hydrogels were still visible at 8 weeks. Regeneration of cartilage and bone was delayed due to the slow degradation of the hydrogel. All groups had more tissue fillings and reduced defect area at 16 weeks compared to 8 weeks. In the blank group, the regenerated surface was rough and unsmooth, with massive fibrous tissue formation. In the N group, despite most of the defect sites being filled, we observed many fat-like structures. The surface cartilage layer of the P group was thin, and the regeneration of cartilage was discontinuous. Notably, the thickness and surface smoothness of regenerated cartilage in the P + KGN group were better than those in the P group. A large number of newly formed, regularly arranged hyaline chondrocytes can be observed. In addition, the cartilage–bone interfaces of the P + KGN group were more regular than those of the P group. The results showed that the bilayer printed scaffolds releasing KGN could further promote the repair of the cartilage layer in osteochondral defect repair.

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