A Bioglass‐Poly(lactic‐co‐glycolic Acid) Scaffold@Fibrin Hydrogel Construct to Support Endochondral Bone Formation

Abstract

Bone tissue engineering using stem cells to build bone directly on a scaffold matrix often fails due to lack of oxygen at the injury site. This may be avoided by following the endochondral ossification route; herein, a cartilage template is promoted first, which can survive hypoxic environments, followed by its hypertrophy and ossification. However, hypertrophy is so far only achieved using biological factors. This work introduces a Bioglass‐Poly(lactic‐co‐glycolic acid@fibrin (Bg‐PLGA@fibrin) construct where a fibrin hydrogel infiltrates and encapsulates a porous Bg‐PLGA. The hypothesis is that mesenchymal stem cells (MSCs) loaded in the fibrin gel and induced into chondrogenesis degrade the gel and become hypertrophic upon reaching the stiffer, bioactive Bg‐PLGA core, without external induction factors. Results show that Bg‐PLGA@fibrin induces hypertrophy, as well as matrix mineralization and osteogenesis; it also promotes a change in morphology of the MSCs at the gel/scaffold interface, possibly a sign of osteoblast‐like differentiation of hypertrophic chondrocytes. Thus, the Bg‐PLGA@fibrin construct can sequentially support the different phases of endochondral ossification purely based on material cues. This may facilitate clinical translation by decreasing in‐vitro cell culture time pre‐implantation and the complexity associated with the use of external induction factors.

Scaffolds for endochondral ossification should support chondrogenesis, hypertrophy, and osteogenesis. This article presents a Bioglass‐Poly(lactic‐co‐glycolic)acid@fibrin hydrogel designed to support these stages sequentially. Chondrogenesis‐primed mesenchymal stem cells seeded in the fibrin gel became hypertrophic and started osteogenesis once they reached the Bioglass‐Poly(lactic‐co‐glycolic)acid scaffold core, without external induction factors. This transformation based on material cues only may facilitate clinical translation.

1. Introduction

Bones can be lost due to injury or disease.^{[ 1 ]} Current methods of treatment include the replacement of the lost bone with a piece of bone from the same person (autograft) or from a donor (allograft). These grafting procedures have several limitations, like non‐union, graft rejection, availability of donors, and risk of infections.^{[ 1} , ² , ^{3 ]} Bone tissue engineering with synthetic bone grafts can be an alternative to current grafting techniques,^{[ 4 ]} especially in large bone defects. In this approach, scaffolds loaded with cells support and guide bone formation and, if degradable, disappear as the regenerated bone grows.^{[ 5} , ⁶ , ^{7 ]}

The two main approaches to bone tissue engineering are based on the two routes of embryonic bone development and fracture repair: direct ossification and endochondral ossification.^{[ 8 ]} During direct ossification, mesenchymal stem cells directly differentiate into osteoblasts and form bone. This process forms the flat bones of our body, and it also occurs during fracture repair at the bone collar (a layer of compact bone surrounding the mid‐shaft of the bone).^{[ 8 ]} On the other hand, endochondral ossification forms bone through a cartilage intermediate. This process forms the long bones of our body and also occurs in the avascular space at the centre of the fracture site during fracture healing.^{[ 8 ]}

Most bone tissue engineering approaches follow direct ossification, due to its simplicity. However, this method often fails due to hypoxia‐induced cell death: mesenchymal stem cells (MSCs) and osteoblasts require oxygen to survive, but at the fracture site, blood vessels are damaged, especially in large defects.^{[ 9} , ^{10 ]} This is usually compensated by adding external factors like exosomes, vascular endothelial growth factor (VEGF), or other growth factors to promote vascularization and osteogenesis, escalating the costs and increasing the complexity of scaffold production.^{[ 11} , ^{12 ]} As the chondrocytes can survive under hypoxic conditions, the endochondral ossification approach can overcome the limitations associated with direct ossification^{[ 13} , ¹⁴ , ^{15 ]} and hence could be a more effective route of ossification than direct ossification, especially for large defects.

During endochondral ossification, MSCs come together to form a cartilage model of the bone to be formed in a process known as mesenchymal condensation, followed by the differentiation of MSCs into chondrocytes (chondrogenesis). The chondrocytes proliferate and as they approach the center of the cartilage model with time, they reach a terminal maturation stage known as hypertrophy, where the chondrocytes enlarge, start calcifying the matrix, and induce vasculature formation hence providing a suitable environment for new bone formation.^{[ 8 ]}

Recently many researchers attempted to regenerate bone through endochondral ossification using cell pellets (i.e., cell aggregates formed by centrifugation of cell suspensions), hydrogels and porous scaffolds.^{[ 16 ]} MSC pellets mimic the mesenchymal condensation step of endochondral ossification^{[ 13} , ¹⁷ , ¹⁸ , ¹⁹ , ^{20 ]} and have promoted endochondral bone formation in animal models.^{[ 19} , ^{21 ]} However, an excessively large concentration of cells would be needed to create pellets for large defects in large animal models and humans; thus, clinical translation of cell pellets is limited. Hydrogels have high cell loading capacity and can be made of fibers that mimic the cartilage matrix, hence promoting chondrogenesis.^{[ 22} , ²³ , ²⁴ , ²⁵ , ²⁶ , ^{27 ]} Hydrogels have promoted endochondral bone formation in several previous studies.^{[ 22} , ²³ , ²⁴ , ²⁵ , ²⁶ , ²⁷ , ²⁸ , ^{29 ]} Their drawbacks however include limited perfusion of media, poor mechanical strength, and poor cell‐cell contact.^{[ 30 ]} Hydrogels have also shown incomplete hypertrophic differentiation of chondrocytes; this was attributed to the nature of hydrogels mimicking the cartilage matrix, hence preserving the chondrogenic phenotype.^{[ 31} , ^{32 ]} Polymeric composites are being widely used as porous scaffolds for bone tissue engineering,^{[ 33 ]} and the porous scaffolds have also been shown to support endochondral bone formation in vivo.^{[ 14} , ¹⁶ , ³⁴ , ³⁵ , ^{36 ]} However, their low cell loading efficiency and the possible need for rotary cell culture limit their application; also, their stiffness promotes more direct ossification than endochondral ossification.^{[ 16} , ³⁷ , ³⁸ , ^{39 ]}

To mimic the endochondral ossification process, it is important to support the three main stages of the process, namely chondrogenesis, hypertrophy and bone formation. As cartilage and bone are two very different tissues with different requirements, a scaffold with two components to sequentially support the cartilage and bone tissues could be a promising approach toward bone regeneration through endochondral ossification.^{[ 40} , ^{41 ]} Some recent approaches involve using hydrogel/polymer composite scaffolds, but this is done to improve the strength of the hydrogel to support endochondral bone formation and not to support the sequential formation of the two tissues.^{[ 42} , ^{43 ]}

A scaffold to support endochondral ossification should also be able to induce hypertrophy as this is the crucial transition stage from cartilage to bone.^{[ 44 ]} Previous studies mostly induced hypertrophy in vitro by adding hypertrophy‐inducing growth factors to the media.^{[ 45 ]} This led to long in vitro cultures, including 21 days for chondrogenic differentiation and 1 to 2 weeks of hypertrophic induction, which increases the cost and complexity for clinical translation. A recent approach tried to reduce the in vitro culture duration by adding growth factor‐releasing particles to the scaffold.^{[ 45 ]} In this study, two growth factors, transforming growth factor (TGF‐ß) to induce chondrogenesis and bone morphogenetic protein‐2 (BMP‐2) to induce hypertrophy, were used to eliminate the need for in vitro chondrogenic and hypertrophic priming.^{[ 45 ]} The use of biological factors led to successful cell differentiation, but it involves a strict regulatory process and also escalates costs.^{[ 45} , ^{46 ]}

Previous studies have shown the potential of material properties, especially stiffness, to direct the differentiation of MSCs.^{[ 16} , ³⁷ , ³⁸ , ^{39 ]} These studies show that stem cells have mechanoreceptors that sense the stiffness of their environment, and they differentiate differently depending on how closely the stiffness of substrates resembles the stiffness of native extracellular matrix.^{[ 16} , ³⁷ , ³⁸ , ^{39 ]} For example, MSCs cultured in polyacrylamide gels matching the stiffness of cartilage undergo chondrogenesis,^{[ 47 ]} whereas polyacrylamide gels with stiffness matching bone tissue were able to direct osteogenic differentiation of MSCs.^{[ 48 ]} Stiff scaffolds were also able to induce hypertrophy in chondrocytes in previous studies.^{[ 32 ]} To the best of our knowledge, the potential of using just material cues to induce hypertrophy and osteogenesis in endochondral bone regeneration has not been explored yet.

We hypothesized that a construct including a fibrin hydrogel containing MSCs to support chondrogenesis, infiltrating and encapsulating a porous Bioglass‐Poly(lactic‐co‐glycolic acid) (Bg‐PLGA) composite scaffold to support hypertrophy and bone formation could sequentially support the endochondral ossification process, and that hypertrophy can be induced in the scaffold solely relying on material cues. This construct will be called Bg‐PLGA@fibrin hereafter to remind the reader of the presence of a Bg‐PLGA core surrounded and encapsulated by a fibrin hydrogel. As schematically shown in Figure  1 , we expect that after inducing chondrogenesis in the MSCs in the fibrin hydrogel, the cells will proliferate and slowly degrade the gel. As they degrade it, they will approach the stiff Bg‐PLGA scaffold; the stiffness of this scaffold and the presence of Bg (a mineralization promoter) will induce hypertrophy with no need for external factors.

Figure 1.

Bg‐PLGA@fibrin construct design. Porous Bg‐PLGA scaffold (A). Bg‐PLGA@fibrin construct showing fibrin gel with MSCs induced into chondrogenesis infiltrating and encapsulating the porous Bg‐PLGA scaffold (B). According to our hypothesis, the chondrocytes degrade the fibrin gel and become hypertrophic upon reaching the Bg‐PLGA scaffold at the core of the gel (C).

To test this hypothesis, we culture MSCs in Bg‐PLGA@fibrin and, as control samples, fibrin hydrogels and porous Bg‐PLGA scaffolds, in chondrogenesis inducing medium or chondrogenic differentiation medium for 21 days; we then switch to basal medium (i.e., no growth factors or induction factors) and prolong the culture for an extra 14 days. We study how the different substrates affect cell differentiation, matrix formation and mineralization in these conditions.

2. Experimental Section

2.1. Sample Preparation

The Bg‐PLGA scaffold core was prepared using a method reported in our previous study with some modifications.^{[ 49 ]} Briefly, a solvent casting and porogen leaching technique using paraffin microspheres of diameter 250–355 µm as the porogen was used. To prepare each scaffold, paraffin microspheres were placed in a syringe, filling half of it, and incubated at 52 °C for 3 h to make the microspheres partially melt and contact each other. This improves the pore interconnectivity upon porogen leaching.

A Bg‐PLGA suspension was prepared by adding 45S5 Bg powder (NovaBone Products, LLC, Alachua, FL) of particle size <25 µm to a 15 wt% PLGA (75:25 lactic acid: glycolic acid, molecular weight 66 000–107 000, Sigma Aldrich) solution in dichloromethane (Sigma‐Aldrich) under continuous stirring for 30 min, at a weight ratio of 1:1 of Bg: PLGA. 400 µL of the suspension was cast inside the syringe containing paraffin microspheres, and infiltrated into the microsphere mold with the help of the syringe piston pushing the suspension into the mold. The resulting materials were then air‐dried and cut into discs of height 2.5 mm and diameter 4.6 mm. These were then placed in 150 mL of CitriSolv (Decon laboratories), a citrus‐based solvent, and stirred for 48 h to leach out the paraffin microspheres. The obtained Bg‐PLGA scaffolds had both macropores created by paraffin sphere removal and micropores created by the partial degradation of the PLGA matrix induced by CitriSolv.^{[ 49 ]} The scaffolds were vacuum dried and stored under vacuum until further use.

To prepare the Bg‐PLGA@fibrin samples, Bg‐PLGA scaffolds were wetted by immersion in 100% ethanol (Fisher Scientific) for 20 min, followed by immersion in phosphate‐buffered saline (PBS) (gibco) for another 20 min (Figure  2A,B). The wetted scaffolds were placed in a syringe and restrained in position with the help of needles (Figure 2C), and a 1:1 solution of 50 mg mL⁻¹ fibrinogen (Fisher Scientific) in 0.9% NaCl (Sigma Aldrich) and 10 U mL⁻¹ (enzyme unit mL⁻¹) thrombin (Sigma Aldrich) solution prepared in 40 mm CaCl₂ (Sigma Aldrich) (50 µL total) was added to the syringe. The infiltration of the solution was facilitated by the vacuum created by pulling the syringe piston down (Figure 2C). Then the syringe was placed at 37 °C for 1 h to allow for gelation, leading to the formation of the Bg‐PLGA@fibrin constructs with fibrin gel infiltrating and encapsulating the porous Bg‐PLGA scaffold core (Figure 2D).

Figure 2.

A schematic representation of the preparation of the Bg‐PLGA/fibrin. Pre‐wetting of Bg‐PLGA porous scaffold by immersion in ethanol followed by immersion in PBS (A,B). Vacuum‐assisted infiltration of fibrinogen and thrombin mixture into the pores of the Bg‐PLGA scaffold restrained with needles in a syringe (C). A photograph of the Bg‐PLGA@fibrin obtained after gelation (D).

The fibrin gel samples were prepared by mixing equal volumes of 50 mg mL⁻¹ fibrinogen solution in 0.9% NaCl and 10 U mL⁻¹ thrombin solution in 40 mm CaCl₂ (50 µL total) and allowing them to gel at 37 °C for an hour.

2.2. Material Characterization

Three replicates each of Bg‐PLGA scaffold, fibrin gel, and Bg‐PLGA@fibrin samples were visualized using a scanning electron microscope (SEM) (Hitachi FlexSEM 1000 II) at an acceleration voltage of 10 kV and a working distance of 10 µm, and the composition was analyzed by multipoint analysis and elemental mapping using integrated energy‐dispersive x‐ray spectroscopy (EDS) (Hitachi FlexSEM 1000 II) at an accelerating voltage of 10 kV and a working distance of 5 µm. Samples with cells (three replicates per type) were analyzed by SEM after being fixed overnight in 2.5 wt% glutaraldehyde solution (Sigma‐Aldrich) at 4 °C followed by serial dehydration by immersion in 50%, 70%, 90%, 95%, and 100% ethanol (20 min each step), and finally drying in a Balzers CPD 030 Critical Point Drier. All the samples were coated with 5 nm of carbon using a Cressington 208 Carbon Evaporator before SEM analysis.

The 3D architecture of the Bg‐PLGA composites was analyzed using a SkyScan 1172 scanner micro‐Computed Tomography (micro‐CT) instrument at 34 kV and 210 µA using 3.0 µm pixel resolution, three‐frame averaging and no filter. Three replicates of the scaffold were analyzed. NRecon software, SkyScan was used to reconstruct the sections; CTAn software, SkyScan was used to create 3D images; CTVol software, SkyScan was used to visualize the 3D scaffold. CTAn software was used to calculate the total open porosity of the scaffolds with three replicates of Bg‐PLGA scaffolds.

The composition of the Bg‐PLGA scaffold was analyzed by thermogravimetric analysis using TA instruments Q500 where the samples were heated up to 600 °C in a nitrogen atmosphere at a rate of 10 °C min⁻¹.

Cyclic compression tests on three replicates each of the Bg‐PLGA scaffold, fibrin gel and Bg‐PLGA@fibrin samples were performed using TA instruments Q800 dynamic mechanical analyzer (DMA). The scaffolds were wetted by immersion in 100% ethanol (Fisher Scientific) and phosphate buffered saline (PBS) (gibco) for 20 min before analysis. The fibrin gels and the Bg‐PLGA@fibrin were stored in PBS until analysis. The tests were performed at room temperature with a static load of 0.01 N at 0.3% strain, and a frequency sweep method was used where the frequency varied from 0.1 to 40 Hz. During these tests, the storage and loss modulus and stiffness were recorded as a function of increasing frequency from the stress‐strain curves obtained from the load‐displacement curves automatically by the software.

2.3. In Vitro Bioactivity

In vitro bioactivity of the samples was evaluated using simulated body fluid (SBF) immersion tests.^{[ 50 ]} SBF was prepared according to the recipe provided in a previous study.^{[ 50 ]} The samples were immersed in SBF to obtain a mass/volume ratio of 1 g mL⁻¹ in SBF for 1, 4, and 7 days (three replicates per type per time point), and the SBF was refreshed every 3 days. After each time point, the samples were removed, rinsed with distilled water, dipped in liquid nitrogen, dried in a freeze dryer (BenchTop K freeze‐dryer, USA) overnight, and stored under vacuum until further characterization.

The surface elemental composition of the samples before and after SBF immersion was analyzed by X‐Ray photoelectron spectroscopy (XPS) using a Thermo Scientific Kα spectrometer with an Al Kα X‐ray source (1486.6 eV, 0.843 nm) and a spot size of 400 µm. Three replicates of the Bg‐PLGA/fibrin, Bg‐PLGA scaffold and fibrin gel were analyzed, taking 6 points per sample, giving a total of 18 points per sample type.

Samples before and after SBF immersion, three replicates per condition, were analyzed by Raman spectroscopy, where the spectra were collected with a Bruker Senterra confocal Raman spectrophotometer with a 785 nm diode laser connected to an Olympus optical microscope. A 20× objective was used to collect the spectra between 400 and 1800 cm⁻¹ at a 3.5 cm⁻¹ resolution and 100 mW laser power. An integration time of 60 s was used and 2 co‐additions were collected for each spectrum. Data was analyzed using OPUS software (OPUS 7.0, Bruker).

2.4. Cell Loading

To prepare the Bg‐PLGA@fibrin samples, the Bg‐PLGA composite was pre‐wetted by immersing in 100% ethanol, followed by immersion in phosphate‐buffered saline for 20 min each. These pre‐wetted Bg‐PLGA scaffolds were immobilized in 1 mL syringes using a method similar to that shown in Figure 2C, and 50 µL of 1:1 suspension of 50 mg mL⁻¹ fibrinogen with 3 × 10⁵ human MSCs (RoosterBio Inc., Frederick, MD 21 703, USA) and 10 U mL⁻¹ thrombin solution was added to the scaffolds and infiltrated with the help of the mild vacuum created by piston pull. Then the syringes were incubated at 37 °C with 5% CO₂ in a cell culture incubator (Forma Scientific, Inc., model 3110) for an hour to allow gelation of the fibrin gel as it infiltrated and encapsulated the Bg‐PLGA porous scaffold core.

Cell loading on the Bg‐PLGA scaffold controls was performed as described in our previous work:^{[ 49 ]} the scaffolds were restrained in a syringe that contained the cell suspension at a concentration of 3 × 10⁵ human MSCs in cell culture media (low glucose Dulbecco's modified Eagle medium (DMEM) media with 10% FBS (gibco), 0.5% gentamicin (gibco) and 1% glutamax (gibco), placed in the cell culture incubator, and periodically rotated in different positions to allow for an even distribution of cells. They were then left overnight in an upright position.

Cell loading in the fibrin gel controls was performed by incubating 1 mL syringes containing 50 µL of 1:1 suspension of 50 mg mL⁻¹ fibrinogen containing 3 × 10⁵ human MSCs and 10 U mL⁻¹ thrombin solution at 37 °C with 5% CO₂ in a cell culture incubator for an hour.

2.5. Cell Culture and Chondrogenic Differentiation

All samples after cell loading were transferred to new syringes with culture media (low glucose Dulbecco's modified Eagle medium (DMEM) media with 10% FBS (gibco), 0.5% gentamicin (gibco) and 1% glutamax (gibco) containing 0.1 m N‐2‐hydroxyethyl piperazine‐N‐2‐ethane sulfonic acid (HEPES) (gibco) to maintain the pH between 7.2 and 7.6 during the culture period and incubated at 37 °C with 5% CO₂ in a cell culture incubator (Forma Scientific, Inc., model 3110) for 2 days. All the samples were then refreshed with chondrogenic differentiation media consisting of high glucose DMEM media (gibco) containing 100 nm dexamethasone (Sigma‐Aldrich), 0.1 m HEPES, 10 ng mL⁻¹ TGF‐ß3 (EMD Millipore), 50 µg mL⁻¹ ascorbic acid (Sigma‐Aldrich), 40 µg mL⁻¹ proline (Sigma‐Aldrich), 1:100 Insulin, Transferrin, Selenium (ITS) premix (gibco), and 0.5% gentamicin (gibco). The media was refreshed every 3–4 days for 21 days. All the samples were then switched to chondrogenic basal media with the same composition as the chondrogenic differentiation media but without TGF‐ß3 for another 2 weeks to study the potential for hypertrophic differentiation of cells without external induction.

2.6. Cell Visualization and Proliferation

Samples cultured with MSCs after 21 and 35 days were visualized by performing live/dead assay in PBS for 30 min at 37 °C using an EVOS FL (Life Technologies) cell imaging system. Control samples without cells were observed first, and the settings were adjusted to obtain a dark background. The samples with cells were observed with the same corrected settings to eliminate autofluorescence from the sample material itself.

Cell metabolic activity was measured by Alamar blue (Invitrogen) assay after 4, 7, and 21 days of cell culture. Briefly, the cell culture media was removed from all samples, and the samples were incubated with fresh DMEM cell culture media (gibco) containing 10 vol% Alamar blue for 4 h at 37 °C. The media from each sample was then transferred to 96 well plates, and the fluorescence intensity was measured at an excitation wavelength of 540 nm and an emission wavelength of 585 nm, respectively. This fluorescence intensity at 585 nm corresponds to the metabolic activity of the cells in the samples as it arises from the conversion of resazurin, the active ingredient of the Alamar blue solution, into a compound called resorcin, by the cells. The fluorescence intensity of control samples without cells was subtracted from the intensity of their respective samples with cells to eliminate background fluorescence.

2.7. Cell Matrix Formation

Cartilage matrix formation by the cells was evaluated by a Dimethylmethylene Blue (DMMB) (Sigma‐Aldrich) assay that quantifies the sulphated glycosaminoglycans (GAGs), a major component of the cartilage matrix. As GAGs are negatively charged, they form a complex with DMMB that can be measured by reading the absorbance at 525 nm. Samples cultured with MSCs in chondrogenic media for 21 days were digested with freshly prepared papain digestion buffer (all components from Sigma‐Aldrich: 125 µg mL⁻¹ papain, 100 mm Na₂HPO₄, 5 mm ethylenediaminetetraacetic acid (EDTA), 5 mm cysteine, pH 6.3) at 60 °C overnight. DMMB was added to the digested samples, and the absorbance was measured at 525 nm to quantify the GAGs present in each sample. The absorbance value measured, indicative of GAG presence, was divided by the Alamar blue fluorescence measured on the same samples. This was done to evaluate cartilage matrix production normalized to the amount of cells present on each sample, and thus have a meaningful comparison between the different sample types.

2.8. Immunohistochemistry Analysis

The samples were prepared for cryo‐sectioning and subsequent histological analysis as follows. The samples were fixed with 10 vol% formalin for 2 h and then immersed in glucose (Sigma Aldrich) solutions of different concentrations to promote better tissue penetration of the embedding compound. The samples were immersed in 5 and 10 wt% glucose for 3 h each, followed by immersion in 20 wt% glucose solution overnight. The next day the samples were immersed in a 1:1 solution of 20 wt% glucose with NEG‐50^T (Richard‐Allan Scientific) embedding compound for 6 h, followed by overnight incubation in NEG‐50. Then the samples were placed in cryomolds with NEG‐50, frozen and stored at −80 °C. The embedded samples were cut horizontally into 10 µm thick sections, collected on glass slides (Fisherbrand Superfrost Plus Microscope Slides, Fisher Scientific) and used for IHC.

The presence of collagen types I, II, and X were analyzed by standard IHC techniques. Briefly, the slides were first fixed with 10 vol% formalin (Sigma‐Aldrich) for 5 min, rinsed with PBS, blocked with 5 wt% bovine serum albumin (BSA) (Sigma‐Aldrich) for 1 h, treated with rabbit primary antibodies collagen type I (ab34710, Abcam), collagen type II (ab34712, Abcam), or collagen type X (A18604, ABclonal) for 1 h, followed by incubation with secondary antibody (BioRad anti‐rabbit antibody conjugated with horseradish peroxidase (HRP)) at room temperature for another hour. Then the slides were treated with DAB (3,3′‐Diaminobenzidine) peroxidase kit (Fisher Scientific) for 15 min and mounted with Vectashield hard set mounting medium, which contains DAPI (4′,6‐diamidino‐2‐phenylindole) as a fluorescent stain for cell nuclei (Vector Laboratories Inc.). All the primary antibodies were prepared in 1% BSA (Sigma Aldrich) at a concentration of 1:100, and the secondary antibodies were prepared in 1% BSA at 1:250. Negative controls for each condition were done by replacing the primary antibody with 1% BSA.

All the slides were visualized under a Leica DMRB fluorescence microscope (20×, 40×), and the optical and fluorescent images for DAPI (excitation: 360 nm, emission: 425 nm) were taken at the same site without moving the slide and merged as layers using a compositing software called Nuke. The exposure and contrast settings remained the same for all the samples, and all samples were analyzed at the same time to make sure there were no variations due to user settings in the microscope as well as in Nuke.

2.9. Statistical Analysis

All the experiments were done with samples in triplicates, and the values are shown as mean ± standard deviation. Prism 8 software for macOS (GraphPad Software, LLC) was used for statistical analysis. Non‐parametric t‐tests using Welch's correction and one‐way ANOVA tests were used to compare the different sample types and time points. In all the tests, a p‐value of <0.05 was considered significant.

3. Results

3.1. Material Characterization

We prepared the control porous Bg‐PLGA scaffolds (Figure  3A) by a solvent casting and particulate leaching method similar to what reported in our previous paper,^{[ 49 ]} with the only modification that we used a higher temperature (52 °C) of incubation of the paraffin sphere mold than the temperature we used earlier (40 °C), to improve the pore interconnectivity by increasing the contact area between the microspheres.^{[ 51} , ⁵² , ⁵³ , ^{54 ]} The fractions of Bg and PLGA in the Bg‐PLGA scaffolds measured by thermogravimetric analysis were found to be 42 ± 5 and 58 ± 5 wt%, respectively (Figure S1A, Supporting Information). MicroCT analysis of the Bg‐PLGA scaffolds showed that most of the pores were interconnected (open porosity of 95.5 ± 0.2%, Figure S1B, Supporting Information).

Figure 3.

Representative photographs of Bg‐PLGA scaffolds (A), fibrin gel (B), and Bg‐PLGA@fibrin (C). SEM images of freeze‐dried Bg‐PLGA scaffold (D), fibrin hydrogel (E), and Bg‐PLGA@fibrin (yellow arrows highlight some fibers showing the presence of fibrin) (F). DMA of all samples under compression showing the storage modulus (G) and loss modulus (H) at increasing frequencies and stiffness (I) under 0.03% compression.

The fibrin gel (Figure 3B) was prepared following a method reported to lead to the production of a gel stable for up to 5 weeks^{[ 55 ]} to ensure it would adequately support the endochondral ossification process as previous studies have established a 3 weeks' time period for chondrogenic induction with additional 1 or 2 weeks for hypertrophic induction of chondrocytes.^{[ 56} , ^{57 ]}

To prepare Bg‐PLGA@fibrin (Figure 3C), Bg‐PLGA porous constructs were pre‐wetted and restrained in a syringe before adding fibrinogen and thrombin solution to facilitate the infiltration of fibrin through the pores of the scaffold.

SEM images of the control Bg‐PLGA scaffold (Figure 3D), fibrin gel (Figure 3E) and Bg‐PLGA@fibrin construct (Figure 3F) show the difference in morphologies between the different sample types. The fibers present in the fibrin gel (Figure 3E) were clearly present inside the pores of the Bg‐PLGA scaffold in the Bg‐PLGA@fibrin construct (Figure 3G), but not in the control Bg‐PLGA scaffold (Figure 3D) confirming the infiltration of fibrin through the macropores of Bg‐PLGA in the Bg‐PLGA@fibrin constructs (Figure 3F).

We studied the mechanical properties of all samples using DMA; all samples were wet and subjected to cyclic compression to mimic the in vivo environment.^{[ 58} , ^{59 ]} Understanding the mechanical properties of the scaffolds is important, as mechanical cues play a major role in driving the MSCs to differentiate into a particular phenotype.^{[ 38 ]} The storage modulus E′ (related to the ability to store energy elastically) increased with increasing frequency in the fibrin hydrogel (Figure 3G). A similar pattern was observed with the fibrin hydrogel for the loss modulus E″ (related to the ability of the material to dissipate stress through heat) (Figure 3H). An increase in E″ for the gel at higher frequencies has been observed in previous studies,^{[ 60} , ^{61 ]} and was attributed to the compressive stiffening of the fibrin gels where the loss modulus increases with higher magnitudes of compression due to the increase in the density of fibrin network and crisscrossing of fibers in the gel. The E′ and E″ of the Bg‐PLGA scaffolds and Bg‐PLGA@fibrin constructs did not show an obvious trend with increasing frequency. A similar non‐monotonous dependency of E′ and E″ with frequency was observed in previous studies for polymer‐Bg composite scaffolds.^{[ 49} , ^{62 ]} The Bg‐PLGA@fibrin constructs showed trends for E′ and E″ very similar to the scaffolds, which indicates that the scaffold core has a dominant influence on the mechanical properties of the Bg‐PLGA@fibrin construct. At any frequency, the E′ of the samples were higher than the E″ showing a viscoelastic behavior of all the samples. The stiffness of the samples at 0.03% strain from DMA was found to be 944 ± 22 N m⁻¹ for Bg‐PLGA scaffolds, 657 ± 23 N m⁻¹ for fibrin hydrogels and 782 ± 5 N m⁻¹ for Bg‐PLGA@fibrin samples (Figure 3I). This further confirms the high stiffness of the Bg‐PLGA composite scaffold compared to the fibrin gel.

3.2. Bioactivity in SBF

We evaluated the samples for their bioactivity (i.e., the ability of the materials to promote deposition of hydroxyapatite (HA) on their surface)^{[ 50 ]} in SBF. SEM analysis of SBF immersed samples after 4 days of immersion highlighted in color‐coded backscattered electron image (BSE) overlapped on secondary electron (SE) images^{[ 63 ]} showed thick, concentrated mineral deposits on Bg‐PLGA scaffolds (Figure  4A), no mineral deposition on fibrin gels (Figure 4B) and dispersed aggregates of mineral deposition on Bg‐PLGA@fibrin constructs (Figure 4C).

Figure 4.

Color‐coded overlay of backscattered electron (BSE) image (red color) and secondary electron (SE) image (green color) from SEM showing calcium phosphate deposits (identified from brighter red regions in the BSE image) on Bg‐PLGA scaffold (A), fibrin gel (B), and Bg‐PLGA@fibrin (C) samples. High magnification SEM secondary electron images (D–F) and EDS analysis (G–I) measured in the same areas of Bg‐PLGA scaffold (D,G), fibrin gel (E,H) and Bg‐PLGA@fibrin (F,I) after 4 days of SBF. XPS surface elemental analysis (J) showing Ca atomic % on the samples after 1, 4, and 7 days of immersion in SBF. Here, solid lines show statistical differences between different time points for the same sample types calculated by t‐tests (differences represented using the “*” symbol). The dotted line shows the statistical difference calculated between the different sample types by one‐way ANOVA analysis (differences represented using the “#” symbol). (****) and (# # # #) represents p‐value < 0.0001.

High magnification SEM SE images showed thick mineral deposits on Bg‐PLGA scaffolds (Figure 4D) confirmed to be calcium phosphate by EDS spectra showing intense Ca and P peaks (Figure 4G). Very few deposits were observed on the fibrin gel (Figure 4E) and Bg‐PLGA@fibrin samples (Figure 4F), containing traces of Ca, P and few other elements (Figure 4H,I). Surface elemental characterization by XPS showed the presence of more Ca (Figure 4J) and P (Figure S3, Supporting Information) on the surface of the Bg‐PLGA scaffolds compared to the other two samples, confirming their higher bioactivity. Fibrin gels showed the lowest amounts of Ca and P, whereas the Bg‐PLGA@fibrin samples showed Ca and P amounts between the scaffold and the gel samples, although closer to the scaffolds.

Raman spectra showed a characteristic HA peak at 960 cm⁻¹ in Bg‐PLGA scaffolds after 4 days of immersion in SBF; this peak was not observed in the Bg‐PLGA@fibrin and the gel samples (Figure S3, Supporting Information).

All these results suggest that the bioactivity of the samples is in the order of Bg‐PLGA> Bg‐PLGA@fibrin > fibrin hydrogel and that the presence of the Bg‐PLGA scaffold contributed to the high bioactivity of the Bg‐PLGA@fibrin

3.3. Cell Culture Results after 21 Days

We first cultured MSCs in the samples in chondrogenic differentiation media for 21 days to induce chondrogenesis. In this section we will be discussing the results of biological assays done at this time point.

We first visualized the cells inside the samples at the top layer under a fluorescence microscope after staining them with the live/dead staining assay. The cells showed a round morphology in all samples (Figure  5A–C). This was further confirmed by SEM analysis (Figure 5D–F).

Figure 5.

Cell morphology, metabolic activity, and GAG production after 21 days of cell culture. Fluorescence images after live/dead staining and SEM images of the Bg‐PLGA scaffold (A), fibrin gel (B), and Bg‐PLGA@fibrin samples (C). Here, live cells are stained in green and dead cells in red. SEM images of Bg‐PLGA scaffold (D), fibrin gel (E), and Bg‐PLGA@fibrin (F) samples. Alamar blue (AB) assay showing cell metabolic activity of the Bg‐PLGA scaffold, fibrin gel and Bg‐PLGA@fibrin samples after 4, 7, and 21 days of cell culture (G). Here (*) represents p < 0.05, (**) represents p < 0.01 and “n.s.” means that the difference was non‐significant. GAG amount quantified from DMMB assay normalized to the metabolic activity of cells from fluorescence measured via AB assay for the samples after 21 days of cell culture (H). Here (*) represents p < 0.05.

We then quantified the metabolic activity of the cells after 4, 7 and 21 days of culture using the Alamar blue assay. The Bg‐PLGA@fibrin and the fibrin gel showed similar cell metabolic activity at all time points (Figure 5G), indicating a similar cell loading in both samples. The Bg‐PLGA scaffolds showed a significantly lower cell metabolic activity at day 4 and 7 than the fibrin gel and Bg‐PLGA@fibrin samples indicating fewer cells were present on these samples (Figure 5C). The metabolic activity decreased in the Bg‐PLGA@fibrin samples at day 21 compared to day 7, whereas no significant differences were observed between day 7 and day 21 for the other sample types.

We analyzed the potential of the samples to support chondrogenic differentiation of MSCs using a colorimetric GAG assay where we quantified the sulphated GAGs, a major component of the cartilage matrix, after 21 days of culture. We normalized the GAG values to the fluorescence measured in the Alamar blue assay at 21 days, to account for differences in cell number on the different samples. We found that all three sample types supported the chondrogenic differentiation of MSCs upon induction with chondrogenic factors in media, as seen from the presence of GAGs in all samples (Figure 5H). The cells in the fibrin gel and Bg‐PLGA@fibrin showed no significant differences in GAG formation, whereas the cells in the Bg‐PLGA scaffold showed significantly lower GAG formation compared to the fibrin gel and Bg‐PLGA@fibrin, showing that the hydrogel plays an important role in promoting the chondrogenesis of MSCs.

3.4. Cell Culture Results after 35 Days

After 21 days of chondrogenic induction, we switched the culture medium to basal medium (i.e., removing chondrogenic induction factor TGF‐ ß3) and continued cell culture for another 14 days, to evaluate the potential of each sample type to induce hypertrophy and osteogenesis without external factors.

Live/dead staining at 35 days of culture showed clear differences in cell morphology among the samples, unlike what observed at 21 days. The cells in the Bg‐PLGA scaffold (Figure  6A) and in the fibrin gel (Figure 6B) still showed a round morphology, while a stretched spindle‐shaped morphology was observed in the samples (Figure 6C).

Figure 6.

Cell morphology and mineralization after 35 days of cell culture. Live/dead images of the Bg‐PLGA scaffold (A), fibrin gel (B), and Bg‐PLGA@fibrin (C) samples showing live cells in green and dead cells in red. Low magnification SEM image of Bg‐PLGA@fibrin (D) showing cells with different morphology in different locations of the sample: spherical cells inside the scaffold core and stretched cells at the interface between the gel and the scaffold. The cyan arrows here highlight the areas where partial degradation of fibrin gel is observed, exposing the Bg‐PLGA core. Zoomed‐in SEM images of the areas highlighted in D showing the stretched cells (E) and spherical cells (H; here, orange arrows highlight examples of the matrix fibers), as well as of the Bg‐PLGA scaffold (F) and fibrin samples (G). The insets in F, G, and H show high‐magnification images of the same areas to highlight the matrix formation (scale bar of insets: 30 µm). EDS analysis showing the map of selected elements identified on the Bg‐PLGA scaffold (I), fibrin gel (J) and Bg‐PLGA@fibrin (K). Scale bars on all images from F to K: 50 µm.

We further analyzed the surface of the samples with SEM. We observed partial degradation of the fibrin gel, exposing the inner Bg‐PLGA core (highlighted by the cyan arrows in Figure 6D); also shown in Figure S4, Supporting Information) and found co‐existence of cells with two different morphologies (Figure 6D): a stretched, spindle‐shaped morphology at the interface between the gel and the scaffold (Figure 6E) and a spherical morphology in the inner pores of the Bg‐PLGA scaffold core (Figure 6H). We found only spherical cells on the surface of the Bg‐PLGA scaffolds (Figure 6F) as well as in the fibrin samples, although embedded in the gel (Figure 6G). SEM images also showed no visible matrix formation on the Bg‐PLGA scaffolds (Figure 6F); matrix formation was not visible in the gel samples (Figure 6G) as the cells were embedded in the gel; whereas extensive matrix formation was observed in the samples (Figure 6H, orange arrows as examples).

EDS analysis on the same areas analyzed by SEM showed intense Ca and P signal on the surface of the Bg‐PLGA samples (Figure 6I), no Ca and P on the fibrin gel (Figure 6J) (Figure S5, Supporting Information) and presence of some Ca and P on the Bg‐PLGA@fibrin samples (Figure 6K). EDS maps of all elements measured on all samples are shown in Figure S5, Supporting Information, shows that only C and O are present in the fibrin gel.

3.5. IHC Analysis of ECM Components at 21 and 35 Days

We did IHC to further investigate the potential of the fibrin gel and Bg‐PLGA@fibrin samples in supporting chondrogenesis, hypertrophy, and osteogenesis, the three main stages of endochondral ossification, by identifying the main matrix components found in these three stages, that is, collagen type II (cartilage), collagen type X (hypertrophic cartilage) and collagen type I (bone), respectively.^{[ 64} , ^{65 ]} We couldn't analyze the Bg‐PLGA scaffolds by IHC as the scaffold sections were washed off from the slides during the different rinsing steps involved in the IHC process. On the other hand, the presence of fibrin in the fibrin gel and Bg‐PLGA@fibrin samples promoted good adhesion of the sections to the slides. So, in this section, we will be comparing only the fibrin gel and the Bg‐PLGA@fibrin samples. We analyzed the samples by IHC at 21 and 35 days to evaluate differences in matrix production at the end of the chondrogenic induction period (21 days) and after the chondrogenic induction was withdrawn (35 days). Negative controls consisting of sections treated with blocking solution instead of primary antibody followed by secondary antibody treatment were analyzed in parallel to confirm the absence of false positive staining.

After 21 days of culture in chondrogenic medium, the fibrin gel and Bg‐PLGA@fibrin showed positive staining for collagen type II (Figure  7A). Since collagen type II is a main component of cartilage, this result confirms MSC chondrogenesis in both samples,^{[ 64 ]} in line with the GAG assay results. Immunostaining for collagen type X showed that MSCs did not produce any collagen X neither in the fibrin gel nor in the Bg‐PLGA@fibrin (Figure 7A). Both fibrin gel and Bg‐PLGA@fibrin showed positive collagen type I staining, an osteogenic marker^{[ 65 ]} (Figure 7A). This staining was more intense in Bg‐PLGA@fibrin compared to the fibrin gel. This result shows that some level of osteogenic differentiation happened in the samples even when they were cultured in chondrogenic differentiation media.

Figure 7.

Optical images of IHC staining for collagen II, collagen X, collagen I, and negative controls. Fluorescent images showing DAPI stained nuclei at the same area are overlapped. The results are shown after 21 (A) and 35 days (B) of cell culture. In both (A) and (B), images on the left column refer to fibrin, and on the right column to Bg‐PLGA@fibrin. Scale bars on all images: 100 µm.

After 35 days we observed positive collagen II staining in the fibrin gel and Bg‐PLGA@fibrin, similar to what was observed after 21 days (Figure 7B). Collagen X was observed on Bg‐PLGA@fibrin, proving the potential of these samples to promote hypertrophic differentiation even in the absence of external induction. Collagen X was not observed in the fibrin gel samples. Collagen I staining in Bg‐PLGA@fibrin and the fibrin gel was similar to what was observed after 21 days showing osteogenesis continuing in both samples.

4. Discussion

A recent shift toward the endochondral approach to bone tissue engineering has been proposed in order to overcome the limitations of the direct ossification approach.^{[ 16 ]} However, scaffolds designed to sequentially support the endochondral ossification process have not been explored before this study. Here, we developed a scaffold design specifically aimed at supporting the endochondral ossification process: MSCs are loaded in a biodegradable fibrin hydrogel able to support chondrogenesis; as the cells degrade the gel, they reach an embedded bioactive porous and stiff Bg‐PLGA scaffold that induces hypertrophy without the need for any external factors, and supports osteogenesis. By evaluating MSC transdifferentiation in the Bg‐PLGA@fibrin sample and in the gel and bioactive scaffold as controls, we were able to assess the contribution of the individual components to the overall scaffold design.

We first tested mechanical properties and bioactivity in SBF; both properties were dominated by the Bg‐PLGA contribution, providing overall stiffness and high bioactivity to the Bg‐PLGA@fibrin samples. We then cultured MSCs in the samples in chondrogenic medium for 21 days. Here, the gel had a dominant effect, as it enabled high cell loading and high cellular metabolic activity in the Bg‐PLGA@fibrin samples, including large amount of GAG production.

After 21 days of cell culture in chondrogenic medium we removed any chondrogenic inducing factor from the medium and prolonged the culture for another 14 days in basal medium, to assess the ability of each sample to induce hypertrophy. These time points were chosen to follow the protocol used in many previous studies that tested scaffolds for endochondral ossification, to match the biological timelines of differentiation.^{[ 14} , ¹⁶ , ²¹ , ^{66 ]}

After 4 and 7 days of cell culture, we observed high metabolic activity in the fibrin gel and Bg‐PLGA@fibrin samples showing that the importance of the hydrogel to ensure high cell loading. The reduction in the metabolic activity of the cells observed in Bg‐PLGA@fibrin samples at day 21 may be related to saturation of cells in the matrix.^{[ 67 ]}

After 21 days of culture, all samples showed cells with a round morphology, which is typical of MSCs differentiated into chondrocytes.^{[ 68} , ⁶⁹ , ^{70 ]} Indeed, GAG production was found in all samples, confirming chondrogenesis; higher GAG formation was found in the gel and Bg‐PLGA@fibrin samples, showing that the presence of hydrogel promotes chondrogenesis of MSCs in the constructs. Hydrogels are known to be ideal candidates for cartilage tissue engineering as they mimic the hydrated nature of the cartilage matrix.^{[ 71} , ⁷² , ^{73 ]} Many natural and synthetic hydrogels have also been used in previous studies attempting endochondral ossification due to their ability to induce chondrogenesis and high cell loading capacity.^{[ 22} , ²⁴ , ²⁵ , ²⁶ , ^{28 ]}

MSC chondrogenesis in the fibrin and Bg‐PLGA@fibrin samples was also confirmed by IHC showing the presence of collagen II in both samples. IHC also showed osteogenesis, evaluated by the presence of collagen I, in both samples. This finding may be explained by the ability of fibrin to induce collagen I production in fibroblasts.^{[ 74 ]} The presence of Ca in the fibrin gels may also have contributed to inducing osteogenesis in both the gel and Bg‐PLGA@fibrin samples,^{[ 75 ]} and the high bioactivity of the Bg‐PLGA@fibrin may have contributed to osteogenesis in the Bg‐PLGA@fibrin samples.^{[ 76 ]} As expected, we didn't observe any hypertrophy in either the fibrin or Bg‐PLGA@fibrin samples at day 21.

At the end of the 35 days of culture, extensive matrix mineralization was observed in the Bg‐PLGA scaffold and Bg‐PLGA@fibrin samples. Mineral formation in the presence of cells on these samples can be due to two factors: mineralization produced by the cells or physicochemical calcium phosphate mineral deposition facilitated by the presence of Bg. EDS analysis showed high levels of Ca and P on the Bg‐PLGA scaffolds, but no cell matrix formation was visible from the SEM images. This suggests that minerals found in these samples may have mostly formed by physicochemical deposition facilitated by Bg. However, in the Bg‐PLGA@fibrin samples, the presence of Ca and P aligned with matrix formation by the cells, suggesting that minerals were produced to a large extent due to cellular activity.

To our surprise, after 35 days, we observed a change in cell morphology only in the Bg‐PLGA@fibrin samples. We could observe distinct spatial differences in the Bg‐PLGA@fibrin constructs: the cells inside the Bg‐PLGA core showed a spherical morphology, while the cells at the interface between the gel and the Bg‐PLGA showed a stretched morphology. MSCs are usually spherical inside soft hydrogels and spread out when in contact with 3D stiff substrates.^{[ 38} , ^{39 ]} However, unexpectedly, in our study, MSCs achieved an elongated morphology only in the Bg‐PLGA@fibrin samples and not in the Bg‐PLGA scaffolds. This suggests that in the Bg‐PLGA@fibrin samples the MSCs changed phenotype upon transition from the gel to the scaffold core, and that the observed change in shape was not just due to spreading upon contact with the scaffold core. Previous studies have shown that hypertrophic chondrocytes assume an elongated morphology when they differentiate to an osteoblast‐like phenotype at the cartilage/bone interface.^{[ 77 ]} A similar phenomenon may be happening at the gel‐scaffold interface in the Bg‐PLGA@fibrin samples, where the hypertrophic chondrocytes transition from the soft fibrin gel gets to the stiffer, mineralized Bg‐PLGA scaffold. This transition could have resulted from the partial degradation of fibrin gels exposing the inner Bg‐PLGA core, allowing cells to transition from the gel to the stiff Bg‐PLGA matrix, as seen in Figure 6D.

The hypertrophic differentiation of chondrocytes (evidenced by collagen X production) in the Bg‐PLGA@fibrin samples was confirmed by IHC. The presence of collagen I, indicative of osteogenesis, at both 21 and 35 days of culture, may have also promoted the osteoblast‐like differentiation of hypertrophic chondrocytes, similarly to what happens during endochondral ossification, where osteogenesis happening at the bone collar promotes the hypertrophic differentiation of chondrocytes at the center of the bone.^{[ 78 ]}

5. Conclusion

We report a Bg‐PLGA@fibrin construct designed to promote endochondral ossification by first supporting MSC chondrogenesis within a biodegradable fibrin hydrogel and then hypertrophy and osteogenesis once the MSCs degrade the gel and get in contact with a stiff bioactive Bg‐PLGA scaffold core.

Results show that the Bg‐PLGA@fibrin samples combine the high cell loading and ability to support chondrogenesis of the fibrin hydrogel with the ability to induce matrix mineralization of the Bg‐PLGA scaffold core. The combination of the two elements uniquely induces a switch in cell morphology at the gel/scaffold interface, which may be related to osteoblast like differentiation of hypertrophic chondrocytes. Hypertrophy was achieved on the Bg‐PLGA@fibrin samples solely based on material cues and without the need for external induction factors.

This work has a few limitations: while fibrin degradation is a crucial parameter in determining hypertrophic differentiation, we only observed it by SEM but did not quantify it. Future work may attempt to quantify fibrin degradation using techniques such as western blotting or fluorescence marking.^{[ 79} , ^{80 ]} Also, we monitored the endochondral ossification process solely through immunostaining. Future work should involve gene expression analysis to further confirm histology results and shed light on the progression of the endochondral ossification process. Special attention should be paid in this study to find suitable housekeeping genes that remain constant among different samples. The effect of scaffolds on cellular activity after complete fibrin gel degradation could be another topic of future investigation. This would have to be addressed in an in vivo model: a 35‐day long 3D in‐vitro culture is at the limit of feasibility due to issues related to contamination.^{[ 81} , ⁸² , ^{83 ]} Also, the final stages of endochondral ossification like vascularization, complete bone formation and remodeling can be studied only in an in vivo model.

With their ability to induce hypertrophy only based on material cues, Bg‐PLGA@fibrin constructs hold great potential as scaffolds for bone tissue regeneration through endochondral ossification. These samples could be implanted in‐vivo after only a short period of chondrogenic induction (we used 21 days in this study, but previous studies showed that as little as 7 days may be necessary to induce chondrogenesis in MSCs cultured in fibrin gels^{[ 84 ]}), thereby facilitating clinical translation.

Conflict of Interest

The authors declare no conflict of interest.

Supporting information

Supporting Information

Jeyachandran D., Murshed M., Haglund L., Cerruti M., A Bioglass‐Poly(lactic‐co‐glycolic Acid) Scaffold@Fibrin Hydrogel Construct to Support Endochondral Bone Formation. Adv. Healthcare Mater. 2023, 12, 2300211. 10.1002/adhm.202300211

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.