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. 2024 Dec 24;17(1):197–210. doi: 10.1021/acsami.4c13246

Effective Bone Tissue Fabrication Using 3D-Printed Citrate-Based Nanocomposite Scaffolds Laden with BMP9-Stimulated Human Urine Stem Cells

Piao Zhao 1,2, Yi Zhu 2,3, Mirae Kim 4,16, Guozhi Zhao 1,2, Yonghui Wang 2,5, Caralyn P Collins 6, Ou Mei 2,7, Yuanyuan Zhang 8, Chongwen Duan 4, Jiamin Zhong 2,9, Hui Zhang 2,10, Wulin You 2,11, Guowei Shen 2,12, Changqi Luo 2,13, Xingye Wu 1,2, Jingjing Li 2,14, Yi Shu 2,15, Hue H Luu 2, Rex C Haydon 2, Michael J Lee 2, Lewis L Shi 2, Wei Huang 1, Jiaming Fan 2,9, Cheng Sun 6,16, Liangyuan Wen 3,*, Guillermo A Ameer 4,16,17,*, Tong-Chuan He 2,16,18,*, Russell R Reid 2,16,18,*
PMCID: PMC11783527  PMID: 39718997

Abstract

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Effective repair of large bone defects through bone tissue engineering (BTE) remains an unmet clinical challenge. Successful BTE requires optimal and synergistic interactions among biocompatible scaffolds, osteogenic factors, and osteoprogenitors to form a highly vascularized microenvironment for bone regeneration and osseointegration. We sought to develop a highly effective BTE system by using 3D printed citrate-based mPOC/hydroxyapatite (HA) composites laden with BMP9-stimulated human urine stem cells (USCs). Specifically, we synthesized and characterized methacrylate poly(1,8 octamethylene citrate) (mPOC), mixed it with 0%, 40% or 60% HA (i.e., mPOC-0HA, mPOC-40HA, or mPOC-60HA), and fabricated composite scaffold via micro-continuous liquid interface production (μCLIP). The 3D-printed mPOC-HA composite scaffolds were compatible with human USCs that exhibited high osteogenic activity in vitro upon BMP9 stimulation. Subcutaneous implantation of mPOC-HA scaffolds laden with BMP9-stimulated USCs revealed effective bone formation in all three types of mPOC-HA composite scaffolds. Histologic evaluation revealed that the mPOC-60HA composite scaffold yielded the most mature bone, resembling native bone tissue with extensive scaffold-osteointegration. Collectively, these findings demonstrate that the citrate-based mPOC-60HA composite, human urine stem cells, and the potent osteogenic factor BMP9 constitute a desirable triad for effective bone tissue engineering.

Keywords: Bone tissue engineering, human urine stem cells, BMP9, citrate-based scaffold, bone formation, osteogenic differentiation, osteointegration

Introduction

As one of a few organs that retain full regenerative potential throughout life, bone is a dynamic tissue and plays important roles in supporting the skeletal system, protecting internal organs, maintaining hematopoietic cells and mineral homeostasis.1 While most fractures and small defects can be effectively self-repaired through bone regeneration and remodeling, large defects that are caused by traumatic injury, severe infection, tumor resection, degenerative disease, congenital defects, or functional atrophy usually exceed the critical size threshold or greater than 50% loss of circumference of bone and may cause nonunion, malunion, or pathological fracture.1,2 These large bone defects are usually fixed by using biologically inert metallic devices and/or bone autografts and allografts.1 Over four million of such orthopedic procedures are performed globally.1,2 However, these so-called gold standard treatments are associated with certain risks. For example, metal devices present infection risks and can often require unplanned surgical removal; bone allografts are associated with possible disease transmission; and bone autografts are limited in supply and can result in donor site morbidity. Thus, more efficacious bone repair strategies are needed, and engineered bone tissue is considered as a potential alternative to the conventional use of bone grafts.3

Bone regeneration during fracture healing recapitulates the well-orchestrated bone formation process during embryonic development,4 which has provided a key inspiration for bone tissue engineering (BTE).1,2 While significant progress has been made since the 1990s, the BTE field still faces unmet needs and technological challenges.1−3 It has been well-established that effective BTE requires optimal and synergistic interactions of the following triad: biocompatible scaffolds, osteogenic factors, and osteoprogenitors (e.g., mesenchymal stem cells, MSCs), which can form a highly osteoconductive and/or osteoinductive microenvironment with extensive vascularization for bone regeneration and osseointegration.3

Bone formation is tightly regulated by numerous growth factors, signaling pathways and even noncoding RNAs.5−8 Through a comprehensive analysis of the 14 types of bone morphogenic proteins (BMPs), we have demonstrated that BMP9 is the most potent osteogenic BMP and superior to the well-recognized, FDA-approved osteogenic BMPs such as BMP2 and BMP7 (i.e., OP-1).9,10 It is conceivable that BMP9 may represent one of the most desirable growth factors for BTE.11 Moreover, BMP9 and relevant gene therapies have been gaining increasing attention within scientific research.12−14

As an important component for effective BTE, MSCs are multipotent stem cells that can self-renew and differentiate into multiple cell types including bone and cartilage.3,15 While bone marrow stromal cells (BM-MSCs) are most commonly used in MSC studies, MSCs have also been identified and isolated from numerous tissues such as adipose tissue, oral cavity, cranial suture, skin basal layer, periosteum, vascular pericytes, and Wharton’s Jelly of umbilical cord.3,15,16 In fact, we and others have identified a more readily available source of highly osteogenic progenitors, called urine stem cells (USCs), isolated from the urine samples of healthy human donors.17,18 The primary advantage of USCs over other stem cells lies in their ease of accessibility. Moreover, the ability to conveniently and noninvasively extract USCs from a patient could address the issue of limited stem cell sources while also reducing the risk of immune rejection. Thus, the use of USCs should provide a novel and convenient osteoprogenitor source for efficacious BTE.

Scaffolds as bone graft substitutes are a key part of the BTE triad and should possess desirable physicochemical-mechanical properties (such as stiffness, biodegradability, and surface chemistry) and biocompatibility (such as cell adhesion, proliferation, differentiation and migration).1,19 Traditionally, scaffolds can be made from natural biomaterials such as collagens, hydroxyapatite (HA), β-tricalcium phosphate (β-TCP), calcium phosphate cement, and ceramic glass,1,2 or synthetic polymers such as poly(lactic acid) (PLA), poly(lactic-co-glycolic acid) (PLGA) and poly(caprolactone) (PCL), or bioceramics, metals and composites that are FDA-approved materials for clinical applications.1,2,20 A citrate-based polymer (CBP) has previously been reported to have novel biodegradable and elastomeric properties, leading to the generation of poly(1,8-octanediol-co-citric acid) (POC).21,22 A composite of POC with hydroxyapatite has recently been used for manufacturing implantable medical devices cleared for marketing in the USA by the FDA for musculoskeletal surgeries. Citric acid is chosen as a polyfunctional monomer since citrate is an endogenous metabolite and can regulate numerous cellular processes.19,23 CBPs have tunable mechanical and physical properties.19,23 In fact, a thermoresponsive poly(polyethylene glycol citrate-co-N-isopropylacrylamide) (PPCN) has been reported for various biomedical applications including the delivery of biologics.24−26

HA, the primary inorganic component of bone and teeth, exhibits excellent bioactivity and biocompatibility. Previous studies have shown that HA enhances osteoconduction, facilitating interactions between cells and tissues.27,28 However, HA alone as a scaffold in bone tissue engineering faces limitations such as brittleness, poor toughness, susceptibility to fatigue fractures, prolonged biodegradation, and structural simplicity, which hinders cell survival and vascularization.29,30 These drawbacks restrict the use of HA as a stand-alone material in bone engineering. Combining HA with other materials, either as a surface coating or in composite form, can integrate the advantages of each material and mitigate their individual limitations.31

In this study, we sought to develop an optimized BTE system via 3D printing of citrate-based POC/HA composites laden with BMP9 transduced USCs. CBP composites of the mPOC with various mass percentages of HA (0%, 40% or 60%; i.e., mPOC-HA, mPOC-40HA, or mPOC-60HA) were prepared to fabricate scaffolds via additive manufacturing using continuous liquid interface production (μCLIP). Our results showed that the 3D-printed mPOC-HA composite scaffolds were compatible with USCs that exhibited high osteogenic capacity in vitro upon BMP9 stimulation. Subcutaneous implantation of mPOC composite scaffolds laden with BMP9-stimulated USCs indicated that BMP9-stimulated USCs effectively formed new bone in all three types of mPOC-HA composite scaffolds. Histologic evaluation revealed that the mPOC-60HA composite scaffold yielded the most mature and highly mineralized new bone resembling native bone tissue with extensive scaffold osteointegration. Collectively, our findings demonstrate that citrate-based mPOC-60HA composite, human urine stem cells, and the potent osteogenic factor BMP9 may constitute a desirable triad for effective bone tissue engineering.

Experimental Section

Chemicals and Cell Culture

Human HEK-293 cells were obtained from the American Type Culture Collection (ATCC, Manassas, VA). 293pTP, RAPA and 293GP cells were derived from HEK-293 cells as previously described.32,33 Human urine stem cells (USCs) were isolated from healthy donors at the Wake Forest Institute for Regenerative Medicine (under the Institutional Review Board protocol number IRB00014033), Winston-Salem, NC, and cultured under the conditions as previously described.34 All other cells were cultured in the DMEM supplemented with 10% fetal bovine serum (FBS, Gemini Bio-Products), 100 U/mL penicillin, and 100 μg/mL streptomycin at 37 °C in 5% CO2 as described.35,36 Unless indicated otherwise, chemicals were obtained from Thermo Fisher Scientific (Waltham, MA) or Millipore Sigma (St. Louis, MO).

Synthesis and Chemical Characterization of mPOC Polymer

The mPOC polymer was synthesized by a two-step process: (1) the synthesis of POC prepolymer and (2) the conjugation of methacrylate groups onto the prepolymer. Briefly, the equal molar of citric acid and 1,8-octanediol was melted at 165 °C, and then transferred into 140 °C oil bath with continuous stirring of 55 min under nitrogen gas. The mixture was subsequently precipitated with Mili-Q (MQ) water and lyophilized to obtain the prepolymer. After lyophilization, the POC prepolymer (66 g) was dissolved in tetrahydrofuran (THF, 540 mL) at 60 °C, and imidazole was added until the mixture became clear. Glycidyl methacrylate was slowly added and allowed to react for 6 h under reflux, followed by concentration in a rotary evaporator. The crude product was obtained by precipitation in excess MQ water. After purifying with MQ water, the final product (namely, mPOC) was obtained by lyophilization. The final product was further characterized by using 1H NMR (X500) spectroscopy and Fourier Transform Infrared (FTIR) spectroscopy analysis (Thermo Nicolet Nexus, ART mode).

Preparation of mPOC-HA Composite with Varied Hydroxyapatite (HA) Concentrations

The mPOC-HA composite with varied HA concentrations was prepared by mixing mPOC polymer and HA powder in various ratios while maintaining an overall concentration of 70 wt % in pure ethanol. Specifically, the mPOC/HA ratios used were 100:0 (i.e., mPOC-0HA), 60:40 (i.e., mPOC-40HA), and 40:60 (i.e., mPOC-60HA). After thorough mixing, Irgacure 819 and ethyl 4-dimethylamino benzoate (EDAB) were added to the mixture at a final concentration of 3% (wt %) as cophoto-initiators, which will not affect the biocompatibility as previous described.37,38

Fabrication of the 3D-Printed mPOC-HA Scaffolds

A custom-built micro-continuous liquid interface production printer was used to fabricate these samples with the same methods as reported in previous work. Briefly, scaffolds were first designed as CAD files in SolidWorks (Dassault Systèmes, Vélizy-Villacoublay, France) to create a printable model. Scaffold design consisted of a repeated hexagonal unit cell with 125 μm strut thickness which was patterned and cut in CAD to a cylindrical scaffold 5.125 mm in diameter and approximately 1.5 mm in height. The CAD models for each scaffold were subsequently sliced into cross-sectional images via the commercial slicer CHITUBOX Basic (CBD-Tech, Shenzhen City, China) with 5 μm layer thickness. The resulting cross-sectional images were then projected onto the resin bath via the use of a digital light projector of wavelength 365 nm with a digital micromirror device (DMD). For this study, a lateral pixel resolution of 3.98 μm x 3.98 μm was utilized during printing. Power delivery on this system was slightly higher than for previous studies, with up to 6.094 mJ/cm2 used in the case of the mPOC-60HA composite. As in previous work, teflon was used as an oxygen permeable membrane in order to prevent polymerization of the resin along the bottom of the resin bath and allow for a continuous printing process. After printing, scaffolds were placed under UV flood for 1 minute, flipped, and UV cured for an additional minute.

Mechanical Characterization of mPOC-HA Scaffolds

The mechanical properties of mPOC-HA structures were assessed by a universal testing machine (Model 5940, Instron, High Wycombe, UK). The mPOC-HA scaffolds were printed in a plug structure with 3 mm diameter and 6 mm height dimensions and were pretreated in PBS at room temperature (RT) overnight before measurement. The compressive crosshead displacement was applied to the plug at a rate of 2 mm/min until it reached the breaking point, whereas the stress versus strain curve was obtained. The compressive modulus was determined by calculating the slope of the stress–strain curve within the initial 10% strain range.

The degradation kinetics of mPOC-HA scaffolds was evaluated by monitoring changes in the mass over time. Specifically, the scaffolds were immersed in PBS at 75 °C, and mass loss was recorded weekly for 13 weeks. The degradation behavior was normalized to the initial mass and then plotted as a mass vs time curve.

Synthesis of the Thermoresponsive PPCN Polymer

The PPCN polymer was synthesized by following the procedure previously described.24 The PPCN gel has a lower critical solution temperature of 26 °C so it remains in liquid status below 21 °C but solidifies above 30 °C. In order to enhance the cell adhesion property of PPCN, gelatin was added to the PPCN solution (100 mg) to a final concentration of 0.1% (wt %) as previously described,25 resulting in the PPCNg solution that was kept at 4 °C before use.

Construction and Amplification of Recombinant Adenoviral Vectors Ad-GFP, Ad-GFP-GLuc, and Ad-BMP9

Recombinant adenoviruses Ad-GFP, Ad-GFP-GLuc and Ad-BMP9 were constructed by using either the AdEasy technology or the Gibson DNA Assembly based OSCA system as previously described.39,40 Briefly, the coding regions of Gaussia luciferase (GLuc),41 and human BMP942 were PCR amplified and subcloned into the adenoviral shuttle vector pAdTrack-CMV, followed by homologous recombination reactions with the adenoviral backbone vector pAdEasy1 in BJ5183 cells.39 The resultant recombinant adenoviral plasmids were used to generate adenoviruses Ad-GFP-GLuc and Ad-BMP9.43 The Ad-GFP was constructed by using the Gibson DNA Assembly based OSCA system as previously described.44 The recombinant adenoviruses were packaged in 293pTP cells, and amplified to high titers in HEK-293, RAPA, 293pTP, or 293GP cells.32,33 The Ad-BMP9 also coexpresses the GFP marker gene. Polybrene (final concentration at 6 μg/mL) was added to enhance adenoviral transduction efficiency in the human USCs as previously described.45,46

Cell and mPOC-HA Coculture Cell Viability Assay

Exponentially growing USCs were seeded at a low density (5 × 104 cells/well) in 12-well plates and cocultured with either mPOC-HA scaffolds or no scaffold control. Viable cells were visualized by using Calcein-AM staining and imaged at indicated time points as previously described.47

mPOC-HA Biocompatibility Assay

Subconfluent USCs were infected with Ad-GFP-GLuc for 16h. Cells were harvested, resuspended in DMEM, and seeded to the pretreated mPOC-HA scaffolds submerged in culture medium (approximately 1 × 104 cells per seeding). At 6 days post seeding, the cell-laden obturator was reinfected with Ad-GFP-GLuc. Both GFP fluorescence and the secreted Gaussia luciferase activity in the culture medium were monitored at the indicated time points as indicators of cell viability. Gaussia luciferase activity was quantified using the Secrete-Pair Gaussia Luciferase Assay Kit (GeneCopoeia, Rockville, MD) as previously described.48,49

Early Osteogenic Marker Alkaline Phosphatase (ALP) Assays

ALP activity was determined both quantitatively and qualitatively as previously described.50 Briefly, exponentially growing USCs were seeded in 24-well cell culture plates and infected with Ad-BMP9 or Ad-GFP. At 3 and 7 days after infection, ALP activity was quantitatively assessed by using a modified Great Escape SEAP Chemiluminescence assay (Takara Bio USA, San Jose, CA) and qualitatively determined by using a histochemical staining assay (using a mixture of 0.1 mg/mL napthol AS-MX phosphate and 0.6 mg/mL Fast Blue BB salt). Each assay condition was performed in triplicate.

Matrix Mineralization Assay (Alizarin Red S Staining)

Exponentially growing USCs were seeded in 24-well cell culture plates, infected with Ad-BMP9 or Ad-GFP, and cultured in the presence of ascorbic acid (50 μg/mL) and β-glycerophosphate (10 mM). At 21 h after infection, the infected cells were fixed with 0.05% (v/v) glutaraldehyde at room temperature for 10 min. After being washed with distilled water, fixed cells were incubated with 0.4% Alizarin Red S (Sigma-Aldrich) for 5 min, followed by extensive washing with distilled water as previously described.43,51 The stained mineralized matrix nodules were recorded by bright field microscopy. Each assay condition was performed in triplicate.

RNA Isolation and TqPCR Analysis of Osteogenic Markers

Total RNA was isolated from the cultured cells using the NucleoZOL reagent (Takara Bio USA Inc.) and subjected to reverse transcription into cDNA with hexamer and M-MuLV reverse transcriptase (New England Biolabs, Ipswich, MA) as previously described.52 The cDNA products were further diluted and used as PCR templates. The PCR primers (Table S1) were designed by using the Primer 3plus program to amplify the genes of interest. SYBR Green-based touchdown qPCR (TqPCR) analysis was carried out as described.53,54 Briefly, the RT products were diluted and used as templates. TqPCR reactions were set up by using the 2x Forget-Me-Not EvaGreen qPCR Master Mix (Biotium, Fremont, CA), and carried out by using CFX-Connect (Bio-Rad Laboratories, Hercules, CA) as previously described.53,55 TqPCR cycling program was as follows: 95 °C × 3′ for one cycle; 95 °C × 20″, 66 °C × 10″, for 4 cycles by decreasing 3 °C per cycle; 95 °C × 20″, 55 °C × 10″, 70 °C× 1″, followed by plate read, for 40 cycles. All TqPCR reactions were performed in triplicate. Gene expression was normalized with GAPDH expression level and calculated by using the 2–ΔΔCq method.

Subcutaneous Implantation of USC-laden mPOC-HA Scaffolds

The use and care of animals in the study followed the guidelines approved by the Institutional Animal Care and Use Committee (ACUP#71445). The subcutaneous implantation experiments were carried out as previously reported.56−58 Briefly, the mPOC-0HA, mPOC-40HA and mPOC-60HA scaffolds were pretreated in sterile PBS and then complete DMEM, respectively. Skeletally mature athymic nude mice (Envigo, Indianapolis, IN; 6–8-week-old, both male and female) were used and divided into the following three groups for each type of scaffold: (1) mPOC-0/40/60HA scaffold only group; (2) mPOC-0/40/60HA scaffolds laden with Ad-GFP transduced USCs group; and (3) mPOC-0/40/60HA scaffolds laden with Ad-BMP9 transduced USCs group. Prior to scaffold implantation, subconflent USCs were infected with Ad-GFP or Ad-BMP9, collected at 16h after infection, and resuspended in ice-cold PPCNg.25 Approximately 5 × 106 cells (in 80 μL of PPCNg, kept at 4 °C) were loaded onto a scaffold, and six implants were set up for each group/scaffold in the flanks of athymic nude mice. Mice were sacrificed 7 weeks after implantation, and the subcutaneous masses were retrieved for histologic evaluation.

H & E Staining

The retrieved specimens were fixed in 10% PBS-buffered formalin, decalcified, and paraffin embedded using Leica TP1020 Automatic Tissue Processor (Leica Microsystems, Deerfield, IL). Sections were deparaffinized, rehydrated and subjected to H & E staining as described.59 New bone formation was quantitatively determined by using the ImageJ analysis of high-power images of multiple samples within the same group and was calculated as% of average osteoid matrix area over the total image area.

Masson’s Trichrome Staining

The retrieved specimens were fixed, decalcified, and paraffin embedded. Sections were deparaffinized, rehydrated and subjected to Masson’s trichrome staining (Newcomer Supply) as described.10,60 Highly mineralized mature bone formation in mPOC-40HA vs mPOC-60HA was quantitatively compared by using the ImageJ analysis of the mature bone regions of multiple high-power images within the same group and was calculated as% of average mature bone area over the total new bone formation area.

Modified Periodic Acid-Schiff (PAS) Staining of Bone-Cartilage Interfaces

Sections of the above paraffin blocks were deparaffinized, rehydrated and subjected to the modified PAS staining protocol as described.61 Briefly, the deparaffinized and hydrated sections were exposed to periodic acid (1g/dL; Sigma-Aldrich, cat. no. 3951–100 ML) for 5 min, rinsed in distilled water, immersed in Schiff’s reagent (pararosaniline HCl 1% and sodium metabisulfite 4% in hydrochloric acid 0.25 mol/L; Sigma-Aldrich, cat. no. 3952–50 ML) for 10 min, and rinsed under running distilled water for 5 min. The slides were then counterstained with Weigert’s iron hematoxylin (Sigma-Aldrich, cat no. 1159730002) for 5 min, rinsed in distilled water, “blued” in PBS for 20 s, briefly rinsed in distilled water, and stained with a 1% light green solution (Sigma-Aldrich, cat. no. L1886) for 30 s to 2 min. The slides were then rinsed in distilled water, allowed to briefly air-dry, then finally dehydrated, and coverslipped. The stained slides were imaged with a bright field microscope.

Statistical Analysis

All experiments were performed at least three times or repeated in three batches of independent experiments. Data were analyzed using GraphPad Prism 7 and presented as the mean ± standard deviations (SD). Statistical significance was determined by one-way ANOVA and the student’s t test for the comparisons between groups. A value of p < 0.05 was considered statistically significant.

Results and Discussion

Hydroxyapatite (HA) Improves Mechanical Properties of 3D-Printed Citrate-Based mPOC Scaffold

mPOC was synthesized first from POC through an esterification reaction between 1,8-octanediol and citric acid as described.21 (Figure 1A-a). Methacrylate moieties were demonstrated via 1H NMR analysis, which identified peaks at 1.9, 5.7, and 6 ppm (Figure 1A-b), while the FT-IR spectrum exhibited a C=C stretching vibration peak at 1636 cm–1 (Figure 1A-c). mPOC was therefore effectively produced through POC methacrylation, and this modification does not prohibit photopolymerization under UV light in the 3D printing process.

Figure 1.

Figure 1

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Characterization and design of 3D-printed mPOC-HA scaffolds. (A) (a) Schematic representation of mPOC synthesis; (b) 1H NMR analysis of mPOC; (c) FT-IR spectra of mPOC; (B) (a) μCLIP 3D printer system and schematic depiction of 3D-printed mPOC-HA scaffolds; (b) Representative stress–strain curves of each mPOC-HA scaffold; *p < 0.05, **p < 0.01 compared with that of the mPOC-0HA group; (c) Degradation behavior of each mPOC-HA scaffold at 75 °C.

To improve the mechanical properties of mPOC, render it more amenable to osteogenesis and achieve a wide range of 3D printable composites, we incorporated HA microparticles at 0%, 40% or 60% of HA. The porous mPOC-HA scaffolds were 3D-printed as disk-like structures for in vivo implantation via a custom-built microcontinuous liquid interface production (μCLIP) 3D printing system62 (Figure 1B-a). Additive-manufacturing of the composite is achieved by projecting cross-sectional images of the 3D-designed scaffold onto the focal plane of the resin using DMD (Figure 1B-a). mPOC-HA composite scaffolds were mechanically tested using plug-shaped samples (3 mm × 6 mm), which is a standard structure (ASTM D695), and measuring the compressive modulus of materials (Figure 1B-b). The stress–strain regimes of mPOC-HA samples were measured using a universal testing machine until they fractured under a compressive load, and we found that mPOC-60HA scaffold modification had the greatest compressive strength profile (Figure 1B-b), signifying compatibility between mPOC and HA. Furthermore, degradation profiles of 3D-printed porous mPOC-HA scaffolds were obtained by immersing the composites in PBS for 13 weeks at 75 °C, mimicking an environment accelerated 16 times compared to body temperature.63 At 12 weeks, while the mPOC-0HA scaffold lost 54.1% of its original mass, the mPOC-60HA scaffold only lost 24.6% of its original mass, indicating that the incorporation of 60% HA delayed the degradation behavior by more than 2-fold (Figure 1B-c). Collectively, these results demonstrate that incorporation of HA significantly improves the compressive strength and modulus of elasticity of 3D-printed mPOC scaffolds.

The mPOC-HA Scaffolds Are Compatible with Human USCs

When the 3D-printed composite scaffolds were examined under a microscope, the HA addition provided more structural rigidity to mPOC scaffolds (Figure 2A, a vs.bc). The mPOC-HA scaffolds were sterilized and pretreated with 70% ethanol and PBS, and then cocultured with human USCs. When the viable cells were stained with calcein-AM, the numbers of positively stained cells counted in mPOC-40HA and mPOC-60HA groups were comparable at day 7 with those of the mPOC-0HA or control group (Figure 2B) even though viable cell numbers were significantly lower in the mPOC-40HA and mPOC-60HA groups than in the control and mPOC-0HA groups at day 3 (Figure 2C). The coculture results indicate that human USCs are well tolerated on the mPOC-HA scaffolds.

Figure 2.

Figure 2

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Coculture of mPOC-HA scaffolds with human urine stem cells (USCs). (A) Microscopic images of mPOC-0HA (a), mPOC-40HA (b), and mPOC-60HA (c) scaffolds. (B) Exponentially growing USCs were seeded in 12-well plates at a low density (5 × 104 cells/well) and cocultured with the mPOC-HA scaffolds or no scaffold control. Viable cells were stained with calcein-AM and imaged at the indicated time points. Representative results are shown. (C) Quantitative analysis of cell viability of USCs cocultured with mPOC-HA scaffolds. The coculture experiments were set up as described in (B). Viable cells were counted on day three and day seven. **p < 0.01 compared with that of the control group.

We further tested the cell adhesion and survival capabilities of the mPOC-HA composite by directly seeding Ad-GFP-GLuc transduced USCs onto the scaffolds, so the cell survival was easily monitored by fluorescence signal visualization and through the quantitative measurement of the secreted GLuc activity in the culture medium (Figure 3A). We found the GLuc activities in the mPOC-40HA and mPOC-60HA groups were higher than that in the mPOC-0HA group, especially at days 3 to 6 after cell seeding although the mPOC-0HA group had higher GLuc activity at day 7 after Ad-GFP-GLuc reinfection (Figure 3B). These findings are consistent with the previous reports in which the surface roughness resulting from HA can promote cell adhesion and differentiation.64 Similar results were obtained from GFP signal analysis (Figure 3C). Collectively, these results demonstrate that the 3D-printed mPOC-HA composite scaffolds exhibited an acceptable biocompatibility with human USCs.

Figure 3.

Figure 3

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Biocompatibility of mPOC-0HA, 40HA, and 60HA with human urine stem cells (USCs). (A) Depiction of the experimental design. Subconfluent USCs were infected with Ad-GFP-GLuc that coexpresses both GFP and secreted Gaussia luciferase. Cells were harvested and loaded onto mPOC-HA scaffold submerged in cell culture. Cell-ladened scaffolds were reinfected with Ad-GFP-GLuc at after 6 days in culture. GFP signal level and GLuc activity served as surrogate of cell survival. (B) Secreted GLuc activities were determined at the indicated time points. *p < 0.05; **p < 0.01, compared with that of the mPOC-0HA group. (C) GFP signal was recorded at the indicated time points. Representative results are shown.

Human USCs Undergo Effective Osteogenic Differentiation upon BMP9 Stimulation

We previously identified BMP9 as the most osteogenic factor among the 14 types of BMPs.10,43,59 We further elucidated the underlying mechanisms of BMP9-inudced osteogenic differentiation, as well as its extensive crosstalk with other major signaling pathways.7,8,11,42,47 We previously reported that human USCs are pluripotent mesenchymal stem cells (MSCs) and possess osteogenic potential.34 When human USCs were transduced with Ad-BMP9, early osteogenic marker ALP activities were induced as early as day 3 and significantly increased at day 7, compared with those of the GFP control (Figure 4A). These results were further confirmed by quantitative ALP assays, in which BMP9 was shown to induce 14-fold and 26-fold increases in ALP activities at day 3 and day 7, respectively (Figure 4B).

Figure 4.

Figure 4

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Human urine stem cells (USCs) exhibit potent osteogenic potential upon BMP9 stimulation. (A, B) Early osteogenic marker alkaline phosphatase (ALP) activity induced by BMP9 in USCs. Subconfluent USCs were transduced with Ad-GFP or Ad-BMP9. Qualitative histochemical analysis (A) or quantitative measurement (B) of ALP activity was determined at days 3 and 7. Representative results are shown. (C) BMP9-induced mineralized matrix formation in USCs. Subconfluent USCs were transduced with Ad-GFP or Ad-BMP9, and cultured in mineralization medium for 14 days, followed by alizarin red S staining (a). Representative results are shown. The stained mineral nodules were dissolved and quantified (b). **p < 0.01, compared with that of the GFP group. (D) BMP9-induced osteogenic regulators and markers in USCs. Subconfluent USCs were infected with Ad-GFP or Ad-BMP9, and collected at 48h after infection. Total RNA was isolated from the infected cells and subjected to RT-PCR reaction, followed by TqPCR analysis of osteogenic regulators RUNX2 and OSX, and late osteogenic markers of OCN, OPN, and COL1A1. **p < 0.01, compared with that of the GFP group.

We also analyzed the capabilities of the USCs in mineralized matrix formation upon osteogenic stimulation. When the USCs were transduced by Ad-BMP9 and cultured in mineralization medium for 14 days, significant numbers of mineralized nodules were formed in BMP9-stimulated USCs, compared with those of the GFP control group (Figure 4C-a). Quantitative analysis showed that BMP9 induced at least a 4-fold increase in mineralized matrix formation, compared to that of the GFP control (Figure 4C-b). Furthermore, we analyzed the expression of osteogenic regulators and markers in BMP9-stimulated USCs. When USCs were transduced by Ad-BMP9 for 48h, we found that osteogenic transcriptional regulators RUNX2 and OSX, along with late osteogenic markers OCN, OPN and COL1A1, were significantly upregulated at the mRNA level by BMP9, compared with that of the GFP control group (Figure 4D). Collectively, these results demonstrate that human USCs exhibit osteogenic potential that can be effectively induced by BMP9. Therefore, human USCs and osteogenic factor BMP9 constitute two important components of the essential triad for bone tissue engineering.

The collective in vitro and in vivo evidence confirms that BMP9 may represent one of the best osteogenic factors for BTE although it is conceivable that BMP9 may synergize with other growth factors to further augment bone regeneration in the context of BTE. In this study, although we used only BMP9 to stimulate USCs, we achieved a desirable osteogenic effect. This result not only demonstrates the strong osteogenic differentiation potential of USCs but also confirms BMP9 as an appropriate osteogenic factor. Future studies should explore the interactions between BMP9 and other growth factors, such as vascular endothelial growth factors (VEGF) and fibroblast growth factors (FGF) and aim to construct a more conducive osteogenic microenvironment through sequential release strategies.

MSCs are considered the most reliable source for bone progenitor cells.3,15,65,66 Since their first discovery in the 1960s, bone marrow stromal cells (or BM-MSCs) are the most commonly used MSCs.67 However, for the past decades MSCs have been isolated from numerous tissues such as adipose tissue, oral cavity, cranial suture, skin basal layer, periosteum, vascular pericytes, and Wharton’s Jelly of umbilical cord.3,15,16,65,66 By comparing four types of commonly used mouse MSCs of mouse embryonic fibroblasts (MEF), mouse bone marrow MSCs, mouse calvarial MSCs and mouse adipose-derived MSCs, we demonstrated that adipose tissue-derived MSCs represent a superior source of progenitor cells for bone regeneration and BTE.26 While among the tested four types of MSCs, adipose tissue-derived MSCs are more abundant and readily available, adipose tissue procurement is invasive, and the recovery of viable MSCs is technically challenging and inefficient, posing major hurdles for effective BTE in clinical settings. The primary advantage of USCs over other stem cells lies in their ease of accessibility. USCs can be noninvasively isolated and collected from human urine, making the process significantly more straightforward compared to the extraction of other stem cell types, which is often more complex. In further clinical applications, the ability to conveniently and noninvasively extract USCs from a patient could address the issue of limited stem cell sources while also reducing the risk of immune rejection. On the other hand, we have demonstrated that the USCs are rich in not only osteoprogenitors but also other types of progenitors such as endothelial cells, smooth muscle cells, uroepithelial cells, and neuronal cells.17,18,34 Thus, human USCs should represent a first-line and reliable osteoprogenitor source for efficacious BTE and multitissue regeneration.

The mPOC-60HA Composite Is a Superior Scaffold for Bone Tissue Engineering with BMP9-Stimulated Human Urine Stem Cells

We further investigated whether mPOC-0HA, mPOC-40HA, or mPOC-60HA scaffold either alone and/or with human USCs and osteogenic factor BMP9, could provide a robust framework for bone tissue engineering. Experimentally, human USCs were transduced with Ad-BMP9 or Ad-GFP control, collected and mixed with thermoresponsive polymer PPCNg, and loaded onto mPOC-HA scaffolds immediately prior to subcutaneous implantation (Figure 5A). Specifically, three groups for each type of scaffold were designed: Group #1, mPOC-0/40/60HA only (scaffold only group or no cells group); Group #2, mPOC-0/40/60HA loaded with Ad-GFP-infected USCs (GFP group); and Group #3, mPOC-0/40/60HA loaded with Ad-BMP9 infected USCs (BMP9 group). As previously reported, the biocompatible thermoresponsive polymer PPCNg was added to further increase cell adhesion and even distribution within scaffolds.25

Figure 5.

Figure 5

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mPOC-60HA is a superior scaffold for in vivo bone tissue engineering. (A) Schematic representation of the subcutaneous implantation experiments of BMP9 or GFP-transduced USCs loaded onto mPOC-HA scaffolds. (B–D) Subconfluent USCs were transduced with Ad-GFP or Ad-BMP9, harvested at 16 h post infection, and loaded onto mPOC-0HA (B, panels b and c), mPOC-40HA (C, panels b and c) and mPOC-60HA (D, panels b and c) scaffolds (at 5 × 106 cells/implantation) immediately prior to subcutaneous implantation into the flanks of athymic nude mice (six implantations per group). Negative control for the mPOC-HA group (panela) contained scaffolds only (i.e., without USCs). Animals were sacrificed at 7 weeks after implantation. Subcutaneous masses containing the implants were retrieved for H & E analysis. Representative results are shown. PS, mPOC-0/40/60HA scaffolds; SC, seeded USCs; NB, new bone; and NBS, newly formed bone reinforced-scaffolds. (E) New bone formation was quantified with the ImageJ analysis of multiple high-power images in each group and is expressed as % new bone area over the total tissue area in the image. *p < 0.05, **p < 0.01, compared with that of the mPOC-0HA “no cells” (scaffold only) group (red asterisk), mPOC-0HA/USCs/GFP (blue asterisk), and mPOC-0HA/USCs/BMP9 (black asterisk) in the mPOC-0HA groups.

Approximately 5 × 106 of the adenovirus-transduced cells were loaded onto each scaffold, and 6 implants were carried out for each group/scaffold in the flanks of athymic nude mice. The animals were sacrificed 7 weeks after implantation. Subcutaneous masses were retrieved for histologic evaluation. H & E staining results indicate that new bone formation was rarely observed in the scaffold only groups although scattered osteoid matrix-like areas could be found in mPOC-40HA and mPOC-60HA groups (Figure 5BCD, panel a; Figure S1A). However, apparent osteoid matrix formation was found in all three types of scaffolds when human USCs were loaded onto the scaffolds, especially a more mature osteoid matrix appeared in the mPOC-60HA group (Figure 5BCD, panel b; Figure S1B), indicating that the mPOC-40/60HA scaffolds are osteoinductive. Lastly, in the scaffolds loaded with Ad-BMP9 transduced USCs, robust new bone formation was readily detected in all three types of scaffolds, and most new bone formation was found in the mPOC-60HA group (Figure 5BCD, panel c; Figure S1C). The new bone formation was further quantitatively determined, and the results showed that the mPOC-60HA/USCs/BMP9 group yielded the highest percentage (22.77%) of new bone formation, followed by the mPOC-40HA/USCs/BMP9 group (17.87%) (Figure 5E). Furthermore, for the mPOC-60HA scaffold, the newly formed bone was shown to extensively integrate into the edges of the scaffold and, in some cases, filled up the void space of the scaffold to form a solid bony structure (Figure S1C).

Modified PAS staining analysis revealed that, while robust bone formation was found in both mPOC-40HA and mPOC-60HA loaded with BMP9-transduced USCs groups, the mPOC-60HA supported most mature bone formation as evidenced by the decrease of purple blue staining along the edges of the scaffolds (Figure 6A panel a vs b). The newly formed bone was shown to extensively integrate into mPOC-60HA (Figure 6A panel b). Masson’s trichrome staining showed similar results, and most mature new bone was found in mPOC-60HA loaded with BMP9-stimulated USCs (stained in dark red in Figure 6B panel a vs b). Highly mineralized mature bone regions were also quantitatively determined in mPOC-40HA vs mPOC-60HA groups by using the ImageJ analysis; and we found that approximately 35% of the new bone formed in the mPOC-60HA/USCs/BMP9 group was mature bone, compared with approximately 18% of that in the mPOC-40HA/USCs/BMP9 group (Figure 6B panel c).

Figure 6.

Figure 6

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Bone mineralization specialty staining. The modified PAS staining (A) and Mason’s Trichrome staining (B) of the retrieved mPOC-40HA (panel a) and mPOC-60HA (panel b) samples from Figure 5. Representative results are shown. CM, chondroid matrix (purple blue in modified PAS staining); MB, mineralized bone (dark red in trichrome staining). (B panel c) Nature bone regions were quantitatively determined in mPOC-40HA vs mPOC-60HA groups by using the ImageJ analysis of multiple high-power images within the same group, and calculated as % of average mature bone area over the total new bone formation area. **p < 0.01, compared with that of the mPOC-40HA/USCs/BMP9 group.

The biomaterial utilized to scaffold and support tissue regeneration is the third essential factor for successful BTE. An ideal scaffold should possess desirable physicochemical-mechanical properties and biocompatibility that can effectively function as bone graft substitutes.1,19 For the past decades, numerous scaffolds have been made either from natural biomaterials such as collagens, hydroxyapatite, β-TCP, calcium phosphate cement, and ceramic glass,1,2 or from synthetic polymers, bioceramics, metals and composites.1,2,20 To fully take advantage of citric acid as an endogenous metabolite, we developed a series of CBPs including a biodegradable elastomer, POC,19,21−23 whose derivatives have recently been approved by FDA for orthopedic fixation applications. Citric acid can exhibit numerous cellular functions such as metabolic regulation, antimicrobial potential, mineralization regulation, anticoagulant effect, neuronal excitability regulation, and renal stone prevention.19,23 Furthermore, the presence of large number of functional overhanging carboxyl and hydroxyl groups renders the CBPs highly tunable, offering multiple functionalities such as incorporation of bioactive moieties, introduction of additional cross-link networks, as well as imparting intrinsic fluorescence, serving as a potent example of utilizing simple chemistry to endow functional complexity.19,23 Thus, more novel materials with various physical properties including elasticity, mechanical strength, and degradation rate can be generated for BTE.19,23

Our modifications to POC proceeded in two stages. First, conjugation of methacrylate groups onto the prepolymer, enhancing its photo-cross-linking properties for easier application in 3D printing. Additionally, we incorporated varying concerntration of HA. Our in vivo studies showed that while mPOC scaffolds mixed with 0%, 40%, or 60% HA, and loaded with BMP9-stimulated USCs, effectively promoted new bone formation, histological evaluation revealed that the mPOC-60HA composite scaffold produced the most mature bone, closely resembling native tissue, with significant osteointegration between new bone and the scaffold. This approach overcomes the limitations of using HA alone as a scaffold—such as brittleness, poor toughness, susceptibility to fatigue fractures, prolonged biodegradation, and structural simplicity—which hinder cell survival and vascularization. It also enhances the properties of the POC scaffold by adding HA, which improves osteoconduction and promotes cell–tissue interactions.

This study has several limitations. First, we did not screen different osteogenic factors to assess their potential in stimulating osteogenic differentiation in USCs, and there may be other individual or combined factors that could be more effective than BMP9. Second, the scaffold size was a small disk structure, which is not suitable for subsequent studies that aim to be more clinically relevant. Third, our study was limited to investigating the osteogenic effect after subcutaneous implantation without addressing the mechanical functions provided by the scaffold such as whether the scaffold could deliver the necessary mechanical stress for fracture defect repair. In future studies, screening for the best osteogenic stimulation combination for USCs and redesign of the scaffold structure will be essential.

Collectively, the in vivo results revealed that while the mPOC-0/40/60HA scaffolds alone exhibited varied and limited osteoinductive capabilities, the addition of Ad-GFP transduced USCs to these scaffolds enhanced ectopic ossification, especially for the mPOC-40/60HA scaffolds. When the Ad-BMP9 infected USCs were loaded onto the mPOC-40HA and mPOC-60HA scaffolds, more robust bone formation and well-mineralized bone matrix were found to fill up the space inside the mPOC-60HA scaffolds, leading to the generation of solid bone tissue and preservation of the scaffold configuration. Thus, these in vivo findings strongly suggest that mPOC-60HA may be a more desirable scaffold for in vivo bone tissue engineering.

Conclusions

The objective of this work was to fabricate and evaluate the osteogenic potential of a 3D-printed polymer-ceramic composite scaffold laden with USCs stimulated with BMP9. CBP composites of mPOC with various mass percentages of HA (i.e., mPOC-HA, mPOC-40HA, or mPOC-60HA) were prepared to fabricate scaffolds via additive manufacturing using mCLIP. Our results showed that the 3D-printed mPOC-HA composite scaffolds were compatible with USCs and USCs exhibited high osteogenic capacity in vitro and in vivo due to BMP9 stimulation. Specifically, subcutaneous implantation of mPOC and composite scaffolds laden with BMP9-stimulated USCs effectively formed new bone in all three types of mPOC-HA composite scaffolds. Histologic evaluation revealed that the mPOC-60HA composite scaffold yielded the most mature new bone resembling native bone tissue with extensive osteointegration between new bone and scaffolds. Collectively, our findings demonstrate that citrate-based mPOC-60HA composite, human urine stem cells, and the potent osteogenic factor BMP9 may constitute an optimal triad for effective bone tissue engineering. Future studies involve implementation of this tripartite strategy in both small and large animal bone defect models.

Acknowledgments

The reported work was supported in part by research grants from the National Institutes of Health (CA226303 to TCH and DE030480 to RRR), and the National Research Foundation of Korea (2021R1A6A3A14039205 to MK). This work made use of the IMSERC NMR and Physical Characterization facility at Northwestern University, which has received support from the Soft and Hybrid Nanotechnology Experimental (SHyNE) Resource (NSF ECCS-2025633), Int. Institute of Nanotechnology, and Northwestern University. This project was also supported in part by The University of Chicago Cancer Center Support Grant (P30CA014599) and the National Center for Advancing Translational Sciences (NCATS) of the National Institutes of Health through Grant Number 5UL1TR002389. TCH was supported by the Mabel Green Myers Research Endowment Fund and The University of Chicago Orthopaedics Alumni Fund. Funding sources were not involved in the study design; in the collection, analysis and interpretation of data; in the writing of the report; and in the decision to submit the paper for publication.

Data Availability Statement

All data are available in the main text or the Supporting Information.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsami.4c13246.

  • List of TqPCR primers; list of abbreviations; representative images of mPOC-60HA scaffold-based bone tissue fabrication (PDF)

Author Contributions

† PZ and YZhu contributed equally to this paper. RRR, TCH, GAA, LW, YZhu, PZ, CS conceived and designed the study. MK, CPC, CD, CS, GAA synthesized and characterized the scaffold materials. YZhang and YZhu isolated and characterized human urine stem cells. PZ, YZhu, GZ, YW, OM, JZ performed in vitro and in vivo experiments and collected data. YZhu, GZ, YW, JZ, TCH performed histologic processing and statistical analysis. HZ, WY, GS, CL, XW, JL, YS, JF participated in experiments, provided essential experimental materials; and/or assisted in data analysis and interpretations. YZhu, RRR, TCH, GAA, SC, HHL, LLS, LW, MJL, RCH drafted and revised the manuscript. All authors reviewed and approved the manuscript.

The authors declare no competing financial interest.

Supplementary Material

am4c13246_si_001.pdf (404.2KB, pdf)

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