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. 2024 Jan 4;16(2):1999–2011. doi: 10.1021/acsami.3c12575

Piezoelectrically and Topographically Engineered Scaffolds for Accelerating Bone Regeneration

Soyun Joo †, Yonghyun Gwon ‡,§,∥, Soyeon Kim †, Sunho Park ‡,§, Jangho Kim ‡,§,∥,*, Seungbum Hong †,⊥,*
PMCID: PMC10798259  PMID: 38175621

Abstract

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Bone regeneration remains a critical concern across diverse medical disciplines, because it is a complex process that requires a combinatorial approach involving the integration of mechanical, electrical, and biological stimuli to emulate the native cellular microenvironment. In this context, piezoelectric scaffolds have attracted considerable interest owing to their remarkable ability to generate electric fields in response to dynamic forces. Nonetheless, the application of such scaffolds in bone tissue engineering has been limited by the lack of a scaffold that can simultaneously provide both the intricate electromechanical environment and the biocompatibility of the native bone tissue. Here, we present a pioneering biomimetic scaffold that combines the unique properties of piezoelectric and topographical enhancement with the inherent osteogenic abilities of hydroxyapatite (HAp). Notably, the novelty of this work lies in the incorporation of HAp into polyvinylidene fluoride-co-trifluoro ethylene in a freestanding form, leveraging its natural osteogenic potential within a piezoelectric framework. Through comprehensive in vitro and in vivo investigations, we demonstrate the remarkable potential of these scaffolds to accelerate bone regeneration. Moreover, we demonstrate and propose three pivotal mechanisms—(i) electrical, (ii) topographical, and (iii) paracrine—that collectively contribute to the facilitated bone healing process. Our findings present a synergistically derived biomimetic scaffold design with wide-ranging prospects for bone regeneration as well as various regenerative medicine applications.

Keywords: ferroelectric scaffolds, piezoelectric and topographical cues, P(VDF-TrFE) composites, hydroxyapatite, bone regeneration

Introduction

Bone regeneration is a complex process that requires a combinatorial approach with multiple modulating factors, including mechanical,1 electrical,2 and biological3 cues. Current strategies for bone regeneration, such as autologous and allogeneic bone grafting4 or growth factor delivery,5 have limitations such as donor site morbidity,6 limited availability,7 and high cost.8 More importantly, such complications have led to unsatisfactory therapeutic results, and it is necessary to develop novel, effective approaches for bone regeneration based on the understanding of the mechanisms that underlie osteogenic cues. Among the multiple strategies, an emerging promising area of research involves the use of piezoelectricity to stimulate the growth of new bone tissue.

Piezoelectric scaffolds, such as polyvinylidene fluoride-co-trifluoro ethylene (P(VDF-TrFE)), have been increasingly investigated as a means to promote bone regeneration due to their ability to generate signals in response to mechanical stress. For example, Adadi et al.9 demonstrated cardiac tissue alignment and differentiation effects on electrospun fibrous P(VDF-TrFE) scaffolds. Similarly, Zhang et al.10 reported that a piezoelectric scaffold made from P(VDF-TrFE) and barium titanate (BTO) could accelerate bone regeneration and improve osseointegration in a rat calvarial defect. Multiple works have additionally highlighted the osteogenic effects of such piezoelectric composites due to their unique combination of mechanical, electrical, and biocompatible properties.11−15 However, novel technologies utilizing piezoelectric materials for bone regeneration still fall short of replicating the intricate cellular environments for bone tissue regeneration. Consequently, there is a need for advanced methods that can effectively optimize and faithfully mimic such environments.16−18 Importantly, current research in this field faces several challenges, including the lack of a comprehensive understanding of the optimal piezoelectric properties required to promote bone growth as well as the difficulties associated with incorporating appropriate fillers to enhance material performance as tissue engineering scaffolds.

Another area of research in bone regeneration focuses on the role of surface topography in stimulating cellular activity. Previous studies have shown that surface modification can influence cell adhesion, proliferation, and differentiation, which are critical factors for the successful integration of implants with the host tissue. For example, Gittens et al.19 demonstrated enhanced osteoblast differentiation on surface-modified Ti surfaces with superimposed nanofeatures compared to pristine Ti. Similarly, Zhang et al.20 revealed improved proliferation and adhesion of rat bone marrow-derived mesenchymal cells on PVDF surfaces with nanostructured topographic features. However, work in this area also has limitations, including the lack of consensus regarding the mechanism by which surface coarseness affects cellular behavior and the challenge of developing materials that combine surface roughness with piezoelectricity.

Regarding this aspect, we may consider hydroxyapatite (HAp), a naturally occurring component of bone that offers various benefits in the context of bone regeneration. Studies have found that HAp enhances osteogenic activity, improves osseointegration, and provides a scaffold for new bone growth.21−25 Its inherent biocompatibility due to its similarity to hard tissue ensures low systematic toxicity and high acceptance by the human body.25 Most importantly, HAp possesses both piezoelectric and surface roughness properties, making it an ideal candidate for use in piezoelectric scaffolds with modulated roughness for bone growth applications. However, although the biocompatibility of HAp in bone regeneration has been extensively studied, its implementation in the form of a flexible, piezoelectric scaffold has not yet been reported.

Here, we present a pioneering approach in fabricating a freestanding scaffold, integrating HAp within the piezoelectric framework of P(VDF-TrFE), marking a departure from prior methodologies primarily focused on surface coatings. Unlike the conventional confines of HAp and the P(VDF-TrFE) combination, which have previously been limited to coatings on metallic prosthetics, our study introduces a technique that develops an independent scaffold, offering a versatile platform for bone regeneration beyond surface-bound applications (Figure 1a). Furthermore, we unveil the underlying factors contributing to the heightened piezoelectric, topographic, and paracrine functionalities within the HAp/P(VDF-TrFE) scaffold, providing a holistic understanding. This elucidation allows us to identify synergistic mechanisms driving the enhanced effects, establishing a foundation crucial for future advancements in biomaterial design. With demonstration of its excellent mechanical, electrical, and biomimetic properties, we present HAp/P(VDF-TrFE) as a new material with potential in various applications for treating bone disorders.

Figure 1.

Figure 1

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Design and characterization of piezoelectrically and topographically originated biomimetic scaffolds. (a) Schematic representation of the enhanced bone regeneration mechanism through electrical and topographical cues provided by HAp-incorporated P(VDF-TrFE) scaffolds. (b) Schematic diagram of the fabrication process. (c) SEM, (d) EDS, and (e) XRD results of composite scaffolds. (f) FT-IR results of HAp/P(VDF-TrFE) of various HAp volume percentages. (g) Piezoelectric coefficients (n = 20) and (h) stress–strain curves of composite scaffolds.

Results and Discussion

Design and Characterization of P(VDF-TrFE)-Based Scaffolds

The design of a piezoelectric bone scaffold involves careful consideration across multiple factors, including the use of piezoelectric materials to stimulate cell growth. Piezoelectric ceramics and polymers have been widely explored in the context of scaffold creation, as highlighted in multiple references.2,11,20,26,27 The piezoelectric coefficients of bones and other bodily components have also been measured,28 although their specific purpose remains largely unknown. One theory suggests that the piezoelectricity of natural materials generates charge under stress, promoting cell growth, while another theory proposes streaming potentials as the main explanation, involving the flow of charged fluid driven by bone movement.29 For preceding studies, ceramic BTO has been most commonly studied in conjunction with PVDF-based polymers.10,26,27,30,31

On the other hand, composite membranes of PVDF-based polymers with HAp have been studied less frequently, mainly as coating materials for metal implants rather than free-standing bone scaffolds. For example, a cytotoxicity assay conducted on PVDF and HAp composites demonstrated successful growth of NCTC clone 929 cells, indicating the absence of toxicity.32 In another study, such a composite was created using a pneumonic spray nozzle, and optimal conditions for mesenchymal stem cell growth were established.33 Notably, these studies did not develop self-standing scaffolds that could be inserted into in vivo environments nor did they examine the causes or effects of enhanced bone regenerative effects. In this context, the investigation of P(VDF-TrFE) in conjunction with HAp is particularly relevant in the design of piezoelectric scaffolds. Biocompatible HAp provides enhanced bioactivity and osteogenesis, while the ferroelectric properties of P(VDF-TrFE) additionally enable the generation of electrical charge and potential in response to exterior material stress, promoting enhanced cellular activity and bone remodeling. To achieve mechanical stimulation, the scaffold is engineered to transmit mechanical forces between the bone and soft tissue.

To confirm the bone regenerative abilities of P(VDF-TrFE) containing HAp compared to previous publications, we fabricated three types of scaffolds: P(VDF-TrFE), BTO/P(VDF-TrFE), and HAp/P(VDF-TrFE). In addition to plain P(VDF-TrFE), we selected BTO as a comparative material in our study based on its well-documented utilization as a piezoelectric material in various biomedical applications, particularly in the realm of tissue engineering.10,26,27,30,31 Notably, BTO has been commonly mixed with P(VDF-TrFE) to form composite materials employed in bone regeneration studies, leveraging its piezoelectric properties to augment the functionality of scaffolds for enhanced cellular response and tissue regeneration. Its widespread usage in previous research endeavors serves as a crucial reference point for evaluating the efficacy and performance of our developed HAp/P(VDF-TrFE) scaffold. The processing sequence of these scaffolds is schematically depicted in Figure 1b, with details regarding the fabrication process outlined in Methods, Supplementary Figure 1, and Supplementary Note 1. We note that both the β phase of P(VDF-TrFE) and excellent dispersion of filler nanoparticles are essential factors in enhancing the bone healing effect of P(VDF-TrFE)-based composites. Rod-like grains—the lamellar structure—were observed in all three scaffold types, and we confirmed the homogeneous dispersion of filler particles for both HAp/P(VDF-TrFE) and BTO/P(VDF-TrFE) (Figure 1c). Energy-dispersive X-ray spectroscopy (EDS) analysis of the points of these circular regions also demonstrates that these particles are indeed incorporated filler particles, as intended (Figure 1d). The elemental set of energy peaks of Ca and P (or Ba and Ti) affirms the presence of HAp (or BTO) nanoparticles in the composite scaffolds. C and F, expressed by the P(VDF-TrFE) matrix and Os, which was sputtered on the surface for conductivity, were commonly observed in all three spectra. We additionally performed XRD analysis to analyze the crystalline phase of the scaffolds for the 2θ range from 10° to 60° at room temperature. The large peak at 19.9° indicates high β-phase crystallinity in all samples, while the set of smaller peaks implies the existence of HAp (or BTO) nanoparticles in HAp/P(VDF-TrFE) (or BTO/P(VDF-TrFE) (Figure 1e). Identification of the smaller peaks, marked by asterisks, is shown in Supplementary Figure 2.

By varying the HAp volume percentage, a set of composite scaffolds (0, 1, 3, 9, and 12 vol %) with an average thickness of 16–18 μm were obtained (Supplementary Figure 3). To check the ferroelectric phase formation and confirm the effects of filler incorporation in the HAp-containing scaffolds, Fourier transform infrared (FTIR) analysis was conducted for wavenumbers from 1600 to 400 cm–1 (Figure 1f). In FTIR, two representative peaks are associated with each of the three axes of the P(VDF-TrFE) crystal. The pair of absorption peaks at 1184 and 890 cm–1 corresponds to the a-axis, that at 1284 and 848 cm–1 corresponds to the b-axis, and the pair of 1400 and 1078 cm–1 corresponds to the c-axis.34−37 Correspondingly, the three peaks at 1400, 1284, and 848 cm–1 are associated with the ferroelectric β phase—the first peak at 1400 cm–1 represents the CH2 wagging vibration with polarization aligned along the c-axis, and the 1284 and 848 cm–1 peaks represent the CF2 symmetric stretching with polarization parallel to the polar b-axis. Peaks at 1284 and 848 cm–1 indicate trans sequences longer than TTT and TTTT, respectively.37 The gray background in Figure 1f highlights these sets of β phase markers. It is notable how peak bands were nearly at the same position with similar peak intensity regardless of the incorporated HAp vol %, indicating that the β-phase molecular chain structure does not change with the addition of fillers. Moreover, the set of spectra obtained from scaffolds containing HAp (1, 3, 9, and 12 vol %) exhibited increasing intensity along peaks of 1048, 600, and 570 cm–1.38 These peaks, highlighted by the yellow background, correspond to the intensive absorption bands formed by the PO43– chemical groups that compose HAp. Based on these results, we may thus confirm that the HAp/P(VDF-TrFE) composite scaffolds consisted of the crystalline β phase with finely integrated HAp filler particles.

Synergistically Enhanced Bone Tissue Regeneration with Piezoelectric and Topographical Cues In Vitro and In Vivo

P(VDF-TrFE) scaffolds mostly composed of the ferroelectric phase would exhibit limited macroscopic piezoelectricity without a proper high-voltage poling treatment. That is, the electromechanical performance of a ferroelectric material—commonly evaluated by the piezoelectric coefficient (d33)—arises from the net dipole moment (remnant polarization) after exposure to a sufficiently strong external field. We thus adopted a corona poling method on the composite scaffolds by locating a high voltage metal needle a few centimeters above the sample with the bottom electrode connected to the ground (Figure 1b). In this process, ionization of the air surrounding the needle tip creates charged ions that are collected along the surface and form a poling electric field.13 As confirmed with measurements on the piezometer, all samples exhibited zero d33 values before poling, whereas specific nonzero values were measured afterward.

The measured d33 values of the poled scaffolds are shown in Figure 1g, where we found that nanoparticle inclusion enhanced electrical polarization in P(VDF-TrFE) for both cases of HAp and BTO (n = 20). This increase may be explained by the inherent piezoelectricity of the fillers or the state of dispersion within the matrix that leads to localized field concentration, which in turn induced enhanced piezoelectricity in the composite scaffolds.39,40 Moreover, the differences between the d33 values of 3, 9, and 12 vol % HAp/P(VDF-TrFE) suggest that there is an appropriate volume percentage at which the polarizability is the most improved. After observing the improved piezoelectric properties of composite scaffolds containing 9 vol % filler particles, subsequent experiments in this study mainly focused on this material, unless stated otherwise. Enhanced piezoelectricity was found in scaffolds containing 9 vol % BTO in comparison to those with 9 vol % HAp. These findings align consistently with prior research, where higher piezoelectric properties have been reported for bulk BTO compared to HAp.41,42 Further investigation regarding these effects is examined again in the latter part of this work. The mechanical properties of the scaffolds were measured using a strain–stress tester. The tensile stress of the HAp/P(VDF-TrFE) scaffold was 2 times and 1.5 times greater than that of the P(VDF-TrFE) and BTO/P(VDF-TrFE) scaffolds, respectively (Figure 1h).

Cell attachment on HAp/P(VDF-TrFE) scaffolds was 15 and 10% higher than that on BTO/P(VDF-TrFE) and P(VDF-TrFE) scaffolds, respectively. In addition, after 5 days of cell culture, cell proliferation was 20 and 30% higher on the HAp/P(VDF-TrFE) scaffolds than on the BTO/P(VDF-TrFE) and P(VDF-TrFE) scaffolds, respectively (Figure 2a). These findings suggest that the topographical origin of HAp can create an environment similar to that of the intricate extracellular matrix (ECM), thereby promoting osteoblast attachment and proliferation. Additionally, we evaluated the osteogenic mineralization of osteoblasts by culturing cells on the scaffolds in an osteogenic induction medium for 7 and 14 days (Figure 2b). Alkaline phosphatase (ALP) staining and Alizarin Red staining revealed approximately 30 and 40% higher osteogenic levels on HAp/P(VDF-TrFE) scaffolds than on BTO/P(VDF-TrFE) and P(VDF-TrFE), respectively, where calcium deposition on HAp-incorporated scaffolds was quantified considering the intrinsic calcium content of HAp (Figure 2c). These results indicate that HAp maximizes the scaffold’s piezoelectric properties and provides a topographical effect akin to the ECM. The synergistic influence of these origins enhances osteoblast attachment, proliferation, and differentiation. A comprehensive examination of these effects will be covered in the following section.

Figure 2.

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Synergistically enhanced bone tissue regeneration with piezoelectric and topographical origin in vitro and in vivo. (a) Cell proliferation, (b) cell osteogenic differentiation, and (c) osteogenic quantification analysis of the three types of scaffolds. (d) In vivo bone regeneration micro-CT analysis and (e) quantification of bone volume and area at 2, 4, and 6 weeks (scale bars: 2.5 mm). (f) Hematoxylin and eosin (H&E) staining and (g) Masson’s trichrome (M&T) staining of bone tissue after 6 weeks.

In addition, we confirmed the effects of piezoelectric and topographical origins by HAp throughout bone regeneration in vivo (Figure 2d). Scaffold patches were placed to cover the defects and intricately created on the calvarial bone using an electrical drill. All mice used in the in vivo studies survived to the date of sacrifice, and no adverse events were observed. No infection or inflammatory response was observed in any mice during the postoperative period, and the implanted scaffolds were maintained for 6 weeks without deformation. To quantitatively evaluate the effects of piezoelectric and topographical origins on bone formation, we performed micro-CT and 3D-image conversion using MIMICS 14.0 software on new bone defects in vivo (Figure 2e). After 2, 4, and 6 weeks, compact bone formation was not observed in the defect group and P(VDF-TrFE) scaffolds, whereas bone regeneration was significantly enhanced in the BTO/P(VDF-TrFE) and HAp/P(VDF-TrFE) scaffolds after 2, 4, and 6 weeks of implantation. To confirm the bone regeneration efficacy from piezoelectric and topographical origins, we conducted H&E and M&T staining at 6 weeks after implantation (Figure 2f,g). As a result, the bone of the defect groups was empty in the defect area, while accelerated bone formation and dense cytoplasm in HAp/P(VDF-TrFE) scaffolds compared with other groups were confirmed.

In the subsequent sections of this work, we delve into a comprehensive discussion of the underlying mechanisms and origins that contribute to the enhanced bone regeneration effects observed with the HAp/P(VDF-TrFE) scaffolds. First, we explore the electrical origin of enhanced regeneration, supported by cell expression analysis and investigations into the piezoelectric properties of the scaffold, enabled by piezoresponse force microscopy (PFM) and Kelvin probe force microscopy (KPFM). We next focus on the topographical origin, analyzing the three-dimensional surface of the composite scaffolds along with cell morphology and vinculin formation. These studies elucidate the influence of surface topography on cellular behavior and tissue regeneration. Last, we explore the paracrine origin, investigating the effects of piezoelectric and topographical origins on the growth factor expression. Our examination of these distinct yet interconnected aspects aim to provide a comprehensive understanding of the multifaceted mechanisms responsible for the synergistic enhancement of bone regeneration observed with HAp/P(VDF-TrFE) scaffolds. Moreover, these findings suggest new avenues for future research, encouraging exploration to leverage the inherent properties of HAp to enhance the functionalities of the commonly utilized P(VDF-TrFE).

Electrical Origin of Biomimetic Scaffolds for Enhanced Bone Tissue Regeneration

To further investigate the underlying mechanism of enhanced bone regeneration by HAp/P(VDF-TrFE) scaffolds, piezoresponse force microscopy (PFM) studies were conducted to examine the dielectric and surface properties. A typical method to examine sample ferroelectricity is to image the PFM amplitude and phase after applying an external switching bias that exceeds the coercive voltage.43 We thus polarized the as-cast composite scaffolds by applying −50 V to a square region and then applying +50 V to a smaller square region, according to the box poling pattern and steps specified in Supplementary Figure 4b. The results shown in Figure 3a indicate the highest electromechanical response (indicated by the strong PFM amplitude within the poled region) and switchability (indicated by the clear contrast in the PFM phase) in 9 vol % HAp/P(VDF-TrFE).

Figure 3.

Figure 3

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Electrical origin of biomimetic scaffolds for enhanced bone tissue regeneration. (a) PFM amplitude and phase images of box-poled composite scaffolds. The white bar represents 2 μm. (b) Histogram of the PFM phase within the innermost box of box-poled HAp-based scaffolds in panel (a). (c) XRD results of scaffolds and (d) schematic representation of filler-derived electrical origins in bone regeneration. (e) Surface potential measurements of scaffolds. (f) SEM and Ca EDS images of cells adhered to sample surfaces. (g) Quantitative EDS analysis of cell Ca expression.

Also worth noting is the improved switchability with increasing levels of HAp incorporation. This trend may be observed by both visual inspection of the PFM phase images and comparison of the PFM phase distribution of the innermost box (Figure 3b), the region that has been once switched up (−50 V) and then down again (+50 V). These results align with the earlier measurements, where higher d33 was measured for increasing amounts of HAp. Meanwhile, the box poling results of P(VDF-TrFE) do not exhibit distinct poled domains. While a common cause of failure to observe switched domains in ferroelectric samples is insufficient switching voltage, this was not the case, as domain switching was not observed even at greater magnitudes of applied bias. Another factor to consider is the scaffold thickness, since the electric field is inversely proportional to distance. Indeed, when thinner scaffolds of a few hundred nm thickness were prepared, switched domains were clearly observed. However, these thin films could not be peeled off the substrate while maintaining mechanical integrity and were therefore inadequate for bone scaffold applications. Further discussions regarding the dielectric properties of P(VDF-TrFE) scaffolds are detailed in Supplementary Note 2.

Box poling was also performed on 9 vol % BTO/P(VDF-TrFE) to determine whether enhanced switching could be observed for composites that contain fillers of the same volume proportion. However, BTO-containing scaffolds did not exhibit poled domains even after the same process (Figure 3a). These results seemingly contradict the XRD results shown in Figure 3c, as scaffolds containing BT fillers exhibit clearer peaks corresponding to the ferroelectric P(VDF-TrFE) β phase. However, these results may be comprehensively explained by our filler model (Figure 3d). HAp nanoparticles exhibit a lower density (3.14 g/mL) than BTO (6.08 g/mL) and result in well-dispersed states within the viscous P(VDF-TrFE) scaffold in the melt state.

Additionally, ferroelectric BTO exhibits a very high dielectric constant (measured to be in the range of 100 to 10,000 depending on frequency and temperature), which would result in randomly aligned dipoles that reduce switchability and the resulting piezoelectric coefficient. On the other hand, induced dipole effects that lead to improved piezoelectricity can be expected from piezoelectric HAp with its relatively lower dielectric constant (measured to be in the range of 10 to 20). The importance of charged surfaces for inducing bone growth has also been repeatedly emphasized.44,45 We thus analyzed the surface potential of the composites by Kelvin probe force microscopy (KPFM), and the results are shown in Figure 3e. Gaussian fitting of the obtained data reveals that 9 vol % HAp/P(VDF-TrFE) exhibits the lowest range of potential distribution centered at approximately −0.39 V compared to P(VDF-TrFE) (approximately −0.24 V). The 9 vol % BTO/P(VDF-TrFE) exhibits a distribution between these values (−0.27 V), although the distribution is much wider. While we will not discuss this broadening in depth, it may be due to the local electric polarization in the interfacial region between BTO and P(VDF-TrFE).46 Nevertheless, these results suggest that the more negative surface of 9 vol % HAp/P(VDF-TrFE) would allow favorable attachment of positively charged ions (Mg2+ and Ca2+) and support the results of accelerated bone growth in Figure 2.

In fact, we observed enhanced Ca expression in HAp-containing scaffolds when cells adhered to the scaffold surfaces (Figure 3f). Quantification of the EDS results further demonstrated the highest degree of Ca expression by cells adhered to the HAp/P(VDF-TrFE) scaffolds (Figure 3g). These results are additionally supported by our model to illustrate the filler-derived electrical origins in bone regeneration. By the effects of induced dipoles and enhanced piezoelectric properties with incorporation of HAp nanoparticles, more negatively charged surfaces would permit preferential ion expression and lead to accelerated bone regeneration, as shown by the existing results.

Topographical Origin of Biomimetic Scaffolds for Enhanced Bone Tissue Regeneration

The surface properties of the scaffolds were examined by topographical scans, which showed significant differences in texture. Three-dimensional (3D) representations of the surfaces and 2D line section representations are shown in Figure 4a. Surface roughness was further quantified by comparing the arithmetic mean height (Ra) and the root-mean-square deviation values (Rq), where HAp/P(VDF-TrFE) (Ra, Rq: 15.65 nm and 18.07 nm) exhibited greater roughness than 9 vol % BTO/P(VDF-TrFE) (Ra, Rq: 8.01 nm, 8.08 nm). We may thus visually and numerically confirm that with the addition of fillers, HAp containing P(VDF-TrFE) increasingly exhibits higher roughness than its counterparts.

Figure 4.

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Topographical origin of biomimetic scaffolds for enhanced bone tissue regeneration. (a) 3D representations of composite scaffolds paired with typical 2D line sections. (b) Cell morphology and (c) quantification analysis of the three types of P(VDF-TrFE) scaffolds. (d) Focal adhesion analysis by vinculin staining. (e) Schematic representation of filler-derived topographical origins in bone regeneration.

As shown in Figure 4b, the change in surface roughness induced by HAp greatly influenced the cells, as evidenced by the spreading cytoskeletal structure on the substrate. The individual cells on HAp/P(VDF-TrFE) exhibited a spreading morphology due to the high density of HAp on the scaffold surface. In addition, the cells adhered closely along the direction of the substrate. Specifically, the long axis and elongation factor of the cells on HAp/P(VDF-TrFE) were reduced compared to those on P(VDF-TrFE) and BTO/P(VDF-TrFE) (Figure 4c). In addition, the cell perimeter and spreading area were 60% higher in cells on HAp/P(VDF-TrFE) than in those without HAp. The elongation factor and shape index showed a negative correlation, indicating that the cytoskeletal structure of cells on the incorporated HAp scaffold can interact closely to regulate the cellular behavior. Furthermore, we observed that the focal adhesions (FAs) of cells were polarized along the high density of HAp on the scaffold surface (Figure 4d), being more polarized with the high density of HAp than the relatively low density. The high density of HAp affected the size of FAs such that larger FA sizes were observed with the low density of HAp.

Here, we again highlight the contrast in the particle densities of HAp and BTO nanofillers. Whereas most of the BTO particles would sink to the bottom of the film during solution casting, lighter HAp particles would be distributed evenly within the films and be expressed on the surface by protrusions that offer roughness suitable for cellular functions, including cell adhesion, proliferation, and osteogenic differentiation (Figure 4e). Based on this model, we can comprehensively understand both the limited switching of 9 vol % BTO/P(VDF-TrFE) found in Figure 3a and the less defined surface roughness in Figure 4a to be the consequence of particle distribution throughout the matrix, mainly affected by filler density. These results are in close agreement with other reports, wherein microrough implant surfaces are suggested to trigger enhanced osseointegration compared to smooth surfaces.47,48 In comparing the results for the particle containing scaffolds versus P(VDF-TrFE), we may additionally note the possible impact of surface morphology changes on the hydrophilic–hydrophobic properties of the scaffolds and their subsequent impact on cell adhesion behavior. The inclusion of hydrophilic components, such as HAp and BTO particles, within the inherently hydrophobic P(VDF-TrFE) matrix can enhance the overall surface hydrophilicity. This alteration is advantageous for promoting cell attachment, given the preferential affinity of cells toward hydrophilic surfaces.

Paracrine Origin of Biomimetic Scaffolds for Enhanced Bone Tissue Regeneration

It is known that cells generate autocrine and paracrine factors, which are crucial molecules for signaling the activation of various cellular functions. The secretion of growth factors by cells cultured on the three types of scaffolds was analyzed. The osteoblasts on the HAp/P(VDF-TrFE) scaffold secreted higher (upregulation) levels of basic fibroblast growth factor (bFGF), bone morphogenetic protein-4 (BMP-4), BMP-7, epidermal growth factor (EGF), epidermal growth factor receptor (EGFR), fibroblast growth factor-4 (FGF-4), FGF-7, glia cell-derived neurotrophic factor (GDNF), hepatocyte growth factor (HGF), osteoprotegerin (OPG), transforming growth factor-α (TGF-α), TGF-β1, TGF-β3, and vascular endothelial growth factor A (VEGF-A) (Figure 5a); however, the osteoblasts on the HAp/P(VDF-TrFE) scaffold secreted lower (downregulation) levels of growth differentiation factor-15 (GFD-15), insulin-like growth factor binding protein-1 (IGFBP-1), IGFBP-3, placental growth factor (PLGF), and VEGF-D (Figure 5b). Specifically, we investigated the relative expression of 6 major growth factors (bFGF, BMP-4, BMP-7, TGF-α, TGF-β1, and TGF-β3) related to bone regeneration. The relative expression of all six growth factors increased on HAp/P(VDF-TrFE), and the relative expression of BMP-4, BMP-7, TGF-α, and TGF-β3 increased more than 1.5 times.

Figure 5.

Figure 5

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Paracrine origin of biomimetic scaffolds for enhanced bone tissue regeneration. (a) Upregulation and (b) downregulation of growth factors on the three types of P(VDF-TrFE) scaffolds. (c) Relative expression of key growth factors during bone regeneration. (d) Schematic representation of the possible mechanism by which Hap/P(VDF-TrFE) affects bone regeneration.

Here, we propose that the mechanism underlying the bone regeneration effect mediated by the HAp/P(VDF-TrFE) scaffold is based on the following key factors: (i) promotion of Ca2+ and growth factor expression through the enhancement of piezoelectric origin by HAp—at this point, the host tissue cells (osteoblast and stem cells) are affected by the piezoelectrical signal, which is a similar environment to bone tissue in vivo, migrating the cells to the defect site; (ii) increase of cell adhesion protein and control of cell morphology through enhancement of topographical origin by HAp—here, the host tissue cells are affected by the surface roughness similar to bone tissue. Through this stimulation, the morphology of host cells is controlled and the expression of cell adhesion protein (i.e., vinculin) is increased, which enhances cell proliferation and osteogenic differentiation. This brings us to the third factor: (iii) increased expression of growth factors related to bone regeneration due to the synergistic effect of the piezoelectric and topographical origins by HAp. The regulation of cellular behavior and functions occurs through multifaceted enhancement of origins, which promote cell growth factor expression related to bone regeneration.

From this standpoint, our in vitro study demonstrates that the morphology of the osteoblast was controlled and the functions of these cells were improved by the piezoelectric and topographical origins in the scaffold. The origins of these improvements in cellular functions lie in the piezoelectric and topographical properties of the scaffolds, which mimic the electrical and topographical environment of bone tissue during bone regeneration.

Conclusions

This study describes the development of piezoelectrically and topographically engineered biomimetic scaffolds that accelerate the process of bone regeneration. Our scaffolds successfully demonstrated enhanced cellular functionalities (cell proliferation and osteogenic differentiation) in vitro and bone regeneration effects in vivo owing to their optimized piezoelectric and topographical characteristics that emulate the native bone tissue microenvironment. Furthermore, we elucidated key factors underlying the bone regeneration effect of our biomimetic scaffolds, which encompasses (i) the acceleration of bone regeneration through elevated Ca2+ expression resulting from the uniform and high piezoelectric properties by HAp, (ii) the promotion of bone regeneration via the regulation of cell behavior and function facilitated by the topographical changes on the scaffold surface by HAp, and (iii) the synergistic modulation of growth factors enabled by the combined piezoelectric and topographical properties of the HAp scaffold. This research not only contributes to a deeper understanding of the impact of piezoelectric and topographical cues on bone regeneration but also proposes a viable strategy for enhancing the performance of synergistically designed biomimetic scaffolds for implementation in bone tissue engineering applications.

Methods

Preparation of Scaffolds

To fabricate P(VDF-TrFE)-based composites, HAp nanoparticles (particle size <200 nm (BET); density: 3.14 g/mL; Sigma-Aldrich) or BTO nanoparticles (particle size <100 nm (BET); density: 6.08 g/mL; Sigma-Aldrich) were dispersed in methyl ethyl ketone (MEK, Sigma-Aldrich) with P(VDF-TrFE) (80/20 mol % VDF/TrFE, 99%, Sigma–Aldrich). The solution was thoroughly mixed using an ultrasonic processor (MJ Research) for 2 h in an ice bath to ensure sufficient mixing and prevent warming. The solution was filtered using a 2 μm syringe filter to remove any impurities or agglomerated particles. The suspension was then cast onto a microscope slide glass and annealed on a hot plate. Annealing was performed in 3 steps, with a preannealing step to evaporate the solvent at 80 °C for 1 h, an annealing step to induce the β phase at 135 °C for another hour, and a postannealing step to control the cooling rate at 80 °C for 30 min. After returning to room temperature, the film was removed from the substrate and poled under an electric field of 7.5 kV for 1 h.

Material Characterization

The morphology of the composite samples was characterized by field emission scanning electron microscopy (Magellan400, FEI Company), and the crystallographic structures were examined by a thin-film X-ray diffractometer (Ultima IV, RIGAKU) and a Fourier transform infrared spectrometer (Nicolet iN10MX, Thermo Scientific). A surface profiler (Alpha-Step IQ, KLA-Tencor) was used to measure the thickness of the samples. A Berlincourt piezometer (PM300, Piezotest) was used to measure the piezoelectric coefficients with an oscillating load of 0.25 nN at a frequency of 110 Hz. For differential scanning calorimetry (Labsys Evo, Setaram) measurements, the temperature was increased from room temperature to 230 °C, with the heating rate maintained at 10 °C/min. The following Scherrer equation was used to calculate the ordered domain size.

graphic file with name am3c12575_m001.jpg 1

Here, τ, K, λ, β, and θ denote the average size of crystalline domains, shape factor, wavelength, width of peak at half the maximum intensity (fwhm), and Bragg angle, respectively. The mechanical tests of all scaffolds were performed using MCT-1150 tensile testers (A&D Company, Japan) at a test speed of 100 mm/min. The tests included the analysis of 10 specimens per sample with the same interval set.

Analysis of Cell Proliferation and Osteogenic Mineralization

Osteoblasts originating from the human bone tissue (adherent phenotype) were seeded onto the samples (1 × 104 cells/samples) and cultured for 6 h (cell attachment) and 5 days (cell proliferation) in DMEM. Quantitative analysis of cell proliferation on the samples was performed using a WST-1 assay (Premix WST-1 Cell Proliferation Assay System, Takara Bio Inc., Kusatsu, Japan). Osteoblasts were cultured for 7 and 14 days on samples (4 × 104 cells/sample) in osteogenic differentiation medium. Alizarin Red S (Sigma-Aldrich, USA) staining and ALP (Sigma-Aldrich, USA) staining were used to confirm the osteogenic differentiation (according to the degree of mineralization) of osteoblasts on sample surfaces. Quantification of Ca deposition on HAp/P(VDF-TrFE) was performed by subtracting the Ca deposition value of the HAp/P(VDF-TrFE) groups without cells from those cultured with osteoblasts. The stained cells were destained with cetylpyridinium chloride (Sigma-Aldrich, USA), and the extracted stains were measured using an absorbance reader (iMarkTM Microplate Absorbance Reader, Bio-Rad, Hercules, CA, USA) at 595 nm to quantify the osteogenic differentiation of hMSCs.

In Vivo Animal Study

The animal study was approved by the Ethics Committee of Chonnam National University. Six week-old male mice (C57Bl/6N) were assigned into four groups of four each: Defect, P(VDF-TrFE), BTO/P(VDF-TrFE), and HAp/P(VDF-TrFE). The mice were fully anesthetized with an intraperitoneal injection of 0.006 cc/10 g of Zoletil and 0.004 cc/10 g of Rumpun, and the heads were shaved and disinfected. The bones were exposed by incising the skin approximately 3.0 cm above the calvaria bone. Bone defects (diameter: 5 mm) were made on one side of the revealed calvarial bone using an electric drill. Prepared patches (diameter: 5 mm) were placed on the calvarial bone defect (Supplementary Figure 5). After the skin was sutured with sutures, the ambient temperature was raised, and mice were awakened from anesthesia. The mice were sacrificed 3 and 6 weeks after surgery to obtain tissues, including the defect region and the calvarial bone.

Histological Observation and Evaluation

Calvarial bone tomography was performed using Skyscan001172 (Skyscan, Konitch, Belgium) microcomputed tomography (micro-CT) at a resolution of 11.38 pixels and an exposure time of 316 ms with an energy source of 80 kV and a current of 124 μA. An average of 488 slices of calvarial bone were scanned. The micro-CT images were analyzed using MIMICS 14.0 3D imaging software (Materialise’s Interactive Medical Image Control System, Leuven, Belgium). The calvarial bone specimens were fixed in 10% formalin and decalcified in a 0.5 M EDTA (pH 7.4) solution at room temperature for 7 days. After the specimens were embedded in paraffin, they were cut into 5 μm-thick sections. Then, they were stained with hematoxylin and Masson’s trichrome stain. Images were obtained by Aperio Images Scope (Leica, CA, USA) software.

Characterizations Based on PFM and KPFM

For material characterization using atomic force microscopy (Cypher-ES, Asylum Research), the solution was spin-coated onto Au/Cr-coated Si wafers (iTASCO). PFM experiments were conducted using a conductive Pt/Ir-coated silicon tip (EFM, Nanosensors) with a spring constant of 2.8 nN/nm, while the tip loading force was set to 40 nN. A dual AC resonance tracking (DART) mode was used for the operation to enhance signal sensitivity.49 Processing to obtain a topological representation and calculation for surface roughness were performed using Igor Pro software (Wavemetrics). KPFM experiments were conducted with an atomic force microscope (Park NX10, Park Systems) using a conductive Au-coated tip (NCSTAu, Nanosensors) with a spring constant of 2.8 nN/nm.

Analysis of Cell Ca

Cells adhered onto the sample surfaces were fixed with modified Karnovsky’s fixative consisting of 2% paraformaldehyde and 2% glutaraldehyde (Sigma-Aldrich) in a 0.05 M sodium cacodylate buffer (Sigma-Aldrich) for 4 h. The samples were washed with 0.05 M sodium cacodylate buffer 3 times for 10 min and fixed with 1% osmium tetroxide (Sigma-Aldrich). The samples were then washed with distilled water and dehydrated with graded concentrations (50, 70, 80, 90, and 100% v/v) of ethanol. Then, the samples were treated with hexamethyldisilazane (Sigma-Aldrich) for 15 min. Finally, the samples were coated with gold prior to cell shape observation with FESEM (JEOL, JSM-5410LV, Japan), while the Ca content in cells was observed with EDS analysis.

Analysis of Cell Morphology and Vinculin

Adhered cells on samples were fixed with a 4% paraformaldehyde solution (Sigma-Aldrich, Milwaukee, WI, USA) for 20 min, permeabilized with 0.2% Triton X-100 (Sigma-Aldrich, WI, Milwaukee, USA) for 15 min, and stained with TRITC-conjugated phalloidin (Millipore, Billerica, MA, USA) and 4,6-diamidino-2-phenylindole (DAPI; Millipore, Billerica, MA, USA) for 1 h. FAs were also stained with a monoclonal antivinculin antibody (1:100; Millipore, Billerica, MA, USA) and a FITC-conjugated goat antimouse secondary antibody (1:500; Millipore, Billerica, MA, USA). Images of the stained cells were taken using a fluorescence microscope (Zeiss, Germany). For quantitative analysis of the body and nuclear shape of osteoblasts on the substrata, the images obtained by fluorescence microscopy were analyzed using a custom-written MATLAB script. To investigate the effects of FAs with variations in BTO and HAp density, we cultured osteoblasts on HAp/P(VDF-TrFE) for 12 h, followed by immunostaining. To further demonstrate the BTO and HAp density-induced changes in FAs, three-dimensional reconstruction images from the normalized fluorescence intensities of vinculin and F-actin were acquired by using custom-written MATLAB 2011b (MathWorks, Natick, MA).

Growth Factor Array

To ascertain whether osteoblasts could secrete growth factors and cytokines on the P(VDF-TrFE), BTO/P(VDF-TrFE), and HAp/P(VDF-TrFE) scaffolds, osteoblasts (3 × 104 cells/sample) were seeded on the scaffolds and cultured for 5 days in proliferation medium. The osteoblasts were then cultured for 3 days in the medium. Finally, the osteoblasts were again cultured for 1 day in DMEM using the RayBio G-Series Human Growth Factor Array 1 Kit (RayBiotech, Norcross, GA) according to the manufacturer’s protocol.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsami.3c12575.

  • Notes and results on the process of fabricating P(VDF-TrFE) scaffolds; peak identification of the XRD spectra for composite scaffolds; morphology and details of HAp-based scaffolds; notes and results of PFM characterization on composite scaffolds (PDF)

Author Contributions

# S.J., Y.G., and S.K. contributed equally to this work. S.J., S.K., and S.H. conceived this study. S.J., Y.G., and S.K. designed this study and conducted experiments. J.K. and S.H. supervised this study. S.J., Y.G., and S.K. wrote the manuscript. S.J., Y.G., S.K., J.K., and S.H. edited this manuscript. All authors contributed to the interpretation of the results.

This research was supported by the Research Promotion Team in Korea Advanced Institute of Science and Technology (KAIST), the KUSTAR-KAIST Institute, KAIST, Korea, and the KAIST-funded Global Singularity Research Program for 2022 and 2023 under award number 1711178180. This work was also supported by National Research Foundation (NRF) grants funded by the Korean government (NRF-2022M3A9E4017151, NRF-2022K1A4A7A04095892, RS-2023-00247245, and NRF-2021R1A4A3025206).

The authors declare no competing financial interest.

Supplementary Material

am3c12575_si_001.pdf (3.6MB, pdf)

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Supplementary Materials

am3c12575_si_001.pdf (3.6MB, pdf)

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