Abstract

   Regulation of the immune response is key to promoting bone regeneration
   by electroactive biomaterials. However, how electrical signals at the
   micro‐ and nanoscale regulate the immune response and subsequent
   angiogenesis during bone regeneration remains to be elucidated. Here,
   the distinctly different surface potential distributions on charged
   poly(vinylidene fluoridetrifluoroethylene) (P(VDF‐TrFE)) matrix
   surfaces are established by altering the dimensions of ferroelectric
   nanofillers from 0D BaTiO[3] nanoparticles (homogeneous surface
   potential distribution, HOPD) to 1D BaTiO[3] nanofibers (heterogeneous
   surface potential distribution, HEPD). Compared to HOPD, HEPD is
   significantly better at inducing the M2 polarization of macrophages and
   promoting neovascularization, which results in accelerated bone
   regeneration in vivo. The transcriptomic analysis reveals that
   macrophages modulated by HEPD display high expression levels of
   pro‐angiogenic genes, which is corroborated by tube‐formation assays,
   RT‐qPCR, and western blot analyses in vitro. Mechanistic explorations
   elucidate activation of the PI3K‐Akt signaling pathway, which in turn
   induces the polarization of macrophages toward a pro‐angiogenic
   phenotype. This study elucidates the cascade of biological processes by
   which heterogeneous electrical signals at the micro‐ and nanoscale
   modulate macrophage functions to promote vascularization and bone
   regeneration. Hence, this study provides new insights into how the
   micro‐ and nanoscale distribution characteristics of electrical signals
   facilitate bone regeneration.

   Keywords: angiogenesis, bone regeneration, heterogeneous surface
   potential distribution, PI3K‐Akt signaling pathway, pro‐angiogenic
   macrophage polarization
     __________________________________________________________________

   The polarized BaTiO[3] nanofibers (BTNF)/P(VDF‐TrFE) nanocomposite
   membrane provides heterogeneous surface potential distribution within
   the bone repair microenvironment, which induces macrophage
   differentiation toward the M2 phenotype and activates the PI3K‐Akt
   signaling pathway in macrophages to enhance their pro‐angiogenic
   functions, thereby enhancing angiogenic differentiation of endothelial
   cells. Ultimately, this cascade of biological processes accelerates
   bone defect repair.

   graphic file with name ADVS-12-2414278-g007.jpg

1. Introduction

   The physicochemical characteristics of biomaterials, including chemical
   composition, structure, stiffness, electrical properties, wettability,
   and pH, are pivotal for modulating cell behavior and maintaining tissue
   homeostasis.^[ [50]^1 , [51]^2 , [52]^3 , [53]^4 , [54]^5 , [55]^6 ^]
   Among the diverse array of biomaterials, electroactive biomaterials are
   particularly effective in enhancing bone regeneration via biomimetic
   recapitulation of the normal electrical microenvironment within the
   bone defect area.^[ [56]^2 , [57]^7 , [58]^8 ^] The electrical
   microenvironment where cells reside, is heterogeneous at the micro‐ and
   nanoscale.^[ [59]^9 , [60]^10 , [61]^11 , [62]^12 , [63]^13 ^] This
   heterogeneity plays a crucial role in cell growth, differentiation,
   migration, intercellular interactions, and various other cellular
   functions. Following this cue, we fabricated macroscopic electroactive
   nanocomposite membranes and further developed the electrical properties
   of these materials by synthesizing nanocomposite membranes with
   heterogeneous surface potential distribution (HEPD) at the micro‐ and
   nanoscale.^[ [64]^10 ^] This innovation mimics the heterogeneous
   electrical microenvironment at the micro‐ and nanoscale within the bone
   regeneration microenvironment. Compared to homogeneous electrical
   signals, our nanocomposite membranes with HEPD were significantly
   better at promoting the osteogenic differentiation and mineralization
   of mesenchymal stem cells by enhancing cellular mechanosensitivity,
   thereby improving the efficacy of bone repair with electroactive
   materials.^[ [65]^10 ^] However, how heterogeneous electrical signals
   at the micro‐ and nanoscale modulate the cascade of biological
   processes including immune response and subsequent angiogenesis during
   bone regeneration remains to be elucidated.

   Studies have demonstrated that immune cells can produce and release a
   variety of pro‐angiogenic factors, to create a microenvironment
   favorable to angiogenesis thereby accelerating bone regeneration.^[
   [66]^14 ^] As the major immune cells that form the first line of
   defense in the innate immune system, macrophages are found in almost
   all tissues and play a crucial role in tissue repair.^[ [67]^15 ,
   [68]^16 , [69]^17 ^] In particular, macrophages have garnered extensive
   attention due to their high plasticity in displaying various
   phenotypes, with the M1 phenotype exhibiting pro‐inflammatory
   functions, while the M2 phenotype contributes to the regression of
   inflammation, promotion of tissue repair, and maintenance of tissue
   homeostasis.^[ [70]^16 , [71]^18 , [72]^19 , [73]^20 , [74]^21 ,
   [75]^22 , [76]^23 ^] Previous studies have confirmed that electroactive
   biomaterials can effectively regulate the polarization of macrophages
   to the M2 phenotype, thereby promoting the regeneration of blood
   vessels that ultimately facilitate the repair of bone defects.^[
   [77]^24 , [78]^25 , [79]^26 ^] However, the majority of published
   research studies on electroactive biomaterials for bone repair have
   focused on the regulation of immune response and vascular regeneration
   by macroscopic homogeneous electrical signals. There is still a lack of
   investigations on the cascade of biological processes involved in
   regulating the immune response, promoting vascular regeneration, and
   ultimately facilitating bone regeneration by heterogeneous
   distributions of electrical signals at the micro‐ and nanoscale. This
   poses a major obstacle in precisely further enhancing the biological
   performance of electroactive biomaterials.

   In this study, we employed ferroelectric nanocomposite membranes
   specifically designed to provide HEPD, which incorporated optimized
   BaTiO[3] nanofibers (BTNF) within poly(vinylidene
   fluoridetrifluoroethylene) (P(VDF‐TrFE)) matrix.^[ [80]^10 ^] This
   facilitated our investigations of the regulatory roles of micro‐ and
   nanoscale heterogeneous electrical signals in immune responses and
   vascular regeneration during bone regeneration. By combining vascular
   microfil‐perfusion and immunofluorescence staining, we demonstrated
   that HEPD can promote angiogenesis and subsequent bone regeneration in
   vivo. Meanwhile, it was also observed that HEPD could better promote
   the M2 polarization of macrophages both in vitro and in vivo, as
   compared to homogeneous surface potential distribution (HOPD). Gene
   ontology (GO) analysis of differentially expressed genes (DEGs)
   revealed that genes highly expressed in macrophages modulated by HEPD
   were significantly enriched in the function associated with
   “angiogenesis”. Consistently, the tube‐formation assay, RT‐qPCR, and
   western blot analyses showed that macrophages modulated by HEPD could
   enhance the angiogenic function of human umbilical vein endothelial
   cells (HUVECs) in vitro. Mechanistically, HEPD promoted the
   pro‐angiogenic functions of macrophages by activating the PI3K‐Akt
   signaling pathway. In summary, this study thus established the
   relationship between micro‐ and nanoscale heterogeneous electrical
   signals, pro‐angiogenic macrophages, angiogenesis, and bone
   regeneration, thereby facilitating a better understanding of the
   underlying mechanisms by which micro‐ and nanoscale heterogeneous
   electrical signal distribution regulate bone regeneration (Scheme [81]1
   ).

Scheme 1.

   Scheme 1
   [82]Open in a new tab

   Schematic illustration of HEPD enhancing osteogenesis by promoting M2
   and pro‐angiogenic polarization of macrophages that facilitate
   angiogenesis. When the polarized BTNF/P(VDF‐TrFE) nanocomposite
   membrane covered the bone defect, the HEPD provided by the membrane
   downregulates the expression of the M1 marker CD86 and inflammatory
   cytokine TNFα in macrophages, and further induces their polarization
   toward the M2 phenotype, thus establishing an anti‐inflammatory
   microenvironment. Meanwhile, the HEPD also activates the PI3K‐Akt
   signaling pathway in macrophages to enhance their pro‐angiogenic
   functions, thereby enhancing subsequent angiogenesis of endothelial
   cells. Ultimately, this cascade of biological processes accelerates
   bone defect repair.

2. Results and Discussion

2.1. The Establishment of Heterogeneous Surface Potential Distribution (HEPD)
on Ferroelectric Nanocomposite Membranes

   Our previous study has demonstrated that BTNF obtained through
   electrospinning, after sintering at 750 °C for 6 h and being subjected
   to ultrasonic treatment for 5 min, could achieve an optimal aspect
   ratio. This results in a heterogeneous surface electric potential
   distribution on the charged nanocomposite membranes after incorporating
   optimized BTNF within the P(VDF‐TrFE) matrix, which facilitated cell
   recognition, adhesion, and osteogenic differentiation.^[ [83]^10 ^]
   Based on this, we fabricated flexible electroactive nanocomposite
   membranes consisting of dopamine‐modified BaTiO[3] nanoparticles (BTNP)
   or BTNF as fillers and P(VDF‐TrFE) as the matrix via the solution
   casting method (Figure [84]1A,B). The two membranes were subjected to
   corona discharge at room temperature to obtain polarized electroactive
   nanocomposite membranes that provided HOPD and HEPD at the micro‐ and
   nanoscale, respectively, thus facilitating the subsequent investigation
   of differences in biological effects between them. The Scanning
   Electron Microscopy (SEM) images revealed that BTNF was anisotropically
   distributed within the polymer matrix, while BTNP was isotropically
   homogeneous (Figure [85]1C). The surface electric potential
   distribution was further characterized by Scanning Probe Microscopy
   (SPM) imaging (Figure [86]1D). It was observed that the surface
   electric potential of the BTNF nanocomposite membrane exhibited a
   line‐like anisotropic distribution, while the BTNP nanocomposite
   membrane displayed a homogeneous high‐potential pitting distribution.
   This difference might be due to the increase in local surface electric
   potential caused by the different shapes of BaTiO[3] fillers after
   polarization.^[ [87]^10 ^] However, there was no significant difference
   in the roughness between the two membranes (Figure [88]1E).
   Additionally, the electrical properties of the membranes were
   characterized, revealing that the piezoelectric constant d[33] , the
   polar 𝛽‐phase content and the polarization‐electric field (P‐E) loop
   were similar between the two membranes (Figure [89]S1A–C, Supporting
   Information). Based on these findings, we confirmed that BTNF and BTNP
   nanocomposite membranes differed in the surface electric potential
   distribution at the micro‐ and nanoscale, with no significant
   differences in other general properties being observed. This conclusion
   was consistent with our previous work.^[ [90]^10 ^] Hence, there is a
   rational basis for utilizing BTNF nanocomposite membranes to
   investigate the effect of micro‐ and nanoscale heterogeneous electrical
   signals in regulating the bone repair process.

Figure 1.

   Figure 1
   [91]Open in a new tab

   HEPD enhances angiogenesis and subsequent bone regeneration in vivo. A)
   Schematic diagram of the fabrication process of BTNF nanocomposite
   membranes. B) Image of a BTNF nanocomposite membrane. C) Representative
   SEM images of BTNF and BTNP nanocomposite membranes. The white arrows
   denote the BaTiO[3] nanofibers (BTO NFs) and BaTiO[3] nanoparticles
   (BTO NPs). Scale bars: 5 µm. D) Surface potentials of BTNF and BTNP
   nanocomposite membranes. Scale bars: 1 µm. E) Roughness of BTNF and
   BTNP nanocomposite membranes, n = 3. F) Schematic illustration of
   angiogenesis and bone regeneration assessment. G) Representative
   micro‐CT images of vasculature within the microfil‐perfused craniums
   showed that compared with the HOPD group, neovessel formation was
   significantly enhanced in the HEPD group. The dashed lines indicate
   bone defect areas. Scale bars: 1 mm. H) Quantitative analysis of
   neovessel volumes in G showed increased vessel volume in the HEPD
   group, n = 6. I) Representative micro‐CT images of mice with
   2mm‐diameter calvarial full‐thickness defects at 12 weeks post‐surgery.
   The red dotted lines denote the boundary between the nascent bone and
   the host bone. Scale bars: 1 mm. J) Quantitative analysis of bone
   volume fraction (BV/TV), n = 6. K) Representative immunofluorescence
   staining images of regenerated tissues at 2 weeks post‐surgery. The
   dashed lines denote the indicated tissues. Scale bars: 100 µm. L)
   Analysis of mean fluorescence intensities representing expression of
   CD34 and osteocalcin (OCN) in K, n = 6. Statistical significance was
   assessed using the unpaired Student's t‐test. ns, not significant; *p <
   0.05; **p < 0.01.

2.2. HEPD Enhances Angiogenesis and Subsequent Bone Regeneration In Vivo

   The vascular network, distributed throughout bone tissue, plays a
   crucial role in bone development and regeneration by delivering oxygen,
   essential nutrients, cells, and regenerative factors.^[ [92]^27 ,
   [93]^28 ^] Following this cue, we first investigated whether HEPD could
   promote angiogenesis during bone regeneration (Figure [94]1F). We
   constructed a mouse calvarial defect model which was covered with
   either polarized BTNF (HEPD group) or BTNP (HOPD group) nanocomposite
   membranes. At 2 weeks post‐surgery, micro‐CT angiography showed that
   BTNF nanocomposite membranes significantly promoted the formation of
   newly regenerated capillaries within the calvarial bone defect area
   (Figure [95]1G). Quantitative analysis showed a significantly increased
   volume of the newly formed capillaries in the HEPD group, with the
   volume being ≈1.5 times that of the HOPD group (Figure [96]1H). At 12
   weeks post‐surgery, micro‐CT analysis showed that the HEPD group
   exhibited a significantly greater degree of regenerated bone tissue in
   the bone defect area compared to the HOPD group (Figure [97]1I). As
   expected, the quantitative analysis results showed a significant
   increase in the BV/TV of the HEPD group, which was ≈25% in the HOPD
   group but >30% in the HEPD group (Figure [98]1J). Furthermore, the
   effects of HEPD on angiogenesis and osteogenesis were assessed by
   immunofluorescence staining for CD34 (a marker of vascular endothelial
   progenitor cells) and OCN (an osteogenesis‐related protein) within the
   regenerated tissues at 2 weeks post‐surgery (Figure [99]1K).
   Quantitative analysis revealed that the intensities of CD34 and OCN
   were significantly higher in the HEPD group compared to the HOPD group
   (Figure [100]1L). Together, these results suggested that HEPD further
   promoted angiogenesis and subsequent bone regeneration in vivo.

2.3. HEPD Promotes the M2 Polarization of Macrophages and the Establishment
of an Anti‐Inflammatory Microenvironment In Vivo

   Next, we endeavored to uncover the mechanisms by which HEPD can further
   promote angiogenesis and osteogenesis. The immune response,
   particularly the polarization of macrophages, has been established as a
   crucial regulatory factor in vascularization during bone
   regeneration.^[ [101]^24 , [102]^25 , [103]^26 ^] Therefore, we
   investigated whether HEPD could modulate the polarization of
   macrophages, to provide a beneficial immune microenvironment conducive
   for tissue regeneration in vivo. Polarized BTNF or BTNP nanocomposite
   membranes were used to cover the calvarial defects in mice (Figure
   [104]2A). On the 3rd and 7th day post‐surgery, we performed
   immunofluorescence staining of the recruited cells on the implanted
   nanocomposite membranes. We utilized CD68 and CD206 to identify M2
   macrophages, while CD68 and CD86 served to label M1 macrophages.^[
   [105]^23 , [106]^29 ^] As shown in Figure [107]2B,C, the HEPD group
   exhibited significantly higher expression of the M2 macrophage surface
   marker CD206 on the 3rd day post‐surgery, when compared to the HOPD
   group. In contrast, the expression of the M1 macrophage surface marker
   CD86 was significantly higher on the BTNP nanocomposite membranes. On
   the 7th day post‐surgery, the expression of CD206 increased in
   macrophages recruited by the BTNP nanocomposite membranes, while
   remaining lower than that on the BTNF nanocomposite membranes, whereas
   the expression of CD86 in the HOPD group was still higher. These
   results thus indicated that the BTNF nanocomposite membranes with HEPD
   promoted M2 polarization of macrophages more effectively than the BTNP
   nanocomposite membrane during the early stage (from day 3) of bone
   defect repair.

Figure 2.

   Figure 2
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   HEPD promotes the M2 polarization of macrophages and the establishment
   of an anti‐inflammatory microenvironment in vivo. A) Schematic
   illustration of the sample source for immunofluorescence staining. B,C)
   Representative immunofluorescence staining images and analysis of
   fluorescence intensity showed that the expression of the M2 macrophage
   surface marker CD206 was higher in the HEPD group compared to the HOPD
   group, while expression of the M1 macrophage surface marker CD86 was
   higher in the HOPD group on the 3rd and 7th day post‐surgery. Scale
   bars: 50 µm, n = 50. D,E) Representative immunofluorescence staining
   images and the analysis of mean fluorescence intensity showed a
   significant increase in IL‐10 expression within the defect area of the
   HEPD group compared with the HOPD group on the 3rd and 7th day
   post‐surgery. Scale bars: 100 µm, n = 3. Statistical significance was
   assessed using the unpaired Student's t‐test. *p < 0.05; ****p <
   0.0001.

   We further conducted immunofluorescence staining for detecting the
   expression of IL‐10, a classic anti‐inflammatory cytokine,^[ [109]^30 ,
   [110]^31 ^] within the regenerated tissue of the bone defect area on
   the 3rd and 7th day post‐surgery. As expected, the results showed a
   significant increase in IL‐10 expression within the defect area of the
   HEPD group compared to the HOPD group (Figure [111]2D,E). Hence, these
   results indicated that HEPD could significantly better induce M2
   polarization of macrophages in vivo, thereby providing an
   anti‐inflammatory microenvironment potentially beneficial for tissue
   repair.

2.4. HEPD Enhances the M2 Polarization of Macrophages In Vitro

   Subsequently, we further conducted in vitro experiments to delve into
   the regulatory role of HEPD on macrophage M2 polarization, thereby
   eliminating interference from the complex in vivo microenvironment and
   providing a more intuitive investigation of the regulatory effects of
   electrical signals on macrophages. Mouse bone marrow‐derived
   macrophages (BMDMs) were cultured on BTNF or BTNP nanocomposite
   membranes. Flow cytometry analysis revealed a significant upregulation
   of CD206 expression in the HEPD group, about threefold that in the HOPD
   group. However, the expression of CD86 in the HEPD group was
   significantly downregulated, about half of that in the HOPD group
   (Figure [112]3A,B). Consistently, immunofluorescence staining revealed
   a significant difference in the expression of both CD206 and CD86
   between the two groups (Figure [113]3C,D). Compared with the HOPD
   group, the fluorescence intensity of CD206 was higher in the HEPD
   group, while the fluorescence intensity of CD86 was lower.
   Additionally, RT‐qPCR analysis showed a significant upregulation of the
   M2 marker genes, Cd206 and Arg1, in BMDMs of the HEPD group, which was
   accompanied by downregulation of the M1 marker gene, Cd86, and the
   inflammatory cytokine gene, Tnfα (Figure [114]3E).^[ [115]^32 ^]
   Furthermore, western blot analysis revealed significantly higher levels
   of CD206 protein expression and significantly lower levels of CD86
   protein expression in the HEPD group compared to the HOPD group
   (Figure [116]3F,G). Hence, our results demonstrated that HEPD can
   better promote the M2 polarization of macrophages in vitro compared to
   HOPD. Together with the in vivo results, they demonstrated the
   regulatory role of HEPD on the immune microenvironment.

Figure 3.

   Figure 3
   [117]Open in a new tab

   HEPD enhances the M2 polarization of macrophages in vitro. A,B)
   Flow cytometry showed that the expression of CD206 was significantly
   upregulated in the HEPD group compared with the HOPD group, whereas
   CD86 expression in the HEPD group was significantly downregulated, n =
   6. C) Representative immunofluorescence staining images showed that the
   expression of CD206 was significantly higher while the expression of
   CD86 was significantly lower in the HEPD versus HOPD group. Scale bars:
   50 µm. D) The immunofluorescence intensity of CD206 in the HEPD group
   was significantly higher while that of CD86 was significantly lower in
   the HEPD versus HOPD group, n = 50. E) RT‐qPCR showed that the gene
   expression levels of M2 markers Cd206 and Arg1 were upregulated in the
   HEPD group, whereas the expression levels of M1 marker Cd86 and the
   inflammatory cytokine Tnfα were downregulated, n = 3. F,G) Western blot
   analysis showed significantly higher levels of CD206 protein expression
   and significantly lower levels of CD86 protein expression in the HEPD
   group compared with the HOPD group, n = 3. Statistical significance was
   assessed using the unpaired Student's t‐test. *p < 0.05; **p < 0.01;
   ***p < 0.001; ****p < 0.0001.

2.5. HEPD Promotes Angiogenesis by Regulating the Pro‐Angiogenic Functions of
Macrophages

   Having established that HEPD can significantly better promote vascular
   regeneration and M2 polarization of macrophages compared to HOPD, we
   hypothesized that the modulation of macrophages by HEPD may positively
   enhance angiogenesis. For this purpose, we collected BMDMs cultured on
   BTNF and BTNP nanocomposite membranes for bulk RNA‐seq analysis. As
   expected, DEGs analysis (Figure [118]4A), showed that “Ramp1”^[
   [119]^33 , [120]^34 , [121]^35 ^] and “Angpt1”^[ [122]^36 , [123]^37 ^]
   genes, known for their key roles in angiogenesis, were highly expressed
   in BMDMs of the HEPD group. The gene enrichment analysis revealed that
   among the top 30 biological processes highly expressed by BMDMs in the
   HEPD group, there were those related to osteogenesis, such as
   “ossification”, “skeletal system development”, and “bone
   mineralization”, as well as angiogenesis‐related biological processes,
   including “angiogenesis”, “artery morphogenesis”, and “artery
   development” (Figure [124]4B). Consistently, osteogenesis and
   angiogenesis‐related gene cluster analysis showed that these genes were
   significantly enriched in the HEPD group (Figure [125]4C). These
   results thus suggested that the BMDMs in the HEPD group display more
   obvious transcriptomic characteristics of promoting angiogenesis and
   bone regeneration than those in the HOPD group. Therefore, we
   hypothesized that the HEPD may play a key role in promoting
   angiogenesis by modulating the pro‐angiogenic functions of macrophages,
   ultimately promoting bone regeneration.

Figure 4.

   Figure 4
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   HEPD promotes angiogenesis by regulating the pro‐angiogenic functions
   of macrophages. A) The volcano plot of DEGs in the HEPD group compared
   with the HOPD group. B) The enriched GO terms of upregulated genes in
   the HEPD group compared with the HOPD group. The green and red
   highlights denote osteogenesis and angiogenesis‐related biological
   processes, respectively. C) Gene cluster analysis of osteogenesis and
   angiogenesis‐related genes in the HEPD and HOPD groups. D) Schematic
   illustration of angiogenic differentiation using conditioned medium
   from BMDMs cultured on the BTNF or BTNP nanocomposite membranes. E)
   Representative images of 2D tube‐formation assay of HUVECs in different
   conditioned mediums of BMDMs cultured on BTNF and BTNP nanocomposite
   membranes showed more angiogenic nodes, meshes, and segments in the
   HEPD versus HOPD group. Arrows denote the nodes, meshes, and segments.
   Scale bars: 200 µm. F) Quantitative analysis of tube‐formation in E
   showed an increased number of nodes, number of meshes, and the number
   and length of segments in the HEPD group, n = 3. G) RT‐qPCR showed that
   the gene expression levels of angiogenesis‐related genes CD31 and VEGF
   were upregulated in the HEPD group compared with the HOPD group, n = 3.
   H,I) Western blot analysis showed significantly higher levels of
   angiogenesis‐related proteins CD31 and CD34 in the HEPD group compared
   with the HOPD group, n = 3. J) Representative micro‐CT images of
   vasculature within the microfil‐perfused craniums showed that
   macrophage depletion significantly reduced the capacity of HEPD to
   promote the formation of newly regenerated capillaries. The dashed
   lines indicate bone defect areas. Scale bars: 1 mm. K) Quantitative
   analysis of neovessel volumes in J showed decreased vessel volume in
   the macrophage depletion group, n = 6. Statistical significance was
   assessed using the unpaired Student's t‐test. **p < 0.01; ****p <
   0.0001.

   To validate the pro‐angiogenic functions of macrophages activated by
   HEPD, we performed a 2D tube‐formation assay (Figure [127]4D). BMDMs
   were cultured on either BTNF or BTNP nanocomposite membranes to obtain
   conditioned media. Subsequently, HUVECs were cultured in the different
   conditioned media for 4 h. As shown in Figure [128]4E,F, the number of
   nodes and meshes, as well as the number and length of segments, were
   significantly increased in the HEPD group compared to the HOPD group.
   We further investigated the expression of classic angiogenesis‐related
   genes in BMDMs in response to different electrical signals at both the
   mRNA and protein levels. The RT‐qPCR results showed that the expression
   levels of CD31 and VEGF in BMDMs were significantly higher in the HEPD
   versus HOPD group (Figure [129]4G). Consistent with these results,
   western blot analysis also revealed significantly higher expression of
   CD31 and CD34 in BMDMs from the HEPD group compared to those from the
   HOPD group (Figure [130]4H,I). To further verify the crucial role of
   macrophages in regulating the process by which HEPD promotes
   angiogenesis, we constructed a mouse calvarial defect model
   and depleted macrophages in vivo using clodronate liposome.^[ [131]^38
   , [132]^39 , [133]^40 ^] At 2 weeks post‐surgery, micro‐CT angiography
   showed that macrophage depletion significantly reduced the capacity of
   HEPD to promote the formation of newly regenerated capillaries
   (Figure [134]4J). The results of the quantitative analysis were
   consistent, and the volume of newly formed capillaries within the
   macrophage depletion group was significantly reduced by ≈28% when
   compared with the HEPD group (Figure [135]4K). Hence, these results
   indicated that HEPD could polarize macrophages toward a pro‐angiogenic
   phenotype, thereby enhancing their pro‐angiogenic effects on HUVECs,
   which in turn promoted subsequent vascular regeneration and bone
   repair.

2.6. HEPD Induces the Pro‐Angiogenic Functions of Macrophages by Activating
the PI3K‐Akt Signaling Pathway

   To investigate the mechanisms by which HEPD modulates the
   pro‐angiogenic functions of macrophages, we further analyzed the bulk
   RNA‐seq results. The pathway enrichment analysis revealed that the
   genes highly expressed in BMDMs of the HEPD group were associated with
   the PI3K‐Akt signaling pathway, whose activation was reported to be
   important for angiogenesis (Figure [136]5A).^[ [137]^41 , [138]^42 ^]
   This finding indicated that the HEPD on BTNF nanocomposite membranes
   modulated the pro‐angiogenic functions of macrophages, which might be
   related to the activation of the PI3K‐Akt signaling pathway.

Figure 5.

   Figure 5
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   HEPD induces the pro‐angiogenic functions of macrophages by activating
   the PI3K‐Akt signaling pathway. A) The pathway enrichment analysis of
   upregulated genes in the HEPD group compared with the HOPD group. B)
   Schematic illustration of angiogenic differentiation using conditioned
   medium from BMDMs cultured on BTNF nanocomposite membranes after adding
   LY294002, the inhibitor of the PI3K‐Akt signaling pathway. C)
   Representative images of 2D tube formation assay of HUVECs in different
   conditioned mediums. Arrows denote the nodes, meshes, and segments.
   Scale bars: 200 µm. D) Quantitative analysis of tube formation in C
   showed decreases in the number of nodes, the number of meshes, and the
   number and length of segments after adding LY294002, n = 3. E) RT‐qPCR
   showed that the gene expression levels of angiogenesis‐related genes
   CD31 and VEGF were downregulated after adding LY294002, n = 3. F,G)
   Western blot analysis showed significantly lower levels of
   angiogenesis‐related proteins CD31 and CD34 after adding LY294002, n =
   3. H) Schematic illustration of the mechanisms by which heterogeneous
   electrical signals induced the pro‐angiogenic polarization of
   macrophages by activating the PI3K‐Akt signaling pathway, thereby
   promoting angiogenesis. Statistical significance was assessed using the
   unpaired Student's t‐test. *p < 0.05; **p < 0.01; ***p < 0.001; ****p <
   0.0001.

   To verify whether HEPD regulates the pro‐angiogenic functions of
   macrophages through the PI3K‐Akt signaling pathway, we utilized
   LY294002, an inhibitor of the PI3K‐Akt signaling pathway.^[ [140]^42 ,
   [141]^43 , [142]^44 , [143]^45 ^] Specifically, when BMDMs were
   cultured on BTNF nanocomposite membranes, we added LY294002 at the same
   time. After stimulating with LY294002 for 24 h, the medium was
   discarded and replaced with fresh complete medium, and the cells were
   further cultured for 48 h. The conditioned medium was then collected
   for subsequent experiments. Following this, HUVECs were cultured in the
   collected conditioned medium for 4 h (Figure [144]5B). The results of
   the 2D tube‐formation assay showed that the addition of LY294002
   significantly inhibited the tube‐formation capacity of HUVECs
   (Figure [145]5C,D). Additionally, we also observed the same results
   from RT‐qPCR and western blot analyses whereby the addition of LY294002
   significantly decreased the gene expression levels of CD31 and VEGF and
   the protein expression levels of CD31 and CD34 (Figure [146]5E–G). In
   summary, these results indicated that HEPD induced the pro‐angiogenic
   polarization of macrophages by activating the PI3K‐Akt signaling
   pathway, which promoted angiogenesis (Figure [147]5H), ultimately
   enhancing bone defect repair.

3. Conclusion

   In this study, we elucidated how electrical signals at the micro‐ and
   nanoscale regulate the immune response to promote bone regeneration. We
   found that, when compared to the HOPD, HEPD could significantly better
   promote the polarization of macrophages toward the M2 phenotype. This
   transition endowed the immune microenvironment with anti‐inflammatory
   properties during the initial stages of bone defect repair, which was
   beneficial to subsequent tissue regeneration. Specifically, the HEPD
   activated the PI3K‐Akt signaling pathway in macrophages to enhance
   their pro‐angiogenic functions, thereby enabling them to promote
   subsequent angiogenesis of endothelial cells. Ultimately, this cascade
   of biological processes accelerated the bone defect repair process.
   Hence, this study provides novel insights into the in‐depth
   understanding of the essence of micro‐ and nanoscale electrical
   signal‐mediated regulation of tissue regeneration. Additionally, cues
   for enhancing the therapeutic efficacy of electroactive biomaterials
   were also provided. Moreover, the focus of this study is on
   macrophages. Following this cue, future research should prioritize
   studies on how heterogeneous electrical signals at the micro‐ and
   nanoscale regulate the interaction between macrophages with other
   immune cells, as well as with non‐immune cells to promote tissue
   regeneration. This will contribute to a more comprehensive and accurate
   understanding of the mechanisms of action of these heterogeneous
   electrical signals, thereby providing a theoretical basis for precise
   improvement of the properties of electroactive biomaterials and
   ensuring the safety of their clinical applications in the future.

4. Experimental Section

Fabrication of BaTiO[3] Nanofibers (BTNF) and BaTiO[3] Nanoparticles (BTNP)
Nanocomposite Membranes

   The BTNF was fabricated as described in previous study.^[ [148]^10 ^]
   Barium acetate, tetrabutyl titanate, and acetylacetone (TONG GUANG,
   China) were dissolved in acetic acid (Macklin, China) at a 1:1:2 molar
   ratio and stirred to yield a homogeneous barium titanate precursor
   solution. The viscosity of the solution was adjusted by adding
   polyvinylpyrrolidone (PVP, M = 1 300 000) (Aladdin, China) to the
   precursor solution. The mixed solution was then subjected to
   electrospinning under an applied electric field of 1.5 kV cm^−1. BTNF
   was obtained by calcining electrospun fibers consisting of PVP and
   barium titanate precursor at 750 °C for 6 h. The BTNF was dispersed by
   ultrasonication for 5 min. BTNP with a diameter of 100 nm was procured
   from Sigma–Aldrich Inc. (St. Louis, USA). Subsequently, BTNF and BTNP
   were surface‐modified based on the protocol of the previous studies.^[
   [149]^2 , [150]^10 ^] Briefly, BTNF and BTNP were mixed with a 0.01 mol
   L^−1 dopamine hydrochloride (Alfa Aesar, USA) solution and stirred for
   12 h at 60 °C. Finally, the dopamine‐modified BTNF or BTNP was
   incorporated into P(VDF‐TrFE) (70/30 mol% VDF/TrFE) (Arkema, France)
   co‐polymer powders dissolved in N,N‐dimethylformamide (DMF) (TONG
   GUANG, China) at a polymer matrix concentration of 5 vol%. The mixture
   was stirred overnight to form a stable suspension and then cast as a
   membrane on a Quartz glass substrate. The membranes were dried at 55 °C
   for 10 h to volatilize the DMF. Importantly, the membranes were
   polarized by subjecting them to a corona discharge at room temperature
   under a 15 kV DC field for 30 min.

Characterization of the BTNF and BTNP Nanocomposite Membranes

   The surface morphology of the nanocomposite membranes after
   polarization was characterized using SEM JSM‐7001F (JEOL, Japan). The
   surface potential and roughness were examined with SPM using a Bruker
   Dimension Icon instrument (Bruker, USA). The piezoelectric coefficient
   d[33] was determined by a Quasistatic d[33] instrument, ZJ‐ 6A (JKZC,
   China).^[ [151]^46 ^] Phase structures were analyzed via X‐ray
   diffraction spectroscopy using a Rigaku D/max 2500VB2t/PC (Rigaku,
   Japan). The P‐E loop was assessed using a Ferroelectric tester,
   Precision 10 kV HVI‐SC (Radiant Technologies, USA).

Animals and Surgical Procedures

   The cranial bone defect model was established using 6‐week‐old male
   C57BL/6 mice in this study. All animal experiments were conducted under
   the approval of the Animal Care and Use Committee of Peking University
   (IACUC No. LA2022682). The mice were anesthetized with 50 mg kg^−1 of
   pentobarbital sodium via intraperitoneal injections, followed by the
   exposure of the dorsal cranium. Two full‐thickness bone defects, each
   2 mm in diameter, were created at the center of each mouse's parietal
   bone using a saline‐cooled trephine drill. The bone defects were
   covered with either polarized BTNF nanocomposite membranes or polarized
   BTNP nanocomposite membranes. The whole calvarias were collected for
   evaluation at multiple time points: 3 days, 7 days, 2 weeks, and 12
   weeks post‐surgery.

   To deplete macrophages, 6‐week‐old male C57BL/6 mice weighing ≈20 g
   were administered clodronate liposome (Yeasen Biotechnology, China) at
   a dosage of 200 µL per mouse, via intravenous injection, at 48 h before
   surgery. Injections were administered every 3 days until the 7th day
   post‐surgery. The control group received the same dose of control
   liposomes (Yeasen Biotechnology, China).^[ [152]^38 , [153]^39 ,
   [154]^40 ^]

Cell Culture

   BMDMs were cultured in dulbecco's modified eagle medium (Procell,
   China) supplemented with 10% (v v^−1) fetal bovine serum (Procell) and
   1% (v v^−1) penicillin/streptomycin solution (Procell). HUVECs
   (Oricell, China) were cultured in endothelial cell medium (Oricell).
   All cells were cultured within a humidified incubator containing 5%
   CO[2] at 37 °C. LY294002 (MedChemexpress, USA) at 10 μм concentration
   was used as an inhibitor of the Pl3K‐Akt signaling pathway.^[ [155]^42
   , [156]^43 , [157]^44 ^]

Flow Cytometry

   BMDMs treated with different nanocomposite membranes were collected,
   fixed, and permeabilized using 100% methanol. Subsequently, the cells
   were incubated with primary antibodies at 4 °C for 30 min. The primary
   antibodies used were as follows: CD86 Polyclonal antibody (1:500;
   Proteintech, USA) and APC‐CD206 (0.5 µg per million cells in 100 µL
   volume; Biolegend, USA). Following incubation with the primary
   antibody, the cells were washed with phosphate‐buffered saline (PBS)
   (Solarbio, China) and incubated with the secondary antibody Goat
   Anti‐Rabbit IgG H&L (Alexa Fluor 488) (Abcam, UK) at 4 °C for 30 min.
   Analysis was conducted using LSRFortessa flow cytometers (BD
   Biosciences, USA), and data were processed using FlowJo software (Tree
   Star, USA).

RT‐qPCR

   RNA from BMDMs or HUVECs was extracted using Trizol reagent
   (Invitrogen, USA). The concentration of RNA was measured at 260 nm
   using a NanoDrop ND‐2000 spectrophotometer (Thermo Fisher Scientific,
   USA). Reverse transcription was performed using a PCR thermal cycler
   (Takara, Japan). PCR was performed on a 96‐well optical reaction plate
   with a total volume of 20 µL, comprising 1 µL of template cDNA, 10 µL
   of FastStart Universal SYBR Green Master Mix, 8 µL of RNase‐free water,
   and 1 µL of primer. PCR amplification was conducted with the following
   cycling parameters: 15 min at 95 °C, followed by 45 cycles of 15 s at
   95 °C and 1 h at 60 °C. Differences in gene expression levels between
   the groups were statistically analyzed after processing the raw data.
   All reactions were conducted in triplicates. Glyceraldehyde‐3‐phosphate
   dehydrogenase (GAPDH) was used as the internal control. The primer
   sequences are listed in Table [158]S1 (Supporting Information).

Western Blot

   Cells were lysed using radioimmunoprecipitation assay buffer (Beyotime,
   China) containing a protease and phosphatase inhibitor cocktail (Thermo
   Fisher Scientific), followed by centrifugation at 12 000 rpm for 20 min
   at 4 °C to collect the supernatants. The protein concentration was
   determined using a bicinchoninic acid protein assay kit (Beyotime).
   Samples were mixed with 6 × sodium dodecyl sulfate loading buffer
   (Beyotime) at a 1:5 ratio and heated at 100 °C for 5 min to denature
   the proteins. Twenty micrograms of protein were separated by sodium
   dodecyl sulfate polyacrylamide gel electrophoresis and transferred onto
   a polyvinylidene fluoride membrane. The membranes were blocked with 5%
   (w v^−1) skimmed milk (diluted with tris buffered saline with tween 20)
   for 1 h at room temperature, followed by overnight incubation with the
   primary antibody at 4 °C, and subsequent incubation with an horseradish
   peroxidase (HRP)‐conjugated secondary antibody for 1 h at room
   temperature. Autoradiograms were obtained using Enhanced
   Chemiluminescence Western Blotting Substrate (Abclonal, China). GAPDH
   served as the internal control. The primary antibodies used were as
   follows: Anti‐GAPDH antibody (1:2000; Abcam), Anti‐Mannose Receptor
   (CD206) antibody (1:1000; Abcam), CD86 Polyclonal antibody (1:1000;
   Proteintech), Anti‐CD31 antibody (1:1000; Abcam), and CD34 Polyclonal
   antibody (1:1000; Proteintech). The HRP‐conjugated secondary antibodies
   used were as follows: HRP‐labeled Goat Anti‐Rabbit IgG(H + L) (1:1000;
   Beyotime) and HRP‐labeled Goat Anti‐Mouse IgG(H + L) (1:1000;
   Beyotime).

Immunohistochemical Staining

   For nanocomposite membranes and BMDMs, the samples were rinsed with PBS
   and fixed with 4% (w v^−1) paraformaldehyde solution (Solarbio) for
   30 min at room temperature. After permeabilizing with 0.1% (w v^−1)
   Triton X‐100 (Solarbio) in PBS for 10 min, the samples were blocked
   with 3% bovine serum albumin (BSA) (Solarbio) in PBS for 1 h at room
   temperature. After washing with PBS, the samples were incubated with
   primary antibodies in 5% (w v^−1) BSA in PBS overnight at 4 °C. After
   rinsing to remove excess antibodies, the samples were incubated with
   secondary antibodies in the dark for 1 h at room temperature. The
   secondary antibody used was goat anti‐rabbit IgG H&L (Alexa Fluor 488)
   (1:1000; Abcam). Phalloidin (1:250; Solarbio) was utilized for staining
   the cytoskeleton, and 4′,6‐diamidino‐2‐phenylindole (DAPI; 1:500;
   Solarbio) was employed for cell nuclei staining, both in the dark for
   30 min at room temperature.

   For calvaria tissues, the samples were fixed with 4% (w v^−1)
   paraformaldehyde solution (Solarbio) for 24 h, and then decalcified and
   dehydrated according to standard protocols, paraffin‐embedded, and
   sectioned at a thickness of 5 µm. The samples were deparaffinized,
   dehydrated, and subjected to antigen retrieval. Then, they were
   permeabilized with 0.1% (w v^−1) Triton X‐100 (Solarbio) in PBS for
   10 min, blocked with 3% (w v^−1) BSA (Solarbio) in PBS for 30 min, and
   then incubated with primary antibodies overnight at 4 °C. After washing
   with PBS, the samples were incubated with secondary antibody for 30 min
   at room temperature. After another round of antigen retrieval
   treatment, the next antibody was stained in the same manner. The cell
   nuclei were stained with DAPI (Solarbio).

   Images were captured with a TCS‐SP8 STED 3X confocal laser scanning
   microscope (Leica, Germany). The primary antibodies used were as
   follows: Anti‐Mannose Receptor (CD206) antibody (1:1000; Abcam), CD86
   Polyclonal antibody (1:500; Proteintech), Alexa Fluor 488 Anti‐CD68
   antibody (1:250; Abcam), Anti‐CD31 antibody (1:1000; Abcam), OCN
   Polyclonal antibody (1:500; Proteintech), and Anti‐IL‐10 antibody
   (1:500; Abcam).

Tube‐Formation Assay

   24‐well plates were coated with 250 µL of Matrigel (BD Biosciences) per
   well, ensuring that no air bubbles were introduced. The plates were
   initially placed in a refrigerator at 4 °C for 10 min to level the
   Matrigel, then transferred to an incubator for at least 30 min to
   solidify the Matrigel. After incubating at 37 °C in a 5% CO[2]
   incubator for 4 h, the tubular structures were observed under confocal
   microscopy. The Image Pro Plus software was used to evaluate the tube
   formation.

Sample Collection for Evaluation of Vascular Formation and Microvascular
Perfusion

   After anesthesia, mice were successively perfused with heparinized
   saline, 4% (w v^−1) paraformaldehyde solution, and microfil (Flow Tech,
   USA)^[ [159]^47 , [160]^48 ^] through the cardiac artery. The culled
   mice were then stored overnight at 4 °C. Subsequently, calvarial
   specimens were collected and fixed in 4% (w v^−1) paraformaldehyde
   solution (Solarbio) for 24 h.

Micro‐CT Scanning Analysis

   The mouse calvaria samples were fixed in 4% (w v^−1) paraformaldehyde
   solution (Solarbio) at 4 °C for 24 h and subsequently scanned using a
   micro‐CT SkyScan 1276 instrument (Bruker). The vascular
   microfil‐perfusion calvaria samples were scanned after decalcification.
   3D reconstruction was performed using the accompanying NRecon software,
   and 3D visualization was achieved using Data Viewer and CTvox software,
   with the volume of regenerated bone or vascular network being
   quantitatively analyzed using the CTAn software.

Bulk RNA‐seq

   RNA was extracted from BMDMs using standard methods. RNA integrity was
   evaluated using the RNA Nano 6000 Assay Kit on the Bioanalyzer 2100
   system (Agilent Technologies, USA). To target cDNA fragments between
   370 and 420 bp in length, library fragments were purified using the
   AMPure XP system (Beckman Coulter, USA). PCR products were purified
   with the same system, and library quality was evaluated with the
   Agilent Bioanalyzer 2100 system. The index‐coded samples were clustered
   using the cBot Cluster Generation System with the TruSeq PE Cluster Kit
   v3‐cBot‐HS (Illumina, USA) following the manufacturer's instructions.
   After cluster generation, the library preparations were sequenced on an
   Illumina Novaseq platform, producing 150 bp paired‐end reads.
   Differential expression analysis between the two groups was performed
   using the DESeq2 R package (1.20.0). GO enrichment analysis of
   differentially expressed genes was implemented by the clusterProfiler R
   package, in which gene length bias was corrected. ClusterProfiler R
   package was also used to test the statistical enrichment of
   differentially expressed genes in the Kyoto Encyclopedia of Genes and
   Genomes pathways.

Statistical Analysis

   Statistical analyses were performed using GraphPad Prism (version 9.0)
   software (USA). All quantitative data were expressed as mean ± standard
   deviation (SD) of at least three independent replicate experiments. The
   Shapiro‐Wilk test was used to assess the normality of distributions.
   Statistical differences were analyzed using the unpaired Student's
   t‐test for independent samples. Statistical significance levels were
   set at *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001. A
   p‐value of less than 0.05 was considered statistically significant.

Conflict of Interest

   The authors declare no conflict of interest.

Supporting information

   Supporting Information
   [161]ADVS-12-2414278-s001.docx^ (4.2MB, docx)

Acknowledgements