Abstract
Current Ti‐6Al‐4V bone implants lack trabecular structure and
pro‑angiogenic cues, both essential for regeneration. Herein, a dual
biomimetic strategy is devised that integrates a 3D‐printed biomimetic
trabecular porous Ti‐6Al‐4V scaffold (BTPS) with exosome‐loaded
PEGDA/GelMA hydrogel microspheres (PGHExo) designed for sustained
release. BTPS is designed using Voronoi algorithms and imaging data,
and replicates the geometry and mechanical properties of natural bone.
Hypoxia‐induced human umbilical vein endothelial cell (HUVEC) derived
exosomes (HExo) are encapsulated in PGHExo microspheres via
microfluidic technology, enabling controlled release of HExo, and
anchored onto BTPS using polydopamine (pDA) modification
(BTPS&pDA@PGHExo). BTPS exhibited an elastic modulus of ≈3.2 GPa and a
permeability of 11.52 × 10^−8 mm^2, mimicking natural bone. In vitro
assays demonstrated that BTPS&pDA@PGHExo significantly enhanced
osteogenesis and angiogenesis. mRNA‐Seq analysis suggested that
BTPS&pDA@PGHExo regulates osteogenic and angiogenic gene expression
through the activation of pathways including MAPK, mTOR, HIF‐1, and
VEGF. In vivo, BTPS&pDA@PGHExo improved bone volume, density, and
neovascularization in a rabbit model. This dual biomimetic strategy
offers a promising clinical solution, addressing the limitations of
conventional Ti‐6Al‐4V scaffolds and providing an innovative approach
for personalized bone defect repair.
Keywords: 3D‐printed; angiogenesis; biomimetic trabecular scaffold;
bone regeneration, exosomes
__________________________________________________________________
This study presents 3D‐printed Ti‐6Al‐4V trabecular scaffolds combined
with hypoxia‐induced exosomes encapsulated in PEGDA/GelMA microspheres
for sustained release. The scaffolds mimic the structure of natural
bone, while the exosome delivery system enhances osteogenesis and
angiogenesis, significantly promoting bone regeneration and offering a
promising strategy for the repair of large bone defects.
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1. Introduction
Large segmental bone defects, arising from severe trauma, tumor
resection, or infection, remain a pressing clinical challenge,^[ [52]^1
^] frequently resulting in nonunion, implant failure, and prolonged
morbidity.^[ [53]^2 ^] Conventional interventions, including autografts
and allografts, are limited by donor scarcity and immunological
risks,^[ [54]^3 ^] while synthetic ceramics and polymers^[ [55]^4 ^]
often fail to provide the requisite mechanical strength and biological
cues for robust, vascularized bone regeneration. Titanium alloys,
particularly Ti‐6Al‐4V, have emerged as widely employed for
load‐bearing implants due to their exceptional mechanical strength,
corrosion resistance, and biocompatibility.^[ [56]^5 ^] Moreover,
advances in additive manufacturing now permit patient‐specific implant
designs with intricate porous architectures that closely mimic native
bone structures.^[ [57]^6 ^] Yet, two critical barriers persist. First,
the elastic modulus of Ti‐6Al‐4V (≈110 GPa) substantially exceeds that
of natural bone (2–30 GPa), causing stress shielding and impaired
osseointegration.^[ [58]^7 ^] Second, while mechanically superior,
Ti‐6Al‐4V surfaces are intrinsically bioinert^[ [59]^8 ^] and lack the
molecular signals essential for the formation of functional vascular
networks that underpin sustained bone healing. Overcoming these dual
challenges requires an integrated approach that marries mechanical
optimization with dynamic biological signaling.
Nature's blueprint—the trabecular architecture of cancellous
bone—offers both mechanical resilience and a porous network conducive
to cellular infiltration and angiogenesis. Despite advances in
computational modeling and 3D printing, replicating the inherent
anisotropy and structural complexity of trabecular bone remains
challenging. Most conventional scaffolds rely on regular geometric
patterns that fail to capture this complexity, resulting in suboptimal
stress distribution and limited osseointegration.^[ [60]^9 ^] By
contrast, biomimetic trabecular porous structures inspired by native
bone can reduce elastic modulus, mitigate stress shielding, and promote
angiogenesis, thereby establishing a biomechanically and biologically
favorable milieu for bone regeneration. Although these refined implants
offer improved mechanical support and accelerated bone tissue
formation, they often lack the endogenous cues necessary to drive
robust vascularization and osteogenesis. Enhancing the bioactivity of
biomimetic trabecular Ti‐6Al‐4V scaffolds is therefore crucial for
achieving functional integration with native bone and advancing their
clinical potential in large bone defect repair.
Among these indispensable biological cues, vascularization stands as a
pivotal orchestrator, ensuring continuous nutrient and oxygen delivery,
modulating local inflammation, and directing osteogenic cascades
integral to sustained bone regeneration. Previous
strategies—incorporating growth factors or pre‐seeding endothelial
cells—have yielded incremental gains but often suffer from immunologic
hurdles, low cell viability, and transient factor release.^[ [61]^10 ^]
HUVEC‐derived exosomes, by contrast, offer stable, cell‐free delivery
of pro‐angiogenic factors. Loaded with microRNAs, proteins, and lipids,
exosomes can enhance endothelial sprouting, regulate local
inflammation, and support osteoblast differentiation.^[ [62]^11 ^] Our
preliminary studies indicate that sustained exosome delivery via
microsphere‐based systems can be temporally synchronized with the
vascularization phase of bone healing, providing extended therapeutic
effects.^[ [63]^12 ^] Exosomes thus represent a scalable,
cost‐effective, and biochemically stable modality for promoting
vascularization in bone repair.^[ [64]^13 ^] However, the integration
of soft exosomes into rigid Ti‐6Al‐4V scaffolds in a manner that
preserves their bioactivity and spatial‐temporal release remains a
significant challenge.
Here, we present a dual‐biomimetic strategy that seamlessly integrates
a trabecular‐inspired Ti‐6Al‐4V scaffold with HUVEC‐derived exosomes
encapsulated in dual‐network hydrogel microspheres engineered via
advanced microfluidic techniques. By precisely tuning microsphere size
and surface characteristics, we achieve sustained, controlled exosome
release within the scaffold's pores, significantly enhancing
exosome‐loading efficiency and prolonging bioactivity at the defect
site. pDA modification establishes a robust adhesive interface,^[
[65]^14 ^] promoting stable exosome immobilization and ensuring
compatibility within the biomimetic microenvironment. This localized
bioactive reservoir continuously supplies exosomal factors, driving
matrix formation and tissue integration at the bone interface.
Taken together, this study pioneers a dual‐biomimetic bone repair
paradigm: a 3D‐printed, trabecular‐mimetic Ti‐6Al‐4V scaffold augmented
with pDA‐anchored HExo microspheres for sustained exosome release.
Critically, this composite design overcomes the conventional
limitations of Ti‐6Al‐4V scaffolds—stress shielding and limited
bioactivity—substantially enhancing osteogenesis and vascularized
tissue regeneration. By integrating personalized treatment principles,
our approach allows scaffolds to be tailored to patient‐specific needs,
offering a safe, efficient, and adaptable platform for large bone
defect repair. Through this synergy of structural and biological
mimicry, we not only optimize scaffold mechanics but also achieve
robust bioactivity, fostering superior vascularized bone regeneration
and scaffold‐tissue integration. Ultimately, this work represents a
significant advance in the clinical application of Ti‐6Al‐4V implants,
underscoring its strong potential for translation into patient care. A
schematic illustration of the scaffold fabrication and its therapeutic
mechanism is shown in Figure [66]1.
Figure 1.
Figure 1
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Schematic illustration of the fabrication process and therapeutic
mechanism of BTPS&pDA@PGHExo scaffolds for large bone defect repair.
2. Results
2.1. Design, Fabrication, and Characterization of BTPS
We developed a BTPS that emulates the anatomical structure of the
femoral cancellous bone (Figure [68]2A). Through the Voronoi algorithm
applied to imaging data, the resulting trabecular structure accurately
mirrored the morphology and spatial organization of natural bone
trabeculae. Following extensive parameter adjustments, an anisotropic
BTPS was achieved with a 600 µm pore diameter and 70% porosity. This
design demonstrated considerable geometric complexity and spatial
distribution patterns, closely aligned with the structure of the
femoral cancellous bone. The Ti‐6Al‐4V scaffold with a biomimetic
porous trabecular design was successfully fabricated using Selective
laser melting (SLM) technology.
Figure 2.
Figure 2
[69]Open in a new tab
Characterization and elemental mapping of BTPS. A) Schematic
illustration of the scaffold fabrication process, involving biomimetic
trabecular design and SLM 3D printing. B) Macroscopic images display
surface morphology of BTPS, scale bar, 5 mm. C) Scanning electron
microscopy (SEM, Thermo Fisher, USA) images highlight microstructural
differences, with scale bars at 400 µm and 1 mm. D) Elemental mapping
via energy‐dispersive X‐ray spectroscopy (EDS, Thermo Fisher, USA) for
BTPS, highlighting the distribution of key elements (Ti, C, Al, and V),
with scale bar at 50 µm. E) Reconstruction process and quantitative
analysis of scaffold microstructure. F) Comparison of designed and
measured porosity (n = 3). G) Comparison of designed and measured pore
sizes (n = 3). H) Comparison of designed and measured trabecular
thickness (n = 3). I) Distribution of pore sizes with an average value
marked by a red dashed line. J) Distribution of trabecular thickness
with the average value indicated by a red dashed line. Data are
presented as the mean ± standard deviation (Mean ± SD). Statistical
analysis was performed using one‐way analysis of variance (ANOVA). ^∗ p
< 0.05, ns: p > 0.05.
The BTPS structure and composition were comprehensively evaluated using
a range of advanced characterization techniques. As shown in SEM images
(Figure [70]2B,C), the BTPS surfaces are smooth and free of microcracks
or defects, with uniform pore wall thickness and a dense architecture.
These characteristics reveal an anisotropic and interconnected porous
structure. Moreover, the irregular pore network conforms well to design
specifications. EDS analysis verified that the BTPS primarily comprises
Titanium (Ti), Aluminum (Al), and Vanadium (V), aligning with Ti‐6Al‐4V
standards, displaying uniform element distribution without impurities
(Figure [71]2D). Furthermore, high‐resolution industrial computed
tomography(CT) scanning verified the BTPS's internal integrity and
connectivity, showing a porosity of 69.26%, closely matching the design
target. Pore size analysis revealed an average pore size of 560 µm,
predominantly centered ≈600 µm, and an average trabecular thickness of
248 µm, further confirming structural stability. Taken together, these
findings align with the design objectives (Figure [72]2E–J),
demonstrating that the BTPS achieves the intended structural precision
and uniformity.
2.2. Finite Element, Mechanical, and Fluid Dynamics Analysis of BTPS
Finite element analysis (FEA) was conducted to assess the mechanical
stability of the BTPS under applied loads (Figure [73]3A–C). The design
of the BTPS scaffold, featuring 600 µm pore size and 70% porosity, was
selected based on literature and prior experimental data,^[ [74]^15 ,
[75]^16 ^] which support that this configuration strikes the optimal
balance between mechanical performance, cellular infiltration, and
nutrient exchange. This pore size and porosity combination was chosen
for its ability to replicate the mechanical properties of natural bone
while providing a biologically favorable environment for cell growth
and nutrient flow, making it a representative scaffold for bone repair.
The FEA focused on this optimized design. The analysis revealed a
maximum stress of 454.26 MPa, indicating a uniform stress distribution
without stress concentrations. The maximum displacement was 0.005 mm,
localized primarily near the load application site, which suggests
robust deformation resistance. Consistent with the results of physical
mechanics experiments, these findings further validate the BTPS's
mechanical stability and structural support capabilities.
Figure 3.
Figure 3
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Mechanical characterization, finite element, and fluid dynamics
analysis of BTPS. A) Finite element model of the BTPS under compressive
loading, with boundary conditions applied. B) Von Mises stress
distribution within the scaffold structure under compression. C) The
global and section views show the distribution of displacement across
the scaffold structure. D) Computational Fluid Dynamics (CFD) model
setup with defined inlet, outlet, and symmetric boundary conditions. E)
Velocity vector field illustrating flow direction and distribution
within the scaffold. F) Velocity contour plot showing the magnitude of
fluid flow across scaffold layers. G) Pressure contour illustrating
pressure distribution throughout the scaffold. H) Wall shear stress
contour indicating shear forces on scaffold surfaces. I) Compression
testing setup for evaluating scaffold mechanical properties. J)
Representative images of scaffolds before and after compression. K)
Stress‐strain curve from compression testing, showing the load‐bearing
capacity of BTPS (n = 3). L) Comparison of elastic modulus between
BTPS, cancellous bone, and cortical bone (n = 3).
Fluid mechanics analysis evaluated the BTPS's permeability and fluid
transport properties (Figure [77]3D–H). Results reveal a uniform flow
velocity within the BTPS, where fluid traverses the pore structure
smoothly, without signs of turbulence or stagnation. The streamline
diagram additionally clarifies the fluid pathway and consistent flow
distribution, demonstrating the BTPS design's optimization of fluid
channels. Shear force distributes evenly along the pore wall surface,
with no excessive localized concentrations observed. Pressure
distribution graphs reveal a notable gradient within the BTPS,
particularly at the top and outlet, which signifies a substantial
pressure differential. Such a gradient strongly suggests the
structure's effective fluid channeling capacity. Permeability (K)
measured at 11.52 × 10^−⁸ confirms BTPS's high fluid transfer capacity,
supporting its suitability for maintaining high permeability and mass
transfer efficiency in biomechanical environments.
Physical mechanical testing demonstrated the BTPS's exceptional
mechanical properties (Figure [78]3I–L). The stress‐strain curves were
consistent, with BTPS showing an elastic modulus of 3.2 GPa, placing it
squarely within the range of cancellous and cortical bone, thus closely
replicating the mechanical properties of natural bone. Moreover, an
average yield compression load of 3085.25 N indicates that BTPS
provides the requisite mechanical strength necessary to support bone
tissue repair and maintain structural integrity.
2.3. HUVECs‐Derived Exosomes Extraction and Characterization
As illustrated in Figure [79]4A, HUVEC‐derived exosomes were
successfully extracted under normoxic and hypoxic conditions via a
multi‐step high‐speed centrifugation protocol. Immunofluorescence
staining confirmed that hypoxia significantly elevated VEGFA expression
in HUVECs (Figure [80]4B). Quantitative analysis of exosomal protein
levels showed that Hypo‐Exos contained significantly higher protein
concentrations than Exos (Figure [81]4C). Western blot analysis further
validated the presence of established exosomal markers ALIX, CD9, and
CD81 in both Exos and Hypo‐Exos, confirming the purity and identity of
the exosomes (Figure [82]4D). Transmission Electron Microscopy (TEM)
imaging revealed that both Exos and Hypo‐Exos exhibited a
characteristic round morphology, uniform structure, and an approximate
diameter of 100 nm (Figure [83]4E). Nanoparticle Tracking Analysis
(NTA) results indicated a greater particle concentration in Hypo‐Exos
compared to Exos, although both maintained similar particle size
distributions predominantly between 100 and 150 nm (Figure [84]4F).
Fluorescence microscopy analysis revealed a time‐dependent
internalization of PKH26‐labeled Exos and Hypo‐Exos by recipient cells,
with a significantly enhanced uptake of Hypo‐Exos observed at 72 h. The
quantitative assessment further demonstrated a peak in exosome
internalization at 72 h for both Exos and Hypo‐Exos, with Hypo‐Exos
consistently exhibiting higher cellular uptake throughout the time
points studied (Figure [85]4G; Figure [86]S2, Supporting Information).
Collectively, these findings suggest that Hypo‐Exos displays superior
concentration and functional properties under hypoxic conditions
compared to normoxia. Subsequently, the pro‐angiogenic potential of
Hypo‐Exos was examined across various concentrations (50, 100, 150, and
200 µg mL^−1). The findings revealed that increased Hypo‐Exos
concentrations were associated with an upsurge in vascular‐like
structures, clearly demonstrating a pro‐angiogenic effect. Notably, the
200 µg mL^−1 Hypo‐Exos group exhibited the most prominent formation of
vascular structures (Figure [87]4H). The quantitative analysis
presented in the figure confirms a dose‐dependent effect of Hypo‐Exos
within the range of 50–150 µg mL^−1, showing pronounced angiogenesis
with rising concentrations. At 200 µg mL^−1, the pro‐angiogenic effect
plateaued, suggesting a saturation threshold and an optimal
concentration for angiogenesis (Figure [88]4I).
Figure 4.
Figure 4
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Isolation, characterization, and bioactivity evaluation of HUVEC
‐derived exosomes. A) Schematic representation of differential
ultracentrifugation protocol employed for the isolation of exosomes. B)
Immunofluorescence staining of VEGFA in HUVECs, visualizing VEGFA
expression (green) with nuclear counterstaining (DAPI, blue), scale
bar, 10 µm. C) Quantitative analysis of exosomal protein levels (n =
3). D) Western blot analysis of exosomal markers CD63, CD9, and Alix in
purified exosomes, confirming exosome identity and integrity. E) TEM
images of exosomes showing typical morphology and nanoscale size, scale
bar, 100 nm. F) NTA of exosomes particle size distribution confirming
homogeneity. G) Time‐dependent internalization of Exos and Hypo‐Exos by
recipient cells (n = 3). H) Representative images showing angiogenic
network formation at Hypo‐Exos concentrations of 50, 100, 150, and 200
µg mL^−1, scale bar, 100 µm. I) Quantitative analysis of total mesh
area across concentrations, indicating a concentration‐dependent
increase in angiogenesis (n = 3). Data are presented as the Mean ± SD.
Statistical analysis was performed using one‐way ANOVA. ns, no
significant difference; ^** p < 0.01; and ^*** p < 0.001.
2.4. Preparation and Characterization of PGHExo Microspheres
Figure [90]5A illustrates the successful fabrication of PGHExo
dual‐network hydrogel sustained‐release microspheres via microfluidic
chip technology. The microspheres exhibited smooth surfaces,
homogeneous internal structures, and well‐defined morphology, with no
apparent defects. Particle size distribution analysis revealed that the
microspheres maintained a consistent size, averaging ≈80 µm within a
range of 60–90 µm(Figure [91]5C,D). Rheological analysis indicated that
the PEGDA/GelMA (PG) gel sustained a stable storage modulus (G′) and
loss modulus (G″) across various shear frequencies, exceeding those of
the GelMA single‐component gel, thereby demonstrating superior
mechanical stability (Figure [92]5B). To evaluate protein release
profiles, PG samples with Hypo‐Exos at various concentrations
(1‐3 mg mL^−1; PGHExo1, PGHExo2, and PGHExo3) and without Hypo‐Exos
were quantified. Notably, PG samples lacking Hypo‐Exos exhibited
minimal protein release throughout the 18‐day period. Conversely,
samples containing Hypo‐Exos demonstrated sustained protein release,
with release rates directly correlated to Hypo‐Exo concentration
(Figure [93]5E). Given that the peak internalization of exosomes
occurred at 72 h, we monitored the release concentration of PGHExo2
every three days. The average release concentration remained ≈200
µg mL^−1 (Figure [94]5F), which is optimal for angiogenesis induction.
These findings confirm PGHExo2 as the most promising candidate for bone
repair applications. To monitor PKH26‐labeled Hypo‐Exos release,
confocal microscopy was employed. The observations revealed that
Hypo‐Exos in PGHExo2 were gradually released, sustaining for up to 18
days (Figure [95]5G). Both immunofluorescence imaging and quantitative
fluorescence analysis of PGHExo2 revealed consistent trends, confirming
effective Hypo‐Exo release within the dual‐network microspheres
(Figure [96]5H). Collectively, these findings underscore that PGHExo
dual‐network microspheres exhibit favorable stability and
controlled‐release capabilities, supporting their suitability for
sustained exosome delivery to HUVECs and enhanced scaffold bioactivity.
Figure 5.
Figure 5
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Fabrication, characterization, and release profile of PEGDA/GelMA
dual‐network hydrogel sustained‐release microspheres loaded with
Hypo‐Exos. A) Schematic of the microfluidic chip setup for generating
uniform PGHExo, alongside a bright‐field microscopy image demonstrating
high monodispersity, scale bar, 100 µm. B) Rheological properties of 1%
GelMA and 1% PGgel (n = 3). C) SEM image revealing surface morphology
and consistent size distribution of microspheres. The inset shows an
enlarged view, scale bar, 50 µm. D) Histogram of microsphere size
distribution. E) Cumulative protein release profile of PGHExo
microspheres over 18 days (n = 3). F) Concentration of protein released
every 3 days (n = 3). G) Confocal microscopy images of PKH26‐labeled
PGHExo2 microspheres showing exosome release at different time points
(0d, 3d, 6d, 9d, 12d, 15d, and 18d). Top row: fluorescence images;
Middle row: light images; Bottom row: merged images. Scale bar, 50 µm.
H) Quantitative analysis of relative fluorescence intensity over time,
indicating a gradual decrease in fluorescence, reflecting sustained
exosome release from PGHExo2 microspheres (n = 3).
2.5. In Vitro Bioactivity Assessment of BTPS&pDA@PGHExo
We initiated the fabrication and performed a detailed characterization
of BTPS&pDA@PGHExo. Figure [98]6A illustrates the fabrication process
and the chemical interactions between the 3D‐printed BTPS scaffold, the
PDA coating, and the externally loaded PGHExo. The BTPS scaffold
underwent plasma treatment to introduce hydroxyl (─OH) groups, which
facilitate the adhesion of the PDA coating. This coating, in turn,
promotes the attachment of the PGHExo microspheres, ensuring that the
exosome‐loaded microspheres are securely anchored to the scaffold,
forming an integrated structure. The untreated BTPS presented a
silvery‐white appearance upon visual inspection (Figure [99]2B). After
pDA modification, the BTPS surface was uniformly coated with a black
film (Figure [100]6B). SEM imaging confirmed the presence of a
continuous pDA coating on the BTPS surface (Figure [101]6C). EDS
analysis (Figure [102]6D) indicated that BTPS&pDA was primarily
composed of carbon (C), nitrogen (N), oxygen (O), and Ti, thereby
validating successful pDA coating. SEM images of BTPS&pDA@PGHExo showed
a uniform distribution of PGHExo microspheres on the BTPS surface,
suggesting favorable adhesion properties. Elemental analysis further
identified C, O, N, Ti, phosphorus (P), sodium (Na), and chloride (Cl)
(Figure [103]6E–H; Figure [104]S3, Supporting Information). Notably,
the detected C, O, N, and P within the exosomes likely originated from
their complex composition, comprising lipids, proteins, and RNA. The
detected Na and Cl likely resulted from saline treatment. Together,
these findings confirm the successful pDA coating modification and
efficient assembly of PGHExo microspheres onto the BTPS surface.
Figure 6.
Figure 6
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Development, characterization, and functional assessment of
BTPS&pDA@PGHExo. A) Schematic illustrating the fabrication of the
BTPS&pDA@PGHExo. B) Macroscopic images display surface morphology of
BTPS&pDA, scale bar, 5 mm. C) SEM images of BTPS&pDA, with scale bars
at 400 µm and 1 mm. D) Elemental mapping via EDS for BTPS&pDA,
highlighting the distribution of key elements (C, N, O, and Ti), with
scale bar at 100 µm. E) Macroscopic images display surface morphology
of BTPS&pDA@PGHExo, scale bar, 5 mm. F) SEM images of BTPS&pDA@PGHExo,
with scale bars at 400 µm and 1 mm. G) Elemental mapping via EDS for
BTPS&pDA@PGHExo, highlighting the distribution of key elements (C, N,
O, and P) and (Ti, Cl, and Na) (Figure [106]S3, Supporting
Information), with scale bar at 50 µm. H) Quantitative analysis of
elemental composition, demonstrating uniform distribution and effective
surface modification, confirming the successful integration of PGHExo
microspheres within the scaffold structure. I) Schematic of MC3T3‐E1
cell co‐culture with each scaffold group. J) Live cell fluorescence
imaging on BTPS, BTPS&pDA, BTPS&pDA@PG, and BTPS&pDA@PGHExo scaffolds.
Scale bar, 200 µm. K) Quantification of live cell count on different
scaffolds (n = 3). L) Fluorescence intensity measurements over 1, 3,
and 5 days, indicated increased cell proliferation on modified
scaffolds, particularly on BTPS&pDA@PG (n = 3). M) Alkaline Phosphatase
(ALP) activity at day 7, showing elevated osteogenic differentiation on
BTPS&pDA@PG scaffolds (n = 3). N) Quantitative analysis of
mineralization at 21 days, with absorbance measured at 620 nm,
indicated significantly higher mineralization on BTPS&pDA@PGHExo (n =
3). O) Macroscopic images of BTPS, BTPS&pDA, BTPS&pDA@PG, and
BTPS&pDA@PGHExo after Alizarin Red staining. Scale bar, 5 mm. P–S) The
Real‐time PCR (RT‐PCR)results of osteogenesis‐associated gene
expression (n = 3). T–W) The RT‐PCR results of angiogenesis‐associated
gene expression (n = 3). Figures [107]6K–N,P–W share a single set of
grouping information. Data are presented as the Mean ± SD. Statistical
analysis was performed using one‐way ANOVA. ^* p < 0.05; ^** p < 0.01;
^*** p < 0.001; and ^**** p < 0.0001.
Cell viability assays demonstrated that all scaffold materials
exhibited strong cytocompatibility, showing no signs of cell death.
Importantly, MC3T3‐E1 cell survival rates in the BTPS&pDA@PGHExo group
were significantly elevated compared to other groups, as evidenced by
an intensified green fluorescence signal. Qualitative results
corroborated the increased cell viability and survival rates in this
group, underscoring the scaffold's potential to promote cell survival
and functional maintenance (Figure [108]6I–K). Cell proliferation
assays further demonstrated the superior biocompatibility of the
BTPS&pDA@PGHExo. Cell proliferation rates increased progressively over
time across all samples (Figure [109]6L). The BTPS&pDA@PGHExo exhibited
significantly higher fluorescence intensity, indicating the most robust
cell proliferation among the groups. In contrast, the BTPS and BTPS&pDA
groups showed comparatively lower proliferation rates, while the
BTPS&pDA@PG group displayed a moderate increase. These findings are
consistent with the live/dead cell data, reinforcing the enhanced
capacity of the BTPS&pDA@PGHExo to support cell growth.
The ALP assay and Alizarin Red S mineralization assay were performed to
evaluate the osteogenic potential of the scaffolds. The BTPS&pDA@PGHExo
group exhibited the highest ALP activity among all groups,
significantly surpassing the BTPS, BTPS&pDA, and BTPS&pDA@PG groups,
indicating an enhanced osteogenic differentiation (Figure [110]6M).
After 21 days of incubation in an osteogenic medium, the
BTPS&pDA@PGHExo also demonstrated substantial mineralized nodule
formation, evidenced by more intense reddish‐orange staining compared
to other groups (Figure [111]6O). Quantitative analysis confirmed that
this group achieved the highest mineralization level, with absorbance
values significantly higher than those of the other groups
(Figure [112]6N). RT‐PCR analysis further supported these findings by
showing a significant upregulation of osteogenesis‐related
markers—including ALP, OCN, COL1, and RUNX2—in the BTPS&pDA@PGHExo
group compared to the other groups (Figure [113]6P–S). This group
consistently demonstrated the highest levels of osteogenic gene
expression, indicating the synergistic effects of the exosome
incorporation. These results suggest that the BTPS&pDA@PGHExo
significantly promotes osteogenic differentiation and mineralization,
potentially by enhancing the mineralization conditions within the
extracellular matrix.
Beyond its osteogenic properties, the BTPS&pDA@PGHExo scaffold
demonstrated significant pro‐angiogenic potential. Angiogenesis was
assessed by quantifying mRNA expression levels of VEGF, CD31, PDGFB,
and FGF2. Among the groups, BTPS&pDA@PGHExo exhibited the highest
expression levels for all markers, with VEGF showing a nearly sixfold
upregulation (p < 0.0001). Similarly, CD31, PDGFB, and FGF2 were
significantly upregulated (p < 0.0001), indicating enhanced angiogenic
activity. These findings underscore the exosome‐enriched scaffold's
ability to establish a pro‐angiogenic microenvironment, which is
critical for effective bone regeneration (Figure [114]6T–W).
2.6. mRNA‐seq Analyses of BTPS&pDA@PGHExo
The mRNA‐seq analyses revealed a total of 28991 genes, with 20678
(73.61%) commonly expressed between the two groups. The volcano plot
demonstrated significant transcriptional changes, including 5102
upregulated and 6049 downregulated genes. Gene Ontology (GO)enrichment
analysis identified key biological processes such as mitochondrial
function, bone morphogenesis, and hypoxia‐induced angiogenesis,
alongside cellular components and molecular functions like
extracellular matrix organization and growth factor binding.
Additionally, Kyoto Encyclopedia of Genes and Genomes (KEGG)pathway
analysis highlighted critical signaling pathways, including MAPK, mTOR,
HIF‐1, and VEGF, as pivotal regulators of osteogenesis and
angiogenesis. The heatmap analysis of DEGs underscored significant
genes, including ALPL, COL18A1, SAMD6, and RUNX2OS1 for bone‐related
functions, as well as VEGFB, PDGFA, and ANGPT2 for angiogenesis. These
findings demonstrate the BTPS&pDA@PGHExo ability to modulate key
molecular pathways, orchestrate osteogenic and angiogenic responses,
and facilitate extracellular matrix remodeling, further supporting its
potential for enhanced bone regeneration and vascularisation in
translational applications (Figure [115]7A–H).
Figure 7.
Figure 7
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mRNA‐seq analyses of gene expression. A) Venn diagram showing the
overlap of DEGs between two scaffold treatment groups. B) Volcano plot
showing DEGs between scaffold treatment groups, with upregulated (red)
and downregulated (blue) genes highlighted. C) Bar chart illustrating
the total number of upregulated and downregulated DEGs. D) Heatmap of
DEGs between BTPS&pDA@PGHExo and Solid Ti‐6Al‐4V scaffold treatments,
with red indicating upregulated genes and blue indicating downregulated
genes. E–G) GO enrichment analysis of DEGs categorized by Biological
Process (E), Cellular Component (F), and Molecular Function (G). H)
KEGG pathway enrichment analysis of DEGs, identifying significantly
enriched pathways such as MAPK, mTOR, HIF‐1, and VEGF signaling, which
are associated with scaffold‐induced cellular responses. The size and
color of each dot represent the count of DEGs in each pathway and the
adjusted p‐value, respectively. I) RT‐PCR analysis of bone‐related DEGs
(n = 3). J) RT‐PCR analysis of angiogenesis‐related DEGs (n = 3). K–N)
Enzyme‐Linked Immunosorbent Assay (ELISA) quantification of protein
levels: The protein concentrations of RUNX2, OCN, PDGF, and VEGF were
measured by ELISA (n = 3). Figure [117]7I–N share a single set of
grouping information. Data are presented as the Mean ± SD. Statistical
analysis was performed using one‐way ANOVA. ^* p < 0.05; ^** p < 0.01;
and ^*** p < 0.001.
To validate the RNA‐seq findings, quantitative Real‐Time
PCR(qRT‐PCR) was performed on key osteogenic and angiogenic DEGs
(Figure [118]7I,J). The results showed a significant increase in ALPL,
COL18A1, SAMD6, and RUNX2OS1 expression in the BTPS&pDA@PGHExo group
compared to the control, confirming RNA‐seq‐predicted upregulation of
bone‐related genes. Similarly, VEGFB, PDGFA, and ANGPT2 expression was
significantly elevated, verifying the angiogenic potential.
Furthermore, ELISA quantification confirmed increased protein
expression levels of RUNX2, OCN, PDGF, and VEGF in the BTPS&pDA@PGHExo
group (Figure [119]7K–N). These findings demonstrate that the
transcriptional changes observed in RNA‐seq data are effectively
translated into protein‐level expression, reinforcing the functional
impact of BTPS&pDA@PGHExo on bone regeneration and angiogenesis.
2.7. In Vivo Bone Defect Repair with BTPS&pDA@PGHExo
The efficacy of various implants in bone defect repair was
comprehensively assessed using Microcomputed Tomography (Micro‐CT) and
histopathological analyses at 4 and 12 weeks post‐surgery. Figure
[120]8A–E demonstrates that the BTPS&pDA@PGHExo group exhibited
significantly higher new bone volume and density compared to other
groups at both time points. Specifically, at 4 weeks, this group
achieved new bone volumes of 19.3 mm^3 and bone densities of
653.9 mg cm^− ^3, which further increased to 28.1 mm^3 and
717.1 mg cm^− ^3 by 12 weeks (p < 0.05). In contrast, the BTPS and
BTPS&pDA@PG groups showed diminished new bone formation and density,
while the blank group presented the least favorable outcomes. The
complementary histopathological analysis presented in Figure [121]8F–I
revealed that the BTPS&pDA@PGHExo implants facilitated significant new
bone formation and a denser vascular network within the defect site at
both 4 and 12 weeks. At the early stage (4 weeks), there was notable
new bone presence and vascularization, which progressed to advanced
bone maturation and vascular densification by 12 weeks. Quantitative
assessments confirmed that both new bone volume and vascular density
were markedly elevated in the BTPS&pDA@PGHExo group compared to
controls at both time points (p < 0.05). Additionally, histological
examinations indicated more uniform cell morphology and a denser tissue
structure in the BTPS&pDA@PGHExo group, underscoring the implant's
enhanced osteogenic and angiogenic potential. Collectively, the
integrated Micro‐CT and histopathological data illustrate that the
BTPS&pDA@PGHExo composite scaffold significantly promotes bone
regeneration and vascularization, demonstrating superior performance in
facilitating bone defect repair relative to other tested implants.
Figure 8.
Figure 8
[122]Open in a new tab
In vivo assessment of bone regeneration efficacy across scaffold groups
in a rabbit femoral defect model. A) Schematic overview and
intraoperative photographs illustrating the creation of femoral defects
and scaffold implantation in a rabbit model. B) Representative Micro‐CT
images showing defect sites treated with Blank, BTPS, BTPS&pDA@PG, and
BTPS&pDA@PGHExo scaffolds, highlighting differences in new bone
formation across groups. C) 3D Micro‐CT reconstructions and sectional
views demonstrating scaffold integration and bone ingrowth within the
defect regions for each scaffold type. D) Bone volume to total volume
ratio (BV/TV%) analysis at 4 and 12 weeks post‐implantation, showed
significantly higher bone formation in BTPS&pDA@PGHExo scaffolds
compared to other groups (n = 3). E) Bone mineral density (BMD)
analysis at 4 and 12 weeks, indicating enhanced mineralization in the
BTPS&pDA@PGHExo scaffold group (n = 3). F) Representative macroscopic
images of femoral defect sites at various time points
post‐implantation, comparing healing progression among the Blank, BTPS,
BTPS&pDA@PG, and BTPS&pDA@PGHExo groups. G) Methylene
blue/fuchsin‐stained sections of BTPS, BTPS&pDA@PG, and BTPS&pDA@PGHExo
scaffolds at 4 weeks (left panels) and 12 weeks (right panels)
post‐implantation. Images captured at different magnifications
highlight differences in new bone formation, cellular infiltration, and
scaffold integration. Yellow arrows denote new bone formation, green
arrows indicate scaffold material and white arrows point to newly
formed blood vessels. Scale bars, 300 and 50 µm. H) Area of new bone
formation for BTPS, BTPS&pDA@PG, and BTPS&pDA@PGHExo scaffolds, with
BTPS&pDA@PGHExo showing significantly increased bone formation at both
4 and 12 weeks (n = 3). I) Vessel count in the region of interest (ROI)
across scaffold groups, indicating enhanced neovascularization in the
BTPS&pDA@PGHExo group (n = 3). Data are presented as the Mean ± SD.
Statistical analysis was performed using one‐way ANOVA. ^* p < 0.05;
^** p < 0.01; and ^**** p < 0.0001.
3. Discussion
In this study, we introduce a dual biomimetic framework that
intricately combines anatomical fidelity and mechanical congruence with
sustained, biologically active signaling to enhance vascularized bone
regeneration. Utilizing a data‐driven 3D‐printing methodology alongside
a Voronoi‐based architectural design, our Ti‐6Al‐4V scaffold
meticulously replicates the porous trabecular structure and mechanical
properties of native cancellous bone, effectively mitigating stress
shielding and facilitating cellular infiltration and tissue remodeling.
Simultaneously, the incorporation of a PEGDA/GelMA dual‐network
hydrogel microsphere enables the controlled, long‐term release of HExo,
providing continuous pro‐osteogenic and pro‐angiogenic stimuli. This
synergistic integration of structural and biochemical elements
establishes a robust regenerative platform that promotes substantial
new bone formation and a well‐organized vascular network, addressing
significant challenges in orthopedic tissue engineering and offering a
promising solution for the repair of large bone defects.
The microstructure and mechanical properties of bone tissue are
essential in supporting effective bone regeneration.^[ [123]^17 ^] The
integration of image data with a Voronoi algorithm facilitated the
construction of a 3D porous structure that closely mimics natural bone
trabeculae. This anatomically‐driven data provides a solid foundation
for precise and tailored scaffold design. The Voronoi algorithm
translated trabecular structural data into a mathematical model,
enabling precise customization of pore geometry and spatial
heterogeneity. This advanced approach closely replicates the mechanical
and biological properties of natural bone, surpassing conventional
isotropic designs by better addressing directional mechanical demands.
Parameters such as pore size, porosity, and trabecular thickness are
precisely modulated by adjusting seed point distribution and density.
To ensure precise microstructural adjustments, geometric
characteristics of natural trabeculae—including morphology, spatial
distribution, anisotropy, and spatial randomness—are meticulously
replicated. As a result, an anisotropic porous scaffold with a 600 µm
pore size and 70% porosity was fabricated successfully. Compared to
conventional porous structures (e.g., cubic, cylindrical, and triply
periodic minimal surfaces), the Voronoi‐based structure demonstrates
enhanced complexity and morphological diversity, more closely mimicking
natural trabeculae. Conventional porous structures are typically
isotropic, limiting their ability to meet the directional mechanical
demands of bone tissue.^[ [124]^18 ^] The anisotropic properties of the
Voronoi structure, optimized through image data integration, support
scaffold designs that satisfy direction‐specific mechanical
requirements. Consequently, this integrated approach enhances the
scaffold's mechanical adaptability and biological compatibility,
rendering it suitable for diverse loading conditions.
We successfully employed 3D printing technology to fabricate a porous
Ti‐6Al‐4V scaffold with a biomimetic trabecular structure. Finite
element analysis and mechanical testing reveal the scaffold's robust
mechanical properties, particularly its high strength and low elastic
modulus. Importantly, the scaffold exhibits an elastic modulus of
3.2 GPa, markedly lower than the conventional Ti‐6Al‐4V modulus of
≈110 GPa. A high elastic modulus in traditional implants commonly
induces stress shielding, thereby hindering bone tissue regeneration.^[
[125]^19 ^] By contrast, the scaffold's elastic modulus approximates
that of natural cancellous bone, thereby enhancing biomechanical
compatibility. This mechanical match minimizes stress shielding,
fosters a conducive microenvironment, and optimizes stress transfer to
new bone tissue, thereby mitigating issues like bone resorption and
implant loosening commonly seen in standard Ti‐6Al‐4V implants. These
findings are consistent with those of Wieding et al.,^[ [126]^20 ^] who
found that optimizing porous Ti‐6Al‐4V scaffold geometry effectively
reduces the elastic modulus, aligning with bone mechanics and enhancing
osseointegration. Additionally, CFD analysis demonstrated favorable
permeability (11.52 × 10^−8 mm^2) and uniform fluid distribution across
the porous scaffold. The scaffold's porous structure not only promotes
cell adhesion, proliferation, and differentiation but also facilitates
vascular and nutrient infiltration. This is supported by enhanced cell
viability, proliferation, and angiogenic marker expression in vitro.
These features are crucial for supporting cellular growth,
neovascularization, and tissue regeneration, aligning with the
principles of effective bone tissue engineering. These attributes
encourage neovascularization and tissue regeneration, upholding the
principle of “structure‐function unity” central to bone tissue
engineering. Uniform fluid shear and pressure distribution simulate the
natural bone marrow environment, further promoting bone regeneration.
This observation aligns with Foroughi et al.,^[ [127]^21 ^] who
emphasized the essential role of fluid mechanics in porous scaffolds
for tissue engineering.
Building upon the optimized structural and mechanical framework, pDA
was employed as a surface modifier to enhance the scaffold's synergy
with bioactive compounds. pDA exhibits adhesion properties akin to
mussel foot proteins, allowing a stable coating to form on the scaffold
surface. This coating supplies a rich density of active functional
groups, which facilitates the binding of bioactive molecules.^[
[128]^22 ^] SEM and EDS analyses confirmed both effective pDA
modification and the successful integration of PGHExo microspheres on
the BTPS surface. The addition of pDA significantly enhances BTPS
biocompatibility, creating an ideal interface for anchoring
exosome‐loaded double‐network hydrogel microspheres. Cell viability and
proliferation assays indicated markedly higher cell survival and growth
rates in the BTPS&pDA group compared to the unmodified BTPS group.
Consistent with findings by Wang et al.,^[ [129]^23 ^] these results
further validate that pDA coatings enhance the biofunctionalization of
scaffold materials.
The bioactive properties of the porous structure in biomimetic
trabecular bone are crucial for facilitating osteogenesis and
angiogenesis.^[ [130]^24 ^] Importantly, the development and
application of PGHExo, a dual‐network hydrogel microsphere engineered
for controlled release under hypoxic conditions in HUVECs, marks a
notable innovation in this study. HUVEC‐derived exosomes were
successfully isolated and analyzed under both normoxic and hypoxic
conditions using a multi‐step, high‐speed centrifugation protocol.
Moreover, the results revealed that hypoxia markedly increased the
protein content, particle concentration, and cellular uptake of
exosomes, thus enhancing their angiogenic potential. Notably, this
pro‐angiogenic effect was dose‐dependent, indicating that higher
exosome concentrations could amplify angiogenic responses. However, at
higher concentrations (200 µg mL^−1), a saturation effect emerged,
constraining further increases in angiogenic activity. Consistent with
prior research, these findings illustrate that hypoxia upregulates VEGF
expression and augments exosome release, reinforcing hypoxia's pivotal
role in angiogenesis.^[ [131]^25 ^] Utilizing microfluidic technology,
we successfully fabricated dual‐network hydrogel microspheres embedding
Hypo‐Exo within a PEGDA/GelMA matrix, enabling controlled, sustained
exosome release. Characterization of PGHExo microspheres revealed a
uniform particle size, high encapsulation efficiency, and controlled
release profile, validating the efficacy of microfluidic technology.
Compared to conventional exosome delivery methods, the PGHExo system
extended exosome release significantly, sustaining release for up to 18
days. This extended‐release sharply contrasts with the brief 5–7‐day
release profiles associated with direct injection or electrospinning
techniques.^[ [132]^26 ^] This controlled‐release system ensured
prolonged osteogenic and angiogenic stimulation in both in vitro and in
vivo environments.
Exosomes are essential mediators of intercellular communication,
transporting various bioactive molecules—such as microRNAs, proteins,
and lipids—that collectively regulate cellular processes.^[ [133]^27 ^]
In our in vitro studies, MC3T3‐E1 osteoblasts exposed to
BTPS&pDA@PGHExo exhibited the highest proliferation, ALP activity,
mineralized nodule formation, and upregulation of osteogenesis‐related
genes compared with all other groups. These effects likely arise from
the activation of osteogenic signaling pathways by specific exosomal
microRNAs and proteins. Moreover, qRT‐PCR revealed significantly
elevated mRNA levels of angiogenesis‐related genes (e.g., VEGF, CD31,
PDGFB, FGF2) in the BTPS&pDA@PGHExo group, suggesting that exosomal
pro‐angiogenic factors such as VEGF and FGF2 enhance endothelial cell
function and augment angiogenic capacity. Given that angiogenesis is
crucial for bone regeneration—providing essential nutrients and
oxygen—these findings support the notion that exosomes effectively
coordinate osteogenic and angiogenic processes. Consistent with this,
mRNA‐seq analyses demonstrated elevated expression of
osteogenesis‐related genes (e.g., ALPL, COL18A1, SAMD6, RUNX2OS1) and
angiogenesis‐related genes (e.g., VEGFB, PDGFA, ANGPT2) associated with
pathways including MAPK, mTOR, HIF‐1, and VEGF. These results were
further validated by the qRT‐PCR and ELISA experiments, which confirmed
that the observed gene expression changes were reflected at the protein
level. Collectively, these data indicate that BTPS&pDA@PGHExo enhances
osteogenic differentiation by orchestrating multiple signaling
pathways. Additionally, Exosomes may also promote tissue repair by
modulating mitochondrial function, inflammation, and macrophage
polarization states.^[ [134]^28 ^]
Crucially, the hypoxia‐induced endothelial exosomes embedded within our
dual‐network hydrogel microspheres contain a rich cargo of bioactive
molecules, including specific microRNAs (e.g., miR‐210, miR‐21) and
proteins (e.g., VEGF, FGF2), known to drive osteogenic and angiogenic
responses.^[ [135]^29 ^] Under hypoxic conditions, stabilized HIF‐1α in
endothelial cells enhances the packaging of these regulatory miRNAs and
proteins into exosomes, which are then internalized by osteoprogenitor
and endothelial progenitor cells. These exosomal miRNAs modulate key
pathways such as HIF‐1, VEGF, and mTOR, facilitating the proliferation,
migration, and differentiation of cells essential for bone and vascular
formation.^[ [136]^30 ^] For instance, miR‐210 upregulates HIF‐1 target
genes, increasing VEGF secretion and promoting neovascularization while
simultaneously influencing osteoblast lineage commitment and matrix
mineralization.^[ [137]^31 ^] Similarly, the protein cargo of these
exosomes can directly activate pro‐angiogenic signaling cascades,
enhancing vascular ingrowth and tissue perfusion.^[ [138]^32 ^] By
integrating these hypoxia‐induced exosomes into PEGDA/GelMA
dual‐network hydrogel microspheres, our approach ensures their
spatially and temporally controlled release, maintaining a sustained,
localized regulatory influence on the host microenvironment. These
findings align with prior evidence that exosomes modulate cellular
metabolism, inflammatory responses, and macrophage polarization—key
elements of the regenerative cascade.^[ [139]^33 ^] This precisely
engineered, molecularly coordinated strategy underpins the enhanced
osteogenesis and angiogenesis observed in our experiments, broadening
our understanding of how biomaterials can harness endogenous signaling
networks to promote functional tissue regeneration. In a preclinical in
vivo rabbit femoral defect model, the BTPS&pDA@PGHExo scaffold
exhibited notable efficacy in promoting bone repair. Micro‐CT and
histological analyses further revealed that, at 4 and 12 weeks
post‐surgery, the BTPS&pDA@PGHExo group displayed significantly greater
bone volume and density than control groups. Additionally, the defect
region in this group exhibited extensive new trabecular bone formation
and an organized vascular network. Taken together, these findings
underscore the pivotal role of the dual‐network hydrogel
sustained‐release microsphere loaded with hypoxia‐induced HUVEC
exosomes in facilitating vascularized bone regeneration. These results
align with previous findings by Hu et al.,^[ [140]^34 ^] which
underscore the critical role of exosomes in bone tissue regeneration.
Hence, this study's scaffold system offers several critical advantages
over conventional Ti‐6Al‐4V scaffolds. Specifically, the biomimetic
design, enabled by advanced 3D printing, allows the scaffold's
mechanical properties to closely resemble natural bone, effectively
minimizing the mechanical mismatch with host tissue. Moreover, the
incorporation of sustained‐release dual‐network hydrogels, driven by
hypoxia‐induced HUVEC‐derived exosomes, markedly enhances scaffold
bioactivity, promoting vascularized bone formation. Finally, the
versatility of 3D printing technology allows precise scaffold
customization, addressing the specific anatomical and functional needs
of individual patients.
While encouraging, these promising results come with certain inherent
limitations. The molecular pathways by which exosomes facilitate bone
regeneration are not yet fully delineated, underscoring the need for
additional studies to achieve a comprehensive understanding.
Furthermore, the limited duration of the animal experiments constrains
our understanding of the scaffold's long‐term efficacy and stability.
Future investigations should focus on elucidating the specific
regulatory mechanisms of exosomes at cellular and molecular levels.
Additionally, larger‐scale animal studies and preclinical evaluations
are planned to thoroughly assess the feasibility and safety of this
therapeutic strategy.
4. Conclusion
Herein, we establish a dual biomimetic paradigm that integrates a
structurally inspired 3D‐printed Ti‐6Al‐4V trabecular scaffold with an
HExo biochemical cue to facilitate vascularized bone regeneration. By
meticulously adjusting both the trabecular anatomical morphology and
mechanical properties of the scaffold to closely mimic native
cancellous bone, and by enabling prolonged exosome release through a
PEGDA/GelMA dual‐network hydrogel, our approach mitigates stress
shielding, enhances cellular infiltration, and sustains osteogenic and
angiogenic signaling over clinically relevant timescales. Beyond
yielding improved bone volume, mineralization, and neovascularization
in vivo, this platform addresses critical hurdles in treating large
bone defects and complex skeletal injuries, and may be extended to
other tissues requiring concurrent mechanical and vascular integration.
Future investigations will delve deeper into the molecular
underpinnings of exosome‐mediated regeneration, harness
patient‐specific anatomies for personalized scaffold design, and ensure
long‐term safety and functional stability. Collectively, our findings
mark a pivotal step toward next‐generation, bioinspired implants that
seamlessly integrate into host tissues, bridging the gap between
engineered constructs and the natural complexity of human bone.
5. Experimental Section
Design of the BTPS
To develop a biomimetic trabecular porous structure reflecting human
anatomical characteristics, this study first employs CT scanning to
obtain high‐resolution images of trabecular bone, thereby enabling the
extraction of microstructural details and subsequent 3D modeling.
Subsequently, Rhino software (McNeal, Seattle, W A, USA) and its
Grasshopper (v.0.9.0076) plug‐in were employed to design a
parameterized porous structure based on the Voronoi algorithm.
Real‐time adjustments to the algorithm parameters ensure that the
porous structure closely replicates the natural anatomical
configuration and spatial organization of the femoral trabecular bone.
With reference to the design methodology of the previous study,^[
[141]^15 ^] finite element and fluid mechanics analyses optimized a
pore size of 600 µm and a porosity of 70% in the anisotropic biomimetic
trabecular porous structure. This design not only replicates the
intricate geometry of natural bone trabeculae but also offers
substantial mechanical and biological benefits, including enhanced
mechanical adaptability and a supportive biological microenvironment.
The detailed optimization process of the biomimetic trabecular porous
structure design is provided in Figure [142]S1 (Supporting
Information).
Preparation and Characterization of BTPS via 3D Printing
The BTPS was fabricated in the United Kingdom using a Renishaw metal 3D
printer with Ti‐6Al‐4V ELI‐0406 powder. To prevent oxidation, the
powder was protected by an argon gas atmosphere throughout the printing
process. SLM technology was employed under precise conditions,
including a 70 µm laser spot diameter, a 170 °C substrate temperature,
30 µm layer thickness, and a 67° rotation per layer. Additionally, the
laser scanning speed was adjusted based on dot spacing, exposure time,
and drill delay to ensure optimal printing quality. Following the
printing process, the excess powder was removed, and post‐processing
produced a BTPS with dimensions of 10 mm in both height and diameter.
For the cell culture, an additional BTPS sample was prepared with a
height of 2 mm and a diameter of 10 mm. The surface morphology of the
scaffold was characterized by SEM, while EDS was used to map the
elemental distribution. The internal structure accuracy was further
verified by high‐resolution industrial CT scanning (RX Solutions, Wuxi
Raider Detection Technology Co., Ltd., Wuxi, China). Quantitative
analysis of porosity, pore size, and trabecular thickness was conducted
using Avizo and VG Studio Max (Wuxi Rui de Inspection Technology Co.,
Ltd., Wuxi, China) software.
Finite Element Analysis of BTPS
The primary aim of this experiment was to evaluate and refine the
mechanical properties of the BTPS. First, a comprehensive 3D solid
model of BTPS was constructed in 3‐Matic software (Materialise NV,
Belgium, Version 11.0), followed by generating a high‐quality surface
and volume mesh. Subsequently, Ti‐6Al‐4V material properties (elastic
modulus 110 GPa, Poisson's ratio 0.3)^[ [143]^35 ^] were assigned in
Mimics (Materialise NV, Belgium, Version 19.0), and the model was
exported in.cbd format to ANSYS Workbench 18.0 (ANSYS, Inc., USA) to
establish the finite element model. To accurately simulate
physiological loading conditions, fixed constraints were applied at the
lower end of the BTPS model, while an axial compression load of 500 N
was introduced at the upper end. Finally, the finite element analysis
sought to analyze the stress distribution and displacement response of
the BTPS under load, thereby identifying areas for optimization.
CFD of BTPS
CFD simulations were performed to investigate the fluid dynamics
characteristics of the BTPS. Simulations were conducted using Ansys
software, employing the Navier–Stokes Equation ([144]1) to model
incompressible fluid flow. Water was selected as the working fluid,
characterized by a density of 1000 kg m^− ^3 and a viscosity of 1.002 ×
10⁻^3 Pa·s.^[ [145]^36 ^] Key fluid dynamic parameters, including shear
force, pressure distribution, and flow velocity, were analyzed
throughout the BTPS. The Reynolds number (Re), calculated using
Equation ([146]2), was employed to distinguish between laminar and
turbulent flow states.^[ [147]^37 ^] Permeability (K), as calculated by
Equation ([148]3), was employed to assess mass transfer capabilities
across distinct porous configurations. To minimize boundary effects, an
additional fluid domain was established above the BTPS model, with an
inlet flow rate set at 1 mm s^−1 and an outlet pressure maintained at
zero. A no‐slip boundary condition was imposed on the BTPS walls to
accurately model realistic flow behavior. The pressure difference (ΔP)
was computed following Equation ([149]4). This simulation approach
accurately captures the fluid dynamics environment within the BTPS,
thereby facilitating design optimization.
[MATH: ρ∂u∂t+u·∇u=−∇p+μ∇2
u+F∇·u=0 :MATH]
(1)
where u is the velocity of the fluid (m s^−1); ρ is the density of the
fluid (kg m^−3); t is the time (s); p is the pressure (Pa); µ is the
dynamic viscosity coefficient of the fluid (Pa s); ∇ is the operator; F
is the acting force (N).
[MATH: Re=ρvL<
/mrow>μ :MATH]
(2)
[MATH: K=v·μ<
mo>·LΔP
:MATH]
(3)
[MATH: ΔP=Pinlet−Poutlet :MATH]
(4)
where Re is the Reynolds number; 𝑣 is the flow velocity(m s^−1); L is
the characteristic length (mm); K is the permeability coefficient
(mm^2); ΔP is the pressure difference (MPa); P [inlet] is the pressure
at the inlet (MPa); P [outlet] is the pressure at the outlet (MPa).
Mechanical Analysis of the BTPS
The mechanical properties of the BTPS were rigorously assessed through
a static axial compression test conducted on a universal testing
machine (WDW‐100Y, Changzhou Geasure Medical Apparatus and Instruments
Co., Ltd). During testing, each sample was securely positioned in a
metallic holder. The pressure was then applied through a hollow push
rod connected by a universal joint, with a loading rate set at
1 mm min^−1, until the sample exhibited either deformation or fracture,
indicating structural failure. For each experiment, a minimum of three
samples were tested, with load‐displacement and stress‐strain curves
recorded to derive the yield load, elastic modulus, and comprehensive
mechanical profile of the BTPS.
Extraction and Characterization of HUVECs‐Derived Exosomes—Extraction of
Exosomes
HUVECs were initially cultured in Dulbecco's Modified Eagle's Medium
(DMEM, Gibco) supplemented with 10% fetal bovine serum under both
normoxic and hypoxic conditions. Upon reaching ≈80% confluence, the
medium was replaced with exosome‐free DMEM and the culture was
continued for another 48 h. Following incubation, exosome‐rich
supernatants were separately collected from both normoxic and hypoxic
cultures. Subsequently, exosomes were isolated through a multi‐step
centrifugation process as previously established.^[ [150]^38 ^]
Specifically, centrifugation steps at 300 g, 2000 g, 10 000 g, and
100 000 g were performed to successively eliminate live/dead cells,
debris, and larger particles. Then, exosomes were resuspended in
phosphate‐buffered saline(PBS, Gibco). Finally, exosomes derived from
normoxic and hypoxic HUVEC cultures were designated as normoxia
exosomes (Exo) and hypoxia exosomes (Hypo‐Exo), respectively.
Extraction and Characterization of HUVECs‐Derived Exosomes—BCA Protein
Quantification
Each exosome sample (20 µL) was combined with BCA (Thermo Scientific)
working solution and incubated at 37 °C for 30 min. Following
incubation, the absorbance at 562 nm was recorded using a microplate
reader. Using a standard curve, the protein concentration was
calculated, and all samples were adjusted to equivalent protein
concentrations for consistent downstream analyses.
Extraction and Characterization of HUVECs‐Derived Exosomes—Exosome Marker
Identification via Western Blot
Adhering to the Minimal Information for Studies of Extracellular
Vesicles (MISEV2018) standards, exosome‐specific markers were
identified by Western blotting. Each 100 µL exosome sample was combined
with 20 µL loading buffer, heated to 95 °C for 5 min, and subsequently
subjected to SDS‐PAGE. After electrophoresis, proteins were transferred
to a PVDF membrane, which was blocked in 5% PBS‐based skim milk for 1 h
at room temperature. The membrane was then incubated with primary
antibodies (anti‐CD81(ab109201, Abcam), anti‐CD9(ab223052, Abcam),
anti‐ALIX(ab117600, Abcam), anti‐β‐actin(ab8227, Abcam)) diluted
1:10 000 at 4 °C overnight, followed by a 1 h incubation with a 1:5000
horseradish peroxidase‐conjugated secondary antibody (ab205718, Abcam)
at room temperature. To visualize, an ECL reagent was applied, and
chemiluminescent signals were imaged after a 3 min exposure.
Extraction and Characterization of HUVECs‐Derived Exosomes—NTA
A 1 mL volume of each diluted exosome sample was injected into the
Nanosight nanoparticle analyzer, and the temperature probe was
positioned accordingly. Once focused, the particle size distribution
and concentration were determined at a wavelength of 405 nm, with data
subsequently recorded for analysis.
Extraction and Characterization of HUVECs‐Derived Exosomes—TEM
Characterization
To examine the morphology of exosomes, TEM imaging was conducted on a
Hitachi H‐7650 microscope set at 80 kV.
Uptake of Exos by HUVECs—Exosome Labeling
For fluorescence labeling, Exos and Hypo‐Exos were treated with PKH26
(Sigma–Aldrich). Specifically, 6 µL of PKH26 dye was added to 1 mL of
diluted exosome solution, mixed gently, and incubated at room
temperature for 5 min. Next, 2 mL of 10% BSA quenching solution was
added, and the total volume was brought to 8.5 mL with serum‐free
medium. The mixture was then transferred to an ultracentrifuge tube and
subjected to centrifugation at 190 000 g for 2 h at 4 °C. Afterward,
PKH26‐labeled exosomes were carefully resuspended in Dulbecco's
phosphate‐buffered saline (DPBS) under dark conditions, quantified
using a BCA kit, and stored at −80 °C.
Uptake of Exos by HUVECs—Exosome Uptake
To evaluate exosome uptake by HUVECs, 200 µg mL^−1 of PKH26‐labeled
exosomes were added to the HUVEC medium and incubated separately for
24 h. After each incubation period, cells were washed with DPBS, fixed
in 4% paraformaldehyde (PFA) for immunostaining, and subsequently
stained with 4′,6‐diamidino‐2‐phenylindole (DAPI, Invitrogen) for
nuclear visualization. For each group, three representative samples
were imaged using an LSM880 confocal laser scanning microscope (Zeiss),
and ImageJ software (National Institutes of Health, USA) was employed
to calculate the relative fluorescence intensity.
Hypo‐Exos Concentration Screening
To determine the optimal Hypo‐Exos concentration for angiogenesis
enhancement, angiogenesis experiments were conducted. Hypo‐Exos were
diluted to final concentrations of 50, 100, 150, and 200 µg mL^−1.
HUVECs were seeded into 96‐well plates pre‐coated with Matrigel and
treated with Hypo‐Exos at each specified concentration. Following
incubation at 37 °C for 6–8 h, the formation of vascular‐like
structures was observed and photographed. Quantitative analysis of
angiogenesis was conducted by calculating the total mesh area in images
using ImageJ software, enabling comparison across Hypo‐Exos
concentrations.
Preparation of PGHExo Microfluidic Devices
To prepare PGHExo double‐network hydrogel sustained‐release
microspheres, microfluidic chip technology was employed. Initially, the
primary channel of the microfluidic device was loaded with a prepolymer
solution composed of 1% PEGDA (Sigma–Aldrich), 1% GelMA
(Sigma–Aldrich), 1‐3 mg mL^−1 Hypo‐Exos, and 0.5% w/v Lithium
phenyl‐2,4,6‐trimethylbenzoylphosphinate (LAP, Sigma–Aldrich).
Concurrently, mineral oil (containing 10% span 80) functioned as the
shear phase. The two‐phase flow was carefully regulated at respective
flow rates of 200 and 1000 µL h^−1 to ensure uniform microsphere
formation. Subsequently, microspheres underwent crosslinking and
solidification within the microfluidic device via 365 nm UV exposure
for 15 s. Following solidification, microspheres were rinsed with
deionized water and transferred to a 1.5 mL Eppendorf tube.
Subsequently, centrifugation at 5000 rpm for 15 min was performed on
the microspheres, repeating this step three times. Finally, light
microscopy assessment was conducted, after which the microspheres were
resuspended in deionized water and stored at 4 °C.
Microstructure and Rheological Properties of PGHExo Microspheres—SEM
The PGHExo microspheres were freeze‐dried and subsequently coated to
enhance electrical conductivity. Next, the surface morphology of the
microspheres was examined under magnifications of 200x and 2000x.
Microsphere particle sizes were analyzed using ImageJ software,
generating a detailed particle size distribution map.
Microstructure and Rheological Properties of PGHExo Microspheres—Rheological
Properties
A rheometer (TA Instruments) was employed to quantify the storage
modulus (G′) and loss modulus (G″), thereby characterizing the PEGDA/
GelMA gel's mechanical properties. Testing frequencies were varied from
0.1 to 10 rad s^−1 while maintaining a constant temperature of 37 °C.
Under these controlled conditions, both GelMA (1%) and PEGDA/ GelMA
(1%) samples were analyzed to ascertain their frequency‐dependent
rheological properties.
Release Characteristics of PGHExo Microspheres—BCA Analysis
The release profile of Hypo‐Exos from PGHExo microspheres was evaluated
in a simulated body fluid (SBF, Leagene, CZ0403) solution. Each
microsphere group was immersed in 1 mL of degradation solution and then
continuously agitated at 37 °C. Protein concentration was measured
every three days up to day 18 by mixing 20 µL of solution with an
equivalent volume of degradation solution. At each measurement
interval, 20 µL of fresh degradation solution was added. BCA assays
were conducted on days 0, 3, 6, 9, 12, 15, and 18. To maintain protein
concentration consistency after each measurement, an equal volume of
BSA solution was added to the SBF. Additionally, exosome release
concentrations were quantified. Protein concentrations were determined
using a BCA kit, with each group measured in triplicate. The release
rate was calculated as follows (Equation [151]5):
[MATH: Releaserate%=Ct−C0/Cl×100 :MATH]
(5)
where C[0] denotes the protein concentration of PG microspheres without
exosomes, C[t] denotes the measured concentration for each group, and
C[l] represents the initial exosome loading concentration.
Additionally, excluding day 0, the protein concentration measured at
the previous time point was defined as C[t‐1]. The three‐day sustained
release concentration was then calculated using the following formula
(Equation [152]6):
[MATH: Proteinconcentrationmg/mL=Ct−Ct−1 :MATH]
(6)
Release Characteristics of PGHExo Microspheres—Fluorescence Analysis
To enable tracking of Hypo‐Exos release from the microspheres,
PKH26‐labeled Hypo‐Exos (2 mg mL^−1) were introduced into the aqueous
phase. Consistent with this approach, PGHExo microspheres were prepared
using the same protocol. Subsequently, a specified quantity of PGHExo
microspheres was introduced into 2 mL of SBF, allowing for individual
monitoring of each microsphere. Fluorescence signals were then recorded
using an LSM880 confocal laser scanning microscope on days 0, 3, 6, 9,
12, 15, and 18. Images were analyzed with ZEN software to quantify both
relative fluorescence intensity and microsphere volume.
Modification of BTPS with Embedded PGHExo Microspheres and Characterization
A pDA (Sigma–Aldrich) solution, prepared at pH 8.5 with a concentration
of 2 mg mL^−1 in 300 mL of 10 mm Tris‐HCl (Sigma–Aldrich), was
synthesized. Subsequently, the 3D‐printed BTPS was immersed in the pDA
solution and agitated overnight in a light‐protected environment.
Afterward, the BTPS was removed, rinsed thrice with deionized water,
and dried at 37 °C for subsequent application. BTPS was then positioned
in a well plate, with 1 mL of PGHExo microspheres resuspended in 500 µL
of PBS. The BTPS surface was sonicated and maintained in a sterile
environment for 24 h, enabling the PGHExo microspheres to embed onto
the pDA film, thus forming the BTPS&pDA@PGHExo composite. The
morphology and elemental composition of BTPS&pDA@PGHExo were
comprehensively characterized using SEM to assess its structural and
compositional properties.
Cell Culture
This study employed the MC3T3‐E1 mouse cranial osteoblast cell line and
HUVECsq, both sourced from the Chinese Academy of Sciences Cell Bank.
For optimal growth, the culture medium for each cell type was
supplemented with 10% fetal bovine serum, 100 µg mL^−1 streptomycin,
and 100 U mL^−1 penicillin. All procedures were conducted aseptically
within a biosafety cabinet to maintain sterile conditions.
Additionally, all reagents, including DPBS, culture medium, and 0.2%
trypsin, were preheated and prepared in advance. Following this, cells
in the logarithmic growth phase were washed twice with DPBS, digested
using 0.2% trypsin and neutralized by adding a complete medium. The
resulting cell suspension was then transferred to a 15 mL centrifuge
tube, centrifuged at 800 rpm for 5 min, after which the supernatant was
discarded. Subsequently, the cells were resuspended, counted, and
adjusted to the target concentration. To prepare for cell seeding,
pre‐sterilized scaffolds were placed in a 24‐well culture plate and
incubated overnight at 37 °C in a 5% CO₂ atmosphere. Finally, MC3T3‐E1
cells were seeded onto the scaffolds at a density of 5 × 10⁴ cells per
scaffold, setting the stage for subsequent experimental procedures.
Cell Viability Assay
To evaluate cell viability, MC3T3‐E1 cells were assessed using a
Live/Dead cell viability kit (Invitrogen, USA), allowing precise
determination of live‐to‐dead cell ratios across groups. Cells were
seeded onto BTPS, BTPS&pDA, BTPS&pDA@PG, and BTPS&pDA@PGHExo scaffolds
and cultured for 24 h. The scaffolds were then carefully rinsed with
DPBS two or three times to remove any residual medium. The staining
solution, containing Calcein AM and Ethidium homodimer‐1 diluted in
DPBS at a 1:1 to 1:1000 ratio, was prepared in darkness to ensure
optimal staining efficacy. A total of 500 µL staining solution was
added to each well and incubated at room temperature for 30 min, after
which the solution was discarded and washed three times with DPBS. The
samples were subsequently visualized under a fluorescence microscope
within a live cell workstation, capturing and documenting the
distribution of live (green fluorescence) and dead (red fluorescence)
cells. Background noise was delineated using a dashed line to identify
noise areas, thereby minimizing the impact of background interference.
Statistical regions were manually defined, and cell viability was
subsequently quantified using ImageJ.
Cell Proliferation Assay
The Alamar Blue assay was employed to quantify MC3T3‐E1 cell
proliferation on each scaffold type. MC3T3‐E1 cells were seeded at a
density of 5 × 10⁴ cells per scaffold across BTPS, BTPS&pDA,
BTPS&pDA@PG, and BTPS&pDA@PGHExo scaffolds, with proliferation measured
at 1, 3, and 5 days. Subsequently, the samples were washed with DPBS
and transferred to new 24‐well plates. At each designated time point, a
1:10 mixture of Alamar Blue solution (Invitrogen, USA) and complete
medium was introduced to each well. The samples were incubated at 37 °C
in darkness for 4 h. Following incubation, the supernatant was
carefully collected and transferred to a 96‐well plate. Finally, cell
proliferation was quantified using a fluorescence detection system
(Cytation5, BioTek, USA) with excitation and emission wavelengths of
530 and 590 nm, respectively.
Cell Osteogenic ALPAssay
The influence of different scaffolds on the osteogenic differentiation
of MC3T3‐E1 cells was systematically evaluated. After seeding cells
onto each scaffold and incubating them for 24 h, cultures were then
maintained in the osteogenic induction medium (high‐glucose DMEM with
10% FBS, 100 µg mL^−1 streptomycin, 100 IU mL^−1 penicillin, 0.1 µM
dexamethasone, 10 mm β‐glycerophosphate, and 50 µg mL^−1 sodium
ascorbate) for 7 days. The medium was refreshed every 2 to 3 days to
sustain optimal conditions for differentiation. On the seventh day,
supernatants from the cell lysates were collected and analyzed for ALP
activity using an ALP detection kit (Beyotime, China), thereby
quantifying the degree of osteogenic differentiation.
Alizarin Red S Mineralization Assay
To assess the mineralization capacity of osteoblasts, cells were
cultured under a standardized osteogenic induction protocol for 21
days. Following the incubation, the medium was discarded, and cells
were washed 2–3 times with DPBS before fixation in 4% paraformaldehyde.
Subsequently, cells were stained at room temperature with a 1% Alizarin
Red S solution (Sigma–Aldrich). After staining, the solution was
discarded, and cells were washed thoroughly with DPBS until no visible
dye residue was observed. The scaffolds were then dried at 37 °C to
document the morphology of mineralized nodules. Next, the scaffolds
were transferred to a 6‐well plate, and a 10% hexadecylpyridinium
chloride solution (Sigma–Aldrich) was added to elute the dye under
shaking conditions at 37 °C and 100 rpm. Finally, absorbance at 620 nm
was recorded using a microplate reader (Thermo Fisher, USA) to quantify
the formation and distribution of calcium‐mineralized nodules.
qRT‐PCR Analysis—Osteogenic Gene Expression
RT‐PCR was employed to quantify osteogenic gene expression. MC3T3‐E1
cells were seeded on each scaffold group and cultured for 7 days. Total
RNA was extracted using Trizol reagent (Beyotime, China) according to
the manufacturer's instructions, and its concentration and purity were
assessed with a NanoDrop spectrophotometer (Thermo Fisher Scientific,
USA). Reverse transcription of RNA into complementary DNA (cDNA) was
performed using a commercially available kit (Beyotime, China).
Quantitative PCR was conducted using a SYBR Green PCR kit (Thermo
Fisher Scientific, USA) on an RT‐PCR system (Beyotime, China). Primers
were designed to target osteogenic markers Runx2, ALP, OCN, and COL1A1,
with GAPDH serving as the internal reference gene. The primer sequences
are listed in Table [153]S1 (Supporting Information). The thermal
cycling conditions were set as follows: initial denaturation at 95 °C
for 5 min, followed by 40 cycles of 95 °C for 30 s, 60 °C for 30 s, and
72 °C for 30 s. Relative gene expression levels were calculated using
the 2^(‐ΔΔCt) method and statistically analyzed to determine the
effects of scaffolds on osteoblast‐related gene expression.
qRT‐PCR Analysis—Angiogenic Gene Expression
Angiogenic gene expression was assessed using qRT‐PCR to evaluate the
effects of the scaffolds on vascularization. HUVECs were seeded onto
each scaffold group and cultured for 3 days. The expression levels of
VEGF, CD31, PDGFB, and FGF2 were quantified by RT‐PCR, with the primer
sequences provided in Table [154]S1 (Supporting Information). The
experimental procedures and data analysis were performed following the
protocol described for osteogenic gene expression, using GAPDH as the
internal reference gene.
mRNA‐seq Analyses
The MC3T3‐E1 cell line was utilized to investigate the underlying
mechanisms. Cells were cultured on scaffolds under standard conditions
until the predefined incubation period was completed. Total RNA was
extracted using a TRIzol reagent following the manufacturer's protocol.
The quality and purity of RNA were assessed with a NanoDrop
spectrophotometer and validated via agarose gel electrophoresis to
ensure suitability for library preparation. High‐quality RNA was
reverse‐transcribed into cDNA, and cDNA libraries were prepared for
high‐throughput sequencing on the Illumina platform (Zhejiang, China).
The sequencing data underwent rigorous quality control, alignment to
the reference genome, gene expression quantification, and
identification of differentially expressed genes. GO and KEGG
enrichment analyses were performed on key genes to uncover essential
osteogenesis‐related pathways, offering insights into the scaffold's
role in regulating bone formation. Furthermore, the expression levels
of bone‐related genes (ALPL, COL18A1, SAMD6, RUNX2OS1) and angiogenic
genes (VEGFB, PDGFA, ANGPT2) were analyzed following the experimental
protocol described under the subhead “qRT‐PCR Analysis” to validate the
differentially expressed genes (DEGs) identified in the mRNA‐seq
analysis. The primer sequences are provided in Table [155]S1
(Supporting Information).
ELISA
The concentrations of osteogenic markers (RUNX2, OCN) and angiogenic
markers (PDGF, VEGF) in the cell supernatant were quantified using
species‐specific ELISA kits (ELK Biotechnology) according to the
manufacturer's instructions. Briefly, centrifuged serum samples (3000 ×
g, 15 min) from each treatment group were incubated in
antibody‐precoated wells, followed by incubation with the corresponding
detection antibodies. The absorbance was measured at 450 nm using a
microplate reader, and concentrations were determined based on standard
curves. To ensure data accuracy and reproducibility, all measurements
were performed in technical triplicates.
Animals and Surgical Procedures
All animal care and experimental procedures were performed in strict
accordance with institutional guidelines, as well as national laws and
regulations. The animal experimental protocol was reviewed and approved
by the Animal Ethics Committee of Southern Medical University
(SMUL2023091), and all animal experiments complied fully with the
committee's ethical requirements. Healthy adult New Zealand white
rabbits (2.5–3.0 kg) were acclimated for one week and then randomly
assigned to one of four groups (n = 6 per group, totaling 24): BTPS,
BTPS&pDA, BTPS&pDA@PG, and BTPS&pDA@PGHExo. Prior to the procedure,
general anesthesia was induced with 3% sodium pentobarbital at a dose
of 1 mL kg^−1. Once anesthesia was confirmed, each rabbit was secured
on the operating table, and the hindlimbs were shaved and disinfected
according to standard protocols. A longitudinal incision above the knee
joint was made to expose the distal femur, ensuring aseptic techniques
throughout. A bone defect model with a diameter and depth of 10 mm was
drilled into the distal femur, carefully avoiding adjacent soft tissues
and vital structures. Following thorough saline irrigation of the
defect, the corresponding scaffold was implanted based on the assigned
grouping. The muscles, subcutaneous tissue, and skin were sutured in
sequential layers. To prevent infection, the wound was irrigated with
saline and an antibiotic solution, supplemented with routine
postoperative care, including antibiotic injections and wound
management. Postoperative monitoring of behavior and wound healing was
conducted to ensure no signs of infection or complications.
Micro‐CTAnalysis
Micro‐CT imaging was performed on the implantation sites at 4 and 12
weeks post‐surgery using a Bruker 1276 Micro‐CT scanner to evaluate the
repair progress of femoral bone defects in rabbits. Following
euthanasia, femoral bone specimens containing implants were carefully
extracted, rinsed thoroughly with saline, and fixed in 10% neutral
formalin for 24 h to preserve tissue structure. The specimens were
subsequently rinsed to eliminate any residual fixative solution before
undergoing Micro‐CT scanning. Scanning parameters were set at 70 kV for
voltage, 200 µA for current, with a resolution of 9 µm and an exposure
time of 200 ms, encompassing the bone defect and adjacent normal bone
tissue. The acquired scan data were processed using 3D reconstruction
software to obtain high‐resolution images of the bone tissue.
Subsequently, image analysis software was applied to quantify new bone
volume and density, providing a comprehensive assessment of bone repair
efficacy.
Histopathological Analysis
To assess histopathological features of bone defect repair in the
rabbit distal femur, samples from the implant site were collected at 4
and 12 weeks post‐surgery. Samples were rinsed in saline and
immediately fixed in 10% neutral formalin for 48 h to ensure tissue
structure preservation. Fixed samples were subsequently dehydrated in
70%, 80%, 90%, 95%, and finally 100% ethanol (each concentration
applied twice) and embedded in polymethyl methacrylate (PMMA,
Cool‐Set‐A, OLYMPUS, Chengdu). Tissue sections, 10–20 µm thick, were
prepared using a diamond microtome (SAT‐001, Olinkem, Chengdu) and
subsequently stained with methylene blue and basic fuchsin (Sigma) to
examine histological structures and cellular morphology. Stained
sections were then observed and imaged using an optical microscope,
with a primary focus on new bone and blood vessel formation. Image
analysis, conducted via ImageJ software, quantitatively assessed new
bone and blood vessel distribution, quantity, and morphology to
comprehensively characterize bone repair outcomes.
Statistical Analysis
Experimental data were reported as the Mean ± SD, with each group
consisting of at least three independent samples. All statistical
analyses were conducted using GraphPad Prism 9 (version 10.0).
Differences between two groups were analyzed using an independent
Student's t‐test, while comparisons among more than two groups were
performed using one‐way ANOVA. Tukey's multiple comparison test was
used for further differentiation after ANOVA. Differences between
groups were considered statistically significant at, ^* p < 0.05; ^** p
< 0.01; ^*** p < 0.001; and ^**** p < 0.0001; “ns” indicated not
significant.
Conflict of Interest
The authors declare no conflict of interest.
Supporting information
Supporting Information
[156]ADVS-12-2500599-s001.docx^ (981.9KB, docx)
Acknowledgements