Abstract
The treatment of infected bone defects (IBDs) needs simultaneous
elimination of infection and acceleration of bone regeneration. One
mechanism that hinders the regeneration of IBDs is the iron competition
between pathogens and host cells, leading to an iron deficient
microenvironment that impairs the innate immune responses. In this
work, an in situ modification strategy is proposed for printing
iron‐active multifunctional scaffolds with iron homeostasis regulation
ability for treating IBDs. As a proof‐of‐concept, ultralong
hydroxyapatite (HA) nanowires are modified through in situ growth of a
layer of iron gallate (FeGA) followed by incorporation in the
poly(lactic‐co‐glycolic acid) (PLGA) matrix to print biomimetic PLGA
based composite scaffolds containing FeGA modified HA nanowires
(FeGA‐HA@PLGA). The photothermal effect of FeGA endows the scaffolds
with excellent antibacterial activity. The released iron ions from the
FeGA‐HA@PLGA help restore the iron homeostasis microenvironment,
thereby promoting anti‐inflammatory, angiogenesis and osteogenic
differentiation. The transcriptomic analysis shows that FeGA‐HA@PLGA
scaffolds exert anti‐inflammatory and pro‐osteogenic differentiation by
activating NF‐κB, MAPK and PI3K‐AKT signaling pathways. Animal
experiments confirm the excellent bone repair performance of
FeGA‐HA@PLGA scaffolds for IBDs, suggesting the promising prospect of
iron homeostasis regulation therapy in future clinical applications.
Keywords: 3D printing, antibacterial activity, bone repair, in situ
growth, iron homeostasis
__________________________________________________________________
Here, an in situ modification strategy is proposed for printing
iron‐active multifunctional scaffolds with iron homeostasis regulation
ability for treating infected bone defects (IBDs). The FeGA‐HA@PLGA
scaffold combats bacterial infection through photothermal effects in
the early stage of bone infection, then helps restore the iron
homeostasis microenvironment, thereby promoting anti‐inflammatory,
vascularization and osteogenic differentiation.
graphic file with name ADVS-11-2407251-g004.jpg
1. Introduction
Infected bone defects (IBDs) is a devastating complication in
orthopedics and maxillofacial surgery, which usually causes persistent
inflammation, delayed bone healing, and sometimes amputation or even
death.^[ [52]^1 ^] To treat IBDs, various antibacterial agents
including antibiotics, noble metal ions, and black materials that
convert light to heat have been introduced to the bone scaffolds to
eliminate infection.^[ [53]^2 ^] However, these antibacterial agents
often cause long‐term safety concerns due to their toxicity at high
doses and/or poor biodegradability.^[ [54]^3 ^] Furthermore, these
antibacterial agents are inert in bioactivity, which potentially
impairs the bone repair effects of the scaffolds. Therefore,
multifunctional bone scaffolds with biodegradable antibacterial agents
with bioactive functions such as the reconstruction of the osteogenic
microenvironment are urgently required but remain challenging.
After bone injury, a profound transformation takes place within the
local microenvironment, characterized by elevated levels of reactive
oxygen species (ROS) and an acute inflammatory response. These
phenomena significantly hinder the efficient healing process of bone
defects.^[ [55]^4 ^] Metal‐phenolic networks (MPNs), as novel compound
materials, capitalize on the intricate coordination chemistry between
phenolic ligands and metal ions to exhibit robust anti‐inflammatory,
antioxidant, and antibacterial properties. This represents a crucial
advancement in the realm of material science.^[ [56]^5 ^] By
synergistically combining the distinct functionalities of metallic ions
and phenolic ligands, MPNs offer tailored benefits that are ideally
suited to meet the specific requirements of orthopedic biomaterial
applications.^[ [57]^6 ^] MPNs composed of bioactive metal ions (e.g.,
Mg^2+,^[ [58]^7 ^] Fe^3+,^[ [59]^8 ^] Sr^2+,^[ [60]^9 ^] etc.)^[
[61]^10 ^] and biocompatible organic small molecule ligands (e.g.,
polyphenols)^[ [62]^11 ^] can be easily degraded under physiological
environment due to their weak binding forces (i.e., coordination
bonds).^[ [63]^12 ^] Their degradation products often provide various
biological activities such as pro‐angiogenesis^[ [64]^13 ^] and free
radical scavenging.^[ [65]^14 ^] Therefore, MPNs are suitable agents
for designing functional scaffolds.^[ [66]^15 ^] Among various
bioactive metal ions, iron‐based MPNs have dual functions of
photothermal antibacterial activity^[ [67]^16 ^] and iron
supplementation.^[ [68]^17 ^] Iron is in a delicate balanced state in
the body, and it is the only hormone‐regulating trace element that
responds to both the host's nutritional status and infected status.^[
[69]^18 ^] At the site of infection, bacteria can compete with host
cells for iron to maintain their survival and proliferation,^[ [70]^19
^] which leads to an imbalance in tissue iron homeostasis, thereby
inhibiting the maturation of neutrophils and their immune defense
capabilities.^[ [71]^18 , [72]^20 ^] One important factor for treating
IBDs is iron supplementation which can reverse the iron‐deficient
microenvironment to restart the immune defense function of cells in the
innate immune system.^[ [73]^18 , [74]^21 ^] In addition, the iron ions
can promote the transformation of proinflammatory macrophages (M1) to
anti‐inflammatory macrophages (M2), ^[ [75]^22 ^] which promotes the
regeneration process.^[ [76]^23 , [77]^24 ^] Therefore, iron‐based
metal organic complex modified scaffolds are expected to have favorable
treatment performance for IBDs, yet are rarely studied.
The porous architecture of the scaffolds is an appropriate
microenvironment that is conducive to cell adhesion, proliferation,
differentiation, and biomineralization.^[ [78]^25 ^] The hydroxyapatite
ultralong nanowires (HA) reported in our previous work have high
biocompatibility, biological activity and thermal stability,^[ [79]^26
^] which stands as a preferred material for bone repair applications.^[
[80]^27 ^] However, due to the intrinsic brittleness of hydroxyapatite
materials, it is not suitable as a base component for 3D printing.^[
[81]^28 ^] To improve its printability, HA is usually mixed in the
polymeric materials such as poly(lactic‐co‐glycolic acid) (PLGA) for
room‐temperature 3D printing that can largely protect the scaffold from
thermal degradation.^[ [82]^29 ^]
In this work, an in situ growth strategy is proposed to prepare iron
gallate (FeGA) metal organic complex modified 3D printing iron‐active
multifunctional bone scaffolds with antibacterial activity,
anti‐inflammatory and iron homeostasis regulation abilities using HA
and PLGA as matrices (Scheme [83]1). The in situ growth strategy uses
the minimum amount of FeGA to turn white HA nanowires into black,
thereby endowing them with excellent photothermal properties while
avoiding potential toxicity. The molecular dynamics (MD) simulation is
used to elucidate the layered structure of FeGA on the surface of HA
nanowires. The as‐obtained multifunctional scaffolds comprising FeGA
modified HA nanowires (FeGA‐HA@PLGA) have designed micro‐scale porous
structure, which facilitates cell adhesion, migration and growth, and
sustained release of bioactive iron ions and GA molecules upon its
degradation. These released iron ions can reverse the iron deficient
microenvironment to reactivate the innate immune defense system in the
early stages of infected bone repair. The photothermal effect endows
the FeGA‐HA@PLGA scaffolds with an excellent antibacterial performance
by enhancing the permeability of bacterial membranes through effective
thermal simulation. The released GA molecules can scavenge ROS while
the released iron ions can promote macrophages to transform to M2
phenotype. The biological activities of GA and iron ions help to
transform the inflammatory microenvironment into a microenvironment
conducive to tissue regeneration. By synergizing antibacterial and
anti‐inflammatory activity, immune system reactivation and
pro‐osteogenic differentiation properties, our FeGA‐HA@PLGA scaffolds
are expected to have favorable therapeutic efficacy for IBDs.
Scheme 1.
Scheme 1
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Schematic illustration of preparation and mechanism of the FeGA‐HA@PLGA
scaffolds for treating IBDs.
2. Results and Discussion
2.1. The In Situ Growth FeGA‐HA Nanowires are Prepared and Characterized
The successful in situ growth of FeGA on the surface of HA nanowires is
confirmed by a transmission electron microscope (TEM, Figure [85]1a–c).
The smooth surface of HA nanowires becomes rough with increasing
reaction time. The process is accompanied by the color change of the
FeGA‐HA samples from white to grey and then to black (Figure [86]1a;
Figures [87]S1–S3, Supporting Information). The energy‐dispersive X‐ray
spectroscopy (EDS) mapping results show the uniform distribution of
iron elements, proving the uniform growth of FeGA on the HA surface
(Figure [88]1c). Compared to directly mixing FeGA nanoparticles with HA
mechanically, in situ growth of FeGA on the surface of HA nanowires has
several advantages including (i) ensuring uniform sustained release of
iron ions and GA molecules; (ii) achieving high photothermal property
at a low FeGA amount; (iii) avoiding potential toxicity caused by FeGA
at a high dose. The relationship between Fe^3+ content in the samples
and reaction time is investigated. The Fe^3+ amount in the samples
prepared at 3, 6 and 12 h is determined to be 2.36, 6.05 and
34.67 mg k^−1g by inductively coupled plasma analysis (ICP,
Figure [89]1f). The increased Fe^3+ content with reaction time results
in the color change from white to black (Figure [90]S2, Supporting
Information). The powder X‐ray diffraction (XRD) and Fourier‐transform
infrared spectroscopy (FTIR) results (Figure [91]1d,e) confirm the
existence of HA (JCPDS 09–0432). The lack of differences observed in
the XRD and FTIR results of HA and FeGA may be due to the thin and
amorphous FeGA layer, which only accounts for a small proportion of the
entire sample.
Figure 1.
Figure 1
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Characterization of the FeGA‐HA. a) Schematic illustration of the
formation mechanism of the FeGA‐HA samples with increasing reaction
time. b) TEM images of FeGA‐HA samples prepared under different
reaction times: (1)–(3) Scale bar: 50 nm, (4) Scale bar: 20 nm. c) TEM
image and corresponding elemental mapping images of FeGA‐HA‐12 h, Scale
bar: 50 nm. d) XRD patterns of HA and FeGA‐HA. e) FTIR curves of HA and
FeGA‐HA. f) The content of Fe^3+ in the FeGA‐HA measured by ICP. g) The
MD simulation results show the adsorption structure of FeGA on the HA
surface. h) The relationship between the density of GA and Fe^3+ in the
system and their distance from the surface of HA. i) Radial
distribution of Fe^3+ and GA in the system before and after MD
simulation.
To investigate the binding mold between FeGA and HA, the MD simulation
is conducted using a system containing Fe^3+, GA, glycerol and HA
crystals with an exposed [001] crystal plane (Figure [93]1g). The
simulation results show that a layer of FeGA with an amorphous
structure is formed on the surface of [001] crystal plane of HA. The
thickness of the FeGA layer is about 2 nm (Figure [94]1h). The radial
distribution function results show that a distinct g(r) peak appears
after the aggregation reaction of Fe^3+ and GA on the surface of HA for
50 ns (Figure [95]1i). The conformation analysis shows that the Fe^3+
ions are initially adsorbed onto the HA surface followed by
coordinating with carboxyl groups of GA to allow the molecular growth
on the crystal plane (Figure [96]1g). Simultaneously, the conjugation
between GA molecules and HA through the benzene rings promotes their
continuous stacking, also facilitating the growth of FeGA on the
crystal surface (Figures [97]S5,S6, Supporting Information). These
results preliminarily confirm that the HA surface exhibits a certain
adsorption capacity for GA and Fe^3+ ions, allowing them to aggregate
and grow on the HA surface.
To further elucidate the interactions among all the molecules in the
system, the binding energy of Fe^3+, GA and glycerol to the HA surface
is calculated (Figure [98]S7, Supporting Information). The results show
that Fe^3+ ions have the strongest interaction with the HA surface with
a binding energy of −397369.2±244.5 kJ mol^−1. In contrast, the binding
energy between GA and HA (i.e., −70455.9±285.5 kJ mol^−1) is much
smaller than that between Fe^3+ and HA. The negative value of binding
energy indicates the spontaneous binding reaction between HA and GA or
Fe^3+. In contrast, the binding energy between glycerol and HA is
calculated to be positive (i.e., 227.5±158.8 kJ mol^−1), indicating
that there is no adsorption between glycerol and HA. The negative
binding energy between Fe^3+ and GA (i.e., −181470.7±541.2 kJ mol^−1)
indicates that the FeGA is formed during the reaction process.
2.2. Fabrication and Characterization of FeGA‐HA@PLGA Scaffolds
3D printing technology is used to fabricate the FeGA‐HA@PLGA scaffolds
with a designed porous structure using PLGA as a matrix. The PLGA with
relatively fast biodegradation characteristics is beneficial for the
exposure of FeGA‐HA and the Fe^3+ release from it. The optical images
of PLGA, HA@PLGA, and FeGA‐HA@PLGA 3D printing scaffolds are shown in
Figure [99]2a. The favorable printability of PLGA is not affected by
the additives of HA or FeGA‐HA while the scaffold color is changed to
white or black by adding HA or FeGA‐HA. The scanning electron
microscope (SEM) images show that the surface of the scaffold becomes a
little rough upon adding FeGA‐HA (Figure [100]2b). The water contact
angle results show that the hydrophilicity of the FeGA‐HA@PLGA
scaffolds is enhanced compared to the PLGA scaffold, which is conducive
to cell adhesion and proliferation (Figure [101]S8, Supporting
Information). EDS mapping results show the uniform distribution of Fe
elements, indicating that the FeGA‐HA nanowires are uniformly
incorporated in the PLGA scaffold (Figure [102]2c). There are no
characteristic diffraction peaks of FeGA that are detected due to its
lower loading content and weak crystallization. This observation also
suggests the effective dispersion of minute FeGA clusters on the HA
(JCPDS 09–0432) surface. In addition, XRD patterns show that the
FeGA‐HA addition can induce the crystallization of PLGA, which helps
enhance the mechanical strength of the scaffolds (Figure [103]S9,
Supporting Information). FTIR measurements show that the typical
vibrational peaks of phosphate (i.e., 1750 cm^−1) are observed in both
FeGA‐HA and FeGA‐HA@PLGA samples, also confirming the successful
incorporation of FeGA‐HA in the PLGA scaffolds (Figure [104]S10).
Figure 2.
Figure 2
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Characterization of FeGA‐HA@PLGA. a) Schematic illustration and digital
photos of PLGA, HA@PLGA and FeGA‐HA@PLGA scaffolds. b) SEM images of
different scaffolds of PLGA (1), HA@PLGA (2) and FeGA‐HA@PLGA (3‐4);
(1–3) Scale bar: 300 µm. (4) Scale bar: 10 µm. c) EDS analysis of C, P,
Ca, and Fe elements in the FeGA‐HA@PLGA scaffolds, Scale bar: 300 µm.
d) The cumulative release of Fe^3+ from the FeGA‐HA@PLGA scaffolds. e)
The thermal gravimetric analysis of HA, FeGA‐HA, PLGA, and
FeGA‐HA@PLGA. f) Compressive stress−strain curves of PLGA, FeGA‐HA, and
FeGA‐HA@PLGA scaffolds.
Iron is the fifth most abundant metal element in healthy adult bones
(i.e., ≈100 ppm, less than calcium, magnesium, zinc, and strontium).^[
[106]^30 ^] Iron maintains bone health from multiple aspects, including
participating in the metabolic process of bones, affecting calcium
absorption, maintaining bone density, and ensuring bone strength.^[
[107]^31 ^] Iron deficiency may lead to issues such as poor bone
development, slow fracture healing, and decreased bone density.^[
[108]^32 ^] Therefore, iron supplementation is crucial for bone repair.
The amount of iron in the FeGA‐HA nanowires is determined to be ≈32 ppm
by inductively coupled plasma atomic emission spectroscopy (ICP‐AES).
The loading of FeGA‐HA nanowires in the FeGA‐HA@PLGA scaffolds is
determined to be ≈14.4% by thermogravimetric analysis (TGA), based on
which iron content in the scaffolds can be estimated to be ≈4.62 ppm
(Figure [109]2e; Figure [110]S11, Supporting Information). To examine
the release kinetics of GA, the FeGA‐HA@PLGA scaffolds were immersed in
PBS, and the absorbance of the supernatant at 259 nm was measured
chronologically. The results show that ≈65% GA can be released from the
FeGA‐HA@PLGA scaffolds after 23 days in a simulated physiological
environment (Figures [111]S12 and [112]S13, Supporting Information).
Benefitting from the biodegradability of PLGA, ≈80% iron can be
released from the FeGA‐HA@PLGA scaffolds after 40 days in a simulated
physiological environment (Figure [113]2d). The released iron can exert
its biological effects to facilitate bone regeneration at the defect
site. Besides the bioactivity, bone implants need to have a certain
level of mechanical strength. Our previous study found that the
ultralong HA nanowires possess high strength and high flexibility.^[
[114]^27a ^] Therefore, the incorporation of HA nanowires into the PLGA
scaffolds is expected to enhance its mechanical properties. The
mechanical strength of PLGA, FeGA‐HA, and FeGA‐HA@PLGA scaffolds at 80%
deformation are measured to be 1.4, 3.2 and 6.1 MPa (Figure [115]2h).
The high strength of FeGA‐HA@PLGA scaffolds is attributed to the
crystalline structure of the PLGA matrix and high flexibility of loaded
HA nanowires. Therefore, the FeGA‐HA@PLGA scaffolds implanted into the
bone defect site can provide certain mechanical support in the process
of bone tissue repair.
2.3. The Photothermal Properties of FeGA‐HA and FeGA‐HA@PLGA
Photothermal heating ability that converts electromagnetic waves into
heat has been intensively used in nanomedicine.^[ [116]^14 , [117]^33
^] For treating IBDs, high‐temperature thermal stimulation can
effectively eliminate pathogenic bacteria, while mild thermal
stimulation can promote bone repair.^[ [118]^14a ^] Therefore, the
regulation of photothermal heating temperature of bone scaffold
materials is crucial for the treatment of IBDs. Our in situ
modification strategy allows one to easily tune the photothermal
performance of modified HA nanowires by controlling the modification
layer thickness (i.e., FeGA layer). Infrared thermographic images and
photothermal curves display that the FeGA‐HA‐12 h sample has the
fastest heating ability with the highest peak temperature compared to
FeGA‐HA‐3 h and FeGA‐HA‐6 h samples under NIR irradiation with a power
density of 0.25 W cm−^2, which is due to its highest iron content
(Figure [119]3a,b). The photothermal performances of the FeGA‐HA
samples under different power densities are investigated (Figure
[120]S14, Supporting Information). These results confirm that the
FeGA‐HA‐12 h sample can be rapidly heated to the antibacterial
hyperthermic temperature (i.e., 49−52 °C) under a low NIR irradiation
power density. Therefore, the FeGA‐HA‐12 h sample is selected to
prepare the FeGA‐HA@PLGA scaffolds for further experiments. The
photothermal performances of scaffolds with and without FeGA‐HA
nanowires are characterized under different NIR irradiation power
densities (Figure [121]3c). The results show that the FeGA‐HA@PLGA
scaffolds is heated up to 31.1, 38.4, 49.2, and 52.1 °C under NIR
irradiation for 10 min with corresponding power densities of 0.5, 1,
1.5 and 2 W cm^− ^2. The lowest irradiation power density (i.e.,
1.5 W cm^− ^2) that can raise the temperature of the scaffold to the
antibacterial temperature (i.e., 49.2 °C) is selected for subsequent
experiments (Figure [122]S15, Supporting Information). At this power
density, the temperature of the FeGA‐HA@PLGA scaffolds can reach above
45 °C after irradiation for only 3 min, whereas the temperatures of the
PLGA and HA@PLGA scaffolds remain below 30 °C under the same conditions
(Figure [123]3d). Furthermore, the FeGA‐HA@PLGA scaffolds has good
photothermal stability, which is proved by the cyclic heating curves
(Figure [124]3e), and the photothermal conversion efficiency of the
FeGA‐HA@PLGA is 45.9%.
Figure 3.
Figure 3
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Characterization of photothermal properties. a,b) Infrared
thermographic maps and photothermal heating curves of HA, FeGA‐HA‐3 h,
FeGA‐HA‐6 h, and FeGA‐HA‐12 h samples under NIR laser irradiation
(808 nm, 0.25 W cm^−2). c) Photothermal heating curves of FeGA‐HA@PLGA
scaffolds under NIR laser irradiations with different power densities.
d) Photothermal heating curves of PLGA, HA@PLGA and FeGA‐HA@PLGA
scaffolds under NIR laser irradiation (808 nm, 1.5 W cm^−2). e) Cyclic
photothermal heating/cooling curves of the FeGA‐HA@PLGA scaffolds under
NIR laser irradiation (808 nm, 1.5 W cm^−2).
2.4. Investigation of Antibacterial Properties with FeGA‐HA@PLGA Scaffolds
At the infected site, bacteria proliferate rapidly and compete for iron
ions from the host to ensure their survival. Therefore, it is essential
not only to provide iron to the host but also to swiftly control
bacterial proliferation for treating IBDs. The antibacterial activities
of our scaffolds are evaluated using both Gram‐positive and
Gram‐negative bacteria under an 808 nm NIR irradiation at 1.5 W cm^−
^2. The bacterial colonies counting results show that the FeGA‐HA@PLGA
NIR group has high bacterial inhibition ratios for both S. aureus
(99.5%) and E. coil (99.8%) compared to other groups (Figure
[126]4a,b). The morphologies of bacteria co‐cultured with different
scaffolds in supernate and on scaffolds are observed by SEM
(Figure [127]3c,d). The bacteria with complete membranes are observed
for PLGA, HA@PLGA and HA@PLGA NIR groups, indicating their low
antibacterial activities, which is consistent with the bacterial
colonies counting results. In contrast, the wrinkling of the bacterial
membrane is observed for the FeGA‐HA@PLGA group, indicating that our
FeGA‐HA@PLGA scaffolds has a certain antibacterial ability. After
applying NIR irradiation, wrinkling and rupture of bacterial membranes
are observed for both S. aureus and E. coil whether on the scaffolds or
in the suspensions, indicating the excellent photothermal antibacterial
activity of our FeGA‐HA@PLGA scaffolds. Our scaffolds possess a high
heating rate at low irradiation power to achieve a photothermal
antibacterial effect (47–50 °C),^[ [128]^34 ^] which avoids the serious
irreversible thermal damage to the surrounding tissue of the defect
site and adverse effects on the integration of bone implants caused by
the high‐power and long‐term photothermal therapy. Simultaneously, at
low irradiation power (47–50 °C) insufficient to kill bacteria, they
can increase the permeability of bacterial membranes, thereby
synergistically promoting the antibacterial action of antimicrobial
drugs or ions.^[ [129]^35 ^] Therefore, to investigate the
antibacterial mechanism of the FeGA‐HA@PLGA scaffolds, the bacterial
membrane permeability is further examined in different treatment
groups. The NPN fluorescence measurement is a typical method to
evaluate the membrane integrity, where the stronger the fluorescence
intensity, the worse the membrane integrity and the better the
permeability. The results show that both E.coli and S. aureus show the
highest fluorescence intensity in the FeGA‐HA@PLGA NIR group compared
to other groups, indicating that the highest permeability of the
bacterial membrane is achieved in the FeGA‐HA@PLGA NIR group
(Figure [130]4e). The results demonstrate that the FeGA‐HA@PLGA
scaffolds possess antibacterial properties, which are further enhanced
under photothermal conditions. This outcome indicates that photothermal
effects can increase the permeability of bacterial membranes and have a
synergistic antibacterial effect with FeGA. The antibacterial mechanism
could be because of the enhanced permeability of bacterial membranes
caused by the photothermal effect, which enhances the antibacterial
effect of FeGA‐HA@PLGA scaffolds (Figure [131]4f).
Figure 4.
Figure 4
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Antibacterial properties of the FeGA‐HA@PLGA scaffolds under 808 nm NIR
irradiation. a) Photographs of bacterial colonies of S. aureus and E.
coli treated by. b) The relative viability of bacteria treated with
different groups. c,d) SEM images of S. aureus and E. coli on scaffolds
or in supernate after treatment with different groups. e) Influences of
different scaffolds of bacterial membrane permeability. *p < 0.05, **p
< 0.01, ***p < 0.001, ****p < 0.0001. f) Schematic diagram of
antibacterial mechanism of the FeGA‐HA@PLGA scaffolds under NIR
irradiation.
2.5. Biocompatibility, Anti‐Inflammatory and Osteogenic Differentiation
Evaluation of FeGA‐HA@PLGA Scaffolds
A prominent advantage of 3D printing scaffolds is their designed porous
structure with adjustable porosity. Such a porous structure is
conducive to the growth of blood vessels and the adhesion and migration
of osteogenic related cells. The biocompatibility and cell adhesion
ability of our scaffolds are evaluated using bone marrow mesenchymal
stem cells (BMSCs). The Live/Dead staining images and OD450 statistical
analysis results indicate that all the different treatment groups have
high cell viability (Figure [133]5a,b). The laser confocal microscopy
results indicate that BMSCs exhibit a healthy growth status on the
scaffolds with unaffected cell morphology (Figure [134]5a; Figure
[135]S16, Supporting Information). Interestingly, BMSCs on the
FeGA‐HA@PLGA scaffolds with or without NIR irradiation demonstrate a
larger spreading area with more pseudopodia, which is an early signal
for enhanced cell proliferation and differentiation. In addition, the
cell adhesion density in the FeGA‐HA@PLGA scaffolds group is
significantly higher than that in the unmodified scaffold group
(Figure [136]5a). These results show that our FeGA‐HA@PLGA scaffolds
has high biocompatibility. The hemolysis experiments show that all the
scaffolds have good blood compatibility (Figure [137]5c).
Figure 5.
Figure 5
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The biocompatibility, anti‐inflammatory, and osteogenic differentiation
of different scaffolds in BMSCs. a) Live/Dead staining and cytoskeleton
staining images (Dapi/Actin, confocal microscopy) of BMSCs cultured
with different groups. Scale bar: 200 µm. b) Cytotoxicity of the
different scaffolds. c) Hemolysis of rat blood treated with different
scaffolds. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. d)
Schematic diagram of cell‐scaffold co‐culture models for Live/Dead and
Cytoskeleton staining. e) The fluorescence images show intracellular
ROS levels of BMSCs after being treated with different scaffolds. ROS
was stained by DCF‐DA. Scale bar: 200 µm. f) Flow cytometry results
showing CD206 and CD86 expression of macrophages with different
treatments and represented as flow histograms. g) Schematic diagram of
cell‐scaffold co‐culture models for ROS staining. h) ALP and ARS
staining images of BMCSs co‐cultured with different groups.
Bacterial infection exacerbates the inflammatory response at the bone
defect, leading to the formation of an intracellular high‐level
endogenous ROS microenvironment, which is the main cause of bone
necrosis in the site of IBDs.^[ [139]^36 ^] Therefore, it is required
to eliminate the excess ROS and its assault on immune cells to
transform the inflammatory microenvironment into one conducive to bone
regeneration. The antioxidant component (i.e., GA) in our scaffolds can
perform the task of clearing ROS. In addition, the binding of GA
molecules with iron ions enhances its stability, which can contribute
to the antioxidant performance of our FeGA‐HA@PLGA scaffolds. The ROS
clearance capability is assessed using the DCF‐DA fluorescence probe
(Figure [140]5d). H[2]O[2] stimulation alone serves as the positive
control and the untreated group serves as the negative control. The
fluorescence images show that the ROS fluorescence intensity in BMSCs
treated with the FeGA‐HA@PLGA+H[2]O[2] is significantly lower than that
in the PLGA+H[2]O[2] or HA@PLGA+H[2]O[2] groups, indicating that the
FeGA‐HA@PLGA scaffolds has outstanding ROS scavenging ability
(Figure [141]5e). In addition, the ROS scavenging performances of the
scaffold against other different free radicals (such as DPPH, ABTS,
PTIO) are also evaluated (Figures [142]S17–S19, Supporting
Information). All these results confirm the excellent antioxidant
properties of our FeGA‐HA@PLGA scaffolds.
The anti‐inflammatory properties of our FeGA‐HA@PLGA scaffolds are not
only achieved by scavenging ROS, but also by supplementing iron ions
into the inflammatory microenvironment. The released iron ions can
reverse the iron deficient microenvironment caused by bacterial
proliferation, thereby activating the innate immune response suppressed
by iron deficiency, and promoting the differentiation of macrophages
toward the anti‐inflammatory M2 phenotype. This differentiation can be
observed by measuring macrophage markers (e.g., CD86 for M1 macrophages
and CD206 for M2 macrophages). As shown in Figure [143]5f, the
percentages of M1 macrophages are measured to be 26.5%, 25.0%, 27.2%,
and 17.1% for the lipopolysaccharide (LPS), PLGA‐LPS, HA@PLGA‐LPS, and
FeGA‐HA@PLGA‐LPS groups. In contrast, the M2 macrophage percentages in
these groups show an opposite trend where the FeGA‐HA@PLGA+LPS group
has the highest M2 macrophage content (i.e., 7.9%). These results
confirm the efficacy of FeGA‐HA@PLGA in alleviating the inflammatory
reaction, which is conducive to maintaining immune balance, combating
inflammatory responses, and inducing tissue repair.
An ideal orthopedic implant scaffold material should have favorable
osteogenesis and bone induction properties.^[ [144]^37 ^] After
completing the anti‐inflammatory process, the next important step of
the scaffolds is to promote osteogenic differentiation. The alkaline
phosphatase (ALP) staining and alizarin red staining of BMSCs are
commonly used to investigate their osteogenic differentiation status.
The ALP activity and calcium deposition are evaluated for different
scaffolds using a Transwell culture method (Figure [145]5g). After 7
days of cultivation, the FeGA‐HA@PLGA group shows increased ALP
activity compared to the PLGA and HA@PLGA groups, indicating that the
FeGA‐HA@PLGA scaffolds can promote proliferation and differentiation of
BMSCs (Figure [146]5h; Figure [147]S20, Supporting Information).
Interestingly, the ALP content is further enhanced when NIR irradiation
is applied, which implies that moderate thermal stimulation is also
beneficial for the proliferation and differentiation of BMSCs. The
deposition of calcium phosphate is considered a hallmark of bone
regeneration. The alizarin red staining results show that both
FeGA‐HA@PLGA and FeGA‐HA@PLGA NIR groups have pronounced calcium
deposition (i.e., red regions), indicating that the BMSCs in these two
groups are undergoing osteogenic differentiation or have already
differentiated into osteoblasts. These results confirm the outstanding
osteogenic efficacy of our FeGA‐HA@PLGA scaffolds, especially under NIR
irradiation. The outstanding pro‐osteogenesis of the FeGA‐HA@PLGA
scaffolds is attributed to the coupled factors including (i) the
regulation of iron homeostasis by the released iron ions, (ii) the
excellent anti‐inflammatory effects of GA molecules, and (iii) the mild
thermal stimulation generated by the FeGA photothermal agent.
2.6. Mechanism Exploration of Anti‐Inflammatory and Osteogenic‐Related of
FeGA‐HA@PLGA Scaffolds
To gain insights into the anti‐inflammatory and pro‐osteogenesis
mechanism of FeGA‐HA@PLGA scaffolds, the RNA transcriptome sequencing
(RNA‐seq) of BMSCs co‐cultured with HA@PLGA or FeGA‐HA@PLGA scaffolds
are measured and analyzed (Figure [148]6 ). The bioinformatics analysis
shows that there is a total of 652 differentially expressed genes
(DEGs) including 514 upregulated genes and 138 downregulated genes by
screening tens of thousands of genes for the two groups of HA@PLGA and
FeGA‐HA@PLGA (Figure [149]6c). Among these DEGs, some can be classified
as osteogenesis related DEGs including inflammation regulation (e.g.,
IL‐10, Bst2, and Cd72), osteogenic differentiation (e.g., Bmp6 and
Bmp2k) and vascular formation (e.g., Hif‐1a), which shows an
upregulation in FeGA‐HA@PLGA group and a downregulation in HA@PLGA
group (Figure [150]6a,b). Quantitative analysis of gene expression
distribution in various samples using TPM (Transcripts Per Million),
and the findings indicated that there was no significant batch effect
observed between the two groups of samples (Figure [151]6f).
Furthermore, PCA revealed the differences in gene expression
variability in BMSCs when comparing cultures with and without the
application of FeGA, specifically between the control group and the
group treated with HA@PLGA and FeGA‐HA@PLGA. (Figure [152]6d).
Conducting a Venn analysis between HA@PLGA and FeGA‐HA@PLGA revealed
12410 genes that are co‐expressed. The results indicate the presence of
shared and uniquely expressed genes between the two groups. In
addition, the effects of FeGA‐HA@PLGA scaffolds on the biological
process (BP), cellular component (CC), and molecular function (MF) in
BMSCs can be gained using the gene ontology (GO) analysis. The GO
analysis results show that DEGs in the FeGA‐HA@PLGA group are enriched
in numerous specific function‐related BPs (e.g., immunity regulation, T
cell‐mediated immunity, and angiogenesis promotion), CC (e.g.,
extracellular collagen and cell adhesion molecules) and MF (e.g.,
cytoskeletal protein binding, growth factor binding and signaling
receptor binding) (Figure [153]S21, Supporting Information). These BP,
CC or MF are guided by several signaling pathways that are closely
associated with immunity regulation and osteogenic differentiation
including KEGG, NF‐κB, PI3K‐AKT and MAPK pathways (Figure [154]6g).
FeGA induced modifications in cellular constituents and molecular
functions, encompassing the extracellular matrix, regions rich in
collagen, and the regulation of signaling receptor activities.
Specifically, the KEGG PI3K‐AKT and MAPK pathways are related to
osteogenesis and the NF‐κB pathway is noted by its regulation of the
immune response, inflammation and cell survival. PI3K‐AKT is also an
important intracellular pathway that can regulate cell survival,
proliferation and protein synthesis. Among these pathways, PI3K‐AKT and
NF‐κB, which are involved in osteogenesis and anti‐inflammatory, were
notably affected by FeGA. In addition, the GO analysis results of
HA@PLGA, FeGA‐HA@PLGA, and FeGA‐HA@PLGA NIR also show that mild heat
stimulation can promote cell proliferation differentiation and
biological regulation (Figure [155]S21–S24, Supporting Information).
Western blot analysis shows increased activity of PI3K‐AKT and MAPK
signaling pathways and decreased activity of NF‐κB signaling pathway
upon treatment of FeGA‐HA@PLGA, which is consistent with the KEGG
enrichment results (Figure [156]S25, Supporting Information). Moreover,
gene set enrichment analysis (GSEA) found that the immunomodulatory
proteins of Tnfrsf11a and Tlr4 were up‐regulated, indicating that
FeGA‐HA@PLGA scaffolds may exhibit a positive influence on
immunomodulation by up‐regulating the response of BMSCs to the iron
homeostasis regulation (Figure [157]6h–j). Intriguingly, the proteins
PIK3CG, VEGFA, and VEGFC were identified as being significantly
upregulated in processes that pertain to cellular reactions to
mechanical stimuli, hinting at a possible cooperative influence of
these pathways when interacting with FeGA‐HA@PLGA. These results reveal
that the FeGA‐HA@PLGA scaffolds can participate in the activation of
processes pertinent to inflammatory regulation, angiogenesis promotion
and pro‐osteogenic differentiation, to transform an infected
microenvironment into one conducive to bone regeneration.
Figure 6.
Figure 6
[158]Open in a new tab
Transcriptome sequencing analysis of BMSCs co‐cultured with HA@PLGA or
FeGA‐HA@PLGA scaffolds. a) Hot map of genes related to inflammation and
osteogenesis for HA@PLGA and FeGA‐HA@PLGA groups (n = 3). b) Volcano
plot showing differently expressed genes of the HA@PLGA and
FeGA‐HA@PLGA groups. c) The number of upregulated and downregulated
genes. d) Principal component analysis (PCA) suggesting the variance
between the HA@PLGA and FeGA‐HA@PLGA groups. e) Venn diagram for the
obtained genes and miRNAs. f) Box plot depicting TPM values. g) Pathway
enrichment of different genes involved in the immune system from gene
ontology analysis. h–j) Gene set enrichment analysis of
anti‐inflammatory and osteogenesis‐related signaling pathways.
2.7. Performance of FeGA‐HA@PLGA Scaffolds in Promoting Bone Regeneration
The infected femoral bone defect model using SD rats is used to
evaluate the efficacy of the FeGA‐HA@PLGA scaffolds for treating IBDs.
Antimicrobial analysis, osteogenic analysis and immunohistochemical
analysis are performed at the time points of 2 weeks, 4 weeks, and 8
weeks (Figure [159]7a). The implantation sites of HA@PLGA NIR and
FeGA‐HA@PLGA NIR groups are irradiated with 808 nm NIR (the power
density is 1.5 W/cm^2) for 5 min once a week after implantation
(Figure [160]7b). As shown in Figure [161]7b,c, the temperature of the
FeGA‐HA@PLGA scaffolds is rapidly increased to 42.0 °C within 30 s then
further to 47.8 °C within 300 s under the NIR irradiation, while the
temperature of the HA@PLGA scaffold shows no significant change under
the same condition. These results show that the FeGA‐HA@PLGA scaffolds
maintain their excellent photothermal effects when implanted into the
femoral bone defect. Intervention for bone infection must be initiated
in the early stages of infection. Otherwise, the infection may spread
to surrounding tissues and form local infection foci, which could lead
to more severe symptoms, such as suppuration, abscess formation, local
exudation, and even functional impairment. Early infection inhibition
effects of scaffolds are evaluated by retrieving and further in vitro
co‐culturing the implanted scaffolds with broth medium. As shown in
Figure [162]7d, the yellowing of the muscle tissues with secretion and
suppuration at the implantation sites for PLGA, HA@PLGA, and HA@PLGA
NIR groups can be visually observed. In contrast, the FeGA‐HA@PLGA
group shows minimal yellowing and suppuration. And it becomes even
better when the NIR irradiation is applied. These observations are
consistent with the turbidity of the culture mediums co‐cultured with
the extracted implants. The culture medium co‐cultured with the
FeGA‐HA@PLGA NIR group is clear while culture media co‐cultured with
other groups become turbid, indicating that there are bacterial
residues on the scaffolds of these groups. The bacterial concentration
is measured using the agar plate counting method (Figure [163]7d,e).
The results show that the FeGA‐HA@PLGA scaffolds has a certain
antibacterial effect (i.e., ≈60%) that can be largely enhanced under
NIR irradiation (i.e., ≈100%). While the other groups have no
antibacterial effects (i.e., ≈0%). These in vivo results confirm the
excellent photothermal antibacterial capability of our FeGA‐HA@PLGA
scaffolds.
Figure 7.
Figure 7
[164]Open in a new tab
Characterization of in vivo antibacterial ability of FeGA‐HA@PLGA
scaffolds. a) Schematic illustration of the animal experiment timeline.
b, c) Thermal graphics (b) and temperature changes (c) of implant sites
for the HA@PLGA and FeGA‐HA@PLGA groups under NIR irradiation (808 nm,
1.5 W cm^−2). d) Top: photographs of the implant sites and media
cultured with the implanted scaffolds; Bottom: photographs of bacterial
colonies co‐cultured with different scaffolds. e) In vivo antibacterial
efficiency of different implanted scaffolds. *p < 0.05, **p < 0.01,
***p < 0.001, ****p < 0.0001. f) The H&E staining images show the
degree of infection in the soft tissues and bone tissues surrounding
the implants (yellow stars represent the neutrophil cells).
To investigate the infection status at the implantation site, H&E
staining is performed on the harvested samples taken at the time point
of week 2. The H&E staining results reveal that inflammatory cells that
gather at the defect site in FeGA‐HA@PLGA and FeGA‐HA@PLGA NIR groups
are much fewer than those in the PLGA, HA@PLGA, and HA@PLGA NIR groups
(indicated by yellow stars), confirming that the FeGA‐HA@PLGA scaffolds
can attenuate early inflammatory responses in the treatment for IBDs.
The anti‐inflammatory effects of the FeGA‐HA@PLGA scaffolds could be
attributed to its iron homeostasis regulation as well as its excellent
antibacterial effects. The bone repair performance of our FeGA‐HA@PLGA
scaffolds is characterized by assessing scaffold degradation and the
formation of new bone at the defect sites using various methods
including Micro‐CT, histological staining, and immunohistochemical
staining (Figures [165]8, [166]9). The reconstructed three‐dimensional
images of the regenerated femur at week 8 are shown in Figure [167]8a,b
(the red region represents new bone and the green region represents the
scaffold). It is evident that the FeGA‐HA@PLGA NIR group has a higher
new bone density at the defect site compared to the HA@PLGA group. The
micro‐CT images from different views and three‐dimensional
reconstruction images show the information on new bone formation at the
defect sites (Figure [168]8b). A large amount of new bone can be
observed from edge to center in the FeGA‐HA@PLGA NIR group. In
contrast, the new bone amount in other groups is less than that in the
FeGA‐HA@PLGA NIR group. The ranking of new bone amount observed by
Micro‐CT is PLGA < HA@PLGA < HA@PLGA NIR < FeGA‐HA@PLGA < FeGA‐HA@PLGA
NIR. The quantitative analysis results are consistent with the Micro‐CT
images. Among all the groups, the values of trabecular thickness (Tb.
Th), bone volume/tissue volume (BV/TV) and bone mineral density (BMD)
in the FeGA‐HA@PLGA NIR group are the highest, while the value of bone
trabecular separation (Tb. Sp) in FeGA‐HA@PLGA NIR group is the lowest,
demonstrating the best bone repair performance of the FeGA‐HA@PLGA NIR
group (Figure [169]8c–f). The excellent bone repair performance is
mainly due to the reconstruction of the bone defect microenvironment
via iron homeostasis regulation combined with moderate photothermal
therapy.
Figure 8.
Figure 8
[170]Open in a new tab
Bone regeneration characterization of FeGA‐HA@PLGA scaffolds for
treating IBDs. a) 3D reconstruction images showing the formation and
distribution of new bones (highlighted in red). b) Micro‐CT images from
different views and three‐dimensional reconstruction images for
different groups at Week 4 and 8 after surgery. c–f) Quantitative
analysis of bone volume/tissue volume (BV/TV), trabecular thickness
(Tb. Th), bone trabecular separation (Tb. Sp) and bone mineral density
(BMD) in defect sites at Week 4 and 8 after surgery. g) H&E staining
and Masson trichrome staining of the scaffold implantation sites. The
yellow stars indicate the distribution of new bone. h,i)
Immunofluorescence staining of OCN (green) at Week 4 and 8 after
scaffold implantation. j,k) Quantitative expression of OCN. *p < 0.05,
**p < 0.01, ***p < 0.001, ****p < 0.0001.
Figure 9.
Figure 9
[171]Open in a new tab
Images of Immunofluorescence staining images of a) CD31, b) CD206, c)
IL‐6, and d) TNF‐α. e–h) Quantitative expression of CD31, CD206, IL‐6,
and TNF‐α. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.
In the H&E staining results, the defect areas are indicated by blue
dashed circles, while the locations of new bone are denoted by pink
regions marked with yellow stars. Upon examination of the first row of
Figure [172]8g, it becomes evident that the defect area progressively
diminishes from left to right. Notably, the PLGA group exhibits the
largest defect area, whereas the FeGA‐HA@PLGA NIR group displays the
smallest defect area. In the Masson staining, collagen and mineralized
bone are visualized by the blue color, as shown in Figure [173]8g. The
results indicate that the FeGA‐HA@PLGA NIR group exhibits the highest
deposition of collagen and mineralized bone area. The FeGA‐HA@PLGA
group without NIR irradiation has the second most collagen deposition
and mineralization, which is much more than the other groups (PLGA,
HA@PLGA, HA@PLGA NIR). As shown in Figures [174]S26,S27 (Supporting
Information), the H&E and Masson staining results collected in Week 4
maintain a similar trend to that in Week 8. These results are
consistent with in vitro osteogenic differentiation results
(Figure [175]5h), demonstrating that the FeGA‐HA@PLGA scaffolds
combined with mild thermal stimulation can promote osteogenic
differentiation both in vitro and in vivo.
Excessive inflammatory response can hinder osteogenesis, eventually
resulting in delayed bone regeneration or even implant failure.
Macrophage's marker for M2 polarization is closely related to
anti‐inflammatory and tissue repair, which promotes the repair and
regeneration of IBDs. To investigate the in vivo antioxidant and
immunomodulatory characteristics of the FeGA‐HA@PLGA scaffolds,
immunohistochemistry assays are conducted to assess the expression of
CD206, TNF‐α and IL‐6, while immunofluorescence staining is performed
to identify macrophage markers (e.g., CD206 as a marker for M2
phenotype). The results show that the expression levels of TNF‐α and
IL‐6 are quite high for the PLGA, HA@PLGA, and HA@PLGA NIR groups but
they are very low for FeGA‐HA@PLGA groups with or without NIR
irradiation (Figure [176]9b,c). The quantitative results are consistent
with the fluorescent staining results (Figure [177]9f,g). The CD206
(green color) immunofluorescence images show that the macrophages in
all groups are predominantly in anti‐inflammatory (M2) phenotype where
the amounts of CD206 in the FeGA‐HA@PLGA group is significantly higher
than that in other groups, showing the anti‐inflammatory effect of
FeGA‐HA@PLGA. The quantitative results of CD206 are consistent with the
fluorescent staining results (Figure [178]9b). It is worth mentioning
that a mild immune response is also essential to fight bacterial
infection and promote osteogenic differentiation of BMSCs. In the early
stage of IBDs, the FeGA‐HA@PLGA NIR group can effectively eliminate
bacteria due to its excellent photothermal properties. The slowly
released iron ions accelerate the immune response by regulating iron
homeostasis, thereby promoting the M2 polarization of macrophages that
is beneficial to osteogenesis. These results are consistent with the
expression levels of pro‐inflammatory and anti‐inflammatory genes in
the RNA‐seq results.
To further investigate the effects of the FeGA‐HA@PLGA NIR group on
angiogenesis and osteogenesis, CD31 and OCN as the representative
markers for angiogenesis and osteogenic differentiation are
investigated using immunofluorescent histochemical staining. As shown
in Figure [179]8j,k, the OCN expression level of the FeGA‐HA@PLGA group
is significantly higher than that of the other groups at both Week 4
and Week 8, which is further enhanced by applying NIR irradiation,
indicating the osteogenic differentiation effects of the FeGA‐HA@PLGA
scaffolds and the mild thermal stimulation. In addition,
immunofluorescence staining results show that the expression of CD31 in
the FeGA‐HA@PLGA NIR group is much higher than that in other groups
(Figure [180]9a), proving its pro‐angiogenic effects, which is
essential for osteoblast function, encompassing proliferation,
differentiation, and bone formation.^[ [181]^38 ^] The quantitative
results of CD31 and OCN are consistent with the fluorescent staining
results (Figures [182]9e and [183]8h,i). After eight weeks of
treatment, important organs are collected for histological examination
using H&E staining. No pathological abnormalities are observed in all
the groups (Figure [184]S28, Supporting Information), suggesting that
all the scaffolds have good biocompatibility in vivo.
3. Conclusions
In this work, we report on an in situ modification strategy for
preparing iron‐active multifunctional scaffolds with photothermal
antibacterial activity, and anti‐inflammatory and iron homeostasis
regulation abilities for infected bone regeneration. The strategy
involves the initial in situ growth of a layer of FeGA on the HA
ultralong nanowires and the subsequent incorporation of the modified
nanowires in the PLGA matrix for 3D printing. The as‐prepared
FeGA‐HA@PLGA scaffolds can effectively eliminate bacterial infection
through its excellent photothermal effects. Additionally, the released
iron ions can regulate dynamic iron homeostasis, reactivate the innate
immune system and promote the polarization of macrophages towards the
M2 phenotype, thereby transforming the infected microenvironment into
one conducive to bone regeneration. The animal experiments confirm that
the FeGA‐HA@PLGA scaffolds significantly promotes the formation of new
bone through combined antibacterial, anti‐inflammatory, pro‐angiogenic,
and osteogenic effects. The transcriptomic analysis shows that
FeGA‐HA@PLGA scaffolds exert anti‐inflammatory and pro‐osteogenic
differentiation by activating NF‐κB, MAPK and PI3K‐AKT signaling
pathways. In summary, this study provides a prospective strategy for
the regeneration of IBDs through the regulation of iron homeostasis by
combining biomimetic microarchitecture and iron‐active ions, and this
iron‐active multifunctional scaffold provides broad implications for
bone defect repair applications.
4. Experimental Section
Fabrication of HA and FeGA‐HA
The ultra‐long hydroxyapatite (HA) nanowires were synthesized using
this previous method.^[ [185]^26a ^] In a typical experiment, 10 mL of
CaCl[2] (22 g) aqueous solution and 10 mL of NaOH (7 g) solution were
each added into a mixture of water (9 mL), methanol (4 mL), and oleic
acid (7 mL) under mechanical agitation (300 r min^−1). Next, 10 mL of
NaH[2]PO[4]·2H[2]O (2.88 g) solution was introduced into the previous
mixture under mechanical agitation (300 r min^−1). The resulting
mixture was then transferred to a 100 mL vessel for solvothermal
reaction at 180 °C for 24 h. Upon cooling the reaction system to room
temperature, the solvothermal product slurry containing ultralong HA
nanowires was obtained. The ultralong HA nanowires were prepared by
dispersing the solvothermal product slurry in absolute ethanol. After
separation, the nanowires underwent three rounds of washing with
ethanol and deionized water, before being dispersed once again in
ethanol for future utilization. Iron gallate (FeGA) was grown in situ
on the surface of HA using a solvothermal method. 1 mL of 0.1 M
Fe(NO[3])[2]·9H[2]O and 1 mL of 0.3 M gallic acid (GA) were added to
13 mL of glycerol and stirred until dissolved. The pH was adjusted to 9
with sodium hydroxide, followed by agitation for 30 min and the
addition of 0.02 g of HA. The mixture was transferred to a 100 mL
vessel for solvothermal reaction at 180 °C for 24 h. Finally, the iron
gallate nanowires (FeGA‐HA) were purified with anhydrous ethanol and
dried at 60 °C.
Characterization of HA and FeGA‐HA
The morphological characterization of HA and FeGA‐HA was examined by
transmission electron microscope (TEM) with a JEOL JEM 2100F. The X‐ray
diffraction (XRD) patterns of HA and FeGA‐HA were acquired using the
Empyrea instrument (Rigaku Smartlab 9KW, Japan), while Fourier
transform infrared (FTIR) spectra were obtained via an FTIR
spectrometer (Thermo Scientific Nicolet iS20, USA). The HA and FeGA‐HA
were captured by a microscope.
The Molecular Dynamics of FeGA‐H
Calcium ion‐rich HA [001] surfaces were constructed respectively, and
the lengths of the HA [001] crystal model in the XYZ direction were
6.2 nm × 8.0 nm × 22.0 nm respectively. Fill the simulation box with
900 gallic acid molecules, 300 Fe^3+ ions, 300 glycerol molecules, and
counter ions respectively. The MD simulation uses the Gromacs 2019.6
program^[ [186]^39 ^] and was performed under constant temperature,
constant pressure, and periodic boundary conditions. The Charmm 36
all‐atom force field was used for small molecules,^[ [187]^40 ^] while
the INTERFACE force field was utilized for HAP molecules.^[ [188]^41 ^]
Parameters for the force fields were sourced from existing literature.
During the MD simulation, all hydrogen bonds involved were constrained
using the LINCS algorithm, and the integration step was 2 fs.
Electrostatic interactions were calculated using the (Particle‐mesh
Ewald) PME method.^[ [189]^42 ^] Finally, a 50 ns MD simulation was
performed on the composite object, and the conformation was saved every
10 ps. The visualization of the simulation results was completed using
the Gromacs embedded program and VMD.
3D Printing of PLGA, HA@PLGA, and FeGA‐HA@PLGA
A 3D porous composite scaffold was created with the assistance of a 3D
biological printer (Regenovo 3D Bio‐Architect Work Station, which was
purchased from Regenovo) under the supervision of a supporting computer
workstation. Before 3D printing, 0.15 g of HA or FeGA‐HA were dispersed
in 15 mL of 1,4‐Dioxane and stirred to form uniform dispersion,
respectively. Subsequently, 0.35 g of PLGA (85:15 lactide to glycolide
ratio, Mw 120000, from Shandong Institute of Biomaterials) was added to
the aforementioned mixed solution and stirred overnight. Then, the
product was evaporated for 1 hour to remove 1,4‐Dioxane. Finally,
computer‐aided design (CAD) software was employed for the 3D modeling
of scaffold materials. A cylindrical structure with a diameter of 3 mm
and a height of 4 mm was designed, configuring the material properties
to a filament diameter of 350 µm and a porosity of 60%.
Characterization of PLGA, HA@PLGA, and FeGA‐HA@PLGA
The morphological characterization of the scaffolds was examined by
scanning electron microscopy (SEM) with a ZEISS Gemini SEM 300. The XRD
patterns of the scaffolds were acquired using the Empyrea instrument
(Rigaku Smartlab 9KW, Japan), while FTIR spectra were obtained via an
FTIR spectrometer (Thermo Scientific Nicolet iS20, USA). The content of
FeGA‐HA within the FeGA‐HA@PLGA scaffolds was assessed using
thermogravimetry (Rigaku, Japan).
Thermogravimetry Analysis
HA, FeGA‐HA, PLGA, and FeGA‐HA@PLGA were performed on a
thermogravimetric analyzer. The temperature range for this study was
from room temperature to 900 °C. The furnace heating rate was 5
°C·min^−1 with a controlled mass flow of air of 60·mL min^−1 as reagent
gas. The sample mass placed in a scaffold inside the balance was around
10 mg for each analysis. Subsequently, the thermogravimetry (TG) curves
were analyzed.
In Vitro Fe Ions Release Measurement
The scaffolds were immersed in 3 mL of PBS (pH 7.4) in a shaker (37 °C,
100 rpm) for 1, 3, 5, 7, 14, 21, 28, and 70 days. The release of Fe
ions was quantified by using an inductively coupled plasma source
spectrometer (Prodigy‐ICP, USA).
In Vitro GA Release Measurement
To determine the GA release profile, five scaffolds of FeGA‐HA@PLGA
were put in 2 mL PBS. The supernatant was taken for absorbance
measurements at 259 nm using a UV spectrometer (UV‐5100, Shanghai,
China) at different times and an identical volume of fresh PBS solution
was added to the release system. Then, the corresponding concentrations
of the GA in the supernatant were calculated from the absorbances using
the calibration curve (Figure [190]S12, Supporting Information). The
release amount was determined using the following equation:
[MATH:
Release%=Ct/<
msub>M0×100% :MATH]
(1)
where C[t] refers to the calculated concentration of GA in the
supernatant at t day and M[0] refers to the mass of GA loaded in the
FeGA‐HA@PLGA scaffold.
Mechanical Characterization
The mechanical characteristics of PLGA, HA@PLGA, and FeGA‐HA@PLGA
scaffolds were assessed using a uniaxial mechanical testing apparatus
(Heng Yu Instrument Co., Ltd. HY‐940FS, Shanghai, China). Both the
inner diameter and the depth of the casting mold were precisely set to
1.0 cm. Dimensions of diameter and height for each specimen were
diligently measured and documented before the commencement of tests.
Compression tests on the scaffolds were performed at a uniform strain
rate of 2 mm per minute until failure was observed.
Photothermal Properties of FeGA‐HA
The photothermal properties of FeGA‐HA‐3 h, FeGA‐HA‐6 h, and
FeGA‐HA‐12 h nanowire were evaluated using NIR 808 nm, and temperature
changes were recorded using an infrared thermal imager. The
photothermal properties of FeGA‐HA nanowires were evaluated at a power
density of 0.25 W/cm^2 for 1 min.
Photothermal Properties of PLGA, HA@PLGA, FeGA‐HA@PLGA scaffolds
The photothermal properties of three scaffolds were evaluated using NIR
808 nm, and temperature changes were recorded using an infrared thermal
imaging camera. The photothermal properties of PLGA, HA@PLGA,
FeGA‐HA@PLGA scaffolds were assessed at various power
density(0.5 W cm^−2, 1 W cm^−2, 1.5 W cm^−2, 2 W cm^−2). The
corresponding temperature stability of the FeGA‐HA@PLGA scaffolds was
evaluated in five on/off cycles.
To determine the photothermal conversion efficiency (𝜂), the equation
derived from the principle of thermal equilibrium was utilized.^[
[191]^43 ^]
[MATH: η=hSTmax−Tsurr−Q0I :MATH]
(2)
where h (%) was the heat transfer coefficient, S (cm^2) was the surface
area of the scaffold, I (W cm^−2) was the effective sunlight power
irradiated upon the scaffold, T[max] was the equilibrium temperature of
the test scaffolds and T[surr] was the ambient temperature (the room
temperature was set at 26.8 degrees Celsius and temperatures were
recorded in degrees Celsius).
In Vitro Antibacterial Assay
The spread plate method was employed to evaluate the antibacterial
properties of different scaffolds against Gram‐negative E. coli
(ATCC25922) and Gram‐positive S. aureus (ATCC25923). Five different
groups were treated, including PLGA, HA@PLGA, HA@PLGA NIR,
FeGA‐HA@PLGA, and FeGA‐HA@PLGA NIR, respectively. For the HA@PLGA NIR
and FeGA‐HA@PLGA NIR groups, additional exposure to an 808 nm laser
(1.5 W cm^−2) was conducted for 10 min. The PLGA group was set as blank
controls. In each group, scaffold specimens were immersed in 1 × 10^8
CFU Ml^−1 bacterial suspensions for 5 h. The bacterial solution was
diluted to a concentration of 1 × 10^4 CFU mL^−1 using PBS.
Subsequently, 100 µL of the diluted bacterial solution was spread onto
LB agar and incubated at 37 °C. Then the bacterial colonies on the
plates were examined after incubation for 16 h. The relative bacteria
viability was assessed using the formula: Survival viability (%) =
N[t]/N[c] × 100%, where N[t] denotes the colonies formed in the
experimental group and N[c] refers signifies colonies formed in the
control group (PLGA group). The bacterial morphology on the different
scaffolds was further investigated by SEM to evaluate the antibacterial
effects. The co‐cultured supernatant and scaffold were fixed with
glutaraldehyde, dehydrated with gradient ethanol, and dried in air.
Subsequently, both the scaffold and supernatant were subjected to gold
sputtering for SEM observation.
NPN Uptake Assays
N‐phenyl‐1‐naphthylamine (NPN) was used to evaluate the permeability of
the bacterial membrane.^[ [192]^44 ^] A 1 mM stock solution of
N‐phenyl‐1‐naphthylamine (NPN) was prepared and then diluted to 40 µM
with 30% DMSO to obtain the working solution. E. coli and S. aureus
were cultured on a shaker for 24 h, after which 10^8 CFU mL^−1 were
transferred into a 24‐well plate. PLGA, HA@PLGA, and FeGA‐HA@PLGA were
introduced into the bacterial suspensions for photothermal or
non‐photothermal treatment. Treated suspensions, 150 µL in volume, were
added to a 96‐well plate and co‐incubated with 50 µL of the 40 µM NPN
solution at 37 °C for 30 min. Fluorescence intensity at 420 nm was
measured upon excitation at 350 nm.
Cell Culture
Bone mesenchymal stem cells (BMSCs) were harvested from the femurs and
tibias of two‐week‐old Sprague‐Dawley (SD) rats, acquired from SLAC
(Shanghai, China). These BMSCs were propagated in α‐MEM supplemented
with 10% fetal bovine serum (FBS, Gibco, USA) and 1%
penicillin/streptomycin (PS, Gibco, USA), and maintained at 37 °C
within a humidified 5% CO[2] incubator. The culture medium was
refreshed every 2–3 days, and cells reaching confluence were
subcultured every 3–4 days following the established protocol. BMSCs
from passages 3–5 were utilized in subsequent experiments.
Cell Viability Test of Scaffolds
In Vitro: First, the sectioned small‐sized scaffolds were irradiated
under ultraviolet light for 1 h, and then placed into α‐MEM culture
medium supplemented with 10% FBS. After scaffold preparation, BMSCs
were seeded at a concentration of 1 × 10^4 cells per well in a 96‐well
plate and cultured with α‐MEM for 24 h, and then put into the scaffolds
and cultured for 24 h. The cells were transferred onto the prepared
scaffolds and further incubated for an additional 24 h to facilitate
cell‐scaffold interactions. The enhanced cell counting kit‐8 (CCK‐8,
Beyotime Biotechnology, Jiangsu, China) was used to assess cell
viability. Then, the optical density (OD) value of the reaction
solution was measured using the microplate reader at a wavelength of
450 nm.
Live/Dead Viability and Cellular Morphology on Scaffolds
In Vitro: First, the scaffolds were sterilized using ultraviolet
irradiation, and BMSCs were then seeded onto the thin‐layer scaffolds
at a density of 2 × 10^4 cells per well within a 24‐well plate. After
incubating for 24 h, cell viability among different treatment groups
was evaluated using the live/dead staining kit (Beyotime Biotechnology,
Jiangsu, China), following the provided protocol. The optical images of
the live (green)/dead (red) staining of BMSCs cocultured with
thin‐layer scaffolds were recorded using a fluorescence microscope
(Olympus Corporation, Japan).
To characterize cellular morphology, a sterile thin‐layer scaffold was
carefully placed at the center of a confocal dish and anchored with a
sterile stainless steel ring to prevent displacement. Cells were then
seeded onto the scaffold at a controlled density of 5 × 10^3 cells per
scaffold. After incubation for 3 days, the cell culture dishes were
washed with PBS twice. Then, the scaffolds were fixed with 4.0%
paraformaldehyde for 20 min and subsequently subjected to three washes
with 0.1% Triton X‐100. For cytoskeletal visualization, Actin‐Tracker
Red‐Rhodamine (Beyotime Biotechnology, China) was diluted by PBS
solution (containing 3% of BSA and 0.1% of Triton X‐100). The scaffolds
were incubated with diluted Actin‐Tracker Red‐Rhodamine solution at a
1:100 dilution for 40 min, followed by nuclear counterstaining with
DAPI provided by Servicebio (Hubei, China) for a brief ten‐minute
period. Subsequently, the cytoskeleton and cell nucleus of the
cocultured BMSCs were stained by fluorescent Actin‐Tracker
Red‐Rhodamine and DAPI, respectively. Optical imaging of the stained
BMSCs was conducted with a confocal laser scanning microscope (ZEISS,
LSM710, Germany).
Hemolysis Assay
The 4% erythrocyte suspension was incubated with PLGA, HA@PLGA,
FeGA‐HA@PLGA, and FeGA‐HA@PLGA NIR at 37 °C for 4 h in a cell culture
incubator. Deionized water and PBS containing purified RBCs were used
as the positive and negative controls, respectively. The samples were
centrifuged at 1500 rpm for 10 min to pellet the blood cells.
Subsequently, 100 µL of the sample supernatant without scaffold was
carefully transferred to a 96‐well plate for further analysis. The
absorbance attributable to hemoglobin present in the supernatant was
quantified using a microplate reader set to a wavelength of 540 nm. The
hemolysis radio of each sample was calculated using the following
equation: Hemolysis% = (A[test] − A[neg])/(A[pos] − A[neg]) × 100%,
where A[test], A[pos], and A[neg] were the absorbance values of the
sample, and the positive and negative groups, respectively.
Antioxidant Effect of Scaffold
In Vitro: The production of intracellular ROS (Reactive Oxygen Species)
was measured using the fluorescent probe 2′,7′‐dichlorofluorescin
diacetate (DCF‐DA). Before the assay, the BMSCs were seeded at a
density of 2 × 10^4 cells per well in a 6‐well plate. Then the
different scaffolds (PLGA, HA@PLGA, FeGA‐HA@PLGA) were incubated with
BMSCs for 48 h (37 °C, 5% CO[2]). After treatment, the cells were
washed twice with PBS and then incubated with 10 µM DCF‐DA in
serum‐free medium for 30 min at 37 °C in the dark. The DCF‐DA solution
was then removed, and the cells were washed with PBS to eliminate any
non‐internalized probe. ROS production was observed using fluorescence
microscopy (ZEISS Axio Observer 3, Germany).
Macrophages Polarization Assessment
In Vitro: To ascertain the effect of various scaffolds on macrophage
polarization, raw 264.7 macrophages were incubated for 24 h, and then
stimulated with lipopolysaccharide (LPS) at 100 ng mL^−1 for 24 h.
After removing the culture medium and washing twice with PBS, the cells
were incubated with PBS, PLGA, HA@PLGA, and FeGA‐HA@PLGA for 48 h. The
positive control for M2 macrophage polarization was treated with
interleukin‐4 (IL‐4) at 20 ng mL^−1. Detailly, raw 264.7 macrophages
were collected and incubated with 100 µL PBS containing
fluorochrome‐conjugated antibodies against CD86 (a marker for M1
macrophages) and CD206 (a marker for M2 macrophages) for 30 min at
4 °C. Flow cytometric analysis was conducted using a BD LSR Fortessa
X‐20 flow cytometer. Results were subsequently analyzed using FlowJo
analysis software.
ALP Staining and ARS Staining
Alkaline phosphatase (ALP) and alizarin red S (ARS) staining assays
were conducted to assess the osteogenic differentiation potential of
BMSCs when co‐cultured with various scaffolds. BMSCs were seeded at a
density of 5 × 10^4 cells per well in a 24‐well plate and cultured for
24 h, and then the medium was changed by the α‐MEM containing scaffold
with NIR or without NIR. The scaffold‐free group was set as the control
group. After incubation for 7 and 14 days, the cells were washed thrice
with PBS and fixed with 4% paraformaldehyde for 20 min. After fixation,
the cells were subjected to an additional three washes with PBS to
eliminate the remaining paraformaldehyde. For staining, the BCIP/NBT
alkaline phosphatase color development kit and a 0.2% ARS staining
solution (Solarbio, Beijing, China) were applied to the respective
wells for 30 min according to the manufacturer's instructions. The
stained cells were then visualized using an inverted fluorescence
microscope. The quantitative analysis of ALP staining was carried out
via Image J software (NIH, USA) by quantifying the stained areas.
Transcriptome Sequencing and Data Analysis
RNA‐sequencing analysis was used to evaluate the expression of mRNA
profiles in BMSCs for the HA@PLGA scaffold group and FeGA‐HA@PLGA
scaffolds group. The RNA sequencing was used to explore the role of
FeGA in regulating immune responses and osteogenic differentiation.
Furthermore, the mRNA profiles in BMSCs were assessed to compare the
photothermal‐treated group (FeGA‐HA@PLGA NIR group) with the
non‐photothermal‐treated group (FeGA‐HA@PLGA group). BMSCs (5 × 10^6
cells/mL) were co‐cultured with the different scaffolds as described
above for 7 days and cultured in an osteogenic medium. Cells in the
five groups were then lysed using TRIzol reagent (Ambion, Carlsbad,
CA), and cell lysates were stored at ‐80 °C pending further analysis.
Eukaryotic mRNA sequencing experiments used the Illumina TruseqTM RNA
sample prep Kit method (Illumina Hiseq2000 platform, Majorbio Biotech,
Shanghai, China) for library construction. The data were analyzed
online with Majorbio Biotech cloud platform, and the cluster Profiler R
package was utilized to conduct Kyoto Encyclopedia of Genes and Genomes
(KEGG) pathway enrichment analysis, aimed at evaluating the molecular
or biological functions of DEGs and identifying enriched metabolic
pathways within DEGs compared to the entire genome background.
Significance was determined by a Corrected P‐value < 0.05 for
enrichment by DEGs.
Western Blot Analysis
For Western Blot (WB) analysis, the BMSCs treated by various scaffolds
(HA@PLGA and FeGA‐HA@PLGA) were lysed using RIPA lysis buffer
(Beyotime, China) contained with proteinase and phosphatase inhibitors
(APExBIO) for 15 min at 4 °C. Then, the mixture was centrifuged at 4 °C
(13,000 rpm, 20 min), mixed with loading buffer (Beyotime, China),
boiled for electrophoresis, and transferred to PVDF membranes
(Millipore, USA). After the membranes were blocked with 5% BSA, the
PVDF membranes were incubated with primary antibodies at 4 °C overnight
and then washed three times by PBST (PBS with Tween). Next, incubated
with secondary antibodies (Beyotime, China) for 1.5 h at room
temperature. Ultimately, the membranes underwent processing with the
Odyssey Scanning system, a product of Li‐Cor, USA, and subsequent
analysis was performed using the Image Studio software.
In Vivo Evaluation of Bone Regeneration of Femoral Defect Infected with
Bacterial
All animal experiments were carried out according to protocols approved
by the Animal Care and Use Committee of Shanghai Tenth People's
Hospital, School of Medicine, Tongji University, Shanghai, China
(Ethical approval number: SHDSYY‐2023‐44850102). Male Sprague Dawley
(SD) rats (12 weeks, 240 g) were employed as the standard animal
models. Before the surgery, the scaffolds (⊝ 3 mm × 4 mm) were
contaminated with S. aureus. The rats were anesthetized, and the
scaffolds contaminated with S. aureus were implanted into the
cylindrical defects of the tibia. In the animal grouping, the
photothermal therapy group was illuminated with NIR light (808 nm,
1.5 W cm^−2) for 5 min each time once a week, continuing until the
sixth‐week post‐treatment. After 2 weeks post‐implantation, 3 rats from
each group were euthanized to evaluate bacterial infection at the
implant sites. The remaining 8 rats in each group were sacrificed after
4 and 8 weeks post‐implantation, and the tibias with different implants
were harvested and fixed before micro‐CT scanning and histological
analysis.
Micro‐CT
To evaluate the effect on bone regeneration, the obtained specimens
were scanned and analyzed by the Micro‐CT system (Hiscan XM Micro CT).
The X‐Ray tube was set to 80 kV and 100 µA, and images were captured at
a resolution of 25 µm. Imaging involved a rotation step of 0.5 through
a 360‐degree angular range, with each step having a 50 ms exposure. The
cylindrical bone defect region (diameter: 3 mm and depth: 4 mm) was set
as the region of interest for quantitative analysis of typical
parameters. Trabecular bone parameters were assessed, encompassing bone
mineral density (BMD, g cm^−3), bone volume fraction (BV/TV, %),
trabecular number (Tb. N, 1/mm), and trabecular thickness (Tb. Th, mm).
Histology and Immunohistochemistry
After decalcification following the Micro‐CT scan, the samples were
dehydrated in a graded ethanol series and embedded in paraffin. Then,
sections of approximately 5 µm were obtained for further staining.
First, H&E and Masson's trichrome staining were performed to evaluate
bone formation and residual materials. Additionally, immunofluorescence
staining (CD86 and CD206, IL‐6 and TNF‐α) evaluated the levels of
inflammatory cytokines. Furthermore, immunofluorescence staining was
carried out to evaluate the expression of osteogenic marker proteins
(OCN). The microvessel density (MVD) was also evaluated by CD31
immunofluorescence staining. The quantification of MVD was determined
by averaging the number of positive cells in three vascularized
regions. Briefly, the deparaffinized sections were first blocked with
5% bovine serum albumin (BSA) solution, followed by incubation with
primary antibodies against CD86, CD206, IL‐6, TNF‐α, CD31and OCN at a
1:100 dilution overnight at 4 °C. In addition, to assess the potential
toxicity of implanted scaffolds in vivo, major organs, including the
heart, liver, spleen, lung, and kidney, were collected at 6 weeks and
stained with H&E. All samples were imaged with an optical microscope.
The staining mentioned above collectively substantiates that the
FeGA‐HA@PLGA NIR group reconstructs the osteogenic microenvironment in
infected bone defects by modulating the immune milieu, promoting
angiogenesis, and enhancing osteogenic differentiation, thereby
facilitating bone regeneration.
Statistical Analysis
All data were expressed as means ± standard deviation (SD). Statistical
analysis was performed by one‐way analysis of variance (ANOVA) with
post‐hoc Tukey's method for multiple comparisons. The values of *p <
0.05, **p < 0.01, ***P < 0.001, and ****P < 0.0001 for all tests were
considered statistically significant.
Conflict of Interest
The authors declare no conflict of interest.
Supporting information
Supporting Information
[193]ADVS-11-2407251-s001.docx^ (5MB, docx)
Acknowledgements