Abstract
Implant materials for bone regeneration necessitate a barrier function
to block bacterial adhesion and fibroblast infiltration, while
maintaining a delicate equilibrium between material degradation and
osteogenesis. Here, a spatiotemporally and hierarchically‐guided bone
regeneration hydrogel with a Janus structure is engineered through a
sequential photocuring protocol, which features full barrier protection
by the outer dense phase and superior osteoconductivity within the
inner loose phase. The Janus hydrogel exhibits stable spatiotemporal
layering, adaptable degradation, asymmetrical combination of network
structures, and mechanical strength. The dense phase, with space
maintenance capacity, completely covers the defective area,
continuously blocking fibroblast infiltration, and preventing bacterial
adhesion. In addition, the loose phase is shape‐adapted to the
defective cavity, allowing osteoblast‐associated cells to migrate and
create a favorable osteogenic microenvironment. In situ implantation of
this Janus hydrogel effectively promoted osteogenesis, angiogenesis,
and neurogenesis in both mouse calvarial and rat periodontal bone
defect models. Furthermore, the osteogenic efficiency achieved by the
Janus hydrogel implanted in mouse calvarial defects and rat periodontal
defects is increased by 42% and 13.7%, respectively, as compared with
previous studies. These findings thus demonstrated the synergy of
protective barrier function, osteoconductive properties, and adaptive
degradation within a single scaffold, which is conducive to bone
regeneration.
Keywords: adaptable degradation, integrated Janus hydrogel,
osteoconductive, protective barrier
__________________________________________________________________
The filler‐barrier hydrogel system developed by dual‐network Janus
structural strategy breaks through the difficulty of a single hydrogel
to combine barrier protection with bone regeneration, and realizes the
regeneration‐material degradation adaptation for the field of
mandibular regeneration. Multi‐omics analysis elucidates the underlying
mechanisms by which the material regulates tissue regeneration and
lipid metabolism pathway.
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1. Introduction
The goal of bone regenerative medicine is to restore the normal
physiological activities and functions of injured or diseased bone
tissues. The optimal strategy for implant biomaterials to achieve this
requires maintaining the delicate balance between material degradation
and tissue regrowth to maximize material function during bone repair^[
[52]^1 ^] However, most studies often only consider the biological
function of the implant material itself to promote bone regeneration.
The infiltration of non‐osteogenic cells and tissues into the bone
defect will upset the degradative–regenerative balance.^[ [53]^2 ,
[54]^3 ^] If bone tissue is infected, the bacterial invasion will cause
further complications, which could result in failed repair or delayed
healing.^[ [55]^4 , [56]^5 ^] Hence, precise coupling of different or
even contradictory material properties and biological characteristics,
such as barrier protection and guided regeneration, as well as
preventing bacterial adhesion and promoting cell‐activation, is much
desired in the field of bone tissue engineering but remains challenging
to achieve.
Hydrogels have been widely used in tissue regeneration due to their
advantages such as, amenability to on‐demand injection, in situ
gelation, and minimally invasive implantation.^[ [57]^6 , [58]^7 ^]
Moreover, hydrogels can also provide a conducive physical
microenvironment for cell growth and differentiation due to their
suitable network structure.^[ [59]^8 , [60]^9 ^] In recent years,
injectable hydrogels with in situ gel‐forming properties upon exposure
to chemical, temperature, pH or light stimuli were designed to simulate
the anisotropic functional and structural characteristics of bone and
have achieved progressive results in promoting osteogenesis, thereby
attracting extensive attention among orthopedic researchers.^[ [61]^10
, [62]^11 ^] Jiang et al. developed a novel injectable in situ forming
composite hydrogel system with physico‐chemical properties and
excellent osteogenic/angiogenic function by introducing fibroin and
sodium alginate for minimally invasive treatment of jaw regeneration.^[
[63]^12 ^] It was reported that a co‐assembly system that integrates
hyaluronic acid tyramine, bioactive peptide amphiphiles, and Laponite
to engineer hydrogels can be fine‐tuned to enhance bone regeneration.^[
[64]^13 ^] However, currently available hydrogels are limited in
clinical application for bone regeneration under physiological
conditions, and in particular, lack the ability of resistance to
fibrous tissue infiltration and bacterial adhesion and adaptable
degradation. Therefore, it is necessary to develop an advanced
hydrogel‐based scaffold with distinct functionalities on opposite
sides, referred to as a Janus hydrogel‐based scaffold to meet the needs
of bone repair under both physiological and pathological conditions.
Silk fibroin (SF) and chondroitin sulfate (CS) are biocompatible and
have been approved by the Food and Drug Administration (FDA) for
biomedical applications.^[ [65]^14 , [66]^15 ^] Here, based on multiple
adaptations including mechanics and pore structure, a spatiotemporally
Janus hydrogel through a simple sequential photocuring was developed,
which can comprehensively overcome various challenging issues during
the regeneration processes (Scheme [67]1 ). The silk‐methacrylate
(SF‐MA) phase exhibits a microporous structure and adhesion to seal the
edge of the defect cavity, so it has a sufficient space maintenance
capacity. The chondroitin sulfate‐methacrylate (CS‐MA) phase exhibits a
macroporous structure and shape plasticity that can adapt to various
bone surface morphologies. Based on the above asymmetric design, we
integrate the framework with plasticity to meet the desirable space
properties of the regeneration proccess. In terms of the hierarchical
function, the SF‐MA dense phase can continuously act as a barrier
against both fibroblasts and bacteria, while the CS‐MA phase is
conducive to the ingrowth of osteogenesis‐associated cells.
Furthermore, we demonstrated that this integrated osteoconductive Janus
hydrogel achieved adaptive degradation with new bone formation in both
mouse calvarial defect and rat periodontal defect models. Mechanistic
study revealed that various signaling pathways associated with cell
proliferation, differentiation, and tissue regeneration were
significantly activated during bone regeneration, with lipid metabolism
being observed to be the most enriched among multiple metabolic
pathways. Combined LC‐MS/MS analysis suggested that the integrated
Janus hydrogel orchestrated cellular lipid metabolism that promoted the
osteogenic differentiation of Rat Bone Marrow Mesenchymal Stem Cells
(rBMSCs). Based on our results, our integrated Janus hydrogel scaffold
achieves both barrier protection and osteogenic enhancement, which
offers much promise in clinical bone regeneration.
Scheme 1.
Scheme 1
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Schematic diagram of the integrated osteoconductive Janus hydrogel in
bone regeneration. a) This integrated osteoconductive Janus hydrogel
was fabricated by sequential application of photocuring, comprising
methacrylated silk fibroin (SF‐MA) hydrogel with barrier protection
properties and methacrylated chondroitin sulfate (CS‐MA) hydrogel with
osteoconductive property. b) After implantation to fill the bone
defect, the integrated osteoconductive Janus hydrogel facilitated
improved new bone formation with adaptable degradation.
2. Results and Discussion
2.1. Design Principles and Application Scenarios of the Integrated
Osteoconductive Janus Hydrogel Scaffold
Functional chemical modifications of injectable hydrogels by grafting
groups or ions, is a reliable method for regulating the physical
properties of injectable hydrogels.^[ [69]^16 ^] To realize the
integration of osteoconductive and protective barrier functions for
synergistically promoting bone defect repair, an integrated
osteoconductive Janus hydrogel was fabricated, which was composed of
SF‐MA hydrogel and CS‐MA hydrogel. Previous studies have shown that CS
is a biologically active polysaccharide that enhances the expression of
osteogenic genes and repair of bone microstructure.^[ [70]^17 ^]
Moreover, CS polymer chains contain a number of sulfate and carboxyl
groups that regulate cytokine recruitment and promote cell adhesion,
migration, proliferation, and differentiation.^[ [71]^18 , [72]^19 ^]
SF is a natural fiber polymer extracted for wound dressings, cartilage
regeneration, and other tissue engineering because of its
biocompatibility, biodegradability, and high strength.^[ [73]^20 ,
[74]^21 , [75]^22 ^] SF can also be used as an important material
source for guiding bone regeneration membranes in bone tissue
reconstruction.^[ [76]^23 , [77]^24 ^] Additionally, a previous study
demonstrated that owing to the presence of carboxyl groups within the
amorphous region of the SF molecular chains, the SF coating increased
the polarity and water contact angle of the materials.^[ [78]^25 ^] The
successful introduction of carbon–carbon double bonds into the
molecular chains of CS and SF through chemical modification techniques
achieves the synthesis of photocurable hydrogel (CS‐MA and SF‐MA)
(Figure [79]1a). Due to the presence of gingival fibroblasts and the
high microbial density of the oral environment, periodontal defect
healing often has a poor prognosis. To overcome the aforementioned
clinical challenges, the integrated osteoconductive Janus hydrogel was
obtained by sequential injection and photocuring (Figure [80]1b). The
chemical structures of CS‐MA and SF‐MA were confirmed by ^1H NMR
spectrum (Figure [81]S1a, Supporting Information), which clearly
exhibited the signals of vinyl protons at 5.6 and 6.2 ppm (─CH[2]). In
this study, the CS‐MA hydrogel was designed with two concentrations of
5% w/v (LCS) and 10% w/v (HCS), which was aimed at investigating the
effects of bonding and regulating cytokines and growth factors involved
in osteogenesis. For the SF‐MA hydrogel, we designed three hydrogels
with concentrations of 5% w/v (SF‐MA 5%), 10% w/v (SF‐MA 10%), and 30%
w/v (SF‐MA) respectively. When the concentration was 30% w/v, the
cross‐section of the SF‐MA hydrogel exhibited a microporous structure
with a smooth and dense outer surface (Figure [82]S1b,c, Supporting
Information), as observed under scanning electron microscopy.
Therefore, we chose 30% (w/v) SF‐MA hydrogel for one phase of the
integrated osteoconductive Janus hydrogel in subsequent experiments.
Figure 1.
Figure 1
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Design principles and application scenarios of the integrated
osteoconductive Janus hydrogel. a) Schematic diagram of the fabrication
process of the SF‐MA and CS‐MA phases. b) The design principles of the
integrated osteoconductive Janus hydrogel.
2.2. Structural and Mechanical Characteristics of the Integrated
Osteoconductive Janus Hydrogel
Owing to the introduction of carbon–carbon double bonds, the hydrogel
exhibited the properties of injectability and rapid photocuring, which
can be adjusted by irradiation time, thus enabling convenient
application. As shown in Figures [84]2a and [85]S1d (Supporting
Information), the precursor solution was able to pass through a
26‐gauge needle and displayed quick curing in molds with various shapes
upon irradiation. By sequentially adding CS‐MA precursor solution to
SF‐MA precursor solution and photocuring respectively, the integrated
osteoconductive Janus hydrogel was thus fabricated (Figure [86]2b),
with the SF‐MA precursor solution displaying fluidity, and being able
to partially penetrate the CS‐MA hydrogel, to form a mechanical
interlock with chemical bonding after photocuring. To investigate the
internal structure of the integrated osteoconductive Janus hydrogel and
the interface bonding between the SF‐MA hydrogel phase and CS‐MA
hydrogel phase, we found that the LCS hydrogel and the HCS hydrogel had
a homogeneous, porous structure, with the pore size of the LCS hydrogel
being 102.78±2.13 µm and that of the HCS hydrogel being 66.72±2.01 µm.
By contrast, the structure of the SF‐MA hydrogel was relatively
microporous with a pore size of just 5.06±0.19 µm (Figure [87]2c;
Figure [88]S1e, Supporting Information). Additionally, the SF‐MA
hydrogel and the CS‐MA hydrogel were closely integrated with no obvious
gap being observed. Longitudinal‐sectional scanning showed a
well‐integrated interface between the SF‐MA hydrogel phase and the
CS‐MA hydrogel phase, with the SF‐MA hydrogel penetrating the CS‐MA
hydrogel such that mechanical interlocking and cohesion could be
observed, which further confirmed the physical integrity of the
integrated osteoconductive Janus hydrogel (Figure [89]2d). Sufficient
mechanical strength and stability of the hydrogel are very important
for functional stability and for creating a conducive pro‐osteogenic
microenvironment.^[ [90]^26 , [91]^27 ^] As shown in Figure [92]2e, all
hydrogels exhibited similar non‐linear rheological behavior. In the
angular frequency range (0.1–10 rad s^−1), the storage modulus (G') is
higher than the loss modulus (G''). This indicated that all hydrogels
had stable 3D network structures, in which they could be able to
maintain the original hydrogel network structure during the functional
state. Hydrogels are thought to better mimic the natural bone ECM in
complex bone defects microenvironments, providing mechanical signals to
promote cell adhesion, proliferation, and osteogenic differentiation.^[
[93]^28 ^] The results in Figure [94]2f and Figure [95]S1f,g
(Supporting Information) showed that the contents of CS‐MA in the
hydrogel matrix exerted a significant effect on the rheological and
mechanical properties of the hydrogel. With an increase of CS‐MA
content, the mechanical properties of the integrated osteoconductive
Janus hydrogel were improved, which could be attributed to their higher
cross‐linking density. Moreover, the SF‐MA hydrogel showed stronger
mechanical properties compared with the CS‐MA hydrogel or the GelMA
(Control). The mechanical strength of our hydrogels greatly exceeded
that of the natural hematoma fibrin clot, which was regarded as the
minimum strength during the bone healing process.^[ [96]^29 , [97]^30
^] These results suggested that the Janus hydrogel met the requirements
in the complex mechanical environment of bone defect sites. Together,
we successfully fabricated an integrated osteoconductive Janus
hydrogel, which exhibited a stable 3D network structure and
considerable mechanical properties.
Figure 2.
Figure 2
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Structural and mechanical characterization of the integrated
osteoconductive Janus hydrogel. a) The injectability and in‐situ
gelation of the integrated osteoconductive Janus hydrogel. b) The
fabrication process of the integrated osteoconductive Janus hydrogel.
c) Representative SEM images of the integrated osteoconductive Janus
hydrogel. The area between the red dotted lines denotes the fusion of
the CS‐MA hydrogel phase with the SF‐MA hydrogel phase. d)
Representative CLSM images of the integrated osteoconductive Janus
hydrogel. The area between the white dotted lines shows the fusion of
the CS‐MA hydrogel phase and SF‐MA hydrogel phase. e) Rheological
properties of the hydrogels after photocuring as measured by the
frequency sweep test at a constant strain of 1% at 25 °C. f)
Stress‐strain curve of the integrated osteoconductive Janus hydrogel.
2.3. In Vitro Swelling Behavior, Degradation Performance and Structural
Stability
The swelling properties of the hydrogel within the in vivo environment
affect its function.^[ [99]^31 , [100]^32 ^] Hydrogels in contact with
blood will rapidly swell without dissolving in vivo, which can mimic
the natural tissue environment to provide support for the defects.
However, too much swelling performance can lead to its original
function and structure being affected. By evaluating the swelling
behavior of the integrated osteoconductive Janus hydrogel within
modified simulated body fluid (SBF), we found that both the LCS‐SF‐MA
(LCS‐SF) hydrogel and the HCS‐SF‐MA (HCS‐SF) integrated osteoconductive
Janus hydrogel rapidly swelled during the 1 h, and gradually reached
swelling equilibrium after 5 h (Figure [101]3a; Figure [102]S2a,
Supporting Information). Hydrogels with high concentrations have been
reported to exhibit higher cross‐linking densities due to having more
carbon–carbon double bonds.^[ [103]^33 ^] As expected, the HCS‐SF
integrated osteoconductive Janus hydrogel displayed less swelling
capacity with an equilibrium swelling percentage of 120% compared to
the LCS‐SF integrated osteoconductive Janus hydrogel, which was mainly
attributed to its higher concentration of CS (Figure [104]3a). Notably,
both the LCS‐SF and HCS‐SF integrated Janus hydrogel stably maintained
the combination during the swelling process, which proved the organic
binding of the two phases (Figure [105]3b). The degradation performance
of the hydrogels in vitro was conducted in SBF over 25 days to mimic
degradation in the biosystem.^[ [106]^34 , [107]^35 ^] Upon exposure to
SBF, all the hydrogels displayed partial degradation behavior in the
first 5 days with different degradation rates. The GelMA hydrogel
showed the fastest degradation rate while the HCS‐SF integrated
osteoconductive Janus hydrogel exhibited the slowest degradation rate
(≈72% remaining after 25 days), which was also attributed to
light‐induced crosslinking of carbon–carbon double bonds and high
concentrations of CS (Figure [108]3c). Moreover, it was revealed that
the integrated osteoconductive Janus hydrogel interfacial bonding
remained stable during the 25‐day in vitro degradation process. With
the prolongation of degradation time, the pores of the CS‐MA hydrogel
gradually increased and the structure disintegrated, while the SF‐MA
hydrogel still maintained a stable structure when the pores increased
(Figure [109]3d; Figure [110]S2b, Supporting Information). Notably, the
average pore size of the SF‐MA hydrogel was still less than 35 µm on
the 25^th day of degradation (Figure [111]3e). Additionally, we also
found that the in vitro weight loss of the SF‐MA hydrogel was only 30%
on the 25^th day, while the in vivo weight loss of SF‐MA hydrogel was
only 50% on the 35^th day (Figure [112]3f; Figure [113]S2c, Supporting
Information). As shown in Figure [114]3g, we observed the adhesion of
SF‐MA hydrogel to the tissue. Early removal of biological barrier
membrane or premature loss of barrier structure due to rapid absorption
will lead to poor osteogenesis.^[ [115]^36 , [116]^37 ^] The above
results demonstrated that the integrated osteoconductive Janus
hydrogels have a relatively stable structure in SBF solution with rapid
swelling equilibrium and appropriate degradation rate adapted to the
needs of neo‐natal bone repair, which are critical for bone
regeneration.
Figure 3.
Figure 3
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The integrated osteoconductive Janus hydrogel maintained a relatively
stable structure in vitro. a) Swelling ratio of the integrated
osteoconductive Janus hydrogel in PBS solution at 37 °C for 25 h (n =
3). b) Representative real‐time images of the integrated
osteoconductive Janus hydrogel swelling over 25 h. c) Degradation of
the integrated osteoconductive Janus hydrogel in vitro (n = 3). d)
Representative SEM images after 1, 5 and 15 days of hydrogel
degradation. The area between the red dotted lines denoted the fusion
of the CS‐MA hydrogel phase with the SF‐MA hydrogel phase. e)
Quantitative analysis of pore size within the integrated
osteoconductive Janus hydrogel after degradation. f) In vitro
degradation properties of the SF‐MA hydrogel. g) Tissue adhesion of the
SF‐MA hydrogel. The black arrow indicated that the SF‐MA adhered to the
tissue. (ns, not significant; ^*** p < 0.001 and ^**** p < 0.0001).
2.4. In Vitro Assessment of the Barrier Protection Functions of the SF‐MA
Hydrogel Phase and In Vitro Assessment of the Pro‐Osteogenic Activity of the
CS‐MA Hydrogel Phase
Figure [118]S3a,b (Supporting Information) showed that the SF‐MA
hydrogel phase was mildly hydrophobic, which made it difficult for
bacteria and cells to form local adhesion sites, as the air layer
formed on the surface kept out bacteria.^[ [119]^38 , [120]^39 ^]
Subsequently, we measured the surface zeta potential of the SF‐MA
hydrogel. We found that the surface potential of the SF‐MA hydrogel
decreased significantly with increasing concentration (Figure [121]S3c,
Supporting Information), factors that are unfavorable for bacterial
adhesion. It was reported that SF has low immunogenicity and causes
mild inflammation during the initial stages of trauma repair, favoring
the destruction of pathogens present at the site of injury.^[ [122]^40
^] As the predominant etiological microbe implicated in oral
infections, Staphylococcus aureus (S. aureus) and Escherichia coli
(E.coli) were selected for in vitro experiments. The results showed
that the activity and proliferation of both S. aureus and E. coli were
inhibited with decreasing surface zeta potential when co‐cultured with
the SF‐MA hydrogel (Figure [123]S3d,e, Supporting Information). As
shown in Figures [124]4a and [125]S3f (Supporting Information), the
bacterial colony‐forming units of S. aureus and E. coli decreased with
decreasing surface potential, as assessed by confocal laser scanning
microscope (CLSM). With the extension of co‐culture time, the number of
bacteria on SF‐MA increased, but it must be noted that the proportion
of dead bacteria also increased (Figure [126]4b,c; Figure [127]S3g,h,
Supporting Information). Biofilm formation is a key mechanism for
bacteria to resist harsh environments and enable drug resistance.^[
[128]^41 ^] We therefore assessed biofilm formation by crystal violet
staining and the same trend was confirmed (Figure [129]4d; Figure
[130]S3i,j, Supporting Information). In brief, the above results
suggested that the dense SF phase was effective in resisting bacterial
adhesion. To verify the feasibility of in vivo applications of the
integrated osteoconductive Janus hydrogel, we first conducted
biocompatibility assays by culturing rBMSCs in the CS‐MA hydrogel and
human gingival fibroblasts (HGFs) in the SF‐MA hydrogel. The cell
viability was assessed by CCK‐8 assay and the results showed that all
groups of hydrogels exhibited biocompatibility, with rBMSCs in the
CS‐MA hydrogels and HGFs in the SF‐MA hydrogel proliferating normally
when cultured for 1, 3, 7 and 14 days (Figure [131]S4a, Supporting
Information). Moreover, for clinical GBR treatment, a protective
barrier to prevent fibroblasts infiltration is of great significance
for occupying the bone defect space and preventing fibroblast
infiltration from the surrounding soft tissue.^[ [132]^42 , [133]^43 ,
[134]^44 ^] Considering the mechanical properties and microporous
structure of the SF‐MA hydrogel, we next assessed the function of
preventing fibroblast infiltration within a simulated in vivo
environment by culturing HGFs on the surface of the SF‐MA hydrogel. The
results showed that HGFs hardly penetrated the SF‐MA hydrogel with a
growth depth of only 12 µm after 14 days of culture (Figure [135]4e,f;
Figure [136]S4b,c, Supporting Information). For comparison, we also
constructed a hydrogel scaffold with a GelMA‐SF‐MA‐GelMA sandwich
structure, and cultured HGFs on the surface of the GelMA hydrogel. HGFs
gradually infiltrated into the GelMA hydrogel, and the depth of growth
reached 38 µm after 14 days, which further proved that the SF‐MA
hydrogel had the effective function of preventing fibroblasts
infiltration (Figure [137]4g). Hence, our data demonstrated that the
SF‐MA hydrogel phase exhibited a protective barrier function, which
provided structural and biological protection for the bone regeneration
process.
Figure 4.
Figure 4
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In vitro barrier protection functions of the SF‐MA hydrogel phase and
in vitro pro‐osteogenic activity of the CS‐MA hydrogel phase. a) The
representative live & dead staining images of S. aureus and E.coli
after 24 h of co‐culture with SF‐MA. b) Quantification of the
fluorescence area based on the live/dead fluorescence staining images
of S. aureus. c) Quantification of the fluorescence area based on the
live/dead fluorescence staining images of E.coli. d)
Semi‐quantification of crystal violet staining after 48 h of co‐culture
with S. aureus and E.coli. e) Representative immunocytochemical
staining images of gingival fibroblasts cultured on the SF hydrogel and
GelMA‐SF‐MA‐GelMA sandwich structure for 1, 7 and 14 days. The red
arrow indicated that the gingival fibroblasts penetrated through GelMA
to the surface of the SF hydrogel. f,g) Quantitative analysis of the
growth depth of gingival fibroblasts cultured on the. SF hydrogel and
GelMA‐SF‐MA‐ GelMA sandwich structure for 1, 7 and 14 days (n = 3). h)
RT‐qPCR quantification of genes related to osteogenic differentiation
(Runx2, Bmp2 and Ocn, Col1al) in rBMSCs cultured on LCS, HCS and GelMA
for 3 days (n = 3). i) RT‐qPCR quantification of genes related to
osteogenic differentiation (Runx2, Bmp2 and Ocn, Col1al) in rBMSCs
cultured on LCS, HCS and GelMA for 7 days (n = 3). j) Representative
immunofluorescence images of osteogenic differentiation protein (RUNX2
green), actin network (Phalloidin, red), and cell nuclei (DAPI, blue)
in rBMSCs cultured on the LCS and HCS for 3 days. k) Representative
immunocytochemical staining images of osteogenic marker protein (BMP2
green), actin network (Phalloidin, red), and cell nuclei (DAPI, blue)
in rBMSCs cultured on the LCS and HCS for 7 days. l,m) The mean
fluorescence intensities were calculated to evaluate protein (RUNX2
and BMP2) expression levels (n = 3). n) Western blot analysis of RUNX2,
BMP2 and OCN in rBMSCs cultured on LCS, HCS and GelMA for 7 days. Error
bars represent the standard error of the mean. (ns, not significant; ^*
p < 0.05, ^** p < 0.01, ^*** p < 0.001 and ^**** p < 0.0001).
To assess the pro‐osteogenic functions of the LCS hydrogel and HCS
hydrogel on rBMSCs, we co‐cultured rBMSCs with hydrogels for 3 and 7
days and measured the expression of osteogenesis‐related genes (Col1al,
Ocn, Bmp2, Runx2) by RT‐qPCR. Compared with the control group,
osteogenesis‐related genes were upregulated in cells cultured in both
the LCS hydrogel and HCS hydrogel (Figure [139]4h,i). Notably,
transcription levels of these genes were much higher in the HCS groups,
thus suggesting the significant stimulatory role of high concentrations
of CS within the hydrogels in promoting osteogenesis.
Immunofluorescence staining (RUNX2 and BMP2) was performed to analyze
the localization and quantity of osteogenic markers in rBMSCs on day 7.
As demonstrated in Figure [140]4j,k, upregulation of RUNX2 and BMP2 was
observed in both the LCS hydrogel and HCS hydrogel, within the nucleus
and cytoplasm, respectively, indicating that the main switch of
osteogenic differentiation was turned on, especially in the HCS
hydrogel. The quantitative analyses of RUNX2 and BMP2 protein
expression shown in Figure [141]4l,m also confirmed the trend. Western
blotting further confirmed the enhancing effects of LCS and HCS on the
osteogenic differentiation of rBMSCs, with upregulated expression of
early osteogenic differentiation‐related markers RUNX2, BMP2, and OCN
in rBMSCs after 7 days of culture (Figure [142]4n). Taken together, our
results revealed that the CS‐MA hydrogel phase can activate osteogenic
differentiation of rBMSCs in vitro.
2.5. Dynamic Adaptation of Material Degradation with New Bone Formation
The adaptive degradation of scaffold materials with new bone formation
is a key indicator in biodegradable material‐mediated bone
regeneration.^[ [143]^45 , [144]^46 , [145]^47 ^] To further evaluate
the degradation behavior of the integrated osteoconductive Janus
hydrogel in vivo, we labeled the hydrogels with fluorescent dyes and
implanted the scaffold into the calvarial defect of mice (Figure
[146]5a). The first stage of bone defect reconstruction is the growth
of periosteum. The complete periosteum covering the defect area is
helpful in preventing the ingrowth of gingival fibrous tissues, thereby
preserving space for new bone generation.^[ [147]^48 , [148]^49 ^]
Before the periosteum is fully repaired, the SF‐MA hydrogel phase needs
to function as a barrier to protect the bone defect from soft tissue
ingrowth. The SF‐MA hydrogel degraded slowly in vivo during the first
28 days, effectively protecting the bone defect area and promoting
periosteal repair, followed by being gradually degraded, with more than
50% remaining at 35 days (Figure [149]S5a,b, Supporting Information).
After the implantation of the materials, the integrated osteoconductive
Janus hydrogel also started to display osteogenic repair function at
the same time. As shown in Figure [150]S5b (Supporting Information),
HCS‐SF and LCS‐SF were gradually degraded after implantation, and the
degradation rate of the HCS‐SF integrated Janus hydrogel was slower
than that of the LCS‐SF integrated Janus hydrogel, with HCS‐SF and
LCS‐SF degrading by 35.8% and 49.6% on the 28th day respectively. In
addition, the periosteum had formed and covered the entire defect area
by 28 days, and new bone had been generated in the bone defect area,
while the integrated osteoconductive Janus hydrogel had been partially
degraded based on micro‐CT scanning and histological analysis (Figure
[151]S5c, Supporting Information). In contrast, the GelMA group was
completely degraded by 21 days with ineffective bone regeneration
(Figure [152]S5b, Supporting Information; Figure [153]5b–d,j), which
indicated that the integrated osteoconductive Janus hydrogel had a more
stable structure and could sustain a longer‐lasting pro‐osteogenic role
in vivo. Moreover, through analysis by synthesis of calvaria neogenesis
and material degradation at 4 weeks post‐implantation, we found that
the bone regeneration and material degradation rate matched better in
the HCS‐SF group, as compared with the GelMA control group
(Figure [154]5e,f). Although the in vivo degradation rate of materials
in the LCS‐SF group was slightly faster than that of bone regeneration,
it still supported bone growth during the early stages of bone
regeneration. At 12 weeks post‐implantation, the defects were almost
completely filled with new bone in the HCS‐SF, LCS‐SF and Bio‐Gide
groups (Figure [155]5g). The new bone volume/total volume (BV/TV) of
the group implanted with HCS‐SF was higher than the control groups
(Figure [156]5h) at 12 weeks post‐implantation. Bone mineral density
(BMD) was also measured and there was a significant difference between
the HCS‐SF group and the Bio‐Gide group at 12 weeks post‐implantation
(Figure [157]5i). These results thus suggested that the integrated
osteoconductive Janus hydrogel mainly played a role in osteogenic
stimulation during the early‐middle process of bone regeneration.
Subsequent H&E staining and Masson's trichrome staining were used to
histologically analyze the osteogenic process (Figure [158]5j; Figure
[159]S5d, Supporting Information). At 12 weeks post‐implantation,
HCS‐SF led to complete healing with flat and consecutive bone‐structure
formation that is characteristic of full bone maturation. Masson's
trichrome staining also revealed mature osteoid tissue after 12 weeks
of implantation of HCS‐SF. In contrast, a small amount of newly‐formed
bone was observed in the GelMA control group, without any complete and
contiguous healing with host tissues, with only fibrous tissues being
detected in the Blank group when observation time was extended to 12
weeks post‐implantation. These results thus indicated that the
coordination of the barrier function of the SF‐MA hydrogel phase and
the osteoconductive function of the CS‐MA hydrogel phase together with
adaptive degradation, contributed to the regeneration of the mouse
calvarial defect. In summary, the integrated osteoconductive Janus
hydrogel has a degradation rate compatible with bone regeneration,
which promoted healing of bone defects and subsequent bone maturation.
Figure 5.
Figure 5
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The integrated osteoconductive Janus hydrogel achieved adaptive
degradation of materials with new bone formation. a) Schematic
representation of adaptation between hydrogel degradation and bone
regeneration within the mouse calvarial defect area. b) Representative
micro‐CT images of mouse calvarial full‐thickness defects at 4 weeks
post‐implantation. Red dotted lines denote the boundaries between the
nascent bone and the host bone. c,d) Quantitative analysis of the bone
volume/ tissue volume and bone mineral density (BMD) at 4 weeks
post‐implantation (n = 4). e) Representative micro‐CT images of mouse
calvarial full‐thickness defects at 4 weeks post‐implantation and
degradation images of the integrated osteoconductive Janus hydrogel in
vivo at 4 weeks post‐implantation. f) Quantification analysis of the
degradation ratio of the integrated osteoconductive Janus hydrogel in
vivo and new bone formation ratio at 4 weeks post‐implantation (n = 4).
g) Representative micro‐CT images of mouse calvarial full‐thickness
defects at 12 weeks post‐implantation. The red dotted lines denote the
boundaries between the nascent bone and the host bone. h,i)
Quantitative analysis of bone volume/ tissue volume and bone mineral
density (BMD) at 12 weeks post‐implantation (n = 4). j) H&E staining
and Masson's trichrome staining of histological sections at 4 & 12
weeks after implantation. (FT, fibrous tissue; NB, nascent bone; OT,
osteoid tissue; MT, mineralized tissue; M, residual materials; P,
periosteum). Error bars represent the standard error of the mean. (ns,
not significant; ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001 and ^**** p
< 0.0001).
2.6. The Integrated Osteoconductive Janus Hydrogel Enhanced Rat Periodontal
Bone Regeneration
To further investigate the osteoconductive functions of the integrated
osteoconductive Janus hydrogel for clinical GBR therapy, a rat
mandibular complete periodontal defect model (with a 3 mm × 1 mm‐sized
defect) was established (Figure [161]6a). Rats treated with Bio‐Gide
and Bio‐Oss (Bio) were assigned as controls; while untreated rats were
assigned as the Blank group. Through micro‐CT and histological
analysis, we found that there was almost no new bone formation in the
Blank group, at 4 weeks and even 12 weeks post‐implantation (Figure
[162]S6a, Supporting Information; Figure [163]6b), thus indicating the
weak self‐regenerative capacity of periodontal tissue. At 4 weeks after
implantation, relatively contiguous and intact newly‐formed bone in the
periodontal defect area was observed in the HCS‐SF group (Figure
[164]S6a, Supporting Information). In contrast, only a small amount of
new bone was formed in the Bio group with much Bio‐Oss residues (Figure
[165]S6a, Supporting Information). The results presented in Figure
[166]S6b–e (Supporting Information) indicated that the bone volume and
mean thickness of the newly formed bone were significantly greater in
the HCS‐SF group versus other groups. The abovementioned results thus
indicated that the integrated osteoconductive Janus hydrogel could
promote stem cell osteogenic differentiation, which was consistent with
the in vitro cell regulation results.
Figure 6.
Figure 6
[167]Open in a new tab
The integrated osteoconductive Janus hydrogel enhanced rat periodontal
bone regeneration. a) Schematic representation of the integrated
osteoconductive Janus hydrogel implantation within the rat periodontal
defect area. b) Representative micro‐CT images of rat periodontal
defects at 12 weeks post‐implantation. Red dotted lines denote the
boundary between nascent bone and host bone. The green fake color
denotes nascent bone. H&E staining and Masson's trichrome staining of
histological sections at 12 weeks after post‐implantation. The parts
circled by the yellow dotted line denote the new bone in the defect
area. (FT, fibrous tissue; NB: nascent bone; OT, osteoid tissue; MT,
mineralized tissue; MC, marrow cavity; D, dentin; PM, periodontium).
c–f) Quantitative analysis of bone volume/ tissue volume, bone mineral
density (BMD), bone thickness and bone coverage at 12 weeks
post‐implantation (n = 5). g) Immunofluorescence images of OCN (Green)
and NF200 (Pink) expression within the rat mandible defect after
implantation of the integrated osteoconductive Janus hydrogel at 12
weeks post‐implantation. DAPI stained cell nuclei (Blue). h,i) The mean
fluorescence intensities were calculated to evaluate protein expression
levels (n = 5). j) Osteogenesis efficiency of HCS‐SF hydrogel (red
asterisk), as compared with hydrogel materials (purple circles),
scaffold materials (green triangles) and membrane materials (orange
rhomboids) within the mouse calvarial defect model and rat periodontal
defect model. Materials are classified according to the material
morphology. The osteogenesis efficiency is represented by the ratio of
bone volume to the total volume (BV/TV). Details and values of the
aforementioned materials are listed in Table [168]S1 (Supporting
Information). Error bars represent the standard error of the mean. (ns,
not significant; ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001 and ^**** p
< 0.0001).
To achieve complete osseous tissue regeneration, the implantation time
was extended to 12 weeks. The micro‐CT results showed that the
newly‐formed jaw bone in the HCS‐SF group was similar to the natural
jaw bone, without an obvious boundary between the defect area and tooth
area (Figure [169]6b). In marked contrast, abundant high‐density
undegraded materials were observed in the Bio group (Figure [170]6b).
The results presented in Figure [171]6c–f indicated that the bone
volume, mean thickness, and coverage of newly‐formed bone were
significantly greater in the HCS‐SF group than in the other groups and
close to complete healing at 12 weeks post‐implantation. The
microstructure of the regenerated osseous tissues after 12 weeks
post‐implantation was further evaluated with H&E and Masson's trichrome
staining (Figure [172]6b). The results showed that HCS‐SF implantation
led to complete healing with contiguous bone‐structure formation at
bone maturity without any hydrogel residue. In contrast, in the Bio
groups, a small amount of newly‐formed bone was observed, but there
were still a lot of Bio‐Oss residues. Masson's trichrome staining
showed that the mature bone in the defect area of the HCS‐SF group was
more than that in the Bio and Blank groups. These results thus
demonstrated that complete regeneration of the jaw bone could be
achieved through HCS‐SF integrated osteoconductive Janus hydrogel
treatment. Furthermore, the canonical osteogenic differentiation marker
OCN, vascular differentiation marker CD31, and early neural
differentiation marker PAX6, were all detectable by immunocytochemical
staining at 4 weeks post‐implantation, with the confocal microscopy
images indicating that HCS‐SF promoted osteogenic differentiation,
vascular differentiation, and neural differentiation at the same time
(Figure [173]S6f–i, Supporting Information). Additionally, as compared
with the control groups, OCN and the mature neural differentiation
marker NF200, were highly expressed in the HCS‐SF group at 12 weeks
post‐implantation (Figure [174]6g–i), which validated mineralization of
bone extracellular matrix and neural maturation respectively. Numerous
studies have been conducted to develop a diverse array of materials for
facilitating bone defect repair, which can induce bone regeneration to
varying degrees. Compared with the performance of previous implant
materials such as hydrogels and scaffolds, HCS‐SF exhibited higher
osteogenic efficiency in the mouse calvarial defect model and in the
rat periodontal defect model, increasing by 42% and 13.7% respectively,
thus demonstrating that the integrated osteoconductive Janus hydrogel
can achieve synergistic osteogenic effects for clinical applications
under normal conditions. (Figure [175]6j; Table [176]S1, Supporting
Information). Additionally, an infected rat periodontal defect model
was employed to further validate the effects of HCS‐SF on bone
regeneration under pathological conditions^[ [177]^50 ^] (Figure
[178]7a). After 4 weeks of implantation, the micro‐CT data clearly
showed increased mass and improved parameters of new bone formation in
the HCS‐SF versus Blank group (Figure [179]7b–d). Notably, the
regenerated bone within the Blank group of the infected periodontal
defect model was much less than that in the normal healthy model,
confirming that the defect site was indeed infected, which inhibited
bone regeneration (Figure [180]7e–j). In sharp contrast, the amount of
newly formed bone in the HCS‐SF group of the infected periodontal
defect model was at the same level as that in the normal healthy model,
suggesting that HCS‐SF can effectively impede bacterial infection and
provide a conducive microenvironment for bone regeneration (Figure
[181]S7a–f, Supporting Information). Moreover, SF and CS used in the
preparation of the integrated osteoconductive Janus hydrogel are
biocompatible and have been approved by FDA for biomedical
applications. Hence, these results demonstrated that the integrated
osteoconductive Janus hydrogel is an effective implant biomaterial with
broad clinical prospects for facilitating craniomaxillofacial bone
repair due to its technical simplicity and convenience of application.
Figure 7.
Figure 7
[182]Open in a new tab
The integrated osteoconductive Janus hydrogel enhanced infected rat
periodontal bone regeneration. a) Schematic representation showing the
creation of rat periodontal defects to assess in vivo osteogenic
ability under pathological conditions. b) Representative micro‐CT
images of infected rat periodontal defects at 4 weeks
post‐implantation. Red dotted lines denote the boundary between nascent
bone and host bone. c) Quantitative analysis of bone volume/ tissue
volume (BV/TV) at 4 weeks post‐implantation (n = 7). d) Quantitative
analysis of bone mineral density (BMD) at 4 weeks post‐implantation (n
= 7). e) Representative micro‐CT images of infected rat periodontal
defects at 8 weeks post‐implantation. Red dotted lines denote the
boundary between nascent bone and host bone. f) Quantitative analysis
of bone volume/ tissue volume (BV/TV) at 8 weeks post‐implantation (n =
7). g) Quantitative analysis of bone mineral density (BMD) at 8 weeks
post‐implantation (n = 7). h) Representative micro‐CT images of
infected rat periodontal defects at 12 weeks post‐implantation. Red
dotted lines denote the boundary between nascent bone and host bone. i)
Quantitative analysis of bone volume/ tissue volume (BV/TV) at 12 weeks
post‐implantation (n = 7). j) Quantitative analysis of bone mineral
density (BMD) at 12 weeks post‐implantation (n = 7). (ns, not
significant; ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001 and ^**** p <
0.0001).
2.7. The Integrated Osteoconductive Janus Hydrogel Promotes Osteogenesis by
Regulating Lipid Metabolism
To further elucidate the the underlying mechanisms driving osteogenesis
within the HCS‐SF integrated osteoconductive Janus hydrogel upon
implantation in situ, we collected healing samples from the rat
periodontal defect for RNA‐seq analysis, at 1 and 4 weeks
post‐implantation. As expected, we found that many genes associated
with tissue regeneration were upregulated in the differential gene
expression volcano map and heat map (Figures [183]S8a,b and [184]S9a,b,
Supporting Information). The Gene Ontology (GO) enrichment analysis and
the Kyoto Encyclopedia Genes and Genomes (KEGG) enrichment analysis of
the functions of differentially expressed genes indicated that the
HCS‐SF mainly had an impact on lipid metabolism and tissue
regeneration‐related pathways, such as “extracellular matrix
organization”, “positive regulation of MSC proliferation” and
“ECM‐receptor interaction” at 1 week post‐implantation, and
“collagen‐containing extracellular matrix”, “PI3K‐Akt signaling
pathway” and “Hippo signaling pathway” at 4 weeks post‐implantation
(Figure [185]S8c–e, Supporting Information; Figure [186]8a–c). The GSEA
analysis of the transcriptional results at 1 week post‐implantation
mainly focused on cell proliferation and protein synthesis (Figure
[187]S8f, Supporting Information). The gene expression heatmap showed
that the levels of Col8a1, Itgb7, and Comp were enhanced after 1 week
of treatment with HCS‐SF (Figure [188]S8g, Supporting Information).
Moreover, the GSEA of the RNA‐Seq results revealed that genes
associated with tissue regeneration including “Wnt signaling pathway
and pluripotency” and “collagen fibril organization” were enriched in
the HCS‐SF group at 4 weeks post‐implantation (Figure [189]8d). The
gene expression heatmap showed that the mRNA levels of Wnt3, Bmpr2, and
Yap1 were increased after 4‐weeks of treatment. These signaling
pathways are crucial for regulating cell proliferation,
differentiation, and migration, which together constituted a complex
cascade of biological processes involved in tissue repair and
regeneration.
Figure 8.
Figure 8
[190]Open in a new tab
The integrated osteoconductive Janus hydrogel promoted osteogenesis at
4 weeks post‐implantation by regulating lipid metabolism. a) The
enriched GO terms of upregulated genes in the HCS‐SF group compared
with the Blank group. The red spectrum denotes osteogenesis‐related
biological processes. b) Representative KEGG pathways of significant
DEGs from the HCS‐SF group versus the Blank group (p ≤ 0.05, |log fold
change| ≥1). c) Enrichment of metabolically related KEGG pathways by
RNA‐seq analysis. d) GSEA analysis of significant DEGs in the HCS‐SF
and Blank groups. e) Gene cluster analysis of osteogenesis‐related
genes in the HCS‐SF and Blank groups. f) Circle plot of differential
metabolite by KEGG enrichment analysis. g) Circle plot of
metabolism‐related pathways by KEGG enrichment analysis.
In addition, RNA‐seq analysis suggested a role for metabolic processes
in the mechanism of bone regeneration promoted by HCS‐SF. To further
explore the characteristics of metabolites in this process, we used
LC‐MS/MS technology to analyze various metabolic components and
enrichment pathways. Clustered heatmaps showed significant differences
in metabolite levels between the HCS‐SF and Blank groups at both 1 and
4 weeks post‐implantation (Figures [191]S8h and [192]S9c, Supporting
Information). KEGG enrichment analysis of differential metabolite
levels showed that metabolic pathways were significantly enriched in
the HCS‐SF group (Figure [193]S8i, Supporting Information;
Figure [194]8f). Further analysis of metabolic pathways indicated that
lipid metabolism accounted for the largest proportion among all
metabolic pathways at both 1 week and 4 weeks post‐implantation (Figure
[195]S8j, Supporting Information; Figure [196]8g), which was consistent
with the enrichment results of differentially expressed genes in the
RNA‐seq analysis, thus revealing the key role of lipid metabolism in
HCS‐SF‐mediated bone regeneration. Previous studies have shown that the
regulation of lipid metabolism contributed to osteogenesis by affecting
several biological processes, including cellular energy supply,
signaling, and cell growth and differentiation.^[ [197]^51 , [198]^52
^] Therefore, the HCS‐SF integrated osteoconductive Janus hydrogel may
promote rBMSCs osteogenic differentiation by modulating lipid
metabolism‐related enzyme molecules or transcription factors.
3. Conclusion
An integrated osteoconductive Janus hydrogel comprising of a SF‐MA
hydrogel phase and a CS‐MA hydrogel phase to promote intensive bone
ingrowth was fabricated by sequential photocuring. The SF‐MA hydrogel
phase with barrier protection function prevented the infiltration and
ingrowth of gingival fibroblasts, and inhibited S. aureus and E.coli
adhesion, thereby providing a relatively closed biological environment
for bone regeneration. Additionally, the CS‐MA hydrogel phase
orchestrated lipid metabolism and thus enhanced osteogenesis,
angiogenesis, and neurogenesis, thereby enhancing extensive bone
regeneration, with osteogenic efficiencies of more than 60% in both the
mouse calvarial defect and rat periodontal defect models. Furthermore,
this integrated osteoconductive Janus hydrogel achieved adaptive
material degradation and facilitated new bone formation with
considerable mechanical strength and early structural stability in
vivo, which might provide an innovative and well‐suited strategy for
bone regeneration therapies. Hence, by combining the advantages of
stability, applicability, and simple fabrication techniques, the
integrated osteoconductive Janus hydrogel has much promising potential
to provide an innovative and well‐suited strategy for bone repair,
thereby maximizing therapeutic efficacy in promoting osteogenesis.
4. Experimental Section
Synthesis of Chondroitin Sulfate‐Methacrylate (CS‐MA)
CS was dissolved in deionized water. After full dissolution, the
methacrylic anhydride (MA) was added dropwise into the CS solution. The
molar ratio of MA versus the hydroxyl groups of CS was 20‐fold. Then,
NaOH solution was carefully added to adjust the pH to ≈8. The reaction
solution was stirred in an ice bath for 24 h. After the reaction
period, the reaction mixture was subjected to dialysis against
deionized water to remove the remaining unreacted MA and any
by‐products. The CS‐MA was then lyophilized and stored at −20 °C.
Synthesis of Silk‐Methacrylate (SF‐MA)
SF‐MA solutions were prepared as previously described. Briefly, 5 g of
sliced cocoons were boiled in 1 L of 0.02 m Na[2]CO[3] solution for
30 min at 100 °C to remove the sericin, and then washed and stirred for
20 min with distilled water several times. Subsequently, the degummed
silk was dried at room temperature and then dissolved in 5 mL of 4.03 g
lithium bromide (LiBr) solution at 60 °C for 1 h. Immediately after the
SF was dissolved by LiBr, 0.3 mL (424 mm) of glycidyl methacrylate
(GMA) solution (Sigma–Aldrich, St. Louis, USA) was added to the mixture
with stirring at 300 rpm for 3 h at 60 °C to create a high‐yield
reaction between GMA and SF. Then, the resulting solution was
centrifuged and dialyzed against distilled water using Slide‐A‐Lyzer
dialysis cassettes (MWCOs of 14000 Da for methacrylated SF solutions)
for 3 days. Finally, those solutions were frozen at ‐20 °C for 12 h and
freeze‐dried for 48 h. The lyophilized SF‐MA powder was stored at
−20 °C before further use.
Fabrication of the Integrated Osteoconductive Janus Hydrogel
Photo‐curable SF‐MA hydrogels were fabricated as follows. Lyophilized
SF‐MA was dissolved in deionized water at a concentration of 30% (w/v),
and the photoinitiator lithium phenyl(2,4,6‐trimethylbenzoyl)
phosphinate (LAP) (0.2% w/v) (Tokyo Chemical Industry, Tokyo, Japan)
was added and mixed. The mixed solution was kept at 4 °C overnight to
ensure complete dissolution. The fabrication of photocurable CS‐MA
hydrogels was prepared as follows. Lyophilized CS‐MA was dissolved in
deionized water and formulated at concentrations of 5% (w/v) and 10%
(w/v), and the LAP (0.2% w/v) (Tokyo Chemical Industry, Tokyo, Japan)
was added and mixed. The mixed solution was then kept at 4 °C overnight
for full dissolution. After the material was completely dissolved,
250 µL of CS‐MA was added into a mold with a diameter of 10 mm, and
then subjected to photocuring via exposure to 30 mW cm^−2 UV light for
5 s at a distance of 1 cm using Architect SV003 (Regenovo). Then 100 µL
of SF‐MA was added, and then photocured via exposure to 30 mW cm^−2 UV
light for 30 s at a distance of 1 cm using Architect SV003 (Regenovo),
to obtain the integrated osteoconductive Janus hydrogel.
Characterization of the Integrated Structure
To determine the molecular structure, CS‐MA and SF‐MA were examined
through ^1H nuclear magnetic resonance (^1H‐NMR) at a frequency of
400 MHz using a Bruker DPX FT‐NMR spectrometer (9.4 T, Bruker, Germany)
and 700 µL of deuterium oxide (D[2]O, Sigma‐Aldrich) as the solvent per
5 mg of sample. The SF‐MA solution was filtered using a 0.45 µm filter
before analysis. To assess the injectability and photocuring property
of the hydrogel, different dyes were added into the precursor solution
of CS‐MA and SF‐MA and then the solution was added to syringes and
extruded through 26‐gauge needles (φ ≈ 260 µm) to observe the
injectability of the hydrogel. The hydrogel was photocured via exposure
to UV light for 30 s at a distance of 1 cm respectively. The gelation
state and the fabrication of the integrated osteoconductive Janus
structure were observed and imaged with a digital camera (Canon
camera). To compare the microstructure and pore characteristics among
the different hydrogels, field emission scanning electron microscopy
(FE‐SEM, S‐4800, HITACHI) was performed after the samples were
embrittled with liquid nitrogen, lyophilized, and gold‐coated. For
further observation of the interface structure, the CS‐MA solution and
SF‐MA solution were mixed with fluorescent dyes of different colors,
and the fabricated integrated osteoconductive Janus hydrogel was
examined with CLSM (Leica).
Mechanical Behavior, Swelling and Degradation of the Integrated
Osteoconductive Janus Hydrogel
Mechanical properties of these cylindrical hydrogels with dimensions of
5 mm in height and 10 mm in diameter were assessed using a universal
mechanical testing machine (WDW3020, China) with a 1 kN load cell at a
cross‐head speed of 5 mm min^−1. Compressive strength and compressive
modulus were determined from stress‐strain curves. Compressive strength
was the stress at which the sample breaks or the stress at a strain of
90% for highly ductile samples, which do not break until very high
strains. Compressive modulus was determined as the slope of the
stress‐strain curve within the initial linear region at low strains
(0–10%). Otherwise, the dynamic mechanical properties of different
hydrogel systems were assessed with a rheometer (TA Instruments‐waters
LLC), and all measurement plates were used with a constant gap (1 mm)
at room temperature. The loss modulus represents the viscous capacities
of the gel, and the resistance of the substance against deformation
under shear was measured by the elastic modulus. Cylindrical hydrogels
were individually weighted (S[0]) and incubated in PBS solution at
37 °C for 24 h. At predetermined intervals, the hydrogels were
carefully taken out, and the residual water on the hydrogel surface was
drained with filter papers and weighed (S[1]). The swelling percentage
(SP) was determined with Equation ([199]1).
[MATH: SP=S1S0×100% :MATH]
(1)
The in vitro degradation of hydrogels was evaluated in a modified
simulated body (SBF) solution (pH = 7.4) at 37 °C for 35 days. The SBF
solution was refreshed daily. At predetermined intervals, the residual
hydrogels were taken out from the solution, carefully washed with
deionized water, and weighed. The weight remaining ratio (WRR) was
determined with Equation ([200]2).
[MATH: WRR=WtW0×100% :MATH]
(2)
Where W[0] and W[t] are the weights of samples before and after
degradation for a specific duration time (t) respectively. To assess
the in vivo degradation rate of the hydrogel, the CS‐MA and SF‐MA
labeled with fluorescent protein were injected into the 3 mm diameter
calvarial defect of C57BL/6J mice. The in vivo degradation of the
hydrogel was tracked and quantified by the in vivo image system
(PerkinElmer).
Bacterial Adhesion‐Related Surface Property Tests
To further examine the properties related to bacterial adhesion, SF‐MA
hydrogels were subjected to water contact angle and surface zeta
potential analyses. The water contact angle was measured by a Kruss
DSC100 (Germany) instrument via a drop of 2 µL on each hydrogel.
Surface zeta potential was measured by a ZETASIZE NANO instrument. The
samples were dispersed in ultrapure water and then filtered through a
filter. The filtered samples were injected into a polystyrene cuvette
and the ZETASIZE NANO instrument automatically calculated the zeta
potential for electrophoretic migration.
Protective Barrier Function Assessment of the SF‐MA Hydrogel
The Gram‐positive bacteria (S. aureus, ATCC 25923) and Gram‐negative
bacteria (E.coli, ATCC 25922) were cultured on SF‐MA hydrogel at 37 °C
for 12 and 24 h respectively. To evaluate bacterial adhesion and
antibacterial activity, the bacteria were stained with SYT09 and PI
fluorescent staining solution (ThermoFisher Scientific) after
co‐culture for 15 min, followed by imaging with CLSM (Leica). MTT assay
was performed according to the manufacturer's protocol (Solarbio) for
the detection of bacterial proliferation. Quantitative analysis of
biofilm formation was carried out by crystal violet (Solarbio)
staining. Briefly, the samples were washed with PBS, stained with 0.1%
(w/v) crystal violet for 20 min, and then washed with deionized water.
The crystal violet granules were eluted with 33% (v/v) acetic acid and
the absorbance at 595 nm was measured. To evaluate the barrier function
of the SF‐MA hydrogel against HGFs, 20 µL aliquots of HGF suspension
were seeded onto the surface of each sample at each time point. After
culturing for 1, 7, and 14 days, cells on the hydrogels were stained
with 4′,6‐diamidino‐2‐phenylindole (DAPI; Sigma) and Phalloidin
(Sigma), and observed under CLSM.
Quantitative Real‐Time Polymerase Chain Reaction (RT‐PCR)
To investigate cell osteogenic differentiation on the integrated
osteoconductive Janus hydrogel in vitro, rBMSCs were cultured
three‐dimensionally onto the CS‐MA hydrogel for 3 and 7 days
separately. Quantitative RT‐PCR was applied to evaluate the expression
of osteogenic differentiation gene markers (BMP‐2, Runx2, Col1a1, and
OCN). Total RNA was extracted with Trizol reagent (Invitrogen) and
synthesis of cDNA was performed using SuperScript III One‐Step RT‐PCR
System with Platinum Taq High Fidelity (Invitrogen). Quantitative
RT‐PCR was performed on a 7500HT Fast Real‐Time PCR using SYBR Green
(Invitrogen). The primer sequences utilized for RT‐PCR were as follows:
Primers Forward Reverse
Rat‐Gapdh TCTCTGCTCCTCCCTGTTC ACACCGACCTTCACCATCT
Rat‐Runx‐2 CTTCCCAAAGCCAGAGCG CAGCGTCAACACCATCATTC
Rat‐Bmp‐2 GAAGCCAGGTGTCTCCA AGTCCACATACAAAGGGTG
Rat‐Col1a1 AGGCAACAGTCGATTCACC GTCCAAGGGAGCCACATC
Rat‐Ocn AGTCTGACAAAGCCTTC AAGCAGGGTTAAGCTCAC
[201]Open in a new tab
Biocompatibility Assessment
The prepared hydrogels were placed in the wells of standard 96‐well
culture plates, and 2 × 10^4 rBMSCs were seeded per well with LCS, HCS
or GelMA, and 2 × 10^4 HGFs were seeded per well with SF‐MA after
ethanol/UV sterilization, and allowed to grow for 14 days. Hydrogel
cytocompatibility was analyzed using a Cell Counting Kit 8 (CCK‐8)
assay, following the manufacturer's instructions (Bimake). The 96‐well
plate was incubated at 37 °C for 20 min in a 5% CO[2] incubator.
Immunocytochemistry
To investigate the biological effects of the CS‐MA hydrogel, CS‐MA was
dissolved and sterilized through a 0.45 µm filter. A condensed cell
suspension at a density of 2 × 10^6 rBMSCs was seeded into each sample.
Subsequently, the samples were slowly added to micro‐tissue 3D petri
dishes (Sigma) and photocured. The samples were cultured in an
incubator for 3 and 7 days, followed by rinsing with phosphate‐buffered
saline (PBS) and fixation with 4% (w/v) paraformaldehyde for 30 min at
room temperature. Then, the samples were permeabilized with 0.1% (w/v)
Triton X‐100 (diluted with PBS) for 10 min and then blocked with 3%
(v/v) bovine serum albumin (BSA; diluted with PBS) for 1 h at room
temperature. The permeabilization solution was removed and the samples
were rinsed with PBS for 5 min at room temperature. The samples were
then incubated respectively with the following primary antibodies in 5%
(w/v) BSA in PBS overnight at 4 °C: polyclonal rabbit anti‐RUNX‐2
(1:100; Abcam), polyclonal rabbit anti‐BMP2 (1:200; Abcam). After
thorough rinsing to remove excess antibodies, the cells were incubated
with the following secondary antibodies for 1 h in the dark: goat
anti‐rabbit IgG H&L Alexa Fluor 488 pre‐adsorbed (1:500; Abcam).
Phalloidin (Sigma) was used for cytoskeletal staining. Cell nuclei were
stained using 4′,6‐diamidino‐2‐phenylindole (DAPI; Sigma). Images were
captured using CLSM (Leica). To investigate the barrier effects of the
SF‐MA hydrogel, SF‐MA and GelMA were dissolved separately, and
sterilized through a 0.45 µm filter. GelMA and SF‐MA were added into
micro‐tissue 3D petri dishes, and a sandwich structure of
GelMA‐SF‐MA‐GelMA, as well as a simple SF‐MA monolayer structure, were
fabricated, which were then subjected to layered photocuring. A
condensed cell suspension at a density of 2 × 10^4 human gingival
fibroblasts was seeded into each sample and the samples were cultured
in an incubator for 1, 7, and 14 days. The samples were rinsed with
phosphate‐buffered saline (PBS) and fixed in 4% (w/v) paraformaldehyde
for 30 min at room temperature. Then, the samples were permeabilized
with 0.1% (w/v) Triton X‐100 (diluted with PBS) for 10 min and blocked
with 3% (w/v) bovine serum albumin (BSA; diluted with PBS) for 1 h at
room temperature. The permeabilization solution was removed and the
samples were rinsed with PBS for 5 min at room temperature. Blocking
with 3% (wv) BSA was used for minimizing non‐specific staining.
Phalloidin (Sigma) was used for cytoskeletal staining, while cell
nuclei were stained using DAPI (Sigma). Images were captured with CLSM
(Leica).
Western Blotting
Briefly, cell lysate proteins were harvested by RIPA Buffer (Thermo
Fisher Scientific), separated by 10% (w/v) SDS–polyacrylamide gel
electrophoresis, and then transferred to polyvinylidene difluoride
membranes and blocked in 5% (w/v) non‐fat milk. The blotted membranes
were separately probed with corresponding primary antibodies against
GAPDH (1:5000, RayAntibody), RUNX‐2 (1:1000, Abcam), BMP‐2 (1:500,
Affinity), or OCN (1:1000, Abcam) overnight at 4 °C. Then the blotted
membranes were washed three times in TBS with 0.1% (v/v) Tween‐20,
incubated with a HRP‐conjugated secondary antibody for 1 h, and imaged
with an Odyssey Imaging System. Quantitative analysis was performed
with the Image J software.
Animals and Surgical Procedures
The experimental protocol was approved by the Animal Care and Use
Committee of Peking University (IACUC number: LA2022604). All the
Sprague‐Dawley rats (280–310 g) were randomly divided into three
groups: HCS‐SF hydrogel (HCS‐SF), Bio‐Oss+Bio‐Gide (Bio), and Blank (n
= 5 for each group). After anesthesia via intravenous injection of 1%
(w/v) pentobarbital sodium (1 mg kg^−1), a 3 mm diameter periodontal
defect was created using a saline‐cooled trephine drill and a 3 mm
outer diameter treble. Then, the materials of each group were implanted
into the defects and the CS‐MA hydrogel and SF‐MA hydrogel were
photocured with UV light for 5 and 30 s respectively. The wound was
closed by suturing the muscle and the skin layer by layer in the normal
healthy model, while those in the infected model were sutured after
inoculation with S. aureus suspension (2 µL, 10^7 CFU·mL^−1) was added
onto the surface of the SF‐MA hydrogel. For establishing the calvarial
defect model, all the C57BL/6J mice (20–25 g) were randomly divided
into five groups: HCS‐SF hydrogel (HCS‐SF), LCS‐SF hydrogel (LCS‐SF),
Bio‐Oss+Bio‐Gide (Bio), GelMA, and Blank (n = 5 for each group). The
mice were anesthetized via intravenous injection of 1% (w/v)
pentobarbital sodium (1 mg kg^−1) and then two bone defects (3 mm
diameter) were prepared in each mouse. Materials were randomly injected
into each calvarial defect and then photocured.
Micro‐CT Scanning Evaluation
At 4, 8, and 12 weeks postimplantation, mandible samples and calvaria
samples were harvested and fixed in 4% (w/v) paraformaldehyde for 24 h
at 4 °C, and the specimens were examined using micro‐CT scanning. After
3D visualization, bone morphometric analyses, including calculation of
BV/TV and BMD measurements, were carried out on the region of interest.
Histological Analysis
Briefly, tissue samples were fixed in 10% (w/v) neutral buffered
formalin for 7 days, decalcified and dehydrated according to standard
protocols, embedded in paraffin, and sectioned at 5 µm thickness. H&E
staining and Masson's trichrome staining were performed separately on
tissue sections, according to the manufacturer's protocols, and images
were captured under a light microscope (CX21, Olympus, Japan). CD31,
OCN, NF200, and PAX6 expression and distribution were observed using
immunocytochemical staining.
RNA Sequencing, LC‐MS/MS Sequencing and Analysis
Periodontal bone defects were created in rats for RNA‐Seq and LC‐MS/MS
analysis. The integrated osteoconductive Janus HCS‐SF hydrogel was
injected to fill into the defects of the HCS‐SF group. Subsequently,
the periodontal bone defects were sampled at 1 and 4 weeks
post‐implantation. For the RNA‐seq analysis, total RNA was extracted
using the Trizol reagent (Invitrogen, CA, USA) according to the
manufacturer's protocol. RNA purity and quantification were evaluated
using the NanoDrop 2000 spectrophotometer (Thermo Scientific, USA). The
RNA integrity was assessed using the Agilent 2100 Bioanalyzer (Agilent
Technologies, Santa Clara, CA, USA). Then the libraries were
constructed using the VAHTS Universal V6 RNA‐seq Library Prep Kit
according to the manufacturer's instructions. This was followed by
sequencing of the libraries on an Illumina Novaseq 6000 platform and
150 bp paired‐end reads were generated. All genes with |log fold
change|>1 and a p‐value of <0.05 were selected as DEGs. Based on the
hypergeometric distribution, GO, KEGG pathway, Reactome and
WikiPathways enrichment analysis of DEGs were performed to screen the
significant enriched term using R (v 3.2.0), respectively. The Gene Set
Enrichment Analysis (GSEA) was performed using the GSEA software. For
the LC‐MS/MS Sequencing, 30 mg of sample was added to a 1.5 mL
Eppendorf tube with 400 µL of methanol‐water (V:V = 4:1) as internal
standard. The mixtures were stored at −40 °C for 2 min and placed in a
grinder (60 Hz, 2 min). The whole samples were extracted by ultrasonic
treatment for 10 min in an ice‐water bath, and stored at −40 °C for
120 min. The extract was then centrifuged at 4 °C (13000 rpm) for
20 min. The supernatants (150 µL) from each tube were collected using
crystal syringes and transferred to LC vials. The vials were stored at
−80 °C until LC‐MS analysis. QC samples were prepared by mixing
aliquots of the samples to form a pooled sample. The metabolomic data
analytical instrument utilized was the ACQUITY UPLC I‐Class plus
instrument (Waters Corporation, Milford, USA). The original LC‐MS data
were processed by the Progenesis QI V2.3 (Nonlinear, Dynamics,
Newcastle, UK) software for baseline filtering, peak identification,
integration, retention time correction, peak alignment, and
normalization. A two‐tailed Student's T‐test was further used to verify
whether the differential metabolite levels between groups were
significantly different. Differential metabolites were selected with
p‐values less than 0.05 and were further used for KEGG pathway
enrichment analysis.
Statistical Analysis
All quantitative data were expressed as mean ± standard deviation (SD)
of at least three independent replicate experiments. The Shapiro‐Wilk
test was used to assess the normality of distributions. Statistical
analyses were carried out using One‐Way analysis of variance (ANOVA)
combined with the Student‐Newman‐Keuls (SNK) multiple comparison
post‐hoc test. The thresholds of statistical significance were set at
^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, and ^**** p < 0.0001.
Conflict of Interest
The authors declare no conflict of interest.
Supporting information
Supporting Information
[202]ADVS-12-e06736-s001.pdf^ (1.8MB, pdf)
Acknowledgements