Abstract
Articular cartilage defects are clinically prevalent yet lack effective
therapeutic solutions. Recent advancements in acellular cartilage
tissue engineering combined with microfracture techniques have shown
promising outcomes. Injectable hydrogels have emerged as particularly
attractive scaffolds due to their minimally invasive implantation and
capacity to conform to irregular cartilage defects. However, their
clinical application remains constrained by inadequate mechanical
strength and insufficient bioadhesion. In this study, we developed a
bioadhesive dynamic hydrogel by integrating catechol-functionalized
chitosan with aldehyde-terminated four-arm polyethylene glycol
(AF-PEG). When combined with KGN-loaded PLGA/PEG nanoparticles, this
hydrogel system enables sustained KGN release while maintaining
injectability, self-healing properties, and a 3D porous architecture.
Mechanical characterization revealed superior bioadhesion strength
(∼1,150 kPa) and compressive modulus (∼195 kPa). The hydrogel
demonstrated excellent biocompatibility, significantly promoting bone
marrow mesenchymal stem cells (BMSCs) proliferation, migration, and
chondrogenic differentiation in vitro. In vivo evaluations showed
superior ICRS and modified O’Driscoll histological scores in defects
treated with the KGN-loaded chitosan hydrogels compared to controls.
Histological analysis confirmed enriched type II collagen deposition in
newly formed cartilage, exhibiting structural organization and
integration with host cartilage comparable to natural tissue. This
novel KGN-loaded bioadhesive dynamic hydrogel provides an optimized
regenerative microenvironment for cartilage repair, demonstrating
substantial translational potential for clinical applications.
Keywords: catechol-modified chitosan, polyethylene glycol, bioadhesive
dynamic hydrogel, kartogenin, endogenous cartilage regeneration
Introduction
Articular cartilage defects caused by trauma, infection, or
osteoarthritis are very common and difficult to repair ([46]Campos et
al., 2019; [47]Richter et al., 2016). Current surgical approaches for
cartilage restoration primarily consist of osteochondral allografts,
autologous chondrocyte implantation, osteochondral autografts, and bone
marrow stimulation techniques such as microfracture and subchondral
drilling ([48]Gracitelli et al., 2016; [49]Sommerfeldt et al., 2016).
Among these interventions, microfracture-pioneered by Steadman in the
late 1990s-has been regarded by some clinicians as the historical gold
standard for treating focal cartilage defects, largely due to its
arthroscopic feasibility and cost-effectiveness ([50]Solheim et al.,
2018). This technique involves creating perforations in the subchondral
bone plate to facilitate the egress of bone marrow constituents,
ultimately generating a fibrocartilaginous repair tissue within the
defect. Numerous clinical studies have documented favorable short-term
clinical outcomes following microfracture ([51]Mithoefer et al., 2009).
However, the newly formed fibrocartilage gradually degenerates over
time, leading to suboptimal long-term outcomes. Additionally, the
microfracture technique struggles to repair large scale cartilage
defects. Therefore, by combining microfracture with cartilage tissue
engineering techniques, it is possible to create a favorable
microenvironment for the retention, proliferation, and chondrogenic
differentiation of locally recruited stem cells ([52]Case and Scopp,
2016; [53]Pueyo et al., 2025). This approach enhances the secretion of
cartilage extracellular matrix and holds promise for promoting
endogenous hyaline cartilage regeneration.
Chitosan (CS), as a natural polysaccharide, exhibits excellent
biocompatibility, making it widely applicable in cartilage tissue
engineering ([54]Comblain et al., 2017). Studies have demonstrated that
chitosan-based hydrogels possess a three-dimensional structure
analogous to the extracellular matrix (ECM) of cartilage, facilitating
the diffusion and exchange of nutrients and metabolic products
([55]Zhang et al., 2021a; [56]Zhang et al., 2021b; [57]Souza-Silva et
al., 2024). The chitosan backbone contains abundant amino groups, which
can dynamically bind with aldehyde groups under physiological
conditions to form reversible imine bonds, enabling the preparation of
dynamic hydrogels ([58]Xu et al., 2025). The breakage and reformation
of imine bonds endow hydrogels with injectability, viscoelastic
adaptability, and self-healing properties. These characteristics mimic
the biomechanical behavior of natural articular cartilage, thereby
promoting the proliferation of chondrocytes and the secretion of
cartilage matrix.
Among the numerous polymers containing aldehyde groups, polyethylene
glycol (PEG) derivatives decorated by aldehyde groups have become
commonly used crosslinking agents for preparing chitosan-based dynamic
hydrogels due to their excellent biocompatibility ([59]Andrade et al.,
2022; [60]Lin et al., 2024). Studies have reported that
dialdehyde-functionalized PEG (DF-PEG), used as a crosslinker, forms
dynamic hydrogels within 60 s when mixed with a chitosan solution,
exhibiting notable self-healing properties ([61]Wang et al., 2020).
Mohrman et al. developed chitosan/DF-PEG hydrogels that promoted
recovery in damaged central nervous systems ([62]Mohrman et al., 2018).
However, the suboptimal mechanical properties of these injectable
hydrogels hinder their application in cartilage defect repair.
[63]Huang et al. (2023) reported an injectable methacrylated
chitosan/PEG hydrogel, whose double-network system endows it with
excellent mechanical strength, making it suitable for the treatment of
intervertebral disc defects. Research indicates that
polyaldehyde-functionalized PEG offers more crosslinking sites,
resulting in hydrogels with superior gelation and mechanical properties
compared to dialdehyde-modified PEG when combined with chitosan
([64]Huang et al., 2016). Aldehyde-terminated four-arm PEG (AF-PEG) is
a star-shaped PEG that is obtained by covalently bonding
4-formylbenzoic acid to the hydroxyl groups at each branch end through
a carbodiimide coupling reaction. Due to its higher aldehyde group
density compared to dialdehyde-functionalized linear PEG, AF-PEG was
selected as the crosslinker in this study to prepare dynamic hydrogels
with chitosan.
The injectability of dynamic hydrogels is particularly advantageous for
repairing articular cartilage defects, as it allows minimally invasive
implantation and precise filling of defects with varying sizes and
geometries ([65]Bertsch et al., 2023). However, it is well known that
during clinical arthroscopic microfracture procedures, the
intra-articular environment presents a highly dynamic aqueous milieu
characterized by continuous irrigation with sterile saline and
transient bleeding from cartilage defects. Traditional hydrogels
struggle to achieve robust adhesion to surrounding cartilage under such
wet conditions, leading to hydrogel detachment and compromised
cartilage repair outcomes. Inspired by the exceptional underwater
adhesion of mussels, studies have revealed that catechol groups can
mediate interfacial adhesion in wet environments ([66]Guo et al.,
2020). Chitosan can be modified with catechol groups via carbodiimide
reactions, yielding chitosan-based polymers that mimic the structure of
mussel adhesive proteins, thereby exhibiting strong bioadhesive
efficacy ([67]Zhang et al., 2019). [68]Zheng et al. (2020) utilized
catechol-modified quaternized chitosan hydrogels to repair skin
defects, demonstrating their excellent dermal adhesion, promotion of
angiogenesis, and acceleration of wound healing. Furthermore, hydrogels
containing catechol groups can immobilize and trap cells and signaling
proteins from blood and bodily fluids after implantation into tissue
defects ([69]Souza Campelo et al., 2020). This enhances cellular
adhesion and further accelerates local tissue regeneration. Therefore,
this study aims to synthesize catechol-functionalized chitosan (CS-HCA)
through carbodiimide-mediated conjugation with hydrocaffeic acid (HCA),
followed by crosslinking with aldehyde-terminated four-arm polyethylene
glycol (AF-PEG) to develop a novel dynamic hydrogel with enhanced
bioadhesive properties. This strategy is expected to simultaneously
improve the adhesive and mechanical characteristics of chitosan-based
hydrogels for cartilage repair applications.
When the dynamic hydrogels adhere to the injured area, they can
gradually release their carried bioactive factors to alleviate local
inflammation, promote stem cell homing, and induce chondrogenic
differentiation, thereby promoting endogenous cartilage repair.
Commonly utilized bioactive factors include transforming growth
factor-beta (TGF-β), insulin-like growth factor-1 (IGF-1), exosomes,
and small-molecule drugs ([70]Olov et al., 2022). Among these,
Kartogenin (KGN), a non-protein small-molecule drug, induces
chondrogenic differentiation of bone marrow mesenchymal stem cells
(BMSCs) via the classical CBFβ-RUNX1 pathway ([71]Cai et al., 2019).
Studies demonstrate that KGN maintains its capacity to drive stem cell
chondrogenesis even under inflammatory conditions. KGN can also
maintain chondrocyte phenotype, alleviate joint inflammation, and
reduce cartilage matrix degradation ([72]Kwon et al., 2018).
Furthermore, KGN exhibits exceptional stability, enabling
room-temperature storage and delivery, and avoids receptor
downregulation or desensitization issues associated with prolonged
TGF-β use, highlighting its clinical potential. KGN has been
encapsulated in nanoparticles due to its hydrophobicity. Researches
also show that encapsulating KGN within poly (lactic acid-co-glycolic
acid) (PLGA) or PLGA-PEG nanoparticles achieves sustained release,
maintaining prolonged bioactivity to support continuous cartilage
regeneration ([73]Zhao et al., 2020; [74]Almeida et al., 2020).
Therefore, this study integrates KGN-loaded PLGA-PEG nanoparticles with
CS-HCA hydrogel to develop a bioadhesive dynamic hydrogel capable of
sustained KGN release. The hydrogel’s physical properties—including
injectability, self-healing capability, microstructure, adhesive
strength—and in vitro biocompatibility will be systematically
evaluated. Subsequently, a rat knee joint cartilage defect model will
be established to assess the hydrogel’s in vivo cartilage repair
efficacy.
Materials and methods
Materials and reagents
Chitosan (deacetylation degree >95%, Jinan Hedebio Marine Biotechnology
Co., Ltd., China); hydrocaffeic acid (HCA, China National
Pharmaceutical Group Chemical Reagent Co., Ltd., China);
1-(3-Dimethylaminopropyl)-3-ethylcarbodiimide (EDC, Acros Organics,
USA); N-hydroxysuccinimide (NHS, Acros Organics, USA);
aldehyde-terminated four-arm polyethylene glycol (AF-PEG, Beijing J&K
Scientific Technology Co., Ltd., China); Kartogenin (analytical grade,
Wuxi Jiehua Pharmaceutical Technology Co., Ltd., China); PEG (China
National Pharmaceutical Group Chemical Reagent Co., Ltd., China); PLGA
(China National Pharmaceutical Group Chemical Reagent Co., Ltd.,
China). Other chemicals and reagents are listed in the
[75]Supplementary Table S1.
Preparation and characterization of KGN-loaded PLGA-PEG nanoparticles
(PLGA-PEG@KGN)
The PLGA–PEG copolymer and PLGA-PEG nanoparticles were synthesized as
previously reported ([76]Almeida et al., 2020). The brief preparation
procedure is illustrated in [77]Figure 1A. 100 mg of PLGA-PEG was
dissolved in 5 mL of dichloromethane (DCM) under magnetic stirring at
500 rpm until complete dissolution. Subsequently, 20 mg of KGN was
added to the PLGA-PEG solution, followed by sonication for 5 min to
ensure homogeneous drug dispersion. The KGN-loaded PLGA-PEG solution
was slowly dripped into a polyvinyl alcohol (PVA) solution (1:5, v/v)
for primary emulsification under ultrasonication, with continuous
cooling in an ice bath. The solvent was rapidly removed using a rotary
evaporator (40°C, 100 rpm, and a vacuum pressure of −0.08 MPa) until
the emulsion transitioned from turbid to semi-transparent, indicating
complete nanoparticle solidification. The emulsion was then transferred
to centrifuge tubes and centrifuged at 15,000 rpm for 30 min to pellet
the nanoparticles, after which the supernatant was discarded. The
precipitate was resuspended in deionized water, and the centrifugation
process was repeated three times to thoroughly remove residual free PVA
and unencapsulated KGN. The washed nanoparticles were freeze-dried for
24–48 h and finally stored in airtight, light-protected containers at
−20°C.
FIGURE 1.
[78](A) Diagram of the synthesis process for PLGA-PEG/KGN
nanoparticles. (B) NMR spectra of PLGA-PEG and PLGA. (C) TEM image of
nanoparticles at 1 µm scale. (D) Close-up TEM image at 100 nm scale.
(E) Bar graph comparing mean size of nanoparticles. (F) Bar graph
comparing mean zeta potential of nanoparticles. (G) Calibration curve
of concentration versus integral area with a linear equation and
R-squared value. (H) Chromatogram depicting retention time. (I) UV-Vis
spectra of various concentrations of a sample. (J) Calibration curve of
concentration versus absorbance with a linear equation and R-squared
value.
[79]Open in a new tab
Characterization of PLGA-PEG nanoparticles. (A) Schematic diagram of
nanoparticle preparation process. (B) ^1H NMR spectra of PLGA and
PLGA-PEG. (C,D) TEM images of PLGA-PEG nanoparticles. (E) Size
distribution of PLGA-PEG and PLGA-PEG@KGN nanoparticles. (F) Surface
Zeta potential of PLGA-PEG and PLGA-PEG@KGN nanoparticles. (G,H)
Standard curve (G) and chromatogram (H) of PLGA-PEG@KGN nanoparticles
analyzed by high-performance liquid chromatography (HPLC). (I,J) UV
spectrophotometer analysis of nanoparticle encapsulation efficiency and
drug loading capacity. Full-wavelength scan (I) and standard curve (J)
of KGN.
Chemical structure of PLGA-PEG was examined by ^1H nuclear magnetic
resonance (^1H NMR) spectroscopy. The size distribution of the
nanoparticles was analyzed using dynamic light scattering (DLS). Zeta
potential was measured via electrophoretic light scattering (ELS).
Morphological characterization of the nanoparticles was conducted using
transmission electron microscopy (TEM). Drug encapsulation efficiency
and loading capacity were determined by ultraviolet-visible (UV-Vis)
spectrophotometry and High Performance Liquid Chromatography (HPLC).
Surface chemical properties of the nanoparticles were examined using
Fourier-transform infrared (FT-IR) spectroscopy. The in vitro release
behavior of KGN was evaluated through a dynamic dialysis method
monitored by UV-Vis spectrophotometry.
Preparation and characterization of catechol-functionalized chitosan polymer
(CS-HCA)
[80]Figure 2A illustrates the grafting of catechol groups onto chitosan
through an EDC/NHS-mediated conjugation reaction. (1) Dissolve 500 mg
of chitosan (CS) in 45.5 mL of deionized water (pH ∼1.6) under
continuous rapid stirring until a transparent light-yellow colloidal
solution is formed. (2) Adjust the pH of the colloidal solution to 5.4
by dropwise addition of NaOH solution, resulting in a turbid colloid.
Then, add 591 mg of hydrocaffeic acid (HCA) and continue stirring until
the colloid becomes transparent again (pH ∼3.86). (3) Dissolve
1,244.8 mg of EDC (1-ethyl-3-(3-dimethylaminopropyl)carbodiimide) and
746.75 mg of NHS (N-hydroxysuccinimide) in 50 mL of an
ethanol/deionized water mixture (1:1, v/v). Add this solution dropwise
to the CS-HCA colloid. Adjust the pH to 4.6 with 1M HCl under vigorous
stirring. (4) Dialyze the mixture using a regenerated cellulose
dialysis membrane (MWCO: 12–14 kDa) against deionized water (pH
3.0–3.5) for 48 h, followed by dialysis against deionized water (pH
5.0) for 4 h. Freeze-dry the purified product to obtain the dried
CS-HCA polymer. Store the final product under inert gas at −40°C.
FIGURE 2.
[81]Chemical synthesis and testing images of chitosan and
hydroxycinnamic acid (HCA) compounds. Panel A shows the chemical
reaction. Panel B displays UV-visible spectroscopy of CS, HCA, and
CS-HCA. Panel C presents FTIR spectra. Panel D features an NMR
spectrum. Panel E exhibits a chitosan sample. Panel F demonstrates
liquid reactions in test tubes. Panel G illustrates a self-healing test
on a material. Panel H shows a strain-stress graph. Panels I and J
compare mechanical properties before and after self-healing. Panels K
and L feature frequency sweep graphs of storage (G') and loss moduli
(G'').
[82]Open in a new tab
Characterization of CS-HCA hydrogels. (A) Synthesis route and chemical
structure of CS-HCA polymer. (B–D) UV-Vis (B) FT-IR (C) and ^1H NMR (D)
spectra of CS-HCA polymer. (E) Image of the injectability of hydrogel
at room temperature. (F) Images of the hydrogels adhesion to glass,
plastic and skin surfaces. (G) Image of the self-healing process of
hydrogel. (H) Adhesive strength of the hydrogels. (I,J) DMA results of
the CS-HCA hydrogels. Compressive (I) and tensile (J) moduli of the
hydrogels before fracture and after self-healing. (K,L) Rheological
analysis of CS-HCA (K) and CS-HCA@PLGA-PEG@KGN (L) hydrogels (G',
storage modulus, G'', loss modulus).
Confirm the successful synthesis of CS-HCA via ^1H NMR, FT-IR, and
UV-Vis spectroscopy.
Preparation of KGN-loaded CS-HCA bioadhesive dynamic hydrogel
(1) CS-HCA Solution Preparation: Weigh 500 mg of CS-HCA powder and
dissolve it in deionized water under continuous stirring to prepare a
1.5% (w/v) homogeneous solution. Filter the solution to obtain a
sterile formulation. (2) AF-PEG Solution Preparation: Weigh 500 mg of
AF-PEG powder and dissolve it in deionized water. Filter the solution
to ensure sterility. (3) Mix 5 mL of the CS-HCA solution with an equal
volume of AF-PEG solution thoroughly. Transfer the mixture into a
12-well plate for molding, resulting in the formation of CS-HCA
hydrogel. (4) Weigh 100 mg of PLGA-PEG@KGN nanoparticles, disperse them
in deionized water, and homogenize via stirring. Add the nanoparticle
suspension to the preformed hydrogel solution and mix rapidly until
uniformity is achieved. Mold the final composite in a 12-well plate to
obtain the KGN-loaded CS-HCA hydrogel (CS-HCA@PLGA-PEG@KGN). FT-IR
spectroscopy analysis was performed to investigate the interactions
between CS-HCA hydrogels and KGN nanoparticles.
Rheological characterization of CS-HCA hydrogels
A 5 wt% CS-HCA hydrogel was prepared in PBS buffer (pH 7.4), and its
viscoelastic properties were evaluated using an Anton Paar rheometer
equipped with a PP50 stainless steel plate (50 mm diameter). The
storage modulus (G′), representing the elastic component, and the loss
modulus (G″), representing the viscous component, were measured to
assess the hydrogel’s mechanical behavior. A hydrogel exhibits dominant
elastic properties (G' > G″) in the gel state, while viscous behavior
prevails (G'' > G′) in the sol state. The sol-gel transition point
occurs when G' = G″, indicating a balance between viscous and elastic
responses. For the frequency sweep test, the following parameters were
applied: a gap distance of 0.5 mm, angular frequency range of
0.1–100 rad/s, constant strain of 1%, temperature of 37°C, and a normal
force F[N] of 0 N.
Evaluation of hydrogel injectability, self-healing, and bioadhesive
properties
(1) Prepare a 5 wt% CS-HCA hydrogel solution. Aspirate 0.5 mL of the
sol into a 1 mL syringe and assess macroscopic injectability via
extrusion-based writing tests. (2) Cut a cylindrical hydrogel into
two-halves. Bring the fractured surfaces into contact and observe
interfacial integration to evaluate macroscopic self-healing
capability. (3) Apply the hydrogel to plastic, glass, and skin
substrates to assess its adhesive performance under varying surface
conditions.
Mechanical properties measurement
The stress/strain sweep of CS-HCA hydrogels before fracture and after
self-healing was performed using a dynamic mechanical analyzer (DMA).
Alternating amplitude strains (large strain to disrupt the gel
structure and small strain to observe structural recovery) were applied
to evaluate the self-healing properties of the hydrogel. The testing
procedure was as follows: the angular frequency was maintained at
10 rad/s, with alternating strains of 1% and 1,000%, the test
temperature was set at 37°C, and the normal stress (F[N]) was 0 N.
Characterization of hydrogel microporous structure
Prepare hydrogel solutions with solid contents of 1 wt%, 2 wt%, and
5 wt% in PBS (pH 7.4). Rapidly quench the gels in liquid nitrogen to
embrittle them, then fracture the samples with tweezers. Freeze-dry the
fractured gels, sputter-coat with gold, and examine cross-sectional
pore structures using scanning electron microscopy (SEM).
Determination of hydrogel porosity
Add anhydrous ethanol (V1) into a clean, dry graduated cylinder.
Submerge a pre-weighed dry hydrogel scaffold in the ethanol, ensuring
complete immersion. Apply negative pressure evacuation until the
scaffold is fully saturated (no bubbles emerge), and record the ethanol
volume (V2). Remove the scaffold and measure the residual ethanol
volume (V3). Calculate porosity using the formula:
[MATH: Porosity %=
V1−V3/
V2−V3×100%. :MATH]
All the experiments were repeated six times.
Adhesion strength measurement
Porcine skin specimens (5.0 cm × 1.5 cm) were cut and fixed onto
transparent glass slides (25.4 mm × 76.2 mm). Two porcine skin
specimens were adhered using the hydrogel solution and maintained at
room temperature for 1 h prior to testing. Specimens were stretched on
a universal testing machine equipped with a CMT 100N force sensor at a
crosshead speed of 5 mm/min under controlled conditions (26°C, 50%
humidity) until separation occurred. Maximum load and displacement were
recorded. Shear strength was calculated as the maximum load divided by
the overlapping contact area. All experiments were performed in
quintuplicate, and the mean values are reported.
In vitro degradation
The initial mass of the hydrogel scaffold was recorded as W0. The
hydrogel was placed into a centrifuge tube prefilled with an equal
volume of PBS (pH 7.4). Complete submersion of the hydrogel in PBS was
ensured, followed by incubation at 37°C for 1–4 weeks. At predetermined
weekly intervals, the hydrogel scaffolds were retrieved, surface
moisture was gently removed, and the remaining mass was recorded as W1.
The degradation rate (%) was calculated using the formula:
[MATH: degradation ratio %=
W0 −<
mrow> W1/W0×100%. :MATH]
Isolation and culture of bone marrow mesenchymal stem cells
All experimental protocols were approved by the Animal Experimental
Ethics Committee of Chongqing Western Biomedical Technology corporation
(No.20240415S0201231 [01]).
SD rats were used to extract BMSCs according to the method described in
the literature ([83]Maridas et al., 2018). The cells were seeded into
culture flasks at a density of 1 × 10^6 cells/mL and cultured in
DMEM/F12 medium supplemented with 10% fetal bovine serum (FBS) and
penicillin-streptomycin. Second-passage (P2) cells with good growth
status were selected for subsequent experiments. The cells were divided
into three groups: 2D conventional culture group (2D), CS/HCA hydrogel
group (CS/HCA), KGN-loaded CS/HCA hydrogel group (CS-HCA@PLGA-PEG@KGN).
The 2D group was cultured under standard 2D conditions, while the other
two groups were cultured on CS/HCA or CS-HCA@PLGA-PEG@KGN hydrogels
respectively.
In vitro BMSCs proliferation
The proliferation of BMSCs in each group was evaluated using the CCK-8
assay at 24, 48, and 72 h post-culture. After termination of
cultivation, cells were treated with trypsin, resuspended in culture
medium, and seeded into 96-well plates. Each well received 100 μL of
cell suspension supplemented with 10 μL of CCK-8 solution. Following
1–4 h of incubation, absorbance (A) at 450 nm was measured using a
microplate reader. Six replicates of each group were studied.
In vitro BMSCs migration
Cell migration ability was evaluated using the scratch assay and Boyden
Transwell chamber assay (24-well plates with polycarbonate membranes,
pore diameter 8.0 μm, membrane thickness 6.5 mm).
Scratch assay: BMSCs from each group were processed as planned,
adjusted to a density of 2 × 10^5 cells/mL, and seeded into 6-well
plates with 2 mL of cell suspension per well. When cell confluence
reached 80%–90%, the medium was aspirated. A sterile 10 μL pipette tip
was used to create a linear scratch perpendicular to the plate surface.
BMSCs were rinsed 3 times with PBS to remove debris, and serum-free
DMEM was added. Scratch closure was observed and quantified at 0h, 12h,
and 24h. ImageJ was used to analyze migration by measuring the initial
scratch area (S[0]) and the healed scratch area (St). Migration
percentage was calculated as:
[MATH: Migration %=
1−St
/S0×100% :MATH]
.
Transwell chamber assay: Cell suspensions were adjusted to 2 × 10^5
cells/mL. Transwell chambers were pre-equilibrated in serum-free DMEM
for 1 h. A total of 100 μL of the cell suspension was added to the
upper chamber, while 500 μL of DMEM containing 10% FBS was introduced
into the lower chamber. Chambers were incubated at 37°C with 5% CO[2]
for 24 h. After incubation, the Transwell chamber was removed, and
cells on the top side of the chamber were gently removed using cotton
swabs. Cells were fixed with 4% paraformaldehyde for 30 min at 37°C and
then washed three times with PBS. Finally, the cells were stained with
a 0.1% crystal violet solution. Afterward, images of five randomly
selected fields of view were captured under an inverted microscope, and
cell migration was quantified.
In vitro BMSCs chondrogenic differentiation
BMSC suspensions were seeded into 6-well plates at a density of 2 ×
10^5 cells/mL and cultured under the previously described grouping
conditions. Each group was supplemented with identical chondrogenic
differentiation induction medium (composed of DMEM/F12 basal medium
containing 50 μg/mL ascorbic acid, 100 nM dexamethasone, 1% ITS
additive, 100 mg/mL sodium pyruvate, 50 μg/mL proline, and 10 ng/mL
TGF-β3). After 7 days, the chondrogenic differentiation potential of
the BMSCs was assessed via qRT-PCR, Western blot, and Alcian blue
staining.
Total RNA was extracted from BMSCs using the RNAiso Plus Kit following
the manufacturer’s protocols. The reference gene was glyceraldehydes
3-phosphate dehydrogenase (GAPDH). cDNA synthesis was performed using
1 μg of total RNA with the RevertAid First Strand cDNA Synthesis Kit.
Quantitative real-time PCR (qRT-PCR) was conducted to analyze the
relative expression levels of chondrogenic markers (Sox9, Acan, and
Col2α1) using the 2^−ΔΔCT method. Primer sequences are detailed in
[84]supplementary Table S2. For protein-level validation, total
cellular proteins were extracted and quantified via Western blot
analysis. β-Actin served as an internal loading control for
normalization.
The cells from each group after treatment were fixed with 4%
paraformaldehyde (PFA), followed by Alcian blue staining for 30 min at
37°C. The cells were then observed by microscopy.
Construction of rat knee joint cartilage defect model
6–8-week-old SD rats (n = 18) were numbered and randomly divided into
three groups: Control group, CS/HCA hydrogel group (CS/HCA), KGN-loaded
CS/HCA hydrogel group (CS-HCA@PLGA-PEG@KGN). Rats were anesthetized
using pentobarbital sodium (0.2 g/mL), followed by shaving and
sterilization of the knee area. A midline incision was made over the
knee joint to expose the femoral trochlea. A full-thickness cartilage
defect (2.0 mm in diameter, 1.5 mm in depth) was created in the center
of the trochlea. After repeated saline irrigation, procedures varied
according to group assignment: the cartilage defect in the control
group was left untreated, and the incision was directly sutured. For
the two experimental groups, the defects were filled with either the
CS/HCA hydrogel or the CS-HCA@PLGA-PEG@KGN hydrogel. Postoperatively,
rats were individually housed with unrestricted knee joint movement.
The incision site was disinfected every 3 days.
Chondrogenic induction in vivo
At 6 weeks postoperatively, the animals were euthanized by intravenous
injection of an overdose of pentobarbital sodium, and the knee joints
were harvested. First, a digital camera was used to capture images of
the osteochondral defects. International Cartilage Repair Society
(ICRS) scoring system was used to score the defect site ([85]Van den
Borne et al., 2007). Three specimens were randomly selected from each
group for histological analysis. The specimens were soaked in 10%
formalin overnight, decalcified, embedded in paraffin, and sectioned
for staining. Hematoxylin and eosin (HE) and Collagen II
immunohistochemical staining were performed for histological analyses.
The sections were scored using a modified O’Driscoll histology scoring
system (MODHS) ([86]O'Driscoll et al., 2001).
Statistical analysis
Statistical analysis was performed using IBM SPSS software 19.0 (IBM
Corp., Armonk, NY, United States). Numerical data are expressed as mean
± standard deviation (M ± SD). One-way ANOVA was used for between-group
comparisons, with statistical significance defined as P < 0.05.
Results
Characterization of PLGA-PEG@KGN nanoparticles
The synthesis route of PLGA-PEG@KGN is shown in [87]Figure 1A, and the
chemical structure of PLGA-PEG was confirmed by ^1H NMR spectroscopy.
As shown in [88]Figure 1B, the PLGA spectrum exhibits a
characteristic–CH3 peak at 1.47 ppm and a CH3CHOCH proton peak in the
range of 5.19–5.21 ppm. After grafting PEG onto PLGA, a
distinct–CH2CH2O- proton peak from PEG is clearly observed at
3.47–3.51 ppm, confirming the successful synthesis of PLGA-PEG.
As shown in [89]Figures 1C–F, the fabricated PLGA-PEG nanoparticles are
spherical with an average diameter of 157.2 nm and a surface Zeta
potential of −1.7 mV. The PLGA-PEG@KGN nanoparticles exhibit an average
diameter of 343.8 nm and a surface Zeta potential of −13.0 mV. The peak
area obtained from the analysis of the sample using high-performance
liquid chromatography (HPLC) was substituted into the standard curve,
resulting in a calculated encapsulation efficiency of 88.86% for the
nanoparticles loaded with KGN ([90]Figures 1G,H). UV-Vis
spectrophotometry analysis ([91]Figures 1I,J) demonstrated an
encapsulation efficiency of 73.2%, which is relatively close to the
result obtained by HPLC. Additionally, the drug loading capacity of KGN
was calculated to be 6.82% based on UV-Vis spectrophotometry analysis.
Characterization of CS-HCA dynamic hydrogel
The synthesis route of CS-HCA polymer is shown in [92]Figure 2A. UV
spectroscopy analysis revealed that CS exhibited no significant
absorption peaks in the range of 200–500 nm. However, after HCA
grafting, distinct characteristic peaks of HCA appeared at 226 and
281 nm ([93]Figure 2B), corresponding to the phenolic structures in
HCA, thereby confirming the successful grafting of CS-HCA.
To verify structural integrity, FT-IR analysis was conducted on CS,
HCA, and CS-HCA ([94]Figure 2C). The CS spectrum originally lacking
benzene ring vibration peaks exhibited distinct benzene skeleton
vibrations and C=C harmonic peaks after HCA grafting, consistent with
UV findings. ^1H-NMR analysis ([95]Figure 2D) identified CS methyl
group (CH3-) at 1.94 ppm, methylene groups (-CH2-) at 3.34 ppm and
-CH2O- at 2.62 ppm. CS aromatic protons (H-1–6) appeared at 4.7, 2.91,
and 3.57–3.81 ppm, while HCA aromatic protons (H-7–9) showed multiplets
at 6.52–6.74 ppm. These spectral analyses collectively confirmed the
successful preparation of CS-HCA. The grafting rate of CS-HCA was
calculated to be 16.67% based on the ratio of the integral area of HCA
characteristic peak in ^1H-NMR to that of H[2] peak in CS, and the
actual yield was 58.33%.
As demonstrated in [96]Figures 2E,G, the CS-HCA hydrogel exhibits
injectability and self-healing properties. The hydrogel solution could
be injected via a syringe to form hydrogels of arbitrary shapes,
enabling seamless filling of irregular cartilage defects. When cut into
halves, the hydrogel exhibited self-healing capability upon contact
without visible cracks. Adhesion tests demonstrated that the hydrogel
adhered firmly to plastic, glass, and skin surfaces ([97]Figure 2F). In
porcine skin adhesion assays ([98]Figure 2H), the hydrogel exhibited
strong bioadhesion, maintaining structural integrity under 80% tensile
deformation with an adhesion strength of 1,150 kPa.
To further verify the self-healing properties of the hydrogel, we
conducted dynamic mechanical analysis. As demonstrated in [99]Figures
2I,J, the compressive modulus of the normally synthesized hydrogel
reaches ∼195 kPa, while that of the self-healed hydrogel after fracture
shows a slight decrease but still achieves ∼170 kPa, indicating its
retained strong compressive performance. In terms of tensile mechanical
properties, the tensile modulus of the intact hydrogel reaches ∼75 kPa,
whereas the self-healed hydrogel maintains a modulus of ∼70 kPa. These
results demonstrate that the CS-HCA hydrogel possesses self-healing
capability, and the healed hydrogel can still maintain robust
compressive and tensile properties.
To analyze the dynamic mechanical properties of the CS-HCA hydrogel, we
conducted rheological analysis. The frequency sweep test results
demonstrated that within the linear viscoelastic region, the storage
modulus (G′) of both the CS-HCA hydrogel and the CS-HCA@PLGA-PEG@KGN
hydrogel was consistently greater than the loss modulus (G″),
indicating a gel-like state. As shown in [100]Figures 2K,L, the
rheological properties of the CS-HCA hydrogel exhibited no significant
changes before and after loading KGN nanoparticles. The small
difference between G′ and G″ for both the CS-HCA hydrogel and the
CS-HCA@PLGA-PEG@KGN hydrogel suggests a certain degree of fluidity,
making them suitable for injectable therapies.
Microstructure of CS-HCA hydrogel
As shown in [101]Figure 3A, hydrogel samples formed at different
concentrations exhibited typical three-dimensional microporous
structures, with porosities ranging between 69% and 80%. The internal
structure displayed a heterogeneous but continuous pore network with
smooth surfaces, where pore diameters primarily ranged from 20 to
200 μm. Notably, the 2 wt% hydrogel exhibited slightly larger pore
sizes compared to the other two groups, whereas the 5 wt% hydrogel
demonstrated the highest porosity of approximately 79%. Nanoparticles
were successfully attached to the surfaces of the internal pores
([102]Figure 3B). This interconnected pore structure facilitates cell
adhesion, nutrient diffusion, and metabolic waste exchange.
FIGURE 3.
[103]Electron microscopy images (A1, A2, A3) show porous structures at
varying concentrations (1wt%, 2wt%, 5wt%). Image B shows a close-up of
a sample surface. Graphs C1, C2, C3 display FTIR spectra for different
materials. Graph D illustrates cumulative release percentages of
substances over time. Graph E shows degradation rates (%) over time for
the samples.
[104]Open in a new tab
(A1–A3) SEM images of CS-HCA hydrogels with concentrations of 1wt%,
2 wt%, and 5wt%. (B) SEM image of CS-HCA@PLGA-PEG@KGN hydrogel. (C)
FT-IR spectra of CS-HCA hydrogel (C1) PLGA-PEG@KGN nanoparticles (C2)
and CS-HCA@PLGA-PEG@KGN hydrogel (C3) (D) Drug release profiles of
PLGA-PEG@KGN nanoparticles and CS-HCA@PLGA-PEG@KGN hydrogels. (E)
Degradation profiles of CS-HCA hydrogels at different concentrations.
Drug release and degradation properties of CS-HCA@PLGA-PEG@KGN dynamic
hydrogel
We first investigated the interaction between CS-HCA hydrogels and
PLGA-PEG@KGN nanoparticles through FT-IR analysis. As shown in
[105]Figure 3C, the FT-IR spectrum of CS-HCA hydrogels displayed
characteristic peaks of -OH and -NH[2] at approximately 3,386 cm^-1.
The characteristic peaks of C=O, -CONH-, and aromatic rings were
observed at 1719, 1,637, 1,604, and 1,528 cm^-1. In the FT-IR spectrum
of PLGA-PEG@KGN nanoparticles, the -OH characteristic peak appeared at
around 3,414 cm^-1, while the C-O-C peak was detected at 1,069 cm^-1.
The C=O peak was observed at 1710 cm^-1, and the characteristic peaks
of the benzene ring from KGN were found at 1,611 and 1,523 cm^-1. The
FT-IR spectrum of CS-HCA@PLGA-PEG@KGN hydrogels revealed that the -OH
and -NH[2] characteristic peaks of CS-HCA and PLGA-PEG@KGN shifted to
3,386 cm^-1, while the C=O peak shifted to 1,111 cm^-1. The benzene
ring and -CONH- characteristic peaks of CS-HCA shifted to 1,539, 1,644,
and 1,600 cm^-1. These results suggest that CS-HCA hydrogels may
achieve effective loading onto PLGA-PEG@KGN nanoparticles through
electrostatic or van der Waals interactions.
UV-Vis spectrophotometry was employed to evaluate the drug release
kinetics of PLGA-PEG@KGN nanoparticles and CS-HCA@PLGA-PEG@KGN
hydrogels. The results ([106]Figure 3D) demonstrated that both systems
exhibited sustained drug release for over 3 weeks. Approximately 75% of
KGN was released from the nanoparticles within the first week, compared
to only 50% from the CS-HCA@PLGA-PEG@KGN hydrogel. By the third week,
cumulative release reached 94% for the nanoparticles and 70% for the
hydrogel. The prolonged drug release from the hydrogel, compared to the
nanoparticles alone, indicates that the hydrogel achieves dual-release
functionality through effective adsorption of nanoparticles.
The degradation profiles in [107]Figure 3E revealed that CS-HCA
hydrogels at different concentrations degraded by approximately 20% in
the first week. The degradation rate accelerated in the second week,
reaching 50%–60%, before slowing again to stabilize between 60% and 75%
by the fourth week. This degradation pattern allowed the hydrogel to
initially provide temporary mechanical support for cell adhesion and
proliferation, followed by gradual degradation during cartilage
regeneration to facilitate the deposition of newly formed tissue.
Effects of CS-HCA@PLGA-PEG@KGN hydrogel on BMSCs proliferation, migration,
and differentiation
In vitro experiments demonstrated that the CS-HCA@PLGA-PEG@KGN hydrogel
significantly enhanced the proliferation, migration, and chondrogenic
differentiation of BMSCs. As shown in [108]Figure 4A, the CCK-8 assay
revealed that BMSCs cultured on the CS-HCA@PLGA-PEG@KGN hydrogel
exhibited a markedly higher proliferation rate compared to those in the
2D group and the CS-HCA group (p < 0.05). Scratch assays and Transwell
migration assays ([109]Figures 4B–D) further indicated that cell
migration rates in the CS-HCA@PLGA-PEG@KGN group were superior to those
in the other two groups, while the CS-HCA group outperformed the
traditional 2D culture group (p < 0.05).
FIGURE 4.
[110]Panel A shows a bar graph comparing mean optical density over time
(24, 48, 72 hours) among three groups: 2D, CS-HCA, and
CS-HCA@PLGA-PEG@KGN, with notable increases in the latter. Panel B
presents a bar chart displaying mean cell numbers for these groups,
with CS-HCA@PLGA-PEG@KGN showing the highest values. Panel C consists
of three images of stained cell cultures: 1) 2D, 2) CS-HCA, and 3)
CS-HCA@PLGA-PEG@KGN, highlighting differences in cell morphology.
Panels D1 to D3 show time-lapse phase-contrast images (0h, 12h, 24h) of
these groups, illustrating cell proliferation and migration trends.
[111]Open in a new tab
In vitro effects of CS-HCA@PLGA-PEG@KGN hydrogels on the proliferation
and migration of BMSCs. (A) Assessment of BMSCs proliferation across
three groups using CCK-8 assay. (B) Quantitative comparison of migrated
BMSCs in three groups at 24-h post-treatment. (C) Assessment of BMSCs
migration across three groups using Transwell chamber assay. (D)
Assessment of BMSCs migration across three groups using Scratch wound
healing assay (D1) 2D group, (D2) CS-HCA hydrogel group, (D3)
CS-HCA@PLGA-PEG@KGN hydrogel group). (☆P < 0.05)
qRT-PCR and Western blot analyses ([112]Figure 5) demonstrated that the
mRNA and protein expression levels of chondrogenic markers (Sox9,
Aggrecan, and COL2A1) in the CS-HCA@PLGA-PEG@KGN group were
significantly upregulated compared to the other groups (p < 0.05).
Alizarin Blue staining showed noticeable blue staining across all three
groups after chondrogenic induction, indicating glycosaminoglycan
synthesis. Notably, the CS-HCA@PLGA-PEG@KGN hydrogel group exhibited
the most intense blue staining, with distinct cell aggregates,
suggesting the highest glycosaminoglycan content.
FIGURE 5.
[113]Four panels labeled A, B, C, and D. Panel A shows a bar graph of
mRNA levels of SOX9, ACAN, and COL2a1 across different conditions.
Panel B displays a bar graph of protein expression for the same
markers. Panel C shows Western blot images for β-actin, Sox-9, ACAN,
and Col2a1 across different samples. Panel D includes histological
images with three samples labeled 1, 2, and 3, showing stained cell
structures at a scale of 100 micrometers.
[114]Open in a new tab
In vitro effects of CS-HCA@PLGA-PEG@KGN hydrogels on the chondrogenic
differentiation of BMSCs. (A) qRT-PCR quantification of Sox9, Acan, and
Col2α1 mRNA expression levels of BMSCs in 2D, CS-HCA,and
CS-HCA@PLGA-PEG@KGN groups. (B) Quantitative comparison of Sox9, Acan
and Col2α1 protein expression levels of BMSCs in three groups. (C)
Western blotting analysis of Sox9, Acan, and Col2α1 protein expression
levels of BMSCs in 2D (1–1,1–2,1–3), CS-HCA (2–1,2–2,2–3) and
CS-HCA@PLGA-PEG@KGN (3–1,3–2,3–3) groups. (D) Alcian blue staining of
sulfated glycosaminoglycan (GAG) content in 2D (D1) CS-HCA (D2) and
CS-HCA@PLGA-PEG@KGN (D3) groups. (☆P < 0.05)
These findings confirm that the CS-HCA@PLGA-PEG@KGN hydrogel provides a
favorable three-dimensional microenvironment for stem cell growth and
differentiation. Its porous structure and sustained KGN release promote
cell adhesion on and within the scaffold, thereby enhancing BMSC
proliferation, migration, and chondrogenic differentiation capacity.
Gross morphological evaluation of cartilage defect repair
[115]Figures 6A-C illustrate the surgical procedure for establishing
the rat cartilage defect model and the ICRS scoring results. At 6 weeks
post-surgery, gross morphological evaluation revealed distinct repair
outcomes across the three groups in the cartilage defect regions
([116]Figure 6E). While all treatments demonstrated new tissue growth
at the defect sites, the CS-HCA@PLGA-PEG@KGN hydrogel group exhibited
the most optimal repair outcome. The control group ([117]Figure 6E1)
displayed incomplete defect filling with localized depressions, and a
clear boundary between the newly formed tissue and surrounding healthy
cartilage. The CS-HCA hydrogel group ([118]Figure 6E2) achieved
approximately 80% defect coverage, though the newly formed tissue
remained slightly depressed compared to the intact cartilage. In
contrast, the CS-HCA@PLGA-PEG@KGN group ([119]Figure 6E3) showed
complete filling of the defect with translucent and smooth tissue,
closely resembling the appearance of adjacent normal cartilage. The
repaired area was well-integrated without visible fissures,
inflammatory infiltration, or residual gel components. ICRS scoring
([120]Figure 6C) confirmed that the CS-HCA@PLGA-PEG@KGN group attained
the highest score, indicating superior cartilage repair efficacy.
FIGURE 6.
[121]A series of images and graphs demonstrating a scientific study on
tissue samples. Image A and B depict close-up photographs of surgical
wounds on a subject, showing different stages of healing with visible
stitches and measurements. Graphs C and D present bar charts comparing
mean CES and MWS values for control and experimental groups,
highlighting significant differences. Images E1, E2, and E3 show
cross-sections of tissue samples with scale bars indicating size.Panels
F, G, H, and I present microscopic images of histological sections,
stained to show tissue structure and cellular detail, with scale bars
for reference.
[122]Open in a new tab
Healing efficacy of in vivo cartilage defects. (A,B) Full-thickness
cartilage defect model of rat femoral trochlea (A) and hydrogel
implantation in osteochondral defects (B). (C) ICRS scores of
neocartilage in three groups. (D) The modified O’Driscoll histological
evaluation of neocartilage in three groups. (E) Gross morphological
observation of neotissue in Control (1), CS-HCA (2) and
CS-HCA@PLGA-PEG@KGN (3) groups. (F,G) HE staining of neocartilage at
×40 (F) and ×200 (G) magnification in three groups. (H,I) Col II
immunohistochemical staining at ×40(H) and ×200(I) magnification in
three groups. (☆P < 0.05).
Histological and immunohistochemical analysis of cartilage defect repair
Tissue samples were sectioned and subjected to histological staining
analyses to further evaluate cartilage regeneration. HE staining
([123]Figures 6F,G) revealed that the control group exhibited fibrous
tissue filling the defect site, with a clear boundary and noticeable
depression compared to surrounding cartilage. Type II collagen staining
([124]Figures 6H,I) demonstrated weaker staining intensity in the
repaired tissue compared to adjacent normal cartilage. The CS-HCA
hydrogel group showed incomplete defect repair, though with reduced
depression depth relative to the control group. While the repaired
tissue was closely integrated with surrounding cartilage, chondrocyte
density was lower, and type II collagen staining intensity remained
inferior to that of normal cartilage. In contrast, the
CS-HCA@PLGA-PEG@KGN hydrogel group achieved complete filling of the
defect with transparent cartilage-like tissue, flush with the
surrounding normal cartilage and exhibiting seamless interface
integration. Although chondrocyte arrangement within the repaired area
partially deviated from natural cartilage morphology, type II collagen
staining was more uniform and robust compared to the other groups.
Modified O’Driscoll histology scoring ([125]Figure 6D) confirmed that
the CS-HCA@PLGA-PEG@KGN hydrogel group attained the highest score,
indicating optimal cartilage regeneration outcomes.
Discussion
The restoration of articular cartilage defects is still a great
clinical challenge because of the limited intrinsic potential for
self-healing ([126]Xiang et al., 2022; [127]Hu et al., 2021). Bone
marrow stimulation techniques include drilling and the microfracture
technique, which aim to recruit BMSCs to repair cartilage defects. In
young patients with cartilage defects, the microfracture technique has
become a first-line clinical treatment due to its cost-effectiveness,
minimally invasive nature, and procedural simplicity. However, due to
insufficient recruitment of BMSCs or the impact of the local
inflammatory microenvironment after cartilage injury on stem cell
differentiation into chondrocytes, the newly formed tissue is
predominantly fibrocartilage with poor integration into the surrounding
cartilage. Fibrocartilage gradually degenerates over time, resulting in
poor long-term outcomes ([128]Mithoefer et al., 2009). Microfracture
combined with tissue engineering technology can improve the
microenvironment around damaged cartilage, promote stem cell migration
and differentiation into chondrocytes, thereby enhancing the repair
effect ([129]Frehner and Benthien, 2018).
In cartilage engineering scaffold materials, injectable hydrogels
exhibit unique advantages and significant clinical potential due to
their ability to repair complex-shaped defects and enable minimally
invasive implantation. The chitosan backbone, rich in amino groups, is
an ideal material for constructing dynamic hydrogels based on imine
bonds. In this study, a bioadhesive dynamic hydrogel was successfully
prepared by crosslinking CS-HCA with aldehyde-terminated four-armed
polyethylene glycol (AF-PEG). AF-PEG contains abundant aldehyde groups
and can rapidly form a gel within 2 min when mixed with CS-HCA under
physiological conditions. This hydrogel demonstrates excellent
injectability and self-healing properties, meeting the requirements for
clinical minimally invasive implantation. Compared to other
aldehyde-based crosslinkers commonly used in chitosan hydrogels, the
CS-HCA hydrogel synthesized in this study exhibits superior toughness,
with a compressive strength of up to 195 kPa. Moreover, the fractured
and self-healed hydrogel retains mechanical properties similar to those
of the original hydrogel, achieving a compressive modulus of 170 kPa
and a tensile modulus of 70 kPa. Research indicates that the phenolic
hydroxyl groups in catechol can also promote hydrogel self-healing
through non-covalent interactions such as hydrogen bonding and π-π
stacking ([130]Li et al., 2019). The CS-HCA hydrogel contains abundant
dynamic imine bonds and catechol groups. These reversible interactions
can break during hydrogel deformation, effectively dissipating energy
and thereby enhancing the hydrogel’s toughness, enabling it to
withstand repeated loads and mimic the biomechanical properties of
natural cartilage.
In addition to its injectability and self-healing properties, the
CS-HCA@PLGA-PEG@KGN hydrogel exhibits robust bioadhesive capabilities.
CS-HCA polymer contains a high density of free phenolic hydroxyl
groups, which can form covalent bonds with amino and thiol groups in
organic tissues, as well as establish non-covalent interactions such as
π-π stacking with various inorganic surfaces, thereby endowing the
CS-HCA hydrogel with superior bioadhesive capabilities ([131]Zhou et
al., 2022). As illustrated in [132]Figure 2, the hydrogel demonstrates
strong adhesion to both organic and inorganic substrates. Even under a
tensile force of up to 1,150 kPa, the hydrogel-bonded porcine skin
samples remained intact, highlighting its exceptional bioadhesive
strength. The catechol groups further mediate interfacial adhesion in
wet environments, with studies confirming their superior underwater
adhesive performance ([133]Guo et al., 2020). Following implantation
into cartilage defects, the CS-HCA@PLGA-PEG@KGN hydrogel forms stable
adhesion with surrounding host tissues, ensuring secure retention
without displacement from the implantation site. Researches indicate
that the catechol groups exhibit high protein affinity, enabling them
to anchor endogenous growth factors from bodily fluids and blood, while
also capturing signaling proteins secreted by adherent cells on the
hydrogel surface ([134]Zhao et al., 2024). These retained growth
factors and signaling proteins synergistically enhance cellular
adhesion, thereby establishing a microenvironment conducive to tissue
regeneration. Its injectability, self-healing capacity, and strong
bioadhesion collectively fulfill the requirements for minimally
invasive arthroscopic procedures, underscoring its significant
translational potential in clinical applications. In vitro degradation
analysis revealed that the hydrogel exhibited minimal mass loss during
the initial 7 days, followed by a progressive degradation profile, with
approximately 30%–40% of the initial mass retained at day 28. This
controlled degradation kinetics not only promoted early-stage cell
adhesion and proliferation but also provided structural support for
neocartilage deposition in subsequent phases.
Studies demonstrate that the three-dimensional porous architecture of
chitosan-based hydrogels mimics the topological structure of natural
cartilage extracellular matrix (ECM). The CS-HCA@PLGA-PEG@KGN hydrogel
features a characteristic 3D porous structure with a porosity of
69%–80% and pore sizes precisely controlled within the range of
20–200 μm. Research indicates that macropores (150–200 μm) facilitate
cell infiltration, while micropores (20–50 μm) increase the surface
area-to-volume ratio for efficient ECM protein adsorption ([135]Lyu et
al., 2022). Kim et al. reported that a macroporous polyvinyl alcohol
(PVA) scaffold facilitated chondrocyte migration from host cartilage
into scaffold and improved interface integration in an in vitro
cartilage defect model ([136]Kim et al., 2017). The microporous
structure also contributes to sustained drug release. Through FT-IR
analysis of CS-HCA hydrogels before and after loading PLGA-PEG@KGN
nanoparticles, our results indicated that the nanoparticles could be
adsorbed into the hydrogel via hydrogen bonds or van der Waals forces.
Drug release profiling revealed that PLGA-PEG@KGN nanoparticles
released approximately 75% of their KGN payload within 1 week, whereas
the CS-HCA@PLGA-PEG@KGN hydrogel released only 50% during the same
period. By week 3, cumulative release reached 94.5% for nanoparticles
compared to 70.6% for the hydrogel. These results confirm the
hydrogel’s capacity to effectively retain nanoparticles and establish a
dual sustained-release mechanism for KGN delivery.
It is well-established that KGN promotes the differentiation of BMSCs
into chondrocytes. Studies have demonstrated that KGN selectively
activates the BMP/Smad1/5 pathway while inhibiting the Wnt/β-catenin
pathway, thereby upregulating the expression of SOX9 and COL2A1 and
downregulating COL10A1 to prevent chondrocyte hypertrophy ([137]Cai et
al., 2019). Researchers have utilized KGN combined with a self-made
double-network hydrogel to repair rabbit knee articular cartilage
defects, achieving favorable outcomes ([138]Chen et al., 2022). RNA
sequencing and Gene Ontology (GO) analysis revealed significant
upregulation of genes involved in cell proliferation, chondrogenic
differentiation, and cartilage matrix synthesis within the regenerated
tissue, accompanied by downregulation of proteolytic and inflammatory
response-related genes. Notably, KEGG pathway enrichment analysis
demonstrated enhanced activation of metabolic pathways linked to
extracellular matrix (ECM) production—including focal adhesion and
ECM-receptor interaction—while pathways associated with matrix
degradation and inflammatory signaling were markedly suppressed. In our
study, the CS-HCA@PLGA-PEG@KGN hydrogel demonstrated excellent
biocompatibility, enhancing BMSC proliferation, migration, and
chondrogenesis. The abundant 3D microporous structure of the hydrogel,
combined with its sustained-release KGN, provides a favorable
microenvironment for BMSC survival and chondrogenic differentiation.
BM-MSCs are abundant yet prone to chondrocyte hypertrophy and
osteogenic differentiation ([139]Zhang et al., 2020). In injured or
degenerative joints, the local inflammatory microenvironment
compromises cell survival and differentiation, severely impairing
repair outcomes ([140]Roseti et al., 2019). Studies demonstrate that
KGN enhances stem cell chondrogenesis and inhibits cartilage
degradation in IL-1β-induced inflammatory joints ([141]Fan et al.,
2020). KGN’s enzymatic metabolite 4-ABP (4-aminobiphenyl) promotes MSCs
proliferation and chondrogenesis via the PI3K-Akt pathway, repairing
osteoarthritis cartilage damage ([142]Zhang et al., 2019).
Additionally, KGN alleviates joint inflammation and prevents
degeneration through the miR-146a/NRF2 axis ([143]Hou et al., 2021). In
our study, animal experiments revealed the CS-HCA@PLGA-PEG@KGN hydrogel
group achieved complete cartilage regeneration in defect zones at 6
weeks post-surgery, with seamless integration into surrounding tissue.
Although chondrocyte arrangement within the repaired area partially
deviated from natural cartilage morphology, type II collagen staining
was more uniform and robust compared to the other groups.
However, this study has several limitations. First, the observation
time points in animal experiments were limited, and the mechanistic
studies on cartilage regeneration remain superficial. Second, the
hydrogel still struggles to replicate the complex hierarchical
architecture of natural cartilage, resulting in structural
discrepancies between the newly formed cartilage and normal tissue. To
address these challenges, future studies could integrate 3D bioprinting
technology with the spatiotemporal synergy of multiple bioactive
factors (e.g., TGF-β, FGF) to engineer biomimetic hydrogels that
recapitulate the structural complexity of articular cartilage, thereby
promoting hierarchical cartilage regeneration.
Conclusion
In this study, we successfully developed a CS-HCA@PLGA-PEG@KGN
bioadhesive dynamic hydrogel based on the design principles of
bioadhesion, dynamic imine bonds, and sustained release of KGN
bioactive factors. In vitro experiments demonstrated that the hydrogel
exhibits excellent bioadhesive properties, injectability, and
self-healing capabilities. Its microporous structure and sustained
release of KGN effectively promoted the proliferation, migration, and
chondrogenic differentiation of BMSCs. Animal studies revealed that the
hydrogel significantly enhanced the repair of cartilage defects in rat
knee joints, with newly formed cartilage resembling hyaline-like tissue
and demonstrating seamless integration with surrounding native
cartilage. Furthermore, the hydrogel displayed rapid gelation under
physiological conditions, adaptability to irregular defect geometries,
and operational simplicity with a controllable fabrication process,
highlighting its promising potential for clinical applications in
cartilage regeneration.
Funding Statement
The author(s) declare that financial support was received for the
research and/or publication of this article. This work was supported by
Knowledge Innovation Project from the Wuhan Science and Technology
Bureau (Grant No. 2023020201010193).
Data availability statement
The datasets presented in this study can be found in online
repositories. The names of the repository/repositories and accession
number(s) can be found in the article/[144]Supplementary Material.
Ethics statement
The animal study was approved by Animal Experimental Ethics Committee
of Chongqing Western Biomedical Technology corporation. The study was
conducted in accordance with the local legislation and institutional
requirements.
Author contributions
ML: Conceptualization, Data curation, Funding acquisition,
Investigation, Methodology, Project administration, Writing – original
draft, Writing – review and editing, Formal Analysis, Resources,
Software, Supervision, Validation, Visualization. FL:
Conceptualization, Formal Analysis, Funding acquisition, Methodology,
Project administration, Resources, Supervision, Validation, Writing –
review and editing, Data curation, Investigation, Software,
Visualization. JXu: Data curation, Formal Analysis, Investigation,
Methodology, Software, Writing – review and editing. LZ: Data curation,
Formal Analysis, Investigation, Methodology, Software, Writing – review
and editing. JXi: Data curation, Formal Analysis, Investigation,
Methodology, Software, Writing – review and editing. CZ: Data curation,
Formal Analysis, Investigation, Methodology, Software, Writing – review
and editing. ZD: Data curation, Formal Analysis, Investigation,
Methodology, Software, Writing – review and editing. ST: Data curation,
Formal Analysis, Investigation, Methodology, Software, Writing – review
and editing. FO: Data curation, Formal Analysis, Investigation,
Methodology, Software, Writing – review and editing. JY: Data curation,
Formal Analysis, Investigation, Methodology, Software, Writing – review
and editing. XH: Data curation, Formal Analysis, Investigation,
Methodology, Software, Writing – review and editing.
Conflict of interest
The authors declare that the research was conducted in the absence of
any commercial or financial relationships that could be construed as a
potential conflict of interest.
Generative AI statement
The author(s) declare that no Generative AI was used in the creation of
this manuscript.
Publisher’s note
All claims expressed in this article are solely those of the authors
and do not necessarily represent those of their affiliated
organizations, or those of the publisher, the editors and the
reviewers. Any product that may be evaluated in this article, or claim
that may be made by its manufacturer, is not guaranteed or endorsed by
the publisher.
Supplementary material
The Supplementary Material for this article can be found online at:
[145]https://www.frontiersin.org/articles/10.3389/fbioe.2025.1606726/fu
ll#supplementary-material
[146]Table1.docx^ (15.1KB, docx)
[147]Table2.docx^ (13.1KB, docx)
References