Abstract
Cell cycle regulation is pivotal for tissue regeneration yet remains
challenging in degenerative microenvironments. We engineered a
sonosensitive conductive hydrogel incorporating
polypyrrole-encapsulated porphyrin derivatives {[Tetrakis
(4-carboxyphenyl) porphyrin (TCPP)]@PPy} to regulate cell cycle
dynamics. Upon ultrasound irradiation, TCPP@PPy generates free
electrons, establishing a controlled microcurrent within degenerative
tissues. This sonoelectric niche induces nucleus pulposus cell (NPC)
membrane depolarization, activating calcium voltage-gated channels
(Ca[v]) to drive Ca^2+ influx. Subsequent calcium- and
calmodulin-dependent protein kinase I activation up-regulates
cyclin-dependent kinases CDK1/CDK2, forming a
sonoelectricity–ion–kinase axis that stimulates NPC proliferation and
anabolism. Concurrently, ultrasound-responsive borate ester bonds in
the hydrogel amplify reactive oxygen species scavenging, counteracting
oxidative stress–induced NPC ferroptosis. In a goat model of
intervertebral disc degeneration, ultrasound-guided hydrogel
implantation alleviated degenerative progression by synergistically
reactivating cell cycle progression and suppressing oxidative damage.
This strategy demonstrates a noninvasive, dual-targeted approach to
regulate degenerative microenvironments through spatiotemporal control
of sonoelectric and biochemical cues, offering a translatable strategy
for tissue regeneration therapies.
__________________________________________________________________
A sonosensitive conductive hydrogel revives cell cycle and blocks
oxidation damage to reverse intervertebral disc degeneration.
INTRODUCTION
During the pathological process of intervertebral disc (IVD)
degeneration (IVDD), avascular nucleus pulposus (NP) tissue at the
center is subjected to persistent abnormal biomechanical loading, which
leads to inflammation and catabolism of NP cells (NPCs), ultimately
resulting in irreversible degenerative changes in the NP tissue ([48]1,
[49]2). Given the deep anatomical location of IVDs, current surgical
interventions remain inherently invasive ([50]3). While regenerative
strategies—including biomaterial-based approaches, stem cell therapies,
gene modulation, and tissue engineering—have predominantly focused on
IVD repair, they fail to induce durable cellular proliferation or
functional extracellular matrix (ECM) restoration ([51]4). Developing
noninvasive strategies to controllably activate endogenous cells
proliferation and rebalance ECM homeostasis thus represents a pivotal
frontier in overcoming pathological IVDD.
The cell cycle is a fundamental biological process that orchestrates
DNA replication, cell growth, and cell division, precisely regulating
tissue homeostasis and regeneration ([52]5, [53]6). Aberrations in cell
cycle control can lead to unchecked cell proliferation, contributing to
cancer ([54]7), while dysregulation of the cell cycle underlies
multiple degenerative disorders. For instance, G[2]-M phase arrest in
renal tubular epithelial cells promotes renal fibrosis ([55]8), and
synergistic interplay between cell cycle dysregulation and oxidative
stress drives neurodegeneration ([56]9). Emerging evidence indicate
that persistent cell cycle arrest initiates a cascade of molecular
events culminating in cellular senescence ([57]10), a biological
process recognized as a key contributor to IVDD ([58]11). The intricate
signaling networks regulate the cell cycle, and their integration with
pathways orchestrates cellular differentiation and tissue regeneration
([59]6). Interfering in these mechanisms may increase the regenerative
potential of cells and promote the development of therapeutic
strategies for regenerative medicine ([60]12). Therefore, targeted
modulation of cell cycle dynamics through specific molecular pathways
thus emerges as a theoretically plausible therapeutic method for IVDD.
Cyclin-dependent kinases (CDKs) form heterodimers with cyclins. In this
complex, CDKs act as catalytic subunits, and cyclins acts as regulatory
subunits that catalyze the phosphorylation of various substrates,
thereby driving the progression through different cell cycle phases and
ensuring that cell division is under stringent control ([61]13,
[62]14). In particular, CDK2 is relevant for DNA replication and
promotes the transition from the G[1] phase to the S phase, and CDK1
has emerged as the dominant determinant of the G[2]-M phase ([63]13,
[64]15). Typically, CDKs become completely active only after being
phosphorylated, combining substrate phosphorylation patterns regulated
by the cyclin-CDK complex, orderly coordinating the transition through
S phase and mitosis ([65]14, [66]16, [67]17). Studies have revealed
that CDK1 and CDK4 overexpression effectively promote cardiomyocyte
division ([68]12). CDKs activation also induces the expansion of
splenic T cell subsets, enhancing immune regulation ([69]18). These
findings imply that activating CDKs to advance the cell cycle may be a
potential therapeutic approach for treating degenerative diseases.
However, a strategy aimed at repairing IVDD by activating the CDKs of
NPCs to regulate the cell cycle has not yet been reported.
CDKs are activated by various upstream factors, including
phosphorylation by CDK-activating kinases; degradation of CDK
inhibitors, which release CDK activity; and regulation by
mitogen-activated protein kinases ([70]18–[71]20). In recent years,
CDKs have also been shown to be activated by calcium- and
calmodulin-dependent protein kinases (CaMKs) ([72]21, [73]22), a class
of protein kinases activated by calcium ions (Ca^2+) and calmodulin
that mediate calcium signal transduction through the phosphorylation of
substrate proteins and play crucial roles in regulating various
cellular functions ([74]23, [75]24). Studies have reported that calcium
voltage-gated channel (Ca[v]) opening results in Ca^2+ influx, which
subsequently activates CaMKs ([76]23–[77]26). Electrical currents can
induce cell membrane depolarization, triggering Ca[v] opening ([78]25,
[79]27). However, the efficacy of electrical signal–based strategies
for tissue regeneration remains unstable and lacks defined molecular
targets ([80]28, [81]29). Moreover, lumbar percutaneous electrical
currents ([82]30, [83]31) cannot accurately target deeply located IVDs.
Therefore, we are seeking a noninvasive and precise local electrical
stimulation strategy to regulate the cell cycle for IVDD therapy.
Tetrakis (4-carboxyphenyl) porphyrin (TCPP) exhibits high molecular
photothermal conversion efficiency and exceptional sonosensitivity,
complemented by its superior aqueous solubility and biocompatibility,
which facilitate homogeneous tissue distribution in vivo and minimize
potential side effects ([84]32, [85]33). In this study, we synthesized
a sonosensitive particle by encapsulating a porphyrin derivative with
polypyrrole (TCPP@PPy). PPy, a highly biocompatible and conductive
polymer ([86]34), was then polymerized with polyvinyl alcohol (PVA) to
form a composite (PPy/PVA). By mixing PPy/PVA, TCPP@PPy, and a reactive
oxygen species (ROS)–sensitive cross-linker
[N^1-(4-boronobenzyl)-N^3-(4-boronophenyl)-N^1,N^1,N^3,N^3-tetramethylp
ropane-1,3-diaminium (TSPBA)] ([87]35), a sonosensitive and conductive
hydrogel (TCPP@PPy-PPy/PVA) was prepared via in situ gelling. We found
that the TCPP@PPy-PPy/PVA hydrogel can generate an ultrasonic current
under ultrasound (US) irradiation; this alters the membrane potential
of the NPCs and promotes Ca[v] opening on the NPC membrane. The influx
of Ca^2+ activates CaMK1, which further activates CDK1 and CDK2. This
cascade promotes the cell cycle, thereby facilitating the regeneration
of NPCs. This ROS-degradable hydrogel subsequently accelerated ROS
clearance under US conditions, thereby mitigating NPC ferroptosis.
Last, we delivered the hydrogel to degenerated IVDs in goats via
puncture and subsequently treated them with US. After customized US
treatment, IVDD in goats was successfully treated, demonstrating the
clinical translational potential of this noninvasive strategy ([88]Fig.
1A).
Fig. 1. Synthesis and identification of sonosensitive hydrogel.
[89]Fig. 1.
[90]Open in a new tab
(A) The schematic illustration encapsulates the salient features and
methodologies in this study: (I) The overall therapeutic strategy
involves the noninvasive administration of a sonosensitive hydrogel
that responds to US for the treatment of IVDD. (II) In the magnified
IVD region, localized administration of the gel, US stimulation,
effectively induced the NPC regeneration. (III) The mechanism by which
US-induced TCPP@PPy sonosensitive particles generate free electrons.
(IV) The intracellular molecular mechanism by which ultrasonic current
facilitates the cell cycle progression in NPCs. (B) A ball-and-stick
model diagram of TCPP@PPy illustrates the structure. (C) SEM and TEM
revealed individual microscopic spherical morphology of PPy and
TCPP@PPy. (D) Dynamic light scattering experiments delineated the size
distribution of particles PPy and TCPP@PPy. (E) The schematic
representation illustrating the constituent elements and architectural
framework of the TCPP@PPy-PPy/PVA hydrogel network. (F)
Photodocumentation of PVA, PPy/PVA, TCPP/PPy/PVA, and TCPP@PPy-PPy/PVA
polymers before (Solution) and after (Hydrogel) cross-linking with the
TSPBA cross-linker. (G) SEM visualized the microporous structures of
PVA, PPy/PVA, TCPP/PPy/PVA, and TCPP@PPy-PPy/PVA hydrogels. (H) The
FTIR spectra of PVA, PPy/PVA, TCPP/PPy/PVA, and TCPP@PPy-PPy/PVA
hydrogels highlight the primary identical peaks, which are highlighted
in gray. (I) Compressive stress-strain curves of TCPP@PPy-PPy/PVA gel
with varying concentrations of PPy (strain rate = 10 mm/min). (J) The
temporal evolution of the swelling rate curves for gels PVA and
TCPP@PPy-PPy/PVA. (K) The degradation rate curves of gels PPy and
TCPP@PPy-PPy/PVA in H[2]O[2] (20 mM) and PBS environments were
analyzed. The error bars in [(J) and (K)] show the means ± SD of three
biological replicates.
RESULTS
Characterization of the TCPP@PPy-PPy/PVA hydrogel
TCPP@PPy particles were prepared via a modified chemical oxidation
polymerization method ([91]36). The long PPy chains enwind TCPP and are
interconnected with them via hydrogen bonds or π–π interactions
([92]Fig. 1B). Scanning electron microscopy (SEM) and transmission
electron microscopy (TEM) indicated that the PPy and TCPP@PPy particles
are relatively uniform spheres ([93]Fig. 1C). Energy-dispersive x-ray
spectroscopy elemental mapping revealed the uniform distribution of C,
N, and O elements in the PPy and TCPP@PPy particles (fig. S1A). Dynamic
light scattering analysis revealed that the peak diameters in the
intensity distribution of PPy and TCPP@PPy particles were approximately
255 ± 80.96 and 396 nm, respectively ([94]Fig. 1D). The polydispersity
index (PDI) of PPy particles (0.227) is consistent with previously
reported values ([95]37). In contrast, the higher PDI of TCPP@PPy
particles (0.353), indicative of moderate dispersity and comparable to
that of various biomedical polymer particles ([96]38), suggests that
surface functionalization may introduce slight heterogeneity (fig.
S1B). The zeta potential of PPy (−7.60 ± 11.5 mV) and TCPP@PPy
(49.0 ± 19.8 mV) indicate the introduction of cationic groups during
TCPP@PPy synthesis, suggesting the possible existence of
charge-interaction binding modes in the TCPP@PPy composite (fig. S1, B
to D). The Fourier transform infrared (FTIR) spectra of PPy, TCPP, and
TCPP@PPy depicted various characteristic bands (fig. S1E)
([97]39–[98]41). The peaks observed in the PPy FTIR spectra at 3445,
1635, and 1400 cm^−1 correspond to the N─H, C─C, and conjugated C─N
stretching vibrations, respectively. The peak at 1050 cm^−1 represents
C─H in-plane deformation vibration. The peaks at 1115 and 932 cm^−1
indicate the PPy doping state. The peaks at 1400, 1228, and 1173 cm^−1
in the TCPP FTIR spectrum represent C─O─H in-plane bending, C─O
stretching, and C─N stretching vibrations, respectively. The peak at
1664 cm^−1 is the stretching vibration of C═O in the carboxylic group.
The peak at 1604 cm^−1 corresponds to the stretching vibration of
aromatic C═C. Various peaks ranging from 1564 to 1472 cm^−1 are
attributed to the stretching modes of the porphyrin ring, specifically
C═C, C[α]─C[β], C[β]─C[β], C[α]─C[m] (m: meso carbon), and C═N bonds.
The peak at 793 cm^−1 is attributed to the C─H out-of-plane bending
vibration of the phenyl ring, indicating p-substitution. The
characteristic peaks of PPy and TCPP are observed in the TCPP@PPy FTIR
spectrum, implying their successful synthesis. Compared with that of
TCPP, the intensity of the C═O peak shifted from 1664 to 1677 cm^−1,
suggesting the formation of hydrogen bonds with the attached carboxyl
groups. At 1604 cm^−1, the C═C aromatic stretching vibration in
TCPP@PPy is weaker than that in TCPP, which may be attributed to π–π
interactions between the TCPP phenyl rings and the pyrrole rings.
Particles incubated in phosphate-buffered saline (PBS) were analyzed by
FTIR at multiple time points. The spectra revealed retention of
TCPP@PPy’s characteristic absorption bands throughout the 42-day
observation period, confirming no significant chemical degradation
occurred under physiological conditions (fig. S1F). The morphology
stability of TCPP@PPy particles in PBS was investigated using TEM. On
day 0, the particles exhibited a relatively well-defined spherical
morphology. From days 14 to 28, the particles gradually formed
irregular aggregates. By day 42, the edges became blurred, and the
aggregated structures became more pronounced (fig. S1G). Overall, the
particles demonstrated relatively good stability throughout the
observation period.
The final hydrogel network was formed by mixing 1 mg of TCPP@PPy with 1
ml of oxidatively polymerized PPy/PVA composite, followed by
cross-linking with TSPBA ([99]Fig. 1E). PVA, PPy/PVA, TCPP mixed with
PPy/PVA (TCPP-PPy/PVA), and TCPP@PPy-PPy/PVA were cross-linked with
TSPBA for comparative analysis ([100]Fig. 1F). SEM revealed that all of
the hydrogel networks exhibited porous structures ([101]Fig. 1G). The
peaks in the FTIR spectrum of the TCPP@PPy-PPy/PVA gel cover the major
peaks of the other intermediate gels, indicating the integrity of its
composition and structure ([102]Fig. 1H, gray box). Given that 9% (w/v)
PVA in hydrogels has optimal flexibility ([103]42), we adjusted the
content of PPy and found that when the concentration of PPy within the
PPy/PVA framework reached 0.1 M, the compressive stress of the gel no
longer substantially increased ([104]Fig. 1I). Simultaneously, as the
concentration of PPy in the gel increases, the stiffness of the gel
rises, while its ductility decreases (fig. S2A). We further investigate
influence of different concentration particles on the biomechanical
properties of hydrogels. Beyond TCPP@PPy (0.1 mg/ml) loading,
compressive strength plateaued (fig. S2B), while ductility exhibited
progressive deterioration with increasing particle content (fig. S2C).
In addition, compared with the PVA gel, the synthesized final gel
exhibited slightly reduced swelling properties. After 240 s, the
aqueous absorption of the TCPP@PPy-PPy/PVA gel did not increase
substantially, indicating that the hydrogel network has both absorbency
and stability ([105]Fig. 1J). The ROS-responsive gel can be
continuously degraded in a H[2]O[2] environment (20 mM). The
degradation assay revealed that the samples degraded slowly in PBS.
Under H[2]O[2] conditions, the PVA gel completely degraded on day 7,
whereas the TCPP@PPy-PPy/PVA gel fully degraded on day 10. This finding
suggested that the TCPP@PPy-PPy/PVA gel with a higher degree of
cross-linking has a longer duration ([106]Fig. 1K).
Sonosensitive properties and mechanism of the TCPP@PPy-PPy/PVA gel
We hypothesized that sonosensitive TCPP@PPy can generate free electrons
to generate a current under US induction. With 0.1 M PPy, we used an
electrochemical workstation to measure the ultrasonic current generated
by TCPP@PPy under US irradiation at different TCPP concentrations. When
the concentration of TCPP increased to 1 mM, the intensity of the
ultrasonic current did not substantially increase, indicating that 1 mM
is an optimal concentration ([107]Fig. 2A). Electrochemistry
measurements were also performed on particles after storage in PBS for
14, 28, and 42 days. Although the ultrasonic current intensity of
TCPP@PPy showed a slight decrease by day 42, it remained substantial
under US stimulation, confirming the stability of its ultrasonic
response (fig. S3A). To further investigate the underlying mechanisms,
photoluminescence (PL) spectra were obtained. The results showed that
the PL intensity of TCPP@PPy was markedly lower than that of TCPP,
indicating that the excited electrons in TCPP@PPy were easier to
transfer. These free electrons generated from TCPP are potentially
captured by PPy, thereby increasing the current intensity ([108]Fig.
2B). Similarly, after immersion in PBS, the particles exhibited a
slight increase in PL peak intensity over time. Even at day 42, the
peak remained substantial different from that of TCPP, demonstrating
the stability of the particles’ ultrasonic excitation–induced electron
migration capability (fig. S3B). As shown in [109]Fig. 2C, TCPP@PPy
exhibited greater absorption than did TCPP according to
ultraviolet-visible (UV-vis) spectroscopy. The corresponding
Kubelka-Munk plots revealed that the bandgaps of TCPP and TCPP@PPy were
estimated as 1.64 and 1.28 eV, respectively, indicating that PPy
decreased the energy barrier of TCPP ([110]Fig. 2D). Collectively,
under US induction, the electrons in the TCPP@PPy particles are more
easily excited from the highest occupied molecular orbital (HOMO),
overcoming the bandgap to transition into the lowest unoccupied
molecular orbital (LUMO) ([111]43). In addition, the conductive PPy
also acted as an electron trap to improve the transfer of free
electrons ([112]Fig. 2E). UV-vis spectra of hydrogels were also
measured, demonstrating that the conductive hydrogel–encapsulated
sonosensitive core approach (TCPP@PPy-PPy/PVA) exhibited the strongest
visible-light absorption with the smallest bandgap ([113]Fig. 2F).
Bandgaps of PVA, PPy/PVA, TCPP/PPy/PVA, and TCPP@PPy-PPy/PVA hydrogels
were calculated from the corresponding Kubelka-Munk plots as 2.36,
2.52, 3.67, and 4.01 eV, respectively ([114]Fig. 2G).
Fig. 2. Sonosensitive mechanism of materials.
[115]Fig. 2.
[116]Open in a new tab
(A) Under US irradiation with a 20 s per switch, TCPP@PPy generates
free electrons that form an electric current, which was detected using
an electrochemical workstation as the ultrasonic current produced by
TCPP@PPy. (B) PL spectra of TCPP and TCPP@PPy. (C) UV-vis adsorption
spectrum of TCPP and TCPP@PPy. (D) Kubelka-Munk plots derived from the
UV-vis adsorption spectrum of TCPP and TCPP@PPy. (E) Schematic
illustrating the mechanism of the generation of free electrons from
sonosensitive TCPP@PPy particles. (F) UV-vis adsorption spectrum of
PVA, PPy/PVA, TCPP/PPy/PVA, and TCPP@PPy-PPy/PVA gels. (G) Kubelka-Munk
plots derived from the UV-vis adsorption spectrum of PVA, PPy/PVA,
TCPP/PPy/PVA, and TCPP@PPy-PPy/PVA gels. (H) The line chart depicts the
correlation between the PPy content in the hydrogel and its
corresponding electrical conductivity. The data plotted represent
individual values and means ± SD of n = 3. (I) Graph comparing
electrical conductivity among PVA, PPy/PVA, TCPP/PPy/PVA, and
TCPP@PPy-PPy/PVA gel. The data plotted represent individual values and
means ± SD of n = 3. P values were determined using one-way analysis of
variance (ANOVA). (J) PL spectra of PVA, PPy/PVA, TCPP/PPy/PVA, and
TCPP@PPy-PPy/PVA gels. (K) Ultrasonic current detection of PVA,
PPy/PVA, TCPP/PPy/PVA, and TCPP@PPy-PPy/PVA gel under US irradiation.
a.u., arbitrary units.
We hypothesized that the generated ultrasonic current can be conducted
within the conductive hydrogel. Under the condition of 9% (w/v) PVA,
electrical conductivity tests demonstrated that the gel was conductive.
Furthermore, when the PPy concentration was increased to 0.1 M, the gel
conductivity did not substantially improve, indicating that 0.1 M is
the optimal concentration for application ([117]Fig. 2H). Similarly,
the synthesized PPy/PVA, TCPP/PPy/PVA, and TCPP@PPy-PPy/PVA hydrogels
exhibited markedly higher conductivities than did PVA, with no notable
differences noted among them ([118]Fig. 2I). The PL spectra revealed
that the TCPP@PPy-PPy/PVA gel is more easily excited to generate free
electrons than the other gels are ([119]Fig. 2J). In addition,
[120]Fig. 2K shows that TCPP@PPy-PPy/PVA produced the strongest
ultrasonic current under US irradiation. In particular, the
sonosensitivity of the TCPP@PPy-PPy/PVA gel is superior to that of the
gel formed by directly mixing TCPP with PPy/PVA (TCPP/PPy/PVA),
highlighting the necessity of introducing sonosensitive TCPP@PPy
particles. In brief, under US induction, the sonosensitive gel is
excited to generate free electrons from many independent sonosensitive
units, resulting in an improved electric current conducted through the
gel. This hydrogel can convert ultrasonic energy into electrical
energy, and the sonoelectric niche may affect the cells or tissues it
contacts.
Biological effects of TCPP@PPy-PPy/PVA gel
The biocompatibility of the TCPP@PPy-PPy/PVA gel was first evaluated
using three-dimensional (3D) cultures of NPCs at a concentration of
2.5 × 10^6 cells/ml ([121]44). Live/dead staining showed that the cells
maintained good viability on the 7th day, indicating that the gel is
relatively biocompatible ([122]Fig. 3A). NPCs (1 × 10^6, 2.5 × 10^6,
5 × 10^6, and 7.5 × 10^6 cells/ml) encapsulated in this hydrogel
demonstrated sustained viability after 7 days of 3D culture, as
evidenced by predominant green fluorescence signals across all
concentration groups, highlighting the hydrogel’s
concentration-independent cytoprotective effects within a certain range
under 3D culture conditions (fig. S4A). Subsequently, we treated NPCs
with US alone of different intensity. It was observed that cell
viability markedly decreased when the US intensity exceeded 0.5 W/cm^2.
To ensure cell viability while maximizing the generation of ultrasonic
current, an US intensity of 0.3 W/cm^2 was found to be suitable
([123]Fig. 3B).
Fig. 3. The effects of TCPP@PPPy-PPy/PVA gel on NPCs’ viability and
anabolic/catabolic metabolism.
[124]Fig. 3.
[125]Open in a new tab
(A) The TCPP@PPy-PPy/PVA gel’s impact on NPC viability within a 3D
culture system was evaluated through live/dead fluorescence staining on
days 1, 3, and 7. (B) The MTT assay was used to determine the impact of
varying US intensity on cell viability, with the culture time plotted
as a line graph. Data shown are means ± SD of n = 3 biological
replicates. (C) NPCs from Control, TBHP, TBHP + US, TBHP + Gel, and
TBHP + Gel + US groups were performed live/dead and phalloidin
fluorescence staining. (D) The PCR assay results from the Control-,
TBHP-, TBHP + US–, TBHP + Gel–, and TBHP + Gel + US–treated NPCs were
normalized and subsequently displayed using a heatmap. Data shown are
means ± SD of n = 3 biological replicates. (E) WB analysis of
inflammatory factor expression in NPCs following treatment with
Control, TBHP, TBHP + US, TBHP + Gel, and TBHP + Gel + US experimental
conditions. (F) The WB assay band images of ECM metabolism–related
protein in NPCs following treatment with Control, TBHP, TBHP + US, TBHP
+ Gel, and TBHP + Gel + US. (G) Control-, TBHP-, TBHP + US–, TBHP +
Gel–, and TBHP + Gel + US–treated NPCs were stained by Alcian. The
darker the blue staining represents the better the catabolism of
proteoglycan in NPCs.
To investigate the effects of gels on degenerative NPCs induced with
tert-butyl hydroperoxide (TBHP), we subjected NPCs to 3D culture using
both TCPP@PPy-PPy/PVA and PVA gels. Notably, only the TCPP@PPy-PPy/PVA
gel noticeably restored the fluorescence intensity of both the
live/dead staining and COL2A1 immunofluorescence following US treatment
(0.3 W/cm^2) compared with the fluorescence intensity induced by
TBHP-mediated inflammation (fig. S4B). Reverse transcription
quantitative polymerase chain reaction (RT-qPCR) and Western blot (WB)
revealed that the TCPP@PPy-PPy/PVA gel, in contrast to the PVA gel,
enhanced anabolism protein (COL2A1) expression and concurrently
suppressed the expression of genes (P16, P21, IL1B, IL6, CCL2, TNF,
ADAMTS5, and MMP13) and a protein (MMP13) related to catabolism (fig.
S4, C and D). These findings suggest that the sonosensitive
TCPP@PPy-PPy/PVA gel has a regenerative effect on cellular activity of
NPCs when subjected to US-induced conditions.
The elimination of US alone and the TCPP@PPy-PPy/PVA gel is essential.
Live/dead staining assays demonstrated that TBHP + Gel marginally
restored the fluorescence intensity relative to TBHP. TBHP + Gel + US
markedly enhanced the fluorescence of live cells. Phalloidin staining
indicated that TBHP + Gel + US treatment effectively restored the
cellular cytoskeletal architecture ([126]Fig. 3C). In addition, the
effects of TCPP@PPy-PPy/PVA hydrogel on the anabolic and catabolic
metabolism of NPCs were further explored. The RT-qPCR results revealed
that after TBHP-induced modeling, the gel slightly alleviated the
expression of a few inflammatory factors, whereas Gel + US
substantially reduced their expression ([127]Fig. 3D). WB analysis
confirmed the expression trend of the aforementioned inflammatory
factors at the protein level ([128]Fig. 3E). Similarly, only Gel + US
markedly restored the expression of the ECM anabolism–related protein
COL2, which was reduced by TBHP, and decreased the expression of the
ECM catabolism–related protein MMP13 and ADAMTS5 ([129]Fig. 3F). Alcian
blue staining of NPCs revealed that TBHP considerably inhibited their
anabolic metabolism. The application of gel alone conferred a modest
protective effect, whereas Gel + US robustly enhanced the proteoglycan
anabolism phenotype of NPCs ([130]Fig. 3G). In conclusion, for NPCs
subjected to inflammatory induction, Gel + US markedly promoted NPC
viability and anabolism, while US alone had no substantial effect.
Molecular mechanism by which the TCPP@PPy-PPy/PVA gel promotes NPC
regeneration under US
We first investigated the impact of the sonoelectric effect from
TCPP@PPy-PPy/PVA gel NPCs during the initial stage. We used
3,3′-dipropylthiadicarbocyanine iodide [DiSC[3] ([131]5)] ([132]45) to
test the membrane potential of NPCs cocultured with the gel. Under US
irradiation (0.3 W/cm^2), a rapid increase in fluorescence intensity
was observed within 5 min, suggesting transient depolarization of the
NPC membrane ([133]Fig. 4A). To investigate the metabolic changes that
occur within NPCs following exposure to ultrasonic current and given
that US alone has no substantial effect on NPC viability, we conducted
mRNA sequencing on NPCs from four treatment groups: the control, TBHP,
TBHP + Gel, and TBHP + Gel + US groups. Kyoto Encyclopedia of Genes and
Genomes (KEGG) pathway enrichment analysis revealed that, compared with
the control, TBHP significantly down-regulated cell cycle–related
pathways in NPCs (fig. S5A). Gene set enrichment analysis (GSEA) also
revealed that the down-regulated genes were significantly associated
with the cell cycle (fig. S5B). Compared with TBHP or TBHP + Gel, TBHP
+ Gel + US significantly up-regulated the cell cycle in KEGG pathways
([134]Fig. 4B and fig. S5C), and the up-regulated enriched genes were
also strongly related to the cell cycle ([135]Fig. 4C and fig. S5D).
The mRNA sequencing data collectively illustrated that Gel + US
effectively reversed the inflammation-induced attenuation of cell cycle
metabolism in NPCs. The heatmap depicts gene expression patterns in the
cell cycle pathway and clearly illustrates that TBHP induced
substantial down-regulation of gene expression. Gel alone did not
sufficiently counteract this suppression, which was able to be
up-regulated by Gel + US ([136]Fig. 4D). These genes regulate the
phenotype of the downstream cell cycle. Cell cycle flow cytometry
analysis of NPCs post–TBHP induction showed a near absence of cells in
the G[2]-M phase relative to the control, with most cells exhibiting
arrest in the G[0]-G[1] phase, which was not reversed by Gel. Gel + US
markedly facilitated the progression of a portion of the TBHP-damaged
NPCs into the G[2]-M phase ([137]Fig. 4E and fig. S6). Regarding cell
cycle regulation of cell proliferation and metabolism, Gel + US not
only markedly restored the viability of TBHP-injured NPCs, as detected
using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide
(MTT) assay ([138]Fig. 4F), but also notably restored anabolism
([139]Fig. 3, D to G).
Fig. 4. Molecular mechanism of cell cycle promotion in NPC.
[140]Fig. 4.
[141]Open in a new tab
(A) Membrane depolarization in NPCs treated with Control, TBHP, TBHP +
Gel, or TBHP + Gel + US (5 min/US cycle over 25 min) was quantified
using membrane potential dye DiSC[3](5). (B) KEGG enrichment analysis
(TBHP + Gel + US versus TBHP) identified the top 10 up-regulated
metabolic pathways, with dot-size reflecting metabolite counts, x axis
as metabolite ratios, and color indicating enrichment significance
(−log[10]P). (C) GSEA confirmed enrichment of cell cycle–related
pathways in TBHP + Gel + US versus TBHP. (D) The mRNA sequencing
heatmap showed the normalized expression of cell cycle–related genes in
groups. (E) Flow cytometry graph of the cell cycle distribution
(G[0]-G[1], S, and G[2]-M phases) demonstrated a higher proportion of
G[2]-M phase in TBHP + Gel + US versus TBHP + Gel (*P = 0.0321). (F)
MTT assay showed enhanced cell viability in TBHP + Gel + US compared to
TBHP or TBHP + Gel. (G) The intracellular Ca^2+ concentrations (Fluo-4
fluorescence) under US exposure (10 min/ US cycle over 40 min) were
determined. (H) Heatmap showed the normalized expression of CAMK
related genes in Control-, TBHP-, TBHP + Gel–, and TBHP + Gel +
US–treated NPCs, based on mRNA sequencing (n = 3). (I) WB images of
CaMK1, P-CaMK1, CDK1/2, and P-CDK1/2 protein expression in NPCs from
all groups. (J) Ben treatment reduced G[2]-M–phase cells (*P = 0.0377
versus TBHP + Gel + US) and suppressed (K) viability compared to TBHP +
Gel + US. (L) WB showed TBHP + Gel + US–rescued COL2 expression and
inhibited MMP13 up-regulation caused by TBHP, with these effects
reversed by Ben. (M) WB confirmed that KN93 blocked CaMK1, P-CaMK1,
CDK1/2, and CDK1/2 expression in TBHP + Gel + US. (N) KN93 reduced
G[2]-M–phase cells (*P = 0.0273 versus TBHP + Gel + US), (O) diminished
viability, (P) abolished COL2 restoration, and weakened MMP-13
suppression compared to TBHP + Gel + US group. (Q) Schematic: The
regeneration is promoted by Ca^2+-CaMK1-CDK1/2-cell cycle axis. Data
represent means ± SD (three biological replicates). Statistical
analysis: one-way ANOVA [(F), (K), and (O)], two-way ANOVA [(E), (J),
and (N)].
Recent studies reported that current induced cell membrane
depolarization, subsequently triggering Ca[v] opening and consequently
facilitating Ca^2+ influx ([142]25, [143]27). NPCs were subjected to a
Fluo-4 assay to detect Ca^2+ influx. The Ca^2+ content in the cytoplasm
of the TBHP-induced NPCs was detected to exclude the influence of TBHP.
The Ca^2+ concentration in NPCs temporarily increased within 12 hours
after the addition of TBHP (fig. S7A). After 24 hours of pretreatment
with TBHP, NPCs were subjected to various treatments, and the Fluo-4
assay results indicated that NPCs treated with Gel + US presented a
transient increase in intracellular Ca^2+ concentration, suggesting
that ultrasonic current stimulates Ca^2+ influx ([144]Fig. 4G and fig.
S7B). Hypothesizing that Ca^2+ influx through Ca[v] activates CaMKs
([145]23–[146]25), on the basis of a gene expression heatmap generated
via mRNA sequencing, we found that CAMK1 expression in NPCs was
down-regulated by TBHP but was observably up-regulated by Gel + US
stimulation ([147]Fig. 4H). Given the potential of CaMKs to activate
CDKs ([148]21, [149]22) and the most pronounced reactivation of CDK1
and CDK2 in NPCs by Gel + US ([150]Fig. 4D), we hypothesized that CaMK1
up-regulation further up-regulates its downstream targets CDK1 and
CDK2. WB assay results initially elucidated that during the promotion
of the cell cycle in NPCs by Gel + US after TBHP preincubation, the
protein expression of CaMK1, CDK1, and CDK2 increased, and their
phosphorylation levels also increased ([151]Fig. 4I).
To initially verify whether ultrasonic current activates CaMK1 by
opening Ca[v] and facilitating Ca^2+ influx, we used the broad-spectrum
Ca[v] blocker benidipine (Ben, 1 μM) ([152]46). The WB results
indicated that in the TBHP + Gel + US + Ben group, the protein
expression of CaMK1 and phosphorylated CaMK1 (P-CaMK1) in NPCs did not
recover as observed in the TBHP + Gel + US group. Furthermore, the
expression of the presumed downstream targets, namely, CDK1,
phosphorylated CDK1 (P-CDK1), CDK2, and phosphorylated CDK2 (P-CDK2),
was also weak ([153]Fig. 4I). After Ben-mediated inhibition, Gel + US
subsequently failed to rescue cells cycle arrest and effectively forced
the cell cycle progression ([154]Fig. 4J and fig. S8A), increasing
cellular viability ([155]Fig. 4K). Compared with Gel + US, Gel + US +
Ben treatment neither effectively restored the expression of the
anabolic protein COL2 nor inhibited the expression of the catabolic
protein MMP13 ([156]Fig. 4L). These results suggest that the increased
CaMK1 activity is regulated by ultrasonic current–induced Ca^2+ influx
via Ca[v].
We used KN93 (30 μM), a nonspecific CaMK inhibitor ([157]23), to
investigate whether downstream CDK1 and CDK2 are regulated by CaMK1.
Compared with that observed in the Gel + US–treated NPCs, the
expression of activated CaMK1 (P-CaMK1) in the KN93-treated NPCs was
substantially lower after TBHP preinduction; subsequently, the activity
of CDK1, P-CDK1, CDK2, and P-CDK2 was also attenuated ([158]Fig. 4M).
Furthermore, KN93 administration mitigated the cells cycle arrest
salvage and cell cycle propulsion induced by Gel + US after TBHP
pretreatment ([159]Fig. 4N and fig. S8B), diminished the rescue of
cellular viability typically achieved with Gel + US ([160]Fig. 4O),
attenuated the up-regulation of COL2 protein expression in NPCs
stimulated with Gel + US, and made the down-regulation of MMP13
expression less pronounced ([161]Fig. 4P). The experimental findings
allow us to elucidate a molecular mechanism: The sonoelectric effect
opens the Ca[v] on the membrane of NPCs, facilitating the influx of
Ca^2+. This event activates CaMK1, which subsequently activates its
downstream targets, CDK1 and CDK2. The activation of these CDKs propels
the cell cycle forward, culminating in subsequent metabolic alterations
and regeneration ([162]Fig. 4Q).
Sonosensitive hydrogel protects NPCs from oxidative stress
Oxidative stress in NPCs triggers ferroptosis, which is recognized as a
pivotal role in the pathogenesis of IVDD ([163]47). Considering the
ROS-scavenging capabilities of the TSPBA hydrogel ([164]35), we
explored its potential to safeguard NPCs against oxidative
stress–induced injury. Titanium oxysulfate was used to investigate the
neutralizing effects of the gel on H[2]O[2] in a PBS solution. The
control and US had minimal effects on H[2]O[2] levels. Compared with
Gel, Gel + US (0.3 W/cm^2) markedly increased the efficiency of
H[2]O[2]neutralization, possibly because ultrasonic cavitation enhanced
the redox reaction between H[2]O[2] and borate ester ([165]Fig. 5A).
Flow cytometry and fluorescence quantification were used to compare the
intracellular ROS content in NPCs. TBHP substantially increased ROS
levels in NPCs. In contrast, US showed minimal efficacy in eliminating
ROS, whereas the gel exhibited a modest mitigating effect. Notably, Gel
+ US was the most effective at reducing the intracellular ROS
concentration ([166]Fig. 5, B and C). Consequently, it can be proved
that the application of US accelerates the clearance of ROS by the Gel
(TCPP@PPy-PPy/PVA).
Fig. 5. Sonosensitive hydrogel regulates oxidative stress and ferroptosis in
NPCs.
[167]Fig. 5.
[168]Open in a new tab
(A) Evaluate the changes in H[2]O[2] (20 mM) content over time using
titanium oxysulfate in four different systems: Control, US, Gel, Gel +
US. (B) The cell flow cytometry assay for ROS compared the
intracellular ROS levels in NPCs following treatments with Control,
TBHP, TBHP + US, TBHP + Gel, and TBHP + Gel + US. (C) The fluorescence
intensity of ROS-positive NPCs was quantified as mean fluorescence
intensity (MFI). (D) In this study’s mRNA sequencing analysis, compared
to group TBHP, the ferroptosis was one of the KEGG pathways that showed
a significant decrease in group Gel + US. (E) GSEA from WikiPathways
(WP) gene sets associated with expression changes in TBHP + Gel + US
versus TBHP NPCs. (F) Heatmap showed the normalized data of
ferroptosis-related gene expression in Control-, TBHP-, TBHP + Gel–,
and TBHP + Gel + US–treated NPCs in mRNA sequencing. (G) In the LPO
fluorescence staining of NPCs, the green positive signal indicates the
presence of peroxidized lipids. In AO staining, the stronger the red
signal, the less damage the endosomal membrane has suffered from
ferroptosis. (H) Representative WB images of GPX4 protein in Control-,
TBHP-, TBHP + Gel–, and TBHP + Gel + US–treated NPCs. Data in [(A) to
(F)] are shown as means ± SD, with n = 3 biological replicates.
Statistical significance in (C) was calculated using one-way ANOVA.
KEGG pathway analysis and GSEA of mRNA sequencing indicates that,
compared with TBHP induction, ferroptosis was related to oxidative
stress and was one of the most down-regulated pathways in NPCs treated
with Gel + US ([169]Fig. 5, D and E). Simultaneously, TBHP markedly
up-regulated the ferroptosis compared to Control, and Gel + US
down-regulated this pathway compared to Gel (fig. S9, A to D), which
indicated that Gel + US can mitigate ferroptosis induced by oxidative
stress. Furthermore, in this mRNA sequencing analysis, the differential
expression of genes in the ferroptosis pathway is clearly illustrated
in a heatmap. TBHP substantially up-regulated ferroptosis-related
genes; Gel slightly ameliorated the up-regulation induced by TBHP,
whereas Gel + US markedly reduced their expression ([170]Fig. 5F). ROS
down-regulates the expression of glutathione peroxidase 4 (GPX4), an
enzyme that plays a crucial role in inhibiting lipid peroxidation (LPO)
([171]48), which is also inhibited by Gel + US ([172]Fig. 5F).
Given the fact that ROS leads to LPO, which is also a major phenotype
of ferroptosis, we proceeded to assess the LPO levels in NPCs via a
lipid droplet fluorescence detection assay. TBHP substantially
increased intracellular LPO levels, an effect that was not effectively
mitigated by Gel, whereas Gel + US effectively eliminated LPO
([173]Fig. 5G). As ROS can damage the endosomal membrane ([174]49),
acridine orange (AO) staining is one of the phenotypes associated with
ferroptosis ([175]50). [176]Figure 5G shows that TBHP substantially
reduced red fluorescence, indicating that the endosomal membrane was
incomplete, with Gel demonstrating limited protective effects. In
contrast, Gel + US provided the most effective protection ([177]Fig.
5G). Ultimately, the key enzyme GPX4 involved in ferroptosis
regulation, which was substantially down-regulated by TBHP, was
markedly restored upon treatment with Gel + US ([178]Fig. 5H). Overall,
the extra protective role of the TCPP@PPy-PPy/PVA gel in NPCs not only
facilitates the cell cycle but also mitigates ferroptosis by
accelerating the clearance of ROS under US.
Therapeutic efficacy against IVDD in goats
Because the size, intradiscal pressure, and range of motion of caprine
lumber spines are similar to those of human lumber spines ([179]51,
[180]52), caprine lumber spines were selected for this study. A cohort
of 12 Boer goats was evenly distributed across four experimental groups
[Control, Decompression (Decomp), Decomp + Gel, and Decomp + Gel + US].
Nine goats underwent IVD decompression on day 0. On the 7th day
postsurgery, x-ray fluoroscopic monitoring revealed that the puncture
needle was accurately localized to the IVD region. Three goats in both
the Decomp + Gel and Decomp + Gel + US groups were injected with
TCPP@PPy-PPy/PVA gel into the IVD using a puncture needle, whereas the
control group underwent a sham operation. On days 7, 14, 21, and 28,
three goats from the Decomp + Gel + US group underwent 30 min of US
irradiation of the lumbar area. By Day 42, all the goats were
euthanized, followed by radiographic and histological evaluations
([181]Fig. 6A).
Fig. 6. Histological and radiographic analysis of goat IVDs.
[182]Fig. 6.
[183]Open in a new tab
(A) Schematic diagram of the experimental process in goats. IVD Decomp
modeling surgery was performed on day 0, followed by gel injection into
the IVD under fluoroscopic guidance on the 7th postoperative day. US
irradiation was administered for 30 min on days 7, 14, 21, and 28, with
euthanasia conducted on day 42. (B) Sagittal and axial MRI images of
the goat IVD. The dark of the outer loop represents the AF, and the
high-density inner area is NP. The yellow arrow points to the leaked
NP. (C) Modified Pfirrmann grade is used to comprehensively evaluated
the MRI images of IVD on day 42. (D) Safranin O, H&E, and Masson
staining of histological sections of IVD tissues on day 42. (E)
Histological grading of IVD was performed on the basis of the degree of
degeneration observed in histological sections from each group on day
42. (F) CT images compare the intervertebral height in Control, Decomp,
Decomp + Gel, and Decomp + Gel + US groups on day 42. (G) Disk height
index collected from CT images compared between all groups on day 42.
Heatmap showed the normalized data of RT-qPCR fold changes of ECM
metabolism factors (H), cell cycle phenotype, and ferroptosis phenotype
(I) related genes in goats’ paravertebral tissues on day 42. Data and
error bars in [(C), (E), and (G)] represent the means ± SD, with n = 3
biological replicates. One-way ANOVA was performed.
On the basis of the sagittal T2-weighted magnetic resonance imaging
(MRI), the control group exhibited strong aqueous disc signals,
indicating healthy and well-hydrated IVDs. In contrast, the Decomp
group presented a significant reduction in signal intensity. The gel
treatment resulted in a slight recovery of the signal, whereas the Gel
+ US therapy resulted in a more pronounced alleviation of signal loss
([184]Fig. 6B). Axial T2 MR images were used to visualize the NP and
annulus fibrosus (AF). Compared with the control treatment,
decompression treatment severely decreased heterogeneity and hydration
in the NP region. The border between the NP and AF appears indistinct,
with the NP seemingly leaking into the AF ([185]Fig. 6B, yellow arrow).
Compared with the Decomp group, treatment with the gel appeared to
mildly preserve the hydration and geometry of the NPs. Gel + US
treatment resulted in goats with IVDs whose hydration and morphology
were most similar to those of the control ([186]Fig. 6B). The Pfirrmann
grade, as determined using MRI, provides a quantitative evaluation of
the aforementioned outcomes ([187]Fig. 6C).
Morphological and histological assessments of IVD tissue were conducted
using the mid-coronal plane section approach. Safranin O staining
revealed the typical native structure of control IVDs, highlighting the
homogeneous, proteoglycan-rich NP with a distinct red hue and intact
lamellar bundles of AF. In the Decomp group, IVDs presented remarkable
heterogeneity in the NP, characterized by the substantial disappearance
of red staining and widespread blue staining. These changes reflect a
marked focal loss of proteoglycans, the deposition of type I collagen,
and advanced calcification. The Gel treatment afforded modest
protection for the IVD, whereas the application of Gel + US markedly
mitigated the depletion of proteoglycans and attenuated the
pathological alterations in disc morphology ([188]Fig. 6D). Masson
staining further confirmed these findings, revealing a loss of
cartilage (blue) at the decompressed IVD, which was replaced by
pathologically deposited red-stained fibrin, scar tissue, and type I
collagen. Gel treatment partially ameliorated this pathology, whereas
Gel + US therapy markedly reversed IVDD, restoring blue staining to
levels more similar to those of the control ([189]Fig. 6D). Hematoxylin
and eosin (H&E) staining revealed that Decomp noticeably induced
inflammatory infiltration and morphological alterations in the IVD. The
Gel treatment offered limited improvement, whereas the Gel + US
treatment had the most pronounced therapeutic effect on the IVD
([190]Fig. 6D). The histological staining grade was used to assess the
severity of IVDD quantitatively, with Decomp markedly inducing IVDD.
Treatment with Gel resulted in a moderate mitigation of degeneration;
however, treatment with Gel + US exhibited considerably enhanced
restoration of IVD compared to Gel alone ([191]Fig. 6E).
Clinically, because of the loss and dehydration of NP tissue, IVDD
progressively diminishes disc height, which compromises the capacity of
the NP to withstand compressive stresses in the spine. Computed
tomography (CT) and the disc height index elucidated that Decomp led to
a substantial narrowing of the IVD space, indicating advanced
degeneration. The gel treatment marginally maintained the disc height,
whereas the Gel + US therapy more effectively preserved the disc height
([192]Fig. 6, F and G).
Last, the IVDs from each group were harvested and subjected to RT-qPCR
analysis to assess the metabolic conditions. The anabolic genes COL2A1,
ACAN, and SOX9 were significantly down-regulated in the IVD tissues of
the Decomp group, with no significant changes observed in the Decomp +
Gel group compared with the Decomp group. In contrast, a noticeable
restoration of the expression of these genes was detected in the Decomp
+ Gel + US group. In the Decomp group, the catabolism genes MMP13,
ADAMTS5, COL1A1, COL10A1, and RUNX2 exhibited pronounced up-regulation
in IVD tissues. Gel treatment alone did not effectively counteract this
increase. In contrast, Gel + US treatment markedly attenuated their
expression ([193]Fig. 6H). In particular, previously validated genes
integral to the cell cycle axis, CAMK1, CDK1, and CDK2, and other cell
cycle phenotype–related genes (CCND1 and CCNE1), were significantly
down-regulated in the IVD tissue of the Decomp and Decomp + Gel groups.
The application of Gel + US led to pronounced up-regulation of these
genes, suggesting the therapeutic modulation of cell cycle–related
pathways by the sonosensitive TCPP@PPy-PPy/PVA gel in vivo ([194]Fig.
6I). Notably, both Decomp and Decomp + Gel groups exhibited marked
down-regulation of GPX4 and SLC7A11 (a cystine transporter critical for
glutathione synthesis) ([195]53), alongside up-regulation of ACSL4, a
key driver of LPO ([196]54). Strikingly, Gel + US intervention restored
GPX4 and SLC7A11 expression while suppressing ACSL4, demonstrating that
the hydrogel therapeutically modulates ferroptosis-associated pathways
in vivo ([197]Fig. 6I).
In conclusion, the sonosensitive TCPP@PPy-PPy/PVA gel has protective
effects on goat IVD tissue, and its application in combination with US
can partially facilitate IVD regeneration. In vivo experiments have
substantiated the efficacy of this noninvasive sonosensitive gel
therapy for IVDD, and its availability in large animal models strongly
highlights its promising prospects for clinical translation.
DISCUSSION
In this study, we present a noninvasive therapeutic strategy for IVDD
using a sonosensitive conductive hydrogel (TCPP@PPy-PPy/PVA) designed
to precisely modulate cell cycle progression and mitigate oxidative
stress damage. The hydrogel leverages US-triggered microcurrent
generation to activate endogenous regenerative pathways within
degenerative IVD. At the micro level, US irradiation induces free
electron excitation in the TCPP@PPy particles, creating a localized
electrical microenvironment that depolarizes NPC membranes. This
depolarization opens Ca[v], driving Ca^2+ influx and activating the
Ca^2+/CaMK1 signaling axis, which subsequently up-regulates and
phosphorylate CDK1 and CDK2 to propel cell cycle progression.
Concurrently, the hydrogel’s ROS-responsive borate ester bonds enhance
scavenging of ROS under US, effectively suppressing ferroptosis in
NPCs. Validated in a caprine IVDD model, this dual-action strategy
restored disc hydration, ECM homeostasis, and structural integrity, as
evidenced by MRI, histology, and molecular profiling. Our findings
establish a paradigm for noninvasive tissue regeneration through
spatiotemporal control of microcurrent and biochemical microenvironment
niche, addressing critical limitations of invasive surgical and
transient pharmacological interventions.
The management of IVDD remains a formidable clinical challenge,
constrained by the limitations of existing therapeutic paradigms.
Surgical interventions, such as discectomy or spinal fusion, provide
symptomatic relief but inevitably disrupt disc biomechanics and
accelerate adjacent segment degeneration, failing to restore native
tissue functionality ([198]3). Regenerative strategies, including stem
cell therapy and biomaterial-based scaffolds, aim to replenish lost
NPCs and ECM ([199]4). However, the harsh microenvironment of
degenerative discs—characterized by hypoxia, nutrient deprivation, and
chronic inflammation—severely compromises transplanted cell viability
and ECM synthesis ([200]1, [201]2, [202]4, [203]55). For instance,
animal and clinical studies have demonstrated that the survival rate of
cells injected into IVD is low (~18%), the therapeutic effects are
inconsistent, and potential complications such as osteophyte formation
and lower back pain may occur ([204]56). Similarly, synthetic hydrogels
designed for mechanical support often lack dynamic responsiveness to
pathological cues, resulting in passive degradation without triggering
endogenous repair ([205]4, [206]57, [207]58). Recent advances in
US-mediated therapies have introduced previously unidentified avenues
for noninvasive modulation. Studies have demonstrated that
sonosensitive biomaterials, under controlled US stimulation, can
release cytotoxic agents (ROS and heat), enhance drug release and
tissue penetration, and generate electrical effects, thereby achieving
therapeutic outcomes such as antimicrobial activity, tumor suppression,
and cell or tissue regeneration ([208]59–[209]62). Concurrently,
emerging bioelectronic strategies—such as triboelectric/piezoelectric
nanogenerators for bone regeneration ([210]63), and implantable
zinc-oxygen batteries for neural repair ([211]64)—demonstrate the broad
potential of electroactive therapies in tissue regeneration. Yet,
percutaneous electrical stimulation’s inability to spatially target
deep spinal structures and maintain stable current delivery limits
efficacy, often causing off-target muscle contractions or tissue damage
([212]65, [213]66). The cell cycle is one of the core molecular
biological processes that drive cell growth ([214]5, [215]6), and
modulating this intricate mechanism may essentially promote tissue
regeneration. However, sufficient evidence indicates that prolonged
cell cycle arrest ultimately leads to reduced viability, senescence, or
cell death ([216]8–[217]10, [218]67, [219]68). Targeting cell cycle
regulation to revitalize quiescent NPCs represents a crucial yet
underdeveloped therapeutic avenue. CDKs, central drivers of
proliferation, have been successfully activated in cardiomyocytes and
immune cells to reverse fibrosis or enhance immunomodulation ([220]12,
[221]18). Here, in IVDD, NPCs are arrested in G[0]-G[1] because of TBHP
oxidative stress and senescence induction ([222]Figs. 4 and [223]5), a
pathology in traditional regenerative approaches that overlook cell
cycle dynamics. This study aims to develop a strategy for controllable,
noninvasive, and efficient deep IVD regeneration by using US to
precisely drive the cell cycle.
Our work delineates a physically biochemically innovative paradigm. In
this research, PPy, a biocompatible and conductive polymer with wide
application potential ([224]69, [225]70), was selected to encapsulate
the sonosensitizer TCPP to manufacture sonosensitive particles denoted
as TCPP@PPy. The TCPP@PPy-PPy/PVA hydrogel generates spatially confined
microcurrents under US, enabling precise Ca[v] channel activation
without off-target effects. This precision current stems from the
tailored electronic structure of TCPP@PPy particles: Compared to
conventional conductive polymers PPy alone, the TCPP@PPy composite
exhibits superior sonoelectronic conversion efficiency ([226]Fig. 2A).
The mechanistic foundation of this technology lies in the unique
electronic structure of the photosensitizer and sonosensitizer. In
molecular orbital theory, HOMO represents the highest energy level
containing electrons at equilibrium state, while LUMO denotes the
lowest energy level available for electron excitation. US waves act
core of sonosensitive materials core and provide sufficient energy
(ΔE = E[LUMO] − E[HOMO]) ([227]43) to trigger interorbital electron
transitions. Spectroscopic analyses revealed that PPy encapsulation
reduces the HOMO-LUMO bandgap from 1.64 eV (pure TCPP) to 1.28 eV
([228]Fig. 2, B to E), optimizing electron excitation and generating an
ultrasonic current with an amplitude of approximately 0.1 μA/cm^2. In
comparison, previously reported TCPP-based sonosensitizers exhibited a
smaller bandgap reduction (1.90 to 1.80 eV) and a lower sonic current
amplitude (~0.021 μA/cm^2) ([229]71), highlighting the markedly
superior performance of this system. The resultant ultrasonic current
induces rapid NPC membrane depolarization ([230]Fig. 4A), triggering
Ca^2+ influx through Ca[v] channels ([231]Fig. 4G). This Ca^2+ surge
activates CaMK1, which phosphorylates CDK1/2 ([232]Fig. 4, I and M),
directly coupling bioelectrical stimuli to cell cycle progression.
Notably, this pathway provides a direct mechanism to override G[0]-G[1]
arrest and enhance cell viability.
Beyond cell cycle modulation, the hydrogel’s ROS-scavenging capability
introduces a dual therapeutic axis. While borate ester–based systems
(TSPBA) ([233]35) passively degrade in oxidative stress environments,
US-enhanced cavitation in our hydrogel accelerates more significant ROS
neutralization and ferroptosis alleviation compared to static
conditions ([234]Fig. 5). Although prior studies using TSPBA-based
hydrogels demonstrated that ROS-responsive degradation plateaued within
2 hours ([235]72), our sonosensitive system, under US stimulation,
notably accelerated the scavenging kinetics to approximately 1 hour
([236]Fig. 5, A and B). This clearance effect not only preserves GPX4
([237]48) and SLC7A11 ([238]53) expression ([239]Figs. 5, F and H, and
[240]6I)—key suppressors of ferroptosis—but also reduces LPO and ACSL4
(A key driver of LPO) ([241]54) expression ([242]Figs. 5G and [243]6I).
Notably, previous studies suggest that Ca^2+-dependent pathways may
activate the Nrf2 antioxidant axis. Ca^2+/CAMKII signaling has been
shown to phosphorylate Nrf2, promoting its nuclear translocation and
transcriptional activation of detoxifying enzymes (e.g., HO-1 and GCLC)
that counteract LPO ([244]73, [245]74). Nrf2 activation is a
well-established suppressor of ferroptosis through GPX4 stabilization
and glutathione synthesis ([246]53, [247]75), aligning with our
observed ferroptosis rescue. These advances merge into a “sonoelectric
niche”—a dynamically tunable biochemical-physical microenvironment
([248]76, [249]77) where US-triggered electrical currents and
ROS-responsive hydrogel properties synergistically regulate cellular
signaling, enhancing endogenous regenerative pathways and enabling
noninvasive spatiotemporal control over tissue repair. While prior
studies leveraged piezoelectric scaffolds ([250]61) to modulate tissue
repair, none achieved spatiotemporal coordination of
electrophysiological and metabolic pathways. Our sonosensitive hydrogel
uniquely bridges this gap, simultaneously driving cell cycle
progression (via CDK1/2) and suppressing ferroptosis (via
GPX4/SLC7A11). This multimodal action explains the superior efficacy of
Gel + US over Gel alone in restoring disc height and ECM anabolism in
vivo ([251]Fig. 6).
Our sonosensitive hydrogel system represents marked advancements over
existing strategies for IVDD treatment in terms of noninvasive drug
delivery and treatment mode, material design, and therapeutic efficacy.
The IVD is located deep within the body. In this study, we developed a
noninvasive therapeutic strategy that involves precise puncture and
injection of an in situ gel-forming agent directly into the IVD under
x-ray fluoroscopic guidance ([252]Fig. 6A). The TCPP@PPy-PPy/PVA gel
responds to external US induction to treat IVDD. Unlike percutaneous
electrical stimulation, which struggles to target deep spinal
structures due to rapid current dissipation, our hydrogel localizes
microcurrents directly within the disc. Animal model outcomes further
validate the translational relevance of this strategy. Caprine lumbar
discs, which closely mimic human spinal biomechanics in terms of size,
intradiscal pressure, and range of motion ([253]51, [254]52), exhibited
approximately a 162.5% disc height index improvement based on MRI, a
46.7% improvement in Pfirrmann MRI score, and a 48.7% recovery in
histological grade based on histological staining, after Gel + US
treatment compared to IVDD ([255]Fig. 6, C to E). These therapeutic
outcomes not only basically matched conventional hydrogel treatments
effect ([256]4, [257]57, [258]58) but also demonstrated statistically
superior disc height index restoration compared to rodents (~118%)
([259]78) and sheep model (~16.5%) ([260]52). Our therapeutic strategy
uses noninvasive US-mediated spatiotemporal control to precisely
activate endogenous molecular pathways, driving histological
analysis–demonstrated substantial restoration of proteoglycan content
([261]Fig. 6, D, H, and I), and achieving functional tissue
regeneration—a critical regenerative metric markedly surpassing
conventional hydrogel-based or ROS-scavenging therapies ([262]52,
[263]72). The feasibility of this strategy in a goat IVDD model
highlights its potential for clinical translation.
To ensure optimal biological performance, biomaterials must exhibit
both in vivo biocompatibility and sufficient stability under
physiological conditions. For example, although glycine-based
sonosensitive piezoelectric biomaterials exhibit excellent
biocompatibility, unencapsulated formulations undergo complete
dissolution within 5 min in aqueous environments. Even after structural
optimization, rapid degradation occurs within hours to a few days,
accompanied by significant functional decay ([264]79–[265]81). In this
study, the hydrogel demonstrated low cytotoxicity ([266]Fig. 3A) and
robust in vivo biosafety ([267]Fig. 6). This biomaterial remained
stable under physiological conditions but underwent efficient
degradation under pathological ROS stimulation ([268]Fig. 1K),
indicating its responsiveness and controlled fate in disease
environments. Time-resolved FTIR and TEM analyses further confirmed the
structural and morphological stability of TCPP@PPy particles (fig. S1,
F and G), while ultrasonic current and PL measurements across time
gradients demonstrated their functional stability (fig. S3, A and B).
Collectively, these findings confirm that our hydrogel maintains
biosafety and stability, thereby ensuring effective therapeutic
performance.
While our study establishes a promising noninvasive strategy for IVDD,
several limitations require further attention. First, the approach does
not involve genetic modification of cells to provide a durable
self-regeneration capacity, implying that degenerated tissues might
necessitate prolonged in vitro US treatment sessions. Second, there is
a pressing need for increased sonoelectric efficiency to increase
treatment effectiveness. Future work should incorporate that
piezoelectric nanoparticles might enable self-sustaining current
generation under mechanical loading, mimicking physiological disc
motion. Furthermore, single-cell RNA sequencing of treated NPCs could
delineate subpopulation-specific responses to sonoelectronic stimuli,
identifying molecular checkpoints for personalized intervention. By
leveraging gene-editing platform, this integrated approach combines
gene-editing tools with tunable acoustic stimulation, enabling the
precise refinement of sonoelectronic parameters to spatiotemporally
regulate key electrochemistry signaling pathways and establishing a
framework for the real-time control of sonoelectric responsive gene
circuits.
In this study, we developed a sonoelectric hydrogel by synthesizing
sonosensitive particles of TCPP@PPy and incorporating them into a
PPy/PVA polymer, followed by cross-linking with the ROS-sensitive agent
TSPBA to form the hydrogel (TCPP@PPy-PPy/PVA). TCPP@PPy overcomes the
HOMO-LUMO gap to excite free electrons under US, resulting in the
formation of an ultrasonic current in hydrogel. This ultrasonic current
alters the membrane potential of NPCs, triggering Ca^2+ influx and
activating the Ca^2+-CaMK1-CDK1/2 signaling axis. This cascade advances
the cell cycle and promotes cell regeneration. Simultaneously, this
ROS-responsive hydrogel accelerates the clearance of ROS under US
exposure, mitigating ferroptosis caused by oxidative stress in cells.
This sonoelectric niche, which creates a dynamically regulated
biochemical-physical microenvironment through the sonoelectrical
effect, has demonstrated promising regenerative therapeutic effects in
a goat model of IVDD, offering a previously unexplored choice for
treating other deep-seated degenerative disorders, such as
osteoarthritis, where invasive interventions remain irreplaceable.
Future iterations of this technology will be enhanced through a
multidimensional optimization framework encompassing bandgap
engineering, genetic engineering, and patient-specific US protocols. In
summary, this convergence of sonosensitive hydrogels with noninvasive
spatiotemporal control establishes a notable therapeutic mode to
precisely orchestrate tissue regeneration, demonstrating transformative
potential for tissue regeneration.
MATERIALS AND METHODS
Synthesis of TCPP@PPy-PPy/PVA hydrogel
The synthesis of TCPP@PPy particles was performed according to a
previous report ([269]36) with minor modifications: A 20-ml aqueous
ethanol solution (50% v/v) was used to dissolve pyrrole (0.1 M) and
TCPP (1 mM). The mixture was subjected to ultrasonic homogenization for
1 hour, followed by the addition of 57 mg of ammonium persulfate (APS).
Then, the solution was mixed at room temperature for 24 hours.
Centrifugation of the mixture at 12,000 rpm for 5 min resulted in the
isolation of a purplish black product. Following a triple wash with
ethanol, the product was dried at 60°C overnight to obtain a powder
product (TCPP@PPy), which was subsequently stored in a desiccator.
The preparation of the TSPBA cross-linker was informed by an
established study ([270]35): Briefly, 0.1 g of
N,N,N′,N′-tetramethyl-1,3-propanediamine and 0.5 g of
4-(bromomethyl)phenylboronic acid were dissolved in 10 ml of
dimethylformamide, and the mixture was stirred overnight at 60°C. The
reaction mixture was transferred into 100 ml of tetrahydrofuran (THF)
and subjected to filtration, followed by successive washes with THF
(three portions of 20 ml each). The product was then dried under vacuum
overnight, yielding TSPBA.
A 10-ml solution of 9% PVA was prepared. Then, pyrrole and the oxidant
APS were added to the PVA solution to achieve a concentration of 0.1 M,
and the mixture was stirred at room temperature overnight to obtain a
PPy/PVA polymer solution. Briefly, 10 mg of TCPP@PPy powder was added
to 10 ml of PPy/PVA solution (1 mg/ml) and thoroughly dispersed via an
ultrasonic homogenizer. Subsequently, dual pipettes with closely fitted
tips were used to simultaneously deliver the TCPP@PPy-infused PPy/PVA
solution and the TSPBA solution into the plate wells, inducing rapid in
situ gelation and producing the TCPP@PPy-PPy/PVA hydrogel. To fabricate
the intermediate control TCPP/PPy/PVA hydrogel, 10 mg of TCPP powder
was added to 10 ml of PPy/PVA solution and thoroughly dispersed,
followed by cross-linking using the same method described previously.
Characterization
SEM (Zeiss Sigma 300)
TCPP@PPy powder samples were placed in a centrifuge tube with ethanol
and sonicated at room temperature for 5 min. A few droplets of the
resulting suspension were deposited onto a silicon wafer and allowed to
dry for analysis. The freeze-dried hydrogel was directly affixed to the
sample stage using conductive adhesive. The chamber was then vented,
and the sample was carefully positioned inside. The chamber was then
evacuated to achieve the required vacuum level. Once the desired vacuum
was attained, the voltage was increased. The sample location was
identified, and an appropriate magnification was selected.
High-resolution images were scanned and saved for further analysis.
TEM (JEOL JEM-F200)
The TCPP@PPy powder was resuspended in ethanol and ultrasonically
dispersed. A suitable amount of the dispersion was dropped onto a
sample grid and dried. The prepared sample was then placed on the
sample holder. Parameters such as the electron beam, aperture, focus,
and astigmatism were adjusted to achieve optimal brightness. The
specific location of the sample was identified for imaging.
FTIR (Nicolet iS 10)
In a dry environment, TCPP@PPy powder was mixed with potassium bromide
powder. The mixture was then pressed to form a pellet for use in
infrared spectroscopy analysis. For the hydrogel, an attenuated total
reflectance (ATR) accessory was positioned in the optical path of the
spectrometer. A background scan of the air was performed. The surface
of the hydrogel sample was pressed firmly against the crystal surface
of the ATR accessory, and the infrared spectrum of the hydrogel was
collected.
Compression test (EUT2000)
The hydrogel samples for testing were fabricated using a mold to
achieve dimensions of 5 mm by 8 mm (radius by thickness). Compression
tests were conducted at a crosshead speed of 10 mm/min, and the
acquired data were used to calculate the compressive stress curve.
Swelling
The initial mass of the hydrogel was recorded as W[d]. The hydrogel is
then immersed in water and weighed at regular intervals until it
reaches swelling equilibrium, with the mass at a given point recorded
as W[s]. The equilibrium swelling ratio is calculated via the following
formula
[MATH: Swelling ratio=(Ws−Wd)Wd×100% :MATH]
Degeneration
The hydrogel is placed in a 20 mM solution of H[2]O[2] or PBS. The
initial mass is recorded as W[d], and the mass at a certain time point
is recorded as W[s]. The degradation rate of the gel can be calculated
using the following formula and plotted as a curve
[MATH: Degradation ratio=(Wd−Ws)Wd×100% :MATH]
Electrochemistry
Under a 1.2 W/cm^2 US irradiation regime, the ultrasonic current
induced in the sample was measured using a standard three-electrode
system in an electrochemical workstation (CH Instruments, 600E).
Briefly, a 1 mM solution of TCPP@PPy was mixed with a 5% Nafion
solution at a volume ratio of 10:1. Aliquots of 200 μl of this mixture
were drop-cast onto indium tin oxide (ITO)–coated glass (15 mm by 30 mm
by 1.1 mm) and allowed to dry. This process was repeated three times.
The hydrogel samples were gelled in situ on ITO glass. The ITO glass,
affixed to the working electrode, was immersed alongside a platinum
electrode and a reference electrode in a 0.5 M Na[2]SO[4] solution. A
US probe was used to irradiate the sample cell with cycles of 20 s per
switch. The electrochemical workstation captured the ultrasonic current
generated within the system, and subsequent data analysis was conducted
using CHI version 22.01 software.
Photoluminescence
The PL properties of the powder and hydrogel samples were characterized
using a fluorescence spectrophotometry (PerkinElmer, LS55). The sample
was prepared by dispersing TCPP or TCPP@PPy in a suitable solvent to
ensure a homogeneous suspension. The hydrogel samples were directly
used for testing. The excitation wavelength was selected on the basis
of the maximum absorption peak identified from the UV-Vis absorption
spectra. The emission spectra were recorded over a range spanning the
characteristic emission profile of the sample. All measurements were
performed at a controlled temperature, and the sample was allowed to
equilibrate for a defined period before data collection. The PL quantum
yield was determined relative to a standard reference compound with a
known quantum yield, following the protocol outlined in the IUPAC
Technical Report ([271]82).
Ultraviolet-visible
The UV-vis absorption properties of the TCPP and TCPP/PPy powders were
characterized via a standard spectrophotometer (UV-3600). A precisely
measured quantity of powder sample was dispersed in a suitable solvent
to ensure a homogeneous solution. The resulting solution was then
placed in a quartz cuvette, and the absorbance was measured over a
wavelength range from UV to the visible spectrum. The spectrophotometer
was calibrated with a blank solvent sample to correct for any
background absorbance. The data were analyzed to determine the
absorption maxima and molar absorptivity, providing insights into the
electronic transitions within the TCPP and TCPP/PPy molecules.
Conductivity
The electrical conductivity of the hydrogel was measured using a
four-point probe method. The hydrogel samples were prepared with a
defined cube and equilibrated in a controlled environment to maintain
consistent hydration. The measured PVA, PPy/PVA, TCPP/PPy/PVA, and
TCPP@PPy-PPy/PVA hydrogels had lengths and widths of 5 mm, and the
thickness (L) was 8 mm. The resistance (R) of the hydrogel was
measured, and the cross-sectional area (S) was calculated. The
resistivity (ρ) of the hydrogel was calculated using the following
formula
[MATH: ρ=RSL :MATH]
The conductivity (σ) of the hydrogel was calculated using the following
formula
[MATH: σ=1ρ :MATH]
NPCs isolation
NPCs were harvested using a previously reported pronase-collagenase
digestion method ([272]83). NP tissues were obtained from discectomy
procedures following informed consent. Under sterile conditions,
surgical specimens were meticulously separated from adjacent
cartilaginous and AF components, followed by mechanical dissociation
into 1- to 2-mm^3 fragments. Tissue digestion was performed using a
0.3% collagenase II/0.2% pronase solution with agitation at 37°C for
3.5 ± 0.5 hours. The obtained cell suspension underwent gradient
centrifugation (1000g, 10 min) and triple PBS washing before
resuspension in F-12 medium supplemented with 10% FBS and
penicillin-streptomycin. Primary cultures were maintained under
standard conditions (5% CO[2] and 37°C) with media changes every
72 hours, limiting cell expansion to passage 2 to preserve phenotype
integrity. This protocol received institutional review board approval
by the Ethics Committee of Tongji Medical College, Huazhong University
of Science and Technology (no. S341), in full compliance with Helsinki
Declaration guidelines.
Cell staining
Live/Dead
The NPCs were resuspended in a solution of PPy/PVA mixed with TCPP@PPy
and then cross-linked with TSPBA to obtain a 3D culture system of NPCs
in a TCPP@PPy-PPy/PVA hydrogel, ultimately reaching a concentration of
2.5 × 10^6 cells/ml, as described in a reported protocol for NPC 3D
culture ([273]44). F12 medium was added to culture the cells. After
various treatments, the NPC-loaded hydrogels were incubated with
calcein AM/propidium iodide (PI) (Beyotime) working solution at 37°C
for 30 min. 3D fluorescence images were obtained using a confocal laser
microscope (Nikon C2Si).
Cell skeleton
The hydrogel was shaped into thin slices via a mold and then overlaid
onto the cells cultured on a six-well plate to facilitate contact with
the cells. Briefly, apoptosis was induced in NPCs upon treatment with
200 μM TBHP for 24 hours. Then, the cells were treated daily with US,
Gel, or Gel + US for 15 min for 3 days. The cells were washed with PBS
three times and fixed for 10 min with 4% paraformaldehyde. The NPCs
were subsequently incubated with fluorescein isothiocyanate–labeled
phalloidin (Solarbio) for 40 min, and the cell nuclei were subsequently
stained with 4′,6-diamidino-2-phenylindole (DAPI). Fluorescence images
were obtained using a fluorescence microscope (Olympus, IX71).
Alcian
The treatment of the cells was consistent with the aforementioned
methods. The cells were pretreated with 3% acetic acid for 3 min at
room temperature to ensure an appropriate acidic environment.
Subsequently, 1% Alcian blue solution was added, and the mixture was
incubated for 30 min. The cells were then washed two to three times
with a 3% acetic acid solution, followed by rinsing with distilled
water. The staining results were observed under a microscope.
Immunofluorescence
We established a 3D culture system for NPCs using PVA or
TCPP@PPy-PPy/PVA hydrogels. After 24 hours of induction with 200 μM
TBHP, the NPC-gel compound underwent daily 15-min treatments with US
for 3 days or was maintained without intervention. After three washes
with PBS, the NPC-gel compounds were fixed with a 4% paraformaldehyde
solution. The samples were blocked with bovine serum albumin (BSA) at
room temperature for 60 min to minimize nonspecific interactions, after
which they were stained with rabbit anti-collagen 2 (Bioss Antibodies,
bs-0709R) overnight at 4°C. The samples were then incubated with an
Alexa Fluor 594–conjugated goat anti-rabbit immunoglobulin G secondary
antibody (Abcam, ab150088) for 60 min. Nuclei were stained with DAPI. A
confocal microscope was used to scan the samples.
Lipid peroxidation
The hydrogel was shaped into thin slices via a mold and then overlaid
onto the cells cultured on a six-well plate to facilitate contact with
the cells. After the induction of NPC apoptosis with 200 μM TBHP for
24 hours, which was followed by daily treatment with Gel or Gel + US
for 15 min for 3 days, the cells were washed with PBS three times.
Next, a Lipid Droplets Green Fluorescence Assay Kit with BODIPY 493/503
(Beyotime) was used to stain the cells for 30 min. Fluorescence images
were obtained using a confocal microscope.
Acridine orange
The treatment of the cells was the same as that described above. To
ensure the integrity of the lysosomal membrane, an AO staining kit
(Beyotime) was used to stain the cells for 20 min, followed by image
acquisition via confocal microscopy.
Fluo-4 assay
Following the aforementioned treatments, for fluorescence microscopy
detection, NPCs were washed once with PBS. Fluo-4 staining solution was
then added. The cells were incubated at 37°C in the dark for 30 min.
After incubation, the cells were washed three times with PBS. The
staining effect was observed under a fluorescence microscope
(λ[ex] = 490 nm, λ[em] = 525 nm). NPCs collected at various time points
were assessed with a microplate reader (λ[ex] = 490 nm, λ[em] = 525
nm).
Reverse transcription qPCR
The hydrogel was shaped into thin slices using a mold and then overlaid
onto the cells cultured on a six-well plate to facilitate contact with
the cells. Following various treatments for NPCs, RNA was extracted
from the cells using TRIzol reagent (Invitrogen) and then reverse
transcribed into cDNA. RT-qPCR was subsequently conducted for
quantitative analysis of gene expression, and
glyceraldehyde-3-phosphate dehydrogenase served as an internal control
to normalize the expression levels. The relative expression levels of
the target genes were determined using the 2^-ΔΔCT method. The primers
used in this study are listed in table S1.
Western blot
Posttreatment, the NPCs were washed twice with PBS and subsequently
incubated with radioimmunoprecipitation assay buffer supplemented with
1% phenylmethylsulfonyl fluoride, a protease inhibitor. The cells were
lysed on ice for 15 min, ultrasonically crushed for 30 s, and
centrifuged at 13,000g for 15 min at 4°C. The lysed supernatant was
aspirated into prechilled EP tubes, and the protein concentration of
the cell lysate was determined using a BCA protein assay kit. Forty
micrograms of protein was separated via 8 to 12% SDS–polyacrylamide gel
electrophoresis and transferred onto 0.22- or 0.45-nm polyvinylidene
difluoride (PVDF) membranes according to the molecular weight of the
proteins. The PVDF membrane was incubated in 5% BSA for 1 hour to block
nonspecific binding sites, followed by washing with 0.1% Tris-buffered
saline with Tween (TBST) four times. The membrane was then incubated
with primary antibodies against P16 (AF5484, Affinity), P21
(10355-1-AP, Proteintech), interleukin -1B (IL-1B) (26048-1-AP,
Proteintech), IL-6 (DF6087, Affinity), CCL2 (DF7577, Affinity), tumor
necrosis factor–α (AF7014, Affinity), COL2 (AF5456, Affinity), MMP13
(18165-1-AP, Proteintech), ADAMTS5 (DF13268, Affinity), CaMK1 (DF7805,
Affinity), phospho-CaMK1 (Thr^177, AF7381, Affinity), CDK1 (10762-1-AP,
Proteintech), phospho-CDK1/CDC2 (Thr^161, AF8001, Affinity), CDK2
(10122-1-AP, Proteintech), or phospho-CDK2 (Thr^160, AF3237, Affinity)
overnight at 4°C and subsequently with secondary antibodies for 1 hour
at room temperature. Following four rinses with 0.1% TBST to eliminate
nonspecific antibody binding, the membrane was subjected to
chemiluminescent detection with a substrate. The resulting luminescent
signals were then accurately captured via a ChemiDoc imaging system
(Bio-Rad).
Membrane depolarization assay
The activity of sonosensitive hydrogel-induced membrane depolarization
was assessed by quantifying the fluorescence emitted by the membrane
potential–sensitive dye DiSC3(5). Following the aforementioned
treatments, the NPC samples from each time point were centrifuged,
washed twice with washing buffer [20 mM glucose and 5 mM Hepes (pH
7.2)], and resuspended in the same buffer [20 mM glucose, 5 mM Hepes
(pH 7.2)] containing 0.1 M KCl to a final optical density at 600 nm of
0.05. The cells (100 μl) were subsequently incubated with 20 nM
DiSC3(5) for 15 min until a stable decrease in fluorescence was
observed, indicating that the dye had integrated into the cell
membrane. Membrane depolarization was monitored by observing the
changes in the fluorescence emission intensity of DiSC3(5) (λ[ex] = 622
nm, λ[em] = 670 nm).
RNA sequencing
Total NPC RNA from the control, TBHP, TBHP + Gel, and TBHP + Gel + US
groups was extracted via TRIzol Reagent (Invitrogen). After quality and
integrity were assessed, the RNA samples were quantified, and RNA
sequencing libraries were constructed. The raw sequencing data were
curated in SeqHealth (Wuhan, China) using STRA software (version
2.5.3a) to map the reads onto the human reference genome under default
parameters. Differential gene expression across groups was analyzed
using the dgeR package (version 3.12.1), which uses a P value threshold
of 0.05 and a fold change magnitude of 2 to define statistical
significance. Subsequent functional enrichment analyses and KEGG
pathway analysis were performed with KOBAS software (version 2.1.1),
and a P value cutoff of 0.05 was used to discern significant
enrichment.
Flow
An ROS assay kit (Beyotime) was used to detect the ROS content in the
NPCs. The hydrogel was shaped into thin slices via a mold and then
overlaid onto the NPCs cultured on a six-well plate to facilitate
contact with the cells. After the induction of NPC apoptosis with 200
μM TBHP for 24 hours, the NPCs were treated daily with US, Gel, or Gel
+ US for 15 min for 3 days. NPCs were washed three times with PBS,
followed by the addition of 2’,7’-dichlorodihydrofluorescein diacetate
(DCFH-DA) diluted in serum-free medium at a 1:1000 ratio to yield a
final concentration of 10 μM. The cells were then incubated at 37°C for
20 min. After incubation, the cells were washed three times with PBS to
remove excess DCFH-DA. The fluorescence was measured using a flow
cytometer (λ[ex] = 488 nm, λ[em] = 525 nm).
Cell cycle
A Cell Cycle and Apoptosis Analysis Kit (Beyotime) was used to assess
the NPC cell cycle. Following treatment, the NPCs were washed and
centrifuged, resuspended in 1 ml of ice-chilled 70% ethanol, and fixed
at 4°C for more than 30 min. After centrifugation at 1000g for 5 min,
the NPCs were washed with 1 ml of ice-chilled PBS and centrifuged again
to remove the supernatant. Briefly, 0.5 ml of PI solution was added to
each tube, and the cell pellets were gently and thoroughly resuspended,
followed by incubation in the dark at 37°C for 30 min. Red fluorescence
was detected using a flow cytometer with excitation at a wavelength of
488 nm.
Animals
The animal experiment protocol was approved by the Ethics Committee of
the Animal Experiment Center, Tongji Medical College, Huazhong
University of Science and Technology (S2895), and the study was
conducted under the principle of blinding. Twelve 6-month-old male Boer
goats, randomly and evenly divided into four groups—Control, Decomp,
Decomp + Gel, and Decomp + Gel + US—were acclimated in a separate room
under quiet conditions for a period of 2 weeks before undergoing Decomp
surgery for IVDD modeling. During the surgical procedure, anesthesia
was induced and maintained via continuous mask administration of
isoflurane gas. The surgery was performed via a posterior approach
through the goats’ lumbar region, involving gradual dissection of the
skin, fascia, and muscles, followed by exploration of the lumbar
vertebrae. The surgical procedure was performed in a manner that
minimized the size of the incision, tissue dissection, and overall
trauma. For goats in the Decomp, Decomp + Gel, and Decomp + Gel + US
groups, a scalpel was used to puncture the L[3]-L[4] and L[4]-L[5]
IVDs, after which NP forceps were used to extract equivalent amounts of
NP tissue, achieving the desired decompression. The control goats
underwent a sham operation. The incised muscles and epithelium of the
goats were sutured at the end of surgery. On the 7th day after surgery,
for the goats in the Decomp + Gel and Decomp + Gel + US groups, two
puncture needles were used to inject the TCPP@PPy-PPy/PVA solution and
the TSPBA cross-linker into the IVD, allowing for in situ gelation. The
procedure was performed under real-time x-ray fluoroscopy (DTP570A,
Anjian, China) monitoring using DXRay Diagnostic software. On days 7,
14, 21, and 28, Decomp + Gel + US goats received 30 min of US
irradiation (1.2 W/cm^2) in the corresponding lumbar region. On day 42
after surgery, all the goats were anesthetized for MRI (WANDONG,
i_Space 1.5 T) and CT (Canon, Tsx303A) analysis, followed by
euthanasia. IVD tissue samples from the goats were collected, and hard
tissue sections were prepared for RT-qPCR and histological examination
(Safranin O, H&E, and Masson staining).
Statistical analysis
The data are expressed as the means ± SDs and were analyzed using
either Origin 8 or GraphPad Prism 8 software. All experiments were
biologically repeated at least three times. The statistical
significance of variance was assessed via one-way or two-way analysis
of variance (ANOVA). P > 0.05 was classified as not statistically
significant (ns), whereas P < 0.05 denoted statistical significance.
Acknowledgments