Abstract
Hepatocellular carcinoma (HCC) is a leading cause of cancer-related
mortality, with high postoperative recurrence rates due to occult
micrometastases or minimal residual disease, markedly worsening the
prognosis for HCC patients. Current therapies lack effective strategies
to prevent recurrence, while traditional Chinese medicine (TCM) shows
potential in delaying HCC progression. Combining a hemostatic hydrogel
with nanoparticle-based delivery of active TCM components provides a
strategy to enhance tumor recurrence prevention. Herein, we develop a
nanocomposite hydrogel (HPS@ZCJ) by encapsulating Jianpi-Huayu
essential oils (JEO) into zein-based nanoparticles (zein@chondroitin
sulfate@JEO, ZCJ) and embedding them in a hydroxymethyl
cellulose/Pluronic F-127/sodium alginate (HPS) hydrogel matrix. HPS@ZCJ
hydrogel enhances cytotoxic T-lymphocyte infiltration, inhibits the
polarization of tumor-associated macrophages to M2 phenotype, induces
tumor cell death, reverses immunosuppression, and inhibits angiogenesis
within the tumor. The antitumor mechanism involves dual downregulation
of GPNMB and DHCR7, key genes in HCC progression and immune evasion. In
vitro and in vivo experiments demonstrate that HPS@ZCJ
hydrogel-mediated targeted comprehensive therapy simultaneously
achieves intraoperative hemostasis, impedes primary tumor growth and
prevents HCC postoperative recurrence. This study provides a promising
postoperative HCC treatment strategy, leveraging TCM's therapeutic
potential with significant clinical translation prospects.
Keywords: Nanoparticle drug delivery, Jianpi-Huayu decoction, Essential
oils, Postoperative treatment, Hepatocellular carcinoma, Hydrogel
Graphical abstract
Image 1
[41]Open in a new tab
1. Introduction
Hepatocellular carcinoma (HCC) ranks as the third-leading cause of
cancer mortality worldwide, with a dismal 5-year survival rate below
18 % [[42]1]. While surgical resection remains the gold-standard
treatment for early-stage HCC [[43]2], up to 70 % of patients,
including those with small, solitary tumor nodules (≤2 cm), experience
recurrence within 5 years [[44]3]. This high recurrence rate stems
primarily from occult micrometastases and minimal residual disease that
enter circulation during intraoperative bleeding [[45]4]. Notably,
current clinical guidelines lack effective pharmacological
interventions for postoperative recurrence prevention [[46]5],
highlighting an urgent unmet need in HCC management.
Traditional Chinese medicine (TCM) has shown promise in managing HCC,
particularly in delaying progression and reducing recurrence, either
alone or in conjunction with other conventional therapies. Among TCM
formulas, Jianpi-Huayu decoction (JHD)-composed of Baizhu (Rhizoma
Atractylodis Macrocephalae), Ezhu (Curcuma zedoaria Roscoe), Fuling
(Poria cocos), Foshou (fingered citron), Kushen (Radix Sophorae
Flavescentis), and Baihuasheshecao (Hedyotis diffusa Willd)-exerts
direct cytotoxicity against HCC cells, inhibits angiogenesis, and
modulates immunity [[47][6], [48][7], [49][8]]. Notably, the essential
oils derived from JHD (JEO)-rich in monoterpenes and sesquiterpenes-are
key bioactive components with demonstrated anticancer and
immunoregulatory properties [[50][9], [51][10], [52][11], [53][12],
[54][13], [55][14]]. However, the clinical translation of JEO is
hindered by poor solubility, which limits its bioavailability and
administration routes. Thus, developing advanced delivery systems for
JEO could improve its bioavailability and enhance HCC therapy.
Nanoparticle (NP) delivery systems have shown promise for overcoming
the poor solubility of JEO [[56]15,[57]16]. Among them, zein NPs
exhibit distinct advantages: (1) high loading capacity for lipophilic
drugs, compatible with JEO's essential oils [[58]17]; (2) γ-zein's
N-terminal proline-rich domain promotes cell membrane interaction,
enhancing cellular uptake [[59]18]. However, conventional intravenous
administration of NPs risks redistribution and side-effects, while
locally injected particles can migrate or prematurely release
therapeutic agents [[60]19,[61]20]. Therefore, engineering zein NPs for
localized JEO delivery with prolonged tumor retention is essential to
maximize therapeutic efficacy while minimizing off-target effects.
Hydrogels have emerged as attractive platforms for local drug delivery
due to their biocompatibility, biodegradability, and ability to adhere
to target sites, yet loading hydrophobic agents remains problematic
[[62]21,[63]22]. This challenge can be addressed through nanocomposite
hydrogels that combine the advantages of drug-loaded NPs with hydrogel
matrices [[64]19]. In the context of HCC treatment, this approach
becomes particularly relevant given the clinical challenges associated
with tumor resection, including intraoperative bleeding and high
postoperative recurrence rates due to potential micrometastasis
[[65]23].
In this study, we introduce an innovative solution through the
development of a thermosensitive composite hydrogel system composed of
hydroxypropyl methylcellulose (HMC), Pluronic F-127 (PF127), and sodium
alginate (SA), which was termed HPS (HMC-PF127-SA). This formulation
was specifically designed to address multiple clinical needs
simultaneously. HMC contributes crucial hemostatic properties through
its branched fiber network and ability to promote thrombus formation at
vascular injury sites [[66][24], [67][25], [68][26]]. PF127 provides
temperature-responsive gelation behavior, enabling convenient in situ
application during surgical procedures [[69]27]. SA complements HMC's
bioadhesive function while strengthening the hydrogel's structural
integrity [[70]28]. In addition, the system was further optimized by
incorporating JEO-loaded zein nanoparticles stabilized with chondroitin
sulfate (CS), designated as Zein-CS@JEO (ZCJ). The CS modification
maintains the NPs'cell-interaction capabilities while improving their
stability within the hydrogel matrix [[71]29].
The complete hydrogel system, incorporating the ZCJ NPs, was termed
HPS@ZCJ ([72]Fig. 1). The in-situ-formed HPS@ZCJ hydrogel enables
intraoperative injection and provides three key clinical benefits for
postoperative HCC management: (1) immediate hemostatic action to reduce
the risk of tumor cell dissemination; (2) precise local delivery of
therapeutic JEO components, minimizing loss and off-target toxicity;
(3) controlled release kinetics through the combined
nanoparticle-hydrogel architecture. This therapeutic strategy
specifically targets early-stage HCC (BCLC stages 0 and A), where
surgical resection remains the primary treatment option but faces
significant challenges from postoperative recurrence [[73]30]. The high
recurrence rates in these stages, often due to occult micrometastases
or minimal residual disease, highlight the need for effective
postoperative therapies to prevent recurrence [[74]31]. In summary, the
HPS@ZCJ hydrogel presents a treatment strategy for HCC based on
TCM-inspired therapy, synergistically combined with physical barriers
against tumor cell spread to effectively prevent post-resection HCC
recurrence.
Fig. 1.
[75]Fig. 1
[76]Open in a new tab
Schematic of preparing HPS@ZCJ hydrogel and combination therapy for
postoperative recurrence of HCC. Created with [77]BioRender.com.
2. Materials and methods
2.1. Preparation and component analysis of JEO
The JHD is formulated from six traditional Chinese medicinal herbs:
Ezhu (Curcuma Zedoaria Roscoe), Baizhu (Rhizoma Atractylodis
Macrocephalae), Foshou (Fingered Citron), Fuling (Poria Cocos), Kushen
(Radix Sophorae Flavescentis), Baihuasheshecao (Hedyotis Diffusa
Willd). These herbs are combined in a proportion of 3:3:3:5:5:5. The
herbs are then cut or ground into small particles using scissors or
herbal grinder. Next, the herbal mixture is placed into a round-bottom
flask and soaked in ultrapure water at a volume ratio of 1:8 (herbs:
water) for 6 h. JEO is subsequently extracted via steam distillation.
The extracted JEO is then sealed with a paraffin film and stored at
4 °C in a light-protected refrigerator for future use.
The chemical constituents of JEO were analyzed using high-resolution
liquid chromatography-mass spectrometry (HRLC-MS) and gas
chromatography-mass spectrometry (GC-MS). Specifically, 100 μL of the
sample was combined with 500 μL of methanol, mixed thoroughly, and
centrifuged at 13,000 rpm for 10 min. The resulting supernatant was
subsequently used for detection.
The liquid chromatography system utilized was an UltiMate 3000 UHPLC.
The chromatographic column was a Thermo Hypersil gold column (1.9 μm,
2.1 mm × 100 mm). The mobile phase was composed of 0.1 % formic acid in
acetonitrile (B) and 0.1 % formic acid in water (A). The flow rate for
chromatographic analysis was maintained at 0.3 mL/min. The gradient
elution program was as follows: starting at 10 % B, the proportion of B
was increased to 100 % over 10 min; 100 % B was held for 7 min, then
returned to the initial condition within 0.1 min, followed by column
equilibration for 2.9 min. The total runtime was 20 min.
The mass spectrometry instrument used was a Q-Exactive (Thermo Fisher
Scientific, CA, USA) equipped with the HESI source. The ion source
temperature was set at 310 °C, the capillary temperature at 320 °C, the
sheath gas flow rate at 30 units, and the auxiliary gas flow rate at 10
units. The spray voltage was 3 kV in positive ion mode and 2.8 kV in
negative ion mode. Data-dependent acquisition (DDA) was employed with a
loop count of 10. The HCD energy was configured with step-wise
normalized collision energy values of 10, 28, and 35 eV. The
first-order mass spectrometry scan range was 100–1500 m/z, with a
resolution of 70,000, an AGC target of 3E6, and an injection time of
200 ms. For the second-order mass spectrometry, the resolution was
17,500, the AGC target was 1E5, and the injection time was 50 ms.
The gas chromatograph employed was an Agilent 7890 B-5977, fitted with
an HP-5MS column (30 m × 0.25 mm × 0.25 μm). Helium (He) was used as
the carrier gas in constant flow mode at a rate of 1 mL/min. The column
temperature was programmed as follows: the initial temperature was set
to 60 °C, then ramped up to 100 °C at a rate of 20 °C/min; after
holding at 100 °C for 1 min, the temperature was further increased to
300 °C at a rate of 12 °C/min and maintained for 1 min. The total
runtime was 20.667 min. The injection port temperature was maintained
at 280 °C. The injection volume was 1 μL, with a split ratio of 5:1.
The mass spectrometer utilized an electron impact ionization source
with an electron energy of 70 eV, an ion source temperature of 220 °C,
and a transfer line temperature of 280 °C. The solvent delay was set to
2.5 min, and the mass spectrometer operated in full scan mode with a
mass range of 10–650 amu.
The HRLC-MS data were analyzed for potential chemical constituents
using the Compound Discoverer software (V 3.2, Thermo Fisher
Scientific, CA, USA) through an automated database search. The GC-MS
data were processed using the Mass Hunter software (VB.07.00) to
identify compounds via integration. The search parameters were set as
follows: signal-to-noise ratio (SNR) at 2; sharpness threshold at 25 %;
absolute height at 500 counts; and relative height at 0.1 %. The NIST
11 database was employed for automatic compound identification, with a
score threshold of 65, absolute height set at 100 counts, and relative
height at 0.5 %. To ensure batch-to-batch reproducibility, the JEO was
extracted by the same steam-distillation protocol and analyzed with
identical HRLC-MS and GC-MS conditions for every preparation. All
approach aligns with the Chinese Pharmacopoeia guidelines for
essential-oil standardization and support reproducible biological
outcomes.
2.2. Preparation of ZCJ NPs
ZCJ NPs were prepared at various zein-to-CS weight ratios using a
solvent displacement method ([78]Table S1). Zein (10 mg, Gibco) and JEO
(1 mg) were dissolved in a co-solvent mixture of ethanol and
double-deionized water (DDW) (3:1 v/v, 2 mL). CS (0, 2, 5, or 10 mg,
Sigma) was dissolved in DDW (2 mL) and added dropwise to the zein
solution under probe sonication at 20 % amplitude for 20 s. The
resulting NP suspensions were dialyzed against DDW using a dialysis bag
with a molecular weight cut-off (MWCO) of 6–8 kDa (Seguin, USA) for
12 h to remove residual ethanol and unloaded JEO. The dialyzed products
were then filtered through a syringe filter with a pore size of 0.45 μm
(Goettingen, Germany) to remove any residual particulate matter or
large aggregates. The final NP compositions are detailed in [79]Table
S1. Blank NPs were prepared using the same method but without adding
JEO.
2.3. Characterization of the ZCJ NPs
The surface morphology of the ZCJ was investigated using Scanning
electron microscopy (SEM, FEI, USA). The core–shell structure was
visualized via Transmission Electron Microscope (TEM, Talos, USA). The
mean particle diameter, PDI, and ζ potential of the nanospheres were
assessed by dynamic light scattering at 25 °C using a Zetasizer Nano
ZS90 (Malvern Instruments, UK). Each sample was analyzed at least three
times, and the results are expressed as the mean ± standard deviation.
Additionally, the chemical structures of the samples were documented
using a Fourier Transform Infrared Spectroscopy (FTIR) spectrometer
(Bruker, Horiba, Germany).
2.4. Preparation of HPS@ZCJ hydrogel
To prepare a 20 % PF127 solution, 10 g of PF127 (Sigma, USA) was
dissolved in 50 mL of sterile deionized water and stirred at 4 °C until
completely dissolved. Subsequently, 10 mL of this solution was mixed
with 400 mg of SA (96 kDa, with an M/G ratio of 1.2 as determined by
FTIR analysis, Sigma), 400 mg of HMC (Macklin, China), and 200 mg of
ZCJ. The mixture was stirred at 300 rpm for 1 h to form the HPS@ZCJ
hydrogel. For the preparation of the blank hydrogel (HPS), the same
procedure was followed, but ZCJ was omitted. In the study, the hydrogel
was administered using a 1 mL syringe equipped with a 21G needle. The
choice of a 21G needle is based on clinical practice, where 10 mL
syringes with 21G needles are commonly used in HCC surgeries. This
ensures compatibility with existing surgical procedures and equipment.
2.5. Characterization of the HPS@ZCJ hydrogel
2.5.1. Microstructure of HPS@ZCJ hydrogel
The samples were freeze-dried with a lyophilizer after preparation.
Following freeze-drying, liquid nitrogen was employed to fracture the
samples, revealing a clean cross-section and the internal structure.
Subsequently, the samples were affixed to a stage using conductive
adhesive tape and coated with gold via a sputter coater at a current of
15 mA for 60 s. Ultimately, the internal structure of the samples was
scrutinized with SEM (FEI, USA).
2.5.2. Rheological test
The rheological characteristics of the samples were assessed utilizing
a rheometer (HAAKE MARS III, Thermo Fisher Scientific, America).
Hydrogel samples were situated between parallel plates (diameter 60 mm,
depth 0.3 mm), and temperature and frequency sweeps were executed on
the samples. The measurement parameters, including amplitude sweep,
were defined according to the linear viscoelastic region of the storage
modulus (G′) and loss modulus (G″). The temperature sweep spanned from
4 to 43 °C at a heating rate of 0.05 °C/s. The frequency sweep ranged
from 0 to 100 rad/s. During viscosity assessments, a shear rate ranging
from 0.01 to 100 s^−1 was applied to evaluate the shear-thinning
characteristics of the gels.
2.5.3. FTIR analysis
The samples underwent freeze-drying via a lyophilizer. Following this,
the samples were pulverized into a fine powder using a mortar and
pestle, after which the powder was gathered. The infrared spectra of
the samples were captured using the FTIR spectrometer (Bruker Vertex
70v, Bruker, Germany) across the range of 4000–400 cm^−1, at a
resolution of 4 cm^−1.
2.5.4. Swelling ratio
The hydrogels were submerged in simulated body fluid (SBF, pH = 7.4) at
a temperature of 37 °C. At set time points (0.5, 1, 2, 4, 8, 16, 24,
and 32 h), the samples were extracted. The surface moisture was dabbed
with filter paper, and the samples were subsequently weighed precisely
to ascertain the swelling ratio.
2.5.5. Adhesive test of the hydrogels
To examine the adhesive strength of the HPS@ZCJ hydrogel, wooden sticks
were chosen as the substrate in this study. The detailed process is
described as follows: Initially, wooden sticks (hydrophilic substrate)
measuring 60 × 5 mm were prepared. Next, at room temperature, 20 μL of
the hydrogel solution was applied to a 5 × 20 mm area at one end of
each wooden stick. The hydrogel-coated ends of the two wooden sticks
were then joined together and maintained at 37 °C for 10 min to allow
curing. The shear strength was subsequently measured using a universal
tensile testing machine (3365 Instron, USA) to evaluate the adhesive
performance of the hydrogel. A PF127 hydrogel (20 %) was used as a
control. Additionally, the experiment was repeated using iron blocks
(hydrophobic substrate). Each test was conducted with three replicates.
2.5.6. In vitro degradation tests
The HPS@ZCJ hydrogel was placed in SBF at 37 °C and immersed
continuously for 21 days. At specified intervals, the hydrogels were
taken out, cleaned with ultrapure water, subjected to freeze-drying,
and weighed afterward. The extent of weight loss for the hydrogels was
quantified using the formula below:
[MATH: Weightlossrate(%)=Wt/Wo×100(%) :MATH]
Here, Wo is the initial dry weight before degradation, while Wt is the
dry weight after degradation.
2.5.7. Encapsulation efficiency (EE) and in vitro drug release test
Precise amounts of JEO were dissolved in PBS (1 % DMSO) to formulate
standard solutions with diverse concentrations. The absorbance of these
solutions at distinct concentrations was assessed using a UV–Vis
spectrophotometer (UV-2600, SHIMADZU, Japan), and calibration curves
were generated. For JEO, the peak absorption wavelength was 274 nm.
Freshly prepared ZCJ NPs were centrifuged at 15,000g in a refrigerated
centrifuge (Thermo, USA) at 25 °C for 30 min. The supernatant was
diluted with PBS containing 1 % DMSO, and the absorbance at 274 nm was
measured using a UV-2600 spectrophotometer. The content of JEO was
quantified against a standard curve of JEO dissolved in PBS (1 % DMSO).
The EE was calculated using the following equations:
[MATH: EE(%)=AmountofencapsulatedJEOofTotalJEOamount×100(%) :MATH]
Moreover, at defined time intervals, 1 mL of the solution was extracted
from the centrifuge tubes and placed into separate Eppendorf tubes,
with 1 mL of fresh PBS (1 % DMSO) replenished into the solution. The
absorbance of the extracted solutions at various time points was
monitored using the UV–Vis spectrophotometer, and the outcomes were
juxtaposed with the calibration curves. Drug release profiles were
constructed to elucidate the kinetics of drug release.
2.6. H22 cancer cells killing effect and biocompatibility assay
To begin with, ZCJ NPs at concentrations of 250, 500, 1000, 2000, and
4000 μg/mL were fully dissolved in corresponding volumes of RPMI-1640
(Gibco) culture medium with 10 % fetal bovine serum (FBS).
Subsequently, the proliferation of H22 cells (3 × 10^3 cells per well)
was evaluated using the Cell Counting Kit-8 (CCK-8) after 1, 2, and 3
days of incubation with ZCJ at these concentrations, with RPMI-1640
alone as the control. Briefly, the CCK-8 reagent was added to the
culture medium and incubated with the cells for 1 h. The absorbance was
then measured at 450 nm using a microplate reader (Bio-Tek, USA). Based
on these findings, the ideal concentration of ZCJ for further cell
proliferation studies was identified.
H22 cells and human umbilical vein endothelial cells (HUVECs) were
plated in 96-well plates (3 × 10^3 cells per well). Once the cells
adhered the following day, they were treated with the growing
concentration of HPS@ZCJ hydrogel for 48 h. In the same way, the IC[50]
values were subsequently assessed using CCK-8 assays.
2.7. Live/dead staining
The viability of H22 cells was further evaluated using the
Calcein-AM/PI Double Stain Kit according to the manufacturer's
protocol. Cells cultured in blank culture medium, ZCJ NPs, HPS, and
HPS@ZCJ hydrogel were resuspended and plated into 96-well plates at a
density of 3 × 10^3 cells per well. Following a 1, 2, 3-day culture
period, the cells were stained with calcein-AM (acetoxymethyl,
indicating live cells) and propidium iodide (PI, indicating dead cells)
for 30 min, and subsequently imaged using confocal laser scanning
microscopy (CLSM).
2.8. Cell apoptosis
H22 cells (1 × 10^5 cells/well) were plated into a 6-well plate and
exposed to 350 μg/mL of ZCJ, HPS hydrogel, and HPS@ZCJ hydrogel. For
controls, cells were cultured in RPMI-1640 with 10 % FBS. Apoptosis in
H22 cells was evaluated using the Annexin-V-FITC Apoptosis Detection
Kit (Beyotime) according to the manufacturer's protocol. Cells were
collected, washed twice with cold PBS (pH = 7.4), centrifuged, and then
incubated with annexin-V-FITC/PI at 37 °C for 15 min as per the kit
instructions. After staining, the cells were analyzed using a
FACS-Calibur flow cytometer and CellQuest software (Becton Dickinson,
San Jose, CA).
2.9. Cell cycle assay
H22 cells (1 × 10^5 cells/well) were plated into 6-well plates and
exposed to ZCJ, HPS hydrogel, and HPS@ZCJ hydrogel (350 μg/mL) at 37 °C
for 48 h. Cells maintained in RPMI-1640 with 10 % FBS were used as
controls. Post-incubation, the cells were dissociated with trypsin,
carefully harvested, and centrifuged at 1000g. The supernatant was
carefully aspirated, and the cells were resuspended in pre-cooled PBS
(pH = 7.4) before being transferred to a 1.5 mL centrifuge tube. The
cells were then fixed in 70 % cold ethanol and stored at 4 °C for 24 h.
The cells were subsequently centrifuged, washed twice with cold PBS
(pH = 7.4), treated with RNase A (0.1 mg/mL) for 1 h at 37 °C, and
stained with PI (0.1 mg/mL) for 30 min in the dark. DNA content was
analyzed by flow cytometry (FACSCalibur, BD, USA), and the distribution
of cells across different cell cycle phases was evaluated using ModFit
software.
2.10. Transwell assay
The migration assay was carried out using transwells (8 μm pore size,
24-well plate, BD Biosciences, USA). H22 cells and HUVECs were deprived
of serum by incubation in serum-free medium for 24 h. Subsequently,
H22 cells and HUVECs (1 × 10^6 cells each) were introduced into the
upper chamber of the transwell, with the lower chamber containing
migration-inducing medium supplemented with 10 % FBS (HPS, HPS@ZCJ or
ZCJ). The cells were allowed to migrate for 24 h. Cells that migrated
to the lower chamber were harvested, fixed with 4 % paraformaldehyde
for 30 min, and stained with 0.5 % crystal violet (Beyotime
Biotechnology, China) for 20 min. Migrated cells were imaged using an
optical microscope, and cell numbers were manually counted in each
microscopic field.
2.11. Scratch assay
HUVECs were plated in 35-mm petri dishes at a density of 1 × 10^6 cells
per dish and cultured at 37 °C for 24 h. Once the cells formed a
confluent monolayer, a uniform gap was introduced using a 200-μL
sterile pipette tip. After PBS rinsing to remove cell debris, the cells
were maintained in either plain medium, ZCJ NPs, or medium supplemented
with HPS or HPS@ZCJ. The closure of the scratch was observed with an
inverted microscope (Olympus IX-73, Japan) at 0-, 12-, and 24-h
post-scratch. The scratch area was quantified using ImageJ software
(version 1.8.0, NIH, USA), and the wound-healing rate was computed to
evaluate the migratory potential of HUVECs. The healing rate was
calculated using the formula: Wound-healing rate
(%) = A[n]/A[0] × 100 %, where A[0] and A[n] denote the initial wound
area and the remaining unhealed area, respectively.
2.12. Cell immunofluorescence staining experiment
HUVECs were plated in confocal dishes (JingAn Biological, China) at a
density of 1 × 10^3 cells/mL and incubated with 350 μg/mL of ZCJ, HPS,
or HPS@ZCJ solutions. After 3 days of incubation, the cells were fixed
using 4 % paraformaldehyde for 15 min. They were then permeabilized
with 0.1 % Triton X-100 (Abcam, USA) and blocked with 3 % BSA/PBS
(Aladdin, China) for 20 and 30 min, respectively, at room temperature.
Following PBS (pH = 7.4) rinsing, the cells were stained with a rabbit
anti-mouse primary antibody specific for CD31 (1:200 dilution, Abcam,
USA) and incubated overnight at 4 °C. The cells were gently washed with
PBS (pH = 7.4) and reacted with Cy3-conjugated anti-rabbit IgG
secondary antibodies (1:200 dilution, Abcam, USA) for 2 h in the dark.
In parallel, cell nuclei were stained with 4,6-diamidino-2-phenylindole
dilactate (DAPI, Abcam, USA) for 15 min in the dark. The stained cells
were imaged using a laser scanning confocal microscope (Nikon, Japan),
and the relative expression levels of VEGF were quantified using ImageJ
software.
2.13. Hemolysis test
Red blood cells were extracted from BALB/c mice, washed with saline,
and diluted. Next, 100 μL of a 4 % (v/v) red blood cell solution was
combined with 0.9 mL of saline containing different concentrations of
HPS@ZCJ hydrogel. In the negative control, 100 μL of 4 % (v/v) red
blood cells was mixed with 0.9 mL of saline, whereas in the positive
control, 100 μL of 4 % (v/v) red blood cells was mixed with 0.9 mL of
deionized water. Following incubation at 37 °C for 2 h, the absorbance
of the supernatant at 540 nm was determined using a microplate reader
(iMark, BIO-RAD, USA). The hemolysis rate was calculated using the
formula:
[MATH: Hemolysisrate(%)=(OD0−OD1)/(OD2−OD1)×100% :MATH]
where OD0 is the optical density (OD) of red blood cells in HPS@ZCJ
hydrogel at various concentrations, OD1 is the OD of red blood cells in
normal saline, and OD2 is the OD of red blood cells in deionized water.
2.14. In vitro and in vivo hemostasis properties
Anticoagulated mouse blood (2000 μL) was combined with PBS, fibrinogen,
or HPS@ZCJ in a test tube, followed by the addition of thrombin to
induce clotting. After a 5-min incubation, the tubes were tilted to
assess clot formation.
To further assess the in vitro hemostatic effect of HPS@ZCJ hydrogel,
an in vitro clotting test was performed with four groups: i) control;
ii) HPS@ZCJ; iii) fibrin glue. The control group was treated with
100 μL of PBS, the HPS@ZCJ group with 100 μL of HPS@ZCJ hydrogel and
the fibrin glue group with 100 μL of fibrin glue. Next, 900 μL of mouse
blood was added to each group. After 1 min, all liquids were gently
removed, and the samples were rinsed three times with PBS. Repeat the
above steps, wait 2–9 min, absorb the liquid, and wash. The first time
a blood clot appeared in each group was photographed and recorded.
The blood clotting index (BCI) was then determined. For the BCI assay,
fibrin hydrogel or HPS@ZCJ was prewarmed at 37 °C for 10 min. Then,
9 mL of anticoagulated blood and 1 mL of 0.1 M CaCl2 were applied to
the hydrogel. Post a 5-min incubation, unclotted blood was dissolved in
5 mL of deionized water, and the optical density (OD) was measured at
540 nm. As a control, 50 μL of anticoagulated blood was mixed with
deionized water. The BCI was calculated using the formula: BCI (%) =
(OD of materials/OD of reference) × 100 %.
Red blood cell (RBC) adhesion was evaluated by preparing 100 μL of
hydrogel with HPS@ZCJ or fibrin in a 96-well plate, to which 50 μL of
anticoagulated whole blood was added. The mixture was agitated at 37 °C
for 10 min, followed by washing with PBS to remove non-adherent RBCs.
The hydrogels were then incubated with deionized water at 37 °C for
30 min to lyse adherent RBCs, after which the OD was measured at
540 nm. RBC attachment was calculated as: RBC adhesion (%) = (OD of
sample/OD of reference) × 100 %.
All animal procedures were sanctioned by the Institutional Animal Care
and Use Committee at Southern University of Science and Technology
(SUSTech-JY202411104). In the standard round liver defect model, a 6 mm
diameter biopsy needle was utilized to induce a circular wound in the
rat liver, reaching a depth of 3 mm. Subsequently, 100 μl of each test
sample was administered directly into the defect. The extent and
duration of blood loss were then systematically evaluated.
2.15. Animals and ethics statement
Male BALB/c mice, 8 weeks old and weighing 16–18g, were sourced from
the Southern University of Science and Technology and maintained under
Specific Pathogen Free (SPF) conditions. These mice were kept in a
specialized facility with controlled temperature and humidity, exposed
to a 12-h light/12-h dark cycle, and provided with unrestricted access
to water and standard rodent diet. The study's protocols were
sanctioned by the Institutional Animal Care and Animal Ethics Committee
of the Southern University of Science and Technology (approval no.
SUSTech-JY202411104). All animal experiments adhered to the local
animal welfare regulations and guidelines set by the Southern
University of Science and Technology.
2.16. In vivo antitumor effects of HPS@ZCJ hydrogel
To assess the in situ antitumor activity of HPS@ZCJ hydrogel, Balb/c
mice were randomly divided into six groups (n = 9): sham operation,
control, ZCJ, HPS, HPS@ZCJ, and doxorubicin (DOX). H22 cells
(1 × 10^6 cells/mL) were suspended in PBS (pH = 7.4), and 100 μL was
injected into the livers of the mice (H22 cells were not injected in
the sham operation group). The mice in each group were then treated as
follows: i) and ii) no treatment; iii) application of 100 μL ZCJ NPs at
the inoculation site; iv) application of 100 μL HPS hydrogel at the
inoculation site; v) application of 100 μL HPS@ZCJ hydrogel at the
inoculation site; vi) application of 100 μL DOX (2 mg/kg) at the
inoculation site. The body weight of the animals was also recorded
every 2 days. On day 7 after treatment, the mice (n = 3 in each group)
were euthanized, and the tumor-bearing livers were photographed and
weighed. After treatment, residual HPS@ZCJ hydrogel remained adhered to
the liver surface. Moreover, the survival rates of mice in each group
(with six mice per group) were tracked for 30 days, and Kaplan-Meier
survival curves were generated.
2.17. In vivo recurrence prevention effect of HPS@ZCJ hydrogel after tumor
resection
To assess the antirecurrence potential of HPS@ZCJ hydrogel in vivo, a
subcutaneous HCC recurrence model was developed using BALB/c mice. H22
cells (1 × 10^6 cells in 0.1 mL PBS) were injected subcutaneously into
the lateral right lower abdominal wall of the mice. The body weight of
the animals was recorded on days 0, 3, and 7. One week after tumor cell
implantation, when the tumor volume reached approximately 150 mm^3,
mice were anesthetized using isoflurane (up to 5 % for induction and
1–3 % for maintenance) in an induction chamber, with anesthesia
maintained via a nose cone. Subsequently, all visible tumors were
surgically excised using sterile instruments, and subsequent treatments
were administered. Mice were randomly allocated into five groups
(n = 12): i) resection only; ii) resection and HPS hydrogel treatment;
iii) resection and ZCJ NPs treatment; iv) resection and DOX treatment;
v) resection and HPS@ZCJ hydrogel treatment. Additionally, the amount
of blood loss during the surgical removal of the tumor was recorded.
Post-surgery, the body weight of the mice was tracked. On day 14
post-surgery, the heart, liver, spleen, lungs, kidneys and recurrent
tumors from all mice (n = 6 in each group) were harvested for follow-up
experiments. The tumors were photographed and weighed. Tumor size was
calculated using the formula: width [[80]2] × length × 0.5. After
treatment, a portion of the HPS@ZCJ hydrogel persisted at the
tumor-resection site; because the hydrogel is fully biodegradable, it
can be left in place, eliminating the need for secondary surgery to
remove residual material and markedly reducing the associated operative
risk. Additionally, the survival of mice in each group (n = 6 in each
group) was monitored over 31 days, and Kaplan-Meier survival curves
were constructed.
2.18. Hematoxylin and eosin (H&E) staining
To determine the potential toxicity of HPS@ZCJ hydrogel on major organs
and the pathological conditions of tumors in mice, histopathological
evaluations were carried out on the heart, liver, spleen, lungs,
kidneys, and tumors. Under a protocol approved by our Institutional
Animal Care and Use Committee, mice from each group were humanely
euthanized at the end of the experiment via cervical dislocation. The
heart, liver, spleen, lungs, kidneys, and tumors were promptly excised
and fixed in 4 % paraformaldehyde for 48 h. The fixed tissues were then
dehydrated with 30 % sucrose solution and embedded in paraffin using
standard protocols. Tissue sections (5 μm thick) were stained with
hematoxylin and eosin (H&E; C0105M, Beyotime). The stained sections
were examined under a light microscope (Leica Microsystems, Germany) to
evaluate tissue morphology changes.
2.19. Immunohistochemical staining
Tumor tissue samples were fixed with 10 % neutral buffered formalin
(Sigma-Aldrich, USA) and embedded in paraffin. Subsequently, the
samples were processed for deparaffinization and rehydration. They were
then incubated overnight at 4 °C with primary antibodies targeting
Ki-67 (1:200, Abcam, USA), TUNEL (1:200, Abcam, USA), DHCR7 (1:200,
SAB, USA), GPNMB (1:200, Proteintech, China), and CD-31 (1:200, Abcam,
USA). On the following day, the samples were treated with biotinylated
secondary antibodies for 30 min at 37 °C. Tissue sections were
visualized using the Pierce™ DAB Substrate Kit (34002, Thermo Fisher,
USA) and examined under an optical microscope.
2.20. Immunofluorescent staining
The tumor microenvironment significantly influences tumor growth.
Consequently, we examined the impact of HPS@ZCJ hydrogel on immune
cells in tumor tissues via immunofluorescence staining. Tumor tissues
were sliced, affixed to slides, and stained with primary antibodies
specific for CD206 and CD8 (1:200, Abcam, USA) for 12 h.
Fluorescence-labeled secondary goat anti-rat antibodies were
subsequently added to enhance signal detection. Nuclei were stained
with DAPI. The sections were then imaged using a Zeiss fluorescence
microscope.
2.21. Human tumor tissue samples
The protocol for obtaining human hepatocellular carcinoma tissue
samples was approved by the Ethics Review Committee of the Guangdong
Provincial People's Hospital (No. KY2024-1105-01). All samples were
collected with informed consent from patients, in accordance with the
International Ethical Guidelines for Biomedical Research Involving
Human Subjects (CIOMS).
2.22. Isolation and culture of human primary HCC cells
Fresh tumor samples were rinsed three times with DMEM to remove blood
and other impurities. The cleaned tumor samples were then minced into
fragments of approximately 1 mm^3 and digested with Hank's Balanced
Salt Solution (HBSS) (Gibco, Carlsbad, CA, USA) and 0.1 % Type IV
collagenase (Gibco, Carlsbad, CA, USA) at 37 °C for 1–2 h. The digested
samples were filtered through a 100 μm nylon filter and centrifuged at
4 °C for 3 min. The supernatant was discarded, and the cells were
washed twice with HBSS and finally resuspended in hepatocyte culture
medium. After seeding, the medium was replaced with fresh medium
containing different concentrations of peptide after 24 h to remove
dead cells and debris.
2.23. RNA sequencing analysis (RNA-seq)
Human HCC cell samples (P1) were randomly divided into two groups and
co-cultured with either blank culture medium or HPS@ZCJ hydrogel for 5
days. Total RNA was then extracted from the samples and its integrity
was assessed using the RNA Nano 6000 Assay Kit (Agilent Technologies,
CA, USA, 5067-1511). mRNA libraries were constructed following the
standard operating procedure with Novogene software. Indexed samples
were clustered on the cBot Cluster Generation System using the TruSeq
PE Cluster Kit v3-cBot-HS (Illumina). After clustering, library
preparation was sequenced on the Illumina Novaseq platform, generating
150 bp paired-end reads. To ensure the reliability and reproducibility
of the results, three independent experiments were performed (n = 3).
Differentially expressed genes (DEGs) were detected from RNA-seq data
using the R package limma (V.3.56.2). DEGs were identified based on a
log2 fold change (log2FC) exceeding 1 or below −1, and an adjusted
p-value <0.05 after Bonferroni correction. Subsequently, these DEGs
were analyzed for enrichment in Gene Ontology (GO), Kyoto Encyclopedia
of Genes and Genomes (KEGG), and Gene Set Enrichment Analysis (GSEA)
using the R package clusterProfiler (V.4.8.3).
2.24. Statistical analysis
Data are presented as individual values with the mean ± standard error
of the mean. Comparisons between multiple groups were made using
one-way ANOVA, while Student's t-test was used for comparisons between
two groups. Survival analysis was assessed by the log-rank test. All
statistical analyses were performed using GraphPad Software, with
P < 0.05 indicating statistical significance.
3. Results and discussion
3.1. Preparation and characterization of ZCJ NPs loaded with JEO
The chemical constituents of JEO were analyzed using HRLC-MS and GC-MS
techniques, with the results presented in [81]Fig. 2A–C. Based on
existing research, Dehydrocostus lactone, Curdione, Germacrone, and
Neocurdione—all identified within JEO—possess significant anticancer
properties [[82][32], [83][33], [84][34]]. Thus, it is indicated that
these compounds are the primary antitumor active components of JEO.
Fig. 2.
[85]Fig. 2
[86]Open in a new tab
Characterization of ZCJ NPs. (A–C) Schematic illustration of JEO's
chemical constituents analyzed via HRLC-MS and GC-MS. (D–F)
Physicochemical properties of ZCJ NPs in different proportions,
including particle size, PDI, and zeta potential. (G) Morphological
examination of ZCJ NPs via SEM and TEM, demonstrating spherical shape
and layered structure (Sscale bar: 120 nm). (H) Physicochemical
properties of zein@JEO, zein-CS, and ZCJ NPs. (I) Particle size
distribution of ZCJ NPs. (J) FTIR spectroscopy confirmed the successful
combination of zein and CS in ZCJ NPs and the encapsulation of JEO.
Zein-based NP delivery systems are widely utilized in tumor therapy due
to their ability to encapsulate hydrophobic compounds [[87]35]. Zein, a
natural protein carrier rich in hydrophobic amino acids, effectively
encapsulates hydrophobic drugs [[88]36]. However, the stability of
these NPs often requires enhancement. To address this, CS, a natural
anionic polysaccharide, is employed as a stabilizer due to its negative
charge, which helps protect the NPs. In the study, we developed a
stable NP system by loading the hydrophobic compound JEO into zein and
coating it with chondroitin sulfate through electrostatic interactions
and hydrogen bonding, forming ZCJ NPs. The physicochemical properties
of JEO-loaded NPs are summarized in [89]Fig. 2D–J. Our preliminary
experiments investigated the effect of varying the zein/CS ratio on NP
size. As shown in [90]Fig. 2D–F and [91]Table S1, the ZCJ-1 formulation
(zein to CS ratio of 5:1) exhibited the smallest particle size
(141.8 ± 1.1 nm) and polydispersity index (PDI, 0.233 ± 0.127). The
negative charge of CS resulted in a negative zeta potential for ZCJ NPs
(−30.11 ± 0.66 mV). SEM and TEM images revealed that the NPs were
spherical, with distinct inner and outer layers ([92]Fig. 2G–I and
[93]S1A). These ZCJ-1 NPs achieved high drug loading efficiency, with
an EE (%) of 94.7 ± 2.8 % ([94]Fig. 2H). On the other, zein and CS had
their characteristic peaks. Zein showed N-H bending vibration and C-N
stretching vibration of the secondary amide at 1529 cm^−1 28, while CS
showed C single bond O single bond C stretching vibration at 1031 cm^−1
37. Notably, the 1031 and 1529 cm^−1 absorption peaks could be detected
in ZCJ and Zein@CS NPs ([95]Fig. 2J). In addition, CS exhibited a peak
at 3244 cm^−1 due to overlapping vibrations of –NH and –OH groups,
while zein showed a narrower peak at 3301 cm^−1 attributed to –OH
vibrations [[96]28,[97]37]. In the spectrum of ZCJ NPs, the –OH groups
exhibited a slight shift to 3282 cm^−1, likely due to the formation of
hydrogen bonds between zein and CS [[98]38]. The peak shape of ZCJ was
similar to that of Zein@CS, indicating that JEO was encased in the
interior.
3.2. Preparation and characterization of HPS@ZCJ hydrogel
In this study, we developed the HPS@ZCJ hydrogel based on PF127, HMC,
and SA for hemostasis of liver hemorrhage, and integrated JEO-loaded
CS-modified zein NPs (ZCJ) into the hydrogel system as a functional
nanoreinforcing filler to prevent postoperative tumor recurrence. To
fulfill the requirements of facile preparation, biocompatibility,
robust drug-loading capacity, and sustained drug release, we selected
PF127 hydrogel as the carrier material. PF127, a polymeric nonionic
surfactant comprising 70 % polyethylene oxide and 30 % polypropylene
oxide, exhibits thermosensitive behavior, transitioning from a sol to a
gel state at its lower critical solution temperature due to
hydrophobic-hydrophilic interactions between its components [[99]27].
To enhance the duration and effectiveness of hemostasis while improving
sustained drug release and stability, we incorporated HMC into the
hydrogel and introduced SA as a double-crosslinked network carrier with
PF127, with HMC also functioning as a thickener.
The HPS@ZCJ hydrogel was specifically designed for injection to ensure
precise delivery to the surgical site. This method allows for
controlled application and ensures that the hydrogel adheres to the
tissue, providing sustained release of the therapeutic agents. The
thermosensitive nature of the hydrogel ensures that it transitions from
a liquid to a solid state at body temperature [[100]39], further
enhancing its ability to remain in place and deliver the JEO
effectively. The as-prepared HPS@ZCJ hydrogel was able to be extruded
from a syringe through a 21G needle ([101]Fig. S1B). Characterization
studies revealed that the HPS@ZCJ hydrogel transitions from a flowing
sol state at 4 °C to a solid gel state at 37 °C. This thermoresponsive
behavior was confirmed in vitro by injecting the HPS@ZCJ solution into
a 37 °C water bath, where it rapidly formed a viscoelastic gel capable
of adhering to surfaces and tissues without flowing ([102]Fig. 3A). To
ensure a stable controlled-release microenvironment, hydrogels must
possess adequate mechanical and adhesive strength. As shown in
[103]Fig. 3B, the HPS@ZCJ hydrogel can adhere the liver tissue of a rat
to the tip of a finger. We further evaluated these properties using a
lap shear test. As shown in [104]Fig. 3C, the adhesive strength of
hydrogels composed solely of PF127 was minimal. In contrast, the
HPS@ZCJ hydrogel demonstrated remarkable adhesive strength, reaching up
to 28.89 kPa on iron blocks (hydrophobic surfaces) and up to 37.17 kPa
on wooden sticks (hydrophilic surfaces). These unique thermal and
adhesive characteristics offer practical advantages. The hydrogel can
be injected at low temperatures, uniformly covering the liver surface.
Upon injection, it solidifies at body temperature, preventing
displacement by wound exudate. This ensures secure mechanical adhesion
and sustained drug release at the wound site, facilitating hemostasis
and anti-tumor effects. [105]Fig. S1C illustrates a real application
scenario. The image shows an isolated rat's liver, with the left side
untreated and the right side uniformly coated with HPS@ZCJ. The bottom
image depicts the in vivo application, with HPS@ZCJ stained red for
visualization. Besides, SEM analysis revealed the microstructural
differences between HPS@ZCJ and HPS. As shown in [106]Fig. 3D, HPS@ZCJ
exhibited a porosity of 43.6 %, compared to 52.1 % for HPS, indicating
smaller and more uniformly distributed pores in HPS@ZCJ, whereas HPS
had larger and more numerous pores. This pore distribution pattern
likely results from the incorporation of ZCJ NPS in HPS@ZCJ, which
enhances intermolecular interactions and increases cross-linking
density, thereby impeding water evaporation during freeze-drying. We
next probed the synergistic interactions among PF127, HMC, and SA
([107]Fig. S2A). In the FTIR analysis of SA, the characteristic band at
1606 cm^−1 is indicative of the asymmetric stretching vibrations of
carboxylate (–COO^-) groups [[108]40]. In the HPS hydrogel, the
hydroxyl group peak of HMC, originally at 3460 cm^−1, broadens and
shifts to 3458 cm^−1, suggesting that these hydroxyl groups are engaged
in hydrogen bonding. Concurrently, the carboxylate ion peak at
1610 cm^−1 in SA is markedly attenuated in the HPS hydrogel spectrum,
which implies the formation of hydrogen bonds between the hydroxyl
groups of HMC and the carboxylate ions of SA. Moreover, PF127 exhibits
a peak at 1103 cm^−1, corresponding to the C-O-C stretching, while HMC
displays this peak at 1056 cm^−1 [[109]41]. Within the HPS hydrogel,
the C-O-C peak shifts to 1095 cm^−1, indicating hydrogen bond formation
between HPMC and PF127, which results in the observed peak shift. The
presence of characteristic peaks for PF127 (1095 cm^−1) and SA
(1610 cm^−1) in the hydrogel confirms the establishment of an
interpenetrating double cross-linked hydrogel network. In addition,
from the spectra of [110]Fig. 3E, it is important to highlight the
absence of a band between 1505 and 1545 cm^−1 characteristic of amide
II in the HPS group [[111]28]. This observation is attributable to the
proteinaceous nature of zein: the sharp, intense band at 1523 cm^−1
present in zein ([112]Fig. 2J), ZCJ and HPS@ZCJ (1505 cm^−1) confirms
successful encapsulation of ZCJ NPs within the hydrogel.
Fig. 3.
[113]Fig. 3
[114]Open in a new tab
Characterization of the HPS@ZCJ hydrogel. (A) Assessed fluidity at 4 °C
and 37 °C to demonstrate temperature-responsive behavior. (B)
Photographs depicting HPS@ZCJ hydrogel adhesion to rat liver. (C)
Schematic and quantitative results of hydrogel adhesion tests. (D) SEM
images detailing the structural and morphological features of HPS@ZCJ
hydrogel. (E) FTIR spectra comparing HPS@ZCJ, ZCJ, and HPS. (F)
Rheological analysis highlighting the viscoelastic properties
influenced by temperature. (G) Frequency scanning outcomes for HPS and
HPS@ZCJ composites. (H) Cumulative in vitro release profile of JEO from
HPS@ZCJ. (n = 3).
Rheological analysis revealed that the critical phase transition
temperature of the HPS@ZCJ hydrogel is approximately 25.6 °C ([115]Fig.
3F). The sol-to-gel transition is rapid and smooth, occurring swiftly
once the gelation temperature is exceeded. Below this temperature, the
hydrogel exhibits liquid-like behavior, characterized by a loss modulus
(G″) greater than the storage modulus (G′). Above 25.6 °C, G′ exceeds
G″, indicating solid-like behavior. Given that operating room
temperatures are typically around 26 °C, the HPS@ZCJ hydrogel, stored
at 4 °C, requires a brief period to reach its critical transition
temperature. This delay prevents premature gelation, facilitating
precise injection during clinical procedures. Frequency scanning
results ([116]Fig. 3G) revealed that both HPS@ZCJ and HPS hydrogels
exhibited G′ higher than their G″, with minimal changes observed with
increasing frequency. This indicates a stable crosslinking network at
room temperature (25 °C). Specifically, HPS@ZCJ had a G′ of
approximately 6.3 kPa and a G″ of 1.3 kPa, whereas HPS, lacking ZCJ
NPs, had lower values of 4.1 kPa and 0.8 kPa, respectively. These
findings suggest that the incorporation of ZCJ NPs significantly alters
the hydrogel structure, enhancing its mechanical properties.
Furthermore, the HPS@ZCJ hydrogel exhibited shear-thinning behavior,
indicating that its viscosity decreased under applied shear stress.
This property facilitates the smooth extrusion of the HPS@ZCJ hydrogel
through medical needles, ensuring ease of injection during clinical
procedures ([117]Fig. S3A). This property complements the hydrogel's
thermosensitive transition to a solid-like state at body temperature,
which ensures stable application and sustained release of therapeutic
agents.
3.3. JEO sustained release, swelling behavior, and in vitro degradation of
HPS@ZCJ hydrogel
To accurately quantify the sustained release capacity of the HPS@ZCJ
hydrogel, standard curves of JEO were established ([118]Fig. S2B).
These curves facilitated the determination that the cumulative release
of JEO from HPS@ZCJ in a pH 7.4 SBF environment reached approximately
89.7 % within 6 days ([119]Fig. 3H). This substantial release profile
confirms the hydrogel's efficacy in achieving sustained drug delivery,
which is critical for maintaining therapeutic concentrations over
extended periods. These results highlight the rational design of
HPS@ZCJ hydrogel with its ideal release kinetics and sustain an
enduring anti-HCC effect. Excessive swelling of hydrogels can compress
surrounding tissues, blood vessels, and nerves, potentially causing
discomfort and severe side effects. Moreover, such swelling can
diminish the hydrogel's cohesion, resulting in inadequate mechanical
and adhesive strength [[120]42]. Therefore, we evaluated the swelling
behavior of the HPS@ZCJ hydrogel. As shown in [121]Fig. S3B, the
swelling ratio of HPS@ZCJ gradually increased, reaching equilibrium at
approximately 131 % after 16 h. This indicated that the HPS@ZCJ
hydrogel has good anti-swelling properties and does not exert pressure
on surrounding tissues post-injection. Furthermore, for optimal in vivo
performance, the hydrogel should exhibit gradual biodegradability to
sustain drug release effectively and be fully absorbed by the body.
[122]Fig. S3C illustrates that the HPS@ZCJ hydrogel underwent gradual
degradation in SBF, with 74.9 % remaining after 7 days and essentially
degrading by day 21. This profile parallels the inflammatory and
proliferative phases of hepatic wound healing, which typically span
7–21 days after partial hepatectomy [[123]43]. Additionally,
degradation releases only naturally occurring metabolites-salts of
alginate, hydroxymethyl cellulose oligosaccharides, and plant-derived
essential-oil constituents-whose benign profiles are well documented.
Consequently, the hydrogel is expected to clear from the surgical site
before the onset of the remodeling phase, ensuring both safety and
compatibility with the normal healing trajectory.
3.4. In vitro antitumor effects of HPS@ZCJ hydrogel
We first studied the antitumor effects of ZCJ NPs on H22 cells by using
CCK-8. ZCJ was at different concentrations (0–4000 μg/mL) and
cocultured with H22 cells for 1–3 days. As shown in [124]Fig. 4A, ZCJ
remarkably inhibited H22 cells in a dose-dependent manner. Moreover,
the cytotoxicity of HPS@ZCJ hydrogel was assessed against H22 cells and
HUVECs. The IC50 values (the dose required to inhibit 50 % cellular
growth within 24 h) were found to be 1848.1 μg/mL for HUVECs and
361.6 μg/mL for H22 cells ([125]Fig. 4B and C). This indicated that
HPS@ZCJ hydrogel is less cytotoxic to normal cells than to the tested
cancerous cells. Consequently, guided by the IC50 results, a
concentration of 350 μg/mL was selected for subsequent cellular
experiments. The live/dead staining outcomes corroborated the
aforementioned findings, indicating that the HPS@ZCJ hydrogel
significantly impedes the proliferation of H22 cells ([126]Fig. 4D and
E). Moreover, for biological materials applied directly to the human
body, biocompatibility is a fundamental requirement. For hemostatic
materials, hemocompatibility testing is crucial to ensure no hemolysis
occurs during hemostasis. As shown in [127]Fig. 4F, the hemolysis ratio
of HPS@ZCJ hydrogel was evaluated over a concentration range of
0–2000 μg/mL. The embedded image reveals that up to 1000 μg/mL, the
solution color remained largely unchanged, indicating minimal hemolysis
and aligning with the negative control. This concentration range is
commonly accepted as indicative of good hemocompatibility. However, at
2000 μg/mL, the hemolysis ratio exceeded 10 %, suggesting a potential
risk at higher concentrations.
Fig. 4.
[128]Fig. 4
[129]Open in a new tab
(A) Cell cytotoxicity of H22 cells treated with ZCJ NPs in different
concentrations by CCK-8 assay. (B, C) The IC50 values for HUVECs and
H22 cells were calculated from nonlinear regression analyses plotting
the percentage of specific cytotoxicity against the Log10 concentration
of the HPS@ZCJ hydrogel. (D) The Live/Dead cell staining result of
H22 cells treated with ZCJ NPs, HPS and HPS@ZCJ hydrogels after 3 days
of culture. (E) Quantitative analysis of the Live/Dead cell staining.
(F) Hemocompatibility evaluations of HPS@ZCJ hydrogel in different
concentrations. (n = 3, ∗ and # represent P < 0.05 by comparing with
the control and HPS groups, respectively).
To further assess the antitumor efficacy of HPS@ZCJ hydrogel, three
groups were established based on their distinct compositions: ZCJ NPs,
HPS, and HPS@ZCJ hydrogel. Flow cytometry was initially employed to
measure apoptosis rates in H22 cells across these groups. The HPS@ZCJ
group exhibited an apoptosis rate of 37.55 ± 3.8 %, whereas ZCJ NPs
achieved a rate of 42.3 ± 4.8 % ([130]Fig. 5A). This difference is
likely due to the sustained-release profile of HPS@ZCJ hydrogel, which
prevents the rapid release of the maximum dose within a short period.
Given that HPS hydrogel alone does not induce apoptosis ([131]Fig. 5B),
the cytotoxicity observed in [132]Fig. 4D and E indicates that HPS
hydrogel may promote tumour-cell death via non-apoptotic pathways. We
further explored the impact of HPS@ZCJ hydrogel on cell cycle
progression in H22 cells ([133]Fig. 5C and D). Following treatment with
HPS@ZCJ hydrogel, the proportion of H22 cells in the G0/G1 phase
significantly increased from 41.01 % to 52.73 % (P < 0.05).
Concurrently, the percentage of cells in the S phase decreased from
34.82 % to 29.18 % (P < 0.05), and that in the G2/M phase decreased
from 24.04 % to 18.09 % (P < 0.05). These findings demonstrate that
HPS@ZCJ hydrogel induces G0/G1 phase arrest in H22 cells. Transwell
chamber assays were employed to evaluate and quantify the migratory
behavior of H22 cells on hydrogels. The results, as shown in [134]Fig.
5, indicated that after 48 h, both the ZCJ and HPS@ZCJ groups
significantly impeded H22 cell migration compared to the control and
HPS groups (P < 0.05, [135]Fig. 5E–G). Collectively, these findings
highlight the significant antitumor potential of HPS@ZCJ hydrogel,
which can induce apoptosis, enhance cell cycle arrest, inhibit cell
proliferation, and impede cell migration.
Fig. 5.
[136]Fig. 5
[137]Open in a new tab
In vitro anti-tumor ability of HPS@ZCJ hydrogel. (A) Flow plot showing
the cell apoptosis rate in different groups with an Annexin V/PI
apoptosis detection kit. (B) Flow cytometry quantification of the
proportion of cell apoptosis rate in different groups. (C) Flow
cytometry analysis of the H22 cells cycle after 2 days of treatment
with HPS@ZCJ hydrogel. (D) The quantification of the cell cycle test.
(E, F) Transwell was used to detect the effect of HPS@ZCJ hydrogel on
the migration of H22 cells and HUVECs, respectively. (G) Quantitative
analysis of the transwell experiment. (H) Representative fluorescence
images of differentially treated HUVECs after CD31 staining. (n = 3,
Data are mean ± SD; ∗ and # represent P < 0.05 by comparing with the
control and HPS groups, respectively).
3.5. HPS@ZCJ hydrogel inhibits endothelial cell functions
Throughout the angiogenesis of tumor progression, the migration and
invasion of endothelial cells are crucial processes [[138]44]. However,
when stimulated by HPS@ZCJ, HUVECs exhibited minimal migration to the
lower compartment of the filter ([139]Fig. 5F and G). In contrast, HPS
hydrogel alone had negligible effects on cell migration. A wound
healing assay also demonstrated that HPS@ZCJ hydrogel inhibited the
migration of HUVECs, as indicated by shorter migration distances
([140]Fig. S4). We further investigated the expression of CD31 in
HUVECs cultured with various samples. CD31, a key marker of
angiogenesis, is implicated in the proliferation, migration, and
vasculogenesis of HUVECs [[141]44]. Our results indicated that HUVECs
cultured with HPS@ZCJ displayed weaker green fluorescence intensity,
signifying a substantial reduction in CD31 protein expression compared
to other groups ([142]Fig. 5H). Semi-quantitative analysis of CD31
expression further corroborated that the CD31 level in the HPS@ZCJ
group was significantly lower than in other groups ([143]Fig. S5).
Collectively, these findings demonstrate that HPS@ZCJ exerts
anti-angiogenic effects by inhibiting CD31-mediated pathways.
3.6. Hemostatic effect of the HPS@ZCJ hydrogel
The hemostatic properties of HPS@ZCJ were evaluated using a tube
tilting experiment ([144]Fig. 6A). Both the fibrin hydrogel and HPS@ZCJ
were able to coagulate blood and maintain their shapes in a solid
state. This hemostatic ability was further confirmed by a clotting
experiment in a 24-well plate ([145]Fig. 6B and C), which demonstrated
significantly shorter clotting times for the fibrin and HPS@ZCJ
hydrogels compared to the PBS group. As expected, the 5-min BCIs of the
fibrin and HPS@ZCJ hydrogels were 26.71 ± 11.39 % and 28.13 ± 10.62 %,
respectively ([146]Fig. 6D; P > 0.05). The red blood cell adhesion
rates were 49.26 ± 2.04 % and 48.64 ± 2.29 %, respectively, further
indicating comparable hemostatic abilities between the fibrin hydrogel
and HPS@ZCJ ([147]Fig. 6D).
Fig. 6.
[148]Fig. 6
[149]Open in a new tab
Hemostatic Evaluation of HPS@ZCJ Hydrogel. (A) Inversion test comparing
PBS, fibrin gel, and HPS@ZCJ hydrogel. (B, C) Time-dependent clot
formation for PBS, fibrin gel, and HPS@ZCJ hydrogel. (D) Blood-clotting
index and red blood cell attachment for PBS, fibrin gel, and HPS@ZCJ
hydrogel. (E, F) Schematic and photographic representation of HPS@ZCJ
hydrogel application in a rat liver resection model. (G) Quantification
of blood loss and hemostasis time across various treatment groups. Data
are mean ± SD; ∗ <0.05. (For interpretation of the references to color