Abstract
MXenzymes, a promising class of catalytic therapeutic material, offer
great potential for tumor treatment, but they encounter significant
obstacles due to suboptimal catalytic efficiency and kinetics in the
tumor microenvironment (TME). Herein, this study draws inspiration from
the electronic structure of transition metal vanadium, proposing the
leverage of TME specific‐features to induce structural transformations
in sheet‐like vanadium carbide MXenzymes (TVMz). These transformations
trigger cascading catalytic reactions that amplify oxidative stress,
thereby significantly enhancing multimodal tumor therapy. Specifically,
the engineered HTVMz, coated with hyaluronic acid, exhibits good
stability and generates a thermal effect under NIR‐II laser
irradiation. The thermal effect, combined with TME characteristics,
facilities a structural transformation into ultra‐small vanadium oxide
nanozymes (VO[x]). The enlarged surface area of VO[x] substantially
enhances ROS regeneration and amplifies oxidative stress, which
promotes lysosomal permeability and induces endoplasmic reticulum
stress. The high‐valent vanadium in VO[x] interacts with intracellular
glutathione, disrupting redox homeostasis and intensifying oxidative
stress further. These amplifications accelerate tumor apoptosis, induce
ferroptosis, and suppress HSP90 expression. Consequently, the
heightened thermal sensitivity of HTVMz synergistically promotes tumor
cell death via multimodal therapeutic pathways. This study presents an
innovative strategy for tumor catalytic therapy by manipulating
MXenzymes structures, advancing the field of catalytic therapy.
Keywords: catalytic therapy, ferroptosis, MXenzymes, NIR‐II PTT,
oxidative stress
__________________________________________________________________
This study introduces a novel strategy that uses the TME to trigger
structural changes in vanadium carbide MXenzymes, converting them into
ultra‐small vanadium oxide nanozymes. These transformations enhance
oxidative stress, promoting tumor apoptosis, ferroptosis, and lysosomal
permeability. The synergy of thermal response and catalytic reactions
boosts antitumor efficacy through thermal, oxidative, and catalytic
mechanisms.
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1. Introduction
The increasing incidence of malignant tumors has underscored the
limitations of traditional therapies in terms of efficacy, spurring the
pursuit of more potent and innovative treatment strategies.^[ [42]^1 ,
[43]^2 ^] Tumors, characterized by high metabolism, rapid
proliferation, and dysregulated metabolism pathways, often exhibited an
increased intracellular redox imbalance, which facilitated the
overproduction of reactive oxygen species (ROS) and hydrogen peroxide
(H[2]O[2]).^[ [44]^3 ^] This accumulation of ROS, combined with the
altered metabolic pathways, created an acidic and oxidative
microenvironment that was distinct from that of normal tissues. This
distinctive tumor microenvironment (TME) not only served as a hallmark
of malignancy but also provided opportunities for the development of
therapeutic strategies based on oxidative stress.^[ [45]^4 , [46]^5 ^]
Nano‐catalytic therapy, particularly involving nanozymes–nanomaterials
that mimic enzymatic functions, has garnered significant attention due
to their selective reactivity toward TME‐specific molecules like
H[2]O[2]. This reactivity facilitated ROS generation and glutathione
(GSH) depletion, consequently intensifying oxidative stress within
tumors.^[ [47]^6 , [48]^7 , [49]^8 , [50]^9 , [51]^10 , [52]^11 ^]
Among these nanozymes, MXenes materials, 2D nanomaterials composed of
transition, have attracted significant interest due to their
exceptional physicochemical properties, including high surface area,
facile functionalization,^[ [53]^12 ^] and enzyme‐like ROS‐generating
activity.^[ [54]^13 ^] Notably, their remarkable photothermal
conversion capability can enhance their catalytic activity, while ROS
generated through catalysis addressed potential resistance to
photothermal therapy, thereby offering a novel strategy for cancer
catalytic treatment.^[ [55]^14 ^] Among the MXenes family,
vanadium‐based MXenes, exemplified by vanadium carbide, exhibit typical
peroxidase‐like (POD) catalytic activity, effectively catalyzing the
conversion of H[2]O[2] into ROS within the TME, thereby intensifying
intracellular oxidative stress.^[ [56]^15 , [57]^16 , [58]^17 ^]
Furthermore, the multivalent nature of vanadium enabled it to deplete
GSH through redox reactions, further amplifying the oxidative stress
and enhancing anticancer effects.^[ [59]^18 , [60]^19 , [61]^20 ^]
Nevertheless, despite its potential, MXene‐based enzymes, termed
MXenzymes, faced a pivotal challenge akin to other nano‐catalytic
nanozymes, such as inherently limited catalytic efficiency and
suboptimal kinetic properties.^[ [62]^21 , [63]^22 ^] Therefore,
improving the catalytic capability of MXenzymes to intensify oxidative
stress within tumoral environments, thereby enhancing anti‐tumor
therapeutic efficacy, emerged as a paramount imperative.
Currently, research efforts have been concentrated on various
strategies to enhance the catalytic performance of MXenzymes, including
surface modification,^[ [64]^23 ^] structural engineering,^[ [65]^24 ^]
doping methodologies,^[ [66]^25 ^] and optimized synthesis methods.^[
[67]^26 ^] While those strategies have shown promise in augmenting the
catalytic activity of MXenzymes, their inherently 2D structure (e.g.,
large size) continued to pose challenges, restricting their catalytic
efficiency. Notably, crucial factors such as surface morphology,
particle size, and shape exerted a profound influence on their
catalytic activity.^[ [68]^27 , [69]^28 ^] Given the distinctive
three‐layered atomic structure of vanadium‐based MXenes, their
exceptional reactivity was accompanied by a vulnerability to structural
transformations, particularly transitioning into vanadium oxides
(VO[x]) under oxidative conditions.^[ [70]^29 ^] This inherent property
presented a unique opportunity to tailor their properties through
environmental cues, offering a pathway to harness their potential in
advanced therapeutic applications.^[ [71]^30 ^] The TME, characterized
by high levels of H[2]O[2] and acidity, served as an ideal milieu to
trigger these structural changes, unleashing novel therapeutic avenues.
Numerous TME‐responsive 2D multifunctional nanomaterials, such as
graphene, layered double hydroxides (LDHs), and transition metal
dichalcogenides (TMDs), have been developed for tumor treatment. When
combined with techniques like photodynamic therapy,^[ [72]^31 ^]
sonodynamic therapy,^[ [73]^32 , [74]^33 ^] and photothermal therapy,^[
[75]^34 ^] those nanomaterials demonstrated significant potential for
enhancing TME‐responsive tumor therapies.^[ [76]^35 ^] Recent studies
have demonstrated that ultra‐small VO[x] nanoenzymes, with their
enlarged surface area and abundant active sites, exhibited exceptional
catalytic performance, significantly enhancing the potential of MXenes
in tumor catalytic therapy.^[ [77]^36 , [78]^37 ^] Nevertheless, these
ultra‐small nanozymes confronted challenges such as rapid in vivo
elimination and suboptimal targeting specificity, limiting their full
therapeutic potential.^[ [79]^38 ^] To address these limitations, we
proposed leveraging the abundant H[2]O[2] and acidic conditions
inherent to the TME as external stimuli to actively manipulate the
structure of vanadium‐based MXenzymes, inducing a controlled
transformation into nano‐sized VO[x] enzymes. This pioneering strategy
sought to overcome the traditional particle size restrictions of
MXenzymes, enhancing catalytic efficiency through the enlarged surface
area of the transformed nano‐sized VO[x] enzymes. Additionally, it
mitigated rapid tissue clearance of these ultra‐small nanozymes.
Expectedly, this will intensify oxidative stress within tumors, thereby
boosting antitumor efficacy. Notably, leveraging tumor‐inherent traits
to amplify MXenzymes catalytic activity was rarely reported.
Drawing inspiration from the above concept, we have employed a
combination of acid etching and ion exchange techniques to synthesize
vanadium carbide (V[2]C) MXenes, designated as TVMz. This design
capitalized on the distinctive features of the TME, particularly its
elevated H[2]O[2] levels and acidic conditions, as triggers for
inducing a structural metamorphosis in TVMz, transforming them into
ultra‐small VO[x] nanoenzymes. This transformation represented a
significant enhancement in catalytic activity, fostering oxidative
stress and potently eradicating tumor cells. Specifically, by coating
TVMz with hyaluronic acid (HA) to construct HTVMz, we have achieved
remarkable stability under physiological conditions while maintaining
the capacity to undergo structural transformation into ultra‐small
VO[x]nanoenzymes within the TME. This process not only amplified ROS
production but also prompted lysosomal membrane disruption to
facilitate the translocation of nano‐catalysts to the endoplasmic
reticulum (ER), thereby exacerbating cellular stress responses and
accelerating apoptosis. Notably, the high‐valent vanadium within VO[x]
engaged in a crucial interaction with intracellular GSH, disrupting
redox hemostasis and profoundly enhancing oxidative stress. This
interplay, coupled with the aforementioned amplification effects,
accelerated tumor apoptosis, triggered ferroptosis, and suppressed
HSP90 expression. As a result, the thermal responsiveness of HTVMz was
fortified, synergistically orchestrating a multi‐faceted attack on
tumor cells through the activation of diverse therapeutic modalities
(Figure [80] 1 ). In all, this investigation introduced a novel
paradigm for tumor catalytic therapy, revolutionizing MXenzymes
structural manipulation and advancing their filed of catalytic therapy.
Figure 1.
Figure 1
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Schematic illustration of tumor microenvironment‐driven structural
transformation of vanadium‐based MXenzymes to amplify oxidative stress
for activating multimodal tumor therapy against breast cancer. A)
Schematic diagram illustrating the transformation of HTVMz structure
simulating the lysosomal microenvironment within tumor cells. B) The
Mechanism of HTVMz in tumor microenvironment‐driven structural
transformation of vanadium‐based MXenzymes amplifying oxidative stress
for multimodal tumor therapeutics.
2. Results and Discussion
2.1. Synthesis and Characterization of Vanadium‐Based MXenzymes
Vanadium carbide MXenzymes (designated as TVMz) were synthesized using
a sophisticated etchant‐assisted ion exchange method (Figure [82] 2A).
After characterized by atomic force microscopy (AFM) and dynamic light
scattering (DLS), we confirmed that TVMz exhibited a sheet‐like
morphology, with a particle size of ≈248.50 nm and an average thickness
of ≈2.50 nm (Figure [83]2B,C). Further transmission electron microscopy
(TEM) elemental mapping scanning results revealed that TVMz was
primarily composed of carbon (C), oxygen (O) corresponding to ‐OH
groups on the TVMz surface, and vanadium (V), showing a well‐defined
structure (Figure [84]2E). Interestingly, the synthesized TVMz
exhibited absorption peaks in the NIR‐II region, which varied with
concentration, indicating their potential for photothermal therapy
(PTT) of tumors, leveraging the superior tissue penetration depth of
NIR‐II laser (Figure [85]2J). X‐ray diffraction (XRD) analysis
corroborated the successful synthesis of TVMz, consistent with the
standard JCPDS card NO. 29‐0101 for the V[2]AlC phase (Figure [86]2F),
with a characteristic peak at 7.03° corresponding to the (002) crystal
plane.
Figure 2.
Figure 2
[87]Open in a new tab
Synthesis and characterization of HTVMz. A) Schematic diagram of the
preparation process for transformable HTVMz. B) AFM image of TVMz and
the thickness of TVMz as determined by AFM analysis. Scale bar =
400 nm. C) Particle size distribution of TVMz and HTVMz. D) Zeta
potential for TVMz and HTVMz (n = 3). E) TEM images of TVMz and
elemental mapping showing the distribution of carbon, oxygen, and
vanadium. Scale bar = 100 nm and 50 nm. F) XRD patterns of V[2]AlC and
TVMz. G) FTIR spectra of HA, TVMz, and HTVMz. H) TGA analysis of HA,
TVMz, and HTVMz. I) UV–vis absorption spectra of HA, TVMz, and HTVMz.
J) UV–vis absorption spectra of HTVMz with different concentrations.
To enhance the biocompatibility and stability of TVMz and mitigate its
oxidation in air, we employed a surface modification strategy using HA,
yielding HA‐modified TVMz (HTVMz). This modification led to a notable
increase in particle size accompanied by a decrease in zeta potential
(Figure [88]2C,D). Moreover, the characteristic peaks observed in
infrared and ultraviolet‐visible spectroscopy provided evidence of the
successful integration of HA (Figure [89]2G,I). Comprehensive
thermogravimetric analysis further revealed that HA comprised ≈30% of
the nanosheet composition (Figure [90]2H). The above findings
demonstrated the effective modification of HA onto TVMz, resulting in
the formation of HTVMz.
2.2. Structure Evolution of HTVMz in Tumor Microenvironment
Inspired by the distinctive three‐layered atomic architecture of V
within HTVMz, which is inherently susceptible to oxidation, we
conducted an assessment of its stability under various environmental
conditions (Figure [91] 3A). Notably, in an acidic environment
mimicking tumor cell lysosomes (pH 5.5) with a high concentration of
H[2]O[2] (100 #x000B5;M), we observed a significant reduction in the
particle size of HTVMz over 24 h, shrinking from 262 to 34 nm
(Figure [92]3E). This remarkable transformation was further validated
by TEM images (Figure [93]3B,C), which starkly contrasted with the
minimal changes observed under neutral conditions (Figure [94]3F). The
stability of HTVMz was further evaluated. The results demonstrated that
the particle size remained relatively stable over a 7‐day period in
both ultrapure water and 1640 medium, with no significant changes
observed (Figure [95]S1, Supporting Information). Intriguingly, the
resulting small nanoparticles exhibited a new peak at ≈400 nm in UV–vis
spectroscopy, with absorption intensity positively correlated with
their concentration. This was accompanied by a noticeable decrease in
intensity within the 500–1000 nm range, indicating a distinct
structural configuration compared to the original HTVMz
(Figure [96]3G). These findings underscored the environmental
responsiveness and potential for controlled structural modulation of
HTVMz.
Figure 3.
Figure 3
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Characterization of structural transformation of HTVMz. A) the
transformation mechanism of HTVMz under simulated intracellular
lysosomal conditions. TEM images of HTVMz before (B) and after (C)
incubation at pH 5.5 with 100 #x000B5;M H[2]O[2] for 24 h (HTVMz 24 h).
Scale bar = 500 and 200 nm. D) TEM image and scanning mapping of HTVMz
24 h. E) Particle sizes of HTVMz before and after incubation at pH 5.5
with 100 #x000B5;M H[2]O[2] for 24 h. F) Particle size variation of
HTVMz over 24 h under neutral conditions. G) UV–vis absorption spectra
of HTVMz 24 h under different concentrations. H) XPS spectra of TVMz
after incubation at pH 5.5 with 100 µm H[2]O[2] for 24 h, including V2p
(I), and O1s (J) spectra. K) XPS spectra of TVMz, including V2p (L),
and O1s (M) spectra.
To further elucidate the structural characteristics of these small
nanoparticles, TEM was employed for elemental analysis, revealing that
the transformed nanoparticles were predominantly composed of V and O
(Figure [98]3D). XRD analysis confirmed strong correlations between the
diffraction peaks of these nanoparticles and the standard PDF cards of
V[2]O[5] (JCPDS card NO. 54‐0513) and VO[2] (JCPDS card NO. 31‐1438)
(Figure [99]S2, Supporting Information), suggesting that TVMz underwent
chemical reactions with H[2]O[2] under acidic conditions, resulting in
the formation of VO[x]. Moreover, X‐ray photoelectron spectroscopy
(XPS) provided detailed insight into the structural composition and
valence state distribution of VO[x] and TVMz, covering the C1s, O1s,
and V2p regions (Figure [100]3H,K). Meticulous analysis of the V2p
(Figure [101]3I,L) and O1s (Figure [102]3J,M) spectra revealed the
presence of V in various valence states, including V^5+, V^4+, V^3+,
and V^2+, as well as distinct oxygen species such as V‐O[x], V‐O, C‐O,
and V‐C‐(OH)[x]. The V2p peak of TVMz 24 h comprised V^5+ and V^4+
states, while TVMz's V2p showed V^4+, V^3+, and V^2+ states, indicating
that after structural transformation, the low‐valent V species were
oxidized to higher valent states of V. Combined with XRD results
analysis, the transformed product was identified as non‐stoichiometric
VO[x] containing V[2]O[5] and VO[2]. These findings validated the
structural transformation of TVMz under simulated lysosomal conditions,
highlighting their potential transformation into VO[x] within the TME.
2.3. Photothermal and Catalytic Performances of HTVMz
Given the absorption characteristics of HTVMz in the NIR‐II region, we
initially evaluated its photothermal performance. The results indicated
that upon NIR‐II laser irradiation for 10 min, the solution temperature
increased from 30.80 to 63.60 °C, showing a direct correlation with
concentration, irradiation power, and duration (Figure [103] 4A,B).
Importantly, after undergoing five irradiation cycles, the HTVMz
solution maintained a consistently stable peak temperature, indicating
remarkable photothermal stability (Figure [104]4C). Further detailed
analysis revealed that the extinction coefficient of HTVMz at 1064 nm
was 10.86 L g^−1 cm^−1 (Figure [105]S3, Supporting Information), with a
corresponding photothermal conversion efficiency of 49.92% at this
wavelength (Figure [106]4D). These findings not only validated the
outstanding photothermal conversion capability of HTVMz but also
highlighted its considerable potential for PTT.
Figure 4.
Figure 4
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NIR‐II photothermal and catalytic properties of HTVMz. A) Temperature
fluctuation diagrams of HTVMz upon irradiation with 1064 nm laser at
varying power densities (2.0, 1.5, and 1.0 W cm^−2). B) Photothermal
heating curves of HTVMz at elevated concentrations (50, 25, and
12.5 ppm) under 1064 nm laser irradiation at a power density of 1.0 W
cm^−2. C) Repeated heating profiles of the HTVMz (25 ppm) aqueous
solution after five cycles of 1064 nm laser irradiation on/off at 1.0 W
cm^−2. D) Calculation of photothermal conversion efficiency at 1064 nm.
E) After incubation under simulated tumor cell lysosomal conditions (pH
5.5 and H[2]O[2] concentration of 100 µm) for different times (0, 12,
and 24 h), UV–vis absorption spectra were recorded for the reactions of
HTVMz at 0, 12, and 24 h with TMB and H[2]O[2]. F) ESR spectra of
H[2]O[2], HTVMz 24 h + H[2]O[2], and HTVMz 0 h + H[2]O[2]. G) UV–vis
absorption spectra of the TMB and H[2]O[2] reaction at various pH
levels for HTVMz 24 h. Absorption spectra of HTVMz 24 h with different
concentrations of H[2]O[2] at pH 5.5 (H); pH 6.5 (I); pH 7.4 (J). K)
Absorption spectra of DTNB and GSH after treatment with HTVMz 24 h. L)
Diagram illustrating the enhanced catalytic activity of HTVMz 24 h.
Furthermore, motivated by the observed structural transformations in
HTVMz within the TME, we conducted a thorough examination of its
catalytic properties before and after transformation. In this study, 3,
3′, 5, 5′‐tetramethylbenzidine (TMB) was employed as a substrate to
probe peroxidase (POD)‐like activity. Notably, the VO[x] particles
derived from HTVMz exhibited a substantial elevation capacity to
generate hydroxyl radicals (·OH), significantly outperforming HTVMz
itself. This enhancement was clearly demonstrated by the emergence of a
blue oxidized product, characterized by a prominent absorption peak at
652 nm. Importantly, the enhancement in catalytic efficiency was
closely correlated with the degree of structural transformation of
HTVMz. Specifically, with increasing incubation time of HTVMz under
simulated tumor cell lysosome condition (100 mM H[2]O[2], pH 5.0) from
0 to 24 h, a notable increase in ·OH generation was observed
(Figure [108]4E). This highlighted that transforming HTVMz into
ultra‐small VO[x] particles, facilitated by the enlarged surface area
and reactive sites, greatly enhanced its catalytic activity. Moreover,
electron spin resonance (ESR) spectra provided additional evidence of
the enhanced catalytic activity in VO[x] particles, as shown in
Figure [109]4F. Notably, both pH and H[2]O[2] concentration also
exerted influence on its catalytic activity. As depicted in
Figure [110]4G, the catalytic efficacy of VO[x] progressively increased
with decreasing pH and increasing H[2]O[2] concentration
(Figure [111]4H–J). To further investigate the ROS generation
capability of HTVMz, the Michaelis‐Menten constant (Km) and maximum
velocity (Vmax) were calculated (Figure [112]S4, Supporting
Information). The initial reaction rates were determined and plotted
against TMB concentrations, with the data subsequently fitted to the
Michaelis–Menten equation (Figure [113]S4A, Supporting Information).
Vmax and Km for HTVMz were determined using the double‐reciprocal
method (Lineweaver‐Burke fitting) (Figure [114]S4B, Supporting
Information). The results obtained were as follows: for HTVMz, Vmax =
1.8392 µm min^−1 and Km = 0.1948 × 10^2 µM. These findings suggested
that the robust oxidative milieu and acidic environment inherent to the
TEM not only facilitated the structural transformation of engineered
HTVMz but also synergistically enhanced their catalytic activity,
ultimately amplifying their potential to induce oxidative stress
against tumor.
Besides, the VO[x] derived from HTVMz exhibited the ability to
disrupting redox homeostasis in tumor cells. This disruption stemmed
from the high valance state of VO[x], which reacted with GSH. To
monitor GSH depletion, DTNB (5,5′‐dithiobis‐(2‐nitrobenzoic acid)) was
utilized as a sensitive indicator. Upon reaction with the sulfhydryl
group of GSH, DTNB yields a yellow product, 5‐thio‐2‐nitrobenzoic acid,
with a characteristic absorption peak at 412 nm (Figure [115]4K),
providing a reliable quantification of GSH level fluctuations. In this
study, it was observed that V^5+ in VO[x] reacts with GSH to be reduced
to V^4+, thereby demonstrating its capability to consume GSH. This
catalytic process led to a decrease in the yellow DTNB‐derived product,
indirectly reflecting a reduction in GSH content, as elucidated in
Equations ([116](2), [117](3), [118](4)). As depicted in
Figure [119]4K, the absorbance at 412 nm gradually declined over time
in the presence of VO[x], confirming its role in GSH depletion. By
decreasing the levels of reduced GSH in tumor cells, VO[x]
significantly intensified intracellularROS generation, thereby
enhancing the efficacy of catalytic therapy, as illustrated in
Figure [120]4L.
2.4. Intracellular Catalytic Performances
Encouraged by the enhanced catalytic activity observed in HTVMz
post‐structural transformation, we conducted further evaluations of its
intracellular performance. Initially, we assessed its cellular uptake
efficiency, revealing a rapid and time‐dependent cellular uptake
profile during co‐culture. Moreover, HTVMz demonstrated progressive
uptake, reaching saturation levels at ≈8 h (Figure [121] 5M,N).
Following internalization, HTVMz initially accumulated within lysosomes
but exhibited notable escape and relocation to the ER within 4 h
(Figure [122]5A,B). This translocation was accompanied by significant
changes in lysosomal permeability at 4 h (Figure [123]5C,D), suggesting
an active response to the intracellular environment. To further
investigate the lysosomal damage mediated by HTVMz, acridine orange
(AO) staining was used as an indicator of lysosomal damage. In both the
control and Laser groups, strong red fluorescence from AO was observed,
signifying intact lysosomal structures. However, in the HTVMz treatment
group, a reduction in AO red fluorescence indicated partial disruption
of lysosomal integrity. Notably, the combination of HTVMz with laser
irradiation resulted in a complete loss of red fluorescence, indicating
severe lysosomal disruption and substantial oxidative stress‐induced
damage to 4T1 cells (Figures [124]S5 and [125]S6, Supporting
Information). Once localized within lysosomes, the structural
transformation of HTVMz was triggered, significantly amplifying their
capacity to generate ROS. This heightened ROS production subsequently
accelerated lysosomal damage, which in turn promoted the migration of
HTVMz from lysosomes to the ER.
Figure 5.
Figure 5
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Intracellular spatiotemporal distribution and in vitro antitumor
efficacy evaluation of HTVMz. A) Colocalization of HTVMz‐FITC (green)
with lysosomes (red) in 4T1 cells after incubation for different time.
Scale bar = 5 µm. B) Colocalization of HTVMz‐FITC (green) with the ER
(red) in 4T1 cells following incubation for different time. Scale bar =
5 µm. C) Live imaging of lysosomes in 4T1 cells using DND‐26 (green)
and Magic Red to illustrate LMP after 0.5 and 4 h of incubation with
HTVMz. Scale bar = 20 µm. D) The mean fluorescence intensity of Magic
Red following treatments at different time (n = 3). E) Schematic
representation of the intracellular trajectories of HTVMz. F)
Cytotoxicity of HTVMz at different concentrations in L929 cells (n =
5). G) Cytotoxicity of HTVMz at varying concentrations in 4T1 cells (n
= 5). H) Comparative toxicity in 4T1 cells of HTVMz 4 h, HTVMz 24 h,
HTVMz + Laser 24 h, Control, and only 1064 nm Laser (n = 5). I)
Cytotoxic effects of HTVMz (50 ppm) at different laser power densities
in 4T1 cells (n = 5). J) CLSM images of 4T1 cells after different
corresponding treatments and subsequent DCFH‐DA staining. Scale bar =
100 µm. K) FCM data quantification analysis of the ROS (n = 3). L)
Relative intracellular GSH content in 4T1 cells flow cytometry (n = 3).
M) FCM patterns of FITC‐labeled HTVMz. N) FCM data quantification
analysis of intracellular uptake of FITC‐labeled HTVMz (n = 3).
To comprehensively assess the ROS‐generating capacity of HTVMz within
4T1 cells, we employed the ROS‐sensitive indicator probe DCFH‐DA. A
comparative analysis of control and laser‐only groups demonstrated that
HTVMz notably induced ROS production following cellular uptake.
Moreover, prolonged incubation with HTVMz led to intensified
ROS‐generating (Figure [127]5J,K), confirming that the structural
transformation of HTVMz within the cell significantly enhances its
ROS‐producing capability. Additionally, the combination of HTVMz with
laser irradiation resulted in a marked increase in ROS fluorescence
intensity, underscoring the enhanced efficiency of the Fenton reaction
facilitated by the photothermal effect.
As ROS levels increased, the cellular redox metabolic system responded
by increasing GSH secretion to neutralize the excess ROS, thereby
maintaining dynamic redox balance equilibrium within the TME.
Remarkably, prolonged exposure to HTVMz for 24 h led to significant
depletion of intracellular GSH compared to a 4 h exposure, further
confirming the potent catalytic activity of VO[x] nanozymes formed
through the structural transformation of HTVMz (Figure [128]5L). These
findings suggested that the structural transformation of HTVMz within
TME not only enhanced its ROS‐generating capacity but also may
intensify oxidative stress, thereby improving its potential to inhibit
tumor cell growth.
To evaluate the role of H[2]O[2] consumption by HTVMz in therapeutic
efficacy, we observed a decrease in, intracellular H[2]O[2]
concentration as the co‐incubation time was extended, while ROS levels
continuously increased, indicating sustained ROS accumulation during
the treatment (Figure [129]S7A,D, Supporting Information). VO[x]
nanozymes contributed to this sustained ROS accumulation by consuming
GSH (Figure [130]S7A,C, Supporting Information). Furthermore, live/dead
staining results showed that HTVMz significantly induced both early and
late apoptosis, suggesting that the therapeutic effect could persist
over a prolonged period (Figure [131]S7E,F, Supporting Information).
Additionally, the catalytic efficiency of VO[x] was calculated (Figure
[132]S4, Supporting Information), confirming its high catalytic
activity. This activity enabled VO[x] to generate substantial ROS even
under limited H[2]O[2] conditions, thereby enhancing its ability to
inhibit tumor cell growth (Figure [133]S7G, Supporting Information).
2.5. In Vitro Antitumor Efficacy Evaluation
Given the promising photothermal properties of HTVMz and its ability to
induce robust intracellular ROS production, we conducted a
comprehensive study to evaluate its potential for suppressing tumor
cell proliferation. MTT assays demonstrated that prolonged incubation
periods (12, 24, and 36 h) with various concentrations of HTVMz had
negligible effects on the viability of normal L929 cells
(Figure [134]5F). In contrast, HTVMz induced significant cytotoxicity
in 4T1 tumor cells (Figure [135]5G). This effect was attributed to the
structural transformation of HTVMz into smaller nanosized VO[x] within
the TME, thereby enhancing ROS production and effectively inhibiting
tumor cell growth. Specifically, a 48‐h incubation of 4T1 cells with
HTVMz containing 100 ppm V resulted in a notable decrease in cell
viability to 53%, highlighting the potent antitumor activity of ROS
mediated by HTVMz.
Further investigations revealed that the photothermal effect of HTVMz,
combined with NIR‐II irradiation, synergistically enhanced its
antitumor efficacy through ROS generation. The increase in laser power
also intensified the phototoxic effect (Figure [136]5I), as evidenced
by the comparative analysis of cell viability (Figure [137]5H).
Specifically, compared to control groups exposed only to laser
irradiation or treated with HTVMz for 4 h, which showed minimal changes
in cell viability. However, the cell viability decreased to 69.67% when
treated with HTVMz prolonged to 24 h. Remarkably, additional exposure
to 1064 nm laser irradiation further increased cytotoxicity,
drastically reducing cell viability to 36.09%. These findings were
supported by laser confocal microscopy observations of cell apoptosis
(Figure [138]S8A, Supporting Information), further reinforcing the
synergistic antitumor efficacy resulting from combined ROS and
photothermal effects. The live/dead staining outcomes provided
compelling evidence that the synergy of HTVMz with NIR laser optimally
suppressed tumor growth (Figure [139]S8B, Supporting Information),
emphasizing the therapeutic potential of HTVMz in tumor treatment
through its photothermal and cascade catalytic properties. The flow
cytometry analysis further confirmed that HTVMz exhibited optimal
therapeutic effects on 4T1 cells after laser irradiation (Figure
[140]S9A,B, Supporting Information).
2.6. Mechanism of Structure‐Transformation‐Induced Oxidative Stress
Enhancement in HTVMz
To elucidate the anticancer mechanism of HTVMz‐induced oxidative stress
through structural transformation combined with photothermal effect, we
performed transcriptomic analysis to investigate the gene expression
changes in tumor cells under different treatment conditions. Principal
component analysis (PCA) and FPKM violin plots (Figure [141]S10A,B,
Supporting Information) clearly showed distinct gene expression
profiles among the treatment groups. Analysis of co‐localization of
HTVMz in lysosomes and the ER revealed that after 4 h, the internalized
material had fully translocated to the ER. Therefore, HTVMz was
co‐incubated with cells for 4 h to simulate its reaction within
lysosomes. Volcano plots (Figures [142] 6A–C and [143]S11, Supporting
Information) highlighted significant alterations in gene expression
between HTMVz treatments lasting 4 h (initial uptake by 4T1 cells) and
24 h (After 4 h of uptake, wash and extend the incubation period).
Notably, the 24‐h treatment obviously downregulated cell cycle genes
while upregulating apoptosis‐related genes, and modulated ER stress
genes (Atf6b, Creb3l3, Cped1, Alox15) and DNA damage genes (Myc, Batf,
Bax, Blk). Gene pathway enrichment analysis (Figure [144]6E,F) revealed
that prolonged exposure to HTVMz (HTVMz 24 h) significantly enhanced
pathways associated with lysosomal membrane permeability (LMP),
ferroptosis, apoptosis, and ER stress compared to shorter treatment
(HTVMz 4 h) (Figure [145]6D). Specially, the Oxidative Stress and Redox
Pathway was prominently upregulated in the HTVMz 24 h group (Figure
[146]S12, Supporting Information), indicating heightened oxidative
stress levels within 4T1 cells. Further validation through Western Blot
(WB) experiments demonstrated significant changes in proteins related
to ferroptosis and ER stress. There was growing evidence indicating
that ferroptosis was an autophagy‐dependent cell death associated with
iron accumulation and lipid peroxidation, which played a crucial role
in anticancer activity.^[ [147]^39 , [148]^40 , [149]^41 ^] As shown in
Figure [150]6J,K, tumor cells treated with HTVMz for 24 h induced
increased LC3B‐I/II conversion and decreased STSQM1 expression,
indicative of enhanced autophagy (Figure [151]S15, Supporting
Information).^[ [152]^42 ^] Additionally, the upregulation of GRP78 and
DDIT3 promoted ER stress,^[ [153]^43 ^] while the degradation of GPX4,
and downregulation of HO‐1 and AKT contributed to ferroptosis
induction.^[ [154]^44 ^] Additionally, the degradation of heat shock
protein HSP90 reduced thermal stress tolerance in tumor cells.^[
[155]^45 ^] Notably, the upregulation of Cleaved caspase‐3 indicated
enhanced apoptosis. Metabolomics analysis (Figure [156]S16A,B,D,
Supporting Information) further supported these findings, showing
significant alterations in pathways including glutathione metabolism,
arginine and proline metabolism, and amino acid metabolism upon HTVMz
24 h treatment (Figure [157]S18, Supporting Information). In contrast,
HTVMz 4 h treatment had a minor impact on tumor cell metabolism (Figure
[158]S19, Supporting Information). Taken together, these results
underscored that HTVMz 24 h treatment enhanced oxidative stress,
markedly disrupted tumor cell metabolism, and promoted pathways leading
to ferroptosis and apoptosis, thereby elucidating its potent anticancer
effects.
Figure 6.
Figure 6
[159]Open in a new tab
The mechanism of TME drove structural transformations of vanadium‐based
MXenzymes to enhance oxidative stress and activated multimodal therapy
against breast cancer. (cz: Control; maz: HTVMz 24 h; mbz: HTVMz for
4 h; mdz: HTVMz + Laser 24 h). A) Volcano plot illustrating DEGs in 4T1
cells treated with mbz versus cz. B) Volcano plot of DEGs in 4T1 cells
following maz versus cz treatment. C) Volcano plot depicting DEGs in
4T1 cells post‐treatment with mdz versus cz. D) Functional enrichment
analysis of upregulated DEGs comparing mbz with cz. E) Functional
enrichment analysis of upregulated DEGs for maz versus cz. F)
Functional enrichment analysis of upregulated DEGs in the comparison of
mdz with cz. G) GSEA map showcasing protein processing in the
endoplasmic reticulum for mdz versus cz. H) GSEA map of lysosome
organization comparing cz with mdz. I) Schematic representation of the
role of HTVMz in regulating apoptosis in tumor cells. J) Immunoblot
analysis revealing proteins associated with intracellular autophagy, ER
stress, heat shock, ferroptosis, and apoptosis, evaluated by WB. K)
Quantification of GPR78, STSQM1, GPX4, LC3B, HSP90, AKT, HO‐1, and
Cleaved‐Caspase3 proteins normalized to GAPDH. Data are presented as
mean ± standard deviation (s.d.).
Surprisingly, when combined with laser irradiation, the HTVMz 24 h
treatment exhibited an enhanced photothermal effect due to decreased
HSP90 protein expression, indicating upregulation pathways associated
with ER stress, ferroptosis, and lysosomal membrane integrity
(Figure [160]6F). Furthermore, this treatment regimen led to
significant impairment of ER processing and lysosomal function
(Figure [161]6G,H; Figures [162]S13 and [163]S14, Supporting
Information). WB results provided additional validation, showing that
HTVMz 24 h combined with laser treatment promoted ferroptosis and
apoptosis through the cascade catalytic reactions involving lysosomes
and ER (Figure [164]6I).^[ [165]^46 ^] Metabolomics analysis revealed
profound disruptions in tumor cell metabolism following HTVMz 24 h
combined with laser treatment, including notable downregulation of
metabolites such as L‐cysteine, pyrrole, and pyruvate (Figure
[166]S16D,G, Supporting Information). Key metabolic pathways, including
tyrosine metabolism, tricarboxylic acid cycle, alanine, aspartate and
glutamate metabolism, pyruvate metabolism, and lysosomal pathways were
also significantly downregulated (Figure [167]S16E,F, Supporting
Information). These findings suggested the substantial impact of HTVMz
24 h combined with laser treatment on tumor cell metabolism, energy
supply, and proliferation. From the above results, it was noted that
HTVMz 24 h treatment significantly boosted LMP through an oxidative
stress induced by enhanced photothermal catalytic reactions and
alterations in lysosomal internal architecture. This process
facilitated the transfer of VO[x] conversion products from lysosomes to
the ER, triggering ER stress. Additionally, VO[x]‐mediated degradation
of GSH reduced GPX4 expression, promoting ferroptosis. Simultaneously,
reduced HSP90 levels enhanced the efficacy of NIR‐II PTT. This
synergistic mechanism harnessed oxidative stress initiated by HTVMz
structural transformations to amplify the therapeutic efficacy of
NIR‐II PTT, ultimately enhancing the multimodal treatment outcomes for
tumors.
2.7. In Vivo Anti‐Tumor Assay
Motivated by encouraging in vitro results, we initiated a preliminary
investigation into the biodistribution of HTVMz in mice bearing 4T1
tumors. The distribution and accumulation of HTVMz at tumor sites were
pivotal for their photothermal and ROS generation capabilities, which
determined their anti‐tumor efficacy. Following the intravenous
injection of HTVMz to tumor‐bearing mice, we utilized inductively
coupled plasma optical emission spectrometry (ICP‐OES) to measure the
vanadium content at tumor sites. Notably, the vanadium concentration
reached a significant 0.64 µg g^−1 after intravenous injection for 8 h,
subsequently decreasing to 0.41 µg g^−1 at 24 h (Figure [168]S20,
Supporting Information). This observation underscored the effective
targeting of HTVMz at tumor sites via systemic circulation, leveraging
the enhanced permeability and retention (EPR) effect.^[ [169]^47 ,
[170]^48 , [171]^49 ^] Furthermore, upon exposing the tumor sites to
NIR‐II laser irradiation, a rapid temperature increased to ≈57 °C was
observed, while tumors in the control group maintained temperatures
≈37 °C (Figure [172] 7B,C). These findings further validated the
tumor‐targeting ability of HTVMz and their potential for effective
photothermal therapy at tumor site.
Figure 7.
Figure 7
[173]Open in a new tab
In vivo antitumor efficacy of HTVMz. A) Schematic of the in vivo
protocol against 4T1 tumors. B) Photothermal heating photos of tumor
mice after injection of HTVMz. C) Photothermal heating of tumor mice of
HTVMz + Laser and only laser. D) Graph showing the body weight of mice
over time (n = 5). E) Tumor volume in mice across different treatment
groups (n = 5). F) Tumor weight in various treatment groups (n = 5). G)
Tumor photos of each experimental group. H) Photographs of mice on days
1, 7, and 13 for each group. I) Heatmap illustrating the tumor
inhibition rate in different groups. J) The tumors in different
experimental groups were stained with H&E, TUNEL, and Ki67. Scale bar =
20 µm. Note: G0: Control group; G1: only Laser group; G2: only HA
group; G3: HA + Laser group; G4: HTVMz group; G5: HTVMz + Laser group.
Subsequently, we assessed the anticancer efficacy of HTVMz using a 4T1
tumor mouse model. As shown in Figure [174]7A, the control group (only
PBS), only Laser group, HA group, and HA + Laser group exhibited a
consistent increase in tumor volume over time without significant
variations. In contrast, treatment with HTVMz alone, especially in
combination with laser treatment, markedly suppressed tumor growth,
with the HTVMz + Laser combination displaying the most significant
antitumor effect. This observation was further supported by analyses
including tumor volume (Figure [175]7E), photographs of tumor‐bearing
mice (Figure [176]7G), images of dissected tumors post‐treatment
(Figure [177]7H), and tumor weight (Figure [178]7F), collectively
indicating the robust antitumor therapeutic efficacy of HTVMz combined
with NIR irradiation (Figure [179]7I). Furthermore, direct detection of
ROS generation within tumor tissues confirmed that HTVMz in combination
with laser treatment elicited significantly elevated ROS levels,
effectively harnessing photothermal synergy to hinder tumor progression
(Figure [180]S21, Supporting Information).^[ [181]^48 , [182]^50 ^]
TUNEL staining of tumor tissue sections revealed the highest apoptosis
rate in the HTVMz + Laser group (Figure [183]7J). Moreover, reduced
expression of the proliferation marker Ki67 also indicated effective
suppression of tumor growth. Hematoxylin and eosin (H&E) staining
revealed more pronounced nuclear damage in the HTVMz + Laser group
compared to other treatments (Figure [184]7J), providing additional
evidence for accelerated tumor cell apoptosis. Overall, these findings
collectively underscored the potent antitumor capabilities of HTVMz in
synergy with laser treatment. Immunohistochemistry (IHC) analysis
(Figure [185]S22, Supporting Information) further demonstrated the
upregulation of active‐caspase 3 and decreased SQSTM1 levels in HTVMz
combined with laser‐treated tumor tissues, indicating the activation of
autophagy and apoptosis pathways. Additionally, IHC analysis of
ferroptosis‐related markers, such as GPX4, confirmed that HTVMz + Laser
treatment induced ferroptosis in vivo (Figure [186]S23, Supporting
Information). H&E staining of tumor‐adjacent tissues after laser
irradiation showed no significant increase in inflammation in the
adjacent skin (Figure [187]S24, Supporting Information), suggesting the
safety of the treatment. Importantly, all treatment groups exhibited
stable body weights (Figure [188]7D), suggesting no adverse effects on
overall health post‐HTVMz treatment. Moreover, H&E staining, along with
blood routine tests and biochemical assessments of main organs
including the heart, liver, spleen, lungs, and kidney, all showed
values within normal physiological ranges (Figures [189]S25 and
[190]S26, Supporting Information), affirming the excellent
biocompatibility and safety profile of HTVMz. These results
collectively highlighted HTVMz as a promising therapeutic agent with
minimal systemic toxicity.
2.8. In Vivo Safety Valuation
To ensure the safety of HTVMz for biological applications, we then
evaluated its blood biocompatibility. The hemolysis assay results
demonstrated minimal hemolytic activity of HTVMz against red blood
cells, indicating good blood compatibility (Figure [191]S27, Supporting
Information). Further evaluation of systemic toxicity in Balb/C mice
involved varying intravenous doses of HTVMz. Mice treated with 5 and
10 mg kg^−1 maintained stable body weights comparable to the control
group, as depicted in Figure [192]8A. Most nanoparticles were primarily
metabolized and cleared through the liver upon entering the animal
body, hence hepatic function was assessed in the experimental mice. As
depicted in Figure [193]8B, biomarkers of liver function such as ALT
and AST showed no significant changes, indicating no adverse effects on
hepatic function. Histological examination using H&E staining of major
organs confirmed the absence of inflammation, underscoring its
excellent biocompatibility (Figure [194]8D). To investigate the
metabolic dynamics of HTVMz in healthy mice in detail, HTVMz were
administered intravenously via the tail vein. Subsequently, ICP‐OES
quantified vanadium levels in the heart, liver, spleen, lungs, and
kidneys on the 1st and 14th days after injection. As shown in
Figure [195]8C, a significant reduction in vanadium content across all
assessed organs was observed by the 14th day, indicating efficient
metabolic clearance of HTVMz by mouse tissues. In conclusion, these
findings underscored the remarkable biocompatibility of HTVMz,
positioning it as a highly promising candidate for further clinical
exploration and potential advancement in therapeutic applications.
Figure 8.
Figure 8
[196]Open in a new tab
Long‐term biotoxicity analysis following intravenous (i.v.) injection
of HTVMz in Balb/C mice. A) Time‐dependent body weight changes in
Balb/C mice following i.v. injection of HTVMz at various concentrations
(n = 3). B) Blood biochemical parameters in mice post i.v. injection of
HTVMz at different concentrations (n = 3). C) Distribution of vanadium
in the heart, liver, spleen, lung, and kidney measured by ICP‐OES 1 and
14 days post‐caudal i.v. injection of HTVMz in healthy mice (n = 3). D)
H&E‐stained microscopic images of major organs from Balb/C mice post
i.v. injection of HTVMz at varied concentrations. Scale bar = 50 µm.
3. Conclusion
In conclusion, our study introduced an approach that leveraged the
unique characteristics of the tumor microenvironment to induce a
structural transformation in sheet‐like TVMz, thereby revolutionizing
MXenzyme‐mediated tumor catalytic therapy. This transformation,
delicately orchestrated by NIR‐II laser irradiation and further
reinforced by the tumor‐specific environment, yield ultra‐small VO[x]
that potently amplified oxidative stress and provoke a diverse range of
intricate cellular stress responses. Encapsulated within hyaluronic
acid‐coated TVMz (HTVMz) to ensure stability, this strategy not only
enhanced lysosomal permeability and endoplasmic reticulum stress but
also disrupted redox homeostasis through intricate interactions
involving high‐valent vanadium species and intracellular glutathione.
The synergistic augmentation of these effects accelerated tumor cell
apoptosis, triggered ferroptosis, and inhibited HSP90 expression,
thereby enhancing thermal sensitivity and facilitating multimodal
therapeutic outcomes. Taken together, this precise engineering of
MXenzyme structures represented a promising avenue for catalytic
therapy, offering significant potential to refine and advance tumor
treatment strategies.
4. Experimental Section
Materials
V[2]AlC powders were purchased from the FoShan Xinxi technology Co.,
Ltd. LTD. Hydrofluoric acid (HF, 40 wt.%) was purchased from Kelong
Chemical (Chengdu, China) Co., Ltd. Tetramethyl ammonium hydroxide
(TMAOH) was purchased from Shanghai Maclean Biochemical Technology Co.,
Ltd. Lithium chloride (LiCl) was purchased from Beijing Leyan
Technology Co., Ltd. Hyaluronic acid, 5, 5′‐Dithiobis (2‐nitrobenzoic
acid) and 3, 3′, 5, 5′‐Tetramethylbenzidine were purchased from
Shanghai Aladdin Biochemical Technology Co., Ltd. Hydrogen peroxide was
purchased from Chengdu Jinshan Chemical Reagent Co., Ltd. Glutathione
was purchased from Shanghai Adamas Reagent Co., Ltd. GSH Kits, Hoechst
33 342 and Reactive oxygen test kit were purchased from Beyotime
Biotechnology. TRIzol was purchased from Thermo Fisher Scientific, USA.
Fluorescein isothiocyanate (FITC) was purchased from TCI Shanghai
Company. 4T1 and L929 were all bought by the Chinese Academy of Science
Cell Bank for Type Culture Collection (Shanghai, China). Lyso‐Tracker
Red and ER‐Tracker Red were purchased from Beyotime Biotechnology.
ROSGreen H[2]O[2] Probe was purchased from Shanghai Maokang
Biotechnology Co., Ltd. ThiolTracker Violet was purchased from Thermo
Fisher Scientific. The Acridine Orange Staining Kit was purchased from
Beyotime Biotechnology. Annexin V‐FITC/PI was purchased from Beijing 4A
Biotech Co., Ltd.
Preparation of 2D HTVMz
First, TVMz were prepared following Yury Gogotsi's literature method,
with modifications specifically tailored for biomedical applications.^[
[197]^29 ^] 1 g of V[2]AlC was dispersed in 20 mL of etching solution
(12 mL HF and 8 mL HCl) under ice bath conditions while stirring the
solution at low speed (150 rpm) for 5 min. Then transferred to an oil
bath and reacted at 400 rpm stirring at 50 °C for 72 h. Subsequently,
centrifuge and add 25 mL of 5 wt.% TMAOH, stirring for 6 h.
Subsequently, the solution was centrifuged and after stirring, the
solution was centrifuged at 3500 rpm for 10 min, the supernatant was
discarded and repeatedly washed to pH > 5.5. The product was prepared
by sonication and centrifugation of the TVMz. The organic intercalator
TMA^+ was removed using an ion exchange process by adding 19.8 m LiCl
solution to the TVMz solution in a volume ratio of 1:5. The resulting
mixture underwent dialysis to remove LiCl. The TVMz solution was
collected and freeze‐dried. Finally, HA (5 mg mL^−1) was weighed and
dissolved in deoxygenated deionized water, added to TVMz solution,
transferred to a hydrothermal reactor, and reacted at 90 °C for 1 h to
construct HTVMz. The HTVMz was then evenly redispersed in
deoxygenated deionized water and stored at 4 °C. FITC (2 mg mL^−1) was
dissolved in the ethanol solution (5 mL) and stirred vigorously
together with HTVMz (10 mg) for 6 h under a nitrogen atmosphere to
obtain FITC‐HTVMz. The product was centrifuged several times to remove
the unlabeled FITC.
Characterization
The phase structure of obtained materials was characterized by using
X‐ray diffraction (XRD, Philips PC‐APD) with Cu Kα radiation (λ =
1.54056 Å, 40 kV and 40 mA), operated at 2θ range from 10° to 80° with
the scanning rate of 10°min^−1. The ζ‐potentials and sizes of the
prepared nanoplatforms were characterized by dynamic light scattering
(DLS, Malven Zetasizer Nano ZS), and the morphology was observed by
atomic force microscopy (AFM Dimension Icon) and transmission electron
microscopy (TEM, Talos FEI 200). The UV–vis spectra were detected by
the UV–vis spectrophotometer (Hitachi U3900). The organs of mice
samples were quantified on the inductively coupled plasma‐optical
emission spectrometry (ICP‐OES, Agilent 725). Electron spin resonance
spectra (ESR) were observed by the Bruker EMXplus X‐ban.
Thermo‐Gravimetric was characterized by the TGA/DSC2. Infrared spectra
were observed using the NEXUS 670. X‐ray photoelectron spectroscopy
(XPS) characterization was performed with the Thermo Kalpha. The
material was freeze‐dried using the FTFDS GX2502 device.
The Photothermal Performance Test
First, the HTVMz solution (1 mL) in ultrapure water was irradiated with
a 1064 nm laser (1.0, 1.5, and 2.0 W cm^−2 for 10 min). HTVMz solutions
at different concentrations (12.5, 25, and 50 ppm) were then irradiated
with a 1064 nm NIR laser (1.0 W cm^−2, 10 min). To further determine
the photothermal stability, the HTVMz solution (25 ppm) was irradiated
at 1064 nm (1.0 W cm^−2, 10 min) and cooled to room temperature after
turning off the laser for five heating‐cooling cycles to record the
temperature change in the solution. The calculation of photothermal
conversion efficiency during the heating‐cooling process was calculated
from other studies reported in the literature.^[ [198]^51 ^]
The Calculation of the Extinction Coefficient
The UV absorption spectra of the material at different concentrations
(0.05, 0.025, 0.0125, and 0.00625 g L^−1) were measured, and the
absorbance at 808 nm (A[808]) was recorded.
The absorption coefficient (𝛼) was calculated using Beer–Lambert Law,
given the known solution concentration (c) and optical path length
(l).^[ [199]^52 ^]
[MATH: α808=A808c·l :MATH]
(1)
where A[808] represented the absorbance at a wavelength of 808 nm, c
was the concentration, and l was the optical path length of the sample
cell.
Finally, the calculation was performed using the following equation:
[MATH: k=α808·λ4π :MATH]
(2)
where the known absorption coefficient (𝛼 [808]) and the wavelength (λ
= 808 ×10^−9 m), the extinction coefficient (k) was obtained.
The Calculation of the Photothermal Conversion Efficiency
The photothermal conversion efficiency (η) of HTVMz was calculated
using the following equation:^[ [200]^53 ^]
[MATH: η=hsΔTHTVMz−ΔTWaterI×<
/mo>1−10−A
:MATH]
(3)
where A was the absorbance of the water at 808 nm, and ΔT[HTVMz] and
ΔT[water] represented the temperature changes of the sample and the
blank, respectively. The heat transfer coefficient was denoted as h,
and s represented the surface area of the container. These parameters
were determined using the following formulas:
[MATH: hs=mcτs :MATH]
(4)
where m was the mass of the solution (≈1 g), c was the specific heat
capacity of the solvent (for water, 4.2 J g^−1 °C^−1), and τ[s] was the
time constant, which could be determined during the cooling period
using the following equation:
[MATH: t=−τs<
mo linebreak="goodbreak">×lnθ :MATH]
(5)
where θ was a dimensionless parameter, which was varied by time and
driven by temperature and is defined as follows:
[MATH: θ=T−T
SurrTMax−TSur
r :MATH]
(6)
where T[Max] and T[Surr] were the maximum steady state temperature and
the environmental temperature.
The Calculation of the Catalytic Efficiency
The conversion rate of absorbance at 658 nm, flowing the reaction of
HTVMz with varying concentrations of TMB (400, 200, 100, 50, 25, and
12.5 µm) and H[2]O[2] (500 µm), was used to determine catalytic
reaction velocity(ν), expressed in micromoles per minute. The
absorbance was plotted against the substrate concentrations to generate
a Michaelis–Menten curve.^[ [201]^54 ^]
The kinetics constants ν[max] and K[m] were determined by fitting the
reaction velocity values to the Michaelis–Menten equation as follows:
[MATH: v=vmax×S/Km+S
:MATH]
(7)
where ν was the initial reaction velocity and ν[max] was the maximal
reaction rate observed at saturating substrate concentrations. [S]
represented the substrate concentration, and K[m] was the Michaelis
constant. K[m] reflected the affinity of the nanozyme for its substrate
and was defined as the substrate concentration at which the reaction
rate was half of V[max].
The molar concentration of the nanozyme in the samples was measured by
a nanoparticle‐tracking analysis system. To ensure accurate
nanoparticle counting, the nanozyme suspension must be monodisperse.
The catalytic constant (k[cat]) was calculated by the following
equation:
[MATH:
kcat
=vmax/E :MATH]
(8)
where k[cat] was the rate constant that defined the maximum number of
substrate molecules converted to product per unit of time. [E] was the
nanozyme concentration.
In Vitro Catalytic Activity Test
To compare the catalytic activity of HTVMz before and after structural
transformation, HTVMz was exposed to simulated tumor cell lysosomal
conditions (pH 5.5, H[2]O[2] concentration of 100 µm), followed by
dialysis, and UV‐vis absorption spectra of the TMB and H[2]O[2]
reaction were recorded at 0, 12, and 24 h. The HTVMz were maintained at
pH 5.5 and under 100 µm H[2]O[2] conditions for 24 h, after which the
products were dialyzed and collected for characterization. Oxidation of
TMB in the presence of H[2]O[2] was carried out to study the peroxidase
activity of the HTVMz structure transformation product VO[x] (HTVMz
24 h). To achieve maximum catalytic effect, the pH was adjusted to 5.5,
6.5, and 7.4. Next, the detailed operations were as follows: first,
HTVMz 24 h (100 ppm), TMB (200 µm), and H[2]O[2] (100 µm) were added to
acetate buffer solutions (pH 5.5, 6.5, and 7.4), incubated for 5 min
and the solutions were transferred to a quartz cell and the UV
absorption at 652 nm was measured with a UV‐2600 spectrophotometer at
different pH conditions. Second, HTVMz 24 h (100 ppm), TMB (200 µm),
and different concentrations of H[2]O[2] (100, 50, 25, 12.5, and 6.25
µm) were added to acetate buffer solutions (pH 5.5, 6.5, and 7.4),
incubated for 5 min and finally the solutions were transferred to a
quartz cell and the UV‐2600 spectrophotometer was used to The UV
absorption at 652 nm was measured for HTVMz 24 h at different H[2]O[2]
concentrations (pH 5.5, 6.5, and 7.4).
A solution of DTNB (100 µm), GSH solution (100 µm), and HTVMz 24 h
solution (100 ppm) were mixed and incubated for 5 min and 10 min. The
absorbance was measured using a UV‐2600 spectrophotometer to
characterize the GSH consumption performance. The change in intensity
at 412 nm was recorded by photographing the color change.
[MATH:
V4++H2O
2+H+→
V5++·OH+H2O
(4−1) :MATH]
(9)
[MATH:
2V5+
+2GSH
→2V4++GSSG+2H+
(4−2) :MATH]
(10)
Cell Cytotoxicity
The cell viability of HTVMz was determined by MTT method. First, cells
(L929 or 4T1) were grown in 96‐well plates for 24 h. Different
concentrations of HTVMz were added to each well and incubated for 12,
24, 36, and 48 h. Then, DMEM or 1640 medium was replaced with
serum‐free medium (100 µL) containing MTT and incubated at 37 °C for
another 4 h. The medium was removed, DMSO (100 µL) was added and shaken
for 5 min to dissolve the blue‐purple crystalline methanogens, and the
peak absorbance (490 nm) was measured using an enzyme marker.
Phagocytose of HTVMz by 4T1 Cells and Co‐Localize in Lysosomes and ER
To detect the uptake of HTVMz by lysosomes in tumor cells, 4T1 cells
were cultured in laser confocal culture dishes. FITC‐labeled HTVMz
suspension was added to the fresh culture medium. At different time
points (0, 1, 3, and 6 h), Lyso‐Tracker Red dye solution (50 nm) and
Hoechst dye solution were added to the culture dishes to label
lysosomes and nuclei. At different time points (0, 2, 4, and 8 h),
ER‐Tracker Red dye solution (50 nm) and Hoechst dye solution were added
to the culture dishes to label lysosomes and nuclei. The localization
of HTVMz in lysosomes and the ER was observed using CLSM, and the data
were analyzed using Image J software.
In Vitro Phototoxic Evaluation
The MTT method was used to characterize the phototoxicity of HTVMz at
1064 nm. 4T1 cells were added to 96‐well plates and cultured for 24 h.
Subsequently, 100 µL of medium containing HTVMz was added at different
time points. Analysis of co‐localization of HTVMz in lysosomes and the
ER revealed that after 4 h, the internalized material had fully
translocated to the ER. Therefore, HTVMz was co‐incubated with cells
for 4 h to simulate its reaction within lysosomes. The experimental
group received 100 µL of medium containing HTVMz, which was incubated
for 4 h, then washed and supplemented with fresh medium prior to laser
irradiation, and further incubated for 24 h (designated as the HTVMz +
Laser 24 h group). Cells treated with medium alone served as the
control group. The cells were irradiated with a 1064 nm (1.0 W cm^−2)
laser for 10 min. The survival rate of 4T1 cells was assessed by MTT
method. 4T1 cells containing HTVMz were irradiated with 1064 nm laser
at different power levels (0.5, 1.0, 1.5, and 2.0 W cm^−2), followed by
the MTT method to evaluate the cell survival rate. Cell apoptosis was
assessed using Annexin V‐FITC/PI staining, and the apoptotic status of
4T1 cells co‐incubated with HTVMz for different time intervals was
observed via confocal microscopy. Additionally, apoptosis in the
experimental groups was quantitatively analyzed by flow cytometry. The
apoptotic status of cells co‐incubated with HTVMz for different time
intervals was assessed to evaluate the sustained effects of the
treatment.
Detection of Intracellular ROS Production
4T1 cells were cultured in glass‐bottom dishes for 12 h. The cells were
divided into 6 groups (n = 3): control, laser 1064 nm, HTVMz 4 h
(treated with medium containing HTVMz for 4 h), HTVMz + Laser 4 h
(treated with 100 µL of medium containing HTVMz for 4 h followed by
laser irradiation), HTVMz 24 h (treated with medium containing HTVMz
for 4 h followed by supplementation with fresh medium and extended
incubation to 24 h), and HTVMz + Laser 24 h group (treated with medium
containing HTVMz for 4 h followed by supplementation with fresh medium,
laser irradiation, and extended incubation to 24 h). After adding the
medium containing HTVMz at different time points, the treated cells
were washed three times with PBS. The DCFH‐DA probe was then added to
the glass‐bottomed dishes, stained for 20 min, and washed three times
for CLSM observation, with the laser irradiation power set at 1 W
cm^−2. A green fluorescent signal representing ROS was observed at an
excitation wavelength of 488 nm. In addition, the amount of
intracellular ROS was quantified by FCM after labeling with a DCFH‐DA
probe (10 mm). HTVMz was co‐incubated with 4T1 cells for different time
periods, and the changes in intracellular ROS fluorescence intensity
over time were evaluated using CLSM.
The Evaluation of Intracellular GSH Depletion
4T1 cells were added to six‐well cell culture plates and after 12 h of
cell wall proliferation, The cells were divided into 5 groups (n = 3):
control, HTVMz 4 h, HTVMz + Laser 4 h, HTVMz 24 h, and HTVMz + Laser
24 h group. Incubation with HTVMz‐containing medium at different time
points was followed by PBS washing, digestion, and transfer into
centrifuge tubes. Samples were weighed before and after transfer. The
subsequent operations were carried out according to the GSH and GSSG
assay kits. HTVMz was co‐incubated with 4T1 cells for different time
intervals, and changes in intracellular GSH fluorescence intensity over
time were evaluated using a GSH probe and CLSM.
Evaluation of Intracellular H[2]O[2]
4T1 cells were cultured in a confocal dish (3 × 10^5 cells per well)
for 12 h and washed with PBS. Cells were incubated with 1× ROSGreen
H[2]O[2] Probe for 30 min to assess changes in intracellular H[2]O[2]
levels after co‐incubation of HTVMz with 4T1 cells for different time
intervals. Subsequently, a microplate reader was used to quantitatively
evaluate the variations in intracellular H[2]O[2] levels following
co‐incubation of HTVMz with 4T1 cells for different time points.
Lysosomal AO Staining Assessment
4T1 cells were cultured in a confocal dish (3 × 10^5 cells per well)
for 24 h and washed with PBS. AO staining solution was then added and
incubated for 10 min. The staining solution was aspirated, and the
cells were washed with PBS. The red and green fluorescence of the cells
under different treatments were evaluated using CLSM.
Imaging Lysosomes in Living Cells Using Magic Red and DND‐26 Probes
4T1 cells were divided into three groups and incubated with culture
medium containing Magic Red (1:1000; ImmunoChemistry Technologies,
Bloomington MN) for 40 min at 37 °C, followed by the addition of
Lyso‐Tracker Green DND‐26 (50 nm; Thermo Fisher, Waltham, MA) for
40 min. The dye‐containing medium was then replaced with fresh cell
culture medium. Two groups of cells were incubated with HTVMz for 0.5
or 4 h, respectively, while a third group was maintained at 37 °C
without treatment as the normal control. Confocal microscopy was used
to image the cells in each group.
Western Blot Test
After treating 4T1 cells with lysis buffer for protein extraction, the
protein concentration was determined using the BCA method.
Subsequently, proteins were separated by sodium dodecyl
sulfate‐polyacrylamide gel electrophoresis (SDS‐PAGE) and transferred
to a nitrocellulose membrane. The membrane was then blocked under
conditions containing 5% nonfat milk. Through WB experiments
characterizing the expression of related proteins: LC3B (Zen‐bioscience
Co., Ltd, 81631), STSQM1 (Huaan Biotechnology Co., Ltd, HA721171),
GAPDH (Huaan Biotechnology Co., Ltd, ET1601‐4), DDIT3 (Huaan
Biotechnology Co., Ltd, ET1703‐05), GPX4 (Cell Signaling Technology,
#59735), HSP90 (Huaan Biotechnology Co., Ltd, SY46‐01), AKT (Cell
Signaling Technology, #4685), HO‐1 (Cell Signaling Technology, #43966),
Cleaved caspase3 (Cell Signaling Technology, #9664), and GPR78 (Huaan
Biotechnology Co., Ltd, ER40402), it was confirmed that the
nano‐carrier can achieve NIR‐II photothermal and ferroptosis
dual‐enhanced therapy for breast cancer through a cascading catalytic
reaction from lysosomes to ER. All WB band results were captured using
the E‐BLOT contact chemiluminescence imaging system.
In Vivo Anti‐Tumor Evaluation
The animal studies were performed according to “The Animal Management
Rules of the Ministry of Health of the People's Republic of China
(Document no. 55, 2001) and the institutional guidelines.” All
experiments were approved by the Animal Care and Use Committee of
Sichuan University. Female Balb/C mice (4–5 weeks old, 18–22 g) were
purchased from Ensiweier Biotechnology Co., Ltd. A 4T1 mammary tumor
model was established, and 100 µL PBS containing 4T1 tumor cells (1.0 ×
10^7 mL^−1) was injected subcutaneously into the right buttock of the
mice. To evaluate their tissue distribution status, Balb/C
tumor‐bearing mice were injected with HTVMz and executed at different
time intervals (1 and 14 days) after intravenous injection, and heart,
liver, spleen, lung, and kidney were collected, and then the V content
in the tissues was determined according to ICP‐OES. Afterward, blood
was taken from the eyes of the mice and hemolysis assays were
performed. When the tumor volume (V = width^2 × length/2) increased to
≈150 mm^3, mice were randomly divided into (G0) Control group (saline,
pH 7.4); (G1) only Laser group (1.0 W cm^−2 for 10 min); (G2) only HA
group; (G3) HA + Laser group (1.0 W cm^−2 for 10 min); (G4) HTVMz; (G5)
HTVMz + Laser group (1.0 W cm^−2 for 10 min). Mice in all experimental
groups were injected with 100 µL PBS (pH 7.4) containing HTVMz (10 mg
kg^−1). All experimental mice received a single treatment session.
Tumor volume and body weight of mice were measured every 2 days.
Finally, mice were executed after 13 days and tumor tissue was excised
for Ki67, H&E, and TUNEL analysis. Immunofluorescence staining was
performed to assess the GSH content in tumor tissues, which served as
an indicator of ferroptosis in vivo. Additionally, H&E staining of the
skin tissue surrounding the tumors in the HTVMz + Laser group was
conducted to evaluate the potential photothermal effects on normal
skin, providing insight into the impact of photothermal therapy on
adjacent healthy tissues. To assess the biosafety of the tissue
surrounding the tumor in the photothermal therapy group, H&E staining
of the tumor‐adjacent tissue after laser irradiation was also
performed. Organs and tumor tissue were fixed by dehydration in an
automatic dehydrator and then embedded in sections. Sliced tissue
sections were stained with standard H&E and then sections were
photographed with an Olympus VS200. Blood was obtained from the eyes of
mice for further blood biochemistry and routine blood analysis.
Evaluation of In Vivo Biocompatibility of HTVMz
Healthy Balb/C mice were used for the study. Different concentrations
of HTVMz (5 and 10 mg kg^−1) were intravenously injected into the tail
vein of the mice. The mice's body weight changes were recorded within 1
month. One‐month post‐treatment, serum samples were collected from the
mice for biochemical analysis, and tissue samples were harvested for
H&E staining to assess inflammation.
Statistical Analysis
The number of samples per group in the study was n ≥ 3. All data
analyses were ultimately expressed as means, and statistical analyses
were performed using SPSS software to analyze and compare data
differences between groups, setting significance levels (*p < 0.05, **p
< 0.01, ***p < 0.001). Inter‐group differences were assessed using
one‐way analysis of variance (ANOVA). All statistical plots were
generated using GraphPad Prism.
Conflict of Interest
The authors declare no conflict of interest.
Author Contributions
H.Z. was responsible for all experiments and the preparation of the
initial draft, serving as the first author. T.L., X.Z., L.L., and X.Z.
participated in some of the cell and animal experiments and were
responsible for statistical data analysis. Q.W. was in charge of part
of the experimental design, and X.P. was responsible for mining
transcriptome data. B.H. was responsible for experimental design. J.C.
and J.Z. were responsible for conceptualization, writing‐review,
validation, supervision, and financial support.
Supporting information
Supporting Information
[202]ADVS-12-2408998-s001.docx^ (20.5MB, docx)
Acknowledgements