Abstract
Retinal neovascularization diseases cause vision impairment due to
abnormal blood vessel growth in the retina. Current treatments,
including repeated intraocular anti–vascular endothelial growth factor
injections, are invasive and often lead to discomfort and complicated
hemorrhages. Here, we developed a noninvasive nanozyme eye drop capable
of penetrating the fundus to eliminate reactive oxygen species (ROS)
and thereby inhibit neovascularization. The nanozyme eye drops consist
of liposomes formed by fluorinated and arginine–glycine–aspartic
acid–modified phospholipids, which enhance the penetration of ocular
barriers. The encapsulated superoxide dismutase–catalase cascade
nanozyme within these liposomes allows for efficient ROS scavenging. In
vitro and in vivo studies demonstrate that these nanozyme eye drops
achieve deep retinal tissue penetration, alleviate oxidative stress,
restore mitochondrial function, and suppress aberrant insulin-like
growth factor binding protein 6 signaling, thereby inhibiting
pathological neovascularization. Enhanced ocular bioavailability and
minimal toxicity further underscore its promise as a safe and effective
noninvasive treatment for retinal neovascularization diseases.
__________________________________________________________________
Nanozyme eye drops penetrate retinal barriers, initiating a catalytic
ROS-scavenging cascade that supports vasculopathy repair.
INTRODUCTION
Retinal neovascularization diseases encompass a spectrum of ocular
conditions characterized by the pathological proliferation of blood
vessels within the retina, posing a notable threat to vision ([44]1).
This category predominantly includes diabetic retinopathy, retinopathy
of prematurity, and wet age-related macular degeneration ([45]2). The
hallmark of these disorders is the abnormal growth of fragile,
rupture-prone blood vessels in the retinal or subretinal tissues,
leading to complications such as retinal edema, hemorrhage, exudate
accumulation, and detachment, severely compromising visual acuity
([46]3). Current therapeutic approaches for retinal neovascularization
diseases primarily focus on antiangiogenic treatments, with
anti–vascular endothelial growth factor (anti-VEGF) therapies at the
forefront ([47]4). While these medications effectively curb abnormal
vessel growth and ameliorate associated symptoms such as edema and
hemorrhage, their administration involves repetitive intraocular
injections, imposing considerable financial and psychological burdens
on patients ([48]5). In addition, the potential for diminished efficacy
over time due to drug resistance remains a critical challenge,
highlighting the imperative for innovative treatment modalities.
In light of these treatment limitations, the role of oxidative stress
in the pathogenesis of retinal neovascular diseases warrants notable
attention. The retina, a metabolically intense tissue, is inherently
susceptible to oxidative injury and the resultant excessive generation
of reactive oxygen species (ROS) ([49]6). This oxidative stress can
deteriorate cellular structures, including membranes, proteins, and
DNA, and incite inflammatory and angiogenic responses ([50]7).
Traditional antioxidants, such as vitamins C and E, provide some
protective effects but are limited by their instability and
insufficient ocular bioavailability when administered orally ([51]4).
Emerging research suggests that nanozymes, a class of nanomaterials
with enzyme-like properties, may offer a robust alternative
([52]8–[53]10). Demonstrating potent antioxidant capabilities in
various biomedical applications such as oncology, gastroenterology, and
neurology, nanozymes mimic the activity of natural enzymes such as
superoxide dismutase (SOD) and catalase (CAT) to efficiently neutralize
ROS ([54]11–[55]13). Their enhanced stability, superior
bioavailability, and ability to function in complex biological settings
for extended durations, combined with their nanoscale properties that
facilitate penetration through biological barriers, position nanozymes
as a promising therapeutic strategy in the management of retinal
neovascular diseases ([56]14, [57]15).
In our previous research, our team used vitreous injections and other
invasive methods to administer nanozymes, effectively suppressing the
formation of pathological neovascularization within the eye ([58]12,
[59]16). Despite the efficacy of these methods, they are associated
with considerable discomfort and potential adverse effects, including
infections and hemorrhages. This necessitates the exploration of
alternative, less invasive drug delivery systems, such as eye drops,
which could potentially mitigate these drawbacks ([60]17). However,
transitioning to eye drop formulations presents distinct challenges due
to the unique anatomical and physiological barriers of the eye, notably
the corneal and blood-retina barriers (BRB). These barriers protect
ocular tissues from external threats but simultaneously restrict the
penetration and absorption of therapeutics, thus limiting their
efficacy in treating posterior eye diseases ([61]18). To overcome these
challenges, this project proposes the development of a liposome-based
eye drop delivery system. Liposomes have inherent cell transcytosis
capabilities that can facilitate drug penetration into the eye,
enhancing bioavailability and targeting efficacy. In addition, coating
liposomes with nanomaterials and chemically modifying them can further
enhance their ability to traverse ocular barriers ([62]19). For
instance, fluorination of liposomes increases their lipophilicity,
allowing them to more readily cross lipid bilayers of cell membranes,
thereby improving drug permeability ([63]20).
Building on this approach, we aim to develop the fluorinated and
RGD-modified ruthenium polymer nanozyme (FR-PolyRu), which involves
encapsulating the antioxidant PolyRu nanozyme within a fluoridated
liposome designed for vascular targeting. The synthesis of the liposome
shell incorporates phospholipid fluoride and arginine–glycine–aspartic
acid (RGD)–modified phospholipids to enhance barrier permeation,
enabling effective drug delivery to the retinal vascular. The core
component, PolyRu nanozyme, maintains robust SOD and CAT activities,
crucial for mitigating oxidative stress by neutralizing ROS and
generating nontoxic oxygen as a byproduct. Through this innovative
noninvasive eye drop formulation, we anticipate targeted therapeutic
effects at sites of retinal neovascularization in murine models,
thereby addressing the oxidative imbalances and inhibiting the
progression of retinal neovascular diseases ([64]Fig. 1).
Fig. 1. Schematic illustration of the synthesis process and therapeutic
effects of FR-PolyRu nanozyme.
[65]Fig. 1.
[66]Open in a new tab
The nanozyme is encapsulated in liposomes made from fluorinated and
RGD-modified phospholipids, enhancing ocular barrier penetration. The
encapsulated SOD-CAT cascade nanozyme efficiently scavenges ROS in the
retina. In vitro and in vivo studies in oxygen-induced retinopathy
(OIR) models show that the nanozyme reduces oxidative stress and
inhibits retinal neovascularization. In addition, transcriptomic
analysis reveals the suppression of the abnormal Igfbp6 signaling
pathway, contributing to the antiangiogenic effect.
RESULTS
Synthesis and characterization of FR-PolyRu nanozymes
The synthesis of Ru-based antioxidant PolyRu nanozyme began by mixing
the polymer polyvinyl pyrrolidone (PVP) with RuCl[3]. Subsequently,
DSPE-PEG-F[7], a derivative synthesized in our laboratory, was prepared
by modifying the amino group of DSPE-PEG
[1,2-distearoyl-sn-glycero-3-phosphoethanolamine-poly(ethylene glycol)]
with heptafluorobutyric anhydride, as confirmed by ^1H nuclear magnetic
resonance (NMR) (fig. S1A) and ^19F NMR spectra (fig. S1B). According
to the ^19F NMR spectra, the characteristic peaks of fluorine atoms
were observed in DSPE-PEG-F[7], indicating that DSPE-PEG-NH[2] was
successfully modified with heptafluorobutyric acid. Next, the PolyRu
nanozymes were encapsulated within liposomes by mixing them with
cholesterol, DSPE-PEG, DSPE-PEG-F[7], and DSPE-PEG-RGD, forming
FR-PolyRu nanozymes ([67]Fig. 2A). In the Fourier transform infrared
spectrum (fig. S1C), distinct characteristic absorption peaks
corresponding to the nanozyme and liposome in FR-PolyRu were observed,
confirming the successful encapsulation of liposomes on the outer layer
of the nanozyme. Transmission electron microscopy (TEM) images revealed
that the PolyRu nanozymes have a spherical structure with a diameter of
~50 nm ([68]Fig. 2B). Negative-stained TEM images further confirmed
that the liposomes effectively coated the surface of the PolyRu
nanozymes ([69]Fig. 2C). Meanwhile, atomic force microscopy (AFM)
indicated that the FR-PolyRu nanozyme exhibited irregular spherical
structures, with diameters and heights measuring around 50 and 7 nm,
respectively ([70]Fig. 2, D and E). Clarity is crucial for eye drop
applications, and as shown in [71]Fig. 2F, the nanozyme solution
exhibited a high degree of clarity.
Fig. 2. The physicochemical characterization of FR-PolyRu nanozymes.
[72]Fig. 2.
[73]Open in a new tab
(A) Schematic illustration of FR-PolyRu nanozymes. (B) TEM
characterization results of PolyRu. Scale bar, 100 nm. (C) TEM
characterization results of FR-PolyRu nanozymes. Scale bar, 200 nm. (D
and E) AFM topographical images of FR-PolyRu nanozymes with a 1 μm–by–1
μm scanning area. Three-dimensional rendering of the topographic image
shown in (D), and the corresponding height profile along the nanozymes
in (E). Scale bar, 200 nm. (F) Images of different concentrations of
FR-PolyRu nanozymes dispersed in deionized water. (G) Elemental
analysis of FR-PolyRu nanozymes by x-ray photoelectron spectroscopy
(XPS). a.u., arbitrary units. (H) High-resolution XPS spectra of C
1s + Ru 3d to compare FR-PolyRu nanozymes. (I) Size distribution of
RuO[2], PolyRu, FR-PolyRu nanozymes by DLS (n = 3). (J) The zeta
potentials of RuO[2], PolyRu, and FR-PolyRu nanozymes by dynamic light
scattering (DLS; n = 3). (K) Pictures of RuO[2], PolyRu, and FR-PolyRu
nanozymes dispersed in deionized water at days 0, 7, 14, and 30.
The physicochemical properties of the FR-PolyRu nanozyme eye drops,
including osmotic pressure, torque, viscosity, shear stress, and shear
rate, were systematically evaluated, as these factors are critical for
determining the efficacy, stability, and user experience of the eye
drops. Table S1 shows that the properties of FR-PolyRu nanozyme eye
drops closely align with those of the commercial Bausch + Lomb
formulation, indicating their suitability for ophthalmic use. The
structure of FR-PolyRu nanozyme was further analyzed using x-ray
photoelectron spectroscopy (XPS). As illustrated in [74]Fig. 2G, the
XPS survey spectrum confirmed the presence of F, O, C, N, and Ru,
indicating the successful incorporation of Ru into the material.
High-resolution Ru 3d XPS spectra ([75]Fig. 2H) revealed binding
energies for Ru 3d 5/2 and Ru 3d 3/2 at 280.47 and 284.64 eV,
corresponding to metallic Ru. Additional peaks at 281.11 and 285.20 eV
were attributed to Ru^4+ 3d 5/2 and Ru^4+ 3d 3/2. These binding
energies suggest that electronic transfer occurs between Ru and the
substrate, resulting in an electron-rich state for Ru, which exhibits
strong reducing properties. This electron-rich state makes it difficult
for adsorbed oxidizing substances to stabilize, thereby balancing the
adsorption and desorption of reaction intermediates and facilitating
reaction progress.
Stability was a critical focus during the development of the eye drops,
as maintaining the nanozyme formulations’ stability during storage is
essential for preserving efficacy. To assess this, we monitored the
particle size, zeta potential, and dispersibility of RuO[2], PolyRu,
and FR-PolyRu nanozymes over 30 days. As shown in [76]Fig. 2 (I and J),
the initial particle sizes of RuO[2], PolyRu, and FR-PolyRu nanozymes
were ~50 nm. Both RuO[2] and FR-PolyRu nanozymes exhibited positive
zeta potentials, which can enhance their retention within the eye and
improve penetration due to the negatively charged ocular surface. The
positive zeta potential of FR-PolyRu is attributed to the composition
of its lipid shell, where the DSPE-PEG-NH[2] contributes to a positive
surface charge despite the presence of negatively charged PolyRu
inside. Over the 30-day period, the particle sizes of RuO[2] and PolyRu
nanozymes increased substantially, reaching around 200 and 100 nm,
respectively, accompanied by a decrease in dispersibility. In contrast,
FR-PolyRu nanozymes maintained a stable particle size and
dispersibility throughout the study, highlighting their superior
stability, which is primarily attributed to the liposomal encapsulation
and the complementary functions of fluorination and RGD modification
that establish a dual stabilization mechanism. Fluorinated lipids
(DSPE-PEG-F[7]) enhance lipid bilayer stability by increasing membrane
rigidity, reducing lipid mobility, and preventing water penetration,
which could otherwise lead to nanoparticle swelling or destabilization,
while also lowering interfacial free energy to prevent lipid fusion and
aggregation. Meanwhile, RGD modification ensures hydrophilic stability
by enhancing steric hindrance through the PEGylated RGD groups, which
prevent close particle interactions that could lead to aggregation.
Moreover, the protective role of the liposomes in shielding the
nanozymes from aggregation further contributes to FR-PolyRu’s
stability, and the grafting of RGD and fluorination enhances the
self-assembly capacity of the liposomes, reinforcing their structural
integrity ([77]Fig. 2K and figs. S2 and S3A). In addition, the zeta
potential of all three nanozymes remained relatively stable (fig. S4).
In addition, the particle size and zeta potential of different batches
of FR-PolyRu eye drops were essentially the same (fig. S5, A and B).
These findings indicate that FR-PolyRu nanozymes provide superior
stability for eye drop applications, particularly in terms of particle
size, zeta potential, and dispersibility.
SOD/CAT cascade catalytic ability of FR-PolyRu nanozymes
The SOD/CAT cascade catalytic ability of FR-PolyRu nanozymes
effectively catalyzes the conversion of cytotoxic superoxide anion
(O[2]^•–) into oxygen (O[2]), thereby reducing ROS levels ([78]Fig.
3A). Compared with the RuO[2] control, PolyRu and FR-PolyRu nanozymes
showed markedly higher SOD-like activity, and liposome encapsulation
did not affect the O[2]^•– scavenging ability of PolyRu nanozymes
([79]Fig. 3B and fig. S6). In addition, measurements of H[2]O[2]
decomposition and O[2] production confirmed that FR-PolyRu nanozyme
exhibits excellent CAT-like activity ([80]Fig. 3C and fig. S7), which
remains effective even at low H[2]O[2] concentrations (fig. S8).
FR-PolyRu nanozymes did not display peroxidase (POD)– or oxidase-like
activity at physiological pH 7.4, demonstrating their selective
ROS-scavenging efficiency (fig. S9). Furthermore, electron spin
resonance (ESR) analysis supported the ROS-scavenging capabilities of
FR-PolyRu nanozymes, showing a notable reduction in O[2]^•–, ^1O[2],
•OH, and •ON ([81]Fig. 3, D to F, and fig. S10).
2,2-diphenyl-1-picrylhydrazyl (DPPH) and
2,2′-azino-bis-3-ethylbenzthiazoline-6-sulphonic acid (ABTS) free
radical inhibition analysis further validated the superior free radical
scavenging ability of FR-PolyRu nanozymes compared to the RuO[2]
control ([82]Fig. 3, G and H). Physical entrapment of nanozymes in
liposomes often faces challenges related to batch-to-batch variability.
In addition to the stability verification mentioned above, we also
assessed the CAT and SOD enzyme activities in three batches of
FR-PolyRu eye drops. The results showed that the different batches
exhibited similar and high enzyme activities (fig. S5, C and D). In
addition, when artificial tears were used as a buffer, FR-PolyRu
maintained high SOD and CAT enzyme activities and remained stable for
up to 30 days (fig. S3, B and C).
Fig. 3. Detection of enzyme-like activity and its theoretical analysis of
FR-PolyRu nanozymes.
[83]Fig. 3.
[84]Open in a new tab
(A) Schematic illustration of the multienzyme-like activities of the
FR-PolyRu nanozymes. (B) The SOD-like activities of RuO[2], PolyRu, and
FR-PolyRu nanozymes detected by the xanthine oxidase/cytochrome c
system at pH 7.4 (n = 3). (C) The CAT-like activities of nanozymes
detected by recording representative dissolved oxygen produced in the
H[2]O[2] solutions containing RuO[2], PolyRu, and FR-PolyRu nanozymes
[Ru equivalent (0.4 μg/ml)] at pH 7.4 (n = 3). (D) FR-PolyRu nanozymes
reduce the generation of superoxide radical demonstrated by ESR
spectroscopy. (E) FR-PolyRu nanozymes reduce the generation of singlet
oxygen demonstrated by ESR spectroscopy. (F) FR-PolyRu nanozymes reduce
the generation of hydroxyl radical demonstrated by ESR spectroscopy.
(G) ABTS radical scavenging ratio of RuO[2], PolyRu, and FR-PolyRu
nanozymes (n = 3). (H) DPPH radical scavenging ratio of RuO[2], PolyRu,
and FR-PolyRu nanozymes (n = 3). (I) The side and top view of the
structural model for PolyRu nanozyme. (J and K) The proposed mechanisms
responsible for the SOD- (J) and CAT-like (K) activities of PolyRu
nanozyme, respectively. (L and M) The calculated energy profiles in
electron volts regarding the SOD- (L) and CAT-like (M) mechanisms of
PolyRu nanozyme, respectively. In addition, the chemical constituents
for the stationary points were listed.
To elucidate the cascade SOD- and CAT-like activities of PolyRu
nanozymes, we performed density functional theory (DFT) calculations.
The dynamic equilibrium structure of PolyRu, based on ab initio
molecular dynamics (AIMD) simulations with a single PVP molecule
adsorbed on its surface, served as the initial model ([85]Fig. 3I) for
investigating the catalytic mechanisms. [86]Figure 3 (J and K) presents
proposed mechanisms underlying the SOD- and CAT-like activities of
PolyRu nanozymes, as informed by prior studies ([87]21–[88]23).
Corresponding Gibbs free energy profiles for these mechanisms are shown
in [89]Fig. 3 (L and M), with the structures of intermediate species
detailed in fig. S11. The Gibbs free energy profiles demonstrate that
PolyRu nanozymes support both SOD- and CAT-like enzyme cascade
reactions, ultimately catalyzing the conversion of O[2]^•– (or •OOH
under acidic conditions) into O[2]. As a highly active reducing metal,
Ru enables the PolyRu nanozyme to effectively adsorb key reactants for
both SOD and CAT reactions, specifically •OOH and H[2]O[2], on its
surface to enable subsequent chemical transformations. For instance, in
the second step illustrated in [90]Fig. 3L, •OOH readily adsorbs onto
the PolyRu surface, capturing hydrogen atoms from H[2]O* (where *
denotes the adsorbed state) to produce H[2]O[2]*, a spontaneous process
with a reaction energy of −1.37 eV. Similarly, in the first step shown
in [91]Fig. 3M, H[2]O[2] undergoes direct decomposition into O* and
H[2]O* upon adsorption on the PolyRu surface, releasing energy of −4.24
eV.
The rate-limiting steps in both SOD and CAT catalytic cycles involve
the release of oxygen, with a desorption energy of 1.63 eV, indicating
a strong affinity for active oxygen on the PolyRu surface. However,
this does not impede the reaction cycle, as competitive adsorption of
•OOH and H[2]O[2] is sufficient to displace surface-bound O[2] for the
next cycle, with adsorption energies reaching −1.62 eV (for
coadsorption of •OOH and H[2]O) and −4.24 eV. These results provide
valuable insights into the molecular mechanisms underlying the cascade
SOD- and CAT-like activities of PolyRu nanozymes.
The antioxidant and oxidative damage protection ability of FR-PolyRu
nanozymes toward human retinal endothelial cells and human umbilical vein
endothelial cells
To assess the antioxidant capacity of FR-PolyRu nanozymes in cellular
systems, we used H[2]O[2]- or O[2]^•–-induced an oxidative damage model
in human umbilical vein endothelial cells (HUVECs) and human retinal
endothelial cells (HRECs). Subsequently, the capacity of pretreated
FR-PolyRu nanozymes to attenuate oxidative stress–induced cellular
injury was assessed ([92]Fig. 4A). To exclude the potential influence
of nanomaterial toxicity on the evaluation of their protective
capacity, we initially incubated cells with different concentrations of
RuO[2], PolyRu, and FR-PolyRu nanozymes. In addition, cell viability
was subsequently determined using the Cell Counting Kit-8 (CCK-8)
assay. The results demonstrated that with increasing concentrations,
neither PolyRu nor FR-PolyRu nanozymes markedly affected cell viability
([93]Fig. 4B and fig. S12A).
Fig. 4. Scavenging of intracellular ROS and reversal of oxidative damage by
FR-PolyRu in vitro.
[94]Fig. 4.
[95]Open in a new tab
(A) Schematic diagram of the in vitro antioxidant experiment of
FR-PolyRu nanozymes. (B) Cell viability of HRECs incubated with
different concentrations of RuO[2], PolyRu, and FR-PolyRu nanozymes for
24 hours via CCK-8 assays (n = 3). (C) Cellular uptake of
Cy5.5–FR-PolyRu was observed through confocal laser scanning
microscopy. Scale bar, 20 μm. (D) Cell viability of HRECs incubated
with 200.0 μM H[2]O[2] and different concentrations of RuO[2], PolyRu,
and FR-PolyRu nanozymes (n = 3). (E) Representative immunofluorescence
of RuO[2], PolyRu, and FR-PolyRu nanozymes–treated HREC for tubulin
(red) and 4′,6-diamidino-2-phenylindole (DAPI; blue) stain. Scale bar,
50 μm. (F and G) Flow cytometry tests of ROS levels in HRECs via
2′,7′-dichlorofluorescein diacetate (DCFH-DA) staining, accompanied by
statistical data [n = 3; one-way analysis of variance (ANOVA) with
Tukey’s multiple comparisons test]. (H and I) Representative
immunofluorescence image and quantitative data of RuO[2], PolyRu, or
FR-PolyRu nanozyme–treated HRECs for DCFH-DA (green) and DAPI (blue)
stain. Scale bar, 200 μm (n = 3; one-way ANOVA with Tukey’s multiple
comparisons test). (J and K) Flow cytometry tests of RuO[2], PolyRu, or
FR-PolyRu nanozyme–treated HREC stained with mitochondrial membrane
potential (MMP) probe JC-1, accompanied by statistical data (n = 3;
one-way ANOVA with Tukey’s multiple comparisons test). (L)
Mitochondrion-specific ROS scavenging activity of RuO[2], PolyRu, or
FR-PolyRu nanozymes. Scale bar, 50 μm. (M) Cellular adenosine
5′-triphosphate (ATP) level in HRECs with different treatments (n = 3;
one-way ANOVA with Tukey’s multiple comparisons test). Data were
presented as mean ± SD; n.s., not significant.
We further examined the cellular uptake efficiency of FR-PolyRu
nanozymes. The intracellular accumulation of FR-PolyRu nanozymes
gradually increased over time, reaching a maximum at 8 hours, which
suggests effective cellular internalization ([96]Fig. 4C and fig.
S12B). Furthermore, we observed that HRECs and HUVECs pretreated with
PolyRu and FR-PolyRu nanozymes effectively resisted H[2]O[2]- or
O[2]^•–-induced reduction in cell viability ([97]Fig. 4D and figs. S12,
C and D, S13, and S14, A and B) and cytoskeletal disruption ([98]Fig.
4E and fig. S12E). Excessive ROS react with unsaturated fatty acids in
cell membranes, triggering lipid peroxidation. Malondialdehyde (MDA), a
key product of lipid peroxidation, is commonly used as an important
marker of oxidative damage. To further evaluate the ability of the
nanozyme to inhibit lipid peroxidation, we conducted additional tests.
As shown in fig. S15, H[2]O[2] stimulation resulted in significant MDA
accumulation in HRECs. However, both PolyRu and FR-PolyRu nanozymes
effectively reversed this abnormal accumulation. This implies that
PolyRu nanozymes may mitigate oxidative cellular damage by neutralizing
ROS through their enzyme-like activity.
We further assessed the impact of PolyRu nanozymes on intracellular ROS
levels, intracellular ROS were labeled using 2′,7′-dichlorofluorescein
diacetate (DCFH-DA) and dihydroethidium (DHE), and flow cytometry
revealed a significant increase in ROS levels following stimulation
with H[2]O[2] ([99]Fig. 4, F and G, and fig. S12, F and G) or O[2]^•−
(fig. S14, C and D). However, ROS levels in cells pretreated with
PolyRu and FR-PolyRu nanozymes were significantly reduced compared to
the H[2]O[2]- or O[2]^•−-treated group, closely resembling the control
group. Confocal imaging of intracellular DCFH-DA fluorescence staining
([100]Fig. 4, H and I, and fig. S12, H and I) corroborated these
results. These findings suggest that PolyRu and FR-PolyRu nanozymes
alleviate cellular oxidative damage by lowering ROS levels.
An abnormal increase in intracellular ROS is frequently linked to
mitochondrial dysfunction. Compromised mitochondrial function typically
results in elevated ROS levels, which subsequently lead to the
degradation of intracellular lipids, proteins, and DNA, further
aggravating mitochondrial damage. To determine whether PolyRu and
FR-PolyRu nanozymes can mitigate mitochondrial impairment, we performed
additional assessments. Using the
5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolylcarbocyanine
iodide (JC-1) probe to measure mitochondrial membrane potential (MMP)
under various conditions, we found that HRECs exposed to H[2]O[2]
exhibited predominantly green fluorescence, indicating low MMP due to
JC-1 monomers ([101]Fig. 4, J and K, and fig. S16). In contrast, cells
treated with PolyRu and FR-PolyRu nanozymes maintained high MMP
following H[2]O[2] stimulation, as indicated by the red fluorescence of
JC-1 aggregates. In addition, using the MitoSOX probe to quantify
mitochondrial ROS, we observed a significant increase in mitochondrial
ROS levels in HRECs exposed to H[2]O[2]. However, cells treated with
PolyRu and FR-PolyRu nanozymes maintained near-normal mitochondrial ROS
levels ([102]Fig. 4, L and M). These findings suggest that PolyRu and
FR-PolyRu nanozymes attenuate mitochondrial oxidative damage by
reducing ROS within mitochondria, thereby preserving mitochondrial
membrane potential.
Tissue penetration of FR-PolyRu nanozymes
The penetration capacity of eye drops within the eyeball is crucial for
determining their concentration and efficacy at the fundus. To evaluate
this, we subsequently assessed the penetration ability of FR-PolyRu eye
drops in the mouse eyeball ([103]Fig. 5A). In a transwell experiment,
we densely inoculated human corneal epithelial cells (HCECs) in the
upper chamber and HRECs in the lower chamber to evaluate the in vitro
penetration of indocyanine green (ICG)–labeled FR-PolyRu nanozymes, as
indicated by their ability to traverse the upper chamber. As
demonstrated in [104]Fig. 5 (B and C), F-PolyRu and FR-PolyRu nanozymes
successfully penetrate the HCECs in the upper chamber and migrate into
the lower chamber, with the extent of penetration progressively
increasing over time. In contrast, PolyRu and R-PolyRu nanozymes
exhibit no such penetrative capability. This ability is strongly
correlated with fluorination modification. Subsequently, we evaluated
the ability of ICG-labeled liposomes prepared with DSPE-PEG-F[5],
DSPE-PEG-F[7], and DSPE-PEG-F[9] to penetrate HCECs. As shown in fig.
S17, liposomes modified with DSPE-PEG-F[7] and DSPE-PEG-F[9] exhibited
significantly enhanced transcellular permeability compared to those
modified with DSPE-PEG-F[5]. Notably, the permeability performance of
DSPE-PEG-F[7] and DSPE-PEG-F[9] liposomes was comparable. On the basis
of literature evidence, experimental validation, and synthetic
feasibility ([105]24), DSPE-PEG-F[7] was selected for formulation
development, as it provided optimal permeability comparable to
DSPE-PEG-F[9] while offering superior synthetic accessibility and cost
effectiveness.
Fig. 5. Penetration of ocular barriers by FR-PolyRu.
[106]Fig. 5.
[107]Open in a new tab
(A) Schematic illustration of FR-PolyRu nanozymes penetrating the
cornea to the fundus. (B) In vitro fluorescence imaging of ICG-labeled
nanozymes’ ability to traverse the cornea was verified by transwell
assay. h, hours. (C) Quantitative analysis of average fluorescence
intensity in the lower compartment at different time intervals when
ICG-labeled nanozymes were added to the upper compartment (n = 3;
one-way ANOVA with Tukey’s multiple comparisons test). (D and E) The
transmembrane system structural model. (E) The force 𝐹 variations
during the translocation process were analyzed as a function of the Z
coordinate, ranging from −1.5 to 1.5 nm. (F and G) Photoacoustic
imaging of intraocular nanozyme distribution at different time
intervals after different nanozymes are dropped into mouse eyeballs,
and their corresponding mean photoacoustic (PA) signal intensity
quantitative analysis (n = 3; two-way ANOVA with Dunnett’s multiple
comparisons test). PAAvr.Thresh, photoacoustic average threshold. (H
and I) Immunofluorescence staining image of Cy5.5 signal at the fundus
site treated with Free-Cy5.5 or Cy5.5–FR-PolyRu nanozymes. Cy5.5 signal
(red) and DAPI (blue). Scale bar, 200 μm. In addition, the
corresponding fluorescence intensity quantitative analysis (n = 3;
two-way ANOVA with Šidák’s multiple comparisons test). MFI, mean
fluorescence intensity. Data were presented as mean ± SD; n.s., not
significant.
To further investigate the mechanisms underlying this enhanced
permeability, we performed steered molecular dynamics (SMD) simulations
to evaluate the membrane permeability of DSPE-PEG-F[7], a fluorinated
lipid designed to enhance nanoparticle stability and facilitate
transmembrane transport, particularly in overcoming electrostatic
barriers such as the tear film, sclera, and vitreous body, which hinder
the diffusion of positively charged substances into the posterior
segment of the eye. The results ([108]Fig. 5, D and E) illustrate force
variations during the translocation process of DSPE-PEG-F[7],
DSPE-PEG-NH[2], and DSPE-PEG-RGD across a phospholipid bilayer. The
maximum force required for DSPE-PEG-F[7] to traverse the membrane was
substantially lower than that of DSPE-PEG-NH[2] (805.7 kJ/mol versus
915.6 kJ/mol) and much lower than that of DSPE-PEG-RGD (1981.5 kJ/mol).
These findings indicate that fluorination substantially enhances
membrane permeability, lowering the energy barrier required for
transmembrane transport. On the basis of these findings, we propose
that the penetration of FR-PolyRu through the ocular barrier is
primarily mediated by transcytosis. To evaluate the transcytosis
capability of different lipid-modified formulations, we performed a
coculture assay using HCECs ([109]25). Specifically, HCECs pretreated
with unmodified Lipo-PolyRu, fluorinated F-PolyRu, or dual-modified
FR-PolyRu were seeded onto glass slides and designated as donor cells.
Separate slides were seeded with untreated HCECs as recipient cells.
The donor and recipient slides were placed in direct contact, and flow
cytometry analysis was conducted to assess nanoparticle transfer. The
results revealed that the ratio of signal intensity in recipient cells
to that in donor cells was significantly higher for the F-PolyRu and
FR-PolyRu groups compared to the unmodified Lipo-PolyRu group (fig.
S18). These results suggest that fluorinated modification enhances the
transcytosis of nanoparticles, thereby promoting their
barrier-penetrating capacity.
To determine whether this increased in vitro permeability translates to
enhanced ocular penetration in vivo, we next evaluated the tissue
penetration of FR-PolyRu nanozymes in mouse eyeballs. Through
photoacoustic imaging, we observed that PolyRu, R-PolyRu, and
DSPE-PEG-NH[2]-PolyRu nanozymes primarily accumulated in the cornea,
demonstrating limited permeability. In contrast, F-PolyRu and FR-PolyRu
nanozymes exhibited robust tissue penetration, with significant
enrichment in the fundus ~4 hours postadministration ([110]Fig. 5, F
and G, and figs. S19 to S21). Besides, frozen section staining of mouse
eyeballs revealed that FR-PolyRu nanozymes could be effectively
enriched in the retinal region of the fundus ([111]Fig. 5, H and I).
To further confirm that FR-PolyRu nanozymes penetrate the fundus as
intact nanoparticles rather than as dissociated ions or oligomers, we
conducted biological tissue TEM imaging (fig. S22). TEM results show
FR-PolyRu nanozymes present in the posterior segment, providing direct
evidence of nanoparticle penetration into the fundus. In addition, we
performed a dialysis stability test in an artificial tear
microenvironment to assess whether ICG dissociates from FR-PolyRu
during penetration. ICG–FR-PolyRu was placed in artificial tear fluid
within a dialysis bag, and ICG fluorescence signal and Ru element
content were monitored over 24 hours (fig. S23). The results
demonstrated no significant changes in ICG fluorescence intensity or Ru
content, confirming that ICG remains stably associated with FR-PolyRu
and does not dissociate in the ocular microenvironment. Furthermore,
DPPH and ABTS free radical scavenging assays confirmed that neither
RuCl[3] nor PVP alone has antioxidant activity (fig. S24), reinforcing
that the therapeutic effect observed is due to intact FR-PolyRu
nanozymes penetrating the fundus rather than dissociated components.
The fluorinated lipid modification plays a critical role in both
enhancing membrane permeability and prolonging nanoparticle retention
in ocular tissues. As observed in our in vivo studies, FR-PolyRu
nanozymes exhibited prolonged accumulation in the fundus compared to
nonfluorinated counterparts ([112]Fig. 5, F and G). This extended
retention can be explained by two key mechanisms. First, fluorination
improves transmembrane transport efficiency by modifying the lipid
bilayer structure. The fluorinated hydrophobic tail of DSPE-PEG-F[7]
enhances membrane rigidity while maintaining structural flexibility,
facilitating efficient penetration across phospholipid barriers. In
addition, fluorine atoms lower surface energy, reducing nanoparticle
aggregation and enabling efficient passage through ocular electrostatic
barriers, such as the tear film and vitreous body, which typically
hinder nanoparticle diffusion into the posterior segment.
Second, fluorination contributes to prolonged retention in retinal
tissues by modifying membrane interactions and clearance dynamics.
Unlike conventional lipophilic molecules, which are rapidly cleared
after crossing the BRB, fluorinated lipids exhibit stronger
interactions with the lipid-rich ocular environment, reducing passive
diffusion–driven clearance and leading to sustained accumulation in
retinal tissues. Furthermore, fluorination may alter nanoparticle
recognition by clearance pathways, thereby potentially reducing rapid
elimination and further supporting prolonged retention ([113]20,
[114]26, [115]27).
Beyond passive diffusion and lipid-assisted penetration, FR-PolyRu uses
an additional, active mechanism to enhance tissue penetration. The
nanozyme core catalyzes the decomposition of H[2]O[2] to generate O[2],
which serves as an active driving force for deeper tissue diffusion.
The locally generated oxygen reduces diffusion resistance and promotes
intraocular penetration and nanoparticle transport.
While DSPE-PEG-F[7] primarily enhances membrane penetration,
DSPE-PEG-RGD contributes to nanoparticle stability by forming a
hydration layer, preventing aggregation, and facilitating
receptor-mediated uptake. The DSPE-PEG-RGD shell stabilizes dispersion
in biological fluids, while DSPE-PEG-F[7] enhances membrane
interactions, ensuring both colloidal stability and efficient
biological transport. Furthermore, the nanozyme-catalyzed oxygen
release introduces an active mechanism that complements the passive
diffusion–enhancing properties of fluorinated lipids, forming a
synergistic strategy for overcoming ocular barriers and enabling
targeted delivery to the posterior segment. These findings suggest that
FR-PolyRu nanozyme eye drops not only have excellent tissue penetration
capabilities but also benefit from prolonged residency in the posterior
segment, facilitating more effective therapeutic delivery to the
retinal region.
Efficacy of FR-PolyRu nanozyme in inhibiting retinal neovascularization in
oxygen-induced retinopathy mice
PolyRu and FR-PolyRu nanozymes exhibit antioxidant properties in
cellular models and can effectively mitigate cellular oxidative damage.
Therefore, we initially assessed the inhibitory effect of intravitreal
injection of PolyRu nanozymes on abnormal retinal blood vessel
formation. As shown in fig. S25A, exposing a 7-day-old mouse to a
high-concentration oxygen environment (75%) for 5 days leads to
vascular degeneration, forming a central avascular zone. Upon
transferring the mouse to a normal 21% oxygen environment at postnatal
day 12 (P12), retinal neovascularization develops as a result of
relative hypoxia. We performed vitreous injections of PolyRu nanozymes
in P12 mice (fig. S25B) to evaluate their inhibitory effect on
neovascularization in the fundus. As shown in fig. S25 (C to E),
treatment with PolyRu nanozymes significantly reduced retinal
neovascularization and the central avascular area in oxygen-induced
retinopathy (OIR) mice, suggesting that PolyRu nanozymes may facilitate
the repair of retinal blood vessels.
Given the robust eyeball penetration capacity of F-PolyRu and FR-PolyRu
nanozymes, we examined the effects of FR-PolyRu nanozyme eye drops
administered at P12 on abnormal retinal blood vessels in mice
([116]Fig. 6, A and B). As illustrated in [117]Fig. 6 (C to E) and fig.
S26, both F-PolyRu and FR-PolyRu nanozyme eye drops effectively
inhibited abnormal retinal blood vessel formation and accelerated the
repair of the central avascular zone, whereas PolyRu eye drops did not
show such effects, likely due to insufficient penetration ability. The
effect of FR-PolyRu nanozyme eye drops is significantly superior to
that of F-PolyRu nanozyme eye drops, while their permeability is
comparable. We hypothesize that this difference may be attributed to
the enhanced vascular targeting capability conferred by the RGD
modification. Coincubation of the two nanozymes with HRECs, followed by
flow cytometry analysis, revealed that RGD-modified FR-PolyRu nanozyme
eye drops were more efficiently taken up by endothelial cells (fig.
S27). This suggests that FR-PolyRu nanozyme eye drops can more
effectively penetrate and target abnormal blood vessels, thereby
repairing defective retinal vasculature. Meanwhile, we assessed ROS
levels in retinal frozen sections from OIR mice using DHE probe
staining. The results revealed a markedly elevated ROS signal in the
retinas of OIR mice compared to normal controls, whereas treatment with
FR-PolyRu eye drops markedly reduced ROS accumulation (fig. S28). In
addition, we performed terminal deoxynucleotidyl transferase–mediated
deoxyuridine triphosphate nick end labeling (TUNEL) staining on retinal
sections from OIR mice treated with FR-PolyRu nanozyme eye drops. The
results demonstrated a significantly higher level of retinal cell
apoptosis in OIR mice compared to normal mice, indicating that
treatment with FR-PolyRu nanozyme eye drops effectively reversed
retinal cell apoptosis ([118]Fig. 6, F and G). These findings suggest
that FR-PolyRu nanozymes, through their tissue penetration, vascular
targeting, and antioxidant properties, can noninvasively repair retinal
vascular abnormalities via eye drop administration.
Fig. 6. Reversal of retinal vasculopathy in OIR mice by FR-PolyRu.
[119]Fig. 6.
[120]Open in a new tab
(A) Schematic diagram of the OIR mouse model. (B) Schematic diagram of
the in vivo antioxidant treatment experiment in OIR mice. (C)
Representative images of different groups of processed whole-mount
retinas are displayed. Purple: avascular area, and red: neovascular
area. Scale bar, 1 mm. (D and E) Quantification of the percentage of
avascular (D) and neovascular area (E) (n = 8; one-way ANOVA with
Tukey’s multiple comparisons test). (F) Immunofluorescence staining
images of TUNEL of fundus parts after different treatments; TUNEL
(green) and DAPI (blue). Scale bars, 100 μm. (G) Quantification of
apoptosis following TUNEL staining (n = 3; one-way ANOVA with Tukey’s
multiple comparisons test). Data were presented as mean ± SD; n.s., not
significant.
To explore the role of retinal neovascularization at the
transcriptional level, we conducted a comprehensive transcriptomic
analysis of treated mouse retinas using mRNA sequencing (mRNA-seq). The
experimental groups consisted of untreated retinas (normal), OIR
treated with phosphate-buffered saline (PBS) (OIR-PBS), and OIR treated
with FR-PolyRu eye drops (OIR–FR-PolyRu). Principal components analysis
(PCA) revealed greater transcriptional similarity between the
FR-PolyRu–treated and normal groups compared to the untreated OIR-PBS
group, suggesting that FR-PolyRu may exert protective effects at the
mRNA transcription level ([121]Fig. 7A). Pathway enrichment analysis
using the Kyoto Encyclopedia of Genes and Genomes (KEGG) identified
significant involvement of the phosphatidylinositol 3-kinase
(PI3K)–Akt, hypoxia-inducible factor–1 (HIF-1), and mitogen-activated
protein kinase (MAPK) signaling pathways in the aberrant
neovascularization observed in the OIR model ([122]Fig. 7B). Gene set
enrichment analysis (GSEA) further indicated that OIR treatment led to
positive regulation of apoptosis, HIF signaling, and PI3K-AKT pathways,
while treatment with FR-PolyRu exhibited a negative regulatory effect
on these pathways ([123]Fig. 7, C and D). Given the substantial impact
on the PI3K-AKT pathway, we conducted further analyses of genes
enriched in this pathway. Heatmaps of differentially expressed genes
showed a marked up-regulation of downstream angiogenesis-associated
genes, including Flt4, Vegfa, Col1a2, Thbs1, and Pdgfd, in the OIR
group. Treatment with FR-PolyRu attenuated the abnormal expression of
these genes, indicating its efficacy in regulating
neovascularization-related gene expression ([124]Fig. 7E and fig. S29).
Fig. 7. FR-PolyRu regulates the dysregulated IGFBP6/PI3K/ERK/AKT signaling
pathway in the retina of OIR mice.
[125]Fig. 7.
[126]Open in a new tab
(A) PCA analysis for the samples in different groups. (B) KEGG-enriched
pathways in the OIR-PBS group compared to the normal group. The
ordinate and abscissa represent the path name and the rich factor,
respectively. The size of the dots indicates the number of
differentially expressed genes (DEGs) in this pathway, and the color of
the dots corresponds to different q values. ECM, extracellular matrix.
(C) GSEA was performed to analyze clusters of genes that belong to
apoptosis, HIF-1 signaling pathway, and PI3K-AKT signaling pathway
(normal versus OIR-PBS). (D) GSEA was performed to analyze clusters of
genes that belong to apoptosis, HIF-1 signaling pathway, and PI3K-AKT
signaling pathway (OIR-PBS versus OIR–FR-PolyRu). (E) Heatmap depicting
representative DEGs of different groups (red, up-regulation; blue,
down-regulation); log[2] fold change ≥ 1, and q < 0.05. (F)
Quantification of Igfbp6 mRNA expression levels (n = 3; one-way ANOVA
with Tukey’s multiple comparisons test). (G) Protein levels of Igfbp-6,
ERK, phospho-ERK (p-ERK), PI3K, phospho-PI3K (p-PI3K), AKT, and
phospho-AKT (p-AKT) in mouse retina determined by Western blotting;
β-actin was used as the loading control. (H) Quantitative statistics
for the protein of Igfbp-6, the ratio of p-ERK to total-ERK, p-PI3K to
total-PI3K, and p-AKT to total-AKT (n = 3; one-way ANOVA with Tukey’s
multiple comparisons test). (I) Schematic diagram of FR-PolyRu
nanozymes alleviating oxidative stress and reducing neovascularization.
Data were presented as mean ± SD; n.s., not significant.
In addition to the aforementioned genes, heatmap analysis revealed
substantial alterations in the expression of insulin-like growth factor
binding protein 6 (Igfbp6), which modulates the PI3K-AKT signaling
pathway ([127]Fig. 7E) ([128]28). This regulation can induce metabolic
changes and oxidative bursts, directly linking Igfbp6 to the previously
mentioned mitochondrial dysfunction ([129]29, [130]30). We further
validated the expression of Igfbp6 at transcriptional and protein
levels, which confirmed its substantial up-regulation in the OIR group,
with expression levels decreasing to near-normal levels in the
FR-PolyRu–treated group ([131]Fig. 7, F to H). We further subjected
HRECs to H[2]O[2]-induced oxidative stress, which resulted in a
significant up-regulation of Igfbp6 mRNA and protein levels. This
abnormal increase was effectively reversed by FR-PolyRu treatment,
suggesting that FR-PolyRu may alleviate oxidative stress by modulating
Igfbp6 signaling (fig. S30). To explore the downstream impact on the
PI3K-AKT signaling pathway, we conducted protein-level validation. The
results demonstrated that abnormal phosphorylation of extracellular
signal–regulated kinase (ERK), PI3K, and AKT was significantly reduced
following FR-PolyRu treatment, which is consistent with the observed
amelioration of abnormal retinal neovascularization ([132]Fig. 7, G and
H, and fig. S31). These findings suggest that FR-PolyRu effectively
modulates both upstream and downstream elements of the PI3K-AKT
pathway, thereby contributing to the restoration of retinal homeostasis
([133]Fig. 7I).
Safety evaluation
To evaluate the biosafety of FR-PolyRu, we administered eye drops to
mice for 14 days, as illustrated in fig. S32A. The results indicated
that the body weight of the mice remained stable throughout the
treatment period (fig. S32B). Furthermore, routine peripheral blood
analysis on day 14 demonstrated that FR-PolyRu eye drops had no adverse
effects on white blood cells, red blood cells, hemoglobin, or platelets
(fig. S33A). In addition, blood biochemical tests confirmed that liver
and kidney function were not compromised by the treatment (fig. S32, C
and D). Histological analysis via hematoxylin and eosin (H&E) staining
after 14 days revealed no notable abnormalities in the number and
arrangement of photoreceptor cells in FR-PolyRu–treated mice, with no
observable pathological changes in other organs (fig. S32E). In
addition, we further evaluated the toxicity of an extremely high
concentration of FR-PolyRu in mice. A 14-day cumulative dose of the eye
drops was administered through a single tail vein injection, with
observations conducted over the subsequent 14 days (fig. S32F). As
shown in figs. S32 (G to J) and S33B, FR-PolyRu at this concentration
did not induce any adverse effects on body weight, blood parameters, or
the organs of the mice.
The biosafety of FR-PolyRu was further evaluated with prolonged
administration over 30 days. Mice received continuous FR-PolyRu eye
drops, with body weight monitored throughout the treatment. After 30
days, routine blood tests and biochemical markers were assessed.
Histological analysis using H&E staining was conducted on major organs,
including the eyes, brain, heart, liver, spleen, lungs, and kidneys. In
addition, optical coherence tomography (OCT) scans were performed to
examine the anterior and posterior segments of the eye, as well as
intraocular pressure (IOP) measurements. The results showed no marked
changes in body weight, blood parameters, or organ histology ([134]Fig.
8, A to E, and fig. S34). OCT imaging of the eyes revealed no
abnormalities in either the anterior or posterior segments, and IOP
measurements remained within the normal range, suggesting that
prolonged treatment with FR-PolyRu does not induce any ocular or
systemic toxicity ([135]Fig. 8, F and G). These results confirm that
continuous 30-day administration of FR-PolyRu maintains excellent
biosafety, with no marked adverse effects observed on systemic health
or ocular function.
Fig. 8. Biosafety analysis of FR-PolyRu.
[136]Fig. 8.
[137]Open in a new tab
(A) Schematic diagram of the in vivo safety assessment experiment of
PBS or FR-PolyRu nanozymes via eye drop administration route. (B) Body
weight was monitored in healthy female C57 mice with continuous eye
drops of PBS or FR-PolyRu nanozymes (5 mg/ml) for 30 days (n = 3). (C
and D) Representative indicators of liver function [aspartate
aminotransferase (AST) and alanine aminotransferase (ALT)] and renal
function [blood urea nitrogen (BU) and creatinine (CREA)] of normal
mice treated with PBS or FR-PolyRu eye drop (n = 3). (E) Representative
H&E staining images of eyeballs and vital organs of normal mice eye
drop treated with PBS or FR-PolyRu eye drop for 30 days (n = 3). Scale
bars, 500, 50, or 100 μm. (F) IOP of normal mice treated with PBS or
FR-PolyRu eye drops for 30 days (n = 5). Each mouse was measured three
consecutive times to calculate the average IOP value. (G)
Representative OCT images of the anterior and posterior segments of the
eyeballs, taken 30 days after treatment with PBS or FR-PolyRu eye
drops. Scale bars, 800 and 500 μm. (H) Development of zebrafish embryos
incubated with FR-PolyRu nanozyme at various time points. Scale bar, 1
mm (n = 10). (I and J) Representative images of zebrafish embryos
incubated with different concentrations of nanozyme for 3 (I) or 7 (J)
days. Data were presented as mean ± SD. Scale bars, 0.5 and 0.2 mm.
Pharmacokinetic behavior and safety of FR-PolyRu eye drops were further
investigated by measuring the Ru metal content in eye and brain tissues
of mice using inductively coupled plasma mass spectrometry analysis at
4, 12, and 24 hours postadministration. The results indicated
detectable levels of Ru in the eyeballs at 4 hours, with levels
gradually decreasing over time and complete metabolism by 24 hours.
Quantitative analysis revealed that ~0.28% of the administered Ru
accumulated in ocular tissue at 4 hours postadministration (fig. S35),
demonstrating effective posterior segment delivery. Notably, no Ru was
detected in the brain tissue or peripheral blood at any time point
(fig. S35).
To further assess the biosafety of FR-PolyRu, we used zebrafish embryos
as a model for evaluating drug toxicity, teratogenicity, and other
safety concerns. Zebrafish embryos incubated with varying
concentrations of nanozyme exhibited normal hatching within 3 days,
with no signs of abnormal edema or deformities in the organs or eyes
([138]Fig. 8, H to J). On day 7, there were no developmental
abnormalities observed in the eyes or other organs. These results
demonstrate that FR-PolyRu nanozyme eye drops do not exhibit marked
toxic side effects in zebrafish, highlighting their good biosafety.
Overall, these comprehensive biosafety evaluations confirm that
FR-PolyRu demonstrates excellent safety profiles both in mice and
zebrafish, with no noticeable adverse effects on systemic health,
ocular function, or development.
DISCUSSION
This study highlights the notable therapeutic potential of FR-PolyRu
nanozyme for treating retinal neovascularization diseases through
noninvasive eye drops. Compared to conventional intravitreal
injections, FR-PolyRu eye drops present a less invasive approach, which
may enhance patient compliance and minimize complications. The
liposome-based formulation, with fluorination and RGD modifications,
effectively traverses ocular barriers to achieve delivery within the
retinal vasculature.
The noninvasive nature of FR-PolyRu eye drops represents a major
innovation in this study, offering substantial advantages over
traditional retinal neovascularization disease therapies. Despite their
effectiveness, intravitreal injections pose several challenges,
including patient discomfort, risk of infection, retinal detachment,
and elevated IOP. These risks necessitate frequent monitoring,
contributing to patient anxiety and affecting treatment adherence. In
contrast, the ease of administration of eye drops enhances patient
comfort and quality of life while also improving treatment
accessibility, particularly in resource-limited settings.
The ability of FR-PolyRu eye drops to overcome anatomical and
physiological barriers to the eye is a key determinant of its success
as a noninvasive treatment. The cornea and the blood-retinal barrier,
while protective, present considerable obstacles to drug delivery to
the posterior segment of the eye. The liposome-based design of
FR-PolyRu, incorporating fluorination and RGD modifications,
facilitates deep tissue penetration by enhancing membrane permeability
and targeted delivery. Fluorination lowers the energy barrier for
transmembrane transport, allowing nanoparticles to effectively diffuse
across ocular barriers, while RGD modification improves vascular
targeting and cellular uptake within retinal tissues. Beyond passive
diffusion, FR-PolyRu introduces an active transport mechanism via
nanozyme-catalyzed oxygen release, which synergizes with
fluorination-enhanced penetration. The oxygen generated from H[2]O[2]
decomposition actively promotes intraocular fluid dynamics,
facilitating nanoparticle diffusion across ocular barriers and
enhancing drug distribution and retention in the posterior segment.
This dual mechanism—fluorination-driven penetration and
oxygen-facilitated transport—enables FR-PolyRu to efficiently overcome
electrostatic barriers, providing a comprehensive strategy for
optimized retinal drug delivery.
In addition to effective delivery, FR-PolyRu nanozyme exhibits potent
antioxidant properties by scavenging ROS through SOD-like and CAT-like
activities. Unlike natural enzymes, FR-PolyRu nanozyme demonstrates
higher stability, superior bioavailability, and sustained activity,
offering advantages in complex biological environments. In vitro and in
vivo studies have shown that it can reduce oxidative damage, maintain
mitochondrial function, and inhibit pathological retinal
neovascularization.
Safety evaluations revealed no systemic toxicity or adverse effects
associated with FR-PolyRu eye drops, even at high concentrations,
supporting its favorable safety profile. This finding further
strengthens its viability as a therapeutic option for ocular diseases,
especially in comparison to invasive anti-VEGF therapies.
Despite these promising results, certain limitations remain. Mouse
models of OIR may not fully mimic human retinal neovascularization
diseases, necessitating further studies, including clinical trials, to
confirm efficacy in patients. In addition, while FR-PolyRu exhibits
considerable antioxidant and antiangiogenic activity, further
investigation is required to elucidate its molecular interactions
during ocular penetration and biodistribution.
In conclusion, FR-PolyRu nanozyme offers a promising noninvasive
therapeutic strategy for retinal neovascularization diseases, with
advantages in antioxidant stability, tissue penetration, and safety.
Its ability to effectively target and mitigate pathological oxidative
stress and neovascularization provides a comprehensive approach to
addressing the core drivers of retinal neovascularization disease
progression.
MATERIALS AND METHODS
Preparation of PolyRu nanozyme
A total of 133.0 mg of polyvinylpyrrolidone (molecular weight: 30 kDa;
Sigma-Aldrich) was dissolved in 180.0 ml of methanol. Subsequently,
20.0 ml of a deionized aqueous solution of RuCl[3] (1.0 mg/ml;
Sigma-Aldrich) was added. The mixture was stirred magnetically at 70°C
and 400 rpm for 3 hours. Upon completion of the reaction, the solvent
was removed by centrifugation, followed by ultrasonication. The
resulting residue was transferred to a centrifuge tube, and an equal
volume of chloroform and five volumes of n-hexane were added. The
mixture was washed repeatedly until a neutral pH was achieved and then
dried in a vacuum oven.
Preparation of fluorinated DSPE-PEG
Fluorinated DSPE-PEG derivatives were synthesized according to
previously reported methods ([139]31). Specifically, perfluoropropionic
anhydride, heptafluorobutyric anhydride, or nonafluoropentanoic
anhydride was reacted with DSPE-PEG-NH[2] in methanol (8.0 ml) at a
molar ratio of 1.1:1. The mixture was stirred at room temperature for
48 hours to ensure complete conversion. The resulting fluorinated
products (DSPE-PEG-F[5], DSPE-PEG-F[7], and DSPE-PEG-F[9]) were then
isolated by precipitation in cold diethyl ether and purified for
subsequent use.
Preparation of F-PolyRu nanozyme
In a 100.0-ml round-bottom flask, 0.5 mg of DSPE-PEG-F[7], 7.0 mg of
lecithin (Ponsure Biotechnology), 2.0 mg of cholesterol
(Sigma-Aldrich), and 3.0 ml of organic solvent
(methanol:chloroform = 1:2) were combined. The solvent was evaporated
to form a thin lipid film. Next, 2.0 ml of PolyRu nanozyme solution
(5.0 mg/ml) was added to hydrate the film. The resulting suspension was
sonicated for 5 min to ensure uniform dispersion.
Preparation of R-PolyRu nanozyme
A total of 1.0 mg of DSPE-PEG-RGD (grafting efficiency: 97.1%; Xi’an
Ruixi Biotech), 6.0 mg of lecithin, 1.0 mg of DSPE-PEG-NH[2], and 2.0
mg of cholesterol were dissolved in 3.0 ml of organic solvent
(methanol:chloroform = 1:2) in a 100.0-ml flask. After solvent
evaporation, the resulting thin film was hydrated with 2.0 ml of PolyRu
nanozyme solution (5.0 mg/ml) and sonicated for 5 min to facilitate
nanozyme encapsulation.
Preparation of FR-PolyRu nanozyme
In a 100.0-ml flask, 0.5 mg of DSPE-PEG-F[7], 1.0 mg of DSPE-PEG-RGD,
6.0 mg of lecithin, 1.0 mg of DSPE-PEG-NH[2], 2.0 mg of cholesterol,
and 3.0 ml of organic solvent (methanol:chloroform = 1:2) were mixed.
After forming a thin film by solvent evaporation, 2.0 ml of PolyRu
nanozyme (5.0 mg/ml) was added to hydrate the film. The mixture was
then sonicated for 5 min to obtain the final FR-PolyRu nanozyme
formulation. Cy5.5–FR-PolyRu nanozymes were prepared using the same
procedure, with the addition of 0.5 mg of Cy5.5-DSPE-PEG during film
formation.
Preparation of ICG-PolyRu, ICG–F-PolyRu, ICG–R-PolyRu, and ICG–FR-PolyRu
nanozymes
To prepare ICG-modified nanozymes, 0.5 mg of DSPE-PEG-F[7], 1.0 mg of
DSPE-PEG-RGD, 6.0 mg of lecithin, 1.0 mg of DSPE-PEG-NH[2], and 2.0 mg
of cholesterol were added to 3.0 ml of organic solvent
(methanol:chloroform = 1:2) in a 100.0-ml flask. The solvent was
evaporated to form a lipid film, which was hydrated with 2.0 ml of
PolyRu nanozyme solution (5.0 mg/ml) and sonicated for 5 min.
Subsequently, 2.0 mg of ICG was added and stirred for 30 min at room
temperature in the dark. ICG-PolyRu, ICG–F-PolyRu, and ICG–R-PolyRu
were prepared using the same method, replacing DSPE-PEG-F[7] or
DSPE-PEG-RGD accordingly.
Characterization
The morphology of the nanozyme in aqueous solution was analyzed using a
Thermo Fisher Scientific Talos L120C transmission electron microscope
operated at 120 kV. To compare the morphology before and after adding
2% photosensitive acid negative staining solution (Solarbio Science &
Technology, Beijing), the sample was subjected to negative staining.
Morphology and height measurements of the nanozyme were performed using
an atomic force microscope (Dimension Icon, Bruker, Germany). The
crystalline structure was analyzed using x-ray diffraction on a D8
Focus diffractometer (Bruker, Germany) with Cu Kα radiation (λ = 1.5406
Å). Elemental and chemical bond information was obtained by XPS
(ESCALAB 250, Thermo Fisher Scientific, USA). ^1H NMR spectra were
recorded on a Bruker Avance III NMR spectrometer (400 MHz) with D[2]O
as the solvent, and chemical shifts (in parts per million) were
referenced to tetramethyl silane. Dynamic light scattering (DLS) and
zeta potential measurements were performed using a Nano ZS90 instrument
(Malvern, UK). Osmotic pressure was measured using a freezing point
osmometer (Advanced Instruments, USA). The viscosity of the liquid was
determined using a rotary viscometer (DV-2 Pro, Brookfield, USA).
Evaluate the stability of FR-PolyRu nanozyme
DLS data of the material dispersed in double-distilled water at a
concentration of 0.1 mg/ml at 0, 7, 14, and 30 days were measured using
a DLS instrument (Nano-ZS90, Malvern, UK). RuO[2], PolyRu, and
FR-PolyRu were dispersed in water, and then their Tyndall effects and
the sedimentation at 0, 7, 14, and 30 days were observed to evaluate
the stability of the nanozymes.
SOD-like activity assays
The level of SOD-like activity of nanozymes was determined by detecting
their elimination rate of O[2]^•–. The principle of the method used can
be summarized as follows: O[2]^•– can be produced in the
xanthine-xanthine oxidase system, and O[2]^•– can reduce a certain
amount of oxidized cytochrome c to reduced cytochrome c, which has the
maximum light absorption at 550 nm. In the presence of SOD, the
reaction of O[2]^•–-reducing cytochrome c is inhibited, so the SOD-like
activity of the nanozymes can be detected indirectly by calculating the
inhibition rate of O[2]^•–.
The total volume of the reaction system was 300.0 μl: xanthine (50.0
μl), cytochrome c (50.0 μl), xanthine oxidase (20.0 μl), and PBS (180.0
μl). The increased absorbance of cytochrome c at 550 nm for 1 min was
determined by an ultraviolet spectrophotometer as ΔA1. Under the
condition of constant total volume, the volume of xanthine oxidase and
PBS was adjusted so that ΔA1 reached 0.0225, and then nanozymes were
added to the system to determine the increased absorbance of cytochrome
c at 550 nm for 1 min, as ΔA2. The elimination rate of O[2]^•– was
calculated as (ΔA1 – ΔA2)/ΔA1 × 100%.
CAT-like activity assays
The CAT-like activity of different nanozymes was evaluated by detecting
the dissolved oxygen concentration of the nanozymes (10.0 μl/ml) in
H[2]O[2] aqueous solution [200.0 mM (pH 7.4)] with a dissolved oxygen
meter (JPBJ-608, Leici). H[2]O[2] decomposition experiment: Different
nanozymes (10.0 μg/ml) were mixed with 3-ml H[2]O[2] solution [10.0 mM
(pH 7.4)] in a cuvette, and the absorbance of the reaction system was
continuously monitored at 240 nm within 10 min by an ultraviolet
spectrophotometer to compare the ability of different nanozymes to
degrade H[2]O[2].
POD- and oxidase-like activity assays
The POD- and oxidase-like activity of the nanozymes was evaluated by
measuring the absorption value of trimethylboron (TMB) at 652 nm for 10
min with a microplate reader. The reaction system for POD-like activity
detection was 10.0 μl of nanozyme [100.0 μg/ml in 50.0 mM phosphate
buffer (pH 7.4)], 1 μl of TMB [20.0 mg/ml in dimethyl sulfoxide
(DMSO)], 5.0 μl of H[2]O[2] (10 M), and 84.0 μl of PBS. The reaction
system for oxidase-like activity was 10.0 μl of nanozyme [100.0 μg/ml
in 50.0 mM phosphate buffer (pH 7.4)], 1.0 μl of TMB (DMSO: 20.0
mg/ml), and 89.0 μl of PBS.
Nanozyme-induced O[2]^•–, ^1O[2], and ·OH production was determined by ESR
Using a Bruker ESR spectrometer (A300-10/12, Germany) at room
temperature, ESR spectroscopy was performed. 5,5-dimethyl-1-pyrroline
N-oxide (DMPO) was used as a O[2]^•– and ·OH catching agent.
2,2,6,6-tetramethylpiperidine (TEMP) was used as a ^1O[2] catching
agent. O[2]^•– was generated via xanthine oxidase–catalyzed oxidation
of xanthine, and its presence was detected using DMPO as a spin trap in
the ESR assay. ^1O[2] was produced via photoexcitation of TiO[2], and
TEMP was used as a spin trap for ESR detection. ·OH was generated via
the Fenton reaction (Fe^2+ + H[2]O[2] → ·OH), and its capture was
detected using DMPO in ESR measurements.
DPPH radical scavenging assay
Different concentrations of RuO[2], PolyRu, and FR-PolyRu were added to
the ethanol solution containing DPPH. The final concentration of DPPH
after mixing was 62.5 μM, and the final concentration of nanozymes was
0.78125, 1.5625, 3.125, 6.25, 12.5, 25.0, 50.0, 100.0, and 200.0 μg/ml.
The above mixture volume was kept the same, and the absorbance value of
DPPH at 517 nm was detected after 30 min of reaction. The radical
scavenging activity of DPPH was calculated using the following formula:
[MATH: DPPH radical cation scavenging
activity=[ADPPH−(ASample−
ABlank)]/ADPPH × 100%
:MATH]
A[DPPH] is the absorbance of DPPH solutions, A[Sample] is the
absorbance of the sample after reacting with DPPH, and A[Blank] is the
absorbance of the blank solutions.
ABTS radical scavenging assay
The radical scavenging activity of ABTS was evaluated using an ABTS
radical scavenging assay kit (Solarbio Science & Technology, BC4770).
Experiments were performed according to the instructions. The test was
conducted according to the detailed steps provided in the kit
instructions. The final concentration of nanozymes was 0.78125, 1.5625,
3.125, 6.25, 12.5, 25.0, 50.0, 100.0, and 200.0 μg/ml. The radical
scavenging activity of ABTS was calculated using the following formula:
[MATH: ABTS radical cation scavenging
activity=[AABTS−(ASample−
ABlank)]/AABTS × 100%
:MATH]
A[ABTS] is the absorbance of ABTS solutions, A[Sample] is the
absorbance of the sample after reacting with ABTS, and A[Blank] is the
absorbance of the blank solutions.
Calculation details
Spin-polarized DFT calculations were performed using the Vienna Ab
initio Simulation Package ([140]32, [141]33) to investigate the
adsorption properties of the synthesized materials. The projector
augmented wave method ([142]33) was used with a cutoff energy of 450
eV, alongside the Perdew-Burke-Ernzerhof functional ([143]34). To
account for van der Waals interactions, the DFT-D3 method ([144]35) was
applied. AIMD simulations were used to construct amorphous ruthenium
(a-Ru). The optimized Ru crystal was heated from 500 to 3000 K using
velocity scaling over 1.25 ps for every 20 molecular dynamics (MD)
steps, followed by equilibration at 2000 K for 0.5 ps with a time step
of 1 fs to obtain the a-Ru structure. Subsequently, a PVP molecule
containing 3 U was placed on the surface to create the a-Ru–PVP model.
All models underwent full relaxation with energy and force convergence
criteria set at 10^−5 eV and 0.02 eV/Å, respectively. The γ point was
used in the K-point mesh, and the adsorption energy (E[ads]) was
calculated using [145]Eq. 1
[MATH:
Eads=Etotal−Esubstrate
−Eadsorbate :MATH]
(1)
The E[total], E[substrate], and E[adsorbate] represent the energies of
the adsorption structure, substrate, and adsorbate, respectively. The
free energies have been calculated using the following [146]Eq. 2
[MATH:
G=EDFT
+ZPE−TS :MATH]
(2)
The G, E[DFT], ZPE, and TS represent the free energy, energy from DFT
calculations, zero-point energy, and entropic contributions,
respectively.
Cell culture
HUVECs and HCECs were obtained from the American Type Culture
Collection. HUVEC cell line was cultured in RPMI 1640 medium (Gibco)
supplemented with 10% fetal bovine serum (FBS) (VivaCell, [147]C04001)
and 1% penicillin/streptomycin (Gibco). HCEC cell line was cultured in
Eagle’s minimum essential medium (BNCC, 360906) supplemented with 10%
FBS (VivaCell, [148]C04001) and 1% penicillin/streptomycin (Gibco).
HRECs (Seattle, WA, USA) were cultured in EGM-2 medium (Lonza,
CC-4176). All the above three cell lines were cultured in a cell
incubator at 37°C with 5% CO[2].
In vitro cytotoxicity
HRECs or HUVECs were seeded at a density of 5000 cells per well in a
96-well cell culture plate and cultured in a cell incubator for 24
hours, and then RuO[2], PolyRu, and FR-PolyRu at different
concentrations (0 to 100.0 μg/ml) were added for another 24 hours.
CCK-8 assay was used to detect cell viability after incubation with
different materials.
Cellular uptake assay
HRECs or HUVECs were inoculated in 12-well plates and cultured in
confocal dishes for 24 hours. The Cy5.5-labeled FR-PolyRu nanozymes
were added to confocal dishes for further incubation for 1, 2, 4, or 8
hours. The cell uptake of the nanozyme after incubation for different
times was observed by confocal microscopy, and it was observed that the
nuclei stained by 4y y,diamidino-2-phenylindole (DAPI; C0065, Solarbio
Science and Technology Co. Ltd.) were blue, and the FR-PolyRu labeled
by Cy5.5 was green.
Cell viability rescue assay
HRECs or HUVECs were inoculated on 96-well plates with a density of
10,000 cells per well and cultured in incubators for 24 hours.
Different concentrations of H[2]O[2] (0 to 400.0 μM) were then added,
and the incubation continued for 6 hours. Last, the toxicity of
H[2]O[2]to the cells was detected by CCK-8 assay.
HRECs or HUVECs were inoculated on 96-well plates with a density of
6000 cells per well and cultured in incubators for 24 hours. Then,
nanozymes of different concentrations (25.0, 50.0, and 100.0 μg/ml)
were added and incubated for 18 hours to preprotect the cells. Then,
the old medium was replaced with a fresh medium of 200.0 μM H[2]O[2]
with or without nanozymes of different concentrations, and the culture
was continued for 6 hours. Last, the cell viability was measured by
CCK-8 assay.
Tubulin immunofluorescence analysis
HRECs or HUVECs were seeded in confocal dishes and cultured for 24
hours. The experiments were divided into four groups, which were
replaced with fresh medium containing PBS, H[2]O[2] (100.0 μM), and
H[2]O[2] (100.0 μM) + PolyRu (100.0 μg/ml) or H[2]O[2] (100.0 μM) +
FR-PolyRu (100.0 μg/ml), and continued for 6 hours. Cells were washed
with PBS, incubated with 4% paraformaldehyde for another 10 min to fix
the cells, and then rinsed again with PBS. HUVECs were incubated with
0.1% Triton X-100 for 10 min at room temperature, followed by a further
1 hour of incubation using 5% bovine serum albumin reagent. Antitubulin
antibody (Beyotime Biotechnology, AT819; 1:500) was added to the
confocal dish and incubated overnight at 4°C. The next day, the cells
were stained with a fluorescent secondary antibody (Alexa Fluor 594),
and the nuclei were lastly stained with DAPI. Cell morphology was
observed under a confocal microscope.
Analysis of intracellular ROS level
We used the fluorescent probe DCFH-DA (Beyotime Biotechnology, S0033S)
to measure intracellular ROS levels. Flow cytometry (fluorescein
isothiocyanate fluorescence channel) was used to detect the changes of
intracellular ROS: HRECs or HUVECs were seeded in six-well plates and
cultured. When the cell density was about 70%, the drug was added and
incubated for 6 hours. The cells were collected and incubated with
DCFH-DA (10 μM) for 30 min at 37°C in the dark, and the changes in
intracellular ROS were analyzed by flow cytometry. The experiment was
divided into five groups: PBS, H[2]O[2] (100.0 μM), H[2]O[2] (100.0
μM) + RuO[2] (100.0 μg/ml), H[2]O[2] (100.0 μM) + PolyRu (100.0 μg/ml),
and H[2]O[2] (100.0 μM) + FR-PolyRu (100.0 μg/ml).
ROS generation in HRECs was observed by fluorescence microscopy: HRECs
were seeded and cultured in 12-well plates. When the cell density was
about 70%, RuO[2] (100.0 μg/ml), PolyRu (100.0 μg/ml), or FR-PolyRu
(100.0 μg/ml) was added and incubated for 6 hours, followed by H[2]O[2]
(800.0 μM) stimulation for 30 min. Last, the cells were incubated with
DCFH-DA (10.0 μM) at 37°C in the dark for 30 min, and the cells were
washed with PBS before being photographed under a fluorescence
microscope.
ROS generation in HUVECs was observed by fluorescence microscopy:
HUVECs were seeded and cultured in 12-well plates. When the cell
density was about 70%, FR-PolyRu nanozymes (25.0 to 100.0 μg/ml) were
added and incubated for 6 hours, followed by H[2]O[2] (800.0 μM)
stimulation for 30 min. Last, the cells were incubated with DCFH-DA
(10.0 μM) at 37°C in the dark for 30 min, and the cells were washed
with PBS before being photographed under a fluorescence microscope.
MMP assay
The MMP of HRECs was detected by MMP assay kit with JC-1 (C2006,
Beyotime Biotechnology). HRECs were inoculated in a six-well plate or
confocal dish. When the cell density reached about 70%, the cells were
incubated with RuO[2] (100.0 μg/ml), PolyRu (100.0 μg/ml), or FR-PolyRu
(100.0 μg/ml) for 6 hours and then stimulated with H[2]O[2] (800.0 μM)
for 30 min. JC-1 and Hoechst 33342 were added to confocal dishes and
incubated at 37°C for 20 min. The cells were cleaned with PBS for
confocal microscope observation in the red channel for J-aggregates and
green channel for J-monomer, separately.
In addition, after the drug incubation, the cells in the six-well
plates were collected, and the cells containing JC-1 were cultured at
37°C for 20 min. After the cells were cleaned twice with PBS, the
changes in intracellular MMP were observed by flow cytometry.
MitoTracker and MitoSOX staining
To analyze changes in mitochondrial membrane potential, HRECs were
stained with MitoTracker Green (100 nM; C1048, Beyotime Biotechnology).
To analyze mitochondrial ROS production, HRECs were stained with
MitoSOX Red (5.0 μM; S0061S, Beyotime Biotechnology).
HRECs were inoculated in confocal dishes, incubated with RuO[2] (100.0
μg/ml), PolyRu (100.0 μg/ml), or FR-PolyRu (100.0 μg/ml) for 6 hours
when the cell density was about 70%, and then stimulated with H[2]O[2]
(800.0 μM) for 30 min. After exchanging fresh medium, MitoSOX Red,
MitoTracker Green, and Hoechst 33342 (1:100; [149]C00031, Solarbio
Science and Technology Co. Ltd) were added and incubated at 37°C for 30
min in the dark light. After washing with PBS, images were obtained
under a confocal microscope.
Evaluation of intracellular adenosine 5′-triphosphate level
HRECs were inoculated on six-well plates, incubated with RuO[2] (100.0
μg/ml), PolyRu (100.0 μg/ml), or FR-PolyRu (100.0 μg/ml) for 6 hours
when the cell density reached about 70%, and then stimulated with
H[2]O[2] (800.0 μM) for 30 min. Then, the adenosine 5′-triphosphate
(ATP) level of each group was detected with the ATP assay kit (S0027,
Beyotime Biotechnology). To put it simply, the medium is removed, 200.0
μl lysate is added, and the cells are repeatedly blown. After cell
lysis, the cells were centrifuged at 12,000 rpm for 10 min, and
supernatant (20.0 μl) was mixed with ATP detection working liquid
(100.0 μl) in an opaque 96-well plate. The luminescence was monitored
at 560 nm using a luminometer.
Experimental animals
C57BL/6J mice used in the experiments were purchased from SPF (Beijing)
Biotechnology Co. Ltd. (Beijing, China). All animal experiments were
conducted in accordance with the ethical standards set by the Animal
Ethics Committee of Zhengzhou University (approval number:
ZZUIRBGZR2024-1521). All experimental animals were housed and fed in a
standard animal house with a 12-hour light/12-hour dark cycle. Mice
were examined and excluded for ophthalmic disease before performing
experiments.
Transwell
HCECs were seeded in the upper transwell chamber at a density of
5 × 10^4 cells per well, and HRECs were seeded in the lower transwell
chamber at a density of 5 × 10^4 cells per well. After 24 hours of
incubation, drugs were added to the upper chamber. The experiments were
divided into five groups: Free-ICG, ICG-PolyRu, ICG–R-PolyRu,
ICG–F-PolyRu, and ICG–FR-PolyRu. At 0, 12, and 48 hours after drug
addition, photographs were taken using the IVIS in vivo imaging system
(Lumina XR series, Perkin Elmer) to observe drug penetration from the
upper chamber to the lower chamber.
SMD simulations
The CHARMM-GUI membrane builder, in conjunction with the CHARMM36m
force field, was used to construct a biomimetic cell membrane bilayer.
The bilayer consisted of six lipid
types—1,2-dimyristoyl-sn-glycero-3-phosphocholine,
1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine,
palmitoylsphingomyelin, cholesterol,
1,2-dimyristoyl-sn-glycero-3-phosphate, and
1,2-dimyristoyl-sn-glycero-3-phospho-l-serine in an equal molar ratio
of 1:1:1:1:1:1. The system comprised a total of 132 lipid molecules per
layer, encapsulated within a water box of dimensions 8.66 nm by 8.66 nm
by 10.00 nm. An isothermal-isobaric (constant number of particles,
pressure, and temperature ensemble, or NPT) MD simulation was
performed, maintaining constant pressure (P = 1 atm) and temperature
(T = 310 K) using the V-rescale thermostat and the Parrinello-Rahman
barostat. Lennard-Jones interactions were evaluated with a cutoff of
1.2 nm, while long-range electrostatic interactions were computed using
the particle mesh Ewald method. Periodic boundary conditions were
applied throughout the simulation. For the SMD simulations, a harmonic
spring was attached to a dummy atom on one end and to the drug molecule
on the other. The dummy atom was pulled in the z direction at a
constant velocity v, covering a displacement Δz = vt, where t denotes
time. The force applied to the drug was determined using the equation
F = k(Δz − vt), where k was set to 600 kJ/(mol·nm^2) and the pulling
velocity to 5.0 nm/ns. These values, previously validated in our
earlier studies, were selected to ensure consistency and reliability.
To prevent membrane drift induced by the applied force, harmonic
restraints with a spring constant of 1000 kJ/(mol·nm^2) were applied to
the phosphate atoms in both lipid layers along the z axis. All
simulations were conducted using the GROMACS 5.1 software package.
In vivo photoacoustic tracking
Healthy 6- to 8-week C57BL/6J mice were randomly divided into five
groups (n = 3): Free-ICG, ICG-PolyRu, ICG–R-PolyRu, ICG–F-PolyRu, and
ICG–FR-PolyRu. The drug was administered topically by eye drop, 10.0 μl
(5.0 mg/ml) per mouse. After administration, the mice were kept in a
dark environment. At 0.5, 3, 4, 6, and 12 hours after administration,
the photoacoustic signal in the eyes of mice was detected by Vevo
LAZR-X.
Biodistribution and penetration of nanozymes in the eyeball
Healthy 6- to 8-week C57BL/6J mice were randomly divided into three
groups (n = 3). Topical administration was performed as eye drops with
Free-Cy5.5 and Cy5.5-PolyRu. At 1, 6, and 12 hours after
administration, eyeballs were removed and snap frozen in liquid
nitrogen, embedded in optimal cutting temperature compound,
cryo-sectioned into 12-μm thickness, and mounted on slides. Slides were
washed three times with PBS and sealed with an antifade mounting medium
containing DAPI. The distribution of nanozymes in the eyeball was
observed by confocal microscopy.
Establishment and treatment of the OIR mouse model
Establishment of OIR animal model: Neonatal mice and their mothers were
placed in a high-oxygen feeding tank (75% O[2]) for 5 days after birth.
Then, the mice at day 12 after birth were kept in normal air (21% O[2])
for 5 days. Last, mice 17 days after birth were analyzed.
In vivo treatment experiment (intraocular injection): OIR mice at day
12 after birth were randomly divided, and the drug was injected into
the eyes of mice with a 30-gauge needle. The experimental groups were
untreated, PBS, RuO[2] (1.0 μg), and PolyRu (0.5, 1.0, and 2.0 μg).
Last, the therapeutic effect of OIR mice 17 days after birth was
analyzed.
In vivo therapy experiment (eye drop): OIR mice at day 12 after birth
were randomly assigned to be treated twice daily with 2.5 μl (1 mg/ml)
eye drops in each eye. The experiments were divided into untreated,
PBS, RuO[2], PolyRu, F-PolyRu, and FR-PolyRu. Last, the therapeutic
effect of OIR mice 17 days after birth was analyzed.
Immunostaining of the whole-mount retinas
The enucleated eyes were fixed in a 4% paraformaldehyde solution for 20
min. The retinal cups were then dissected and divided into four petals.
Following dissection, the retinas were washed three times with PBS (5
min each wash) and subsequently blocked in PBS containing 5% FBS for 30
min at room temperature. The tissues were then purified with isolectin
GS-IB4 antibody ([150]I21411, Thermo Fisher Scientific; 1:200)
overnight at 4°C and washed three times with PBS (5 min each wash).
Afterwards, the retinal cups were flat mounted, and the whole-mount
retina images were acquired using a confocal microscope.
TUNEL assay
TUNEL assay kit (Beyotime Biotechnology, C1086) was used to analyze the
death of mouse retinal cells after different treatments. Briefly,
frozen tissue sections of the eyeball were fixed in 4% paraformaldehyde
and washed twice with PBS. The slides were further incubated in PBS
containing 0.5% Triton X-100 for 5 min at room temperature and lastly
incubated with TUNEL assay solution for 60 min at 37°C in the dark.
Slides were washed three times with PBS and sealed with an antifade
mounting medium containing DAPI. Cell apoptosis was observed under a
confocal microscope.
Transcriptomics sequencing and data analysis
Mice in the normal group, the PBS group (OIR), and the FR-PolyRu group
(OIR) were randomly selected, and their retinal tissues were taken
(n = 3). Total RNA was separated from retinal tissues by TRIzol reagent
(Solarbio Science & Technology, R1100) and then quickly frozen in
liquid nitrogen. The whole RNA-seq analysis was performed by Biomarker
Technologies (Beijing).
RNA extraction and reverse transcription quantitative polymerase chain
reaction analysis
Mice in the normal group, the PBS group (OIR), and the FR-PolyRu group
(OIR) were randomly selected, and their retinal tissues were taken
(n = 3). The RNA of the retinas was extracted using TRIzol reagent
(Solarbio Science & Technology, R1100) according to the manufacturer’s
protocols. cDNA was obtained by reverse transcribing total RNA with
HiScript III All-in-one RT SuperMix Perfect for quantitative polymerase
chain reaction (qPCR) kit (R333-01, Vazyme) according to the
manufacturer’s protocols. Reverse transcription (RT)–qPCR was conducted
using the Taq Pro Universal SYBR qPCR Master Mix kit (Q712-02, Vazyme)
according to the manufacturer’s instructions. The relative mRNA
expression was calculated by the comparative cycle threshold (CT)
method (ΔΔCT method). All experiments were repeated three times. Primer
sequences used were as follows: Igfbp6 gene, CCTTCTCTGTCCTCCCCTT
(forward) and CTCCGCCGCTGTTTACTT (reverse), and β-actin gene,
TGTGTCCGTCGTGGATCTGA (forward) and TTGCTGTTGAAGTCGCAGGAG (reverse).
Western blot
Mice in the normal group, the PBS group (OIR), and the FR-PolyRu group
(OIR) were randomly selected, and their retinal tissues were taken
(n = 3). Total protein was extracted from the retinal tissue of mice
(n = 3) in the normal, PBS, and FR-PolyRu groups using a total protein
extraction kit (BC3710, Solarbio Science & Technology). The protein
concentration was determined by bicinchoninic acid protein assay
(P0009, Beyotime Biotechnology). The extracted proteins were separated
by SDS–polyacrylamide gel electrophoresis and transferred to
polyvinylidene fluoride membranes. Membranes were incubated with the
indicated primary antibody overnight at 4°C, followed by incubation
with the indicated secondary antibody for 1 hour at room temperature.
Signals were detected by a hypersensitive enhanced chemiluminescence
kit (P10018S, Beyotime Biotechnology).
The information we used for primary antibodies and secondary antibodies
is as follows: rabbit anti-Igfbp6 (GB114043-100, Servicebio), rabbit
anti-ERK (#4695T, Cell Signaling Technology), rabbit anti–phospho-ERK
(p-ERK; #4370T, Cell Signaling Technology), rabbit anti-PI3K (ab191606,
Abcam), rabbit anti-Phospho-PI3K (#4228T, Cell Signaling Technology),
rabbit anti-AKT (ab179463, Abcam), rabbit anti–phospho-AKT (p-AKT;
ab192623, Abcam), mouse anti–β-actin (AF0003, Beyotime Biotechnology),
anti–rabbit–immunoglobulin G (IgG) conjugated to horseradish peroxidase
(HRP; #7074, Cell Signaling Technology), and anti–mouse-IgG conjugated
to HRP (A0208, Beyotime Biotechnology). The primary antibody was
diluted 1:1000, and the secondary antibody was diluted 1:5000.
In vivo biosafety evaluation
Healthy C57BL/6J mice were randomized (n = 3) to detect the safety of
FR-PolyRu nanozyme in mice after administration by eye drop or tail
vein. Mice administered by tail vein were divided into three groups:
PBS, FR-PolyRu (7.0 mg/kg), and FR-PolyRu (35.0 mg/kg). Mice given eye
drops were divided into three groups: normal, PBS, and FR-PolyRu (5.0
mg/ml); each mouse was given once a day, and each eye was given 5.0 μl
of eye drops. After the start of administration, the status of the mice
was continuously observed for 30 days, and the weight change of the
mice was recorded. On day 30, IOP and OCT were measured. Following
this, the mice were euthanized, blood was collected for biochemical
analysis, and H&E staining was performed on the eyeballs, brain, heart,
liver, spleen, lungs, and kidneys.
Statistical analysis
All data were expressed as mean ± SD. All data were analyzed for
statistical differences using GraphPad Prism 8.0. One-way analysis of
variance (ANOVA) with Tukey’s multiple comparisons was used when
comparing three or more groups within one factor. Two-way ANOVA with
Šidák’s multiple comparisons was used when comparing three or more
groups of two factors. In all statistical analysis methods, a
difference of P < 0.05 was considered statistically significant.
Acknowledgments