Humanin reduces damage to eye support cells caused by beta-amyloid through energy control and cell cleanup processes

To investigate the effects of Aβ on mitochondrial dysfunction and oxidative stress, ARPE19 cells and primary marmoset RPE cells were exposed to oligomeric Aβ1‐42. Exogenously treated FITC‐labeled Aβ entered adapted ARPE19 cells, disrupting the intercellular junctions of ZO‐1, a critical tight junction protein necessary for maintaining the structure and integrity of the RPE. This disruption of tight junction led to an increase in both cell area and perimeter. However, treatment with HN mitigated these Aβ‐induced morphological alterations, including the increases in cell area and perimeter (Figure 1A). In primary marmoset RPE cells, Aβ treatment induced multinucleation with nuclear enlargement, in addition to the morphological changes observed in ARPE19 cells, and these alterations were restored by HN treatment (Figure 1B).

In addition to morphological assessments, we measured the substantial transepithelial electrical resistance (TEER) in mature primary marmoset RPE cells to confirm the tight junction integrity and barrier function (Srinivasan et al. 2015). Aβ exposure compromised the TEER value, indicating disrupted barrier function, but this effect was significantly restored by HN co‐treatment (Figure 1C). Aβ increased cytosolic reactive oxygen species (Figure 1D) and mitochondrial superoxide (Figure 1E), both of which were prevented by the cotreatment with HN. As an indicator of mitochondrial activity, oxygen consumption, particularly protonophore (FCCP)‐induced maximal respiration, was markedly deteriorated by Aβ (Figure 1F). HN partially recovered Aβ‐induced suppression in mitochondrial respiratory capacity. The fluorescence ratio of potentiometric dye, JC‐1, which reflects mitochondrial membrane potential, was reduced by Aβ, indicating mitochondrial depolarization. Aβ exposure abolished the hyperpolarizing response to oligomycin due to the accumulation of ATP‐consuming dysfunctional mitochondria (Figure 1G). Consistently, HN restored mitochondrial electrical gradient as well as the hyperpolarizing response to oligomycin. Taken together, HN protects against oxidative stress, mitochondrial dysfunction, and degenerative changes with damaged barrier integrity elicited by pathogenic Aβ exposure.

Concurrently reducing intracellular Aβ levels, HN also decreased the abundance of p62 and increased the LC3II/I ratio, indicating enhanced autophagic processes. Aβ promoted the phosphorylation of Transcription factor EB (TFEB), leading to its retention in the cytosol. In contrast, HN reduced the phosphorylated (inactive) form of TFEB, accentuating its nuclear actions as the master regulator of lysosomal biogenesis, as evidenced by increased level of Lysosomal Associated Membrane Protein 1 (LAMP1) (Figure 2A) (Martina et al. 2014; Sardiello et al. 2009). Immunofluorescence analysis further showed that HN enhanced nuclear localization of TFEB in Aβ‐treated ARPE19 cells (Figure S1A,B). Additionally, HN significantly decreased the abundance of mitochondrial proteins such as TIM23, COX IV, and TOM20, suggesting the potential elimination of mitochondria through autophagy (Figure 2B). To determine whether Aβ‐accumulated mitochondria were undergoing elimination, live cell imaging showed the colocalization of mitochondria (Mitotracker) with FITC‐labeled Aβ or with lysosome (Lysotracker). In response to HN co‐treatment, the colocalization of FITC‐labeled Aβ with mitochondria decreased (Figure 2C), while the colocalization of lysosome with mitochondria increased (Figure 2D).

Next, we investigated whether HN enhances the lysosomal degradation of Aβ‐accumulated mitochondria by analyzing the colocalization of mitochondria with LAMP1 or FITC‐labeled Aβ. HN decreased the colocalization of mitochondria with Aβ and increased their colocalization with LAMP1 (Figure 2E,F). Bafilomycin A1 (Baf.A1) abolished HN‐induced removal of colocalized Aβ and mitochondria (Figure 2E). This suggests that HN‐induced Aβ removal might be mediated by autophago‐lysosomal degradation. Moreover, the colocalization between LAMP1 and mitochondria also notably increased with Baf.A1, indicating a potential accumulation of autophagosome‐engulfed mitochondria (Figure 2F).

Analysis of mitochondrial and cytoplasmic fractions revealed that HN reduced Aβ accumulation within mitochondria while promoting the translocation of LAMP1 from the cytosol to the mitochondrial fraction (Figure 2G,H). In the presence of Baf.A1, HN increased Aβ accumulation in both mitochondrial and cytosolic fractions, suggesting that HN enhances the clearance of Aβ through lysosomal degradation. Consistently, time‐dependent analysis revealed that HN transiently reduced mitochondrial mass‐related proteins, including TIM23, COX IV, and TOM20, at earlier time points, with restoration at 24 h (Figure S2A,B), indicating a dynamic and regulated turnover of mitochondria rather than sustained mitochondrial loss, supporting the notion that mitochondrial function is preserved.

We investigated whether HN affects AMPK‐ULK1 signaling related to the clearance of Aβ‐accumulated mitochondria. HN increases the phosphorylation levels of AMPK and its downstream target, acetyl‐CoA carboxylase (ACC), followed by phosphorylation of ULK1 and Beclin1 in ARPE19 cells (Figure 3A). These phosphorylations were entirely abrogated by preincubation with compound C (10 μM), an inhibitor of AMPK activation (Figure 3B). Genetic suppression of AMPK using RNA interference also substantially attenuated HN‐activated AMPK, ACC, ULK1, and Beclin1, demonstrating that HN induces AMPK‐dependent phosphorylations of ULK1 and Beclin1 (Figure 3C).

To further understand the dependency of HN's actions on AMPK, we examined the effect of pretreatment with compound C on Aβ‐induced ROS generation. As shown in Figure 3D,E, blocking AMPK activation with compound C effectively prevented Aβ‐triggered production of cytosolic ROS and mitochondrial superoxide. HN's abilities to increase the mitochondrial electrical gradient and to induce a hyperpolarizing response to oligomycin were blunted by co‐treatment with compound C (Figure 3F). Consistently, HN‐induced recovery of mitochondrial respiratory activities was abolished by compound C treatment (Figure 3G). These results suggest that HN induces AMPK‐dependent phosphorylation of ULK1 and Beclin1, which are responsible for the protective actions of HN against Aβ‐induced oxidative stress and mitochondrial dysfunction in the RPE.

We investigated whether HN‐induced phosphorylation of Beclin1 could facilitate the mitochondrial translocation of Parkin. To assess this, we analyzed the expression levels of Beclin1 and Parkin in mitochondrial and cytosolic fractions. As shown in Figure 4A,B, Aβ inhibited the mitochondrial translocation of both Beclin1 and Parkin, which was recovered by HN treatment. Imaging experiments confirmed that HN increased the mitochondrial localization of Parkin (Figure 4C) and Beclin1 (Figure 4D). Furthermore, the colocalization between Beclin1 and Parkin was also significantly increased by HN treatment (Figure 4E). Co‐immunoprecipitation studies revealed that Aβ suppressed the physical interaction between Beclin1 and Parkin, while HN restored their binding (Figure 4F).

We used siRNA to downregulate Parkin and Beclin1 and investigated their roles in the colocalization of LAMP1 or FITC‐labeled Aβ with mitochondria. Genetic suppression of Parkin using siRNA abolished HN‐induced augmentation of colocalization between LAMP1 and mitochondria. Similarly, Beclin1 knockdown reduced this colocalization, preventing HN‐stimulated mitochondria‐lysosome interaction. Time‐course analysis revealed that HN reduced Aβ‐mitochondria colocalization at 6 h, whereas no significant difference was observed at 24 h, suggesting progressive clearance of Aβ‐accumulated mitochondria at earlier time points (Figure S3A). LAMP1 recruitment to mitochondria appeared to remain elevated in cells treated with both Aβ and HN compared to Aβ alone throughout the time course, peaking at 6 h, reflecting continued lysosomal association with mitochondria (Figure S3B). Concurrent knockdown of Parkin and Beclin1 did not further inhibit colocalization beyond the effect of Beclin1 knockdown alone (Figure 5A). In addition, knockdown of Beclin1, either alone or in combination with Parkin, led to increased accumulation of Aβ in mitochondria, negating the action of HN (Figure 5B). However, knockdown of PINK1 did not affect HN‐induced mitochondria‐lysosome colocalization, LAMP1 recruitment to mitochondria, or the clearance of Aβ‐accumulated mitochondria (Figure S4A,B), indicating that HN‐mediated mitophagy is largely independent of the canonical PINK1 pathway. These findings suggest that HN's action on the degradation of Aβ‐accumulated mitochondria depends on both Parkin and Beclin1.

Immunoblot analysis for Aβ and LAMP1 in cytosolic and mitochondrial fractions, consistent with the immunostaining results, showed that HN‐induced removal of mitochondrial Aβ was decreased by Parkin downregulation, and further abolished by Beclin1 downregulation (Figure 5C). Moreover, downregulation of either Beclin1 or Parkin significantly hindered the mitochondrial translocation of the other protein (Figure 5D). Consistently, phosphorylated Beclin1 exhibited a similar pattern of change in relation to mitochondrial localization (Figure S5). Immunostaining results also demonstrated that mitochondrial localization of Parkin is markedly suppressed by Beclin1 knockdown, with no noticeable change observed in response to HN (Figure 5E). These results indicate that both Beclin1 and Parkin are required for their mitochondrial translocation and the recruitment of lysosome to mitochondria, leading to the removal of Aβ‐accumulated mitochondria. To further determine whether Beclin1‐dependent mitophagy contributes to mitochondrial functional recovery, we assessed mitochondrial functional parameters, including oxygen consumption rate and mitochondrial membrane potential (ΔΨm). HN treatment significantly increased maximal respiratory capacity, which was markedly attenuated upon Beclin1 knockdown. In addition, the recovery of mitochondrial membrane potential induced by HN was significantly reduced in Beclin1‐deficient cells. These findings provide functional evidence linking Beclin1‐dependent mitophagy to the restoration of mitochondrial function (Figure 5F,G).

To assess the role of HN in modulating Aβ accumulation and distribution, FITC‐labeled Aβ1‐42 (10 μM) was subretinally injected into mice. Three days later, the RPE exhibited degenerative changes, losing its characteristic cobblestone‐like morphology and disruption of ZO‐1, accompanied by intracellular localization of FITC‐labeled Aβ (Figure S6). Quantitative morphological analysis showed enlarged and irregular cells. Notably, Aβ also induced nuclear enlargement and multinucleation, consistent with the observations in primary marmoset RPE cells. Intravitreal co‐administration of HN effectively inhibited the Aβ‐induced disruption of the external blood‐retinal barrier, significantly restoring normal RPE morphology (Figure 6A). HN reduced the colocalization of Aβ with TOM20 while enhancing the colocalization of TOM20 and LAMP1 (Figure 6B), suggesting that HN enhances the removal of Aβ‐accumulated mitochondria.

The overall deposition of FITC‐labeled Aβ throughout the whole RPE was reduced by HN treatment (Figure 6C). Additionally, HN downregulated Aβ levels and upregulated LAMP1 expression across the RPE, preventing the decrease in ZO‐1 and F‐actin expression (Figure 6D). Electroretinography (ERG) measurements taken 3 days after subretinal Aβ injection showed subnormal amplitudes in both dark‐adapted and light‐adapted retinas. HN co‐treatment substantially improved these retinal responses, particularly under dark‐adapted conditions, improving retinal function (Figure 6E).

All animal procedures were approved by the Institutional Animal Care and Use Ethics Committee of Seoul National University (IACUC No. 231103–3), and all efforts were made to minimize animal suffering and reduce the number of animals used. Six‐week‐old male C57BL/6J mice (18–22 g, Orient Bio Inc., Sungnam, Korea) were housed in a pathogen‐free barrier environment throughout the study. The animals were anesthetized by intraperitoneal injection of a mixture of Zoletil 50 (50 mg/kg) and xylazine (5 mg/kg). Before the injection, the animals received tropicamide (Tropherin) eye drops for dilation. A step incision in the pars plana region of the sclera was made using a 26G needle, and subretinal (SR) and intravitreal (IVT) injections were carried out using a Hamilton syringe (Hamilton, Bonaduz, Switzerland) under a surgical microscope (Leica Microsystems Ltd., Wetzlar, Germany). The animals received an SR injection of 1 μL (10 μM) of oligomeric FITC‐labeled Aβ1‐42 (#50–194‐6621, Bachem) and an IVT injection of 1 μL of HN (20 μM) or PBS.

Three days after the injection, the animals were sacrificed, and their eyes were fixed by immersion in 4% paraformaldehyde for 15 min at room temperature. The RPE‐choroid‐scleral complexes (RCSs) were carefully separated from the eyeball using a dissecting microscope (Nikon SM2745T, Nikon Corporation, Japan) and incubated with Perm/Block solution (0.2% Triton‐X 100 and 0.3% BSA in PBS). Subsequently, they were incubated with primary antibodies in the same solution overnight at 4°C, followed by incubation with secondary antibodies for 1 h at room temperature in a dark room. Nuclei were counterstained with 1 μg/mL 4′,6′‐diamidino‐2‐phenylindole (DAPI, Sigma Aldrich) for 5 min at room temperature. After washing with PBS, the RCSs were mounted using Fluoromount Aqueous Mounting Medium (DAKO, Glostrup, Denmark) and observed under a confocal microscope (Leica TCS STED, Leica Microsystems Ltd., Wetzlar, Germany). For ARPE19 cells and primary marmoset RPE cells, fixation, permeabilization, blocking, and antibody incubation were performed as described above. All primary, conjugated, and secondary antibodies used in this study are listed in Table. S1

Six z‐stack images were taken in a clockwise direction from between the central and equatorial regions within a single RCS, as well as from differentiated ARPE19 cells and primary marmoset RPE cells, as previously described (Kim et al. 2021). For morphological analysis of individual cells in each image, we segmented RPE cells by extracting the image that captured cell borders stained with ZO‐1. Using a machine‐learning‐based ImageJ plugin (National Institutes of Health, Bethesda, MD, USA), Trainable Weka Segmentation, the image was binarized. Following a sequence of steps including despeckling, skeletonization, and removal of abnormally segmented cells, including those at the cut edge, we performed several morphometric analyses on the skeletonized image with thin cell borders. We calculated the morphological characteristics of each cell, including cell area, perimeter, aspect ratio (major axis length/minor axis length), and circularity ([4 × area × π]/perimeter (Biscetti et al. 2017)).

Colocalization analysis was performed on images after deconvolution and thresholding using ImageJ software (National Institutes of Health, Bethesda, MD, USA) on a minimum of 20 cells per slide and a total of at least 50 cells from 3 to 5 biologically independent experiments. Thresholding was applied to each image, and the adequacy of the threshold was manually validated by comparing with the original image. The same threshold value was applied to all images within the same experiment. The particle number was quantified using the ’analyze particles’ function in thresholded single sections with the size (pixel²) setting from 0.1 to 10 and circularity from 0 to 1. Cellular fluorescence intensity was expressed as the mean integrated density as a function of individual cell size. The percentage of colocalization was calculated in single z‐stack sections of deconvolved images (Turco et al. 2021).

Protein samples from cell lysates were precleared using 50 μL of packed protein A‐crosslinked 4% beaded agarose (Cell Signaling Technology, Beverly, MA, USA) at 4°C for 1 h. The beads were removed using centrifugation, and the supernatant was collected. The supernatant was incubated with Parkin (#39–0900, Invitrogen, 1:50), Beclin1 (#PA5‐96649, Invitrogen, 1:50), and PINK1 (#6946, Cell Signaling Technology, 1:50) antibodies at 4°C with constant mixing for 12 h. The immune complex was captured following the addition of packed agarose protein A beads (50 μL/500 μL supernatant) for 2 h at 4°C. Bead separation was achieved using centrifugation (13,000 × g for 25 min), and the supernatant was removed. Subsequently, the samples were subjected to immunoblot analysis with primary and appropriate secondary antibodies. The specificity of immunoprecipitation was confirmed using negative control reactions performed with a rabbit IgG or mouse IgG control without a primary antibody. The primary and secondary antibodies used in this study are listed in Table S1.

RCSs and ARPE19 cells were lysed in RIPA buffer (1% Triton X‐100) containing the protease inhibitor (GenDEPOT) and the phosphatase inhibitor (GenDEPOT). Additionally, cells were fractionated into mitochondrial and cytoplasmic components using a mitochondrial/cytoplasmic fractionation kit (ab65320, Abcam), according to the manufacturer's instructions. Briefly, cells were homogenized using a pestle homogenizer and then centrifuged at 700 × g for 10 min. The supernatant was re‐centrifuged at 10,000 × g for 30 min to collect the supernatant (cytoplasmic fraction) and pellet (mitochondrial fraction). The purity of the cytosolic and mitochondrial fractions was confirmed by immunoblot analysis using anti‐GAPDH and anti‐COX IV antibodies, respectively. Protein content was measured using the BCA protein assay kit (Pierce, Rockford, IL, USA). Proteins were separated using SDS‐PAGE and transferred onto polyvinylidene fluoride (PVDF) membranes (Merck Millipore Ltd., Tullagreen, Ireland). The PVDF membrane was blocked using non‐fat milk and incubated with TBST, followed by sequential incubation with primary and appropriate secondary antibodies. Signals were visualized with SuperSignal West Pico Chemiluminescent Substrate (Thermo Fisher Scientific). Immunoreactive bands were visualized using ImageQuantTMLAS4000 (GE Healthcare, USA) according to the manufacturer's instructions. Densitometric analysis was performed using ImageJ software, and the results were normalized to those of GAPDH. The primary and secondary antibodies used in this study are listed in Table S1.

The ARPE19 cell line obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA) was cultured in a 1:1 (vol/vol) mixture of Dulbecco's Modified Eagle Medium (DMEM) and Ham's F‐12 (Welgene, Daegu, Korea) containing 10% FBS (Gibco, Waltham, MA, USA) and 1% penicillin/streptomycin (Gibco). Primary RPE cells were isolated from the common marmoset (Callithrix jacchus), a small New World monkey, at the Marmoset Model Network Center in Seoul National University Hospital (IACUC No. 22‐0069), as previously described (Jang et al. 2023). Cells were cultured in DMEM supplemented with N1 supplement (1%; Sigma Aldrich), penicillin/streptomycin (1%; Gibco), nonessential amino acids (1%; Sigma Aldrich), taurine (0.25 g/L; Sigma Aldrich), hydrocortisone (20 μg/L; Sigma Aldrich), triiodothyronine (13 ng/L; Sigma Aldrich), and 10% FBS (Gibco). Both ARPE19 cells and primary marmoset RPE cells were cultured on Transwell membranes with 0.4 μm pores (Corning Costar) to facilitate differentiation for morphological analysis and TEER measurement. ARPE19 cells were maintained in DMEM containing 1% FBS for 3 months (Ahmado et al. 2011), while primary marmoset RPE cells were cultured in serum‐free medium supplemented with B27 (Gibco) for at least 4 weeks (Jang et al. 2023). Cells grown on Transwell permeable supports were maintained with 200 μL of media in the apical chamber and 700 μL in the basal chamber. Human induced pluripotent stem cells (iPSCs) were generated from human fibroblasts (ATCC CRL‐2097) by reprogramming with plasmid vectors encoding OCT4, SOX2, KLF4, and c‐MYC, as previously described (Kim et al. 2017). iPSC‐derived RPE cells were generated using a previously described differentiation protocol (Surendran et al. 2022). Cells were maintained at 37°C and 5% CO₂ and were regularly tested for mycoplasma.

Oligomeric Aβ1‐42 (# 50‐194‐6585, Bachem, 10 μM) with or without HN (20 μM) was added at the time points for immunoblot analysis and immunofluorescence. To assess the colocalization of mitochondria and Aβ by immunostaining, cells were treated with FITC‐labeled Aβ1‐42 (#50‐194‐6621, Bachem, 10 μM) with or without HN (20 μM). Bafilomycin A1 (10 nM; Sigma Aldrich) was used as a lysosomal function by selectively inhibiting vacuolar‐type H⁺‐ATPase (V‐ATPase), thereby preventing lysosomal acidification and blocking lysosomal degradation.

ARPE19 cells were transfected with Silencer Select siRNA targeting Parkin (ID: s502575; Invitrogen) or Beclin1 (IDL 137198; Invitrogen) using the Lipofectamine RNAiMAX reagent (Invitrogen) when the cells reached 60%–80% confluence, according to the manufacturer's instructions. Silencer Select Negative Control No. 1 siRNA (Invitrogen) served as the negative control. Briefly, 3 μL of RNAiMAX transfection reagent was mixed with 50 μL of Opti‐MEM medium (Thermo Fisher Scientific). Then 1 μL of siRNA (10 μM) was diluted with 50 μL of Opti‐MEM medium. The diluted siRNA was mixed with transfection reagent at room temperature for 5 min, and then added to ARPE19 cells. Following 24 h of transfection, cells were treated with Aβ (10 μM) with or without HN (20 μM).

Mice were anesthetized with an intraperitoneal injection of a mixture of Zoletil 50 (2.25 mg/kg) and xylazine (0.7 mg/kg) after dark adaptation for 16 h. Pupils were dilated with an eye drop of 0.5% tropicamide. Full‐field ERGs were recorded using the universal testing and electrophysiologic system 200 (UTAS E‐2000, LKC Technologies, Gaithersburg, MD, USA), as previously described (Gresh et al. 2003). The responses were recorded at 2 K gain, using a 69 Hz notch filter and bandpass filtering between 0.1 and 1500 Hz. In the light‐adapted state (photopic), cone responses were isolated and recorded in response to a single flash at 0 dB and a flicker sequence of 30 Hz with 30 cd/m² background illumination to desensitize the rods. A‐wave amplitude was measured from baseline to the lowest negative voltage, while peak b‐wave amplitude was measured from the lowest point of the a‐wave to the highest peak of the positive b‐wave.

TEER measurements were taken using an EVOM3 epithelial volt‐ohmmeter and STX4 electrode (World Precision Instruments, Sarasota, FL, USA) according to the manufacturer's instructions. The electrodes were sterilized with 70% ethanol, rinsed in ddH₂O, and equilibrated in pre‐warmed culture medium. To minimize the influence of temperature, measurements were performed in the hood within 5 min after the cells were taken out of the incubator. Measurements were repeated in five independent experiments, with at least three separate wells recorded per experiment to obtain an average value. Net TEER (Ω·cm²) was calculated by subtracting the value of a blank insert from the experimental value and multiplying it by the area of the insert membrane (0.33 cm² for 25‐well insert).

For live‐cell imaging, ARPE19 cells were seeded in glass bottom dishes (Nunc, Thermo Fisher Scientific) at a density of 5 × 10⁴ cells per dish in a growth medium. After an overnight incubation, cells were treated with Aβ (10 μM) with or without HN (20 μM) in DMEM/F‐12 without phenol red (Welgene) containing 10% FBS (Gibco) and 1% penicillin/streptomycin (Gibco). Cells were stained with LysoTracker Green (#8783, Cell Signaling Technology, 50 nM) and MitoTracker Red (#9082, Cell Signaling Technology, 50 nM) for 30 min in a humidified CO₂ incubator and observed under a confocal microscope (Leica STED, Leica Microscope Ltd., Wetzlar, Germany) for 1 h. Z‐stacks ranging from 80 to 100 μM were taken with a step size of 4–5 μM at intervals of 3 min. Temperature and CO₂ levels were controlled with a stage‐top incubator.

To measure intracellular reactive oxygen species (ROS) production, ARPE‐19 cells were incubated with chloromethyl‐H2‐dichlorofluorescein diacetate (CM‐H2DCF‐DA), a cell‐permeable, non‐fluorescent probe. Upon entering the cell, intracellular esterases cleave the diacetate groups, and the chloromethyl group facilitates intracellular retention via binding to glutathione and thiol‐containing proteins. The deacetylated probe is then rapidly oxidized by intracellular ROS to produce the highly fluorescent 2,7‐dichlorofluorescein (DCF), detected at excitation/emission wavelengths of 485/535 nm. Briefly, cultured ARPE‐19 cells, treated with or without TGF‐β1 and specific inhibitors as specified, were exposed to a 5 μM concentration of CM‐H2DCF‐DA for 20 min at 37°C. Following incubation, the cells were washed with Krebs‐Ringer bicarbonate (KRB) solution (135 mM NaCl, 3.6 mM KCl, 2 mM NaHCO3, 0.5 mM NaH2PO4, 0.5 mM MgSO4, 1.5 mM CaCl2, and 10 mM HEPES; pH 7.4). Fluorescence images (excitation/emission: 490/535 nm) were captured using an IX81 fluorescence microscope (Olympus, Tokyo, Japan) with a confocal spinning disk, and fluorescence intensity was quantified using Metamorph 6.1 software (Molecular Devices, Sunnyvale, CA). for the selective detection of mitochondrial superoxide, ARPE‐19 cells were treated with MitoSOX Red mitochondrial superoxide indicator. MitoSOX reagent was prepared to a 5 μM concentration in KRB solution and used to treat ARPE‐19 cells for 15 min at 37°C. Fluorescence intensity was then measured by confocal microscopy and analyzed using MetaMorph software.

An Extracellular Flux Analyzer XF‐96 (Seahorse Bioscience, Agilent, Santa Clara. CA, USA) was used to determine cellular oxygen consumption rate (OCR) in the cells XF Base Media was used as an assay medium including glucose 22 mM, pyruvate I mM and glutamine 2 mM, which was used to incubate with cells for 45–60 min before basal measurement. After 20 min of measurement for the baseline, the effect of oligomycin (2uM), an ATP synthase inhibitor that blocks mitochondrial ATP production, FCCP (2 μM), rotenone/antimycin A (0.5 μM/1.5 μM) were recorded during OCR measurement.

Mitochondrial membrane potentials (Δψm) were measured using 5,5′,6,6′‐tetrachloro‐1,1′,3,3′‐tetraethylbenzimidazolylcarbocyanine iodide (JC‐1, catalog no. T3168, Molecular Probes, Thermo Fisher Scientific), a lipophilic cationic dye. Due to its positive charge and lipophilic nature, JC‐1 monomers accumulate in mitochondria where the membrane potential is highly negative (approximately −150 to −200 mV), causing them to aggregate into polymers. JC‐1 monomers emit green fluorescence (excitation at 490 nm, emission at 535 nm), while the polymers emit red fluorescence (excitation at 540 nm, emission at 590 nm). The fluorescence signals were recorded using a fluorescence microplate reader (FlexStation III, Molecular Devices). The Δψm was estimated by calculating the ratio of red to green fluorescence (590 nm/535 nm).

The results are presented as the mean ± SD. One‐way ANOVA followed by Tukey's post hoc test was used for comparisons among three or more groups, and unpaired two‐tailed Student's t‐test was used for comparisons between two groups. No statistical methods were used to predetermine the sample size. Mice were randomly allocated to experimental groups. No blinding method was used for SR and IVT injection. There were no animal exclusion criteria. The variance was similar between groups that were statistically compared. One‐way ANOVA and Student's t‐test analysis were performed using GraphPad Prism (San Diego, CA, USA) version 10.0 software. Data was considered statistically significant at p < 0.05.