Oxidized-LDL Induces Metabolic Dysfunction in Retinal Pigment Epithelial Cells

Abstract

Recently, mitochondrial dysfunction has gained attention as a causative factor in the pathogenesis and progression of age-related macular degeneration (AMD). Mitochondrial damage plays a key role in metabolism and disrupts the balance of intracellular metabolic pathways, such as oxidative phosphorylation (OXPHOS) and glycolysis. In this study, we focused on oxidized low-density lipoprotein (ox-LDL), a major constituent of drusen that accumulates in the retina of patients with AMD, and investigated whether it could be a causative factor for metabolic alterations in retinal pigment epithelial (RPE) cells. We found that prolonged exposure to ox-LDL induced changes in fatty acid β-oxidation (FAO), OXPHOS, and glycolytic activity and increased the mitochondrial reactive oxygen species production in RPE cells. Notably, the effects on metabolic alterations varied with the concentration and duration of ox-LDL treatment. In addition, we addressed the limitations of using ARPE-19 cells for retinal disease research by highlighting their lower barrier function and FAO activity compared to those of induced pluripotent stem cell-derived RPE cells. Our findings can aid in the elucidation of mechanisms underlying the metabolic alterations in AMD.

INTRODUCTION

Age-related macular degeneration (AMD) is a disease characterized by the deterioration of the macula in the retina, leading to the loss of central vision and distortion of the visual field.¹⁾ AMD ranks as the fourth leading cause of blindness worldwide in 2020.²⁾ Accumulation of drusen in the subretinal pigment epithelial space is characteristic of early-stage AMD. As the disease advances, it irreversibly transitions to dry AMD (d-AMD), which is characterized by geographic atrophy and loss of retinal pigment epithelial (RPE) and photoreceptor (PR) cells. Effective treatment methods for d-AMD are scarce, necessitating the elucidation of its disease progression mechanisms.

Among the cells constituting the retina, RPE cells are the primary site of AMD pathology, and RPE cell dysfunction is suggested to contribute to the pathogenesis of AMD.³⁾ RPE cells play key roles in maintaining the normal function of the retina, providing nutrients to PR cells, and acting as the blood-retinal barrier via the formation of intercellular tight junctions. Recently, mitochondrial dysfunction of RPE cells has been recognized as a contributing factor to the onset and progression of d-AMD.^4–6) Mitochondria are the primary site of cellular energy production, fatty acid β-oxidation (FAO), tricarboxylic acid cycle, and oxidative phosphorylation (OXPHOS). Hence, mitochondrial dysfunction is suggested to alter these metabolic activities in AMD. Fluctuations in the balance between OXPHOS and glycolytic activity are observed in the primary RPE cells derived from AMD donors.^7,8) However, the specific mechanisms underlying these metabolic alterations remain unclear.

In this study, we focused on lipoproteins as the causative factors of metabolic changes. Low-density lipoprotein (LDL), a type of lipoprotein, is responsible for cholesterol and lipid transport; it becomes a source of fatty acids, the substrate for FAO, when taken up by RPE cells. LDL is rich in polyunsaturated fatty acids⁹⁾ and easily oxidized. Oxidized (Ox)-LDL, generated via the oxidative modification of LDL, accumulates in the drusen¹⁰⁾ and choroidal neovascular membranes¹¹⁾ of patients with AMD. Moreover, plasma ox-LDL concentrations are increased in patients with AMD.^12,13) These studies suggest that ox-LDL is associated with the onset and progression of AMD.

The persistence of chronic oxidative stress due to aging has been proposed as a risk factor for the onset of AMD.^14,15) To date, studies investigating the effects of oxidative stress on cultured RPE cells have focused on transient oxidative stress. However, these experimental conditions do not accurately reflect the pathophysiology of AMD, in which the cells are chronically exposed to oxidative stress. Therefore, in this study, we investigated whether chronic ox-LDL treatment induces metabolic alterations in mature RPE cells.

MATERIALS AND METHODS

Reagents

Most reagents were purchased from FUJIFILM Wako Pure Chemical Corporation (Osaka, Japan) or Nacalai Tesque (Kyoto, Japan). LDL (360-10) and ox-LDL (360-31) were purchased from Medix Biochemica U.S.A. (St. Louis, MO, U.S.A.). Ox-LDL was produced by oxidizing LDL with copper sulfate. The oxidation of LDL was validated by electrophoresis. MitoROS 580 was purchased from AAT Bioquest (Pleasanton, CA, U.S.A.; 16052).

Cell Culture

Induced pluripotent stem cell-derived retinal pigment epithelial (iPSC-RPE) cells (FUJIFILM Wako Pure Chemical Corporation) were seeded in a 6-well plate containing a 1 : 1 mixture of SFRM-B27 and PA6 media. SFRM-B27 medium contained Dulbecco’s modified Eagle’s medium (DMEM)-low glucose (Nacalai Tesque), 30% F-12 (Nacalai Tesque), 2% B-27 Supplement (50×; Thermo Fisher Scientific, Waltham, MA, U.S.A.), 100 u/mL penicillin, 100 µg/mL streptomycin (Nacalai Tesque), and 2 mM L-glutamine (Nacalai Tesque). PA6 medium contained DMEM/Ham’s F-12 (Nacalai Tesque), 10% fetal bovine serum (FBS), 100 u/mL penicillin, and 100 µg/mL streptomycin (Nacalai Tesque). After the cells were attached, the medium was changed to the SFRM-B27 maintenance medium with 0.5 µM SB431542 (transforming growth factor β (TGFβ) receptor kinase inhibitor; Nacalai Tesque) and 10 ng/mL basic fibroblast growth factor (FUJIFILM Wako Pure Chemical Corporation). The cell culture was expanded through one passage in maintenance medium, and the cells were stored in liquid N₂ using Cell Reservoir One (Nacalai Tesque). For each experiment, the stocked cells were thawed and seeded with the 1 : 1 mixed medium, after which the medium was changed to the maintenance medium every two days until the cells were used.¹⁶⁾

Adult retinal pigment epithelial cell line-19 (ARPE-19; American Type Culture Collection) cells were cultured in DMEM/Ham’s F-12 (Nacalai Tesque) supplemented with 1% FBS, 100 u/mL penicillin, and 100 µg/mL streptomycin (Nacalai Tesque) for two weeks before experiments.

Measurement of Transepithelial Electrical Resistance (TER)

iPSC-RPE (8.0 × 10⁴ cells/well) or ARPE-19 (1.0 × 10⁵ cells/well) cells were seeded into Transwell inserts (0.33 cm²; Corning, Corning, NY, U.S.A.) coated with laminin (FUJIFILM Wako Pure Chemical Corporation). Their barrier function was measured using EVOM3 and STX2-PLUS (World Precision Instruments, Sarasota, FL, U.S.A.), according to the manufacturer’s instructions. The values for the laminin-coated cell-free inserts were defined as the background, and individual resistances were calculated using the following equation¹⁷⁾:

  

Immunofluorescence Staining

iPSC-RPE (8.0 × 10⁴ cells/well) or ARPE-19 (1.0 × 10⁵ cells/well) cells were seeded in a laminin-coated 96-well plate. The cells were cultured for two weeks and treated with each reagent. After washing thrice with phosphate-buffered saline (PBS), the cells were fixed with 4% paraformaldehyde-phosphate buffer (163-20145; FUJIFILM Wako Pure Chemical Corporation) for 15 min at room temperature. After washing thrice with PBS, the cells were blocked with a blocking solution (PBS with 0.3% Triton X-100 containing 5% goat serum; 005-000-121; Jackson ImmunoResearch, West Grove, PA, U.S.A.) for 30 min at room temperature. After washing thrice with PBS, the cells were further incubated with a polyclonal anti-zonula occludens (ZO)-1 antibody (1 : 50; 61-7300; Invitrogen, Waltham, MA, U.S.A.) dissolved in a blocking solution for 1.5 h at room temperature. After washing thrice with PBS, the cells were incubated with Alexa Fluor 488 AffiniPure Goat Anti-Rabbit immunoglobulin G (IgG) (1 : 200; 111-545-003; Jackson ImmunoResearch) dissolved in a blocking solution for 30 min at room temperature. Then, after washing thrice with PBS, the cells were sealed in the VECTASHIELD PLUS Antifade Mounting Medium with 4′-6-diamidino-2-phenylindole (DAPI) (H-2000; Vector Laboratories, Newark, CA, U.S.A.), and fluorescence images were captured using the BZ-X800 Analyzer (KEYENCE, Osaka, Japan) with a GFP filter (Ex/Em = 470/525 nm).

Lipoprotein Treatment

iPSC-RPE (8.0 × 10⁴ cells/well) or ARPE-19 (1.0 × 10⁵ cells/well) cells were cultured for two weeks, serum-starved for 24 h, and treated with (ox-)LDL for 24 h or 1 week (the medium was replaced with a fresh medium containing (ox-)LDL on day 4).

Cell Viability Assay

iPSC-RPE cells were seeded at a density of 8.0 × 10⁴ cells/well in a 96-well plate, cultured for two weeks, and treated with each reagent. After stimulation, 15 µL of 5 mg/mL 3-(4,5-dimethylthiazol-2-yl)-2-5-diphenyltetrazolium bromide (MTT) reagent (23547-21; Nacalai Tesque) was added to each well and incubated at 37 °C and 5% CO₂ for 2 h. After aspirating the medium, 100 µL of dimethyl sulfoxide was added to each well, and the absorbance was measured at 570 and 650 nm using the EnSpire Multimode Plate Reader (PerkinElmer, Inc., Waltham, MA, U.S.A.). Then, cell viability of each reagent-treated group was calculated by converting the absorbance of the control group to 100% viability.

Fluorescence Detection of FAO

iPSC-RPE (8.0 × 10⁴ cells/well) or ARPE-19 (1.0 × 10⁵ cells/well) cells were seeded in a 96-well plate, cultured for two weeks, and treated with each reagent. FAOBlue, a fluorescent probe used to measure FAO activity, was kindly provided by A. Ojida and S. Uchinomiya (Kyushu University, Japan). Additionally, 5-aminoimidazole-4-carboxamide riboside (AICAR; AG-CR1-0061-M010; AdipoGen) was used as a positive control to increase the FAO activity. After washing twice with Hank's balanced salt solution (+) (HBSS (+); 17459-55; Nacalai Tesque), the cells were incubated with 5 µM FAOBlue at 37 °C and 5% CO₂ for 30 min. Fluorescence of FAOBlue was detected at a specific wavelength (Ex/Em = 405/460 nm) using the BZ-X800 Analyzer (KEYENCE). Then, fluorescence images were quantified using the Fiji software.

RNA Extraction and Quantitative RT-PCR Analysis

iPSC-RPE cells were seeded at a density of 8 × 10⁴ cells/well in a 96-well plate and cultured for two weeks. After treatment with ox-LDL, total RNA was extracted using ISOSPIN Cell & Tissue RNA (314-08211; NIPPON GENE, Tokyo, Japan), according to the manufacturer’s instructions. RNA content was measured using the NanoDrop One UV/visible spectrophotometer (Thermo Fisher Scientific). Then, RNA was reverse-transcribed into cDNA using the ReverTra Ace qPCR RT Master Mix (FSQ-201; TOYOBO, Osaka, Japan). Then, THUNDERBIRD SYBR qPCR Mix (QPS-201; TOYOBO) was used for DNA amplification and fluorescence emission in cDNA samples. Subsequently, qPCR was performed using the CFX Connect Real-Time PCR Detection System (Bio-Rad, Hercules, CA, U.S.A.), according to the manufacturer’s protocol. Relative gene expression levels were calculated using the ΔCq method with glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as the internal control. All primers were synthesized by Eurofins Genomics. All primer sequences used in this study are listed in Table 1.

Table 1. Primers (5′–3′) Used for RT-qPCR Analysis

Gene	Primer	Sequence
GAPDH	Forward	CGTGGAAGGACTCATGACCA
	Reverse	CATCACGCCACAGTTTCCC
CPT1A	Forward	CGTCTTTTGGGATCCACGATT
	Reverse	TGTGCTGGATGGTGTCTGTCTC
CPT2	Forward	TGCCATCCACTTTGAGCACT
	Reverse	GGGGTCTGAGTGCTGTCTTT
ACADM	Forward	ATGCCCTGGAAAGGAAAACT
	Reverse	AACCTCCCAAGCTGCTCTCT
ACADL	Forward	TTGGCAAAACAGTTGCTCAC
	Reverse	ACATGTATCCCCAACCTCCA
PPARA	Forward	GATGCGCTGACAGATGGAGA
	Reverse	TAGAGACGGCTCTTCTGCCT
HK	Forward	GGCTCATTTCCACCTCACCA
	Reverse	GGAGGGCAGCATCTTAACCA
PFKFB3	Forward	TTGGCGTCCCCACAAAAGT
	Reverse	AGTTGTAGGAGCTGTACTGCTT
PKM2	Forward	CAAAGGACCTCAGCAGCCATGTC
	Reverse	GGGAAGCTGGGCCAATGGTACAGA
LDHA	Forward	GGCCTGTGCCATCAGTATCT
	Reverse	GGAGATCCATCATCTCTCCC

Seahorse Metabolic Analysis

Oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) were measured using the XFp Extracellular Flux Analyzer kindly provided by T. Ide, M. Ikeda, and M. Watanabe (Kyushu University, Japan). iPSC-RPE cells were seeded in a Seahorse XFp Cell Culture Mini plate (Agilent Technologies, Santa Clara, CA, U.S.A.) at a density of 3.2 × 10⁴ cells/well. After treatment with (ox-)LDL, the cells were washed twice with and incubated in the XF Base Medium with 10 mM glucose, 1 mM pyruvate, and 2 mM glutamine for OCR or with 2 mM glutamine for ECAR at 37 °C and 5% CO₂ for 1 h. After injecting 1.5 µM oligomycin (Oligo), 2 µM carbonyl cyanide-4-[trifluoromethoxy]phenylhydrazone (FCCP), and 2 µM rotenone and antimycin A (Rot/AA) into the Seahorse XFp Extracellular Flux Cartridge (Agilent Technologies), OCR was measured. ECAR was measured with 10 mM glucose (Glc), 2 µM oligomycin, and 100 mM 2-deoxy-D-glucose (2-DG).

Measurement of Cellular ATP Levels

iPSC-RPE cells were seeded at a density of 8 × 10⁴ cells/well in a 96-well plate and cultured for 2–3 weeks. After treatment with ox-LDL, the cellular ATP levels were measured using CellTiter-Glo 2.0 Assay (G9241; Promega, Madison, WI, U.S.A.), according to the manufacturer’s instructions.

Measurement of Mitochondrial Reactive Oxygen Species (MitoROS) Levels

MitoROS 580 stock solution (1000×) was prepared, according to the manufacturer’s instructions. iPSC-RPE cells were seeded at a density of 8 × 10⁴ cells/well in a 96-well plate and cultured for two weeks. After treatment with each reagent and washing twice with HBSS (+), 100 µL of MitoROS 580 working solution (1×) diluted in HBSS (+) was added to each well and incubated at 37 °C and 5% CO₂ for 30 min. Tert-butyl hydroperoxide (tBHP) was used as a positive control for oxidative stimulation. After washing thrice with HBSS (+), the medium was replaced with HBSS (+) for fluorescence microscopy. Fluorescence images were acquired using the BZ-X800 Analyzer (KEYENCE) with a TRITC filter (Ex/Em = 545/605 nm). Finally, fluorescence images were quantified using the Fiji software.

Statistical Analyses

Data are expressed as the mean + standard deviation (S.D.). Statistical analyses were conducted using the Student’s t-test or one-way ANOVA followed by Tukey’s multiple comparison tests. Statistical significance was set at p < 0.05. GraphPad Prism software (version 9.0; GraphPad Software, San Diego, CA, U.S.A.) was used for all analyses.

RESULTS

Cell Morphology and Barrier Functions Differ between iPSC-RPE and ARPE-19 Cells

Several types of RPE cells, such as human iPSC-RPE and ARPE-19 cells, are commonly used for in vitro experiments.¹⁸⁾ Here, we compared their utility as RPE models.

In general, cultured RPE cells mature and exhibit epithelial cell characteristics similar to those of physiological RPE cells after two or more weeks in culture.^17,19) Indeed, after two weeks of maturation, iPSC-RPE cells exhibited a uniform hexagonal shape, characteristic of RPE cells, with the deposition of melanin pigments (Fig. 1a). In contrast, ARPE-19 cells exhibited irregular shapes, including spindle shape, with no pigmentation. Immunofluorescence staining for ZO-1, an indicator of tight junction formation in epithelial tissues, revealed its continuous expression in iPSC-RPE cells. In contrast, ZO-1 expression was low and intercellular adhesion was discontinuous in ARPE-19 cells (Fig. 1a). TER, a measure of barrier function, exceeded 200 Ω·cm² in iPSC-RPE cells and was comparable to the reported TER of human RPE cells (150–200 Ω·cm²).^20,21) However, the TER of ARPE-19 cells was approximately 60 Ω·cm² (Fig. 1b).

Fig. 1. Barrier Functions and Fatty Acid β-Oxidation (FAO) Activities of Retinal Pigment Epithelial (RPE) Cells

(a) Bright-field and zonula occludens (ZO)-1 immunofluorescence staining images of induced pluripotent stem cell-derived RPE (iPSC-RPE) and adult RPE cell line-19 (ARPE-19) cells matured for two weeks. Scale bar, 50 µm. (b) Transepithelial electrical resistance (TER) of each cell seeded in the Transwell inserts (0.33 cm²) and matured for two weeks. Mean + standard deviation (S.D.). n = 3. *** p < 0.001 via Student’s t-test. (c) iPSC-RPE cells were cultured for two weeks. Then, oxidized low-density lipoprotein (ox-LDL) was added twice a week at respective concentrations. After one week, 3-(4,5-dimethylthiazol-2-yl)-2-5-diphenyltetrazolium bromide (MTT) assay was performed to evaluate its cytotoxicity. Mean + S.D. n = 3. *** p < 0.001 via one-way ANOVA followed by Tukey’s multiple comparison test. (d) Left: iPSC-RPE cells were cultured in the presence of ox-LDL for one week. To evaluate the effect of 5-aminoimidazole-4-carboxamide riboside (AICAR), cells were pretreated with 500 µM AICAR for 3 h. Treated cells were stained with FAOBlue, and FAO activity was analyzed via fluorescence microscopy. Scale bar, 150 µm. Right: Quantified data from fluorescence images. Mean fluorescence intensity is shown as a fold-change. Mean + S.D. n = 3. * p < 0.05 and ** p < 0.01 via one-way ANOVA followed by Tukey’s multiple comparison test. (e) FAO activity of ARPE-19 cells treated with ox-LDL for 24 h or 500 µM AICAR for 3 h (positive control). Scale bar, 150 µm. (f) Bright-field images for (e). Scale bar, 150 µm.

These results indicate the differences in the cell morphology and barrier functions of iPSC-RPE and ARPE-19 cells cultured for two weeks. Interestingly, ARPE-19 cells did not fully exhibit the characteristics of physiological RPE cells.

Metabolic Activities Are Different between iPSC-RPE and ARPE-19 Cells

Mitochondrial FAO is a major pathway for fatty acid degradation and a source of energy production in RPE cells.²²⁾ Therefore, we analyzed the effect of oxidation of LDL, a supplier of fatty acids, on FAO activity in this study. To mimic chronic oxidative stress, mature iPSC-RPE cells cultured for two weeks were treated twice with ox-LDL on days 1 and 4, and the cells were evaluated on day 7. The addition of ox-LDL at concentrations below 50 µg/mL did not affect the viability of iPSC-RPE cells (Fig. 1c). Moreover, the addition of 25 µg/mL ox-LDL increased the FAO activity, as measured by the FAOBlue probe,²³⁾ whereas the addition of 50 µg/mL ox-LDL did not increase the FAO activity (Fig. 1d). AICAR, an AMP-activated protein kinase activator, was used as a positive control for FAO activity. Compared to iPSC-RPE cells, when treated with ox-LDL for 24 h, mature ARPE-19 cells showed weak FAOBlue fluorescence regardless of ox-LDL treatment. Evaluation of FAO activity in ARPE-19 cells was difficult using the FAOBlue probe (Figs. 1e, f). Therefore, we evaluated the metabolic activity in iPSC-RPE cells hereafter.

Chronic Exposure to Ox-LDL Changes the Metabolic Profile of RPE Cells

Next, we measured the mRNA levels of FAO-related genes, including carnitine palmitoyl-transferase 1A (CPT1A) and carnitine palmitoyl-transferase 2 (CPT2), which are responsible for the transport of fatty acids into mitochondria, and acyl-CoA dehydrogenase medium chain (ACADM) and acyl-CoA dehydrogenase long chain (ACADL), which are enzymes involved in the initial steps of FAO, using RT-qPCR. Additionally, we assessed the transcriptional regulation of FAO-related genes by measuring the mRNA levels of peroxisome proliferator-activated receptor alpha (PPARA). Addition of 25 µg/mL ox-LDL significantly increased the expression levels of several mRNAs, whereas the addition of 50 µg/mL ox-LDL reversed this effect (Fig. 2a).

Fig. 2. Effects of Chronic Ox-LDL Treatment on the Metabolic Activities of RPE Cells

(a) iPSC-RPE cells were treated with ox-LDL for one week. Then, the expression levels of FAO-related genes were determined using RT-qPCR. The expression of each gene was normalized to that of glyceraldehyde 3-phosphate dehydrogenase (GAPDH). (b, c) iPSC-RPE cells were cultured in the XFp Cell Culture Miniplate for two weeks, followed by treatment with ox-LDL for one week. Oxygen consumption rate (OCR) (b) and extracellular acidification rate (ECAR) (c) were measured using the XFp Extracellular Cell Flux Analyzer. Bar graphs show the quantified data for OCR (basal, ATP-linked, and maximal respiration) (b) and ECAR (glycolysis and glycolytic capacity) (c). (d) iPSC-RPE cells were treated with ox-LDL for one week. Then, the expression levels of glycolysis-related genes were determined using RT-qPCR. The expression of each gene was normalized to that of GAPDH. (e) iPSC-RPE cells were treated with ox-LDL for one week. Then, the cellular ATP levels were measured with a luciferase assay. Mean + S.D.. n = 3 (a–d), n = 4 (e). * p < 0.05, ** p < 0.01, and *** p < 0.001 via one-way ANOVA followed by Tukey’s multiple comparison test.

As long-term stimulation with ox-LDL for one week caused fluctuations in FAO activity, we evaluated the OCR, an indicator of OXPHOS activity, using an extracellular flux analyzer. Basal, ATP-linked, and maximal respiration were significantly decreased (Fig. 2b), indicating a reduction in OXPHOS activity.

Treatment of macrophages with ox-LDL inhibits OXPHOS and increases the glycolytic activity via a compensatory switch to glycolysis.²⁴⁾ Here, we assessed the glycolytic activity of ox-LDL-treated iPSC-RPE cells using an extracellular flux analyzer to measure ECAR. iPSC-RPE cells stimulated with 50 µg/mL ox-LDL showed enhanced glycolysis (Fig. 2c), suggesting a metabolic shift from OXPHOS to glycolysis. Moreover, mRNA expression levels of glycolytic genes, including hexokinase (HK), 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase-3 (PFKFB3), pyruvate kinase M2 (PKM2), and lactate dehydrogenase A subunit (LDHA), were decreased under these conditions (Fig. 2d).

Since ox-LDL altered OXPHOS and glycolytic activities, the cellular ATP levels were assessed using a luciferase assay. The results indicated that chronic treatment with 50 µg/mL ox-LDL led to a reduction in ATP levels in iPSC-RPE cells (Fig. 2e).

These results indicate that prolonged exposure to ox-LDL induced metabolic alterations in RPE cells. Furthermore, 25 µg/mL ox-LDL enhanced the FAO activity, whereas 50 µg/mL ox-LDL triggered a shift toward glycolysis, suggesting a considerable difference in metabolic effects based on the amount of ox-LDL added.

Acute Exposure to Ox-LDL Does Not Alter the Metabolic Profile of Cells

Next, we investigated the effects of acute ox-LDL exposure for 24 h on cells to determine the timing of metabolic changes (Fig. 3). Treatment with 50 µg/mL ox-LDL significantly decreased the ATP-linked respiration (Fig. 3b). Notably, ECAR, mRNA levels of genes related to FAO and glycolysis, and ATP content were not affected by the addition of either 25 or 50 µg/mL ox-LDL (Figs. 3a, c–e).

Fig. 3. Effects of Acute Ox-LDL Treatment on the Metabolic Activities of RPE Cells

(a) iPSC-RPE cells were treated with ox-LDL for 24 h. Then, the expression levels of FAO-related genes were determined using RT-qPCR. The expression of each gene was normalized to that of GAPDH. (b, c) iPSC-RPE cells were cultured in the XFp Cell Culture Miniplate for two weeks, followed by treatment with ox-LDL for 24 h. OCR (b) and ECAR (c) were measured using the XFp Extracellular Cell Flux Analyzer. Bar graphs show the quantified data for OCR (basal, ATP-linked, and maximal respiration) (b) and ECAR (glycolysis and glycolytic capacity) (c). (d) iPSC-RPE cells were treated with ox-LDL for 24 h. Then, the expression levels of glycolysis-related genes were determined using RT-qPCR. The expression of each gene was normalized to that of GAPDH. (e) iPSC-RPE cells were treated with ox-LDL for 24 h. Then, the cellular ATP levels were measured with a luciferase assay. Mean + S.D. n = 3 (a–d), n = 4 (e). Mean + S.D. n = 3. ** p < 0.01 via one-way ANOVA followed by Tukey’s multiple comparison test.

Ox-LDL Induces MitoROS Production in RPE Cells

Mitochondria play a vital role in energy metabolism and are a major source of reactive oxygen species (ROS) in RPE cells. Here, we examined whether ox-LDL affects MitoROS production in RPE cells. Mature iPSC-RPE cells were subjected to long- or short-term stimulation with 25 or 50 µg/mL ox-LDL. tBHP was used as a positive control for oxidative stimulation. We found that ox-LDL increased MitoROS production after both long- and short-term stimulation (Fig. 4).

Fig. 4. Reactive Oxygen Species (ROS) Production in the Mitochondria of Ox-LDL-Treated RPE Cells

(a, b) Left: iPSC-RPE cells were treated with ox-LDL for one week (a) or 24 h (b). As a positive control, cells were treated with 200 µM tBHP for 3 h. Treated cells were then stained with MitoROS 580, and mitochondrial superoxide levels were analyzed via fluorescence microscopy. Right: Quantified data from the fluorescence images. Mean fluorescence intensity is shown as a fold-change. Scale bar, 150 µm. Mean + S.D. n = 3. * p < 0.05, ** p < 0.01, and *** p < 0.001 via one-way ANOVA followed by Tukey’s multiple comparison test.

LDL Affects Metabolic Pathways in Different Ways

Finally, we examined the effects of unoxidized LDL on the metabolic pathway. iPSC-RPE cells cultured for two weeks were treated twice with LDL, similar to ox-LDL treatment. The addition of LDL did not affect the cell viability (Fig. 5a). FAO activity did not change with 50 µg/mL LDL (Fig. 5b). On the other hand, OCR was decreased and ECAR was increased by LDL similar to ox-LDL (Figs. 5c, d). In addition, MitoROS levels did not vary with 50 µg/mL LDL (Fig. 5e).

Fig. 5. Effects of Chronic LDL Treatment on the Metabolic Activities of RPE Cells

(a) iPSC-RPE cells were cultured for two weeks. Then, low-density lipoprotein (LDL) was added twice a week at respective concentrations. After one week, MTT assay was performed to evaluate its cytotoxicity. (b) Left: iPSC-RPE cells were cultured in the presence of 50 µg/mL LDL for one week. Treated cells were stained with FAOBlue, and FAO activity was analyzed via fluorescence microscopy. Right: Quantified data from fluorescence images. Mean fluorescence intensity is shown as a fold-change. Scale bar, 150 µm. (c, d) iPSC-RPE cells were cultured in the XFp Cell Culture Miniplate for two weeks, followed by treatment with 50 µg/mL LDL for one week. OCR (c) and ECAR (d) were measured using the XFp Extracellular Cell Flux Analyzer. Bar graphs show the quantified data for OCR (basal, ATP-linked, and maximal respiration) (c) and ECAR (glycolysis and glycolytic capacity) (d). (e) Left: iPSC-RPE cells were treated with 50 µg/mL LDL for one week. Treated cells were then stained with MitoROS 580, and mitochondrial superoxide levels were analyzed via fluorescence microscopy. Right: Quantified data from the fluorescence images. Mean fluorescence intensity is shown as a fold-change. Scale bar, 150 µm. Mean + S.D. n = 3. * p < 0.05, ** p < 0.01, and *** p < 0.001 via one-way ANOVA followed by Tukey’s multiple comparison test.

DISCUSSION

In this study, we showed that, to assess the functionality of RPE cells, it is desirable to use mature iPSC-RPE cells cultured for two weeks to exhibit the required physiological characteristics. Furthermore, we showed that the effects on metabolic alterations varied depending on the concentration and duration of ox-LDL stimulation. Specifically, exposure to 50 µg/mL ox-LDL for 24 h decreased the OXPHOS levels (Fig. 3b), whereas stimulation with 25 µg/mL ox-LDL for one week increased the FAO activity (Fig. 1d). In addition, treatment with 50 µg/mL ox-LDL for one week increased the glycolytic activity (Fig. 2c).

In many studies, RPE cells are used for experiments immediately after reaching confluence during cultivation. In contrast, we cultured RPE cells for a long period in this study. Prolonged culture of RPE cells results in their acquisition of characteristics similar to those of in vivo epithelial cells, such as increased TER, expression of ZO-1, cobblestone morphology, pigment deposition, and RPE cell-specific gene expression.^17,19,25) Here, compared to iPSC-RPE cells, ARPE-19 cells exhibited inadequate morphological features and barrier functions as epithelial cells, even after long-term culture (Figs. 1a, b), with low metabolic activity (Fig. 1e). This is consistent with previous reports comparing the OCR of ARPE-19 and primary human fetal RPE cells.²⁶⁾ Therefore, iPSC-RPE cells should be cultured for at least two weeks prior to their use in research. Furthermore, it is necessary to note that ARPE-19 cells did not exhibit physiological cell morphology or barrier function even after prolonged culture.

Here, ATP-linked respiration of iPSC-RPE cells decreased after 24 h stimulation with 50 µg/mL ox-LDL (Fig. 3b). After one week of ox-LDL stimulation, the OXPHOS activity further decreased in iPSC-RPE cells. Subsequently, low concentrations of ox-LDL shifted the metabolism toward FAO (Fig. 1d), while high concentrations of ox-LDL shifted the metabolism toward glycolysis (Fig. 2c). A previous study has reported that acute ox-LDL treatment of macrophages within 24 h shifts metabolism from OXPHOS to glycolysis, accompanied by an increase in the expression of rate-limiting enzymes of glycolysis.²⁴⁾ In contrast, our results indicate that 24-h ox-LDL treatment of iPSC-RPE cells had no effect on ECAR and glycolysis-related gene expression, whereas 1-week ox-LDL treatment increased ECAR and decreased glycolytic gene expression. One month of stimulation with chewing tobacco extract increases OXPHOS and decreases glycolysis in the human esophageal squamous epithelial cell line, Het1A cells. In contrast, eight months of treatment decreases OXPHOS and increases glycolysis in Het1A cells.²⁷⁾ These findings suggest that the cellular phenotype varies depending on the cell type and the duration of stimulation, which should be considered to understand the pathology in living organisms.

After one week of treatment, the expression of glycolysis-related genes was decreased, while ECAR measurement suggested increased glycolytic activity (Figs. 2c, d). In this study, we examined only the mRNA expression of these genes, and it may take longer for the protein expression to decrease. Therefore, after one week, the compensatory increase in glycolytic activity due to the decrease in OXPHOS activity was observed, but over a more extended period, a reduction in glycolytic activity would be expected.

As described above, chronic treatment with 50 µg/mL ox-LDL attenuated OCR and enhanced ECAR, indicating a metabolic shift from OXPHOS to glycolysis. Under these conditions, total ATP production was reduced in ox-LDL-treated cells (Fig. 2e), suggesting that the relative contribution of OXPHOS is greater than that of glycolysis.

In this study, short-term ox-LDL treatment increased the MitoROS levels (Fig. 4b), suggesting that MitoROS production precedes the metabolic changes in RPE cells. Typically, ROS are generated by electron leakage from complexes I and III in the mitochondrial electron transport chain.²⁸⁾ Under hypoxic conditions, ROS shift the metabolism of human umbilical vein endothelial cells from mitochondrial respiration to glycolysis by activating the hypoxia-inducible factor-1α.^28–30) Additionally, enhanced glucose uptake induced by ox-LDL treatment is suppressed by the addition of MitoTEMPO, a mitochondrial superoxide scavenger, in macrophages,²⁴⁾ suggesting that the attenuation of MitoROS production contributes to the inhibition of metabolic alterations. Furthermore, decreased FAO activity leads to fatty acid accumulation, which may contribute to increased MitoROS levels. In fact, treatment of macrophages with ox-LDL leads to the accumulation of long-chain fatty acids in the mitochondria, thereby promoting intracellular ROS production.²⁴⁾ These findings indicate that ox-LDL increases the MitoROS levels and induces metabolic changes, which further elevate the ROS levels, leading to a vicious cycle. On the other hand, unoxidized LDL did not increase MitoROS, while altered OCR and ECAR (Fig. 5). This suggests that LDL affects to the metabolic pathway in RPE cells through a different pathway than ox-LDL.

The key factor regulating OXPHOS activity in ox-LDL remains unknown. Oxidized 1-palmitoyl-2-arachidonyl-sn-glycero-3-phosphocholine (ox-PAPC) is a representative oxidized phospholipid that is also an active component of ox-LDL.³¹⁾ Serbulea et al. reported a decrease in the OCR of bone marrow-derived macrophages (BMDMs) treated with 10 µg/mL ox-PAPC for 4 h.³²⁾ Meanwhile, Gioia et al. reported an increase in the OCR of BMDMs treated with 100 µg/mL ox-PAPC for 24 h.³¹⁾ Thus, the effect of ox-PAPC on metabolic activity varied depending on the stimulation conditions, highlighting the need for further investigation to elucidate the mechanisms underlying the metabolic changes induced by ox-LDL.

In this study, we investigated the effects of long-term stimulation to mirror the pathophysiology of AMD, in which RPE is chronically exposed to causative substances. We found that the pattern of metabolic changes varied with the ox-LDL concentration and stimulation time. Nevertheless, the ATP-producing capacity was apparently reduced, which may lead to cell fragility and death. ox-LDL is taken up by macrophages via the scavenger receptor CD36, and contributes to plaque formation in atherosclerosis.^33,34) Similarities in the risk factors and pathophysiology between AMD and atherosclerosis,^11,35,36) and their epidemiological correlations, strongly suggest ox-LDL as a key factor in AMD progression.

Overall, this study showed that ox-LDL induced metabolic changes and increased MitoROS production in RPE cells, shedding light on the cause of metabolic alterations in mitochondrial dysfunction. Further elucidation of the effects of oxidative lipid species in ox-LDL on metabolic changes can aid in the identification of key molecules and pathways responsible for metabolic alterations.

Acknowledgments

This study was partly supported by the AMED-CREST (JP22gm0910013 to KY) and JSPS KAKENHI (23H05481, 22H05572, and 20H00493 to KY) Grants. We thank Midori Sato (Kyushu University, Japan) for providing excellent experimental technical support.

Conflict of Interest

The authors declare no conflict of interest.

REFERENCES

• 1) Thomas CJ, Mirza RG, Gill MK. Age-related macular degeneration. Med. Clin. North Am., 105, 473–491 (2021).
• 2) Steinmetz JD, Bourne RRA, Briant PS, et al. Causes of blindness and vision impairment in 2020 and trends over 30 years, and prevalence of avoidable blindness in relation to VISION 2020: the right to sight: an analysis for the Global Burden of Disease Study. Lancet Glob. Health, 9, e144–e160 (2021).
• 3) Wong JHC, Ma JYW, Jobling AI, Brandli A, Greferath U, Fletcher EL, Vessey KA. Exploring the pathogenesis of age-related macular degeneration: a review of the interplay between retinal pigment epithelium dysfunction and the innate immune system. Front. Neurosci., 16, 1009599 (2022).
• 4) Kaarniranta K, Uusitalo H, Blasiak J, Felszeghy S, Kannan R, Kauppinen A, Salminen A, Sinha D, Ferrington D. Mechanisms of mitochondrial dysfunction and their impact on age-related macular degeneration. Prog. Retin. Eye Res., 79, 100858 (2020).
• 5) Handa JT, Bowes Rickman C, Dick AD, Gorin MB, Miller JW, Toth CA, Ueffing M, Zarbin M, Farrer LA. A systems biology approach towards understanding and treating non-neovascular age-related macular degeneration. Nat. Commun., 10, 3347 (2019).
• 6) Fisher CR, Ferrington DA. Perspective on AMD pathobiology: a bioenergetic crisis in the RPE. Invest. Ophthalmol. Vis. Sci., 59, AMD41–AMD47 (2018).
• 7) Golestaneh N, Chu Y, Xiao YY, Stoleru GL, Theos AC. Dysfunctional autophagy in RPE, a contributing factor in age-related macular degeneration. Cell Death Dis., 8, e2537 (2017).
• 8) Ferrington DA, Ebeling MC, Kapphahn RJ, Terluk MR, Fisher CR, Polanco JR, Roehrich H, Leary MM, Geng Z, Dutton JR, Montezuma SR. Altered bioenergetics and enhanced resistance to oxidative stress in human retinal pigment epithelial cells from donors with age-related macular degeneration. Redox Biol., 13, 255–265 (2017).
• 9) Colas R, Sassolas A, Guichardant M, Cugnet-Anceau C, Moret M, Moulin P, Lagarde M, Calzada C. LDL from obese patients with the metabolic syndrome show increased lipid peroxidation and activate platelets. Diabetologia, 54, 2931–2940 (2011).
• 10) Koirala D, Beranova-Giorgianni S, Giorgianni F. Early transcriptomic response to oxldl in human retinal pigment epithelial cells. Int. J. Mol. Sci., 21, 8818 (2020).
• 11) Kamei M, Yoneda K, Kume N, Suzuki M, Itabe H, Matsuda KI, Shimaoka T, Minami M, Yonehara S, Kita T, Kinoshita S. Scavenger receptors for oxidized lipoprotein in age-related macular degeneration. Invest. Ophthalmol. Vis. Sci., 48, 1801–1807 (2007).
• 12) Javadzadeh A, Ghorbanihaghjo A, Bahreini E, Rashtchizadeh N, Argani H, Alizadeh S. Serum paraoxonase phenotype distribution in exudative age-related macular degeneration and its relationship to homocysteine and oxidized low-density lipoprotein. Retina, 32, 658–666 (2012).
• 13) Javadzadeh A, Ghorbanihaghjo A, Bahreini E, Rashtchizadeh N, Argani H, Alizadeh S. Plasma oxidized LDL and thiol-containing molecules in patients with exudative age-related macular degeneration. Mol. Vis., 16, 2578–2584 (2010).
• 14) Ruan Y, Jiang S, Gericke A. Age-related macular degeneration: Role of oxidative stress and blood vessels. Int. J. Mol. Sci., 22, 1296 (2021).
• 15) Bellezza I. Oxidative stress in age-related macular degeneration: NRF2 as therapeutic target. Front. Pharmacol., 9, 1280 (2018).
• 16) Nishida M, Tanaka Y, Tanaka Y, Amaya S, Tanaka N, Uyama H, Masuda T, Onishi A, Sho J, Yokota S, Takahashi M, Mandai M. Human iPS cell derived RPE strips for secure delivery of graft cells at a target place with minimal surgical invasion. Sci. Rep., 11, 21421 (2021).
• 17) Hazim RA, Karumbayaram S, Jiang M, Dimashkie A, Lopes VS, Li D, Burgess BL, Vijayaraj P, Alva-Ornelas JA, Zack JA, Kohn DB, Gomperts BN, Pyle AD, Lowry WE, Williams DS. Differentiation of RPE cells from integration-free iPS cells and their cell biological characterization. Stem Cell Res. Ther., 8, 2171 (2017).
• 18) Bharti K, den Hollander AI, Lakkaraju A, Sinha D, Williams DS, Finnemann SC, Bowes-Rickman C, Malek G, D’Amore PA. Cell culture models to study retinal pigment epithelium-related pathogenesis in age-related macular degeneration. Exp. Eye Res., 222, 109170 (2022).
• 19) Ablonczy Z, Dahrouj M, Tang PH, Liu Y, Sambamurti K, Marmorstein AD, Crosson CE. Human retinal pigment epithelium cells as functional models for the RPE in vivo. Invest. Ophthalmol. Vis. Sci., 52, 8614–8620 (2011).
• 20) Quinn RH, Miller SS. Ion transport mechanisms in native human retinal pigment epithelium. Invest. Ophthalmol. Vis. Sci., 33, 3513–3527 (1992).
• 21) Ferrer M, Corneo B, Davis J, Wan Q, Miyagishima KJ, King R, Maminishkis A, Marugan J, Sharma R, Shure M, Temple S, Miller S, Bharti K. A multiplex high-throughput gene expression assay to simultaneously detect disease and functional markers in induced pluripotent stem cell-derived retinal pigment epithelium. Stem Cells Transl. Med., 3, 911–922 (2014).
• 22) Reyes-Reveles J, Dhingra A, Alexander D, Bragin A, Philp NJ, Boesze-Battaglia K. Phagocytosis-dependent ketogenesis in retinal pigment epithelium. J. Biol. Chem., 292, 8038–8047 (2017).
• 23) Uchinomiya S, Matsunaga N, Kamoda K, Kawagoe R, Tsuruta A, Ohdo S, Ojida A. Fluorescence detection of metabolic activity of the fatty acid beta oxidation pathway in living cells. Chem. Commun. (Camb.), 56, 3023–3026 (2020).
• 24) Chen Y, Yang M, Huang W, et al. Mitochondrial metabolic reprogramming by CD36 signaling drives macrophage inflammatory responses. Circ. Res., 125, 1087–1102 (2019).
• 25) Lakkaraju A, Umapathy A, Tan LX, Daniele L, Philp NJ, Boesze-Battaglia K, Williams DS. The cell biology of the retinal pigment epithelium. Prog. Retin. Eye Res., 78, 100846 (2020).
• 26) Iacovelli J, Rowe GC, Khadka A, Diaz-Aguilar D, Spencer C, Arany Z, Saint-Geniez M. PGC-1α induces human RPE oxidative metabolism and antioxidant capacity. Invest. Ophthalmol. Vis. Sci., 57, 1038–1051 (2016).
• 27) Datta KK, Patil S, Patel K, et al. Chronic exposure to chewing tobacco induces metabolic reprogramming and cancer stem cell-like properties in esophageal epithelial cells. Cells, 8, 949 (2019).
• 28) Forrester SJ, Kikuchi DS, Hernandes MS, Xu Q, Griendling KK. Reactive oxygen species in metabolic and inflammatory signaling. Circ. Res., 122, 877–902 (2018).
• 29) Kim JW, Tchernyshyov I, Semenza GL, Dang CV. HIF-1-mediated expression of pyruvate dehydrogenase kinase: A metabolic switch required for cellular adaptation to hypoxia. Cell Metab., 3, 177–185 (2006).
• 30) Lum JJ, Bui T, Gruber M, Gordan JD, DeBerardinis RJ, Covello KL, Simon MC, Thompson CB. The transcription factor HIF-1alpha plays a critical role in the growth factor-dependent regulation of both aerobic and anaerobic glycolysis. Genes Dev., 21, 1037–1049 (2007).
• 31) Di Gioia M, Spreafico R, Springstead JR, Mendelson MM, Joehanes R, Levy D, Zanoni I. Endogenous oxidized phospholipids reprogram cellular metabolism and boost hyperinflammation. Nat. Immunol., 21, 42–53 (2020).
• 32) Serbulea V, Upchurch CM, Ahern KW, Bories G, Voigt P, DeWeese DE, Meher AK, Harris TE, Leitinger N. Macrophages sensing oxidized DAMPs reprogram their metabolism to support redox homeostasis and inflammation through a TLR2-Syk-ceramide dependent mechanism. Mol. Metab., 7, 23–34 (2018).
• 33) Frostegård J. Immunity, atherosclerosis and cardiovascular disease. BMC Med., 11, 117 (2013).
• 34) He F, Zuo L. Redox roles of reactive oxygen species in cardiovascular diseases. Int. J. Mol. Sci., 16, 27770–27780 (2015).
• 35) Friedman E. The role of the atherosclerotic process in the pathogenesis of age-related macular degeneration. Am. J. Ophthalmol., 130, 658–663 (2000).
• 36) Wang J, Zhang H, Ji J, Wang L, Lv W, He Y, Li X, Feng G, Chen K. A histological study of atherosclerotic characteristics in age-related macular degeneration. Heliyon, 8, e08973 (2022).