High glucose-induced phospholipase D activity in retinal pigment epithelium cells: New insights into the molecular mechanisms of diabetic retinopathy

Experimental Eye Research 184 (2019) 243–257

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Experimental Eye Research journal homepage: www.elsevier.com/locate/yexer

High glucose-induced phospholipase D activity in retinal pigment epithelium cells: New insights into the molecular mechanisms of diabetic retinopathy

T

Paula E. Tenconia,b, Vicente Bermúdeza, Gerardo M. Orestia,b, Norma M. Giustoa,b, Gabriela A. Salvadora,b, Melina V. Mateosa,b,∗ a

Instituto de Investigaciones Bioquímicas de Bahía Blanca (INIBIBB), Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), 8000, Bahía, Blanca, Argentina Departamento de Biología, Bioquímica y Farmacia (DBByF), Universidad Nacional del Sur (UNS), 8000, Bahía, Blanca, Argentina

b

A B S T R A C T Keywords: Phospholipase D (PLD) Retinal pigment epithelium (RPE) Inflammation Diabetic retinopathy Nuclear factor kappa B (NFκB) Cyclooxygenase-2 (COX-2) Interleukin-6 (IL-6)

Chronic hyperglycemia, oxidative stress and inflammation are key players in the pathogenesis of diabetic retinopathy (DR). In this work we study the role of phospholipase D (PLD) pathway in an in vitro model of high glucose (HG)-induced damage. To this end, we exposed human retinal pigment epithelium (RPE) cell lines (ARPE-19 and D407) to HG concentrations (16.5 or 33 mM) or to normal glucose concentration (NG, 5.5 mM) for 4, 24 or 72 h. Exposure to HG increased reactive oxygen species levels and caspase-3 cleavage and reduced cell viability after 72 h of incubation. In addition, short term HG exposure (4 h) induced the activation of early events, that involve PLD and ERK1/2 signaling, nuclear factor kappa B (NFκB) nuclear translocation and IκB phosphorylation. The increment in pro-inflammatory interleukins (IL-6 and IL-8) and cyclooxygenase-2 (COX-2) mRNA levels was observed after 24 h of HG exposure. The effect of selective pharmacological PLD1 (VU0359595) and PLD2 (VU0285655-1) inhibitors demonstrated that ERK1/2 and NFκB activation were downstream events of both PLD isoforms. The increment in IL-6 and COX-2 mRNA levels induced by HG was reduced to control levels in cells pre-incubated with both PLD inhibitors. Furthermore, the inhibition of PLD1, PLD2 and MEK/ERK pathway prevented the loss of cell viability and the activation of caspase-3 induced by HG. In conclusion, our findings demonstrate that PLD1 and PLD2 mediate the inflammatory response triggered by HG in RPE cells, pointing to their potential use as a therapeutic target for DR treatment.

1. Introduction Diabetes and obesity are nowadays recognized worldwide as chronic epidemic diseases and diabetic retinopathy (DR) is one of the main causes of visual dysfunction and blindness in working-age adults (Chen and Ma, 2017; Chen et al., 2015; Simo and Hernandez, 2015; Tarr et al., 2013; Simo et al., 2010). DR is characterized by microvascular lesions, impaired blood flow regulation, increased vasopermeability, microaneurysm formation, and eventually widespread non-perfusion and ischemia (Ahsan, 2015; Safi et al., 2014). Neo-

vascularization due to severe hypoxia is characteristic of proliferative DR whereas vascular leakage produced by blood-retinal barrier (BRB) breakdown is the main event involved in the pathogenesis of diabetic macular edema (Simo et al., 2010). Chronic hyperglycemia, oxidative stress (OS), accelerated formation of advanced glycation end-products and inflammation are key players in the pathogenesis of DR (Tarr et al., 2013; Xie et al., 2012; Simo et al., 2010). The retinal pigment epithelium (RPE) is located between the choroid and the neural retina. RPE cells are essential for the integrity and function of the retina since they protect against photo-oxidation

Abbreviations: COX-2, cyclooxygenase-2; DAG, diacylglycerol; DR, diabetic retinopathy; ERK, extracellular signal-regulated kinase; HG, high glucose; HRP, horseradish peroxidase; IL, interleukin; IκB, kappa B inhibitor protein; LPS, lipopolysaccharide; LPPs, lipid phosphate phosphatases; Man, mannitol; NG, normoglucose; NFκB, nuclear factor kappa B; OS, oxidative stress; PA, phosphatidic acid; PC, phosphatidylcholine; PEth, phosphatidylethanol; PGs, prostaglandins; PKC, protein kinase C; PLA2, phospholipase A2; PLD, phospholipase D; POS, photoreceptor outer segments; PVDF, polyvinylidene fluoride; RasGRP, Ras guanine-releasing protein; ROS, reactive oxygen species; RPE, retinal pigment epithelium; WB, western blot ∗ Corresponding author. Instituto de Investigaciones Bioquímicas de Bahía Blanca, CONICET-Bahía Blanca and Universidad Nacional del Sur, Edificio E1, Camino La Carrindanga km 7, 8000, Bahía, Blanca, Argentina. E-mail address: [email protected] (M.V. Mateos). https://doi.org/10.1016/j.exer.2019.04.028 Received 12 November 2018; Received in revised form 5 April 2019; Accepted 30 April 2019 Available online 03 May 2019 0014-4835/ © 2019 Elsevier Ltd. All rights reserved.

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Probes (Eugene, OR, USA). All other chemicals were of the highest purity available.

and mediate the re-isomerization of all-trans-retinal and the renewal of photoreceptor outer segments by phagocytosis (Strauss, 2005, 2016; Simo et al., 2010). Furthermore, RPE cells form the outer BRB, secrete several growth factors and cytokines, and transport nutrients and water to the retina (Simo et al., 2010; Strauss, 2005). The highest rate of glucose utilization in mammal tissues occurs in the retina where the RPE tissue is critical for glucose supply to retinal neurons (Senanayake et al., 2006). To satisfy the retina's large requirement of glucose, RPE cells express high levels of glucose transporters 1 and 3 (Senanayake et al., 2006; Ban and Rizzolo, 2000). Our previous studies demonstrated for the first time the participation of classical phospholipase D isoforms (PLD1 and PLD2) in the inflammatory response of lipopolysaccharide (LPS)-challenged RPE cells (Tenconi et al., 2016; Mateos et al., 2014). Classical PLDs catalyze phosphatidylcholine (PC) hydrolysis to generate phosphatidic acid (PA) which can be further dephosphorylated by lipid phosphate phosphatases (LPPs) to diacylglycerol (DAG) (Tang et al., 2015; Peng and Frohman, 2012). Thus, through the generation of these lipid messengers PLD pathway can modulate the activity of PA-responding proteins, such as mTOR (mammalian target of rapamycin), phospholipase Cγ and sphingosine kinase-1 (Brindley et al., 2009), as well as DAG-responding proteins, such as classical and novel protein kinases C (PKCs), protein kinases D (PKDs) and Ras guanine-releasing protein (RasGRP), among others (Newton, 2010; Caloca et al., 2008; Carrasco and Merida, 2007; Yang and Kazanietz, 2007; Wang, 2006). With the development of pharmacological selective PLD1 and PLD2 inhibitors, these enzymes have been recently postulated as possible therapeutic targets in hypertension, cancer, and autoimmune, thrombotic, neurodegenerative and infectious diseases (Brown et al., 2017; Henkels et al., 2016; Frohman, 2015; Scott et al., 2009). In line with this, our previous studies pointed to the potential use of PLD inhibition for the treatment of ocular inflammatory diseases (Tenconi et al., 2016; Mateos et al., 2014). Our results constituted the first evidence that classical PLDs participate in the LPS-induced inflammatory response of RPE cells through extracellular signal-regulated kinase (ERK1/2) activation and cyclooxygenase-2 (COX-2) expression and prostaglandins (PGs) production (Mateos et al., 2014). In addition, we demonstrated that PLD1, through PKCε activation, also mediates cell survival by preventing LPS-induced increase in caspase-3 cleavage and LPS-induced decrease in Bcl-2 expression and Akt activation (Tenconi et al., 2016). The aim of the present work was to study the role of classical PLDs in RPE cells exposed to an in vitro model of high glucose (HG)-induced damage. This paper reports for the first time the participation of PLD1 and PLD2 in the inflammatory response of RPE cells exposed to HG concentrations.

2.2. Antibodies Anti-phospho-p44/42 MAPK (ERK1/2) (Thr202/Tyr204) (#9101), anti-p44/42 MAPK (ERK1/2) (#9102), anti-IκBα (#4814), antiphospho-IκBα (Ser32/36) (#9246) and anti-cleaved caspase-3 (#9661) were from Cell Signaling (Beverly, MA, USA). Anti-α Tubulin (CP06) was from EMD/Biosciences-Calbiochem (San Diego, CA, USA). AntiNFκB p65 (sc-109), anti-superoxide dismutase 1 (SOD1) (sc-11407), anti-peroxiredoxin (PRX) (sc-25591), polyclonal horse radish peroxidase (HRP)-conjugated goat anti-rabbit IgG (sc-2004), polyclonal HRPconjugated goat anti-mouse IgG (sc-2005) were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Alexa Fluor®546 goat anti-rabbit (A11035) and Alexa Fluor®488 goat anti-rabbit (A11008) were from Life Technologies Corporation (Grand Island, NY, USA). 2.3. Cell culture and treatments Human RPE cell line ARPE-19 from the American Type Culture Collection (ATCC, Manassas, VA, USA) was generously donated by Dr. L. Politi and Dr. N. Rotstein (INIBIBB, Bahía Blanca, Argentina) and human RPE cell line D407 was a generous gift from Dr. E. RodriguezBouland (Weill Medical College of Cornell University, New York, NY, USA). Both types of cells (in passage 5–12) were propagated in T-25 cm2 culture flacks and maintained in Dulbecco's Modified Eagle's Medium (DMEM) supplemented with 10% fetal bovine serum (FBS, Natocor, Argentina) and antibiotic-antimycotic (Anti-Anti 100X, Gibco by Life Technologies, Grand Island, NY, USA) at 37 °C under 5% CO2. Except for the microscopy experiments, for the remaining experiments cells were grown to 100% confluence on plastic 35 mm diameter culture dishes. Confluent ARPE-19 cells were serum-starved for 1 h and subsequently exposed to normal glucose concentration (Control condition or NG, 5.5 mM) or to high glucose concentrations (HG, 16.5 or 33 mM) for 4, 24 or 72 h in order to mimic a peak of and a sustained hyperglycemia. To maintain glucose concentrations, the medium was replaced every 24 h. Osmotic controls were performed with mannitol (Man), an impermeable hexose that does not undergo facilitated transport (Henry et al., 1993, 2000), in order to reach the same osmolarity as that under HG conditions, but maintaining NG concentration. For confirmation, some experiments were also performed using D407 RPE cell line. To study the role of classical PLDs, cells were preincubated with 0.15 μM PLD1 inhibitor (PLD1i) or with 0.5 μM PLD2 inhibitor (PLD2i). To study the role of MEK/ERK pathway and COX-2, U0126 (10 μM) and celecoxib (10 μM) were used, respectively. Cells were pre-incubated with these pharmacological inhibitors for 1 h at 37 °C in serum free DMEM prior to HG exposure. Inhibitors were readded in the HG-containing medium, in each medium replacement, in order to maintain their concentration during all the experimental procedure and DMSO (vehicle) was added to all conditions to achieve a final concentration of 0.005%.

2. Materials and Methods 2.1. Reagents Sterile dextrose (50% w/v in water) was from ROUX- OCEFA S.A. (Buenos Aires, Argentina). Triton X-100 (octyl phenoxy polyethoxyethanol), dimethyl sulfoxide (DMSO), U0126 (1,4-diamino-2,3dicyano-1,4-bis[2-aminophenylthio] butadiene), celecoxib (4-[5-(4Methylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl] benzenesulfonamide) and MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) were from Sigma-Aldrich (St. Louis, MO, USA). VU0359595 (PLD1 inhibitor), VU0285655-1 (PLD2 inhibitor) and oleic acid (OA) were from Avanti Polar Lipids, Inc. (Alabaster, AL, USA). DAPI (4′,6diamidino-2-phenylindole dihydrochloride) was from Life Technologies Corporation (Grand Island, NY, USA). Radiolabeled oleic acid [9,10-3H (N)] ([3H]-OA) (15–60 Ci/mmol) was purchased from New England Nuclear-Dupont (Boston, MA, USA). Preblended dry fluor 2a70 (98% PPO and 2% bis-MSB) was obtained from Research Products International Corp. (Mount Prospect, IL, USA). DCDCDHF (5(6)-carboxy-2′7′-dichlorodihydrofluorescein diacetate) was from Molecular

2.4. MTT reduction assay ARPE-19 cells (1,5 × 104 cells/well) were seeded in 96-well plates and after 24 or 72 h HG exposure, mitochondrial function was assessed by MTT reduction assay. MTT is a water-soluble tetrazolium salt which is reduced by mitochondrial dehydrogenases of metabolically viable cells to a colored, water-insoluble formazan salt. Briefly, MTT (5 mg/ ml) was prepared in sterile phosphate buffer saline (PBS) and added to the cell culture medium to reach a final concentration of 0.5 mg/ml. Cells were subsequently incubated for 1 h at 37 °C in a 5% CO2 atmosphere, washed twice with PBS and lysed with 100 μl of a buffer containing 10% Triton X-100 and 0.1N HCl in isopropanol. The extent of MTT reduction was measured spectrophotometrically (570 nm 244

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Giusto, 1998).

absorbance - 650 nm absorbance) using a Multiskan™ 60 microplate spectrophotometer (Thermo Fisher Scientific). Results are expressed as arbitrary units with respect to the control condition.

2.8. Immunocytochemistry, wide-field fluorescence and confocal microscopy

2.5. Measurement of reactive oxygen species (ROS) production For immunocytochemistry assays, 60,000 cells were seeded onto 12 mm coverslips on plastic 35 mm diameter culture dishes. After 4 h exposure to HG (33 mM), cells were washed twice with ice cold PBS, fixed with 2% paraformaldehyde in PBS at room temperature for 40 min, permeabilized with 0.1% Triton X-100 in PBS for 15 min, and blocked with 2% BSA for 15 min. Cells were then incubated with the primary antibody anti-p65 NFκB subunit (1:100 in blocking solution) or anti-cleaved caspase-3 (1:200 in blocking solution) for 1 h and Alexa Fluor®546-conjugated secondary antibody or Alexa Fluor®488-conjugated secondary antibody (1:500 in blocking solution) for 1 h. Finally, nuclei were stained with DAPI for 10 min at room temperature. The whole immunocytochemical method was performed at room temperature and three washes with ice-cold PBS were performed between each step of the procedure and after nuclei staining. Coverslips were mounted for wide-field fluorescence or confocal microscopy. For wide-field microscopy a Nikon Eclipse TE2000-S microscope coupled to a Nikon DS-Qi2 camera (1608 × 1608 pixels) and a 60x Plan Apo (1.4 N.A.) oil-immersion objective. Confocal microscopy was performed using a TCS-SP2 confocal microscope (Leica MikrosystemeVertrieb GmbH, Wetzlar, Germany) equipped with an acousto optical beam splitter using a 63x (1.2 N.A.) objective. Fluorescence intensity values were determined using ImageJ 1.46 software.

ROS production was measured using the probe DCDCDHF (Molecular Probes, Eugene, OR). This probe can cross the membrane, and, after oxidation, it is converted into a fluorescent compound. 60,000 ARPE-19 cells were seeded onto 12 mm coverslips and exposed to NG or HG (16.5 and 33 mM) or to Man (11.0 and 27.5 mM) for 72 h. After the experimental treatment, cell culture medium was removed and replaced by medium containing 10 μM DCDCDHF and cells were incubated for 30 min at 37 °C. Cells were subsequently washed three times with PBS and coverslips were mounted for examination with a Nikon Eclipse TE2000-S microscope coupled to a Nikon DS-Qi2 camera (1608 × 1608 pixels) and a 60x Plan Apo (1.4 N.A.) oil-immersion objective. Fluorescence intensity values were determined using ImageJ 1.46 software. 2.6. Western blot (WB) assays After experimental treatments, the medium was removed from confluent 35 mm dishes, cells were washed three times with PBS and scraped off with 80 μl ice-cold RIPA lyses buffer [10 mM Tris-HCl (pH 7.4), 15 mM NaCl, 1% Triton X-100, 5 mM NaF, 1 mM Na2VO4 and the complete protease inhibitor cocktail]. Protein content of total cell lysates was determined by Bradford method (Bradford, 1976) (Bio-Rad Life Science group, #500–0006) and samples were denatured with Laemmli sample buffer at 100 °C for 5 min (Laemmli, 1970). Equivalent amounts of proteins (30 μg) were separated by sodium dodecylsulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to polyvinylidene fluoride (PVDF) membranes (Millipore, Bedford, MA, USA). WB assays were performed as previously described (Tenconi et al., 2016; Mateos et al., 2014). Briefly, after being blocked with 10% BSA in TTBS buffer [20 mM Tris–HCl (pH 7.4), 100 mM NaCl and 0.1% (w/v) Tween 20] at room temperature for 2 h, membranes were subsequently incubated overnight at 4 °C with primary antibodies, washed three times with TTBS and exposed to the appropriate HRP-conjugated secondary antibody for 2 h at room temperature. Immunoreactive bands were detected by enhanced chemiluminescence (Pierce® ECL Western Blotting Substrate, #32209, Thermo Scientific) using UltraCruz® Autoradiography Film, Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Densitometry values of the immunoreactive bands were determined using ImageJ 1.46 software. The molecular weight of bands was determined using the spectra multicolor broad range protein ladder (26634, Thermo Scientific).

2.9. Real-time quantitative PCR (qPCR) assay Total RNA was isolated from treated ARPE-19 cells (confluent 35 mm dishes) using 500 μl TRIzol™ reagent (Invitrogen, Carlsbad, USA) following the manufacturer's instructions. The RNA was resuspended in RNase-free water and its concentration and purity were assessed from the A260:A280 absorbance ratio in a PicoDrop Spectrophotometer. Total RNA (2 μg) was used to synthesize cDNA by reverse transcription (RT) in a final volume of 25 μl containing 1 μg Random Primers hexamers (Biodynamics, Buenos Aires, Argentina), 1x M-MLV RT Reaction Buffer, 0.5 mM of each dNTP, 25 UI RNase inhibitor (Promega, Madison, WI, USA) and 200 UI M-MLV Reverse Transcriptase (Promega, Madison, WI, USA). The cDNA resulting from the RT was amplified by real-time quantitative PCR (qPCR). The qPCR assays were performed in a final volume of 10 μl using 0.2 μM of each primer and KAPA SYBR® FAST qPCR Kit Master Mix (Kapa Biosystems, Boston, Massachusetts, USA). Primer combinations for the specific amplification of the genes analyzed were designed in ‘Primer BLAST’ (https://www.ncbi.nlm.nih.gov/tools/primer-blast/) and purchased from Invitrogen Life Technologies. Primer sequences for the specific amplification of human COX-2, IL-6, IL-8 and GADPH are listed in Table 1. Gene expression levels were determined using RotorGene 6000 Series Software machine (Corbett Research, Australia) with the following conditions: 35 cycles of denaturation at 94 °C for 20 s, annealing and extension at 58 °C for 30 s and a final extension step at 72 °C for 30 s. Ct values of COX-2, IL-6 and IL-8 mRNA obtained from 3 different experiments were normalized using GAPDH as reference gene, and relative quantification was performed applying 2−ΔΔCt method (Livak and Schmittgen, 2001).

2.7. PLD activity assay To assay PLD activity, transphosphatidylation reaction was measured in ARPE-19 cells exposed to HG (33 mM), Man (27.5 mM) or to NG condition as previously described (Mateos et al., 2014). Briefly, confluent 35 mm dishes of ARPE-19 cells were pre-labeled with radiolabeled oleic acid ([3H]-OA) mixed with unlabeled OA (final concentration of 2.2 μM and 0.5 μCi/dish) in the presence of lipid-free bovine serum albumin (BSA) (4 mol OA/mol BSA) in serum-free DMEM. Cells were incubated with this medium for 24 h at 37 °C to allow [3H]OA incorporation. After 24 h, the medium was removed, cells were washed three times with PBS and were exposed to NG, HG (33 mM) or Man (27.5 mM) for 4 h in the presence of 0.4% ethanol. After the experimental treatment, the medium was removed, cells were washed three times with PBS and scraped off with 800 μl PBS. Lipids were extracted and radiolabeled phosphatidylethanol ([3H]-PEth), an unequivocal PLD activity marker, was isolated and quantified as previously described by our laboratory (Mateos et al., 2010, 2014; Salvador and

2.10. Statistical analysis Statistical analysis was performed using ANOVA followed by Bonferroni's test to compare means. p-values lower than 0.05 were considered statistically significant. Data represent the mean value ± SD of three independent experiments. The WBs and microscopy images shown are representative of three independent 245

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Table 1 Primer sequences for the specific amplification of human COX-2, IL-6, IL-8 and GADPH. Gene Name

Gene Symbol

Sequence accession number

Primer sequences 5′-3′

Amplicon (pb)

Interleukin-6

HUMAN_IL-6

NM_000600.4

130

Interleukin-8

HUMAN_IL-8

NM_000584.3

Cyclooxygenase-2

HUMAN_COX-2

NM_000963.3

Glyceraldehyde-3-phosphate dehydrogenase

HUMAN_GAPDH

NM_001289746.1

GATGAGTACAAAAGTCCTGATCCA CTGCAGCCACTGGTTCTGT TCTGTGTGAAGGTGCAGTTTT GTGTGGTCCACTCTCAATCAC CCCATGTCAAAACCGAGGTG AAATTCCGGTGTTGAGCAGTT CACTGAATCTCCCCTCCTCACA TGATGGTACATGACAAGGTGCG

127 106 87

Fig. 1. ROS generation in HG-exposed RPE cells. ARPE-19 cells were seeded onto 12 mm coverslips and exposed to NG (5.5 mM), HG (16.5 and 33 mM) or Man (11.0 and 27.5 mM) for 72 h. A). ROS production was measured using DCDCDHF as described in Materials and Methods. Representative images from three independent experiments are shown. Scale bar = 10 μm. B) Bar graph shows fluorescence intensity expressed as arbitrary units (AU) with respect to control condition (NG). C) WB assays showing SOD1 and PRX expression in ARPE-19 cells exposed to NG, HG (16.5 mM) or Man (11 mM) for 72 h. The bar graph shows the densitometry values of each protein with respect to α-tubulin expressed as arbitrary units with respect to NG condition. Numbers to the right indicate molecular weights. For B and C asterisks (*) indicate significant differences with respect to each control condition (***p < 0.001; **p < 0.01; *p < 0.05).

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Fig. 2. HG-effect on caspase-3 cleavage and RPE cell viability. A) Cleaved caspase-3 detection. ARPE19 cells were seeded onto 12 mm coverslips and exposed to NG (5.5 mM), HG (16.5 mM) or Man (11 mM) for 72 h. After experimental treatment immunocytochemistry assays were performed as described in Materials and Methods. Representative images from three independent experiments show cleaved caspase-3 staining (first column), DAPI staining (second column) and the merge image (third column). Scale bar = 10 μm. B) Bar graph shows nuclear cleaved caspase-3 fluorescence intensity expressed as arbitrary units (AU) with respect to control condition (NG). C) Cell viability was evaluated using MTT reduction assay in ARPE-19 cells exposed to NG or HG (33 mM) for 24 h and to NG, HG (16.5 and 33 mM) or Man (11 and 27.5 mM) for 72 h. Data represent the mean value ± SD of three independent experiments. Results are expressed as arbitrary units (AU) with respect to control conditions (NG). Asterisks (*) indicate significant differences with respect to each control condition (***p < 0.001; **p < 0.01).

and NG (Fig. 1 A and B) indicating that ROS generation induced by HG treatment is not due to increased osmolarity under HG conditions. In accordance with the enhanced ROS generation induced by HG, WB assays showed decreased peroxiredoxin (PRX) expression in ARPE19 cells exposed to HG (16.5 mM) for 72 h (Fig. 1C). On the contrary, no changes in superoxide dismutase 1 (SOD1) expression were observed (Fig. 1C). Caspase-3 cleavage was studied using immunocytochemistry assays, followed by wide-field fluorescence microscopy and mitochondrial function was assessed using the MTT reduction assay. Fig. 2 shows that HG (16.5 mM) exposure for 72 h increased caspase-3 cleavage by 100% with respect to NG (Fig. 2 A and B). In addition, MTT reduction was reduced by 30% in ARPE-19 cells exposed to both HG concentrations (16.5 and 33 mM) for 72 h with respect to the control condition (Fig. 2C). No differences were observed in MTT reduction (Fig. 2C) and caspase-3 activation (Fig. 2 A and B) between NG and osmotic controls after 72 h, thus indicating apoptosis is not induced by increased osmolarity but by increased glucose concentrations. After 24 h exposure to HG (33 mM), no differences in mitochondrial function were observed with respect to NG (Fig. 2C). Our results demonstrate that sustained HG exposure induces enhanced OS, cell damage and caspase-3 activation in ARPE-19 cells. Since the loss of mitochondrial function and the increment in ROS generation was similar in the cells exposed to both HG

experiments. 3. Results 3.1. Sustained HG exposure increases reactive oxygen species (ROS) generation and caspase-3 cleavage and reduces mitochondrial viability in ARPE-19 cells To study the effect of sustained HG exposure on RPE cells, ARPE19 cells were exposed to normal glucose concentration (NG, 5.5 mM) or to both HG concentrations (16.5 mM or 33 mM glucose) for 72 h. Both HG concentrations could be associated with possible in vivo hyperglycemias. Whereas 33 mM HG represents an extremely high hyperglycemia, 16.5 mM HG mimics a hyperglycemia that can be present in diabetic patients even without producing symptoms. Osmotic controls were performed with mannitol (Man) as described in the experimental procedures section. OS was evaluated by measuring ROS generation, using the probe DCDCDHF. Once introduced into cells, this probe is deacetylated and easily oxidized in the presence of ROS to 2′,7′-dichlorofluorescein, a highly fluorescent compound. ROS generation rate was significantly increased in ARPE-19 cells exposed to 16.5 mM HG (174%) and 33 mM HG (230%) with respect to NG (Fig. 1 A and B). No differences were observed in ROS generation between osmotic controls 247

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Fig. 3. ERK1/2 activation in HG-exposed ARPE-19 cells. ARPE-19 cells were exposed to NG, HG (16.5 mM) or Man (11 mM) for 4 h (A) and 72 h (C) or to NG, HG (33 mM) or Man (27.5 mM) for 4 h (B). ERK1/2 activation was evaluated by WB assays using anti-phosphoERK1/2 (pERK1/2) antibody. The bar graph shows the densitometry values of pERK1/2/ERK1/2 expressed as arbitrary units with respect to control conditions (NG). Asterisks (*) indicate significant differences with respect to NG (***p < 0.001; **p < 0.01; *p < 0.05). Numbers to the right indicate molecular weights. Fig. 4. PLD activation in HG-exposed RPE cells. A) Schematic view of the transphosphatidylation reaction carried out by PLD1 and PLD2 in the presence of ethanol (EtOH). B) ARPE-19 cells were pre-labeled with [3H]-OA as described in Materials and Methods. PLD activity was measured as [3H]-PEth formation in the presence of 0.4% EtOH in cells pre-incubated with 0.05% DMSO (vehicle), 0.15 μM PLD1i or 0.5 μM PLD2i for 1 h before HG (33 mM) or NG exposure for 4 h. PLD activity was also measured in cells exposed to Man (27.5 mM) for 4 h. Data represent the mean value ± SD of at least three independent experiments. Results are expressed as percentage of PLD activity considering as 100% PLD activity in NG condition (7300 dpm [3H]-PEth/mg protein). Asterisks (*) indicate significant differences with respect to NG (**p < 0.01; ***p < 0.001), number signs (#) indicate significant differences with respect to HG (#p < 0.05; ##p < 0.01).

concentrations, the following 72 h treatment experiments were performed using 16.5 mM HG. The latter mimics sustained hyperglycemia in vivo more accurately.

to HG for 48 h (Che et al., 2016; Yuan et al., 2009). Furthermore, previous findings from our laboratory demonstrated that ERK1/2 plays a key role in the inflammatory response of ARPE-19 cells to LPS mediating COX-2 expression, PGE2 production and cell viability loss (Mateos et al., 2014). We therefore decided to study not only the effect of sustained (72 h) but also acute (4 h) HG exposure on ERK1/2 activation. To this end, ERK1/2 activation was evaluated by WB in cells exposed to HG for 4 or 72 h. Fig. 3 shows that after 4 h incubation with

3.2. HG exposure modulates ERK1/2 activation in ARPE-19 cells It has been reported that ERK activation mediates iNOS expression, cell damage and epithelial-mesenchymal transition in RPE cells exposed 248

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Fig. 5. Role of PLD1 and PLD2 on ERK1/2 activation. ARPE-19 (A and C) and D407 (B and D) cells were pre-incubated with 0.05% DMSO (vehicle conditions), 0.15 μM PLD1i or 0.5 μM PLD2i for 1 h before 33 mM HG (A and B) or 27.5 mM Man (C and D) exposure for 4 h. ERK1/2 activation was evaluated by WB as described in Fig. 3. Numbers to the right indicate molecular weights. The bar graph shows the densitometry values of pERK1/2/ERK1/2 expressed as arbitrary units with respect to NG condition. Asterisks (*) indicate significant differences with respect to NG (***p < 0.001; **p < 0.01; *p < 0.05).

pathway in the inflammatory response of RPE cells to LPS (Tenconi et al., 2016; Mateos et al., 2014). In the present work, we studied whether or not classical PLDs are involved in the modulation of the inflammatory response induced by HG in RPE cells. On account of the fact that in the presence of primary alcohols only classical PLDs are able to catalyze transphosphatidylation reaction yielding phosphatidylalcohols instead of PA (Fig. 4A) (Frohman, 2015; Mateos et al., 2006, 2008, 2014; Kobayashi and Kanfer, 1987), radiolabeled phosphatidylethanol ([3H]-PEth) generation was measured to determine the effect of HG on PLD activity. To this end, ARPE-19 cells were pre-labeled with [3H]-OA for 24 h and subsequently exposed to NG, HG (33 mM) or Man (27.5 mM) for 4 h in the presence of 0.4% ethanol. [3H]-PEth generation was measured as previously described (Mateos et al., 2014). Fig. 4B shows that PLD activity was increased by 80% after 4 h treatment with HG (33 mM) while no changes in PLD activity were detected in cells exposed to Man (27.5 mM), with respect

both HG concentrations (16.5 and 33 mM) ERK1/2 activation (phosphorylation) was increased by 35 and 82%, respectively, with respect to NG (Fig. 3 A and B). Immunocytochemistry assays showed that HG (33 mM) exposure for 4 h increased ERK1/2 nuclear translocation (Supplementary Fig. 1). Similar effects on ERK1/2 phosphorylation (Fig. 3 A and B) and nuclear translocation (Supplementary Fig. 1) were observed in cells exposed to Man (osmotic controls) for 4 h, thus suggesting that this effect is mediated by the increment in osmolarity under HG conditions. After 72 h HG (16.5 mM) exposure, ERK1/2 phosphorylation was reduced by 20% with respect to NG, and no differences in ERK1/2 activation were observed between the osmotic control and NG (Fig. 3C).

3.3. PLD activity is increased in ARPE-19 cells exposed to HG Our previous studies demonstrated the participation of PLD 249

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Fig. 6. NFκB activation in ARPE-19 cells exposed to HG and Man. ARPE-19 cells were seeded onto 12 mm coverslips and were treated for 4 h with HG (16.5 and 33 mM), Man (11 mM and 27.5 mM) or NG (A) or for 24 h with HG (33 mM), Man (27.5 mM) or NG (B). For A and B, after experimental treatment immunocytochemistry assays were performed as described in Materials and Methods. Representative fluorescence images from three independent experiments show p65 NFκB staining (first row), and the merge image (second row) between p65 NFκB (green) and DAPI (blue). Scale bar = 10 μm. C) Bar graph shows nuclear p65 NFκB fluorescence intensity expressed as arbitrary units (AU) with respect to control conditions (NG). Asterisks (*) indicate significant differences with respect to each control condition (***p < 0.001; **p < 0.01). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

to cells exposed to NG. The incubation with selective pharmacological inhibitors of PLD1 or PLD2 (PLD1i or PLD2i), prior to and during HG (33 mM) treatment, partially reduced the increased [3H]-PEth generation induced by HG (Fig. 4B). These results demonstrate that both classical PLD isoforms are activated in ARPE-19 cells exposed to HG.

conditions and that both PLDs can mediate ERK activation induced by HG. In addition, the activation of only one PLD isoform seems to be insufficient to induce ERK activation in the presence of HG in RPE cells. On the contrary, ERK1/2 activation induced by osmotic stress was found to be dependent neither on PLD1 nor on PLD2.

3.4. HG-induced ERK1/2 activation in RPE cells depends on PLD1 and PLD2

3.5. HG-induced nuclear factor kappa B (NFκB) activation in RPE cells depends on PLD1, PLD2 and ERK1/2

Previous work from our laboratory demonstrated that PLD2 mediates LPS-induced ERK1/2 activation in ARPE-19 cells (Mateos et al., 2014). In order to study the role of classical PLDs in HG-induced ERK1/ 2 activation, ARPE-19 and D407 cells were incubated with PLD1i or PLD2i prior to and during HG (33 mM) or Man (27.5 mM) treatment for 4 h. In ARPE-19 cells either PLD1i or PLD2i, reduced HG-induced ERK1/2 activation to control condition levels (Fig. 5A). These results demonstrate that HG-induced ERK1/2 activation is dependent on both PLD isoforms in ARPE-19 cells. Similar results were obtained in D407 RPE cells, in which HG exposure for 4 h increased ERK1/2 activation by 67% with respect to NG (Fig. 5B). As observed in ARPE-19 cells, PLD1i and PLD2i also reduced HG-induced ERK1/2 activation to control levels in D407 cells (Fig. 5B). As expected, and in agreement with the results derived from the PLD activity assay, the activation of ERK1/2 induced by Man was not prevented with PLD1i or PLD2i neither in ARPE-19 (Fig. 5C) nor in D407 cells (Fig. 5D). Taken together, our results demonstrate that in RPE cells PLD activity is increased under HG

NFκB is a ubiquitous transcription factor that is well known for its role in transcriptional activation of inflammatory mediators, such as cytokines and COX-2. NFκB comprises a small family of inducible transcription factors that form homodimers and heterodimers and that are kept inactive in the cytoplasm through the association with inhibitory proteins (IκBs). Activation of NFκB occurs after IκBs phosphorylation by the multiprotein IκB-kinase (IKK). Phosphorylated IκBs (pIκBs) are then polyubiquitinylated and subsequently degraded by the proteasome. Following IκB degradation, NFκB dimers are released and are then able to translocate into the nucleus to activate gene transcription (Jost and Ruland, 2007; Jacobs and Harrison, 1998). To study the effect of HG on NFκB activation, immunocytochemistry assays, followed by wide-field fluorescence and confocal microscopy, were performed as described in the experimental procedures section. Fig. 6A shows that a 4 h treatment with HG (16.5 and 33 mM) induced NFκB translocation to the nucleus in ARPE-19 cells. In addition, NFκB was also observed to translocate to the nucleus in ARPE-19 cells 250

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Fig. 7. Role of PLD1, PLD2 and ERK1/2 on HG-induced NFκB activation in ARPE-19 cells. A) Subcellular localization of p65 NFκB. ARPE-19 cells were seeded onto 12 mm coverslips and were preincubated with 0.05% DMSO (NG and HG conditions), 0.15 μM PLD1i, 0.5 μM PLD2i or 10 μM U0126 for 1 h before HG (33 mM) or NG exposure for 4 h. After experimental treatment immunocytochemistry assays were performed as described in Materials and Methods. Representative confocal images from three independent experiments show p65 NFκB staining. Scale bar = 20 μm. B) Bar graph shows nuclear p65 NFκB fluorescence intensity expressed as arbitrary units (AU) with respect to control condition (NG). Asterisks (*) indicate significant differences with respect to NG (***p < 0.001; **p < 0.01; *p < 0.05). C) WB assays were performed to detect pIκBα, total IκBα and αTubulin in ARPE-19 cells exposed to the experimental conditions described in A. Numbers to the right indicate molecular weights. D) Bar graph shows the densitometry values of pIκBα/IκBα expressed as arbitrary units with respect to control condition (NG). Asterisks (*) indicate significant differences with respect to NG (**p < 0.01; *p < 0.05).

of osmotic stress.

exposed to Man (11 and 27.5 mM) for 4 h (Fig. 6 A and C). After a 24 h treatment, NFκB nuclear translocation was still observed in ARPE19 cells exposed to HG (33 mM) but not in cells exposed to Man (27.5 mM) (Fig. 6 B and C). These results demonstrate that the osmotic stress effect on NFκB is an early but transient event while HG-induced NFκB activation persists at least for 24 h. NFκB nuclear translocation induced by a 4 h HG exposure was inhibited in ARPE-19 cells pre-incubated with PLD1i, PLD2i and MEK/ ERK pathway inhibitor (U0126) (Fig. 7 A and B). The same was also observed in D407 cells (Fig. 8). In agreement with the enhanced NFκB nuclear translocation, WB assays showed an increased phosphorylation of IκBα (pIκBα) in ARPE-19 cells exposed to HG (Fig. 7 C and D). Furthermore, PLD1, PLD2 and ERK pharmacological inhibition reduced pIκBα and increased total IκBα (Fig. 7C). In addition, PLD1i, PLD2i and U0126 also prevented NFκB nuclear localization after 24 h of HG treatment, at this time of incubation HG-induced effect on the transcription factor is not due to changes in extracellular osmolarity (Supplementary Fig. 2). Taken together, these results demonstrate that in RPE cells, HG induces NFκB activation through a signaling mechanism that involves PLD1, PLD2 and ERK1/2 and that is independent

3.6. PLD pathway modulates HG-induced increment in COX-2, interleukin6 (IL-6) and IL-8 mRNA levels In view of the activation of NFκB observed after a 4 h exposure to HG, we next studied the level of expression of two pro-inflammatory interleukin (IL-6 and IL-8) mRNA and of COX-2 mRNA. To this end, qPCR assays were performed in ARPE-19 cells exposed to HG (33 mM) for 4 and 24 h. Whereas as after a 4 h HG exposure, neither IL-6 nor IL-8 transcript accumulation was detected and COX-2 transcript accumulation was increased by 230% (data not shown), after a 24 h HG treatment, a significant higher transcript accumulation of IL-6 (340%), IL-8 (56%) and COX-2 (350%) was observed with respect to cells maintained in NG (Fig. 9 A, B and C, respectively). In cells exposed to HG for 24 h and also pre- and co-incubated with PLD1i or PLD2i, IL-6 and COX-2 mRNA levels were reduced to control condition levels (Fig. 9 A and C). Interestingly, the HG-induced increment in IL-8 mRNA levels was only reduced in cells incubated with PLD1i and was unaffected in cells incubated with PLD2i (Fig. 9B). In ARPE-19 cells exposed to Man 251

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Fig. 8. Role of PLD1, PLD2 and ERK1/2 on HG-induced NFκB activation in D407 cells. A) D407 cells were seeded onto 12 mm coverslips and were preincubated with 0.05% DMSO (NG and HG conditions), 0.15 μM PLD1i, 0.5 μM PLD2i or 10 μM U0126 for 1 h before HG (33 mM) or NG exposure for 4 h. After experimental treatment, immunocytochemistry assays were performed as described in Materials and Methods. Representative fluorescence images from three independent experiments show p65 NFκB staining (first row), and the merge image (second row) between p65 NFκB (green) and DAPI (blue). Scale bar = 10 μm. B) Bar graph shows nuclear p65 NFκB fluorescence intensity expressed as arbitrary units (AU) with respect to control condition (NG). Asterisks (*) indicate significant differences with respect to NG (***p < 0.001). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

RPE cells. In this work we demonstrate that HG exposure induces early triggered (after 4 h treatment) and concatenated events that involve PLD and ERK1/2 activation, IκB phosphorylation and NFκB nuclear translocation (Figs. 3–8). The increase in PEth generation undoubtedly confirms the activation of classical PLDs in the RPE response to HG exposure (Fig. 4B). Moreover, results obtained in the presence of PLD1 and PLD2 selective inhibitors demonstrate that both classical PLD isoforms are activated under HG conditions (Fig. 4B). In addition, in ARPE-19 and in D407 cells HG-induced ERK1/2 activation depends on both PLD isoforms, since the inhibition of either PLD isoform prevents HG-induced ERK1/2 activation (Fig. 5 C and D). HG-induced ERK1/2 activation was also reported after 48 h treatment by other authors in ARPE-19 cells (Xie et al., 2012; Yuan et al., 2009). However, we demonstrate that ERK1/2 and NFκB activation can be triggered by a shorter HG-exposure time. Similar results are obtained under osmotic control conditions, suggesting that these early events can be mediated by the increment in osmolarity either under HG conditions o under other hyperosmolar stress, such as high salt diet. On the contrary, our results show that after a sustained HG exposure (72 h), ERK1/2 phosphorylation was reduced with respect to control condition, possibly by a decreased MEK activity or by an increased MAP kinase phosphatase activity (Fig. 3C). Taking into account that ERK1/2 pathway has been shown to mediate RPE cell damage under HG conditions (Xie et al., 2012; Yuan et al., 2009 and our results presented in this work), we hypothesize that downregulation of ERK1/2 signaling observed after a 72 h HG exposure could be a cellular protective mechanism elicited in RPE cells exposed to sustained hyperglycemia. Our results constitute the first evidence that HG induces ERK1/2 and NFκB activation in a PLD1 and PLD2 dependent manner in RPE cells. The fact that either PLD1i or PLD2i has the ability to prevent HGinduced ERK1/2 and NFkB activation led us to conclude that both PLDs must be activated simultaneously to mediate these cellular events (Figs. 5, 7 and 8). On the contrary, osmotic controls showed that although increased osmolarity can activate ERK1/2 and induce an early but transient NFκB nuclear translocation, the activation of ERK1/2 induced by Man is a PLD-independent event (Fig. 5 C and D). In agreement with the latter, PLD activity was found not to be induced by Man

(27.5 mM) for 24 h no changes were detected in IL-6, IL-8 and COX-2 mRNA levels with respect to cells exposed to NG (Fig. 9 A, B and C). 3.7. PLD1, PLD2, MEK/ERK pathway and COX-2 inhibitors prevent the increased caspase-3 activation and the reduced cell viability induced by HG in RPE cells In order to study the effect of PLD1, PLD2 and MEK/ERK pathway on the increased caspase-3 cleavage and the reduced RPE cell viability induced by HG, ARPE-19 cells were exposed to HG (16.5 mM) for 72 h in the presence of PLD1i, PLD2i or U0126 and caspase-3 cleavage and cell viability was assayed as previously described. The effect of COX-2 inhibitor celecoxib (10 μM) was also studied. As shown in Fig. 10, the inhibition of PLD1, PLD2, the MEK/ERK pathway and COX-2 prevented the activation of caspase-3 (Fig. 10 A and B) and the loss in cell viability (Fig. 10 C) induced by HG treatment for 72 h. 4. Discussion The incidence of diabetes increases every year and most patients with type 1 or type 2 diabetes eventually develop some degree of DR (Chen and Ma, 2017). Therefore, DR is considered a significant threat to global health. One of the most important tissues for the proper retina function is the RPE, which maintains the structural integrity of photoreceptors by secretion of neurotrophic factors and by the phagocytosis of photoreceptor outer segments (POS) (Strauss, 2016). Furthermore, the RPE constitutes the interface between the retina and the body system and is essential to glucose supply to neural retina (Senanayake et al., 2006). Our previous findings described for the first time the participation of classical PLDs in the LPS-induced inflammatory response of RPE cells through ERK1/2 activation, COX-2 expression and PGs production (Mateos et al., 2014). In LPS-challenged ARPE-19 cells, ERK1/2 activation was dependent only on PLD2 while COX-2 induction required PLD1 and PLD2 activation (Mateos et al., 2014). We also demonstrated that PLD1, through PKCε activation, also mediates cell survival by preventing LPS-induced apoptotic signals (Tenconi et al., 2016). In addition, findings from the present work indicate that PLD1 and PLD2 also mediate the inflammatory response triggered by HG in 252

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The mRNA levels of these pro-inflammatory mediators were found not to be modified in cells exposed to Man (Fig. 9), thus indicating that the inflammatory response is glucose-specific and not induced by osmotic stress. In agreement with our results, it was previously reported that linoleic acid induces COX-2 expression in ARPE-19 through ERK1/2 activation, IκB degradation and NFκB activation (Fang et al., 2007). Furthermore, IL-17A-induced production of IL-6 by ARPE-19 cells also involves ERK1/2 and NFκB pathways (Chen et al., 2011). As stated above, regarding ERK1/2 and NFκB activation, the results obtained with PLD1i and PLD2i showed that activation of both isozymes is necessary to mediate IL-6 and COX-2 expression in HG-exposed RPE cells. In a different way, HG-induced IL-8 expression was observed to be reduced only by PLD1i, thus suggesting that a transcription factor, different from NFκB and not PLD2-regulated, is involved in the expression of this pro-inflammatory cytokine. In line with this, Catarino and collaborators reported that in ARPE-19 cells treated with 25-hydroxycholesterol IL-8 expression was increased in an ERK1/2 and AP-1 dependent manner (Catarino et al., 2012) and Grab and co-workers demonstrated that COX-2 and IL-8 expression was increased only in HEK-293 cells overexpressing PLD1 but not PLD2 (Grab et al., 2004). Recently, and in agreement with our findings, a decrease in IL-6 mRNA levels was observed in rheumatoid arthritis synovial fibroblast treated either with PLD1 or with PLD2 inhibitors (Friday and Fox, 2016). PLD isoforms inhibition was also reported to have anti-inflammatory effects and to be a potential therapeutic target for the treatment of periodontitis since PLD1 and PLD2 inhibitors and siRNA inhibited nicotine- and LPS-induced ERK1/2 and NFκB activation and COX-2 expression in human periodontal ligament cells (Shin et al., 2015). PLD1 and PLD2 inhibitors and siRNA were also found to inhibit mRNA levels and secretion of tumor necrosis factor-α, IL-1β and IL-8 in human periodontal ligament cells (Shin et al., 2015). Taken together, these previous evidences and the findings reported herein, postulate PLD isoenzymes as promising targets for controlling pro-inflammatory gene expression in several tissues as well as in diverse retinal inflammatory diseases, such as DR and aged-related macular degeneration. Nevertheless, results obtained in RPE cells exposed to HG or to LPS demonstrate that the role of PLD1 and PLD2 in the inflammatory response of the RPE differs depending on the nature of the inflammatory stimulus, thus highlighting the importance of elucidating the role of classical PLDs in different inflammatory contexts. Since the development of selective inhibitors of PLD1 and PLD2, these enzymes have been postulated as possible therapeutic targets for several diseases (Brown et al., 2017; Henkels et al., 2016; Frohman, 2015; Scott et al., 2009). The effectiveness of the inhibitors used was previously assayed in our laboratory (Mateos et al., 2012, 2014) as well as by other authors (Burkhardt et al., 2015; Shin et al., 2015). On account of the fact that, to our knowledge, no clinical trials have been performed for these inhibitors to date, their possible side effects are still unknown. However, selective inhibitors of PLD enzymes are derivatives from the antipsychotic drug halopemide, which has been used clinically for prolonged periods of time at high doses which have the potential to block PLD activity. Furthermore, PLD1 and 2 single- and doubleknockout mice are viable (Burkhardt et al., 2015), suggesting that PLD pharmacological inhibition has a reasonable chance of being well tolerated, especially if PLD inhibition is achieved locally, for example with intravitreal injections. In agreement with this, our previous results showed that the concentrations of the PLD1 and PLD2 inhibitors used in the present work do not affect RPE cell viability under control conditions (Mateos et al., 2014). Furthermore, non-specific effects on astrocyte proliferation were observed when these inhibitors were used at concentrations 10 times higher than those used in our work (Burkhardt et al., 2015). Unlike PLDs, whose role in DR-associated inflammation is postulated for the first time in the present work, phospholipases A2 (PLA2) have been shown to play key roles in DR pathophysiology (Giurdanella et al., 2015, 2017; Gong et al., 2014; Lupo et al., 2013). PLA2s are

Fig. 9. HG-effect on IL-6, IL-8 and COX-2 mRNA levels. ARPE-19 cells were preincubated with 0.05% DMSO (NG and HG conditions), 0.15 μM PLD1i or 0.5 μM PLD2i for 1 h before HG (33 mM) or NG exposure for 24 h. Cells were also exposed to Man (27.5 mM) for 24 h qPCR assays for the quantification of IL-6 (A), IL-8 (B) and COX-2 (C) mRNA levels were performed as described in Materials and Methods. Results are expressed as 2−ΔΔCt relative fold mRNA accumulation using GAPDH as internal reference gene. Asterisks (*) indicate significant differences with respect to NG (***p < 0.001; **p < 0.01; *p < 0.05).

(Fig. 4B). The effect of MEK inhibitor U0126 on NFκB nuclear translocation and IκB phosphorylation demonstrates that NFκB activation depends not only on both PLDs but also on HG-induced ERK1/2 activation in ARPE-19 and in D407 cells (Figs. 7 and 8). The RPE is highly specialized not only in phagocytosis of POS, but also in secretion of a number of cytokines, chemokines, angiogenic and anti-angiogenic factors. Thus, the RPE secretome is crucial in DR development (Ponnalagu et al., 2017). In agreement with this, elevated levels of IL-6, IL-8 and monocyte chemo-attractant protein-1 (MCP-1) were observed in the vitreous of patients with proliferative DR who did not respond to the anti-VEGF treatment, suggesting the role of these cytokines and chemokines in neovascularization (Ponnalagu et al., 2017). Our results show that NFκB activation induced by HG correlates with previous reports in ARPE-19 cells and in type 2 diabetic retinas (Chen et al., 2013; Lim et al., 2012) and with the increment in proinflammatory cytokines (IL-6 and IL-8) and COX-2 mRNA levels (Fig. 9). 253

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Fig. 10. Effect of PLD1, PLD2, MEK/ERK and COX-2 inhibitors on caspase-3 cleavage and cell viability in HG-exposed RPE cells. ARPE-19 cells were pre-incubated with 0.05% DMSO (NG and HG conditions), 0.15 μM PLD1i, 0.5 μM PLD2i, 10 μM U0126 or 10 μM celecoxib for 1 h before HG (16.5 mM) or NG exposure for 72 h. A) Caspase-3 cleavage was assayed as described in Fig. 2. Representative images from three independent experiments show cleaved caspase-3 staining (first row), DAPI staining (second row) and the merge image (third row). Scale bar = 10 μm. B) Bar graph shows nuclear cleaved caspase-3 fluorescence intensity expressed as arbitrary units (AU) with respect to control condition (NG). C) Cell viability was evaluated using MTT reduction assay. Data represent the mean value ± SD of three independent experiments. Results are expressed as arbitrary units (AU) with respect to control condition (NG). Asterisks (*) indicate significant differences with respect to NG (***p < 0.001; **p < 0.01; *p < 0.05), Number signs (#) indicate significant differences with respect to HG (###p < 0.001; ##p < 0.01).

classified into several groups depending on their cellular location, calcium dependency and substrate specificity (Vasquez et al., 2018). PLA2 catalyzes the hydrolysis of membrane phospholipids releasing lysophospholipids and free fatty acids, such as arachidonic acid (ARA, 20:4 n-6) and docosahexaenoic acid (DHA, 22:6 n-3). ARA and DHA are substrates for the synthesis of more potent lipid mediators, named eicosanoids and docosanoids respectively (Tallima and El, 2018; Asatryan and Bazan, 2017; Bazan, 2007). Differential activation of PLA2 isozymes induced by OS promoted either protection or damage in an in vitro model of retinal macular degeneration (Rodriguez et al., 2012, 2013). It has been reported that HG can directly damage pericytes through activation of PLA2/COX-2/VEGF-A pathway (Giurdanella et al., 2015). In a similar manner, in human retinal endothelial cells HG and advanced glycation end-products (AGEs) induce cell damage via ERK/cPLA2/COX-2/PGE2 inflammatory pathway (Giurdanella et al., 2017). Although the role of PLA2 has not been elucidated in RPE exposed to HG, PLA2 may participate in the inflammatory response either through the release of ARA, substrate for the PLD-induced COX-2, or through the release of DHA and the consequent docosanoid synthesis. Regarding PLA2 in RPE, a significant role of intracellular PLA2-VIA subtype in POS phagocytosis and RPE cell survival has been demonstrated (Kolko et al., 2007, 2014). Furthermore, in RPE cells the

Fig. 11. Schematic view of signaling events elicited by HG in RPE cells. Dashed arrows and question marks indicate possible but unstudied mechanisms. 254

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pathway in RPE cells exposed to HG. In addition, our results underline the importance of studying the role of classical PLDs and the effect of their selective inhibitors in other retina cell types exposed to inflammatory conditions.

docosanoid neuroprotectin D1 (NPD1) is formed through various enzymatic steps that involve PLA2 followed by a 15-lipoxygenase-like enzyme and NPD1 inhibits OS-induced RPE apoptosis and cytokinetriggered COX-2 induction (Bazan, 2006; Mukherjee et al., 2004). Results from the present work also show that sustained exposure (72 h) to HG significantly increases cleaved caspase-3 content and reduces mitochondrial functionality accordingly with increased ROS generation (Figs. 1 and 2). Once again, these effects are glucose-specific since they are not induced by Man. In response to elevated intracellular glucose levels the polyol pathway is activated for glucose metabolism in RPE cells. In this pathway the enzyme aldose reductase reduces glucose to sorbitol which is metabolized to fructose by the enzyme sorbitol dehydrogenase (Lorenzi, 2007). Thus, intracellular sorbitol accumulation can induce osmotic damage of RPE cells (Willermain et al., 2018), although the accumulation of intracellular sorbitol may be accompanied by reciprocal decreases in other intracellular osmolytes, such as myo-inositol, betaine and glycerophosphorylcholine in order to maintain osmotic equilibrium (Henry et al., 1993). On the contrary and as stated above, mannitol is considered an impermeable hexose, that does not undergo facilitated transport and is therefore not metabolized by the cell (Henry et al., 1993, 2000). The lack of effect after prolonged periods of incubation with Man (24 and 72 h) indicates that long term events induced by HG are mediated by mechanisms different to extracellular hyperosmolarity, possibly through AGE receptor-triggered signaling pathways. PLD1 and PLD2 inhibitors were observed to have the ability to prevent caspase-3 cleavage and reduction in cell viability induced by HG (Fig. 10). The same protective effect was achieved with MEK/ERK1/2 pathway inhibitor (U0126) and with COX-2 inhibitor (celecoxib) (Fig. 10), thus demonstrating once again that classical PLDs are upstream ERK1/2 phosphorylation and COX-2 expression. Furthermore, the protective effect of celecoxib suggests that COX-2 products affect RPE cell viability in an autocrine or paracrine manner. A similar protective effect of celecoxib was observed in LPS-challenged RPE cells (Mateos et al., 2014). Another essential function of the RPE is to form the outer BRB. Although inconsistent results regarding this barrier function were obtained when transepithelial electrical resistance (TEER) or dextran permeability was measured in ARPE-19 cells treated with HG concentrations mimicking severe hyperglycemia (Pavan et al., 2014; Trudeau et al., 2011), the early role exerted by PLDs in the modulation of the inflammatory response and the secretome in RPE cells exposed to HG suggest their participation in the long-term injury effects promoted by hyperglycemia such as BRB disruption and DR development. A very limited number of pharmacological tools are currently available for DR treatment. Even though results obtained in RPE cell cultures cannot be directly extrapolated to what occurs in the retina in vivo and further experiments in diabetic animal models are certainly needed to fully elucidate the role of classical PLDs in DR, our findings contribute to the understanding of the molecular mechanisms underlying the pathogenesis of this retinal disease and open new avenues for the development of future pharmacotherapies.

Funding This work was supported by grants from Fundación Florencio Fiorini (Subsidio para investigación en ciencias biomédicas 2016), Comisión de Investigaciones Científicas de la Prov. de Buenos Aires (CIC), Universidad Nacional del Sur [UNS, PGI 24/B226 8388], Consejo Nacional de Investigaciones Científicas y Técnicas [CONICET, PIP 112-201101-00437] and Agencia Nacional de Promoción Científica y Tecnológica [ANPCYT; PICTs 2013-0987, 2013-2317 and 2014-3352]. MVM, GMO, GAS and NMG are research. Conflicts of interest The authors declare that they have no conflicts of interest with the contents of this article. Author contributions PET, VB, GMO and MVM performed the experiments. MVM and GAS designed the experiments and supervised the study. MVM wrote the manuscript. NMG and GAS provided equipment and reagents and revised the manuscript. Acknowledgements Authors want to thank Dr. E. Rodriguez-Bouland (Weill Medical College of Cornell University, New York, USA) for kindly providing D407 cells. Authors are also grateful to Translator Viviana Soler for her technical assistance in controlling the use of the English language. Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.exer.2019.04.028. References Ahsan, H., 2015. Diabetic retinopathy–biomolecules and multiple pathophysiology. Diabetes Metab Syndr 9, 51–54. https://doi.org/10.1016/j.dsx.2014.09.011. S18714021(14)00088-5 [pii]. Asatryan, A., Bazan, N.G., 2017. Molecular mechanisms of signaling via the docosanoid neuroprotectin D1 for cellular homeostasis and neuroprotection. J. Biol. Chem. 292, 12390–12397. https://doi.org/10.1074/jbc.R117.783076. R117.783076 [pii]. Ban, Y., Rizzolo, L.J., 2000. Regulation of glucose transporters during development of the retinal pigment epithelium. Brain Res.Dev.Brain Res. 121, 89–95 S0165380600000286 [pii]. Bazan, N.G., 2006. Survival signaling in retinal pigment epithelial cells in response to oxidative stress: significance in retinal degenerations. Adv. Exp. Med. Biol. 572, 531–540. https://doi.org/10.1007/0-387-32442-9_74. Bazan, N.G., 2007. Homeostatic regulation of photoreceptor cell integrity: significance of the potent mediator neuroprotectin D1 biosynthesized from docosahexaenoic acid: the Proctor Lecture. Investig. Ophthalmol. Vis. Sci. 48, 4866–4881. https://doi.org/ 10.1167/iovs.07-0918. 48/11/4866 [pii]. Bradford, M.M., 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254. https://doi.org/10.1016/0003-2697(76)90527-3. S0003269776699996 [pii]. Brindley, D.N., Pilquil, C., Sariahmetoglu, M., Reue, K., 2009. Phosphatidate degradation: phosphatidate phosphatases (lipins) and lipid phosphate phosphatases. Biochim. Biophys. Acta 1791, 956–961. https://doi.org/10.1016/j.bbalip.2009.02.007. S1388-1981(09)00061-4 [pii]. Brown, H.A., Thomas, P.G., Lindsley, C.W., 2017. Targeting phospholipase D in cancer, infection and neurodegenerative disorders. Nat. Rev. Drug Discov. 16, 351–367. https://doi.org/10.1038/nrd.2016.252. nrd.2016.252 [pii]. Burkhardt, U., Beyer, S., Klein, J., 2015. Role of phospholipases D1 and 2 in astroglial proliferation: effects of specific inhibitors and genetic deletion. Eur. J. Pharmacol. 761, 398–404. https://doi.org/10.1016/j.ejphar.2015.05.004. S0014-2999(15) 30002-9 [pii].

5. Conclusion Summing up, the present work demonstrates that HG exposure induces PLD1 and PLD2 activation in RPE cells, leading to ERK1/2 activation, IκB degradation, NFκB nuclear translocation and expression of pro-inflammatory ILs and COX-2 and reduced cell viability (Fig. 11). Our findings constitute the first evidence that classical PLDs participate in the inflammatory response of RPE cells exposed to HG. Furthermore, the fact that PLD1 and PLD2 inhibition was observed to prevent HGinduced RPE cell damage and caspase-3 cleavage leads us to postulate these signaling pathways as potential therapeutic targets for treatment of DR and of other retinal inflammatory diseases. Further experiments are certainly needed to elucidate the upstream mechanisms leading to PLD activation and to fully understand the role of this signaling 255

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