Erythropoietin Protects Retinal Cells in Diabetic Rats Through Upregulating ZnT8 via Activating ERK Pathway and Inhibiting HIF-1α Expression

Abstract

Purpose: Zinc transporter 8 (ZnT8) was downregulated in hypoxic retina, which could be rescued by hypoxia-inducible factor-1α (HIF-1α) inhibition. Erythropoietin (EPO) protects retinal cells in diabetic rats through inhibiting HIF-1α as one of its mechanisms. We hence tried to explore the effect of EPO in regulating ZnT8 and protecting retinal cells in diabetic rats and possible mechanisms.

Methods: Diabetes was induced in Sprague-Dawley rats. Intravitreal injection of EPO was performed 1 month after diabetes onset. The CoCl₂-treated rat Müller cell line (rMC-1) was cotreated with EPO, soluble EPO receptor (sEPOR), digoxin, or U0126. Cell viability, cell death, and intracellular zinc level were examined. The expression of ZnT8, HIF-1α, AKT, and ERK was studied.

Results: In diabetic rat retinas, EPO significantly decreased HIF-1α expression and increased ZnT8 expression. In CoCl₂-treated rMC-1 cells, EPO increased cell viability and decreased intracellular zinc. Erythropoietin or digoxin could activate ERK pathway, downregulate HIF-1α, and upregulate ZnT8. The effect of EPO was abolished by sEPOR and U0126. Transient knockdown of ZnT8 increased intracellular zinc level, but not to a degree that would decrease cell viability or cause cell death.

Conclusions: In diabetic retinas, EPO maintains zinc homeostasis through activating the ERK pathway and downregulating HIF-1α, and thus upregulating ZnT8 expression. This work proposed a possible new protective mechanism for EPO in, and indicated a potential target for, the treatment of diabetic retinopathy.

Diabetes mellitus (DM), one of the most common chronic diseases, continues to increase in incidence in nearly all countries.¹ As a major complication of diabetes, diabetic retinopathy (DR) is one of the leading causes of legal blindness in the working population, yet there is a lack of effective interventions. Further understanding of the mechanisms for DR would be of great help in the development of better therapies. To date, DR has been regarded as both a neurodegenerative and a microvascular disease,^2–6 and the mechanisms might be related to oxidative stress, in which zinc plays a role.^7–9 

Zinc, an important nutrient and cofactor of numerous enzymes and transcription factors, plays an important role in maintenance of the structural and functional integrity of eukaryotic cells and tissues.^10–13 Zinc deficiency results in abnormal dark adaptation and age-related macular degeneration (AMD).^14–17 Zinc supplements, in clinical trials, show beneficial effects on patients with AMD by reducing visual loss, improving visual acuity and contrast sensitivity, and inhibiting complement activation, etc.^17–20 However, excessive accumulation of intracellular zinc is toxic to cells, possibly through overproduction of reactive oxygen species (ROS) and reactive nitrogen species (RNS) due to mitochondria dysfunction.^21–23 Cellular zinc homeostasis is maintained by three systems, that is, transporters of Zrt/Irt-like protein (ZIP) and zinc transporter (ZnT) families involved in uptake/extrusion of zinc; metallothioneins, which bind zinc for storage; and organelles (e.g., mitochondrion) that sequestrate zinc.^7,24–26 In contrast to the functions of ZIP proteins, ZnT proteins move zinc out of the cell or into organelles to decrease the cytosolic zinc concentration. 

As one of the zinc transporters, zinc transporter 8 (ZnT8) was first reported to be expressed in the pancreatic cells, playing a key role in mediating insulin secretion.^27–30 Loss of ZnT8 from pancreatic β cells reduces insulin content and compromises insulin release, which contributes to the pathophysiology of DM. The polymorphism in a genetic variant of ZnT8 (SLC30A8) is associated with increased risk of type 1 and 2 DM.^31,32 Furthermore, in patients with type 1 DM, ZnT8 is reported as an autoantigen, and ZnT8 autoantibodies, causing decreased ZnT8 levels in DM, have been detected in 60% to 80% of patients, making it a good diagnostic marker for the disease.^33,34 In addition to the pancreas, ZnT8 is also expressed in other tissues, such as human adipose tissue,³⁵ blood lymphocytes,³⁶ and retina.^37,38 

Deniro et al.³⁸ have demonstrated that ZnT8 expression is reduced in both the retina of mice with oxygen-induced retinopathy and in rat Müller cells (rMC-1) under hypoxia, while the retina injury could be rescued when ZnT8 is restored to its basal homeostatic level by hypoxia-inducible factor-1 (HIF-1) inhibitor YC-1. The increase of ZnT8 via HIF-1 inhibition has prompted us to test the effect of erythropoietin (EPO) on ZnT8 expression, since EPO has been found to downregulate HIF-1α via negative feedback when treating DR in rats.³⁹ Previous studies have shown that EPO is protective for the retinal pigment epithelium (RPE) cells, retinal neurons, and vascular cells in early diabetes,^40,41 and it is safe even when high doses are delivered into the eyes both in experimental and clinical studies.^42–44 In fact, the clinical trial of intravitreal injection of EPO into eyes with severe diffuse diabetic macular edema produces favorable visual outcomes and anatomic improvement.⁴⁵ 

Although protective mechanisms of EPO have been extensively studied,^{39,41,46–49} the changes of zinc level and ZnT8 expression in diabetic retina, as well as the effect of EPO on such systems, have not been reported yet. The present study demonstrated that ZnT8 expression in diabetic rat retina and hypoxic rMC-1 cells was downregulated at both the mRNA and protein levels, accompanied by overload of intracellular zinc. Erythropoietin rescued retinal cells and tissue under such stress or disease by maintaining the homeostasis of intracellular zinc, and its mechanism might involve the increase of ZnT8 expression via inhibiting HIF-1α and activating the ERK pathway. 

Materials and Methods

Reagents and Antibodies

Recombinant human erythropoietin (rHuEPO, S20010001) was purchased from 3S Bio (Shenyang, China). Soluble EPO receptor (sEPOR, 307-ER-050) was purchased from R&D (Shanghai, China). Digoxin (D102298) was purchased from Aladdin (Shanghai, China). Streptozotocin (STZ, S0130) and Thiazolyl Blue Tetrazolium Bromide (M2128) were purchased from Sigma-Aldrich Corp. (St. Louis, MO, USA). The DMEM Low Glucose Medium (SH30021.01B) was purchased from HyClone (Logan, UT, USA). Penicillin/streptomycin (15140155) and FluoZin-3 AM (F24195) were purchased from Invitrogen (Carlsbad, CA, USA). Pierce BCA Protein Assay Kit (23225) was purchased from Thermo Scientific (Shanghai, China). The primary antibodies against ZnT8 (16169-1-AP) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH; 60004-1-Ig) were purchased from Proteintech (Shanghai, China). The HIF-1α antibody was purchased from Abcam (ab463; Cambridge, UK). The primary antibodies against phospho-p44/42 MAP kinase (Thr202/Tyr204, 4370) and p44/42 MAP kinase (9107) were purchased from Cell Signaling Technology (Universal Biotech Company, Shanghai, China). Goat anti-rabbit IgG (H+L) (111-165-003) was purchased from Jackson ImmunoResearch Laboratories, Inc. (West Grove, PA, USA). Anti-mouse IgG (610-431-002) and anti-rabbit IgG (611-131-002) antibodies were purchased from Rockland Immunochemicals, Inc. (Limerick, PA, USA). The protein extraction Radio-Immunoprecipitation Assay (RIPA) buffer (P0013B) and U0126 (S1901) were purchased from Beyotime Institute of Biotechnology (Jiangsu, China). The TUNEL Apoptosis Detection Kit (Alexa Fluor 488, Cat. 40307ES60) was purchased from Shanghai Yeasen Biotechnology Co. Ltd. (Shanghai, China). Zinc chloride (ZnCl₂, A501003) was purchased from Shuji Biotechnology Co. Ltd. (Shanghai, China). 

Experimental Animals and Intravitreal EPO Injection

Male Sprague-Dawley rats weighing 120 to 160 g (Slaccas, Shanghai, China) were used. They were treated in accordance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and The Guides for the Care and Use of Animals (National Research Council and Tongji University). The protocol was approved by the Committee on the Ethics of Animal Experiments of Tongji University (Permit No. TJmed-010-32). The rats were randomly divided into three groups: normal control (N), diabetic rats (D), and diabetic rats treated with EPO (D+E). Diabetes was induced by intraperitoneal injection of STZ (60 mg/kg body weight dissolved in citrate buffer, pH 4.5) and the control rats received an equal volume of citrate buffer. The rats with blood glucose level exceeding 300 mg/dL for 3 consecutive days were considered as diabetic rats and were included in this study. 

Erythropoietin (16 U/eye, 2 μL) was injected, 4 weeks after diabetes onset, into the vitreous with a microsyringe (Hamilton, Reno, NV, USA) through a 30-gauge, 0.5-in. needle (BD Biosciences, Franklin Lakes, NJ, USA), which was inserted into the eye 2 mm posterior to the limbus at the temporal side. For normal control and untreated diabetes rats, the same volume (2 μL) of normal saline was injected. Four days after the injections, the rats were examined and killed, and the retinas were isolated for the assay. 

Rat Müller Cell (rMC-1) Culture

Transformed retinal Müller cell line (rMC-1) was kindly provided by Sarthy et al.⁵⁰ (Northwestern University, Chicago, IL, USA). The cells were maintained in normal glucose (1.0 g/L) Dulbecco's modified Eagle's medium (DMEM; Corning, Corning, NY, USA) containing 10% fetal bovine serum (Gibco, Shanghai, China) and 1% penicillin/streptomycin (Invitrogen) at 37°C with 5% CO₂ in a humidified incubator. When the cells reached ∼80% confluence in 10-cm dish, they were divided into three groups, that is, normal control (N), CoCl₂ (200 μM)–treated group (CC), and CoCl₂ (200 μM) + EPO (40 U/mL)–treated group (CC+E). Pretreatments of rMC-1 cells with sEPOR (250 ng/mL), digoxin (100 nM), or U0126 (10 μM), when needed, were carried out 1 hour before the incubation with CoCl₂. 

Cell Viability Assay

Cell viability of rMC-1 cells under different treatments was measured by using the methyl-thiazol-diphenyltetrazolium (MTT) assay. The rMC-1 cells were seeded on 96-well plates at a density of 1.0 × 10⁴ cells per well. To measure cell viability of rMC-1 cells incubated with CoCl₂ with or without EPO, the cells were transferred into the medium containing CoCl₂, with or without EPO (0, 10, 20, 40, 80, and 160 U/mL) for 24 hours. The cells were washed with 1X phosphate-buffered saline (PBS), and then incubated with MTT (0.5 mg/mL) in PBS for 4 hours. The media were then removed and the formazan crystals produced in the wells were dissolved in 100 μL dimethylsulfoxide (DMSO). The absorbance was measured at 570 nm with background subtraction at 630 nm by using a microplate spectrophotometer (Tecan, Crailsheim, Germany). For the examination of cell viability in rMC-1 cells treated with ZnCl₂ or in combination with EPO, the same procedure was used and the concentration of ZnCl₂ ranged from 1 to 200 μM. The cell viability was expressed as the percentage of the untreated control, which was defined as 100% for each experiment. 

RNA Extraction and Real-Time PCR

Total RNA was extracted from rMC-1 cells and rat retinas. Reverse transcription (RT) was performed and real-time PCR was carried out by using SYBR Green Real-Time PCR master mix (Toybo, Osaka, Japan). The primers were designed by using Primer Premier Version 5.0 software and were ordered from Shanghai DNA Biotechnology Co. Ltd. (Shanghai, China). The primers for ZnT8 were 5′-AAGTGGAGACTCTGTGCTGCTTCA-3′ (sense) and 5′-GGCCTCGATGACAACCACAAAGAA-3′ (antisense). The primers for HIF-1α were 5′-ACTATGTCGCTTTCTTGG-3′ (sense) and 5′-GTTTCTGCTGCCTTGTAT-3′ (antisense). The primers for β-actin were 5′-GTAAAGACCTCTATGCCAACA-3′ (sense) and 5′-GGACTCATCGTACTCCTGCT-3′ (antisense). The primers were verified by RT-PCR; the products were fractionated electrophoretically in 1.5% agarose gel in 1X Tris-acetate-EDTA buffer and imaged by gel image system (Tanon-3500; Tanon, Shanghai, China). 

Protein Extraction and Western Blot

The retinas and rMC-1 cells were lysed in RIPA buffer on ice. After 15-second ultrasonic wave treatment, the samples were placed on ice for 30 minutes before centrifugation. Protein concentrations were determined with Pierce BCA Protein Assay Kit (Thermo Scientific). Equal amounts of protein were resolved on 10% SDS–polyacrylamide gels and transferred electrophoretically onto nitrocellulose membranes (Bio-Rad, Shanghai, China). The membranes were blocked in 5% PBS-buffered nonfat milk at room temperature for 30 minutes, and then separately incubated with antibodies against ZnT8 (1:500), HIF-1α (1:500), phospho-p44/42 MAP kinase (1:1000), p44/42 MAP kinase (1:1000), or GAPDH (1:5000), overnight at 4°C. After being washed three times with 0.1% PBS-buffered Tween-20 (PBST), the membranes were incubated with the corresponding secondary antibodies (anti-rabbit or anti-mouse) at room temperature for 2 hours, followed by other washes with PBST (three times), and then visualized by chemiluminescence or Odyssey infrared imaging system (LI-COR Biosciences, Lincoln, NE, USA). After detection with ERK phospho-specific antibody, the blot was stripped with striping buffer (Beyotime stripping buffer) and reprobed with p44/42 MAP kinase antibody (1:1000). The optical density of each band was determined by using Quantity One software (Bio-Rad), and the densitometric values for the proteins were normalized by GAPDH. 

Immunofluorescence of ZnT8 in rMC-1 Cells

The rMC-1 cells were fixed in cold methanol for 1 minute, permeabilized with 0.5% Triton X-100 in PBS for 10 minutes, and blocked with 1% BSA and 0.2% Triton X-100 in PBS for 30 minutes. Then, the cells were first incubated with anti-ZnT8 antibody (1:100) overnight at 4°C, and then, after three 5-minute washes in PBS, incubated with the corresponding secondary antibody (1:1000, anti-rabbit CY3) for 1 hour at room temperature. The cells were further incubated with 4′,6-diamidino-2-phenylindole (DAPI, 100 ng/mL) for 3 minutes, and washed three times (5 minutes each wash). The coverslides were mounted by using Dako fluorescent mounting medium (Shanghai, China) and visualized with confocal microscope (488 nm excitation, 495–532 nm emission band; LSM 710; Zeiss Microsystems, Inc., Wetzlar, Germany). 

Measurement of Intracellular Zinc

Intracellular zinc was detected with zinc indicator FluoZin-3 AM (excitation 494 nm/emission 516 nm; Invitrogen) using microscopy, and its stock solution (5 mM) was reconstituted in DMSO, stored at −20°C, and protected from light. The rMC-1 cells were first seeded on coverslides for 24 hours, and then incubated with FluoZin-3 AM (diluted to the final concentration 5 μM) at 37°C for 1 hour, followed by a brief wash in PBS. The coverslides were switched to fresh medium to continue the incubation for 30 minutes at 37°C and then incubated with DAPI for 5 minutes. Finally, the coverslides were washed extensively three times in PBS (5 minutes each wash) and then mounted by using Dako fluorescent mounting medium and visualized with confocal microscope equipped with ×60 objective (488 nm excitation, 495–532 nm emission band; LSM 710). 

In Situ Detection of Cell Death

After rMC-1 cells were incubated with ZnCl₂ (125 μM) alone or in combination with EPO (40 U/mL) for 24 hours, in situ cell death was determined with TUNEL Apoptosis Detection Kit (Alexa Fluor 488) according to the manufacturer's instruction. Positive controls were treated with grade I DNase-I for 10 minutes at room temperature before the labeling procedure. Negative controls were treated with 10 μL label solution, but incubated in the absence of the terminal transferase. The coverslips were rinsed three times with PBS after incubation and analyzed under fluorescence microscope (Leica, Wetzlar, Germany) with an excitation wavelength in the range of 450 to 490 nm. 

ZnT8 Knockdown With Small Interfering RNA (siRNA) in rMC-1 Cells

Six siRNAs targeting ZnT8 (siRNA-1 to siRNA-6) and a nontargeting siRNA (scrambled siRNA, served as the control) were synthesized by GenePharma (Suzhou, China). The sequence information for both the scrambled siRNA and the six ZnT8 siRNAs is listed in the Table. The transfection was performed by using Lipofectamine 2000 (Invitrogen) according to the manufacturer's instruction. Briefly, the rMC-1 cells, seeded in 12-well plates, were cultured overnight (30%–50% confluence) and then transfected with either scrambled siRNA or one of the six ZnT8 siRNAs (20 μM) in serum-free medium. Six hours later, the cells were transferred to the culture under DMEM containing 10% fetal bovine serum and 1% penicillin/streptomycin at 37°C with 5% CO₂ in a humidified incubator. Two days after transfection, the rMC-1 cells were used for subsequent experiments, including Western blot, light microscopy, MTT assay, TUNEL assay, and intracellular zinc detection. The intracellular zinc intensity was quantified with the software ImageJ (http://imagej.nih.gov/ij/; provided in the public domain by the National Institutes of Health, Bethesda, MD, USA). 

Table

 

Information for siRNA

 

Statistical Analysis

Data were expressed as the mean ± SE. The statistical analysis was performed by using least significant difference test after 1-way ANOVA; a P value of 0.05 or less was considered statistically significant. 

Results

Erythropoietin Upregulates ZnT8 Expression and Downregulates HIF-1α Expression in Retina of Diabetic Rat

Increased HIF-1α in retina is one of the important pathophysiological changes in DR and is one of the targets through which EPO exerts its protection.³⁹ The HIF-1 inhibitor is also reported to rescue retina injury and restore ZnT8 homeostasis.³⁸ To better understand the changes of ZnT8 and their relationship with HIF-1α in diabetic rat retinas and EPO for its effects, we examined the retinas for both ZnT8 and HIF-1α in diabetic rats with or without EPO treatment. As shown in Figure 1, the mRNA level of HIF-1α in diabetic retinas was significantly upregulated by 24.0% (n = 4, P < 0.01) when compared with that in normal control, but downregulated by 16.94% (Fig. 1A; n = 4, P < 0.05) after EPO treatment. The protein level of HIF-1α was also upregulated in diabetic retinas by 69.20% (n = 3, P < 0.05) in comparison with the control. It was downregulated by 68.42% (Fig. 1B; n = 3, P < 0.01) after EPO treatment, consistent with the trend of the change in mRNA. 

Figure 1

 

Expression of HIF-1α (A, B) and ZnT8 (C, D) in retinas of 4-week diabetic rats with or without intravitreal EPO treatment. The mRNA (A) and protein (B) expression levels of HIF-1α were upregulated; they were downregulated after EPO treatment. While the mRNA (C) and total protein (D) expression levels of ZnT8 were downregulated, both were upregulated after EPO treatment. Data were expressed as mean ± SE (n = 3). GAPDH was used as the loading control (*P < 0.05, **P < 0.01).

Opposite to the changes in HIF-1α, both the mRNA and protein levels of ZnT8 in diabetic retina were significantly decreased when compared with those in normal control, and they were increased after EPO treatment when HIF-1α was downregulated. The mRNA level of ZnT8 was downregulated by 32.25% (n = 6, P < 0.01) in diabetic retinas in comparison with the control, and upregulated by 26.28% (n = 6, P < 0.05) after EPO treatment as compared to nontreated diabetic group (Fig. 1C). The total ZnT8 protein was downregulated by 30.48% (n = 3, P < 0.01) in diabetic rat retina and was upregulated by 26.47% (n = 3, P < 0.05) after EPO treatment (Fig. 1D). Hence, like increased HIF-1, decreased ZnT8 might be also closely involved in the development of DR; upregulation of ZnT8 might be another important mechanism for EPO in protecting retinal functions in this disorder. 

Erythropoietin Upregulates ZnT8 Expressions in CoCl₂-Treated rMC-1 Cells and Rescues the Cells From Apoptosis

Hypoxia is an early event in diabetic retina.⁵¹ To confirm if hypoxia affects ZnT8 expression in diabetic retina, rMC-1 cells were incubated with CoCl₂ to mimic the hypoxic condition of the diabetic retina. The rMC-1 cells were treated with CoCl₂ (200 μM) for 3, 6, and 12 hours, and ZnT8 mRNA expression was examined with real-time PCR. As shown in Figure 2A, after CoCl₂ treatment, ZnT8 level decreased by 45.20% at 3 hours (n = 5, P < 0.01), 36.88% at 6 hours (n = 5, P < 0.05), and 49.90% at 12 hours (n = 5, P < 0.01). The cell viability was reduced by 11.85% (n = 8, P < 0.01) after CoCl₂ treatment for 24 hours, but it was significantly increased by EPO at concentrations of 20 U/mL, 40 U/mL, and 80 U/mL (n = 8, P < 0.05) (Fig. 2B). Erythropoietin at 40 U/mL was used in this study, since it increased the cell viability to a relative consistent level by 26.85% (n = 8, P < 0.05) as compared with CoCl₂-treated cells, which gave enough room to monitor the changes under different treatments. 

Figure 2

 

The ZnT8 mRNA levels and cell viability of CoCl₂-treated rMC-1 cells with or without EPO intervention. (A) ZnT8 mRNA expression levels, as examined by real-time PCR, in rMC-1 cells that were treated with CoCl₂ (200 μM) for different times. (B) The cell viability, as detected with the MTT method, of rMC-1 cells that were treated with 200 μM CoCl₂ and different concentrations of EPO for 24 hours. Data were expressed as mean ± SE (n = 6 in [A], and n = 8 in [B]); *P < 0.05, **P < 0.01). N, rMC-1 cells without treatment.

To verify the in vivo results as showed in Figure 1, the change in ZnT8 was examined in the CoCl₂-treated rMC-1 cells with or without EPO treatment. As compared with the normal control, the mRNA and protein levels of ZnT8 in CoCl₂-treated cells were significantly decreased by 40.81% (n = 9, P < 0.01) and 38.77% (n = 3, P < 0.01), respectively. They were respectively increased by 38.54% (n = 9, P < 0.01) and 34.46% (n = 3, P < 0.01) after EPO treatment when compared with those in the CoCl₂-treated group (Figs. 3A, 3B). 

Figure 3

 

Effects of EPO on ZnT8 expression in CoCl₂-treated rMC-1 cells and the possible involvement of ERK signaling. (A) ZnT8 mRNA expression in CoCl₂-incubated rMC-1 cells treated with or without EPO. (B) Protein level of ZnT8 in different treatment groups of cells. The concentrations of CoCl₂ and EPO were 200 μM and 40 U/mL, respectively. (C) Examination of ZnT8 in the cells with confocal microscope. Data were expressed as mean ± SE from nine (A) and three (B) independent experiments. GAPDH was used as the loading control (*P < 0.05, **P < 0.01). CC, CoCl₂-treated rMC-1 cells; CC+D, CoCl₂ + digoxin (100 nM) treatment; CC+E, CoCl₂ + EPO treatment; CC+E+Es, CoCl₂ + EPO + sEPOR (250 ng/mL) treatment; CC+E+U, CoCl₂ + EPO + U0126 (10 μM); N, normal control.

To further substantiate these observations, ZnT8 distributions in the three groups of rMC-1 cells were studied with confocal microscopy. As showed in Figure 3C, ZnT8 staining was strong at the cell membrane and visible in the cytoplasm in normal control cells. In CoCl₂-treated cells, the intensity of ZnT8 staining was significantly decreased and aggregated in the nucleus or close to the nuclear envelope. But such CoCl₂-induced intracellular ZnT8 reduction was evidently reversed by EPO treatment. 

It was also clear that the effect of EPO in upregulating ZnT8 in CoCl₂-treated cells could be abolished by U0126 and sEPOR (Fig. 3B), indicating the involvement of ERK signaling in the pathology and its intervention in this system. As shown in Figure 3B, when compared with the effect of EPO in the CoCl₂-treated rMC-1 cells (CC+E), ZnT8 expression was decreased by 34.20% when these cells were cotreated with U0126 (CC+E+U) (n = 3, P < 0.01) and 26.75% with sEPOR (CC+E+Es) (n = 3, P < 0.01). Another interesting observation was that digoxin, a HIF-1α inhibitor, was also able to rescue the rMC-1 cells from CoCl₂-induced injury. These data indicate that EPO/EpoR upregulated ZnT8 expression via the ERK pathway in diabetic or CoCl₂-induced hypoxic retina. 

Erythropoietin Decreases Intracellular Zinc Level in CoCl₂-Treated rMC-1 Cells and Protects the Cells From Zinc Ion–Induced Cell Death

Considering that ZnT8 was decreased under hypoxia and upregulated by EPO, and a function of ZnT8 is to maintain cellular zinc homeostasis, the intracellular zinc levels in these three groups of cells were measured with zinc indicator FluoZin-3 AM as shown in Figure 4; consistent with the changes of ZnT8 (Fig. 3C), intracellular zinc intensity was low in normal controls but was significantly increased when the cells were incubated with CoCl₂. When EPO was added to the CoCl₂-treated cells, the intracellular zinc intensity was significantly reduced. The intracellular zinc intensity was also reduced by EPO or digoxin or in combination. The effects of both EPO and digoxin were largely abolished by ERK inhibitor U0126 (Fig. 4), providing further support to the conclusion that EPO upregulated ZnT8 through ERK pathway. 

Figure 4

 

Examination of intracellular zinc in the rMC-1 cells with FluoZin-3 AM. The concentrations of CoCl₂, EPO, and digoxin were 200 μM, 40 U/mL, and 100 nM, respectively. Scale bars: 50 μm. CC+U, CoCl₂ + U0126 (10 μM); CC+D+U, CoCl₂ + digoxin + U0126 (10 μM); CC+E+D, CoCl₂ + EPO + digoxin; CC+E+U, CoCl₂ + EPO + U0126 (10 μM).

The abovementioned result indicated that hypoxic condition would result in the overdose of zinc in rMC-1 cells, which might cause cell death. To test the causal effect of the overdose of zinc on rMC-1 cells, the cells were treated with different concentrations of ZnCl₂ as a stressor, for 24 hours, and the cell viability was measured with the MTT method. As shown in Figure 5A, when compared with nontreated rMC-1 cells, ZnCl₂ treatment decreased the cell viability by 48.58% (125 μM; n = 4, P < 0.01), 93.28% (150 μM; n = 5, P < 0.01), and 99.93% (175 μM; n = 6, P < 0.01). Other concentrations of ZnCl₂ (1–100 μM) were also tested, but no obvious effect on cell viability (data not shown) was observed. When EPO (40 U/mL) was administered, the cell viability was significantly increased (Fig. 5A), for example, the cell viability of ZnCl₂ (125 μM)–treated cells was increased by 39.38% (n = 4, P < 0.05) by EPO. We also used ATP assay to estimate the number of viable cells because this assay is the fastest and most sensitive cell viability assay, and it is less prone to artifacts than other viability assay methods. As shown in Supplementary Figure S1, as compared with normal control, the viability in ZnCl₂-treated rMC-1 cells was reduced by 53.34% (n = 6, P < 0.05), while EPO treatment increased the viability by 27.73% (n = 6, P < 0.05) over the zinc-treated cells. These data showed similar pattern of cell viability as measured with the MTT method (Fig. 5A) and confirmed the results of cell viability obtained with the MTT method. 

Figure 5

 

The effect of exogenous zinc (ZnCl₂) on the rMC-1 cells and the intervention with EPO. (A) Cell viability assay, as measured with the MTT method, of the rMC-1 cells treated with different concentrations of ZnCl₂ or ZnCl₂ + EPO (40 U/mL). (B) Detection of apoptosis with TUNEL. The rMC-1 cells were treated with ZnCl₂ (125 μM) or ZnCl₂ (125 μM) + EPO (40 U/mL). The TUNEL-positive cells were labeled as green and the nucleus was counterstained with DAPI. Data were expressed as mean ± SE from four to six independent experiments (*P < 0.05, **P < 0.01). N, rMC-1 cells without treatment.

To further consolidate the effect of zinc and EPO on rMC-1 cells, cell apoptosis was detected with the TUNEL method (Fig. 5B). It was obvious that apoptosis increased in the ZnCl₂ (125 μM)–treated group but decreased after EPO treatment, indicating that EPO could protect rMC-1 cells from death caused by exogenous zinc ion. 

Erythropoietin Downregulates HIF-1α Expression in CoCl₂-Treated rMC-1 Cells

Our in vivo study showed that HIF-1α was significantly upregulated in diabetic retinas and its downregulation was an important mechanism for EPO to protect retinal cells in diabetic rats.³⁹ In this study, the effects and mechanism of EPO on HIF-1α were further explored in an in vitro hypoxic cell system, namely, CoCl₂-treated rMC-1 cells. The results showed that the mRNA and protein levels of HIF-1α in CoCl₂-treated cells (for 3 hours) were increased by 64.58% (Fig. 6A; n = 9, P < 0.01) and 47.0% (Fig. 6B; n = 3, P < 0.05), respectively, as compared with the control. When EPO was included in such CoCl₂-treated cell culture, the elevated mRNA and protein levels of HIF-1α were decreased by 33.43% (n = 9, P < 0.01) and 33.19% (n = 3, P < 0.01), basically back to normal levels. When digoxin (100 nM) was used in this system, the HIF-1α protein was also decreased by 42.18% (Fig. 6B; n = 3, P < 0.05). There was no significant difference between EPO and digoxin in the efficacy of downregulation of HIF-1α under this condition. However, EPO's effect on HIF-1α was abolished when sEPOR (250 ng/mL) was added into the culture, that is, the HIF-1α protein was increased by 42.55% (Fig. 6C; n = 3, P < 0.01) to a level similar to that of the CoCl₂ treatment group. U0126 was also tested for its blocking effect to HIF-1α expression, but it had no effect on HIF-1α expression (data not shown). 

Figure 6

 

Erythropoietin downregulates the expressions of HIF-1α in CoCl₂-treated rMC-1 cells. (A) The mRNA levels of HIF-1α in the rMC-1 cells under different treatments. (B, C) The protein levels of HIF-1α in those cells. The cells were preincubated with CoCl₂ for 3 hours before addition of EPO, digoxin, or EPO plus sEPOR. Data were expressed as mean ± SE from three independent experiments. GAPDH was used as the loading control *P < 0.05, **P < 0.01). CC, CoCl₂ (200 μM)–treated rMC-1 cells; CC+D, CoCl₂ (200 μM) + digoxin (100 nM) treatment; CC+E, CoCl₂ (200 μM) + EPO (40 U/mL) treatment; CC+E+Es, CoCl₂ (200 μM) + EPO (40 U/mL) + sEPOR (250 ng/mL) treatment.

To detect the DNA-binding activity of HIF-1α among three groups, we used the electrophoretic mobility shift assay (EMSA). As shown in Supplementary Figures S2A and S2B, when compared with negative control (NC, wild-type probe only, lane 1), the shift bands appeared in normal control group (N, lane 4), CC group (lane 5), and CC+E group (lane 6). The binding ability in CC group (lane 5) was blocked by the addition of a 100-fold molar excess of unlabeled probe (cold probe, lane 2), but was not blocked when a similar excess of mutant probe was added (lane 3). Interestingly, the intensity of shift band in CC group was much stronger than that in N group (lane 4, approximately 2.45-fold) but was reduced in EPO-treated group (lane 6, reduced by 21.78%). The EMSA revealed a marked increase in HIF-1α transcriptional activity in the rMC-1 cells exposed to CoCl₂, which was reduced by EPO. To further confirm the changes of HIF-1α in the above three groups, immunostaining of HIF-1α was performed. The data showed that the intensity of HIF-1α was greatly increased and most HIF-1α protein translocated into the nucleus in the CC group compared with N group. After EPO treatment, the intensity of HIF-1α was reduced in both the nucleus and cytoplasm (Supplementary Fig. S2C). 

These findings indicate that, under CoCl₂ treatment, more HIF-1α protein translocated to the nucleus with increased transcriptional activity, while EPO reduced not only the level of total HIF-1α protein (Fig. 6), but also the amount of nuclear HIF-1α, and decreased its transcriptional activity (Supplementary Fig. S2). 

Erythropoietin and Digoxin Upregulate Phospho-ERK Expression in CoCl₂-Treated rMC-1 Cells

To understand the molecular mechanisms involved in ZnT8 regulation, we examined both AKT and ERK pathways, since they are the key signaling pathways in EPO/EPOR-mediated retinal protection.^41,46,50 The present result ruled out the involvement of AKT pathway in the response of rMC-1 cells to CoCl₂ treatment (for 3 hours) with or without EPO treatment because no difference was found among the three groups in term of the ratio of phospho-AKT to total AKT (Fig. 7A). However, the level of activated phospho-ERK (p-ERK) was reduced in the CoCl₂-treated rMC-1 cells but maintained at normal level when the cells also were treated with EPO (Fig. 7B). In comparison with the normal control, the ratio of p-ERK to total ERK (t-ERK), p/t-ERK, in CoCl₂-treated cells was reduced by 31.26% (n = 6, P < 0.01) for p/t-ERK-44 and 29.68% (n = 6, P < 0.01) for p/t-ERK-42. When the CoCl₂-treated cells were cotreated with EPO, the ratio of p/t-ERK was increased by 39.13% (n = 6, P < 0.01) for ERK-44, and 39.82% (n = 6, P < 0.05) for ERK-42 (Fig. 7B). The action of EPO on ERK pathway was further confirmed by ERK inhibitor U0126 (10 μM), which significantly decreased the level of p/t-ERK (Fig. 7B). 

Figure 7

 

Effects of EPO on p-ERK and phosphor-AKT (p-AKT) in CoCl₂-treated rMC-1 cells. (A) The change of p-AKT and total AKT (t-AKT) in CoCl₂-treated rMC-1 cells with or without EPO intervention (n = 2). (B, C) The changes of p-ERK and t-ERK in rMC-1 cells under different treatments. Data were expressed as mean ± SE from six (B) and three (C) independent experiments. Total AKT and t-ERK were used as the loading control (**P < 0.01). CC+D, CoCl₂ + digoxin (100 nM) treatment; CC+E, CoCl₂ + EPO (40 U/mL) treatment; CC+E+U, CoCl₂ + EPO (40 U/mL) + U0126 (10 μM).

It should not be ignored that the decreased level of p-ERK in the CoCl₂-treated rMC-1 cells was also reversed by digoxin, which increased the ratio of p/t-ERK by 49.27% for ERK-44 (n = 3, P < 0.01) and 49.20% for ERK-42 (n = 3, P < 0.01) (Fig. 7C). 

ZnT8 Knockdown Results in the Increase of Intracellular Zinc Level

To learn more about the function of ZnT8, we knocked down endogenous ZnT8 in rMC-1 cells with siRNAs. As showed in Fig. 8A, in comparison with the scrambled siRNA–treated group, the expression of ZnT8 protein in five of the six ZnT8 siRNA–treated groups was reduced by 52.70% (siRNA-1; n = 3, P < 0.05), 43.83% (siRNA-3; n = 3, P < 0.01), 47.15% (siRNA-4; n = 3, P < 0.01), 50.26% (siRNA-5; n = 3, P < 0.01), and 46.17% (siRNA-6; n = 3, P < 0.01). ZnT8 in siRNA-2 was reduced by 29.49%, but not to a statistically significant level (n = 3, P > 0.05). In ZnT8 siRNA knockdown groups, the intensities of intracellular zinc in rMC-1 cells were also increased (Fig. 8B). The quantified examination of the fluorescence intensity of intracellular zinc partially supported such a trend. When compared with those in the scrambled siRNA–treated cells, the fluorescence intensities in the six ZnT8 siRNA–treated groups were increased by 6.59% to ∼62.62%, but they only reached statistically significant levels in two of the six groups owing to the relative larger variation (data not shown). These results indicated that ZnT8 reduction should be related to the increase in intracellular zinc level, or at least as one of the contributors. Silencing ZnT8 in vitro for only a short time may not be enough to increase the zinc level to a toxic level, since the significant reduction of ZnT8 was confirmed only in some of the siRNA-treated cells, and no evident changes in cell morphology and cell viability were observed (data not shown). 

Figure 8

 

Effect of ZnT8 siRNA knockdown on intracellular zinc level in rMC-1 cells. (A) Western blot analysis of ZnT8 protein expression in the cells treated with scrambled siRNA (scrambled) and six ZnT8 siRNAs (siRNA-1 to siRNA-6). The quantitative data were plotted from three independent experiments (*P < 0.05, **P < 0.01). (B) The representative image of intracellular zinc detection (green) with FluoZin-3 AM in both scrambled siRNA–treated and six ZnT8 siRNA–treated groups. The nucleus was counterstained with DAPI (blue). Scale bars: 50 μm.

Discussion

Diabetic retinopathy has been considered as a microangiopathy and neuronopathy, involving both blood–retinal barrier (BRB) breakdown and retinal neuron apoptosis.^2–6 Our previous studies have demonstrated that EPO has protective effects on both BRB and retinal cells, and its mechanisms involve neuroprotection, neurotrophic effect, anti-inflammation, and BRB maintenance.^{39,41,46–49} In addition to these mechanisms, whether EPO is involved in the regulation of intracellular zinc homeostasis in DR⁵³ merited the present study. 

Zinc is the second most abundant trace element in the human body and is present at high levels in the RPE/choroid and neural retina.⁷ Zinc plays important roles in both retinal physiology and pathology, and is involved in various retinal functions such as phototransduction, visual cycle, and the process of neurotransmission.⁷ Even though zinc treatment is effective for diabetic rats at an early stage,^9,53 deregulation or overload of cytosolic zinc, especially at late stage of diabetic retinopathy or under hypoxic conditions, might be harmful to the cells owing to excess ROS and RNS generation and mitochondrial dysfunction.^21–23 Therefore, intracellular zinc homeostasis is the key to maintain normal functions of cells. In retina, zinc homeostasis is maintained by its transporters as well as zinc-binding proteins.^7,38 Zinc importers (Zip1, Zip2, and Zip12), as well as transporters ZnT6 and ZnT7, are detected in RPE cells, and ZnT3 is found in Müller cells, RPE cells, outer limiting membrane, inner segment, outer plexiform layer, inner nuclear layer, inner plexiform layer, and ganglion cell layer. ZnT7 is also found in photoreceptors, amacrine, and ganglion cells, while ZnT8 is widely distributed throughout the retina. As one of the zinc transporters, ZnT8 plays an important role in regulating zinc homeostasis, especially in pancreas islet cells.^28–30 In a mouse model of oxygen-induced retinopathy, ZnT8 expression is downregulated in the retina and is reversed by HIF-1α inhibition, accompanied by alleviated retinal insult.³⁸ Our previous work with diabetic rats has demonstrated that EPO reduces VEGF level via its feedback inhibition to HIF-1α in the retina.³⁹ These findings encouraged us to further explore whether EPO could maintain ZnT8 and if its mechanism would involve HIF-1α regulation in a diabetic rat model as well as in CoCl₂-treated rMC-1 cells. In this study, we found that ZnT8 expression was decreased in both diabetic rat retina and CoCl₂-treated rMC-1 cells, while EPO prevented ZnT8 reduction in such systems via HIF-1α inhibition and ERK pathway activation, thus maintaining intracellular zinc homeostasis. 

In 4-week diabetic rat retinas, we detected decreased expression of ZnT8 mRNA and protein, as well as increased HIF-1α expression (Fig. 1). To mimic the hypoxic condition in DR and examine whether hypoxia affects ZnT8 expression, we treated rMC-1 cells with CoCl₂, since CoCl₂ can stabilize HIF and thus mimics hypoxia.⁵⁴ Hypoxia-inducible factor-1 is a transcription factor for cellular and tissue adaptation to low oxygen tension and has been implicated in regulating angiogenic and metabolic response to ischemia. Growing evidence indicates that HIF-1α plays a causative role in the pathogenesis of DR.^55–57 In both diabetic retina and CoCl₂-treated rMC cells, we found upregulated HIF-1α expression and downregulated ZnT8 expression. The increased HIF-1α was downregulated by either EPO (Figs. 1, 3, 6), consistent with our previous study,³⁹ or by the HIF-1α inhibitor digoxin. The decreased ZnT8 expression in diabetes was also reversed after EPO or digoxin treatment (Figs. 1, 3, 6). ZnT8 expression is also reported to decrease in both a mouse model of oxygen-induced retinopathy and in CoCl₂-treated rMC-1 cells, but it increases by treatment with another HIF inhibitor, YC-1.³⁸ Therefore, HIF-1α was possibly also involved in the regulation of ZnT8. Digoxin, having similar effect as EPO, also increased ZnT8 expression in CoCl₂-treated rMC-1 cells (Fig. 3B). These findings indicated that HIF-1α might be one of the targets for regulating ZnT8 expression. 

ZnT8 protein participates in zinc homeostasis maintenance by moving zinc out of the cell or into organelles, and thus decreases the cytosolic zinc concentration. Therefore, ZnT8 reduction would cause intracellular zinc accumulation. Even though zinc plays a vital role in many biological processes,^58,59 elevated zinc ion is toxic to neurons, glia, and other cells,^60–62 and has been reported to result in the overproduction of reactive oxygen species, cell cycle arrest at G2/M phases, disrupted homeostasis of calcium in photoreceptors, and thus it induces cell death.⁶³ Hence, it is easy to believe that the reduced ZnT8 expression in the retinal cells under hypoxia could lead to overload of intracellular zinc, and then cell death. Such a hypothesis was further confirmed by ZnT8 knockdown and detection of intracellular zinc, using FluoZin-3 AM, a zinc-selective indicator with high zinc-binding affinity and good selectivity.^64,65 When ZnT8 was transiently knocked down with siRNA in rMC-1 cells under normal culture conditions, the intracellular zinc level increased as expected (Fig. 8). It is understandable that no obvious phenotypes, such as lowered cell viability and increased cell death, could be observed after transient treatment. However, we believe that prolonged stimulation in the retina of a diabetic individual could significantly increase intracellular zinc through persistent ZnT8 silencing, and finally cause cell death. Under hypoxic condition, zinc intensity was significantly increased in CoCl₂-treated rMC-1 cells (Fig. 4), accompanied with decreased cell viability (Fig. 2), indicating that higher concentrations of intracellular zinc might contribute to the decreased cell viability. Direct stimulation of the rMC-1 cells with exogenous zinc (ZnCl₂) confirmed it could decrease cell viability, resulting in cell death (Fig. 5). Erythropoietin protected these cells from insults caused by zinc overdose, as shown by increased cell viability and reduced cell apoptosis (Figs. 2, 5), possibly through the increase of ZnT8 and thus the reduction of intracellular zinc levels (Fig. 4). 

In addition to HIF-1α inhibition, EPO also play roles through activating ERK and AKT pathways,^41,46 as based on our previous study in another in vitro system. In our previous study, we have used glyoxal to treat rat retinal organ culture, primary retinal neuron culture, as well as retinal cell line (R28 cell) culture, and have found that exogenous EPO could protect the retinal neuronal cells from glyoxal-induced cell death by regulating Bcl-xL/Bax and BAD via the ERK and Akt pathways.⁴⁶ However, in the present study with cultures exposed to hypoxic conditions, we found that only ERK, but not AKT, was activated after EPO treatment (Fig. 7). Western blot analysis showed that the ratio of phospho-ERK-1/2 to total ERK-1/2 was significantly decreased in hypoxic rMC-1 cells, while EPO treatment reversed this ratio. The upregulation of ZnT8 by EPO was abolished by ERK inhibitor U0126 (Fig. 3B), which was further supported by the increased intracellular zinc intensity when U0126 was added into the EPO and CoCl₂–cotreated rMC-1 cells (Fig. 4). Surprisingly, digoxin also had a similar effect to EPO, in terms of ERK activation, ZnT8 upregulation, as well as in lowering intracellular zinc levels (Figs. 3, 4, 7). 

Taken together, our work demonstrated a new complementary mechanism for DR development and a possible new protective mechanism for EPO in DR. In DR and CoCl₂-treated rMC-1 cells, increased HIF-1α decreased ZnT8 and caused the overload of intracellular zinc, which is toxic to cells. Erythropoietin, after binding to EPOR, possibly exerts its protective effect through downregulation of HIF-1α expression and activation of the ERK pathway, which partially promotes ZnT8 expression, thus lowering intracellular zinc level. It is certain that other molecules are also involved in the pathogenesis of DR and EPO's protective role; however, this finding suggested a novel complementary mechanism for EPO as a therapeutic agent for DR treatment. To fully understand the function of ZnT8 in the retina, an animal model of ZnT8 conditional knockout merits further study. 

Acknowledgments

We thank Debasish Sinha, PhD, and J. Samuel Zigler, PhD, from Wilmer Eye Institute (The Johns Hopkins University School of Medicine, Baltimore, MD, USA) and Weiye Li, MD, PhD, from the Department of Ophthalmology (Drexel University College of Medicine, Philadelphia, PA, USA) for critical reading and discussion during the manuscript preparation. 

Supported by the Key State Basic Research Development Program of China (2012CBA01308 and 2013CB967501), National Natural Science Foundation of China (81570852 and 31171419), National High Technology Research and Development Program of China (2012AA020906), and Shanghai Pujiang Program (15PJ1408700). 

Disclosure: G. Xu, None; D. Kang, None; C. Zhang, None; H. Lou, None; C. Sun, None; Q. Yang, None; L. Lu, None; G.-T. Xu, None; J. Zhang, None; F. Wang, None