Association of HMGB1 with oxidative stress markers and regulators in PDR

Vitreous samples and epiretinal membrane specimens

We studied undiluted vitreous fluid samples obtained from 47 patients with PDR during pars plana vitrectomy. The indications for vitrectomy were tractional retinal detachment or non-clearing vitreous hemorrhage. The severity of retinal neovascular activity was graded clinically at the time of vitrectomy using previously published criteria [51]. PDR was considered active if there were visible perfused new vessels on the retina or optic disc and within the tractional epiretinal membranes. PDR was considered inactive (involuted) if only non-vascularized, white fibrotic epiretinal membranes were present. Active PDR was present in 25 patients, and inactive PDR was present in 22 patients. The diabetic patients were 32 males and 15 females whose ages ranged from 27 to 72 years, with a mean of 52.1 ± 11.4 years. The duration of diabetes ranged from six to 30 years, with a mean of 16.2 ± 5.4 years. The total PDR group consisted of a subgroup of 17 patients who had insulin-dependent diabetes mellitus and another group of 30 patients who had non-insulin-dependent diabetes mellitus. The control group consisted of 19 patients who had undergone vitrectomy for the treatment of rhegmatogenous retinal detachment with no proliferative vitreoretinopathy. Controls were free from systemic disease. Epiretinal fibrovascular membranes were obtained from 11 patients with PDR during pars plana vitrectomy for the repair of tractional retinal detachment. Membranes were fixed for 2 h in 10% formalin solution and embedded in paraffin.

The study was conducted according to the tenets of the Declaration of Helsinki. All patients were candidates for vitrectomy as a surgical procedure, and all patients signed preoperative informed written consent and approved of the use of the excised epiretinal membranes and vitreous fluid for further analysis and clinical research. The study design and the protocol were approved by the Research Centre and Institutional Review Board of the College of Medicine, King Saud University.

Human postmortem eyes

Human postmortem eyes were obtained from 16 diabetic and 16 non-diabetic (control group) donors matched by age (67 ± 8 years and 68 ± 6 years, respectively). The time elapsed from death to eye enucleation was 3.4 ± 1.9 h. After enucleation, one eye of each donor was snap-frozen in liquid nitrogen and stored at -80 °C until they were assayed for mRNA and western blot analyses. The other eye was fixed in 4% paraformaldehyde and embedded in paraffin for the immunofluorescence study.

After thawing, the neuroretina and RPE were quickly harvested. The vitreous and neuroretina were removed and the RPE layer was then carefully peeled off from Bruch’s membrane using forceps (Dumont No 5, Sigma, Madrid, Spain) under the dissecting microscope (Olympus SZ61, Barcelona, Spain).

All ocular tissues were obtained and used in accordance with applicable laws of the Diabetes and Metabolism Research Unit and CIBERDEM (ISCIII), Vall d’Hebron Research Institute, Barcelona, Spain, and following the Declaration of Helsinki for research involving human tissue. In addition, this study was approved by the Hospital Ethics Committee of Vall d’Hebron, Barcelona, Spain.

Induction of diabetes and glycyrrhizin treatment

All procedures with animals were performed in accordance with the Association for Research in Vision and Ophthalmology (ARVO) statement for use of animals in ophthalmic and vision research and they were approved by the institutional animal care and use committee of the College of Pharmacy, King Saud University. Adult male Sprague Dawley rats 7–10 weeks of age (200−220 g) were overnight fasted and a single bolus dose of streptozotocin (STZ) 55 mg/kg in 10 mM sodium citrate buffer, pH 4.5 (Sigma, St. Louis, MO), was injected intraperitoneally. Equal volumes of citrate buffer were injected in age-matched control rats. Rats were considered diabetic if their blood glucose was greater than 250 mg/dl.

Diabetic rats were divided into two groups: the rats in group I received normal drinking water without any supplementation and those in group II (curative treatment scheme, after disease induction) received drinking water supplemented with the potent HMGB1 inhibitor glycyrrhizic acid [52] (150 mg/kg/day, Santa Cruz Biotechnology Inc., Santa Cruz, CA) immediately after the establishment of diabetes [32-35]. After fou r weeks of diabetes, the rats were euthanized by an overdose of chloral hydrate, the eyes were removed, and the retinas were isolated and frozen immediately in liquid nitrogen and stored at -80 °C until analyzed.

Intravitreal injection of HMGB1

Sprague Dawley rats (220–250 g) were kept under deep anesthesia, and a sterilized solution of recombinant HMGB1 (5 ng/5 μl; R&D Systems, Minneapolis, MN) was injected into the vitreous of the right eye as previously described [32-35]. For the control, the left eye received 5 μl of sterile PBS (10 mM sodium phosphate ,150 mM sodium chloride, pH 7.8±0.2). Each group had 7–10 rats. The animals were sacrificed four days after intravitreal administration, and the retinas were carefully dissected, snap frozen in liquid nitrogen, and stored at -80 °C until analyzed.

Cell culture

HRMECs were purchased from Cell Systems Corporation (Kirkland, WA) and maintained in complete serum-free media (Cat. No. SF-4Z0–500, Cell System Corporation) supplemented with “Rocket Fuel” (Cat No. SF-4Z0–500, Cell System Corporation), “Culture Boost” (Cat. No. 4CB-500, Cell System Corporation), and antibiotics (Cat. No. 4Z0–643, Cell System Corporation) at 37 °C in a humidified atmosphere with 5% CO₂. We used HRMECs up to passage 8 for all the experiments. About 80% confluent cells were starved in a minimal medium (medium supplemented with 0.25% “Rocket Fuel” and antibiotics) overnight to eliminate any residual effects of growth factors. Following starvation, HRMECs were either left untreated or treated with 100 ng/ml of cytokine-HMGB1, lipopolysaccharide free (Cat. No. REHM120, IBL International Corporation, Toronto, ON). Cells were harvested after 24 h and lysed in radioimmunoprecipitation assay (RIPA) lysis buffer (sc-24948, Santa Cruz Biotechnology, Dallas, TX) for western blot analysis.

Enzyme-linked immunosorbent assays for vitreous samples

Enzyme-linked immunosorbent assay (ELISA) kits for human sVAP-1 (Cat No: ab-119564) and human 8-OHdG (Cat No: ab-101245) were purchased from Abcam, Cambridge, UK. The ELISA kit for human HO-1 (Cat No: KCB3776) was purchased from R&D Systems and the ELISA kit for human HMGB1 (Cat No: ST5101) was purchased from IBL International GMBH, Hamburg, Germany. The ELISA plate readings were done using a Stat Fax-4200 microplate reader from Awareness Technology, Inc., Palm City, FL. The levels of human sVAP-1, 8-OHdG, HO-1, and HMGB1 in vitreous fluid samples were determined based on the standards and protocols provided by the manufacturers.

Western blot analysis of human vitreous fluid and rat retinas

Retinas and cells were homogenized in a western lysis buffer (30 mM Tris-HCl; pH 7.5, 5 mM EDTA, 1% Triton X-100, 250 mM sucrose, 1 mM sodium vanadate and protease inhibitor cocktail), the lysates were then centrifuged at 14,000 ×g for 15 min at 4 °C, and the supernatant was used for protein estimation and further analysis. Equal amounts of protein were subjected to sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) in 8%–10% gels and transferred onto nitrocellulose membranes (Bio-Rad Laboratories). To determine the presence of VAP-1 in the vitreous samples, equal volumes (15 μl) of vitreous samples were boiled in Laemmli’s sample buffer (1:1, v/v) under reducing conditions for 10 min and analyzed as described (6,10,11). Immunodetection was performed with the use of the anti-VAP-1 antibody (1:1000, ab196739, Abcam), anti-ICAM-1 antibody (1:500; sc 8439, Santa Cruz, Biotechnology Inc.), and anti-vascular cell adhesion molecule-1 (VCAM-1) antibody (1:500, sc 13,160, Santa Cruz, Biotechnology Inc.). Membranes were stripped and re-probed with an antibody against β-actin to evaluate sample processing and lane loading. Bands were visualized using a high-performance chemiluminescence machine (G: Box Chemi-XX8 from Syngene, Synoptic Ltd. Cambridge, UK), and the intensities were quantified using the GeneTools software (Syngene by Synoptic Ltd.).

Immunohistochemical staining of epiretinal membranes

For CD31 and α-SMA, antigen retrieval was performed by boiling the sections in citrate-based buffer (pH 5.9–6.1; BOND Epitope Retrieval Solution 1; Leica) for 10 min. For CD45, VAP-1, and 8-OHdG detection, antigen retrieval was performed by boiling the sections in Tris/EDTA buffer (pH 9; BOND Epitope Retrieval Solution 2; Leica) for 20 min. Subsequently, the sections were incubated for 60 min with mouse monoclonal anti-CD31 (ready-to-use; clone JC70A; Dako, Glostrup, Denmark), mouse monoclonal anti-CD45 (ready-to-use; clones 2B11+PD7/26; Dako), mouse monoclonal anti-α-SMA (ready-to-use; clone 1A4; Dako), rabbit polyclonal anti-VAP-1 antibody (1:100; ab135695, Abcam), and mouse monoclonal anti-8-OHdG (1:50; ab26842, Abcam). Optimal working concentrations and incubation times for the antibodies were determined earlier in pilot experiments. The sections were then incubated for 20 min with a secondary IgG conjugated with alkaline phosphatase. The reaction product of the enzyme was visualized by incubation for 15 min with the Fast Red chromogen, resulting in bright-red immunoreactive sites. The slides were then faintly counterstained with Mayer’s hematoxylin (BOND) Polymer Refine Red Detection Kit (Leica). Negative controls were identified by omission of the primary antibody from the staining protocol. Instead, the ready-to-use DAKO Real Antibody Diluent (Agilent Technologies product code S2022) was applied. To identify the phenotype of stromal cells expressing VAP-1, sequential double immunohistochemistry was performed as described [5,6,10,11].

Immunofluorescence for VAP-1 in the retinas of human subjects with diabetes mellitus

Paraffinized eyes were cut at a thickness of 7 μm. Sections were deparaffinized with xylene, rehydrated in ethanol, washed in PBS, and placed in an antigen-retrieval solution (Dako A/S, Glostrup, Denmark). Permanganate bleaching was used to eliminate the autofluorescence of the RPE. Sections were then incubated for 1 h with 2% BSA in 0.05% Tween in PBS to block unspecific binding and then incubated overnight at 4 °C with the anti-VAP-1 antibody (1:500; ab115574, Abcam, Madrid, Spain) or anti-collagen IV (1:200; ab6586, Abcam, Madrid, Spain). After washing, sections were incubated with Alexa Fluor 488 and 594 secondary antibodies (Molecular Probes, Invitrogen, Madrid, Spain) at room temperature for 1 h. Slides were coverslipped with a mounting medium containing 4′, 6-diamidino-2-phenylindole (DAPI) for the visualization of cell nuclei (Vector Laboratories, Palex, Sant Cugat del Vallés, Spain). Images were acquired with a confocal scanning microscope (FV1000, Olympus, Hamburg, Germany).

Protein extraction and western blot analysis of the VAP-1 expression in the retinas of human subjects with diabetes mellitus

RPE and neuroretina samples were extracted with a RIPA buffer (1% Triton X-100, 100 mmol/l Tris-HCl, pH 7.5; 150 mmol/l NaCl; 1.5 mmol/l EDTA, pH 8.0; 25 mmol/l NaF; 0.5 mmol/l Na₃VO₄; 0.1 mmol/l phenylmethylsulfonyl fluoride; and Complete protease inhibitors [Roche, Basel, Switzerland]) and homogenized by sonication. Protein concentrations were determined using bicinchoninic acid assay (Bio-Rad Laboratories, Madrid, Spain). Equivalent amounts (20 µg) of total protein extracts were resolved by 10% SDS-PAGE and transferred to ECL nitrocellulose membranes (Hybond, Amersham Pharmacia Biotech). The membranes were incubated with a primary antibody against VAP-1 (1:1000; ab115574, Abcam, Madrid, Spain) and further incubated with a peroxidase-conjugated secondary antibody (Bio-Rad Laboratories, Madrid, Spain). Membranes were stripped and re-probed with β-actin to evaluate the lane-loading control. Proteins were visualized using the enhanced chemiluminescence detection system ECL (Advansta, CA).

RNA extraction and quantitative real-time PCR of the VAP-1 expression in the retinas from human subjects with diabetes mellitus

The human neuroretina (NR) and RPE of isolated eyecups from human donors were harvested by dissection under the microscope. Total RNA was extracted from isolated retinal tissues using the Trizol reagent (Invitrogen, Barcelona, Spain) according to the manufacturer’s instructions. RNA quantity was measured on a NanoDrop spectrophotometer and integrity was determined on an Agilent 2100 Bioanalyzer. Reverse transcription was performed with a High Capacity Kit (Applied Biosystems, Madrid, Spain) with random hexamer primers. RT-PCR was performed with the use of the SYBR Green PCR Master Mix (Applied Biosystems, Warrington, UK) in the 7.900 HT Sequence Detection System with 384-well optical plates. The specific primers were for hVAP-1: 5′-CAA TGA GAC CAT TGC TGG AA-3′ and 5′-TGT CCT CTG CAT GTG GGA TA-3′. For each sample, qPCR was performed in triplicate and relative quantities were calculated, and the ABI SDS 2.0 RQ software and the 2^−ΔΔCt analysis method with human β-actin (as the endogenous control; β-actin forward primer, 5′-TGG AGA AAA TCT GGC ACC AC-3′ and reverse primer, 5′-GAG GCG TAC AGG GAT AGC AC-3′) were used.

Cell adhesion assay

The adherence of human leukocytes to stimulated HRMEC monolayers was assessed. We used the CytoSelect Leukocyte-Endothelium Adhesion Assay kit (Cat. No. CBA-210, CELL BIOLABS, INC.) and followed the provided protocol exactly. Briefly, 1×10⁵ HMRECs were seeded into each well of attachment factor-coated 96-well plates in 200 μl of a completely serum-free medium and grown for 48 h. Confluent monolayers of endothelial cells were starved overnight in a minimal medium. Starved HRMECs were left untreated or treated with different doses (2 ng/ml to 200 ng/ml) of cytokine-HMGB1, lipopolysaccharide free (Cat. No. REHM120, IBL International Corporation) or with 1 ng/ml recombinant human tumor necrosis factor-α (TNF-α, positive control; Cat. No. 210-TA, R&D Systems) for 24 h.

In the meantime, leukocytes were isolated from healthy volunteers by Ficoll-HISTOPAQUE-1077 (Cat. No. 10,771, Sigma-Aldrich) density gradient centrifugation. Briefly, in 15 ml conical tubes, 5.4 ml of the Ficoll HISTOPAQUE-1077 reagent was added and on top of this, 7.5 ml of 1:1 blood-PBS was slowly layered. Thereafter, the contents were centrifuged at 595 ×g for 30 min at room temperature and the buffy coats were carefully pipetted off into other conical tubes. Cells were pelleted by centrifugation at 393 ×g for 10 min and they were resuspended in Dulbecco’s Modified Eagle Medium (DMEM) with 10% fetal bovine serum (FBS) and antibiotics. Peripheral blood leukocytes were labeled with the fluorescent LeukoTracker for 60 min at 37 °C, then counted by trypan blue exclusion assay, and immediately used for adhesion assay.

In total, 3×10⁵ labeled leukocytes were added to each well of the control or treated HRMEC monolayers for 30 min at 37 °C. After treatment with the washing buffer provided in the kit, the remaining adherent leukocytes were lysed in the lysis buffer of the kit. Fluorescence was measured using a spectraMax Gemini XPS (Molecular Devices, CA) with excitation and emission wavelengths of 485 nm and 538 nm, respectively.

Immunofluorescence staining and microscopy of HRMECs

HRMECs were cultured in coverslips in six-well plates and starved in minimal media overnight. They were left untreated or were treated with 100 ng/ml of cytokine-HMGB1, lipopolysaccharide free (Cat. No. REHM120, IBL International Corporation) for 24 h and were fixed with ice-cold acetone or 4% paraformaldehyde followed by blocking with 10% goat serum for 1 h. Without being washed, the cells were incubated with anti-HO-1 (1:100, sc-10789, Santa Cruz Biotechnology, Inc.), anti-VAP-1 (1:100, ab196739, Abcam), or anti-8OHdG (1:50, ab62623, Abcam) primary antibodies overnight at 4 °C. After washing, cells were incubated with secondary antibodies conjugated with appropriate fluorophores (Alexa Fluor 488 or 555) diluted in 10% goat serum (1:1000) for 1 h at room temperature. After washing, the cell-containing coverslips were counterstained with DAPI before being mounted on a glass slide with the use of the UltraCruz^TM Mounting Medium (sc-24941, Santa Cruz Biotechnology Inc.). Immunofluorescence microscopy was performed using an Olympus motorized system microscope (BX61, Olympus Corporation, Tokyo, Japan) at 20X or 40X magnification.

Statistical analysis

Data are presented as the mean ± standard deviation (SD). The non-parametric Kruskal–Wallis test was used to compare means among active PDR patients, inactive PDR patients, and non-diabetic control patients. The non-parametric Mann–Whitney U test was used to compare means from two independent groups. Spearman’s correlation coefficients were computed to investigate correlations between variables. A p value less than 0.05 was considered statistically significant, and SPSS version 20.0 for Windows (IBM Inc., Chicago, IL) was used for statistical analysis.

ELISA levels of HMGB1, 8-OHdG, sVAP-1, and HO-1 in vitreous samples and their relation with PDR activity and correlations

HMGB1, 8-OHdG, sVAP-1, and HO-1 were detected in all vitreous samples from patients with PDR and non-diabetic control patients. The mean HMGB1 level in PDR patients (7.59 ± 3.68 ng/ml) was significantly higher than the mean level in non-diabetic control patients (3.89 ± 3.22 ng/ml; p = 0.001). The mean 8-OHdG level in PDR patients (7.81 ± 4.86 ng/ml) was significantly higher than the mean level in controls (3.34 ± 1.43 ng/ml; p<0.0001). The mean sVAP-1 level in PDR patients (1,217 ± 590 pg/ml) was significantly higher than the mean level in controls (421 ± 144 pg/ml; p<0.0001). The mean HO-1 level in vitreous samples from PDR patients (236 ± 205 RFUs) was higher than that in control patients (178 ± 69 RFUs). However, the latter difference was not statistically significant (p = 0.805; Table 1).

Comparisons of the mean levels of HMGB1, 8-OHdG, sVAP-1, and HO-1 among active PDR patients, inactive PDR patients, and non-diabetic control patients were conducted with the Kruskal–Wallis test and the results are shown in Table 1. The mean levels differed significantly among the three groups for HMGB1 (p<0.0001), 8-OHdG (p<0.0001), sVAP-1 (p<0.0001), and HO-1 (p = 0.035). Pairwise comparisons (Mann–Whitney U test) indicated that the mean HMGB1 level was significantly higher in patients with active PDR than in inactive PDR and control patients (p = 0.025; p<0.0001, respectively). In addition, the mean HMGB1 level in patients with inactive PDR was significantly higher than the mean level in control patients (p = 0.019). For 8-OHdG, the mean level was significantly higher in patients with active PDR than in inactive PDR and control patients (p<0.0001 for both comparisons). For sVAP-1, the mean level was significantly higher in patients with active PDR than in inactive PDR and non-diabetic patients (p<0.0001 for both comparisons). In addition, the mean sVAP-1 level in patients with inactive PDR was significantly higher than the mean level in control patients (p = 0.002). For HO-1, the mean level in patients with active PDR was significantly higher than in inactive PDR patients (p = 0.012).

Significant positive correlations (Spearman’s correlation test) were found between the vitreous fluid levels of HMBG1 and the levels of 8-OHdG (r = 422; p = 0.001) and sVAP-1 (r = 0.354, p = 0.004). A significant positive correlation was observed between the vitreous fluid levels of sVAP-1 and 8-OHdG (r = 0.598; p<0.0001; Figure 1). In contrast, the correlations between the levels of HO-1 and levels of HMGB1, 8-OHdG, and sVAP-1 were not significant.

To obtain information about the molecular form of sVAP-1 and to compare its concentrations, we analyzed equal volumes (15 µl) of vitreous samples with the use of western blot analysis, and we confirmed that sVAP-1 was present in vitreous samples. Although the signal was rather weak, the protein in the samples migrated at the expected molecular weight, and the relative levels in PDR patients versus non-diabetic controls were in line with the ELISA data (Figure 2).

Immunohistochemical analysis of fibrovascular epiretinal membranes from patients with PDR

To identify the cell source of vitreous fluid sVAP-1 and 8-OHdG, fibrovascular epiretinal membranes from patients with PDR were studied by immunohistochemical analysis. No staining was observed in the negative control slides (Figure 3A). All membranes showed blood vessels that were positive for CD31 (Figure 3B), stromal cells expressing CD45 (Figure 3C), and stromal spindle-shaped cells expressing the myofibroblast marker α-SMA (Figure 3D). VAP-1 immunoreactivity was present in all membranes and was noted in vascular endothelial cells and stromal cells (Figure 4A-C). In addition, VAP-1 immunoreactivity was detected in stromal cells expressing CD45 and in spindle-shaped cells. In serial sections, the distribution and morphology of spindle-shaped cells expressing VAP-1 (Figure 4C) were similar to those of cells expressing α-SMA (Figure 3D). A double immunohistochemistry analysis confirmed that stromal cells expressing VAP-1 co-expressed CD45 (Figure 4D, E). Strong 8-OHdG immunoreactivity was detected in all cell types of the PDR fibrovascular epiretinal membranes, including endothelial and stromal cells. As well, 8-OHdG immunoreactivity was observed in the nuclear and cytoplasmic compartments of the cells (Figure 4F).

Effect of diabetes on the VAP-1 expression in retinas of human subjects with diabetes mellitus

The VAP-1 mRNA expression was higher in diabetic donors than in non-diabetic donors in the RPE (p = 0.03). On the other hand, the expression of VAP-1 mRNA did not differ significantly between non-diabetic donors and diabetic donors in the neuroretina (p = 0.81; Figure 5A). These findings were also confirmed at the protein level by western blot analysis (p = 0.04 for the RPE; p = 0.07 for the neuroretina; Figure 5B). VAP-1 immunofluorescence showed a higher expression of VAP-1 in the RPE of diabetic donors compared to non-diabetic donors (Figure 6A). A double immunofluorescence analysis revealed that VAP-1 immunostaining was co-localized with the basal lamina marker collagen IV in retinal vessels (Figure 6B).

Effect of diabetes and intravitreal administration of HMGB1 on the retinal expression of adhesion molecules

We tried to corroborate the findings in a preclinical rat model of streptozotocin-induced diabetes. The body weights of the diabetic rats were lower and their blood glucose levels were more than fourfold higher compared to the age-matched normal control rats (171 ± 20 versus 269 ± 27 g and 487 ± 31 versus 107 ± 10 mg/dl, respectively). Treatment of the diabetic rats with glycyrrhizin for one month did not change these metabolic variables in the diabetic rats (189 ± 29 versus 171 ± 20 g and 492 ± 28 versus 487 ± 31 mg/dl, respectively).

Western blot analysis of homogenized retinal tissue demonstrated that diabetes and an intravitreal injection of HMGB1 induced the significant upregulation of ICAM-1. Diabetes and an HMGB1 injection resulted in about 45% and 51% increases, respectively, in the ICAM-1 expression (Figure 7). On the other hand, the expressions of VCAM-1 and VAP-1 were not affected by diabetes or an intravitreal injection of HMGB1. Constant glycyrrhizin intake from the onset of diabetes significantly attenuated the diabetes-induced upregulation of ICAM-1 by about 44% (Figure 7), confirming that this compound acted in vivo.

HMGB1 induces leukocyte adhesion to HRMECs and the expression of cell adhesion molecules, 8-OHdG, and HO-1 in HRMECs

In the optimizing experiment, HRMECs were treated with different doses of HMGB1 (2 ng/ml to 200 ng/ml) for 24 h to activate the endothelial cells. Thereafter, LeukoTracker-labeled leukocytes were added and allowed to interact with the endothelial cells. The cultures were then gently washed and the remaining adhered leukocytes were quantified by measuring the amount of fluorescent labels in total cell lysates. We found that treatment of HRMECs with 100 ng/ml HMGB1 resulted in maximum adhesion of the leukocytes (Figure 8A). Therefore, we used this dose for all other in vitro experiments. We also used TNF-α (1 ng/ml) as a positive control. Our data revealed that 100 ng/ml HMGB1 significantly upregulated the adherence of human leukocytes to HRMECs (Figure 8B).

With the use of western blot analysis, we demonstrated that the treatment of HRMECs with HMGB1 (100 ng/ml) for 24 h resulted in an increased expression of ICAM-1 compared with the control medium. On the other hand, the expressions of VCAM-1 and VAP-1 were not affected by HMGB1 treatment (Figure 8C).

Immunofluorescence microscopy showed both cytoplasmic and membranous VAP-1 immunoreactivity in the untreated cells. The HMGB1-treated cells showed predominant membranous immunoreactivity (Figure 9A).

HO-1 and 8-OHdG are intracellular molecules that can be detected in extracellular body fluids, such as urine [39,50]. Above, we described changes in the levels of these molecules in vitreous fluids. To demonstrate the in situ regulation and production, we aimed to extend our findings with the use of immunohistochemical analysis. Immunofluorescence microscopy showed both nuclear and cytoplasmic 8-OHdG immunoreactivity in the untreated cells. HMGB1 treatment induced the upregulation of cytoplasmic and nuclear immunoreactivities (Figure 9B). Similarly, HMGB1 treatment induced the upregulation of HO-1 immunoreactivity in the cytoplasm of HRMECs (Figure 9C).