Receptor Protein Tyrosine Phosphatase Sigma (RPTP-σ) Increases pro-MMP Activity in a Trabecular Meshwork Cell Line Following Oxidative Stress Conditions

Abstract

Purpose: To elucidate the role of phosphatases in the eye drainage system by overexpressing the receptor tyrosine phosphatase sigma (RPTP-σ) in a human normal trabecular meshwork (NTM) cell line.

Methods: The efficacy, expression, and location of RPTP-σ were evaluated following its transfection in NTM cells (NTM^T) and in NTM control cells. The cells were also analyzed for viability, matrix metalloproteinase (MMP) activity, and phosphatase activity following oxidative stress conditions. Assays were conducted in the presence or absence of a specific RPTP-σ inhibitor.

Results: Transfection efficacy measurements revealed that RPTP-σ expression measured via GFP fluorescence was significantly higher (×3.8) in NTM^T cells than in control cells. Western blot analyses showed that RPTP-σ expression was significantly higher (×2.25) in NTM^T cells than in control cells. No significant differences were observed in cell viability between NTM^T and control cells after oxidative stress. We found that pro–MMP-2 and pro–MMP-9 showed a significantly higher activity (×2.18 and ×1.9; respectively) in NTM^T cells than in control cells. Serine/threonine phosphatase activity in NTM^T cells was significantly increased following oxidative stress. The specific phosphatase inhibitor PTP-IV inhibited 15% of the RPTP-σ expression in NTM cells and 31% in NTM^T cells. The activity of pro–MMP-9, pro–MMP-2, and MMP-9 was significantly inhibited (48%, 35%, and 78% respectively).

Conclusions: The findings indicate that RPTP-σ is expressed constituently in NTM cells and that oxidative stress changes the general phosphatase balance in NTM cells. In addition, the results show that expression levels of RPTP-σ affect the activity of various forms of MMP.

Today, glaucoma is the second leading cause of blindness worldwide.^1,2 In light of the expansion of the aged population in recent years, the demand for novel treatment approaches to primary open angle glaucoma (POAG) has increased. However, although POAG has been investigated for decades, its pathophysiology is not yet fully understood. 

Changes in various factors have been reported in the aqueous humor (AH) of POAG patients, including in the levels of cytokines,^3–5 matrix metalloproteinases (MMPs), and their inhibitors,⁶ endothelin, and nitric oxide (NO),⁷ to name a few. In addition, several markers of oxidative stress have been found in the AH of POAG patients^8–11 and in many animal models.^12–14 In the presence of oxidative stress, it has been shown that tumor necrosis factor-alpha (TNFα) and NO activate various cellular signaling pathways, including the mitogen-activated protein kinase (MAPK) pathway.¹⁵ Our group has previously described changes in MAPK-related proteins in the AH of a rat model of induced elevated ocular pressure^16,17; we found active kinase proteins in the AH, a finding that has led us to search the AH for phosphatases, which are known to be in balance with their kinase counterparts in various biological systems.¹⁸ 

In eukaryotic cells, protein phosphorylation plays a key role in regulating cellular processes that are initiated by various external effectors, including hormones, neurotransmitters, growth factors, and cytokines, to name a few. Phosphoregulation—the reversible phosphorylation and dephosphorylation of proteins by kinases and phosphatases—is a major part of cell metabolism, gene expression, and cell differentiation. In general, phosphoregulating enzymes are either specific to the protein residues serine/threonine or tyrosine, or they exhibit a dual specificity and can phosphorylate or dephosphorylate both serine/threonine residues and tyrosine residues. 

Protein tyrosine phosphatases (PTPs) are usually divided into four sub-groups: receptor PTPs (RPTPs), nonreceptor PTPs (nrPTPs), dual-specificity PTPs, and low M_r PTPs. The receptor PTPs are a family of transmembrane receptors that includes three subtypes (LAR, PTP-δ, and PTP-σ). Unlike nrPTPs, which are usually intracellular, RPTPs are typically found in the cell membrane. They possess an extracellular domain attached to two different PTP sites via a trans-membrane protein, suggesting that they play a role in mediating extracellular and intracellular signaling. The extracellular domain of RPTPs has an Ig-like and fibronectin-like structure, which is similar to that of adhesion molecules and suggests that they may be involved in regulating the connection between the cell and its extracellular matrix. For example, cadherin adhesion activity is highly sensitive to the phosphorylation of the tyrosine residue of β-catenin units. Following its phosphorylation, β-catenin dissociates from cadherin and the cell-cell adhesion is terminated. As cadherin/catenin phosphorylation is central to the adhesion functionality of cadherin, it is not surprising that RPTPs are involved in the regulation of the adhesion complex. 

Our understanding of the role that phosphatases play in the ocular system, in general, and in glaucoma, in particular, is limited. Serine/threonine phosphatases (namely, PPase2A and PPase2C) and tyrosine phosphatases were detected in the AH of POAG patients,¹⁹ and two studies demonstrated that phosphatase regulation depends on the redox state of the cell.^20,21 Taken together, these findings have led us to assume that the oxidative stress conditions that are found within the ocular anterior chamber affect parallel signaling pathways, in which kinases and phosphatases may participate. 

The aim of the present study, therefore, was to elucidate the effect of RPTP-σ on the activity of gelatinases (subtypes of MMPs) in the human normal trabecular meshwork (NTM) cell line under oxidative stress conditions. Such data will increase our currently limited understanding of the possible roles of phosphatases in the drainage system of the eye; furthermore, the data may help elucidate the changes in activity that phosphatases undergo as part of the signaling pathway activated by oxidative stress (as is evident in POAG patients), which ultimately changes the ability of the trabecular meshwork to resist AH outflow. 

Materials and Methods

Chemicals

Unless stated otherwise, all chemicals were obtained from Sigma-Aldrich (Jerusalem, Israel) and were of reagent grade. Chemicals used in this study included a 20% sodium dodecyl sulfate (SDS) solution (Amresco LLC, Solon, OH, USA), dimethyl sulfoxide (DMSO), isopropyl alcohol, methanol (Bio-Lab Ltd., Jerusalem, Israel), diethylpyrocarbonate-treated water, Dulbecco's modified Eagle's medium (DMEM) with 4.5 g/mL D-glucose without L-glutamine, fetal bovine serum (FBS), L-glutamine, streptomycin/penicillin, trypsin-EDTA 0.05% with phenol red, trypsin-EDTA 0.02% without phenol red (Biological Industries Israel Beit-Haemek Ltd., Beit-Haemek, Israel), an acrylamide:Bis solution (29:1), 30% ammonium persulfate, blotting-grade blocker, nonfat dry milk, glycine, Laemmli sample buffer, protein standards (Precision Plus; Bio-Rad Laboratories, Inc., Hercules, CA, USA), protein assay reagent, TEMED (N,N,N′,N′-tetramethylethylenediamine), tris-buffered saline, β-mercaptoethanol (Bio-Rad Laboratories, Inc.), PTP inhibitor IV (CAS 329317-98-8, Calbiochem, San Diego, CA, USA), luminol, gelatin, Coomassie Brilliant Blue G250 (Fluka BioChemika, Jerusalem, Israel), acetic acid glacial, sodium chloride (Frutarom, Haifa, Israel), OptiMEM-I reduced serum medium (GIBCO, Israel), Lipofectamine 2000 Transfection Reagent (Invitrogen, Israel), 6,8-difluoro-4-methylumbelliferyl phosphate (DiFMUP), DiFMU (Invitrogen), peroxidase-conjugated affinity pure bovine anti-goat IgG secondary antibody, β-actin secondary antibody (anti-mouse; both from Jackson ImmunoResearch Laboratories, PA, USA), goat anti-human RPTP-σ antibody and Northern Lights anti-goat IgG-NL493 (R&D Systems, Minneapolis, MN, USA) ethanol, hydrogen peroxide (Merck, Kenilworth, NJ, USA), Escherichia coli (JM109) competent cells (Promega, Beit Haemek, Israel), bacteriological agar, LB broth (Pronadisa; Laboratorios Conda, Madrid, Spain), and DAPI mounting medium (Fluoromount-G; Southern Biotech, AL, USA). 

Cell Cultures

Transformed normal (NTM-5) and glaucomatous (GTM-3) trabecular meshwork cell lines were derived from a 72-year-old man who had a medically controlled POAG at the time of death (GTM-3) and from an 18-year-old man without eye disease (NTM-5), both donated by A. F. Clark (Alcon Research, Ltd., Fort Worth, TX, USA).²² Cells up to the 40th passage were cultured in DMEM supplemented with 10% FBS, 1% penicillin/streptomycin, and 1% glutamine. All cells were maintained in a humidified incubator at 37°C in an atmosphere of 95% air and 5% CO₂. 

Transient Transfection of Plasmid DNA

Normal trabecular meshwork cells underwent a transient cotransfection with pcDNA3.1/Zeo(+) with the RPTP-σ sequence (a gift from Michel L. Tremblay, McGill University, Canada).²³ Two days prior to transfection, 2 × 10⁵, 2 × 10⁶, and 3 × 10⁶ NTM cells were seeded onto 96-, 24-, and 6-well plates, respectively, in a growth medium that did not contain antibiotics. As a control, similar numbers of untransfected NTM cells and GTM cells were seeded in different plates in growth medium. Normal trabecular meshwork cells were transfected for 6 hours with a transfection reagent (Lipofectamine 2000; Life Technologies, Carlsbad, CA, USA) according to the manufacturer's instructions. For the 24-well plate, a DNA:transfection reagent mixture (8 μg:2 μL; Life Technologies) in a 100 μL reduced serum medium (Opti-MEM I; Life Technologies) was added to 500 μL of medium without antibiotics (DNA content was 9 × pcDNA3.1:1 × Gr GFP in a plasmid, a gift from A. David; Ben-Gurion University of the Negev, Beer-Sheva, Israel). After 6 hours, the medium was replaced with a fresh original medium and the cells were incubated for additional 48 hours. The transfection and post-transfection incubation was in a humidified atmosphere of 95% air and 5% CO₂ at 37°C. Successfully transfected NTM cells are designated transfected NTM (NTM^T). 

Flow Cytometry

After 48 hours of post-transfection incubation, the cells were washed with PBS, detached with white trypsin, and centrifuged (1200g, 5 minutes) at room temperature. The supernatant was removed and the cells were suspended in 0.5 mL PBS. To estimate transfection efficacy, the percentage of fluorescent cells in the NTM^T and in the NTM cells cultures was measured by using flow cytometry (Guava easyCyte; EDM Millipore, Billerica, MA, USA) with 418-nm excitation and 488-nm emission. 

Protein Extraction

Cells were washed three times with ice-cold saline and scraped with a rubber policeman into a lysis buffer. The composition of the lysis buffer was altered based on the specific assay for which the cells were designated: for immune-blotting and zymography analysis: Tris-HCl 50 mM (pH = 7.5), EDTA 1 mM (pH = 7.5), EGTA 1 mM (pH = 7.5), NaF 50 mM, Triton X-100 0.1%, sodium vanadate 1 mM, and protease inhibitor cocktail 10 μL/1 mL buffer; for DiFMUP: Tris-HCl 50 mM (pH = 7.5), EGTA 0.1 mM, EDTA 1 mM, dithiothreitol 0.5 mM, and protease inhibitor cocktail 10 μL/1 mL buffer. Cells were homogenized at 450 rpm and sonicated for 15 seconds at 4°C in 40% amplitude (Vibra-Cell, Sonics, Newton, CT, USA). The resulting suspension was centrifuged at 12,000 g for 15 minutes at 4°C. The supernatant was collected and kept at −80°C for later analyses. 

Protein Content and Sample Preparation

Protein concentrations were measured by using the Bradford assay (Bio-Rad Laboratories) with bovine serum albumin as a standard.²⁴ For immunoblotting analyses, the samples were diluted with a sample buffer 1:2 (95% vol/vol Laemmli sample buffer, 5% vol/vol β-mercaptoethanol) and heated to 95°C for 5 minutes. For zymography analysis, samples were diluted with a sample buffer 1:3 (51% vol/vol 0.5 M Tris pH = 6.8, 0.08% wt/vol SDS, 40% vol/vol glycerol, 10% vol/vol H₂O, and a few drops of bromophenol blue). 

Western Blot Analysis

The assessment of RPTP-σ activity by the immunoblotting assay²³ involved seeding of NTM cells in two different 6-well plates. Protein aliquots (20–30 μg/lane) of the tested cells were separated with a 7.5% SDS-PAGE gel and transferred to nitrocellulose membranes (0.45 μm, Trans-Blot transfer medium; Bio-Rad Laboratories). The membranes were blocked with 5% BSA in TTBS (Tween Tris-buffered saline) for 1 hour at room temperature and then incubated with the primary antibody affinity-purified goat anti-human RPTP-σ antibody (1 μg/mL in TTBS plus 5% BSA) overnight at 4°C. After three washes with TTBS, the membranes were incubated for 1 hour at room temperature with a peroxidase-conjugated affinity purified bovine anti-goat IgG secondary antibody at 1:1000 in TTBS plus 3% BSA. Immunoreactivity was detected by using the EZ-ECL chemiluminescence detection kit (Biological Industries) followed by exposure to X-ray film (Fuji medical X-ray film; Fujifilm, Tokyo, Japan). As a control, actin levels were measured by using a mouse monoclonal anti-actin antibody diluted 1:50,000. The secondary antibody was an anti-mouse antibody diluted 1:20,000 in TTBS + 3% BSA. A semiquantitative analysis was performed by using the EZQuant-Gel 2.1 Software (Israel). 

Immunofluorescence Assay for RPTP-σ Activity

Cell medium was aspirated and the cells were washed with PBS, fixed for 8 minutes with methanol at −20°C, and blocked with 3% BSA for 1 hour. The cells were then exposed to a human RPTP-σ antibody antigen affinity purified polyclonal goat IgG (R&D Systems) that was diluted (15 μg/mL in 0.1% vol/vol Tween 20 in ×1 PBS containing 1% BSA) and incubated for 1 hour at room temperature. The cells were then washed carefully and incubated with anti-goat IgG-NL493 (1:200; Northern Lights, R&D Systems) containing 1% BSA for 1 hour at room temperature. Finally, the cells were washed and mounted in a mounting medium containing DAPI. Images were obtained with a confocal laser–scanning microscope with 360 nm (for DAPI, Olympus FluoView; Olympus Corp., Sinjuku, Tokyo, Japan) or 493 nm (for Cy2) excitation wavelengths. 

Exposure of NTM and NTM^T to H₂O₂

To model oxidative stress, the cell cultures were exposed to H₂O₂. Specifically, NTM and NTM^T cells (48 hours post-transfection) were incubated in a starvation medium (1% FCS) for 24 hours at 37°C in a 5% CO₂ atmosphere. The medium was then replaced with starvation medium containing 100 μM H₂O₂ and the cells were again incubated for 15 minutes, after which they were returned to the original growth medium. Protein extraction was performed at different times (0, 0.5, 1, 2, 4, and 24 hours) after the H₂O₂ exposure. Control cells were not exposed to H₂O₂, but were otherwise treated identically. 

MTT Assay for Cell Viability

The viability of NTM and NTM^T cells was assessed following exposure to oxidative stress by using the MTT assay.²⁵ The absorbance of each sample was measured at 570 nm by a microplate reader (model 680; Bio-Rad Laboratories). 

Zymography

For gelatin zymography, cell lysates (7 μg protein/well) were electrophoresed through a 12.5% SDS-PAGE gel polymerized with 0.1% gelatin. Gels were washed twice (1 hour each time) in 2.5% Triton X-100 and incubated in a developing buffer (50 mM Tris, 200 mM NaCl, 5 mM CaCl₂, and 0.02% Brij-35) for 18 hours at 37°C. Gels were stained with 30% methanol, 10% acetic acid, and 0.5% wt/vol Coomassie brilliant blue for 30 minutes, after which they underwent destaining.²⁶ Gelatinolytic activity was manifested as horizontal white bands on a blue background. 

Inhibition of RPTP-σ Activity

Normal trabecular meshwork and NTM^T cells were exposed to 20 μM of RPTP-σ phosphatase inhibitor (PTP Inhibitor IV) 48 hours after transfection, according to the manufacturer's recommendations. Briefly, after 48 hours of incubation in 24-well plates (37°C, 5% CO₂), the medium of the NTM and NTM^T cells was replaced with a starvation medium (1% FCS) and the cells were incubated for an additional 24 hours at 37°C, 5% CO₂. The medium was then replaced with a different starvation medium, with a final RPTP-σ phosphatase inhibitor concentration of 20 μM. 

DiFMUP

Protein phosphatase activity in cell lysates was assayed in black, flat-bottomed, 96-well plates (transparent Greiner plate). The mixture of cell lysates in a buffer was incubated for 10 minutes at 37°C, after which the fluorescent phosphatase substrate DiFMUP, at a concentration of 200 μM, was added to each well with intermittent mixing. Phosphatase activity was assessed in a fluorescent plate reader with excitation and emission filters of 358 and 455 nm, respectively. 

Statistical Analysis

Differences between the experimental and control cells were analyzed by a one-way ANOVA followed by Bonferroni corrections, by using the statistics software (InStat; GraphPad, La Jolla, CA, USA). Results with P values lower than 0.05 were considered significant. 

Results

To evaluate transfection efficacy, GFP expression after transient co-transfection of plasmid DNA with 9 × pcDNA3.1: 1 × GFP was followed by flow cytometry. A significant increase (P < 0.0001) in GFP fluorescence was observed in the NTM^T cells, compared with the controls (Table). 

Table

 

RPTP-σ Expression in NTM Cells Based on GFP Fluorescence 48 Hours Following Transfection

 

Transfection Efficiency–RPTP-σ Activity

Proteins were extracted 48 hours after transfection and the activity of RPTP-σ was analyzed by Western blotting. Following an SDS-PAGE gel electrophoresis of the extracted proteins, RPTP-σ activity was assessed by using a specific antibody that detects the endogenous cleaved extracellular domain of human RPTP-σ.²³ Compared with its activity in NTM cells (which was considered as 100% activity; 100% ± 12.50%) and in GTM cells (68.03% ± 17.69%), the activity of RPTP-σ was significantly increased in NTM^T cells (226.55% ± 45.54%, P < 0.001; Figs. 1A, 1B). 

Figure 1

 

Activity of RPTP-σ phosphatase in NTM, NTM^T, and GTM cells, as measured by Western blot analysis. Activity of RPTP-σ in NTM cells was considered as 100% and was normalized to the β-actin band. (A) Mean ± SD of three independent experiments. **P < 0.01 (one-way ANOVA). (B) Representative Western blot results.

Location of RPTP-σ in NTM^T Cells–Confocal Microscopy

RPTP-σ is a trans-membrane receptor that is normally involved in cell-cell and cell–extra-cellular matrix communication. To determine the location of RPTP-σ in the cells, we used confocal microscopy after staining the NTM^T cells with DAPI (for the nucleus) and with an RPTP-σ antibody. RPTP-σ proteins, marked with specific antibody, were spread across the entire cytosol, including on the NTM cell extensions (Fig. 2; Supplementary Fig. S1). 

Figure 2

 

Fluorescence imaging of RPTP-σ expression in NTM^T cells. (A) RPTP-σ and DAPI overlay. (B) NTM^T cells bright field picture.

NTM^T Viability Following H₂O₂ Exposure

Published data suggest that RPTP-σ is involved in redox cell regulation.²⁷ To test whether RPTP-σ influences the viability of NTM^T cells following oxidative stress, the cells were exposed to different H₂O₂ concentrations for 15 minutes and their viability was evaluated 24 hours later. No significant changes in cell viability were observed in the RPTP-σ transfected cells following exposure to 100 μM H₂O₂ (NTM cells: 120.81% ± 17.52%, NTM^T cells: 111.70% ± 9.47%; compared with the viability of cells not exposed to H₂O₂). Subsequent experiments in the present study were conducted with 15-minutes exposures to 100 μM H₂O₂. 

Effect of RPTP-σ Transfection on MMP Activity in NTM Cells

Activity of MMP in NTM cells was assessed by a zymography analysis 48 hours following transfection. Using gelatinase activity assay, in human samples only pro–MMP-9, MMP-9, pro–MMP-2, and MMP-2 can be detected. Four different bands were detected; based on their molecular weights (pro–MMP-9, ∼90 KDa; MMP-9, ∼86 KDa; pro–MMP-2, ∼72 KDa; and MMP-2, ∼66 KDa), gel shifts, and values in the literature,^28,29 they were identified (Figs. 3, 4). The activities of pro–MMP-9 (P < 0.05) and pro–MMP-2 (P < 0.01) were significantly increased in NTM^T cells compared with NTM controls. 

Figure 3

 

Activity of pro–MMP-9, pro–MMP-2, MMP-9, and MMP-2 in NTM and in NTM^T cells. A representative stained zymography gel of protein extracts from NTM and NTM^T cells.

Figure 4

 

Average activity of the pro- and active forms of MMP-2 and MMP-9 in NTM and in NTM^T cells, based on zymography analysis of (A) pro–MMP-9, (B) MMP-9, (C) pro–MMP-2, and (D) MMP-2. Cell activity of NTM was considered as 100%. The graph shows mean ± SD of three independent experiments, each performed in duplicates. **P < 0.01 (one-way ANOVA).

Serine/Threonine Phosphatase Activity in NTM^T Cells

We tested NTM^T cells transfected with tyrosine receptor phosphatases for serine/threonine phosphatase activity to verify that the phosphatase activation was not limited to the transfection. Activity was analyzed by DiFMUP at different time points following a 15-minute exposure to 100 μM H₂O₂. Serine/threonine phosphatase activity increased significantly within 24 hours in the H₂O₂-exposed NTM^T cells, compared with the NTM^T cells that were not exposed to H₂O₂. Across all time points that were examined, the activity of the serine/threonine phosphatase was significantly higher in the NTM^T cells than in the NTM cells (Fig. 5). 

Figure 5

 

Serine/threonine phosphatase activity in NTM (solid gray bars) and in NTM^T (solid black bars) cells exposed to H₂O₂, measured by using the DiFMUP method. Cells were incubated for 15 minutes in 100 μM H₂O_2, after which they were returned to the original growth medium. Protein extraction was performed at different times after the H₂O₂ exposure. Control cells were not exposed to H₂O₂ (NTM, spotted gray bar; NTM^T, spotted black bar). Data show the mean ± SD of three independent experiments performed in six repetitions. Analysis was performed with a two-way ANOVA: A = significantly different from the NTM^T control group, P < 0.001; B = significantly different from the NTM^T group at the same time after exposure to H₂O₂, P < 0.001.

Effect of RPTP-σ Inhibition on Phosphatase and MMPs Activity

To establish a link between the expression and activity of RPTP-σ and the activity of MMPs, we treated the NTM^T cells with PTP-IV, a specific RPTP-σ inhibitor. Cell proteins of NTM^T were then extracted and tested by Western blotting for RPTP-σ (Fig. 6A). As was shown previously, the NTM^T cells exhibited a significantly higher RPTP-σ activity compared with the control NTM cells and with GTM cells. Treating the NTM^T cells with PTP-IV significantly reduced RPTP-σ activity (namely, by 31%, P < 0.0001; Fig. 6B), whereas no change in RPTP-σ activity was observed in the NTM and GTM controls. The increase in RPTP-σ expression in NTM^T cells was not completely abolished by PTP-IV in the concentration used in the present study. The inhibition of RPTP-σ attenuated the activity of MMP; in NTM^T cells, the PTP-IV treatment resulted in a 48% inhibition of pro–MMP-9 (P < 0.05), in a 35% inhibition of the active MMP-9 (P < 0.001), in a 78% inhibition of pro–MMP-2 (P < 0.001), and in a nonsignificant change in active MMP-2. In the control NTM cells, a smaller reduction (namely, of 27%) was observed in the activity of pro–MMP-9 (Fig. 7). 

Figure 6

 

Activity of the RPTP- σ phosphatase in NTM^T cells in the presence of an RPTP-σ inhibitor (In). (A) A representative Western blot analysis of RPTP-σ activity in NTM, NTM^T, and GTM cells in the presence (+) and abscence (−) of the RPTP-σ inhibitor. (B) The mean ± SD of three independent experiments. **P < 0.01, ***P < 0.001 (one-way ANOVA).

Figure 7

 

Activity of pro–MMP-9, pro–MMP-2, MMP-9, and MMP-2 in RPTP-σ–transfected NTM cells in the presence of an RPTP-σ inhibitor. Shown are representative examples of zymography results of NTM, NTM^T, and GTM cells in the presence of the RPTP-σ inhibitor (A). Graphs (B1–4) show the mean ± SD of three independent experiments. **P < 0.01 (one-way ANOVA).

Discussion

The aim of the present research was to elucidate a new role for phosphatases in the control of the ocular drainage system. A trabecular meshwork cell line was successfully transfected transiently with RPTP-σ, a subtype of the tyrosine phosphatase receptor. The expression levels and activity of RPTP-σ in transfected cells, as measured by GFP fluorescence and by a direct antibody, respectively, in a Western blot analysis, was significantly higher than in control cells, thus allowing us to investigate part of the potential role of RPTP-σ in TM cells. By using confocal microscopy, we show that RPTP-σ is distributed throughout the cytosol, including in the cell extensions. Previously, RPTP-σ was found to be a trans-membrane protein²³; but, in our results, it appeared to be cytosolic. The possibility that RPTP-σ expression in TM cells is not exclusively cytosolic needs further investigation. 

Subtypes of RPTP (namely, RPTP-σ, RPTP-α, RPTP-δ) have been found on the surface and cytosol of different neuronal and fibroblast cells.^23,30,31 To the best of our knowledge, the current study is the first to show that RPTP-σ is generally expressed in NTM and in GTM cells. The activity of RPTP-σ in NTM and GTM cells was found to be similar and significantly lower than in NTM^T cells. Thus, in the current study, we enhanced an already functioning phosphatase receptor and its consequent signaling, which increases the relevance of the presented data. In studies employing NTM and GTM cells, results have suggested similar³² or a different³³ response to some stimuli of the two cell lines. While comparing the NTM cell line to TM primary cell culture, a similarity in response to some stimuli³⁴—while a poor response was exhibited in others³⁵—was found. Further studies that compare primary TM cells derived from glaucoma patients to cells from a normal TM tissue are needed to verify the activity of RPTP-σ. 

The findings of the present study highlight the importance of the signaling pathway mediated by phosphatases. We were able to present PTRP-σ–mediated changes in the activity of MMPs following oxidative stress applied to the TM cells. These findings suggest a novel pathway in TM cells, which might be involved in outflow resistance changes occurring in the ocular drainage system. In the delicate control of the drainage system, several signaling pathways that link MMP changes with TM exposure to proteins or stress have been demonstrated; the current study suggests that phosphatases, in general, and PTRP-σ, in particular, are another important pathway. 

Since the introduction of prostaglandin analogues, more than a decade ago, the activation of MMPs has been a well-known means by which to treat glaucoma patients. Trabecular meshwork cells treated with the prostaglandin analogue latanoprost responded with a significant elevation in MMP-2 mRNA and with a significant elevation in the expression of MMPs and tissue inhibitors MMPs (TIMMPs),³⁶ but not of MMP-9. Similar to our findings, a study by Pang et al.³⁷ found basal levels of pro–MMP-2 and pro–MMP-9 and others MMPs in TM cells.³⁷ The study by Fleenor et al.³⁸ successfully linked between TM exposure to cytokines—including IL-1α and phorbol ester—and changes in the expression of MMPs and TIMMPs.³⁸ In another investigation, treating human TM cell culture with TGF-β2 significantly increased the expression of pro–MMP-2 without changing the expression of MMP-9. Moreover, the significant increase in the expression of PAI-1 decreased MMPs activity.³⁹ Treating TM cells with an adenosine-1 agonist resulted in an increase in MMP-2 excretion, which was found to be mediated by Erk1/2.⁴⁰ Activation of adenosine A1 receptors also increased the activity of MMP-9 secreted by TM cells.⁴¹ The literature is rich with reports of the effects of mechanical stress on changes in MMPs and TIMMPs expression and activation in TM cells and in TM tissue culture; those changes in expression patterns suggest the involvement of NFκB, AP, MAPK, and PLA2.⁴² These studies, and many others, suggest a complex response of the TM to stress, with different levels of expression and activation for different MMPs. Among the many MMPs examined, MMP-2 was found to be involved in many cases.^36–39 Exposure to chronic oxidative stress is a well-known characteristic of POAG pathophysiology,⁹ which we mimicked by exposing NTM^T and control cells to moderate, nonlethal levels of H₂O₂.^13,43 The link between oxidative stress and MMP activity was suggested in several previous studies.^9,44,45 We examined the changes that occurred in MMP activity following overexpression of RPTP-σ under oxidative stress. The matrix metalloproteinases are secreted from cells as proenzymes that must be activated by proteolytic cleavage. We found that the activities of pro–MMP-2 and pro–MMP-9 were significantly increased in the NTM^T cells. Basal levels of these two gelatinases and of other MMPs have been demonstrated previously in human TM cells.³⁷ Data in the literature regarding the factors that affect the TM and that result in the changes in MMPs were obtained mainly in AH analysis⁴⁶ or in cell cultures by medium analysis.^47,48 To the best of our knowledge, our study is the first to measure the activity of MMPs within TM extracts. As such, the changes in MMPs reported in this study occurred before the TM cell secreted these enzymes, and additional controls that exist in the TM cell may attenuate the final MMP activity measured post secretion. We were able to directly link the oxidative stress to which the different types of NTM cells were exposed, with changes in phosphatase activity. By using PTP-IV, a specific RPTP-σ inhibitor, part of the signaling pathway mediated by RPTP-σ was inhibited. We assume that by prolonging PTP-IV incubation and increasing its dose, a further inhibition of the mediated RPTP-σ, MMPs activity could be achieved. Notably, RPTP-σ transfection affected both total tyrosine phosphatase activity and serine/threonine phosphatase activity. Elucidating the mechanism underlying this phenomenon will require additional research. 

The present study demonstrated two different effects: changes in tyrosine phosphatase levels in NTM cells are linked to gelatinase activity, and hydrogen peroxide–induced stress causes changes in NTM cell phosphatases activity. The degree of protein phosphorylation and dephosphorylation is one means by which cell signaling is controlled. Phosphatases and kinases are the enzymes responsible for the phosphorylation and dephosphorylation of specific protein sites in the cells. Disruptions of the homeostatic balance between the activities of phosphatases and kinases can lead to a variety of human diseases.^49,50 Kar et al.⁵¹ showed a direct link between oxidative stress, tyrosine phosphatase inhibition, and pro–MMP-1 activation. Phosphatase activity is thought to allow unbalanced kinase signaling activation—for example, by oxidative stress—leading to the recruitment of proteins to the promoter region and to an increased transcription of MMP. The use of Erk and JNK inhibitors decreased MMP-1 promoter activation.^50,51 Based on these data, we suggest that the exposure of NTM cells to H₂O₂ led to modifications in the phosphatase activity in those cells, which resulted in changes in the activity of MMPs. Further understanding of the role of phosphatases in the ocular drainage system will hopefully suggest new targets for intervention for POAG patients. 

Acknowledgments

Disclosure: M. Zaiden, None; E. Beit-Yannai, None