Lyophilized Aβ_1–42 peptide (Sigma-Aldrich) was dissolved to a concentration of 1.5 M in hexafluoroisopropanol (HFIP) on ice and aliquoted at −20°C. Aβ monomers were spin-vacuumed just prior to the experiment, diluted to 250 μM in HFIP solution, and maintained at room temperature (RT) for 3 days to synthesize Aβ oligomers. To note, In the present study, references to Aβ or OAβ are to the oligomeric form of Aβ_1–42, unless otherwise stated.

Several lines of evidence have demonstrated that soluble oligomers of Aβ may be better correlated with the severity of the disease than are monomers or insoluble amyloid fibrils (3,27). Therefore, a dot blot assay was performed to determine the oligomeric form of Aβ_1–42. Five microliters of oligomeric Aβ_1–42 solution and monomers of Aβ_1–42 were spotted onto a nitrocellulose membrane. The membrane was then blocked with 5% dry milk in TTBS (50 mM Tris, 0.05% Tween-20) for 1 h at RT, and washed three times before being incubated for 1 h at RT with the A11 (27), an anti-oligomer antibody (1:1,000; Invitrogen, Carlsbad, CA, USA).The membrane was then washed and incubated for 1 h at RT with a HRP-conjugated secondary antibody (1:1,000 goat anti-rabbit IgG; Santa Cruz Biotechnology, Inc. Santa Cruz, CA, USA). The blots were washed three times in TTBS and incubated with chemiluminescent reagent, and finally exposed to ImageQuant LAS 4,000.

Informed consent for tissue donation was obtained from the relatives of the donors, and the protocol of the study was approved by the local ethics committee and adhered to the tenets of the Declaration of Helsinki for experiments involving human tissue.

Five human donor eyes were obtained from the eye bank of the Eye and ENT Hospital of Fudan University, Shanghai, China. The donors ranged in age from 30 to 40 years. None of the donors had a history of eye disease. In brief, whole eyes were cleansed in 0.9% NaCl solution, immersed in 5% polyvinylpyrrolidone iodine and rinsed again in NaCl solution. Then the anterior segment from each eye was removed. The neural retina was peeled away from the RPE-choroid-sclera. The eyecup was rinsed with Ca²⁺ and Mg²⁺-free Hank’s balanced salt solution and treated with 0.25% trypsin for 1 h at 37°C. The trypsin was aspirated and replaced with DMEM/F12 (HyClone) supplemented with 20% fetal calf serum (FCS).

To measure cytotoxicity, haRPE cells seeded in a 96-well plate were treated with 0.3 μM of OAβ (Sigma-Aldrich, China), 20 μg/ml of GM6001 (Calbiochem, Germany), 100 μM of APMA (Sigma-Aldrich) for 24 h. Cell viability was measured by the addition of 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenol)-2-(4-sulphophenyl)-2H-tetrazolium (MTS; Promega, USA) for 3 h at 37°C. The optical density was measured spectrophotometrically at 490 nm on a microplate reader.

The protein concentrations of IL-8 and IL-6 were determined using ELISA kits (R&D Systems, Minneapolis, MN, USA) and were calculated based on standard curves. The optical density was measured at 490 nm on a microplate reader.

Human MMP-9-specific siRNAs were purchased from Santa Cruz Biotechnology, Inc. For RNA interference experiments, cells were plated at a density of 4×10⁵ cells/well in a 6-well plate. After 24 h, cells were transfected with siRNA using Lipofectamine 2000 (Invitrogen), according to the manufacturer’s instructions.

To examine whether Aβ induces MMP secretion, gelatin zymography was carried out as previous described (19). Briefly, the supernatant was collected after treatment and subjected to SDS-PAGE in 10% polyacrylamide gels with 1 mg/ml gelatin. After electrophoresis, gels were incubated in 2.5% Triton X-100 (1 h, 37°C) followed by overnight incubation in 50 mM Tris-HCl (pH 7.8), 5 mM CaCl₂, 0.02% NaN₃, 0.02% Brij gels, and were stained with 2.5% Coomassie Blue R-250 (Bio-Rad) for 45 min followed by destaining in deionized water with 10% acetic acid and 20% methanol. Gels were scanned and the density analyses of the bands were performed using Photoshop CS4.0.

The barrier function of RPE cells is considered highly dependent on the integrity of TJ proteins and F-actin. To investigate the effects of OAβ, RPE cell morphology, structures of F-actin, and location of TJ proteins were examined by light microscopy and confocal microscopy, respectively. The haRPE cells were treated with Aβ (0.3 μM) for 72 h. We hypothesized that Aβ-induced MMP-9 secretion causes disruption of epithelial barrier integrity. To test this hypothesis, cells were treated with APMA (100 μM) for 6 h to induce MMP activation. For inhibitory studies, MMP-9 activity was inhibited by MMP-9 siRNA or by the MMP inhibitor GM6001. Then cells were fixed in 4% paraformaldehyde (PFA) for 30 min and blocked with 1% BSA in TBS for 1 h, then incubated with rabbit anti-occludin antibody or mouse anti-ZO-1 antibody (Abcam, Hong Kong) for 1 h. After washing, they were incubated with AlexaFluor 488-conjugated secondary antibody (1:500; Invitrogen) and DAPI (1:1,000) for 1 h. Changes in F-actin structures were detected by FITC-labeled phalloidin (1:200; Beyotime, China). Then slides were viewed using a Leica SP5 scanning confocal microscope.

RPE cell cultures were grown on a microporous filter to form monolayers. TER was measured using Millicell-ERS (Millicell; Millipore, Bedford, MA, USA) and calculated by subtracting the value of a blank filter without cells from the experimental value. Final resistance-area products (Ω × cm²) were obtained by multiplying the TER by the effective growth area. Fifteen days after TER stabilization, the monolayers were incubated with or without one of the stimuli: Aβ (0.3 μM), APMA (100 μM), GM6001 (20 μg/ml). To demonstrate the key role of MMP-9 in regulating barrier function, MMP-9 silenced haRPE cells were exposed to Aβ (0.3 μM). Measurements were repeated at least four times for each filter, and each experiment was repeated at least four times.

The permeability assays were performed by measuring the passive permeation of FITC-dextran (4 kDa; Sigma-Aldrich) across confluent cells grown on filters. Fifteen days later, the monolayers were treated as previously described. FITC-dextran (500 mg/ml) was added to the upper chamber on day 21. Samples (100 μl) were taken from the upper and lower chamber 24 h after addition of FITC-dextran. The concentration of FITC-dextran in these samples was quantified by a microplate reader (ex1800; Biotek, Winooski, VT, USA). Each experiment was repeated four times.

Data were analyzed with the software SPSS 11.5. Results were expressed as means ± SEM. Values were processed for statistical analysis (unpaired t-test or by ANOVA), and differences were considered statistically significant at P<0.01 and P<0.05.

Samples were examined after incubation for 12 or 72 h. Dot blot assay with the anti-oligomeric specific antibody (A11) further confirmed the presence of oligomeric structures as it reacted positively with our OAβ sample (Fig. 1A). Furthermore, the A11 antibody did not react against a monomer (Fig. 1B) confirming the specificity of the antibody to the oligomeric form.

Structural integrity is the basis of good function. To confirm that the isolated haRPE cells displayed classical morphology (uniform hexagonal arrays of cells), confluence and uniform pigmentation as in native tissue, the morphology of the cells was evaluated using fluorescence microscopy (Fig. 1C). Images captured by fluorescence microscopy were analyzed using image editing software (Photoshop CS4) in the ‘a’ channel of LAB color mode. Apparent regular hexagonal morphology of the cells was observed (Fig. 1D). This result confirmed that the isolated haRPE cells exhibited heavy pigmentation (Fig. 1C) and hexagonal epithelial morphology (Fig. 1C and D) similar to these features in native tissue (17).

Effects of GM6001 (20 μg/ml) and APMA (100 μM) on cell viability were demonstrated at 24 h. None of the measurements showed significant cytotoxicity for the treatments at the concentrations used in the present study (data not shown).

Since the inflammatory response is speculated to be one of the primary causative events that contributes to OAβ-induced RPE degeneration, haRPE cells were treated with 0.3 μM of OAβ for 24 h, and the conditioned medium was collected. Expression of IL-6 and IL-8 was analyzed using ELISA (Fig. 2A and B), and the secreted levels of MMP-2 and MMP-9 were detected by gelatin zymography (Fig. 2C). The protein levels of IL-6 and IL-8 in the culture media of unstimulated RPE cells were 123.6±62.5 and 190.9±68.2 pg/ml, respectively. In OAβ-stimulated cells, the protein level of IL-6 increased to 1086.2±153.3 pg/ml (Fig. 2A), and the concentration of IL-8 increased to 1493.2±182.3 pg/ml (Fig. 2B). MMP release from the control cells was characterized by low levels of pro-MMP-9, absent levels of active-MMP-9, considerable output of pro-MMP-2, and absence of active-MMP-2. Exposure to OAβ significantly increased secreted levels of pro-MMP-9 (3.8±0.8-fold, P<0.05) and active-MMP-9 (5.1±1.2-fold, P<0.05), while treatment with OAβ did not induce any change in the secreted levels of pro-MMP-2. Active foms of MMP-2 were not observed in the OAβ-stimulated cells. These results suggest that OAβ induces haRPE cells to secrete higher concentrations of IL-6 and IL-8, which are critical chemoattractants responsible for immune-cell recruitment. In addition, the results revealed that OAβ induced the release of pro-MMP-9 and active-MMP-9 from haRPE cells.

Aβ has been shown to affect the TJs of ARPE-19 cells (16). Uncontrolled increase in MMP-9 activity has been causally linked to epithelial barrier disruption as the result of abnormal degradation of TJ proteins (21). Increased MMP-9 activity on the ocular surface disrupted corneal epithelial barrier function due to proteolytic cleavage of occludin (26). We hypothesized that Aβ-induced RPE disruption is mediated by activation of MMP-9. Therefore, MMP agonist APMA (100 μM) and MMP inhibitor GM6001 (20 μg/ml) were used to stimulate haRPE cells for 6 h. The MMP-9 gene was silenced by transfection of haRPE cells with 40 nM of MMP-9 siRNA for 24 h. Release of MMP-9 and MMP-2 from haRPE cells was then detected by gelatin zymography (Fig. 3A), and the secreted levels of MMP-2 and MMP-9 were quantified by densitometry (Fig. 3B and C). Secretion of MMPs from control cells was characterized by low levels of pro-MMP-9, absent levels of active-MMP-9, considerable output of pro-MMP-2 and absence of active-MMP-2. Treatment with APMA significantly increased secretion of pro-MMP-2 and pro-MMP-9, as well as the activated form of MMP-2 and MMP-9. Treatment with GM6001 or silencing of MMP-9 reduced the increase in pro-MMP-9 and active-MMP-9 resulting from Aβ treatment.

Recent studies involving several CNS diseases suggest that MMP-9 is involved in the permeability of the blood-brain barrier by disrupting junction complexes (15,20). Giebel et al (28) found that RPE cells treated with purified MMP-9 displayed alterations in tight junction function as shown by decreased TER. Therefore, we hypothesized that OAβ-induced RPE barrier disruption is mediated by activation of MMP-9. To test this hypothesis, haRPE cells were treated with OAβ (0.3 μM) or with APMA (100 μM) to induce MMP activation, and the barrier structure was evaluated by immunostaining of TJ proteins. The barrier function was evaluated by measuring TER and passive permeation of FITC-dextran. In the haRPE monolayers cultured with DMEM/F12 as control (Fig. 4A), no morphological change was observed; actin filaments were regular with no breaks in the staining pattern, and the distribution of occludin and ZO-1 was continuous and regular around the cells. Exposure to Aβ (Fig. 4B) resulted in an irregular morphology and a disturbed distribution of F-actin, ZO-1 and occludin; the staining of F-actin was interrupted in the intercellular space. The staining of occludin showed a diffuse cytoplasmic distribution, and the abnormal distribution of ZO-1 was typically manifested as fragmental staining. The alterations caused by Aβ (Fig. 4B) suggest that Aβ may be associated with RPE dysfunction in AMD. Notably, the deleterious effects of Aβ were reproduced by stimulation with APMA (Fig. 4C) which is known as an MMP agonist. Further study was performed to examine whether this abnormality was associated with a compromised barrier function following exposure to Aβ or APMA. The TER (Fig. 5A) was recorded to determine the stability of TJs, and the transepithelial diffusion rate of FITC-dextran (Fig. 5B) was measured to evaluate the permeability of the monolayers. The result revealed that the TER increased rapidly during the initial 12 days of standard culture and reached a plateau within the following 3 days (Fig. 5A). A mean level of 138±8.5 Ω × cm² TER was recorded on day 15, and then the monolayer was stimulated as previously described and the TER was measured every day. We observed that exposure of haRPE cells to Aβ or APMA gradually decreased the TER (Fig. 5A) and increased the diffusion rate of FITC-dextran (Fig. 5B). These results suggest that activation of MMP-9 by Aβ or APMA caused disruption of TJ proteins and transepithelial permeability dysfunction.

To confirm whether MMP-9 is responsible for disruption of epithelial barrier integrity, haRPE cells were transfected with 40 nM MMP-9 siRNA or were pretreated with GM6001 (20 μg/ml) to inhibit MMP activation prior to Aβ stimulation. Pretreatment with GM6001 inhibited the deleterious effects of Aβ on TJ proteins in the RPE cells (Fig. 4D), and partially attenuated the decrease in TER (Fig. 5A) and the increase in permeability resulting from Aβ treatment (Fig. 5B). These results suggest that inhibition of MMP activation may reverse the deleterious effects of Aβ. Exposure of MMP-9-silenced haRPE cells to Aβ did not induce abnormal staining of TJ proteins (Fig. 4E). Furthermore, transfection with MMP-9 siRNA in haRPE cells attenuated Aβ-induced transepithelial permeability dysfunction (Fig. 5). We demonstrated for the first time that inhibition of MMP-9 activity using RNA interference strategy attenuated Aβ-induced disruption of TJ proteins and permeability dysfunction.