Molecular Vision 2017; 23:1029-1038
Received 30 January 2017 | Accepted 27 December 2017 | Published 29 December 2017
1Balgeunsesang Eye clinic, Seoul, South Korea; 2Department of Ophthalmology, Keimyung University School of Medicine, Dongsan Medical Center, Daegu, South Korea; 3Institute for Cancer Research, Keimyung University, Dongsan Medical Center, Daegu, South Korea; 4Department of Microbiology, Keimyung University School of Medicine, Daegu, South Korea; 5Department of Oral Biochemistry, School of Dentistry, IHBR, Kyungpook National University, Daegu, South Korea; 6Department of Ophthalmology and Visual Science, Seoul St. Mary’s Hospital, College of Medicine, The Catholic University of Korea, Seoul, South Korea
Correspondence to: Jong Hwa Jun, Department of Ophthalmology, Keimyung University School of Medicine, Dongsan Medical Center, Daegu, South Korea, Phone: 82-53-250-7708; FAX: 82-53-250-7705; email: firstname.lastname@example.org
Purpose: Nuclear factor kappa B (NF-κB) plays an important role in the epithelial–mesenchymal transition (EMT) of RPE cells. We investigated the effects of a proteasome inhibitor, bortezomib, on the EMT in RPE cells. In addition, we assessed the influence of bortezomib on regulation of the NF-κB pathway during this process.
Methods: After treatment with various concentrations of bortezomib, cell viability was analyzed with the water-soluble tetrazolium salt-8 assay, cell-cycle regulation was evaluated with flow cytometry, and cell migration was monitored with in vitro wound healing and Transwell migration assays. To induce fibroblastoid transformation, the RPE cells were treated with recombinant human transforming growth factor (TGF)-β1 (10 ng/ml), and western blot and immunocytochemical analyses were performed to evaluate altered expression of EMT markers after treatment with bortezomib. To verify the effect of bortezomib on shrinkage by myofibroblastic transformation, a contraction assay of the RPE–collagen gel lattice was performed.
Results: Treatment with bortezomib decreased RPE viability in a dose-dependent manner, and flow cytometry revealed that these effects were due to arrest of the G2/M phase cell-cycle. In the in vitro wound healing and Transwell migration assays, treatment with 20 nM bortezomib significantly impeded RPE migration. Treatment with bortezomib also significantly inhibited TGF-β1-induced transdifferentiation of the RPE cells. The effects on proliferation, migration, and the EMT were mediated by regulation of the NF-κB signaling pathway. In addition, bortezomib inhibited contraction of the RPE–collagen gel lattices.
Conclusions: Bortezomib inhibits myofibroblastic transformation of RPE cells by downregulating NF-κB expression and prevents contraction of the RPE–collagen gel matrix. Thus, bortezomib represents a candidate putative therapeutic agent for management of retinal fibrotic diseases.
Human RPE cells are essential for visual functions that support photoreceptor function and regulate the blood–retina barrier (BRB) by forming mature tight junctions composed of zonular occludens-1 (ZO-1), occludin, and claudin family proteins [1,2]. After the completion of development in the early gestational period, RPE cells become dormant, undergoing minimal proliferation throughout life . Dissociation of epithelial cells from their original locations promotes alterations in phenotypes and physiology, often accompanied by development of cellular plasticity, acquisition of migratory properties, loss of cell type-specific gene expression, and cytoskeletal rearrangements [3-5].
In ocular pathophysiology, such cellular processes are implicated in various fibrotic diseases. In proliferative vitreoretinopathy (PVR), dispersion of RPE cells through the retinal breaks in rhegmatogenous retinal detachment (RRD) provides the opportunity to disseminate RPE cells into the subretinal space or vitreous cavity and induce inflammatory processes caused by breakdown of the BRB [6,7]. In addition, the epiretinal/subretinal membrane (ERM), or tractional membrane formations in proliferative diabetic retinopathy (PDR) accompanying RRD, habitual retinal cells, RPE cells, and glial cells are major sources of the ERM [8,9]. Moreover, RPE cells may play pathological roles in idiopathic ERM, although this remains controversial .
In these diseases, contraction of the fibrotic membrane is crucial for ERM or PVR pathology, and myofibroblastic transformation by epithelial–mesenchymal transition (EMT) of RPE cells is a hallmark of the associated pathobiology [11,12]. RPE cells are stimulated by various cytokines, including transforming growth factor (TGF)-β, platelet-derived growth factor (PDGF), vascular endothelial growth factor, tumor necrosis factor α, and epidermal growth factor [6,7,13-15]. RPE cells that undergo dedifferentiation are transformed into myofibroblasts with accelerated proliferation and migration.
Nuclear factor kappa B (NF-κB) is a key transcriptional regulator of various responses and plays a pivotal role in regulation of inflammatory signaling cascades. NF-κB is activated by stress or injury, inflammatory cytokines, virus, and bacterial toxins, and downstream effectors of the NF-κB are involved in cellular proliferation, differentiation, and apoptosis . Several drugs are used to regulate NF-κB in inflammatory diseases and tumors. Furthermore, fibrogenesis in multiple tissues can be inhibited by blockade of NF-κB [17,18]. Ocular fibrotic diseases, such as posterior capsule opacification, are associated with similar EMT pathobiology. Therefore, inhibition of NF-κB signaling by RNA silencing, sulfasalazine, or antisense oligodeoxynucleotide prevents these diseases [19-21]. NF-κB plays essential roles in induction and maintenance of EMT, and loss of NF-κB activity decreases EMT-related gene expression [22-24]. Together, these observations suggest that inhibition of NF-κB represents a promising therapeutic strategy for the prevention or control of ocular fibrotic diseases arising from EMT pathobiology.
Bortezomib (Velcade®; Millennium Pharmaceuticals Inc., Cambridge, MA) is a cancer drug approved by the U.S. Food and Drug Administration (FDA) for the treatment of multiple myeloma and mantle cell lymphoma as a proteasome inhibitor [25,26]. Bortezomib was originally developed as a blocker of proteasomal degradation of nuclear factor of kappa light polypeptide gene enhancer in B-cells inhibitor, alpha (IκBα), which acts by binding to the catalytic site of the 26S proteasome with high affinity and specificity; however, the precise mechanisms have not been fully elucidated [25-27].
In this study, we focused on the therapeutic effects of bortezomib on the EMT of RPE cells induced by TGF-β1, a potent inhibitor of the NF-κB signaling pathway . We evaluated the inhibitory effects of bortezomib in an in vitro EMT model system consisting of RPE cells.
The immortalized human RPE line ARPE-19 was purchased from American Type Culture Collection (ATCC, Manassas, VA). Passage 19 cells were received and were used in experiments after five to ten more passages. ARPE-19 was maintained in 1:1 Dulbecco’s modified eagle’s medium/Ham’s F12 nutrient (DMEM/F12; Welgene, Daegu, South Korea) supplemented with 10% (v/v) fetal bovine serum (FBS; Welgene) and 1% (v/v) penicillin/streptomycin (Welgene) in a humidified incubator at 37 °C in 5% CO2. To induce EMT, RPE cells at 90% confluence were treated with human recombinant TGF-β1 (10 ng/ml) in serum-free medium. Bortezomib was dissolved in dimethyl sulfoxide (DMSO), and the final concentration of DMSO in samples did not exceed 0.0002% (v/v). RPE cells were cultured with 20 nM bortezomib or the corresponding volume of DMSO (control).
Seventeen short tandem repeat (STR) loci plus the gender-determining locus, amelogenin, were analyzed in Cosmo genetech (Seoul, Korea) using GeneMapper® software 5(Applied Biosystems, Foster City, CA). Appropriate positive and negative controls were run and confirmed for each sample submitted. The STR analyses are presented in Appendix 1.
Reagents and kits were obtained from the following suppliers: bortezomib, Selleckchem (Houston, TX); human recombinant TGF-β1, PeproTech (Rocky Hill, NJ); Vectashield Mounting Medium with 4',6-diamidino-2-phenylindole (DAPI), Vector Labs (Burlingame, CA); a Cell Counting Kit-8 (CCK-8), Dojindo Molecular Technologies (Rockville, MD); inserts for Transwell migration assays, Corning (Corning, NY); a Collagen Gel Contraction Assay Kit, Cell Biolabs (San Diego, CA); and a Propidium Iodide Flow Cytometry Kit, Abcam (Cambridge, UK). Primary antibodies were as follows: anti-RHOG, Sigma (St. Louis, MO); anti-RAC1, EMD Millipore (Billerica, MA); anti-α-smooth muscle actin (SMA) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH), Abcam; anti-NF-κB, phospho-NF-κB, IκBα, vimentin, ZO-1, N-cadherin, and β-catenin, Cell Signaling Technology (Danvers, MA); and anti-occludin, Santa Cruz Biotechnology (Dallas, TX). All secondary antibodies for the immunoblot assays were acquired from Jackson Laboratory (Bar Harbor, ME), and fluorescence-conjugated secondary antibodies for immunocytochemistry and Alexa Fluor 488 phalloidin were obtained from Thermo Fisher Scientific (Waltham, MA). All other chemicals were obtained from Sigma.
To determine the effective concentrations of bortezomib for regulation of RPE physiology, we plated ARPE-19 cells in four sets of 12-well plates and tested the effects of various concentrations of bortezomib (control [vehicle], 0.2, 2.0, and 20 nM) on cell viability at various time points (12, 24, 36, and 48 h). Every 12 h, the culture medium was aspirated, and the cells were rinsed with PBS (1X; 9 g/l NaCl, 0.21 g/l KH2PO4, 0.726 g/l Na2HPO4·7H2O, pH 7.2). CCK-8 reagent (100 µl) mixed with serum-free medium (900 µl) was added to each well. After 4-h incubation, 100 µl of the reagent/medium mixture was transferred to a 96-well plate. Colorimetric detection at a wavelength of 450 nm was performed on a microplate reader (SPECTROstar Nano, BMG LABTECH GmbH, Ortenberg, Germany).
RPE cells were cultured in the presence or absence of 20 nM bortezomib for 24 or 48 h, and flow cytometric analysis was performed at each time point. For this purpose, culture medium was aspirated and rinsed with PBS, and 0.25% trypsin-EDTA solution was added to the cell monolayer to harvest the RPE cells as a single-cell suspension. After the cell pellets were spun down, the supernatant was aspirated, and the pellets were resuspended in PBS. The RPE cells were then fixed with 70% cold ethanol on ice and rehydrated with PBS. The fixed cells were stained with the propidium iodide and RNase mixture. Measurements were performed on a benchtop flow cytometer (NovoCyte Flow Cytometer Systems, ACEA Biosciences, San Diego, CA), and the results were analyzed using the NovoExpress software (ACEA Biosciences).
After the culture medium was removed followed by three rinses with cold PBS, the cells were lysed using radioimmunoprecipitation assay (RIPA) buffer (ELPIS-Biotech Inc., Daejeon, South Korea). Equal quantities of each protein sample were loaded in lanes of 8–15% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS–PAGE) gels, and the proteins were separated with electrophoresis. Resolved proteins were transferred to nitrocellulose membranes and blocked with 5% nonfat milk in a mixture of Tris-buffered saline (TBS) and Tween 20 (TBST) for 1 h. After overnight incubation with the indicated primary antibodies, the membranes were incubated with secondary antibodies for 1 h. Specific proteins were detected with enhanced chemiluminescence on a ChemiDoc™ MP System using Image Lab™ Software (Bio-Rad, Hercules, CA). GAPDH was used as a loading control for blot band densitometry. ImageJ software (freely available from the National Institutes of Health, Bethesda, MD) was used for densitometric analyses.
ARPE-19 cells were seeded in four-well glass cell-culture slides (SPL Life Sciences, Pocheon, Korea). The RPE cells were cultured
in the presence or absence of 20 nM bortezomib at 24 h after plating and cultured to 50–60% confluence. After the medium was
removed followed by three rinses with PBS, the cells were fixed with 4% paraformaldehyde for 30 min. For observations of filamentous
actin, the cells were stained with Alexa Fluor 488–conjugated phalloidin/Vectashield mounting solution
ARPE-19 cells were placed on 12-well plates and cultured to 90% confluence. Crosshair linear wounds were made with a standard 200-µl pipette tip. After three rinses with PBS, medium containing 1% FBS was added, and the cells were cultured in the presence or absence of 20 nM bortezomib. The remaining wound area was observed under a phase-contrast microscope every 12 h. The wound area at each time point was measured using ImageJ.
ARPE-19 cells were trypsinized and resuspended in serum-free medium in the presence or absence of 20 nM bortezomib. Medium containing 10% FBS was added to each well (the bottom well), and the cell suspension was poured into polycarbonate membrane inserts with 8-µm pores (the top well). After 24-h incubation, cells that had migrated to the bottom side of the polycarbonate membrane were fixed with 4% paraformaldehyde and stained with hematoxylin. Migratory cells were counted under a phase-contrast microscope at three different locations on each insert.
ARPE-19 cells were cultured in 100-mm dishes and harvested using 0.25% trypsin-EDTA solution. Two parts of cell suspension were mixed with eight parts of cold collagen gel solution, and then 100-μl aliquots of the cell/collagen gel mixture was dispensed into 48-well culture plates. After polymerization of the cell/gel mixture for 1 h at 37 °C, 100 μl of medium containing 20 ng/ml TGF-β1, with or without 20 nM bortezomib, was added to the top of the collagen lattice, and the samples were incubated for up to 72 h. Each gel was observed under a phase-contrast microscope, and the collagen gel area in each well was measured using images taken after 24 h of incubation. To evaluate cell spreading and cell-to-cell adhesion, each cell/collagen gel mixture was observed on an inverted microscope (Ti-U, Nikon Instruments Inc.). Images were acquired using the NIS-Elements imaging software (Nikon Instruments Inc.). The collagen gel area was measured using ImageJ.
Representative data were collected from at least triplicate experiments. All experiments were evaluated statistically with an independent t test or a one-way ANOVA (ANOVA) test, and the Tukey HSD post-hoc test was applied to determine significant differences between results. A p value of less than 0.05 was regarded as statistically significant.
Treatment of RPE cells with 20 nM bortezomib for 48 h resulted in significantly lower cell density relative to the control samples (Figure 1A). Therefore, to determine the effective working concentration of bortezomib, we treated the RPE cells with various concentrations of bortezomib (0, 0.2, 2.0, and 20 nM) and repeatedly measured cell viability every 12 h for 48 h. Bortezomib decreased cell viability in a dose-dependent manner; 20 nM bortezomib significantly decreased cell viability or density after 24 h, and this effect was maintained at 48 h (Figure 1B).
After we treated the RPE cells with 20 nM bortezomib, we analyzed the cell cycle with flow cytometry. Treatment with bortezomib for 24 or 48 h increased the population of G2/M-arrested cells (Figure 1C,D). However, the proportion of sub-G1 cells was not significantly increased by treatment with bortezomib in comparison with vehicle-treated cells (data not shown). These results support that the decrease in cell viability, especially the abrupt change at the 20 nM concentration, was due to arrest in the G2/M phase.
To assess the effects of bortezomib on RPE migration, we performed in vitro wound-healing and Transwell migration assays. After 24 h in the presence of bortezomib, the remaining wound area was significantly larger than in the control wells. After treatment with bortezomib for 36 or 48 h, the wound area remained larger than in the control samples (Figure 2A,B). The results of the Transwell migration assay were in accordance with those of the wound-healing assay: The number of migrated cells was significantly lower in the bortezomib-treated samples (Figure 2C,D).
Based on the results of the migration and in vitro wound-healing assays, we hypothesized that treatment of RPE cells with bortezomib would affect factors related to cell motility. To test this idea, we performed phalloidin staining in the presence or absence of bortezomib. As we expected, bortezomib decreased the number of migratory organelles, including lamellipodia and stress fibers within the cytoplasm (Figure 2E). Western blot analysis showed that bortezomib downregulated expression of RHOG/RAC in RPE cells at early time points (Figure 2F). In addition, immunocytochemical staining revealed reduced expression of RHOG and RAC1 in RPE cells after treatment with bortezomib (Figure 2G).
Various concentrations of bortezomib were applied to RPE cells in which fibroblastoid transdifferentiation had been induced by treatment with TGF-β1. At 20 nM bortezomib, epithelial markers such as ZO-1 and occludin were upregulated, whereas N-cadherin, vimentin, α-SMA, and β-catenin were downregulated (Figure 3A); band densitometry in the triplicate samples revealed these differences were statistically significant (Figure 3B). Similar patterns of expression were displayed in immunocytochemical analysis (Figure 3C). Thus, bortezomib effectively reversed the EMT in RPE cells.
Matrix contraction plays a crucial role in the pathology of myofibroblastic transformation, especially in PVR or the ERM. Therefore, we investigated the effect of bortezomib on the contraction of the RPE–collagen gel lattices induced by TGF-β1. In the presence of 20 nM bortezomib, contraction of the collagen gel was significantly reduced, whereas the control-treated gels shrunk to approximately half of their initial area (Figure 4A). After 24 h, bortezomib effectively reduced shrinkage of the collagen gel lattices, and this effect remained statistically significant for 72 h (Figure 4B). Microscopic observations revealed that treatment with bortezomib reduced RPE spreading and cell-to-cell adhesion, which is the basis of the fibrous contraction band (Figure 4C). Filamentous actin staining with phalloidin revealed that the formation of stress fibers, which contributes to contraction of the RPE–collagen gel lattice, was diminished by treatment with bortezomib (Figure 4D).
To determine whether treatment with the 20 nM bortezomib concentration could regulate the NF-κB signaling pathway in RPE cells stimulated or unstimulated with 10 ng/ml TGF-β1, we performed western blot analyses in the presence or absence of bortezomib. As shown in Figure 5A,B, 20 nM bortezomib inhibited expression of NF-κB (p65) and decreased the level of NF-κB phosphorylation. Conversely, bortezomib downregulated IκBα at early time points (Figure 5A,B). These regulatory patterns were observed regardless of whether the RPE cells were stimulated or unstimulated with TGF-β1. Accordingly, immunocytochemical staining 1 h after treatment with vehicle or bortezomib revealed decreased cytoplasmic expression of NF-κB but increased expression of IκBα in RPE cells, with or without application of TGF-β1 (Figure 5C,D).
Bortezomib, a modified dipeptidyl boronic acid analog, is the first proteasome inhibitor drug approved for relapsed refractory multiple myeloma. Since bortezomib was approved in 2003, it has been used as a first-line drug for this indication, either alone or in combination with other drugs . Current clinical usage of bortezomib has been extended to treatment of thrombotic thrombocytopenic purpura and lymphoproliferative neoplasms, and recent reports showed that bortezomib used in concert with other agents promotes long-term survival of solid organ transplants [30-33]. In addition, several studies reported benefits of treatment with bortezomib in ophthalmic pathologies, such as experimental uveitis, ischemia-reperfusion injury, postoperative fibrosis in glaucoma filtration surgery, and retinoblastoma [34-38]. In this study, we investigated the inhibitory role of bortezomib in TGF-β1-induced EMT in RPE cells, which mimics the PVR of rhegmatogenous retinal detachment or tractional ERM in diabetic retinopathy.
Initially, we tried to measure the minimal inhibitory concentration of bortezomib in the regulation of RPE physiologies. Cell viability analysis with the water-soluble tetrazolium salt-8 (WST-8) assay and microscopic observations showed that the 20 nM concentration of bortezomib clearly decreased cell density. However, because bortezomib can act as a cell-cycle regulator or an inducer of apoptosis [37,39], we cannot be sure whether the effects of the drug are mediated by arrest of the cell cycle or apoptosis. Therefore, we monitored the effect of the drug on cell-cycle progression, using flow cytometry. The results revealed that 20 nM bortezomib did not induce significant arrest in the sub-G1 phase in RPE cells but instead induced only arrest of the G2/M phase (data not shown).
In addition, we sought to determine with an in vitro cell scratch assay whether bortezomib could inhibit wound-healing ability. Although we limited the serum content of the culture medium, the results of the in vitro wound-healing assays reflect the proliferation and migration of cells. Therefore, to focus on migration, we also performed the Transwell migration assay. We observed that bortezomib effectively inhibited wound healing in RPE monolayers and that pure migration of RPE cells was also limited. For a detailed evaluation of RPE migration, we investigated alterations of the cytoskeleton by visualizing filamentous actin. Interestingly, bortezomib distinctively altered cytoplasmic actin and diminished lamellipodium formation at the cell border, and consequently, the RPE cells were elongated with the loss of polarity. Because such cytoskeletal regulation is governed by the RHOG/RAC1 signaling pathway, we assessed the effect of treatment with bortezomib on these factors and found that bortezomib downregulated the expression of RHOG and RAC1. It remains unclear whether the proteasome/NF-κB and RHOG/RAC1 pathways engage in crosstalk, and further evaluations are needed to confirm this relationship.
When activated, the NF-κB signaling pathway stimulates the production of various cytokines, including TGF-β [40,41]. TGF-β1 itself can also induce NF-κB signaling transduction by downregulating expression of IκBα . Downregulation of NF-κB expression by proteasome inhibitors blocks transdifferentiation of human lens epithelial cells (LECs) [20,43]. Therefore, we predicted that bortezomib would also inhibit transdifferentiation of RPE cells via downregulation of NF-κB signaling transduction. Treatment with bortezomib downregulated NF-κB/phospho-NF-κB and upregulated IκBα expression in the presence or absence of TGF-β1. Furthermore, this agent also downregulated mesenchymal factors, such as N-cadherin, vimentin, α-SMA, and β-catenin, but upregulated epithelial markers, such as ZO-1 and occludin. Together, these results showed that 20 nM bortezomib reversed the EMT process in a statistically significant manner.
Alterations in myofibroblasts promote pathologic contractile force during the EMT process. Therefore, blockade of matrix contraction is a crucial pharmacologic feature in the evaluation of drug effectiveness in the EMT process. We confirmed that bortezomib strikingly inhibited this pathologic process, preventing progression of matrix contraction even after 24 h. Microscopic observations supported this observation. RPE spreading and cell-to-cell adhesion were also significantly perturbed by treatment with bortezomib. Moreover, the migration assays indicated that NF-κB activity is suppressed by proteasome inhibition and that RHOG/RAC1 signal transduction plays an important role in this pathway.
In conclusion, bortezomib effectively inhibits proliferation, migration, and EMT induced by TGF-β1 in RPE cells. In particular, results showed that bortezomib has potential regulatory effects that inhibit tractional membrane formation and its pathologic contraction, which are the main pathologic processes underlying PVR or ERM. These results support the idea that bortezomib could be repositioned for treatment of intractable retinal diseases.
Appendix 1. STR analysis.
This work was supported by a National Research Foundation of Korea (NRF) grant funded by the Korean government (Ministry of Science, ICT & Future Planning; NRF-2015R1C1A1A02037062 and 2014R1A5A2010008).