Research ArticleArticle

The Nuclear Factor κB Family Member RelB Facilitates Apoptosis of Renal Epithelial Cells Caused by Cisplatin/Tumor Necrosis Factor α Synergy by Suppressing an Epithelial to Mesenchymal Transition–Like Phenotypic Switch

Giulia Benedetti, Michiel Fokkelman, Kuan Yan, Lisa Fredriksson, Bram Herpers, John Meerman, Bob van de Water and Marjo de Graauw
Molecular Pharmacology July 2013, 84 (1) 128-138; DOI: https://doi.org/10.1124/mol.112.084053

Abstract

Cis-diamminedichloroplatinum(II) (cisplatin)–induced renal proximal tubular apoptosis is known to be preceded by actin cytoskeleton reorganization, in conjunction with disruption of cell-matrix and cell-cell adhesion. In the present study, we show that the proinflammatory cytokine tumor necrosis factor α (TNF-α) aggravated these cisplatin-induced F-actin and cell adhesion changes, which was associated with enhanced cisplatin-induced apoptosis of immortalized proximal tubular epithelial cells. TNF-α–induced RelB expression and lentiviral small hairpin RNA (shRNA)-mediated knockdown of RelB, but not other nuclear factor κB members, abrogated the synergistic apoptosis observed with cisplatin/TNF-α treatment to the level of cisplatin-induced apoptosis. This protective effect was associated with increased stress fiber formation, cell-matrix, and cell-cell adhesion in the shRNARelB (shRelB) cells during cisplatin/TNF-α treatment, mimicking an epithelial-to-mesenchymal phenotypic switch. Indeed, gene array analysis revealed that knockdown of RelB was associated with upregulation of several actin regulatory genes, including Snai2 and the Rho GTPase proteins Rhophilin and Rho guanine nucleotide exchange factor 3 (ARHGEF3). Pharmacological inhibition of Rho kinase signaling re-established the synergistic apoptosis induced by combined cisplatin/TNF-α treatment of shRelB cells. In conclusion, our study shows for the first time that RelB is required for the cisplatin/TNF-α–induced cytoskeletal reorganization and apoptosis in renal cells by controlling a Rho kinase–dependent signaling network.

Introduction

Acute renal failure refers to a sudden deterioration of kidney function and is caused by either ischemia/reperfusion (I/R) or exposure to nephrotoxic xenobiotics such as cis-diamminedichloroplatinum(II) (cisplatin) (Thadhani et al., 1996). The proximal tubular epithelial cells (PTECs) are the primary target. After ischemic or nephrotoxic injury, morphologic alterations occur in these PTECs. Disruption of the actin cytoskeleton of the PTECs is an early event, leading to loss of adhesion of cells from the extracellular matrix (ECM) as well as neighboring cells, ultimately resulting in loss of cell function, proapoptotic signaling, and cell death (Gailit et al., 1993; van de Water et al., 1994; Kruidering et al., 1998; Zuk et al., 1998; Cordes, 2006; Pabla and Dong, 2008; Qin et al., 2011). Given the central role of the actin cytoskeletal network in cell adhesion maintenance and PTEC injury, it is important to better understand the molecular signaling networks that regulate the F-actin cytoskeleton organization in PTECs and, thereby, cell adhesion and cell survival during renal cell injury stress responses.

The actin cytoskeleton reorganization that takes place prior to and during PTEC injury is directly associated with changes in cell-matrix and cell-cell adhesion. We have previously shown that exposure to nephrotoxic compounds, including cisplatin, affects the actin cytoskeleton network accompanied by disruption of focal adhesions followed by subsequent cell detachment and cell death in renal epithelial cells (van de Water et al., 1994, 1999, 2000, 2001; Kruidering et al., 1998). In addition, cisplatin treatment induces a loss of the tight junction and adherens junction proteins β-catenin and zona-occludens 1 at the cell membrane in mouse proximal tubular cells, and preservation of the cell-cell junctions protects against cisplatin-induced apoptosis (Imamdi et al., 2004; Qin et al., 2012). Understanding which PTEC signaling programs can control the actin signaling network may lead to new therapeutic avenues to protect against cell detachment and cell death during renal injury. Here we focus on the regulation of actin organization during cisplatin-induced nephrotoxicity.

The proinflammatory cytokine tumor necrosis factor α (TNF-α) is involved in cisplatin-induced nephrotoxicity in vivo (Ramesh and Reeves, 2002, 2003). Recently, we showed that cisplatin-induced cell death of PTECs is aggravated by pre-exposure to TNF-α (Benedetti et al., 2012). This enhancement was associated with inhibition of TNF-α–induced nuclear factor κB (NF-κB) activation by cisplatin. Interestingly, TNF-α–mediated apoptosis has been associated with activation of the actin regulatory protein family of Rho GTPases in several cell lines (Mathew et al., 2009). This Rho GTPase family, including, among others, Cdc42, Rac, RhoA, and regulating the Rho kinase [Rho-associated protein kinase (ROCK)] (Leung et al., 1995) and myosin light chain kinase (Amano et al., 1996), is the most important family of proteins responsible for the actin cytoskeletal network changes that occur during PTEC injury (Kroshian et al., 1994; Raman and Atkinson, 1999). TNF-α has specifically been shown to induce the activation of the GTPases Cdc42, Rac, and Rho (Puls et al., 1999; Mathew et al., 2009).

Here we investigated the involvement of TNF-α and downstream NF-κB signaling in the regulation of cisplatin toxicity through the control of actin cytoskeletal changes that occur during cisplatin cytotoxicity. We show that TNF-α aggravates cisplatin-induced disruption of actin stress fibers in association with reduction of focal adhesions and loss of cell-cell contacts, ultimately resulting in enhanced apoptosis. Intriguingly, knockdown of the NF-κB subunit RelB, but not other NF-κB family members, protected the cells against this synergistic apoptosis. This was associated with inhibition of the morphologic changes induced by TNF-α. Gene array analysis revealed that knockdown of RelB was associated with an epithelial mesenchymal-like transition (EMT) via upregulation of Snai2. Moreover, during cisplatin/TNF-α treatment, RhoA-pathway–related gene expression, including Rhophilin and Rho guanine nucleotide exchange factor 3 (ARHGEF3), was increased. Inhibition of RhoA/ROCK signaling re-established the synergistic apoptosis with the cisplatin/TNF-α treatment in shRNARelB (shRelB) cells, indicating that TNF-α aggravated cisplatin-induced PTEC apoptosis via RelB-mediated RhoA/ROCK inactivation.

Materials and Methods

Reagents and Antibodies.

Mouse recombinant TNF-α was acquired from R&D Systems (Abingdon, UK). The selective ROCK inhibitor Y27632 [(R)-(+)-trans-N-(4-pyridyl)-4-(1-aminoethyl)-cyclohexanecarboxamide] was from Bio-Connect (Huissen, The Netherlands). The myosin light chain kinase inhibitor ML-7 was from Enzo Life Sciences (Antwerpen, Belgium). Cisplatin was from Sigma-Aldrich (Zwijndrecht, The Netherlands). AnnexinV-Alexa488 was made as previously described (Puigvert et al., 2010). The antibodies against β-actin, RelA, NF-κB1, and RelB were from Santa Cruz (Tebu-Bio, Heerhugowaard, The Netherlands), and the phospho-specific c-Jun N-terminal kinase antibody (Thr183/Tyr185) was from New England Biolabs (Leusden, The Netherlands). The antibody against NF-κB2 was from Cell Signaling (Bioké, Leiden, The Netherlands). The antibody against β-catenin was from BD Biosciences (San Jose, CA), the antibody against P-paxillin-Y118 was from Biosource (Bleiswijk, The Netherlands), and rhodamine phalloidin was from Molecular Probes (Breda, The Netherlands).

Cell Culture.

Immortalized proximal tubular epithelial cells (IM-PTECs), described previously (Stokman et al., 2011), were cultured at 33°C in human-kidney 2 medium (Dulbecco’s modified Eagle’s Medium/F12 medium; Invitrogen, Breda, The Netherlands) with 10% fetal bovine serum (HyClone, Etten-Leur, The Netherlands), 5 µg/ml insulin and transferrin, 5 ng/ml sodium selenite (Roche, Basel, Switzerland), 20 ng/ml triiodothyrionine (Sigma-Aldrich), 50 ng/ml hydrocortisone (Sigma-Aldrich), and 5 ng/ml prostaglandin E1 (Sigma-Aldrich) with L-glutamine and antibiotics (both from Invitrogen) and mouse interferon-γ (1 ng/ml; R&D Systems) in 5% CO2 and 95% air between passages 3 and 20. Prior to each experiment, the cells were differentiated into proximal tubular cells by culturing them for 4 days under restrictive conditions (at 37°C in the absence of mouse interferon-γ). The cells were then plated in the appropriate assay plates and cultured for an additional 2 days. In total, IM-PTECs were cultured under restrictive conditions for 6 days, allowing the disappearance of SV40 activity and completion of differentiation (Stokman et al., 2011).

Exposures.

IM-PTECs were exposed to 20 µM cisplatin or vehicle in the presence or absence of TNF-α (8 ng/ml). Exposure to TNF-α occurred 30 minutes prior to cisplatin, as previously described (Benedetti et al., 2012). Cells were preincubated with Y27632 and ML-7 inhibitors 30 minutes prior to TNF-α.

Lentiviral Knockdowns.

Stable IM-PTECs with knockdown of each NF-κB member were generated using lentiviral shRNA vectors (Sigma-Aldrich, in collaboration with Rob Hoeben, Leiden University Medical Centre, The Netherlands) and selection with puromycin (1 µg/ml). The plasmid encoding nontarget control SHC002 was used as a control, and the plasmids encoding mouse RelA number TRCN55343 (CCGGGCGAATCCAGACCAACAATAACTCGAGT TATTGTTGGTC TGGATTCGCTTTTTG) and TRC55346 (CCGGCTGTCCTCTCACATCCGATTTCTCGAGAAATCGGATGTGAGAGGACAGTTTTTG), mouse RelB number TRC095314 (CCGGCGGTTCTCTTTGAGCCCATTTCTCGAGAAA TGGGCTCAAAGAGAACCGTTTTTG) and TRC095318 (CCGGCGTGACTGTCAATGTGTTCTTCTCGAGAAGAACACATTGACAGTCACGTTTTTG), mouse c-Rel number TRC042549 (CCGGCCATTGTTTCTAACCCAATTTCTCGAGAAATTGGGTTAGAAACAATGGTTTTTG) and TRC042550 (CCGGGCGACTTGAGTGCATCTAATTCTCGAGAATTAGATGCACTCAAGTCGCTTTTTG), mouse NF-κB1 number TRC009514 (CCGGCACTGCTTTGACTCACTCAATCTCGAGATTGAGTGAGTCAAAGCAGTGTTTTT), mouse NF-κB2 TRC012343 (CCGGCCTGTCTAATCGAAATCTTATCTCGAGATAAGATTTCGATTAGACAGGTTTTT), and TRC012346 (CCGGCGTCATTTATCACGCTCAGTACTCGAGTACTGAGCGTGATAAATGACGTTTTT) were used.

Live Apoptosis.

Real-time induction of apoptosis was quantified using a live cell apoptosis assay that has been previously described (Puigvert et al., 2010). Briefly, binding of annexin V-Alexa488 conjugate to phosphatidyl serine present on the membranes of apoptotic cells was followed in time by imaging the cells every hour after drug exposure with a BD Pathway 855 imager (Becton Dickinson, Erembodegem, Belgium). The total area of annexin V-Alexa488 fluorescence per image was quantified using Image Pro (Media Cybernetics, Bethesda, MD).

Western Blot.

Cells were harvested as previously described (de Graauw et al., 2005). The samples were subjected to protein separation and blotted on Immobilon-P membranes (Millipore, Amsterdam, The Netherlands).The membranes were blocked overnight at 4°C in milk powder 5% (w/v) in Tris-buffered saline/Tween 20. Primary antibody incubation was performed for 1 hour at room temperature followed by incubation with horseradish peroxidase–conjugated or Cy5-labeled secondary antibodies (Jackson ImmunoResearch, Newmarket, UK) in Tris-buffered saline/Tween 20 for 1 hour at room temperature. Protein signals were detected with enhanced chemiluminescence (GE Healthcare, Diegem, Belgium) followed by film detection for the NF-κB members or visualization on the Typhoon 9400 imager (GE Healthcare) for β-actin.

Immunofluorescence.

Cultured cells were immunostained for β-catenin, P-paxillin-Y118, and F-actin (rhodamine/phalloidin) followed by secondary antibody labeling (goat antimouse Alexa488-labeled, goat antirabbit Alexa488-labeled (Molecular Probes, Breda, The Netherlands), and goat antimouse Cy3-labeled (Jackson ImmunoResearch). Hoechst 33258 (2 µg/ml) was used to visualize the nuclei. Cells were imaged using a Nikon TiE2000 confocal microscope (Nikon, Amstelveen, The Netherlands). All antibodies were used with a 1:1000 dilution.

Quantitative Analysis of Cell-Cell Junctions and Focal Adhesions.

β-Catenin segmentation and quantification were performed using Image J 1.44i software (NIH, Bethesda, MD), as previously described (Qin et al., 2012), by calculating the average area occupied by β-catenin fluorescence signal in the cell-cell junction region of each cell and expressing it as β-catenin junction size per nucleus. Focal adhesion quantification was performed using Image J 1.43h software. Segmentation of the two signals P-paxillin-Y118 and Hoescht was followed by artificial cell region border creation. With the segmented P-paxillin-Y118 and nucleus masks together with the reconstructed cell region border, several phenotypic measurements were derived, including focal adhesion size, dispersion, orientation, and number. A minimum of 3400 focal adhesions and 100 cells per condition was used for the quantification. The focal adhesions size distribution was represented as empirical cumulative distribution function curves using the cdfplot function of the MATLAB software (MathWorks, Eindhoven, The Netherlands). The empirical cdf F(x) is defined as the proportion of X values less than or equal to x.

Gene Array Analysis.

IM-PTECs wild-type (wt), control shRNA (shCtrl) cells, and shRNA RelB (shRelB) cells were exposed to vehicle, cisplatin (20µM), TNF-α (8 ng/ml), or cisplatin in combination with TNF-α for 8 hours. Three replicates for each treatment were used. Total RNA was isolated from cells using an RNeasy Mini Kit (Qiagen, Venlo, The Netherlands) according to the manufacturer’s protocol. RNA concentration was determined by absorbance at 260 nm (NanoDrop Technologies, Montchanin, DE), and the integrity of the RNA was checked using an Agilent 2100 Bioanalyzer (Agilent Technologies, Palo Alto, CA). The synthesis of labeled complement RNA (cRNA) and hybridization steps were performed by Service XS (Leiden, The Netherlands). Briefly, 100 ng of RNA was used to synthesize biotin-labeled cRNA with the Affymetrix 3′ IVT Express Labeling Kit (Affymetrix, Santa Clara, CA). The resulting labeled cRNA was quantified by absorbance at 260 nm (NanoDrop), and 6 μg was hybridized on an Affymetrix 24 all-in HT MG 430 PM Array Plate following the procedures recommended by the manufacturer. After hybridization, the microarray slide was washed, dried, and scanned using the Hybridization, Wash and Stain Kit #901530 compatible with Affymetrix protocols. The Affymetrix Command Console (version 3.0) and Expression Console software (version 1.1) were used to analyze the washing, staining, and scanning of the chips.

Subsequently, the raw intensity data were normalized with BRB-ArrayTools software (developed by Dr. Richard Simon and the BRB-ArrayTools Development Team; http://linus.nci.nih.gov/BRB-ArrayTools.html) using the robust multichip average method. Significantly differentially expressed genes [false discovery rate (FDR) ≤ 0.1) between the various experimental conditions were identified with a one-way blocked analysis of variance test. A randomized blocked design was chosen to control for the different days that the replicate experiments were run. The analysis of variance test was followed by calculation of the FDRs according to Benjamini and Hochberg (1995). Since only one differentially expressed gene was found between wt and shCtrl cells, they were pooled to increase the statistical power of the analysis. Only probes with expression levels displaying a 1.5-fold change or more relative to wt+shCtrl cells were selected for further analysis.

Commonly deregulated genes between the different groups were identified by Venn diagrams. Pathway analysis of the selected genes was performed using the GeneGo MetaCore pathway analysis software (GeneGo, St. Joseph, MI). Heat map representations were performed using the MultiExperiment Viewer software (Dana-Farber Cancer Institute, Boston, MA). The complete data set will be submitted to ArrayExpress (EMBL-European Bioinformatics Institute, Hinxton, UK; http://www.ebi.ac.uk/arrayexpress) and will be available for public download July 2013. The accession number referencing this data set will be available on the Gene Expression Omnibus database (http://www.ncbi.nlm.nih.gov/geo/).

Statistical Procedures.

All data are expressed as the mean ± S.E.M. Statistical significance was determined by GraphPad Prism (GraphPad Software, La Jolla, CA) using an unpaired two-tailed t test. Data from focal adhesion quantification were analyzed using the nonparametric Kolmogorov-Smirnov test via the MATLAB software (MathWorks), which computes test statistics that are derived from the empirical cdf. The cumulative distribution function curves are represented, and the level of confidence is indicated by P values in the figures.

Results

TNF-α Aggravates Cisplatin-Induced Disruption of Stress Fibers, Cell-ECM, and Cell-Cell Adhesion.

In a recent study, we showed that TNF-α enhanced cisplatin-induced cell death in IM-PTECs (Benedetti et al., 2012). As cisplatin alone is known to induce actin reorganization and loss of cell-cell and cell-ECM adhesion prior to apoptosis of renal tubular cells (Kruidering et al., 1998; van de Water et al., 2000; Imamdi et al., 2004; Qin et al., 2012), we determined whether TNF-α could promote these events. IM-PTECs were exposed to cisplatin (20 µM) and/or TNF-α (8 ng/ml) for 8 hours, and thereafter immunostained to visualize the actin cytoskeleton, focal adhesions, and cell-cell adhesions. Treatment of the cells with TNF-α alone enhanced actin fiber formation in particular at cell-cell junctions (Fig. 1A), but diminished the average size of focal adhesions (Fig. 1, B and D) and reduced the accumulation of β-catenin molecules at the cell membrane (Fig. 1, C and E). This effect of TNF-α aggravated the reorganization and disruption of the stress fibers (Fig. 1A), reduced the size of the focal adhesions (Fig. 1, B and Di), and mildly aggravated the loss of β-catenin at the cell-cell contacts (Fig. 1, C and E) caused by cisplatin treatment alone. TNF-α only mildly affected the reduction in the number of focal adhesions induced by cisplatin alone (Fig. 1Dii). These data show that, prior to the onset of apoptosis, TNF-α aggravates the cytoskeletal and cell adhesion reorganization induced by cisplatin alone.

Fig. 1.
Fig. 1.

TNF-α aggravates cisplatin-induced disruption of stress fibers, cell-ECM adhesion, and cell-cell contacts. IM-PTECs were exposed to cisplatin (20 µM) and/or TNF-α (8 ng/ml) for 8 hours and immunostained to visualize the actin cytoskeleton, focal adhesions, and cell-cell adhesions. Cells were imaged using a Nikon TiE2000 confocal microscope. The effect of treatment on the actin cytoskeleton was visualized by staining of F-actin with rhodamine phalloidin (red) (A). The effect of treatment on focal adhesions (FA) was assessed by staining of P-paxillin-Y118 (green) (B). Quantification of P-paxillin was performed. The cumulative distribution function curve indicates the direction of the changes in focal adhesion size with the different treatments. A Kolmogorov-Smirnov test was performed relative to control treatment and indicates the differences observed in P-paxillin size between cisplatin and/or TNF-α–treated cells and vehicle-treated cells. The P values obtained are indicated in red. The number of focal adhesions was also assessed by quantifying P-paxillin fluorescence per cell (D). Cell-cell contact disturbances after treatment were assessed by staining the adherens junction protein β-catenin (green) (C), which was quantified by assessing the average area occupied by β-catenin per cell (E). For all images, the nuclei were stained with Hoechst (blue) (A–C). The scale is indicated in the images in the bottom-left corner. The data are representative of three independent experiments and expressed as the mean ± S.E.M. *P ≤ 0.05; #P ≤ 0.05; and ##P ≤ 0.01 compared with vehicle-treated cells. Cispt, cisplatin.

RelB Knockdown Suppresses Cisplatin/TNF-α Synergistic Apoptosis.

TNF-α activates NF-κB signaling. Furthermore, NF-κB signaling can affect the actin cytoskeletal organization (Nakamichi et al., 2007). The NF-κB transcription factor comprises five different members: RelA, RelB, NF-κB1, NF-κB2, and c-Rel (Hoffmann and Baltimore, 2006). Our next goal was therefore to determine which NF-κB subunit could contribute to the TNF-α–mediated aggravation of both cisplatin-induced cytoskeletal reorganization and cell death. Therefore, we first systematically generated knockdown IM-PTEC cell lines using five individual lentiviral shRNA targeting each individual NF-κB family member. Knockdown was confirmed using Western blotting (Supplemental Fig. 1, top panel). For NF-κB1, only one, instead of two, out of five shRNA lentiviral vectors resulted in a sufficient and reliable protein knockdown. Due to its very weak expression in our IM-PTECs determined from previous Affymetrix array data (Benedetti et al., 2012), c-Rel was not considered for further investigation.

Next, we investigated the effect of the knockdown of each NF-κB member on cisplatin- and cisplatin/TNF-α–induced apoptosis. All cell lines were exposed to cisplatin (20 µM) and/or TNF-α (8 ng/ml), and cell death was measured using live apoptosis imaging. RelA and NF-κB1 knockdown increased cisplatin- and cisplatin/TNF-α–induced apoptosis (Fig. 2A), suggesting a role for the RelA/p50 NF-κB dimer in survival signaling. Interestingly, for both NF-κB2 knockdown cell lines, an increased cisplatin-induced apoptosis was observed, whereas cisplatin/TNF-α–induced apoptosis was unaffected (Fig. 2A). This observation indicates that, in our IM-PTECs, NF-κB2–based NF-κB signaling protects the cells from cisplatin-induced apoptosis. Importantly, knockdown of RelB using two independent shRNAs suppressed the cisplatin/TNF-α–induced synergistic apoptosis without affecting cisplatin-induced apoptosis (Fig. 2A). This suggests that RelB is the only NF-κB family member that is key in the regulation of the TNF-α–mediated enhancement of cisplatin-induced apoptosis. Of critical relevance, TNF-α itself increased expression of RelB at both the gene as well as the protein expression level, which was not affected by cisplatin (Fig. 2, B and C). Together, these data indicate that the NF-κB family member RelB is directly involved in the proapoptotic activity of TNF-α and, thereby, enhances cisplatin-induced apoptotic cell death.

Fig. 2.
Fig. 2.

RelB knockdown suppresses cisplatin/TNF-α synergistic apoptosis. IM-PTEC cell lines with knockdown for each NF-κB family member (NF-κB1, NF-κB2, RelA, and RelB) were generated using lentiviral shRNA (see Materials and Methods). Knockdown cells were exposed to cisplatin (20 µM) and/or TNF-α (8 ng/ml). Cell death was followed over time using Annexin V-Alexa488 staining and automated imaging. Both apoptosis over time and endpoints are represented. The dashed orange and red lines indicate the amount of apoptosis observed with shCtrl cells after cisplatin or cisplatin/TNF-α treatment, respectively (A). After cisplatin (5 µM) and/or TNF-α (8 ng/ml) treatment for 8 hours, the gene expression (B) and protein (C) levels of RelB were assessed. Gene expression values were obtained from previous Affymetrix gene array data (Benedetti et al., 2012) (B) and protein levels were assessed by Western blotting (C). Β-actin was used as a loading control, and the quantification indicated is the mean of three independent experiments (C). The data are representative of three independent experiments and expressed as the mean± S.E.M. *P ≤ 0.05; **P ≤ 0.01; #P ≤ 0.05; and ##P ≤ 0.01 compared with vehicle-treated cells.

RelB Knockdown Inhibits Cisplatin/TNF-α–Induced Cell Adhesion and Actin Cytoskeleton Changes.

Next, we determined whether depletion of RelB also affected the TNF-α- and cisplatin-mediated changes in the actin cytoskeletal network, cell-matrix adhesions, and cell-cell contacts. Cisplatin caused loss of F-actin organization and cell matrix and cell-cell adhesions in shCtrl cells, as expected (Fig. 3, A–E), which was further enhanced by TNF-α. Knockdown of RelB resulted in some morphologic changes compared with both wt and shCtrl cells. shRelB cells were more spread with a reorganization of stress fibers around the nucleus, larger focal adhesions, and a reduced cortical-actin network. In addition, a loss in cell-cell contact formation was observed as indicated by the disappearance of β-catenin at the cell-cell border (Fig. 3, C and E). Although we expected these changes to influence cisplatin-induced renal cell death, no significant changes were observed between dimethylsulfoxide and cisplatin-treated cells. However, in sharp contrast to shCtrl cells, treatment of shRelB cells with cisplatin/TNF-α did not result in drastic morphologic changes: cisplatin/TNF-α–treated shRelB cells maintained their actin organization and did not show significant changes in focal adhesion size and number (Fig. 3, A, B, and D; Supplemental Fig. 2A), coinciding with the protection of the cells. Furthermore, although slightly diminished, β-catenin containing cell-cell adhesions remained largely intact during cisplatin/TNF-α treatment of shRelB cells compared with untreated cells (Fig. 3, C and E). Together, these data indicate that RelB-mediated signaling is essential in the nephrotoxic cytoskeletal reorganization in association with the onset of cell death during cisplatin/TNF-α treatment.

Fig. 3.
Fig. 3.

RelB knockdown inhibits cisplatin/TNF-α–induced cell adhesion and actin cytoskeleton changes. shCtrl and shRelB cells were exposed to cisplatin (20 µM) and/or TNF-α (8 ng/ml) for 8 hours and immunostained to visualize the actin cytoskeleton, focal adhesions, and cell-cell adhesions. Cells were imaged using a Nikon TiE2000 confocal microscope. The effect of RelB knockdown on actin cytoskeleton was visualized by the staining of F-actin with rhodamine phalloidin (red) (A). The effect of RelB knockdown on focal adhesions was assessed by staining of P-paxillin-Y118 (green) (B). Cell-cell contact disturbances after treatment were assessed by staining the adherens junction protein β-catenin (green) (C). Quantification of P-paxillin (D) and β-catenin (E) was performed as described in Fig. 1 and Materials and Methods. For all images, the nuclei were stained with Hoechst (blue) (A–C). Images of shCtrl cells and shRelB cells have the same scale as indicated in the images in the bottom-left corner. Cispt, cisplatin; cispt+ T, cisplatin+TNF-α. The data are representative of three independent experiments and expressed as the mean ± S.E.M. *P ≤ 0.05; #P ≤ 0.05; and ##P ≤ 0.01 compared with vehicle-treated cells.

RelB Knockdown Leads to EMT-Like-Driven Cytoskeletal Reorganization.

To obtain detailed insight into the pathways involved in the actin cytoskeleton changes and the suppression of the synergistic apoptosis observed in shRelB cells, transcriptomic analysis was performed. Wt, shCtrl, and shRelB cells were exposed to cisplatin (20 µM) and/or TNF-α (8 ng/ml) for 8 hours, a time point at which cell death did not yet occur (Fig. 2). Since wt cells and shCtrl cells only had one statistically different gene (Neurog3) with an FDR ≤ 0.1, we pooled the data sets to increase the statistical power of our gene array analysis. We first analyzed the effect of RelB knockdown on gene expression changes under control conditions. Gene expression changes with ≥1.5-fold change and FDR ≤ 0.1 for shRelB #1 and #2 cells compared with wt + shCtrl cells were considered for further analysis. A total of 800 genes for shRelB #1 and 698 genes for shRelB #2 were identified with our selection criteria, and 388 genes were common for both shRelB cell lines (Fig. 4A). A pathway analysis was then performed on these common genes under control conditions using the GeneGo MetaCore pathway analysis software. The top seven significantly different pathways were all related to cytoskeletal and cell adhesion changes as well as induction of an EMT (Fig. 4B). The most prominent cytoskeleton and cell adhesion–related genes that were upregulated by RelB knockdown included actin, myosin IIB, the gene encoding the tight junction-related protein cingulin, the cell adhesion–related genes collagen type IV α1 and serpine 2, as well as the genes encoding the Cdc42 regulating proteins Cdc42 small effector 1 and Par6 (Fig. 4C). In contrast, Vcan, encoding the cell adhesion protein versican, was downregulated in shRelB cells (Fig. 4C). In addition, one of the key EMT-inducing genes, Snai2, encoding the protein Snai2, was more than 2-fold upregulated in both shRelB cell lines (Fig. 4C), suggesting that the cytoskeletal changes described earlier could well be a consequence of a Snai2-induced EMT-like switch of the PTEC cells. Other EMT-related transcription factors, including Twist, were not affected.

Fig. 4.
Fig. 4.

RelB knockdown leads to EMT-like-driven cytoskeletal rearrangements in vehicle-treated cells. shCtrl and shRelB cells were exposed to cisplatin (20 µM) and/or TNF-α (8 ng/ml) for 8 hours. Following hybridization and robust multichip average (RMA) normalization, the genes with ≥1.5-fold change and FDR ≤ 0.1for shRelB #1 and #2 cell lines compared with wt + shCtrl cells in control conditions were further selected. A Venn diagram was made comparing both shRelB cell lines in control conditions (A). Using the MetaCore software, pathway analysis was performed on the genes commonly regulated in both shRelB cell lines in control conditions. The top pathways are represented in a heat map (B). The gene expression changes of genes involved in cytoskeleton organization and EMT are represented for wt and shCtrl as well as both shRelB cell lines in control conditions. The data are representative of three independent experiments and expressed as the mean ± S.E.M. *P ≤ 0.05 (C). Actb, actin; Cdc42se1, Cdc42 small effector 1; Cgn, cingulin; Col4a1, collagen type IV α1; ILK, integrin-linked kinase; Myh10, myosin IIB; Pard6g, Par6; PI3K, phosphoinositide 3-kinase; TGF, tumor growth factor; Vcan, versican.

RelB Knockdown Leads to Rho Kinase Pathway Activation During Cisplatin/TNF-α Treatment.

We next evaluated which pathways were affected by RelB knockdown under TNF-α, cisplatin, or TNF-α/cisplatin conditions. Gene expression changes with ≥1.5-fold change and FDR ≤ 0.1for shRelB #1 and #2 cells compared with wt + shCtrl cells were considered for further analysis. Comparing differentially expressed genes between treatments resulted in identification of 21 and 17 uniquely changing genes for cisplatin and TNF-α treatments, respectively, compared with 118 and 70 uniquely changing genes for vehicle and cisplatin/TNF-α treatments, respectively (Supplemental Fig. 3; Supplemental Table 1).

To identify pathways by which RelB controlled the synergistic apoptosis induced by cisplatin/TNF-α treatment, pathway analysis was performed on the common genes in cells treated with the combined cisplatin/TNF-α treatment using the GeneGo MetaCore pathway analysis software. A total of 782 genes for shRelB #1 and 462 genes for shRelB #2 were identified under these treatment conditions and 278 genes were common for both shRelB cell lines (Fig. 5A) comprising the 70 unique genes (Supplemental Fig. 3; Supplemental Table 1). Pathway analysis showed that, also under cisplatin/TNF-α conditions, the top significantly different pathways were all related to cytoskeletal and cell adhesion changes as well as induction of an EMT (Fig. 5B). The main genes involved in cell adhesion included the Ras-related protein 1a (Rap1a), the Eph receptor B3, and the gap junction protein β2 (Gjb2) (Fig. 5C), of which expression of Rap1a and Gjb2 was specifically changed under cisplatin/TNF-α conditions. Interestingly, the EMT gene Snai2 was slightly induced in shCtrl cells; yet, depletion of RelB allowed an even further induction of Snai2 expression after cisplatin treatment, which was approximately four times higher than under shCtrl conditions and independent of TNF-α (Fig. 5C). In addition, Rho GTPase pathway activation was observed in shRelB IM-PTECs (Fig. 5B), including upregulation of Rho guanine nucleotide exchange factor 3 (Arhgef3) and rhophilin 1 (Fig. 5C). Together, these data suggest a key role for RelB to suppress a stress-induced cytoskeletal reorganization response, which is likely part of an EMT-like program.

Fig. 5.
Fig. 5.

RelB knockdown is associated with upregulation of RhoA pathway–related genes during cisplatin/TNF-α treatment. shCtrl and shRelB cells were exposed to cisplatin (20 µM) and/or TNF-α (8 ng/ml) for 8 hours. Following hybridization and robust multichip average (RMA) normalization, the genes with ≥1.5-fold change and FDR ≤ 0.1 for shRelB #1 and #2 cell lines compared with wt + shCtrl cells after cisplatin/TNF-α treatment were further selected. A Venn diagram was made comparing both shRelB cell lines (A). Using the MetaCore software, pathway analysis was performed on the genes commonly regulated in both shRelB cell lines with cisplatin/TNF-α treatment. The top pathways are represented in a heat map (B). The gene expression changes of genes involved in cell adhesion, EMT, and the RhoA pathway are represented for wt-shCtrl cells combined and both shRelB cell lines combined in cisplatin/TNF-α treated conditions. The statistical significance is indicated only for the conditions under which the fold-change was ≥1.5 for both shRelB #1 and #2. The data are representative of three independent experiments and expressed as the mean ± S.E.M. *P ≤ 0.05; and **P ≤ 0.05 (C). Ephb3, Eph receptor B3; ILK, integrin-linked kinase; PI3K, phosphoinositide 3-kinase; Rhpn1, rhophilin 1; TGF, tumor growth factor.

RelB Knockdown Protects Against Cisplatin/TNF-α Treatment in a Rho Kinase–Dependent Manner.

The Rho GTPase family is the most important family of proteins responsible for the regulation of the actin cytoskeletal network in cells, and it is known to be involved in PTEC injury (Kroshian et al., 1994; Raman and Atkinson, 1999). Given the changes observed in the expression of some Rho GTPase family members in shRelB cells, we determined whether RelB knockdown protects the IM-PTECs against cisplatin/TNF-α–induced synergistic cell death via activation of the Rho GTPase pathway. Therefore, we inhibited one of the key downstream components, Rho kinase using the Y27632 inhibitor. Cells were pre-exposed for 30 minutes with the ROCK inhibitor and thereafter treated with cisplatin (20 µM) and/or TNF-α (8 ng/ml). Y27632 caused disruption of stress fiber formation in both shCtrl and shRelB cells (Fig. 6A). This disruption of the stress fibers was accompanied by an enhancement of apoptosis in shCtrl cells after both cisplatin and cisplatin/TNF-α treatment (Fig. 6B). In shRelB cells, apoptosis was only increased with cisplatin/TNF-α treatment and not with cisplatin treatment alone (Fig. 6B). These data are indicative of a mechanism whereby RelB knockdown confers protection against cisplatin/TNF-α treatment in a Rho kinase–dependent manner.

Fig. 6.
Fig. 6.

RelB knockdown protects against cisplatin/TNF-α treatment in a Rho kinase–dependent manner. shCtrl and shRelB cells were pre-exposed to the ROCK inhibitor Y27632 for 30 minutes prior to cisplatin (20 µM) and/or TNF-α (8 ng/ml). After 8 hours of exposure to cisplatin and/or TNF-α in combination with the ROCK inhibitor, actin was visualized by staining of F-actin with rhodamine phalloidin (red). The nuclei were visualized with Hoechst staining (blue). Images of both shCtrl cells and shRelB cells have the same scale as indicated in the bottom-left corner (A). Cell death after ROCK inhibition was followed over time using Annexin V-Alexa488 staining and automated imaging (B). cispt, cisplatin; cispt+T, cisplatin+TNF-α; ctl, control. The data are representative of three independent experiments and expressed as the mean ± S.E.M. *P ≤ 0.05; #P ≤ 0.05; and ##P ≤ 0.01 compared with vehicle-treated cells.

Discussion

In the present study, we demonstrate that TNF-α aggravated the cisplatin-induced changes in the actin cytoskeleton and cell adhesions that occur prior to the onset of apoptosis. Knockdown of the NF-κB family member RelB abrogated both the synergistic cisplatin/TNF-α–induced apoptosis as well as the changes in actin cytoskeleton and cell adhesions. RelB knockdown resulted in an enhancement of the expression of Rho GTPase–related actin cytoskeletal regulators in association with a Snai2-related EMT-like program. Likewise, a Rho kinase inhibitor, Y27632, reversed the cytoprotective effect of the RelB depletion.

Although the transcription factor NF-κB has been associated with morphologic changes that occur prior to apoptosis (Nakamichi et al., 2007), the role of the family member RelB in Rho kinase regulation in relation to cell death has never been described before. Our study showed that upregulation of RelB is responsible for the synergistic apoptosis induced by cisplatin/TNF-α treatment. This observation is contradictory to studies using thymocytes and cancer cells, in which RelB is involved in antiapoptotic signaling (Guerin et al., 2002; Mineva et al., 2007; Wang et al., 2007); these effects were independent of TNF-α signaling. Moreover, different transcriptional programs may be activated by RelB in immune and cancer cells, compared with renal cells. Jacque et al. (2005) showed in mouse embryonic fibroblasts that, although RelB entered the nucleus in response to TNF-α, it could not bind to DNA due to repression by RelA. In our IM-PTECs, TNF-α–induced RelA translocation was inhibited by cisplatin/TNF-α treatment (Benedetti et al., 2012). Furthermore, RelB expression was upregulated during TNF-α treatment (Fig. 2, B and C). It could be that the increase in apoptosis in our cells is due to the combination of RelB upregulation and RelA translocation inhibition, thereby resulting in a different transcriptional program than the one occurring in immune and cancer cells. Additionally, other kinases regulating the cytoskeletal organization, such as Src, could be involved. v-Src has been shown to regulate the activation of RelB via accelerating nuclear factor of κ light polypeptide gene enhancer in B-cells inhibitor (IκB) proteolysis in fibroblasts (Shain et al., 1999), and TNF-α increased Src activity in intestinal epithelial cells (Kawai et al., 2002). It could be that, in our cells, the increased RelB activity observed with TNF-α and cisplatin/TNF-α treatments is due to increased Src activation.

Transcriptomic analysis revealed activation of the RhoA/Rho kinase pathway in shRelB cells during treatment of the cells with cisplatin/TNF-α via increased gene expression of ARHGEF3 and rhophilin 1. ARHGEF3 has been shown to activate RhoA and RhoB specifically leading to assembly of stress fibers and thereby enlargement of focal adhesions in a Rho kinase–dependent manner (Arthur et al., 2002). Rhophilin 1 interacts strongly with GTP-bound RhoA, but its exact role in cytoskeletal organization remains elusive (Watanabe et al., 1996; Peck et al., 2002). Their role in apoptosis is unknown. Using the Rho kinase inhibitor, we showed that RhoA pathway activation was crucial for cisplatin/TNF-α–induced synergistic apoptosis. It is well known that, depending on the cell type and stimulus used, activation of the RhoA/ROCK pathway can lead to either apoptosis or cell survival. For example, ROCK inhibition enhanced cisplatin-induced cytotoxicity of lung carcinoma cells through a focal adhesion kinase suppression-independent mechanism (Igishi et al., 2003), whereas it attenuated I/R-induced renal injury in an in vivo model (Song and Gao, 2011). In addition, several other studies indicated a protective role of ROCK in several cell types, including epithelial, cancer, and endothelial cells, as well as in vivo (Amano et al., 2010). Yet, to the best of our knowledge, the role of ROCK signaling in cisplatin-induced apoptosis in renal cells remains thus far unknown.

In addition to RhoA and ROCK pathway activation, our transcriptomic analysis demonstrated upregulation of the EMT-regulatory gene Snai2 in shRelB cells, which corresponded with the EMT-like switch observed in these cells. Since TNF-α strongly induces RelB expression (Fig. 2C), this may suppress a cytoprotective Snai2-based EMT-like switch initiated by cisplatin. RhoA pathway activation was shown to be a key step in renal EMT, leading to transdifferentiation of renal tubular cells to fibroblasts (Liu, 2004, 2010). It has been shown that renal regeneration during and after toxicant- or I/R-induced renal failure occurred by transformation of epithelial cells into fibroblasts via EMT. During an EMT, cells may acquire new mesenchymal markers such as vimentin, α smooth muscle actin, S100 calcium binding protein A4, the transcription factor Snai2, and N-cadherin (Liu, 2010). Although Snai2 was upregulated more than 2-fold, no changes occurred in gene expression of the mesenchymal markers vimentin, desmin, collagen I, fibronectin, and N-cadherin. α Smooth muscle actin gene expression was only increased in the shRelB #2 cells showing the highest knockdown, but not in the shRelB #1 cells, whereas the S100 calcium binding protein A4 gene expression was downregulated in both shRelB cell lines. Therefore, RelB knockdown did not induce a complete traditional EMT in IM-PTECs, but this partial EMT accompanied by RhoA/Rho kinase activation was sufficient to confer protection against the synergistic apoptosis induced by cisplatin/TNF-α treatment. In contrast to our data, Wang et al. (2007) showed that RelB expression was required to maintain the mesenchymal phenotype of breast cancer cells. The link between RhoA/ROCK pathway activation and RelB in this process is not yet clear.

Last, our transcriptomic analysis revealed specific and significant changes in the expression of the cell-cell proteins Rap1a and Gjb2 in cisplatin/TNF-α–treated shRelB cells. We have previously shown that Rap1a preserved cell-cell junctions and protected against cisplatin-induced apoptosis in proximal tubular cells (Qin et al., 2012). Gjb2 is part of the connexins that could be involved in the proximal tubular cells in the formation of hemichannels regulating renal sodium, potassium, and water handling (Hanner et al., 2010). In mice, knockout of GJB2 has been shown to be responsible for cochlear cell apoptosis, so it could be that this connexin also has a protective role in renal cells (Zhang et al., 2012).

In conclusion, our observations suggest that TNF-α–induced upregulation of RelB expression is responsible for the cisplatin/TNF-α–induced synergistic apoptosis and show, for the first time, the role of RelB in renal EMT induction and RhoA/ROCK-mediated cytoskeletal regulation as a protective mechanism.

Authorship Contributions

Participated in research design: Benedetti, Fredriksson, de Graauw, van de Water.

Conducted experiments: Benedetti.

Contributed new reagents or analytic tools: Yan, Herpers.

Performed data analysis: Benedetti, Fokkelman, Meerman, de Graauw.

Wrote or contributed to the writing of the manuscript: Benedetti, de Graauw, van de Water.

Footnotes

    • Received November 26, 2012.
    • Accepted April 26, 2013.

  • This work was performed as part of the Netherlands Toxicogenomics Center/Netherlands Genomics Initiative supported by the Dutch Organization for Scientific Research [Grant 050-06-510]; and the Dutch Top Institute Pharma project [Grant D3-201].

  • dx.doi.org/10.1124/mol.112.084053.

  • ↵Embedded ImageThis article has supplemental material available at molpharm.aspetjournals.org.

Abbreviations

ARHGEF3
Rho guanine nucleotide exchange factor 3
cisplatin
cis-diamminedichloroplatinum(II)
cRNA
complement RNA
ECM
extracellular matrix
EMT
epithelial mesenchymal-like transition
FDR
false discovery rate
Gjb2
gap junction protein β2
I/R
ischemia/reperfusion
IM-PTEC
immortalized proximal tubular epithelial cell
NF-κB
nuclear factor κB
PTEC
proximal tubular epithelial cell
Rap1a
Ras-related protein 1a
ROCK
Rho-associated protein kinase
shCtrl
control shRNA
shRelB
shRNARelB
shRNA
small hairpin RNA
TNF-α
tumor necrosis factor α
wt
wild-type
Y27632
(R)-(+)-trans-N-(4-pyridyl)-4-(1-aminoethyl)-cyclohexanecarboxamide

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RelB in Cisplatin/TNF-α–Induced Synergistic Apoptosis

Giulia Benedetti, Michiel Fokkelman, Kuan Yan, Lisa Fredriksson, Bram Herpers, John Meerman, Bob van de Water and Marjo de Graauw
Molecular Pharmacology July 1, 2013, 84 (1) 128-138; DOI: https://doi.org/10.1124/mol.112.084053
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