December 2009
Volume 50, Issue 12
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Retinal Cell Biology  |   December 2009
Dual Involvement of Caspase-4 in Inflammatory and ER Stress-Induced Apoptotic Responses in Human Retinal Pigment Epithelial Cells
Author Notes
  • From the Department of Ophthalmology, University of Michigan, Ann Arbor, Michigan. 
  • Corresponding author: Victor M. Elner, Department of Ophthalmology, University of Michigan, 1000 Wall Street, Ann Arbor, MI 48105; velner@med.umich.edu
Investigative Ophthalmology & Visual Science December 2009, Vol.50, 6006-6014. doi:10.1167/iovs.09-3628
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      Zong-Mei Bian, Susan G. Elner, Victor M. Elner; Dual Involvement of Caspase-4 in Inflammatory and ER Stress-Induced Apoptotic Responses in Human Retinal Pigment Epithelial Cells. Invest. Ophthalmol. Vis. Sci. 2009;50(12):6006-6014. doi: 10.1167/iovs.09-3628.

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      © ARVO (1962-2015); The Authors (2016-present)

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Abstract

Purpose.: To investigate the functional involvement of caspase-4 in human retinal pigment epithelial (hRPE) cells.

Methods.: Expression and activation of caspase-4 in hRPE cells were measured after stimulation with proinflammatory agents IL-1β (2 ng/mL), TNF-α (20 ng/mL), lipopolysaccharide (1000 ng/mL), interferon-γ (500 U/mL), or monocyte coculture in the absence or presence of immunomodulating agent cyclosporine (3 or 30 ng/mL), dexamethasone (10 μM), or IL-10 (100 U/mL) and endoplasmic reticulum (ER) stress inducer thapsigargin (25 nM) or tunicamycin (3 or 10 μM). The onset of ER stress was determined by expression of GRP78. The involvement of caspase-4 in inflammation and apoptosis was further examined by treating the cells with caspase-4 inhibitor Z-LEVD-fmk, caspase-1 and -4 inhibitor Z-YVAD-fmk, and pan-caspase inhibitor Z-VAD-fmk.

Results.: Caspase-4 mRNA expression and protein activation were induced by all the proinflammatory agents and ER stress inducers tested in this study. Caspase-4 activation was blocked or reduced by dexamethasone and IL-10. Elevated ER stress by proinflammatory agents and ER stress inducers was shown by increased expression of the ER stress marker GRP78. The induced caspase-4 and caspase-3 activities by tunicamycin and the stimulated IL-8 protein expression by IL-1β were markedly reduced by caspase-4 inhibitor Z-LEVD-fmk. Although caspase-4 inhibitor Z-LEVD-fmk and caspase-1 and -4 inhibitor Z-YVAD-fmk reduced tunicamycin-induced hRPE apoptotic cell death by 59% and 86%, respectively, pan-caspase inhibitor Z-VAD-fmk completely abolished the induced apoptosis.

Conclusions.: Caspase-4 is dually involved in hRPE proinflammatory and proapoptotic responses. Various proinflammatory stimuli and ER stress induce hRPE caspase-4 mRNA synthesis and protein activation. ER stress-induced hRPE cell death is caspase and, in part, caspase-4 dependent.

Caspases are a family of cytosolic, aspartate-specific, cysteine proteases involved in apoptosis, inflammation, proliferation, and differentiation. 14 At least 17 members of the caspase family have been identified, of which 13 are found in humans. 5 Human caspase-4 was cloned independently in three laboratories and designated as ICH2, 6 ICErelII, 7 and TX. 8 The caspase-4 gene is expressed ubiquitously in various tissues with the exception of brain. 6,7 Although human caspase-4 has no corresponding mouse orthologue, 1 human caspase-4 and -5 are possibly the orthologues of mouse caspase-11. 1 Caspase-4 cDNA exhibits 68% sequence homology with human caspase-1. 7 As with caspase-1, caspase-4 is composed of a large prodomain (p22) and two small domains (p20 and p10) that are cleaved on activation. 7 Transient expression of the cloned caspase-4 gene causes apoptotic cell death in fibroblasts, 7 Sf9 insect cells, 6 and COS cells. 8 Subsequent studies have confirmed the apoptotic role of caspase-4 in endoplasmic reticulum (ER) stress-induced cell death. 912  
The ER is responsible for folding, maturation, and storage of membrane and secreted proteins. ER is also the major organelle that stores second-messenger calcium ions, which sense and respond to changes in cellular homeostasis. ER stress occurs when the cellular demand for ER function exceeds its capacity. Overloading of unfolded protein aggregates triggers a signaling cascade of events, called unfolded protein response (UPR). Excess UPR leads to irreversible commitment to cell death. There is accumulating evidence to suggest the involvement of caspase-4 in ER stress-induced apoptosis. First, caspase-4 is localized mainly to the ER. 9 Second, caspase-4 is closely associated with many essential proteins in ER stress-induced cell death pathways, including GRP78, a well-known marker of ER stress 10 ; CARD-only protein (Cop or pseudoICE), a regulator of procaspase-1 11 ; Apf1, a protein involved in death protease-mediated cell death 12 ; and TRAF6, a member of the TNF receptor-associated factor. 13 Third, caspase-4 inhibitor Z-LEVD-FMK (z-LEVD-fmk) selectively and effectively blocks ER stress-induced apoptosis in many cancer cells, such as neuroblastoma cells, 14 lung and esophageal cancer cells, 15 Jurkat cells, 16 and melanoma cells. 17 Fourth, knocking down caspase-4 expression by siRNA in multiple myeloma cells, 18 leukemia cells, 19 glioma cell lines, 20 and neuroblastoma cells, 9 introducing caspase-4 antisense oligonucleotides to lymphoblastoid AHH-1 cells, 21 expressing catalytically inactive caspase-4, and microinjecting anti–caspase-4 antibodies into HeLa cells 22 abolishes ER stress-induced cell death. Conversely, overexpression of caspase-4 in COS-7 cells induces activation of caspase-3 and -9, the two well-known death proteases. 23  
Chromosomal mapping reveals that the human caspase-4 gene is colocalized within a cluster of functionally related genes caspase-1, -5, and -12 and caspase-1 pseudogenes ICEBERG, COP, and INCA in human chromosome 11q22–23. 24 The chromosomal colocalization of caspase-4 with inflammatory caspases implies that these caspases are derived from a common ancestor through gene multiplication and that they share common functions in innate immunity and inflammation, despite the common acceptance that caspase-4 is a member of the inflammatory caspase family. Most previous functional studies have focused on the role of caspase-4 in apoptosis. Thus far, only one study has shown, by having demonstrated its role in lipopolysaccharide (LPS)-induced inflammatory responses, that caspase-4 is involved in inflammation. 13  
In this study we investigated the functional involvement of caspase-4 in hRPE cells. Our data showed that caspase-4 is involved in both inflammation and apoptosis in hRPE cells. 
Materials and Methods
Materials
Recombinant human IL-1β, TNF-α, interferon-γ (IFN-γ), and IL-10 were purchased from R&D Systems (Minneapolis, MN). Dexamethasone, cyclosporine, and tunicamycin were purchased from Sigma-Aldrich (St. Louis, MO). The cell-permeable general caspase inhibitor Z-VAD (OMe)-fluoromethylketone was from Bachem Americas, Inc. (Torrance, CA). The caspase-1 and -4 inhibitor Z-YVAD (OMe)-fmk and the caspase-4 inhibitor z-LEVD-fmk were from R&D Systems and BioVision (Mountain View, CA), respectively. The mouse monoclonal antibody (4B9) against caspase-4 was from Abcam (Cambridge, MA). The goat polyclonal antibody against GRP-78 was from Santa Cruz Biotechnology (Santa Cruz, CA). A shredding system (QIAshredder) and a purification kit (RNeasy Mini Kit) were purchased from Qiagen (Valencia, CA). Reverse transcription system was obtained from Invitrogen (Carlsbad, CA). RQ1 RNase-free DNase was purchased from Promega (Madison, WI). Detection kits (Cell Death Detection ELISA and In Situ Cell Death Detection) were purchased from Roche Molecular Biochemicals (Indianapolis, IN). A visualization kit (EnVision G/2 System/AP [Permanent Red]) was obtained from DAKO (Carpinteria, CA). All other reagents were obtained from Sigma-Aldrich. 
Cell Isolation and Culture
The hRPE cells were isolated within 24 hours of death from the donor eyes, as previously described. 2528 In brief, the sensory retina tissue was separated gently from the hRPE monolayer, and the hRPE cells were removed from Bruch's membrane with papain (5 U/mL). The hRPE cells were cultured in Dulbecco's modified Eagle's/Ham's F12 nutrient mixture medium, containing 15% fetal bovine serum, penicillin G (100 U/mL), streptomycin sulfate (100 μg/mL), and amphotericin B (0.25 μg/mL) in culture plates (Falcon Primaria; Becton Dickinson, San Jose, CA) to inhibit fibroblast growth. The hRPE monolayers exhibited uniform immunohistochemical staining for cytokeratin 8/18, fibronectin, laminin, and type IV collagen in the chicken-wire distribution characteristic for these epithelial cells. The identity of hRPE cells in the culture was confirmed by apical immunohistochemical staining of Na+-K+ ATPase. Cells were subcultured, grown to reach confluence, and exposed to the same medium but contained reduced serum (5%) for further experiments. The cells were in culture up to four to six passages. Three donors were used in this study. For each experiment at least two donors were used, and a typical result is shown. 
Monocyte Isolation and hRPE-Monocyte Coculture
Human monocytes were freshly isolated from the peripheral blood of healthy volunteers, as described previously. 26 In brief, peripheral blood was drawn into a heparinized syringe and 1:1 diluted in 0.9% saline. Mononuclear cells were separated by density gradient centrifugation. The cells were washed and then layered onto density gradient (Fico-Lite monocytes, 1.068 g/mL) for the enrichment of monocytes. The isolated cells were then washed, cytospun onto a glass slide, stained (Diff-Quick; Dade-Behring, Newark, DE), and differentially counted. The purity of the cell was >97%. For hRPE cells in monocyte coculture, enriched monocyte populations (3 × 105) were overlaid onto untreated or pretreated near-confluent hRPE cultures (2 × 105) for 6 hours. After coculture, the monocytes were removed as previously described, 26 and hRPE cells were subjected to further analyses. 
RNA Isolation and Reverse Transcription-Polymerase Chain Reaction
The total cellular RNA was isolated from hRPE cells (QIAshredder and RNeasy Mini Kit; Qiagen) according to the manufacturer's protocol. The cDNA synthesis reaction was set up according to the protocol for a reverse transcription system. Briefly, 5 μg RNA was added to the reaction mixture with a first-strand synthesis system (RT Superscript III [Invitrogen, Carlsbad, CA]; 200 U/μL) and 1 μL primer (Oligo(dT)20 [Invitrogen]; 0.5 μg/μL) in a total volume of 20 μL. Linear range of the β-actin PCR reaction was predetermined by using a series of 15 to 35 cycles. The mid-linear portion of the response curve was selected as the condition for semiquantitative PCR. The primer and condition for caspase-4 PCR were as described by Lin et al. 29 and confirmed by first examining three cycles (15, 25, and 35) and then cycles 30 and 32. The reaction was initiated by adding 0.15 μL Taq DNA polymerase (5 U/μL) to a final volume of 20 μL. Resultant cDNAs were amplified though 32 and 20 cycles for caspase-4 and β-actin, respectively. Primer sequences for human caspase-4 genes were 5′-CAGACTCTATGCAAGAGAAGCAACGTATGGCAGGA-3′ (forward) and 5′-CACCTCTGCAGGCCTGGACAATG ATGAC-3′ (reverse). To ensure that an equal amount of templates was used in each amplification reaction, human β-actin sense (5′-GTGGGGCGCCCCAGGCACCA-3′) and antisense (5′-GCTCGGCCGTGGTGGTGAAGC-3′) primers were used in parallel. The following conditions were used in RT-PCR reaction for caspase-4 and β-actin: denaturation at 95°C for 45 seconds (caspase-4) or 1 minute (β-actin), annealing at 65°C for 1 minute (caspase-4) or at 62°C (β-actin) for 1 minute, and extension at 72°C for 1 minute (caspase-4) or 2 minutes (β-actin) for 32 (caspase-4) or 20 (β-actin) cycles. RT-PCR products were analyzed by electrophoresis on a 2% agarose gel and stained with ethidium bromide. 
Assays for Apoptosis
ELISA (Cell Death Detection kit; Roche Molecular Biochemicals) was performed according to the manufacturer's protocol. Briefly, the hRPE cells were seeded and grown in 96-well plates until cells were close to confluence. After treating the cells with or without varieties of inducers or inhibitors for 24, 48, or 72 hours, apoptosis was quantified. Cultures were lysed with the lysis buffer of the ELISA kit (Cell Death Detection kit; Roche Molecular Biochemicals), and then the cytoplasmic fraction was transferred to the wells coated by streptavidin in the microplate modules for further analysis. Next, the immunoreagent was added to each well, which contained anti-histone-biotin and anti-DNA-peroxidase. The immunoreagents bonded to or reacted with the histone and DNA part of the mononucleosomes and oligonucleosomes that were out of the cytoplasm of cells during apoptosis. After removal of the unbound components with wash, the substrate 2,2′-azino-di-[3-ethylbenzthiazoline sulfonate] was added to determine the amount of peroxidase by reading absorbance difference between A405nm and A490nM in an ELISA reader as a measure of apoptosis. 
TUNEL Staining
The cells were stained with TdT-mediated dUTP nick end labeling (TUNEL) according to the manufacturer's protocol. Briefly, hRPE cells were fixed and incubated with TUNEL mixtures for 1 hour at 37°C. The incorporated fluorescein was detected by sheep anti-fluorescein antibody conjugated with horseradish peroxidase using the substrate diaminobenzidine. hRPE cells were distinguished by subsequent labeling with anti-vimentin antibody, alkaline phosphatase-labeled polymer, and permanent red substrate (EnVision G/2 system; DAKO). The stained cells were analyzed by light microscopy. Apoptotic cells in the cultures were quantified by counting the number of TUNEL-positive cells in five random microscope fields. 
Caspase-3 Activity
Caspase-3 activity was assayed using a cellular caspase-3 activity assay kit (Biomol, Plymouth Meeting, PA), according to the manufacturer's protocol. Briefly, cell extracts were added to the microtiter wells, and the reaction was initiated by adding 200 μM Ac-DEVD-pNA substrate. In parallel, the samples were reacted with this substrate in the presence of 0.1 μM Ac-DEVD-CHO, a specific caspase-3 inhibitor, to measure the nonspecific hydrolysis of the substrate. Absorbance was read at 405 nm in a microtiter plate reader at the indicated time intervals. 
Statistical Analysis
Various assay conditions were compared using ANOVA and t-tests (Stat-View; SAS Institute, Cary, NC), and P < 0.05 was considered statistically significant. Values represent mean ± SEM. 
Results
Expression and Activation of Caspase-4 in Response to Proinflammatory Stimulation
To determine the involvement of caspase-4 in proinflammatory response by hRPE cells, a group of known proinflammatory agents was selected for this study, including IL-1β, TNF-α, LPS, IFN-γ, and monocyte coculture. The concentrations of IL-1β (2 ng/mL), TNF-α (20 ng/mL), LPS (1000 ng/mL), and IFN-γ (500 U/mL) and the conditions for monocyte-hRPE coculture used in this study have been shown to maximally stimulate proinflammatory responses in hRPE cells. 2628,30 After hRPE cells were treated with these agents for 6 hours, total cellular mRNA was isolated and subjected to RT-PCR analysis. To compare caspase-4 mRNA levels, expression of the housekeeping gene β-actin was used to monitor gel loading. As shown in Figure 1A, treatment of hRPE cells with IL-1β, TNF-α, LPS, or monocyte coculture increased caspase-4 mRNA synthesis by 0.7-, 0.5-, 0.5-, or 0.6-fold, respectively. 
Figure 1.
 
Stimulation of human RPE caspase-4 mRNA synthesis (A) and protein activation (B) and blockade of IL-1β–induced IL-8 production by caspase-4 inhibitor Z-LEVD (C). hRPE cells were cultured either without (untreated; Ctl) or with IL-1β (IL-1; 2 ng/mL), TNF-α (TNF; 20 ng/mL), LPS (1000 ng/mL), IFN-γ (500 U/mL), or overlaid monocytes (RM) and were incubated for 6 hours (A) or 24 hours (B). The data shown represent results from a typical experiment. (A) Steady state caspase-4 mRNA determined by RT-PCR. The fold changes were calculated by normalization against β-actin and comparison with untreated control. (B) Western blot analysis of caspase-4 and actin proteins. The arrow-pointed bands are presumably either nonspecific bands or intermediately cleaved caspase-4. 10 (C) hRPE cells were pretreated with caspase-4 inhibitor Z-LEVD-fmk (2 μM) for 30 minutes and then coincubated with IL-1β for another 24 hours. Proteins from whole hRPE cell lysates were subjected to Western blot analysis by anti–IL-8 antibody.
Figure 1.
 
Stimulation of human RPE caspase-4 mRNA synthesis (A) and protein activation (B) and blockade of IL-1β–induced IL-8 production by caspase-4 inhibitor Z-LEVD (C). hRPE cells were cultured either without (untreated; Ctl) or with IL-1β (IL-1; 2 ng/mL), TNF-α (TNF; 20 ng/mL), LPS (1000 ng/mL), IFN-γ (500 U/mL), or overlaid monocytes (RM) and were incubated for 6 hours (A) or 24 hours (B). The data shown represent results from a typical experiment. (A) Steady state caspase-4 mRNA determined by RT-PCR. The fold changes were calculated by normalization against β-actin and comparison with untreated control. (B) Western blot analysis of caspase-4 and actin proteins. The arrow-pointed bands are presumably either nonspecific bands or intermediately cleaved caspase-4. 10 (C) hRPE cells were pretreated with caspase-4 inhibitor Z-LEVD-fmk (2 μM) for 30 minutes and then coincubated with IL-1β for another 24 hours. Proteins from whole hRPE cell lysates were subjected to Western blot analysis by anti–IL-8 antibody.
Next, whole lysates from hRPE cells treated under the same conditions, but for 24 hours, were subjected to Western blot analysis. Activation of caspase-4 protein was determined by the appearance of cleaved caspase-4 products (Fig. 1B). Data showed that IL-1β, TNF-α, LPS, IFN-γ, and monocyte coculture each activated caspase-4 protein. 
The activation of caspase-4 by proinflammatory agents suggested that caspase-4 may participate in proinflammatory responses in hRPE cells. To prove this is the case, IL-1β–induced IL-8 protein production was examined by Western blot analysis of the whole cell lysates from the hRPE cells treated with 2 ng/mL IL-1β in the presence or absence of caspase-4 inhibitor Z-LEVD-fmk. As we have shown previously, 27 IL-1β induced IL-8 protein production significantly above basal levels. As shown in Figure 1C, the presence of caspase-4 inhibitor completely eliminated the induced IL-8 protein production. 
Effect of Dexamethasone, IL-10, and Cyclosporine on IL-1β– and IFN-γ–Induced Activation of Caspase-4
Given that expression and activation of caspase-4 were induced by IL-1β and IFN-γ, we next investigated whether anti-inflammatory agents could neutralize this induced caspase-4 activation by these two proinflammatory agents. As expected, treatment with dexamethasone (1 μM) and IL-10 (100 U/mL) reduced IL-1β– and IFN-γ–induced cleavage of procaspase-4 by 60% and 33% and by 15% and 50%, respectively (Fig. 2). On the other hand, cyclosporine (3 ng/mL) inhibited IFN-γ–induced caspase-4 activation by 47% but markedly increased IL-1β–induced caspase-4 activation by more than twofold. 
Figure 2.
 
The effect of dexamethasone (Dex), cyclosporine (CsA), and IL-10 on caspase-4 activation by IL-1β (A, C) and IFN-γ (B, C) in hRPE cells. The hRPE cells were pretreated with Dex (1 μM), CsA (3 ng/mL), or IL-10 (100 U/mL) for 30 minutes and then coincubated with IL-1β (2 ng/mL) and IFN-γ (500 U/mL) for an additional 24 hours. Proteins from whole hRPE cell lysates were detected by anti–caspase-4 antibody specific for pro-caspase-4 and cleaved caspase-4. The fold changes of the cleaved caspase-4 were calculated by relative density between treated and untreated samples, as determined by densitometry after normalization with actin protein.
Figure 2.
 
The effect of dexamethasone (Dex), cyclosporine (CsA), and IL-10 on caspase-4 activation by IL-1β (A, C) and IFN-γ (B, C) in hRPE cells. The hRPE cells were pretreated with Dex (1 μM), CsA (3 ng/mL), or IL-10 (100 U/mL) for 30 minutes and then coincubated with IL-1β (2 ng/mL) and IFN-γ (500 U/mL) for an additional 24 hours. Proteins from whole hRPE cell lysates were detected by anti–caspase-4 antibody specific for pro-caspase-4 and cleaved caspase-4. The fold changes of the cleaved caspase-4 were calculated by relative density between treated and untreated samples, as determined by densitometry after normalization with actin protein.
Induction of Caspase-4 mRNA Synthesis and Protein Activation by ER Stress-Inducer Tunicamycin or Thapsigargin
Tunicamycin and thapsigargin are the two well-known ER stress inducers that act by blocking, respectively, N-glycosylation of newly synthesized proteins and Ca2+-ATPase, which maintains Ca2+ homeostasis in the ER. We used these two agents to study apoptotic involvement of casapse-4. Similar to the results from using proinflammatory agents, as described, tunicamycin (3 μM) and thapsigargin (25 ng/mL) both moderately increased caspase-4 mRNA by 0.3- and 0.4-fold, respectively (Fig. 3A). Tunicamycin (10 μM) induced caspase-4 cleavage that appeared as early as 24 hours after stimulation, and activation remained up to 72 hours (Fig. 3B). The induced caspase-4 activation was reduced by coincubation with caspase-4 inhibitor Z-LEVD-fmk (2 μM) and caspase-1 and -4 inhibitor Z-YVAD-fmk (2 and 20 μM) (Fig. 3C). 
Figure 3.
 
Stimulation of hRPE caspase-4 mRNA synthesis (A), protein activation (B, C), and caspase-3 activity (D) by tunicamycin (Tu) or thapsigargin (Tha). The hRPE cells were cultured either without (untreated; Ctl) or with tunicamycin (3 or 10 μM) or thapsigargin (25 ng/mL) for 6 (A), 24 (B, D), 48 (B, C), or 72 (B) hours. (C, D) In hRPE, cells were pretreated with or without caspase-4 inhibitor Z-LEVD and caspase-1 and -4 inhibitor Z-YVAD. (A) To determine the steady state levels of caspase-4 mRNA, total RNA was isolated and subjected to semiquantitative RT-PCR. The fold changes were expressed as ratios between treated and untreated samples after normalization by β-actin. (B, C) Caspase-4 protein production and activation. Western blot analysis of proteins from the whole cell lysates treated or untreated were detected by anti–caspase-4 antibody. (D) Caspase-3 activity was determined by cleavage of substrate Ac-DEVD-pNA, and the absorbance was read at 405 nm. Protein content of each sample was determined with a BCA assay. Values represent mean ± SEM; n = 4; ***P < 0.001.
Figure 3.
 
Stimulation of hRPE caspase-4 mRNA synthesis (A), protein activation (B, C), and caspase-3 activity (D) by tunicamycin (Tu) or thapsigargin (Tha). The hRPE cells were cultured either without (untreated; Ctl) or with tunicamycin (3 or 10 μM) or thapsigargin (25 ng/mL) for 6 (A), 24 (B, D), 48 (B, C), or 72 (B) hours. (C, D) In hRPE, cells were pretreated with or without caspase-4 inhibitor Z-LEVD and caspase-1 and -4 inhibitor Z-YVAD. (A) To determine the steady state levels of caspase-4 mRNA, total RNA was isolated and subjected to semiquantitative RT-PCR. The fold changes were expressed as ratios between treated and untreated samples after normalization by β-actin. (B, C) Caspase-4 protein production and activation. Western blot analysis of proteins from the whole cell lysates treated or untreated were detected by anti–caspase-4 antibody. (D) Caspase-3 activity was determined by cleavage of substrate Ac-DEVD-pNA, and the absorbance was read at 405 nm. Protein content of each sample was determined with a BCA assay. Values represent mean ± SEM; n = 4; ***P < 0.001.
Given that executioner caspase-3 is the downstream target of caspase-4, 10,23 we next assessed caspase-3 activity in the presence or absence of 2 μM caspase-4 inhibitor Z-LEVD-fmk. Caspase-3 activity was determined by caspase-3-specific cleavage of substrate Ac-DEVD-pNA after hRPE cells were treated with 3 μM tunicamycin. Consistent with previous reports, 10,23,31 caspase-4 inhibitor completely abolished the tunicamycin-induced activation of caspase-3 (Fig. 3D). 
ER Stress in Response to Apoptotic and Inflammatory Stimuli
Tunicamycin has been widely used to induce ER stress. To confirm the existence of ER stress when caspase-4 was activated by this agent, expression of GRP78, a specific marker of ER stress, was examined. In untreated hRPE cells, GRP78 protein was barely detectable by Western blot analysis (Fig. 4A). Induction of GRP78 production appeared at 24 hours after hRPE cells were treated with 10 μM tunicamycin. The induced GRP78 expression continued to increase up to 48 hours. 
Figure 4.
 
Time-dependent effects of tunicamycin (Tu) on GRP78 (A), Bcl-2, and Bax (B) protein expression. The hRPE cells were cultured either without or with tunicamycin (10 μM) for 24 and 48 hours. The hRPE whole cell lysates were subjected to Western blot analysis for GRP-78, Bcl-2, and Bax expression. The data shown represent results from a typical experiment. The fold changes were calculated by normalization of band density with actin and assigned control value as 1.
Figure 4.
 
Time-dependent effects of tunicamycin (Tu) on GRP78 (A), Bcl-2, and Bax (B) protein expression. The hRPE cells were cultured either without or with tunicamycin (10 μM) for 24 and 48 hours. The hRPE whole cell lysates were subjected to Western blot analysis for GRP-78, Bcl-2, and Bax expression. The data shown represent results from a typical experiment. The fold changes were calculated by normalization of band density with actin and assigned control value as 1.
Tunicamycin treatment also triggered a significant increase in expression of the antiapoptotic protein Bcl-2 as early as 24 hours after stimulation, and the increase was sustained up to 2.5-fold at 48 hours after treatment (Fig. 4B). On the other hand, tunicamycin modestly (by 0.7-fold) increased the proapoptotic protein Bax. As a result, the Bcl-2/Bax ratios were enhanced 1.1-fold by tunicamycin. 
Because GRP78 has been shown to have immunosuppressive activity, 31,32 we then examined the expression of GRP78 after treating hRPE cells with IL-1β, TNF-α, LPS, or overlaid monocytes. As shown in Figure 5A, these treatments increased hRPE GRP78 expression. In monocyte coculture, for example, the enhanced GRP78 protein levels appeared as early as 2 hours and reached steady state in 8 hours. 
Figure 5.
 
Expression of GRP78 protein in hRPE cells that were stimulated by proinflammatory agents (A) or in combination with anti-inflammatory agents (B). The hRPE cells were cultured either without (Ctl, control) or with IL-1β (IL-1, 2 ng/mL), TNF-α (TNF, 20 ng/mL), LPS (1000 ng/mL), IFN-γ (500 U/mL), or overlaid monocytes (RM) for 2 to 24 hours (A, middle and bottom) or 24 hours (A, top; B). (B, C) Cultures were pretreated with or without dexamethasone (Dex, 1 μM), cyclosporine (CsA, 3 or 30 ng/mL), or IL-10 (100 U/mL) for 30 minutes and then were coincubated with IL-1β and IFN-γ for an additional 24 hours. Whole hRPE cell lysates were subjected to Western blot analysis for GRP-78 expression. The fold changes of cleaved caspase-4 were determined by the ratios between treated and untreated samples of the band densities, which were quantified by densitometry and normalized by actin.
Figure 5.
 
Expression of GRP78 protein in hRPE cells that were stimulated by proinflammatory agents (A) or in combination with anti-inflammatory agents (B). The hRPE cells were cultured either without (Ctl, control) or with IL-1β (IL-1, 2 ng/mL), TNF-α (TNF, 20 ng/mL), LPS (1000 ng/mL), IFN-γ (500 U/mL), or overlaid monocytes (RM) for 2 to 24 hours (A, middle and bottom) or 24 hours (A, top; B). (B, C) Cultures were pretreated with or without dexamethasone (Dex, 1 μM), cyclosporine (CsA, 3 or 30 ng/mL), or IL-10 (100 U/mL) for 30 minutes and then were coincubated with IL-1β and IFN-γ for an additional 24 hours. Whole hRPE cell lysates were subjected to Western blot analysis for GRP-78 expression. The fold changes of cleaved caspase-4 were determined by the ratios between treated and untreated samples of the band densities, which were quantified by densitometry and normalized by actin.
The GRP78 expression induced by IL-1β was modestly inhibited by dexamethasone by 38% (Fig. 5B). In contrast, cyclosporine (3 ng/mL) and IL-10 markedly enhanced the induced GRP78 protein production by nearly 2-fold (Fig. 5B). IFN-γ, either alone or in combination with dexamethasone, cyclosporine (3 ng/mL), or IL-10, did not affect GRP78 expression. Of note, cyclosporine at 30 ng/mL appeared to be cytotoxic and caused strong ER stress. The latter was demonstrated by a 5-fold increase in GRP78 protein production in IL-1β treatment and a 3-fold increase in IFN-γ treatment. 
Involvement of Caspase-4 in Tunicamycin-Induced Apoptotic Cell Death
ELISA cell death detection kits were used to measure apoptotic cell death at the same concentrations used for caspase-4 activation. At 72 hours after treatment, tunicamycin induced substantial hRPE apoptotic cell death compared with untreated control cells, which had undetectable levels of cell death. Arbitrarily taking the ELISA readings under tunicamycin treatment as 100% cell death, the induced hRPE apoptosis was inhibited by caspase-4 inhibitor Z-LEVD (2 μM) and caspase-1 and -4 inhibitor Z-YVAD-fmk (20 μM) by 59% and 86%, respectively (Fig. 6). The pan-caspase inhibitor Z-VAD-fmk (50 μM) completely abolished the induced hRPE cell death. 
Figure 6.
 
ER stress-induced hRPE apoptotic cell death. (A) HRPE cells were cultured either without or with 10 μM tunicamycin in the presence or absence of caspase-4 inhibitor Z-LEVD, caspase-1 and -4 inhibitor Z-YVAD, or pan-caspase inhibitor Z-VAD for 72 hours. (B) The hRPE cells were cultured either without (Ctl, control) or with IL-1β (IL-1, 2 ng/mL), TNF-α (TNF, 20 ng/mL), LPS (1000 ng/mL), and tunicamycin (3 μM) for 24 hours (left) or with cyclosporine (CsA, 3 ng/mL), IL-10 (100 U/mL), dexamethasone (Dex, 1 μM), and tunicamycin (10 μM) for 72 hours (right). Apoptosis was determined by the absorbance difference between A405nm and A490nM using an ELISA cell death detection kit. For comparisons, the stress inducer-treated cells were arbitrarily assigned as 100% apoptotic cell death by tunicamycin. ***P<0.001; **P < 0.01; *P < 0.05 compared with tunicamycin treatment without inhibitors.
Figure 6.
 
ER stress-induced hRPE apoptotic cell death. (A) HRPE cells were cultured either without or with 10 μM tunicamycin in the presence or absence of caspase-4 inhibitor Z-LEVD, caspase-1 and -4 inhibitor Z-YVAD, or pan-caspase inhibitor Z-VAD for 72 hours. (B) The hRPE cells were cultured either without (Ctl, control) or with IL-1β (IL-1, 2 ng/mL), TNF-α (TNF, 20 ng/mL), LPS (1000 ng/mL), and tunicamycin (3 μM) for 24 hours (left) or with cyclosporine (CsA, 3 ng/mL), IL-10 (100 U/mL), dexamethasone (Dex, 1 μM), and tunicamycin (10 μM) for 72 hours (right). Apoptosis was determined by the absorbance difference between A405nm and A490nM using an ELISA cell death detection kit. For comparisons, the stress inducer-treated cells were arbitrarily assigned as 100% apoptotic cell death by tunicamycin. ***P<0.001; **P < 0.01; *P < 0.05 compared with tunicamycin treatment without inhibitors.
To validate the tunicamycin-induced apoptosis, TUNEL staining was performed. Normal hRPE cells in the culture exhibited typical hexagonal arrays (Fig. 7A, upper left). First we conducted microscopic analysis of the hRPE cells in the presence (Fig. 7A, lower panels) or absence (Fig. 7A[b], upper right) of 10 μM tunicamycin. As shown in Figure 7A, TUNEL staining is dark brown and hRPE vimentin staining is red. Nuclear condensation and cell shrinkage were evident after hRPE cells were treated with tunicamycin for 24 hours (Fig. 7A, lower panels). Next we determined the relative levels of the tunicamycin-induced apoptotic cell death by TUNEL assays in the presence of caspase-4 inhibitor Z-LEVD or pan-caspase inhibitor Z-VAD for 24 or 48 hours (Fig. 7B). After hRPE cells were treated with tunicamycin (10 μM) for 24 and 48 hours, 13% and 24% of the cells exhibited apoptotic cell death as detected by TUNEL assay. In the presence of the caspase-4 inhibitor Z-LEVD-fmk (2 μM), tunicamycin-induced apoptotic cell death at 24 and 48 hours was reduced by 62% and 53%, respectively. In contrast to Z-LEVD-fmk, pan-caspase inhibitor Z-VAD-fmk almost completely blocked the induced apoptotic cell death. Furthermore, treatment of hRPE cells with IL-1β (IL-1; 2 ng/mL), TNF-α (TNF; 20 ng/mL), and lipopolysaccharide (LPS; 1000 ng/mL) for 24 hours (Fig. 6B, left) or with cyclosporine (CsA; 3 ng/mL), IL-10 (100 U/mL), and dexamethasone (Dex; 1 μM) for 72 hours (Fig. 6B, right) did not result in apparent cell death compared with tunicamycin control. 
Figure 7.
 
Quantification of the effects of ER stress-induced hRPE cell death by TUNEL assays. HRPE cells were cultured either without or with 10 μM tunicamycin in the presence or absence of caspase-4 inhibitor Z-LEVD or pan-caspase inhibitor Z-VAD for 24 or 48 hours. (A) Dark TUNEL staining, 400×. The hRPE cells stained darkly for vimentin. Top: unstimulated hRPE cells (cultures and in TUNEL assay). Bottom: hRPE cells treated with tunicamycin showing nuclear condensation and cell shrinkage. (B) Data are expressed as percentage of TUNEL-positive hRPE cells. Values represent mean ± SEM. ***P < 0.001; **P < 0.01 compared with tunicamycin treatment without inhibitors.
Figure 7.
 
Quantification of the effects of ER stress-induced hRPE cell death by TUNEL assays. HRPE cells were cultured either without or with 10 μM tunicamycin in the presence or absence of caspase-4 inhibitor Z-LEVD or pan-caspase inhibitor Z-VAD for 24 or 48 hours. (A) Dark TUNEL staining, 400×. The hRPE cells stained darkly for vimentin. Top: unstimulated hRPE cells (cultures and in TUNEL assay). Bottom: hRPE cells treated with tunicamycin showing nuclear condensation and cell shrinkage. (B) Data are expressed as percentage of TUNEL-positive hRPE cells. Values represent mean ± SEM. ***P < 0.001; **P < 0.01 compared with tunicamycin treatment without inhibitors.
Discussion
Human caspase-12 was originally thought to play distinctive roles in inflammation and apoptosis, corresponding to those of caspase-12 in mice. However, given that most human populations express only the C-terminal truncated form of caspase-12, the role of this caspase in human apoptosis has been challenged. 2,33,34 The short form of human caspase-12 is not a null product in hRPE cells; we have recently shown that it may play an immunomodulatory role in these cells. 35 Phylogenetic analyses suggest that caspase-4 and -5 may be the functional counterparts of caspase-12 in humans. 4 The data from the present study support the notion that human caspase-4 may correspond to murine caspase-12, involved in both inflammation and apoptosis. 36  
Despite the accumulating evidence supporting an apoptotic role of caspase-4, negative results have been reported in tunicamycin- and thapsigargin-induced apoptosis in human multiple myeloma cell lines, 37 homoharringtonine-induced apoptosis in MUTZ-1 cells, 38 and Z α-1 antitrypsin-induced apoptosis in HEK293 cells. 39 Although apoptosis usually does not require de novo synthesis of caspases, ER stress may increase mRNA expression of certain caspases, including caspase-4. 40 Our study demonstrated that proinflammatory stimuli (IL-1β, TNF-α, LPS, IFN-γ, and monocyte coculture) induced both mRNA synthesis and activation of caspase-4 in hRPE cells. IL-1β–induced IL-8 protein production was reduced by the caspase-4 inhibitor Z-LEVD-fmk, suggesting that caspase-4 is involved in the proinflammatory responses of hRPE cells. These results are consistent with a previous study in which LPS-induced IL-8 mRNA synthesis and protein production are reduced by caspase-4 knockdown in human THP1 monocytic cell lines. 13 On the other hand, the ER-stress inducer tunicamycin induced expression of GRP78, a specific marker of ER stress. Although tunicamycin increased the expression of proapoptotic Bax, this study also found increased expression of the antiapoptotic Bcl-2 protein, resulting in an increased Bcl-2/Bax ratio. In most cases, the ER stress-induced apoptotic response is regulated by or dependent on the Bcl-2 family of proteins. 41 The Bcl-2/Bax ratio has been clinically used as an indicator of the susceptibility of tumor cells to the induction of apoptosis by chemotherapeutic agents. 42 However, the involvement of Bcl-2 and Bax in tunicamycin-induced apoptosis has not been well characterized. Increased Bcl-2 protein expression by tunicamycin has been reported in one previous report. 43 The increased Bcl-2 expression shown in our study, however, might not have affected caspase-4 activation because a previous study has shown that activation of caspase-4 by tunicamycin is only slightly affected by overexpression of Bcl-2. 9 The increased Bcl-2/Bax ratio by tunicamycin may suggest an early protective response of hRPE cells to apoptotic stimuli by enhanced expression of the antiapoptotic protein Bcl-2 to counteract the increase in proapoptotic protein Bax. 
Furthermore, in response to ER stress, hRPE cells increased the production and activation of caspase-4. The latter is consistent with a recent report in ARPE19 cells, an immortalized hRPE cell line. 40 These authors showed that treatment of ARPE19 cells with tunicamycin and thapsigargin stimulated the activation of caspase-4, though the effect of tunicamycin on caspase-4 mRNA expression was negligible. In agreement with this recent report, apoptotic hRPE cell death by staurosporine was insensitive to caspase-4 inhibitor; thus, caspase-4 is unlikely to be involved in the staurosporine-induced mitochondrial apoptotic pathway (data not shown). We further demonstrated in this study that tunicamycin-induced caspase-4 activation and apoptotic hRPE cell death were highly sensitive (59%) to inhibition by caspase-4 inhibitor Z-LEVD-fmk, implying that ER stress-induced apoptosis in hRPE occurs in part through the activation of caspase-4. The difference in potency between caspase-4 inhibitor Z-LEVD-fmk (59% inhibition) and pan-caspase inhibitor Z-VAD (98% inhibition) suggests the existence of parallel caspase-4-independent apoptotic pathways that are also induced by ER stress. Although apoptotic cell death can be caspase independent, 44 the hRPE apoptotic cell death induced by ER stress was totally caspase dependent because this induction was completely abolished by a pan-caspase inhibitor. The caspase inhibitor Z-YVAD-fmk has been used as a caspase-4 inhibitor, 45,46 but it is more frequently used as a caspase-1 inhibitor. 47 In fact, Z-YVAD-fmk inhibits both caspase-1 and -4. 48 The stronger inhibition (86%) of apoptosis by Z-YVAD-fmk suggests that caspase-1 is likely to be responsible for the ER stress-induced, caspase-4-independent hRPE cell death. Caspase-1 and -4 have been shown to coordinate in TNF-α–induced, but not in tunicamycin-induced, cell death in Cop-transfected HeLa cells. 11 Our data also show that caspase-4 inhibitor Z-LEVD-fmk abolished both tunicamycin-induced activation of caspase-3 and apoptotic hRPE cell death, suggesting that caspase-3, the central effector of apoptosis, acts downstream from caspase-4 in the ER stress-induced RPE apoptotic pathway, as reported in other cell types. 10,23,31,46,49 Taken together, our hRPE data support the proposal by Momoi et al. 36 that caspase-4 may be a functional surrogate of the truncated human caspase-12. 
The proinflammatory agents tested in this study, including IL-1β, TNF-α, and LPS, upregulated the ER stress marker GRP78. However, when treating hRPE cells with these three agents at the concentrations used for other experiments in this study for up to 24 hours, none of these agents caused apoptotic cell death. Upregulation of GRP78 in response to inflammation suggests its immunosuppressive and protective role. 31,32 Moreover, the drugs dexamethasone and cyclosporine and the anti-inflammatory cytokine IL-10, when added together with proinflammatory agent IL-1β, exerted differential consequences on the inductions of GRP78 expression. Although dexamethasone reversed IL1-β–induced increases in hRPE GRP78, cyclosporine at 3 and 30 ng/mL enhanced the already elevated GPR78 production caused by IL-1β. The different impact by dexamethasone and cyclosporine, as indicated by the ER stress marker GRP78 response, suggests potential risks of long-term treatment with some anti-inflammatory drugs, such as cyclosporine. Indeed, when we treated hRPE cells with cyclosporine (30 ng/mL) for 72 hours, all hRPE cells died. In contrast, neither dexamethasone (10 μM) nor IL-10 (100 U/mL) induced noticeable apoptosis under the same conditions. The cyclosporine concentration used in this study (3–30 ng/mL) was well within the drug concentrations used clinically (5–100 ng/mL), 50 and 30 ng/mL cyclosporine appeared to be cytotoxic in this study. Indeed, cyclosporine itself has been shown to induce ER stress and GRP78 causing nephrotoxicity and apoptotic cell death, 51 whereas dexamethasone has been shown effective in treating acquired glomerular diseases by suppressing ER stress and GRP78. 52  
The signaling pathway mediating caspase-4 proinflammatory responses remains elusive. Caspase-4 does not appear to act via caspase-1 activation. Activation of caspase-4 requires both dimerization and proteolysis, a feature that combines the requirement for activation of initiator caspases such as caspase-1 (dimerization) and effector caspases such as caspase-3 (interdomain cleavage). 53 Caspase-4 does not promote the maturation of caspase-1 substrate pro-IL-1β, 6 and no specific substrates have been identified for caspase-4. These observations suggest that the proinflammatory mechanisms involving caspase-4 differ from those involving caspase-1. 6,8 It has been proposed that caspase-4 may function mainly via the NF-κB signal pathway in inflammatory responses. 13 Our previous studies have shown that the NF-κB pathway is essential for the expression of IL-8 and other cytokines in hRPE cells, and the inhibition of NF-κB activation effectively blocks IL-1β–induced cytokine production in hRPE cells. 30 Because knockdown of the caspase-4 gene by siRNA significantly reduces NF-κB activation and nuclear translocation, 13 this mechanism for caspase-4 in the IL-8 pathway is likely. Caspase-4 has been associated with activation of signal transducer TRAF6. 13 In response to stimulation by proinflammatory cytokines, TRAF6 activates IκB kinase in the NF-κB pathway. 54 Thus, TRAF6 may also be involved with ER stress response-induced hRPE IL-8 expression. 
This study shows that caspase-4 appears to be a key mediator of apoptosis and inflammation in hRPE cells, underscoring its potential value as a novel therapeutic target. The pathophysiological relevance of this dual role in hRPE responses warrants further investigation. Inflammatory cytokines are essential mediators of the innate immune response. Given that hRPE cells play important roles in ocular functions under normal and diseased conditions, 27,55 caspase-4–mediated cytokine expression could be relevant to many noninfectious and infectious retinal diseases, such as proliferative vitreoretinopathy, 56 age-related macular degeneration, 57 uveitis, 58 and endophthalmitis. 59 On the other hand, apoptotic cell death is an established response in many ocular diseases, such as age-related macular degeneration, diabetic retinopathy, retinitis pigmentosa, retinal ischemia, photoreceptor degeneration, and glaucoma. 6063 Blockade of caspapse-3, the target downstream of caspase-4, may represent a therapeutic strategy in the protection of retinal degeneration. 64,65 Therefore, it is of interest to investigate whether caspase-4 is involved under those diseased conditions. Further delineating the signaling pathway, regulation, and functional roles of caspase-4 may suggest novel strategies for developing therapies for ocular disease. 
Footnotes
 Supported by National Institutes of Health Grants EY09441 and EY007003 and by a Research to Prevent Blindness–Olga Keith Weiss Award (VME).
Footnotes
 Disclosure: Z.-M. Bian, None; S.G. Elner, None; V.M. Elner, None
Footnotes
 The publication costs of this article were defrayed in part by page charge payment. This article must therefore be marked “advertisement” in accordance with 18 U.S.C. §1734 solely to indicate this fact.
References
Martinon F Tschopp J . Inflammatory caspases and inflammasomes: master switches of inflammation. Cell Death Differ. 2007;14:10–22. [CrossRef] [PubMed]
Scott AM Saleh M . The inflammatory caspases: guardians against infections and sepsis. Cell Death Differ. 2007;14:23–31. [CrossRef] [PubMed]
Cornelis S Kersse K Festjens N Lamkanfi M Vandenabeele P . Inflammatory caspases: targets for novel therapies. Curr Pharm Des. 2007;13:367–385. [CrossRef] [PubMed]
Lamkanfi M Kalai M Vandenabeele P . Caspase-12: an overview. Cell Death Differ. 2004;11:365–368. [CrossRef] [PubMed]
Eckhart L Ballaun C Hermann M . Identification of novel mammalian caspases reveals an important role of gene loss in shaping the human caspase repertoire. Mol Biol Evol. 2008;25:831–841. [CrossRef] [PubMed]
Kamens J Paskind M Hugunin M . Identification and characterization of ICH-2, a novel member of the interleukin-1 beta-converting enzyme family of cysteine proteases. J Biol Chem. 1995;270:15250–15256. [CrossRef] [PubMed]
Munday NA Vaillancourt JP Ali A . Molecular cloning and pro-apoptotic activity of ICErelII and ICErelIII, members of the ICE/CED-3 family of cysteine proteases. J Biol Chem. 1995;270:15870–15876. [CrossRef] [PubMed]
Faucheu C Blanchet AM Collard-Dutilleul V Lalanne JL Diu-Hercend A . Identification of a cysteine protease closely related to interleukin-1 beta-converting enzyme. Eur J Biochem. 1996;236:207–213. [CrossRef] [PubMed]
Hitomi J Katayama T Eguchi Y . Involvement of caspase-4 in endoplasmic reticulum stress-induced apoptosis and Abeta-induced cell death. J Cell Biol. 2004;165:347–356. [CrossRef] [PubMed]
Jiang CC Chen LH Gillespie S . Inhibition of MEK sensitizes human melanoma cells to endoplasmic reticulum stress-induced apoptosis. Cancer Res. 2007;67:9750–9761. [CrossRef] [PubMed]
Wang X Narayanan M Bruey JM . Protective role of Cop in Rip2/caspase-1/caspase-4-mediated HeLa cell death. Biochim Biophys Acta. 2006;1762:742–754. [CrossRef] [PubMed]
Hu Y Benedict MA Wu D Inohara N Núñez G . Bcl-XL interacts with Apaf-1 and inhibits Apaf-1-dependent caspase-9 activation. Proc Natl Acad Sci U S A. 1998;95:4386–4391. [CrossRef] [PubMed]
Lakshmanan U Porter AG . Caspase-4 interacts with TNF receptor-associated factor 6 and mediates lipopolysaccharide-induced NF-κB-dependent production of IL-8 and CC chemokine ligand 4 (macrophage-inflammatory protein-1). J Immunol. 2007;179:8480–8490. [CrossRef] [PubMed]
Oda T Kosuge Y Arakawa M Ishige K Ito Y . Distinct mechanism of cell death is responsible for tunicamycin-induced ER stress in SK-N-SH and SH-SY5Y cells. Neurosci Res. 2008;60:29–39. [CrossRef] [PubMed]
Pataer A Hu W Xiaolin L . Adenoviral endoplasmic reticulum-targeted mda-7/interleukin-24 vector enhances human cancer cell killing. Mol Cancer Ther. 2008;7:2528–2535. [CrossRef] [PubMed]
López-Antón N Rudy A Barth N . The marine product cephalostatin 1 activates an endoplasmic reticulum stress-specific and apoptosome-independent apoptotic signaling pathway. J Biol Chem. 2006;281:33078–33086. [CrossRef] [PubMed]
Chen LH Jiang CC Watts R . Inhibition of endoplasmic reticulum stress-induced apoptosis of melanoma cells by the ARC protein. Cancer Res. 2008;68:834–842. [CrossRef] [PubMed]
Nawrocki ST Carew JS Maclean KH . Myc regulates aggresome formation, the induction of Noxa, and apoptosis in response to the combination of bortezomib and SAHA. Blood. 2008;112:2917–2926. [CrossRef] [PubMed]
Rahmani M Davis EM Crabtree TR . The kinase inhibitor sorafenib induces cell death through a process involving induction of endoplasmic reticulum stress. Mol Cell Biol. 2007;27:5499–5513. [CrossRef] [PubMed]
Pyrko P Kardosh A Wang W Xiong W Schönthal AH Chen TC . HIV-1 protease inhibitors nelfinavir and atazanavir induce malignant glioma death by triggering endoplasmic reticulum stress. Cancer Res. 2007;67:10920–10928. [CrossRef] [PubMed]
Lin R Sun Y Li C Xie C Wang S . Identification of differentially expressed genes in human lymphoblastoid cells exposed to irradiation and suppression of radiation-induced apoptosis with antisense oligonucleotides against caspase-4. Oligonucleotides. 2007;17:314–326. [CrossRef] [PubMed]
Kamada S Washida M Hasegawa J Kusano H Funahashi Y Tsujimoto Y . Involvement of caspase-4 (-like) protease in Fas-mediated apoptotic pathway. Oncogene. 1997;15:285–290. [CrossRef] [PubMed]
Yukioka F Matsuzaki S Kawamoto K . Presenilin-1 mutation activates the signaling pathway of caspase-4 in endoplasmic reticulum stress-induced apoptosis. Neurochem Int. 2008;52:683–687. [CrossRef] [PubMed]
Nadiri A Wolinski MK Saleh M . The inflammatory caspases: key players in the host response to pathogenic invasion and sepsis. J Immunol. 2006;177:4239–4245. [CrossRef] [PubMed]
Elner SG Elner VM Pavilack MA . Modulation and function of intercellular adhesion molecule-1 (CD54) on human retinal pigment epithelial cells. Lab Invest. 1992;66:200–211. [PubMed]
Bian ZM Elner SG Yoshida A Elner VM . Human RPE-monocyte co-culture induces chemokine gene expression through activation of MAPK and NIK cascade. Exp Eye Res. 2003;76:573–583. [CrossRef] [PubMed]
Elner VM Strieter RM Elner SG Baggiolini M Lindley I Kunkel SL . Neutrophil chemotactic factor (IL-8) gene expression by cytokine-treated retinal pigment epithelial cells. Am J Pathol. 1990;136:745–750. [PubMed]
Elner VM Strieter RM Pavilack MA . Human corneal interleukin-8: IL-1 and TNF-induced gene expression and secretion. Am J Pathol. 1991;139:977–988. [PubMed]
Lin XY Choi MS Porter AG . Expression analysis of the human caspase-1 subfamily reveals specific regulation of the CASP5 gene by lipopolysaccharide and interferon-gamma. J Biol Chem. 2000;275:39920–39926. [CrossRef] [PubMed]
Bian ZM Elner SG Yoshida A Kunkel SL Su J Elner VM . Activation of p38, ERK1/2 and NIK pathways is required for IL-1beta and TNF-alpha-induced chemokine expression in human retinal pigment epithelial cells. Exp Eye Res. 2001;73:111–121. [CrossRef] [PubMed]
Wang M Wang P Liu YQ . The immunosuppressive and protective ability of glucose-regulated protein 78 for improvement of alloimmunity in beta cell transplantation. Clin Exp Immunol. 2007;150:546–552. [CrossRef] [PubMed]
Panayi GS Corrigall VM . BiP regulates autoimmune inflammation and tissue damage. Autoimmun Rev. 2006;5:140–142. [CrossRef] [PubMed]
Saleh M Vaillancourt JP Graham RK . Differential modulation of endotoxin responsiveness by human caspase-12 polymorphisms. Nature. 2004;429:75–79. [CrossRef] [PubMed]
Fischer H Koenig U Eckhart L Tschachler E . Human caspase 12 has acquired deleterious mutations. Biochem Biophys Res Commun. 2002;293:722–726. [CrossRef] [PubMed]
Bian ZM Elner SG Elner VM . Regulated expression of caspase-12 gene in human retinal pigment epithelial cells suggests its immunomodulating role. Invest Ophthalmol Vis Sci. 2008;49:5593–5601. [CrossRef] [PubMed]
Momoi T . Caspases involved in ER stress-mediated cell death. J Chem Neuroanat. 2004;28:101–105. [CrossRef] [PubMed]
Obeng EA Boise LH . Caspase-12 and caspase-4 are not required for caspase-dependent endoplasmic reticulum stress-induced apoptosis. J Biol Chem. 2005;280:29578–29587. [CrossRef] [PubMed]
Jie H Donghua H Xingkui X . Homoharringtonine-induced apoptosis of MDS cell line MUTZ-1 cells is mediated by the endoplasmic reticulum stress pathway. Leuk Lymphoma. 2007;48:964–977. [CrossRef] [PubMed]
Miller SD Greene CM McLean C . Tauroursodeoxycholic acid inhibits apoptosis induced by Z alpha-1 antitrypsin via inhibition of Bad. Hepatology. 2007;46:496–503. [CrossRef] [PubMed]
Koyama Y Matsuzaki S Gomi F . Induction of amyloid beta accumulation by ER calcium disruption and resultant upregulation of angiogenic factors in ARPE19 cells. Invest Ophthalmol Vis Sci. 2008;49:2376–2383. [CrossRef] [PubMed]
Hetz CA . ER stress signaling and the BCL-2 family of proteins: from adaptation to irreversible cellular damage. Antioxid Redox Signal. 2007;9:2345–2355. [CrossRef] [PubMed]
Cory S Huang DC Adams JM . The Bcl-2 family: roles in cell survival and oncogenesis. Nat Rev Cancer. 2002;2:647–656. [CrossRef] [PubMed]
Zhu L Xiang R Dong W Liu Y Qi Y . Anti-apoptotic activity of Bcl-2 is enhanced by its interaction with RTN3. Cell Biol Int. 2007;31:825–830. [CrossRef] [PubMed]
Bröker LE Kruyt FA Giaccone G . Cell death independent of caspases: a review. Clin Cancer Res. 2005;11:3155–3162. [CrossRef] [PubMed]
Liang B Song X Liu G . Involvement of TR3/Nur77 translocation to the endoplasmic reticulum in ER stress-induced apoptosis. Exp Cell Res. 2007;313:2833–2844. [CrossRef] [PubMed]
Li J Xia X Ke Y Nie H Smith MA Zhu X . Trichosanthin induced apoptosis in HL-60 cells via mitochondrial and endoplasmic reticulum stress signaling pathways. Biochim Biophys Acta. 2007;1770:1169–1180. [CrossRef] [PubMed]
Guo H Petrin D Zhang Y Bergeron C Goodyer CG LeBlanc AC . Caspase-1 activation of caspase-6 in human apoptotic neurons. Cell Death Differ. 2006;13:285–292. [CrossRef] [PubMed]
Perche O Doly M Ranchon-Cole I . Transient protective effect of caspase inhibitors in RCS rat. Invest Ophthalmol Vis Sci. 2008;86:519–527.
Kim SJ Zhang Z Hitomi E Lee YC Mukherjee AB . Endoplasmic reticulum stress-induced caspase-4 activation mediates apoptosis and neurodegeneration in INCL. Hum Mol Genet. 2006;15:1826–1834. [CrossRef] [PubMed]
Kurtz RM Elner VM Bian ZM Strieter RM Kunkel SL Elner SG . Dexamethasone and cyclosporin A modulation of human retinal pigment epithelial cell monocyte chemotactic protein-1 and interleukin-8. Invest Ophthalmol Vis Sci. 1997;38:436–445. [PubMed]
Pallet N Bouvier N Bendjallabah A . Cyclosporine-induced endoplasmic reticulum stress triggers tubular phenotypic changes and death. Am J Transplant. 2008;8:2283–2296. [CrossRef] [PubMed]
Fujii Y Khoshnoodi J Takenaka H . The effect of dexamethasone on defective nephrin transport caused by ER stress: a potential mechanism for the therapeutic action of glucocorticoids in the acquired glomerular diseases. Kidney Int. 2006;69:1350–1359. [PubMed]
Karki P Dahal GR Park IS . Both dimerization and interdomain processing are essential for caspase-4 activation. Biochem Biophys Res Commun. 2007;356:1056–1061. [CrossRef] [PubMed]
Wajant H Henkler F Scheurich P . The TNF-receptor-associated factor family: scaffold molecules for cytokine receptors, kinases and their regulators. Cell Signal. 2001;13:389–400. [CrossRef] [PubMed]
Hecquet C Lefevre G Valtink M Engelmann K Mascarelli F . Activation and role of MAP kinase-dependent pathways in retinal pigment epithelium cells: JNK1, P38 kinase, and cell death. Invest Ophthalmol Vis Sci. 2003;44:1320–1329. [CrossRef] [PubMed]
Charteris DG Hiscott P Grierson I Lightman SL . Proliferative vitreoretinopathy: lymphocytes in epiretinal membranes. Ophthalmology. 1992;99:1364–1367. [CrossRef] [PubMed]
Lopez PF Grossniklaus HE Lambert HM . Pathologic features of surgically excised subretinal neovascular membranes in age-related macular degeneration. Am J Ophthalmol. 1991;112:647–656. [CrossRef] [PubMed]
Koizumi K Poulaki V Doehmen S . Contribution of TNF-alpha to leukocyte adhesion, vascular leakage, and apoptotic cell death in endotoxin-induced uveitis in vivo. Invest Ophthalmol Vis Sci. 2003;44:2184–2191. [CrossRef] [PubMed]
Petropoulos IK Vantzou CV Lamari FN Karamanos NK Anastassiou ED Pharmakakis NM . Expression of TNF-alpha, IL-1beta, and IFN-gamma in Staphylococcus epidermidis slime-positive experimental endophthalmitis is closely related to clinical inflammatory scores. Graefes Arch Clin Exp Ophthalmol. 2006;244:1322–1328. [CrossRef] [PubMed]
Nickells RW Zack DJ . Apoptosis in ocular disease: a molecular overview. Ophthalmic Genet. 1996;17:145–165. [CrossRef] [PubMed]
Heathcote JG . Apoptosis and oncosis in ocular disease. Can J Ophthalmol. 1995;30:298–300. [PubMed]
Campochiaro PA . Potential applications for RNAi to probe pathogenesis and develop new treatments for ocular disorders. Gene Ther. 2006;13:559–562. [CrossRef] [PubMed]
Algvere PV Marshall J Seregard S . Acta Ophthalmol Scand. 2006;84:4–15. [CrossRef] [PubMed]
Bode C Wolfrum U . Caspase-3 inhibitor reduces apoptotic photoreceptor cell death during inherited retinal degeneration in tubby mice. Mol Vis. 2003;9:144–150. [PubMed]
Chen TA Yang F Cole GM Chan SO . Inhibition of caspase-3-like activity reduces glutamate induced cell death in adult rat retina. Brain Res. 2001;904:177–188. [CrossRef] [PubMed]
Figure 1.
 
Stimulation of human RPE caspase-4 mRNA synthesis (A) and protein activation (B) and blockade of IL-1β–induced IL-8 production by caspase-4 inhibitor Z-LEVD (C). hRPE cells were cultured either without (untreated; Ctl) or with IL-1β (IL-1; 2 ng/mL), TNF-α (TNF; 20 ng/mL), LPS (1000 ng/mL), IFN-γ (500 U/mL), or overlaid monocytes (RM) and were incubated for 6 hours (A) or 24 hours (B). The data shown represent results from a typical experiment. (A) Steady state caspase-4 mRNA determined by RT-PCR. The fold changes were calculated by normalization against β-actin and comparison with untreated control. (B) Western blot analysis of caspase-4 and actin proteins. The arrow-pointed bands are presumably either nonspecific bands or intermediately cleaved caspase-4. 10 (C) hRPE cells were pretreated with caspase-4 inhibitor Z-LEVD-fmk (2 μM) for 30 minutes and then coincubated with IL-1β for another 24 hours. Proteins from whole hRPE cell lysates were subjected to Western blot analysis by anti–IL-8 antibody.
Figure 1.
 
Stimulation of human RPE caspase-4 mRNA synthesis (A) and protein activation (B) and blockade of IL-1β–induced IL-8 production by caspase-4 inhibitor Z-LEVD (C). hRPE cells were cultured either without (untreated; Ctl) or with IL-1β (IL-1; 2 ng/mL), TNF-α (TNF; 20 ng/mL), LPS (1000 ng/mL), IFN-γ (500 U/mL), or overlaid monocytes (RM) and were incubated for 6 hours (A) or 24 hours (B). The data shown represent results from a typical experiment. (A) Steady state caspase-4 mRNA determined by RT-PCR. The fold changes were calculated by normalization against β-actin and comparison with untreated control. (B) Western blot analysis of caspase-4 and actin proteins. The arrow-pointed bands are presumably either nonspecific bands or intermediately cleaved caspase-4. 10 (C) hRPE cells were pretreated with caspase-4 inhibitor Z-LEVD-fmk (2 μM) for 30 minutes and then coincubated with IL-1β for another 24 hours. Proteins from whole hRPE cell lysates were subjected to Western blot analysis by anti–IL-8 antibody.
Figure 2.
 
The effect of dexamethasone (Dex), cyclosporine (CsA), and IL-10 on caspase-4 activation by IL-1β (A, C) and IFN-γ (B, C) in hRPE cells. The hRPE cells were pretreated with Dex (1 μM), CsA (3 ng/mL), or IL-10 (100 U/mL) for 30 minutes and then coincubated with IL-1β (2 ng/mL) and IFN-γ (500 U/mL) for an additional 24 hours. Proteins from whole hRPE cell lysates were detected by anti–caspase-4 antibody specific for pro-caspase-4 and cleaved caspase-4. The fold changes of the cleaved caspase-4 were calculated by relative density between treated and untreated samples, as determined by densitometry after normalization with actin protein.
Figure 2.
 
The effect of dexamethasone (Dex), cyclosporine (CsA), and IL-10 on caspase-4 activation by IL-1β (A, C) and IFN-γ (B, C) in hRPE cells. The hRPE cells were pretreated with Dex (1 μM), CsA (3 ng/mL), or IL-10 (100 U/mL) for 30 minutes and then coincubated with IL-1β (2 ng/mL) and IFN-γ (500 U/mL) for an additional 24 hours. Proteins from whole hRPE cell lysates were detected by anti–caspase-4 antibody specific for pro-caspase-4 and cleaved caspase-4. The fold changes of the cleaved caspase-4 were calculated by relative density between treated and untreated samples, as determined by densitometry after normalization with actin protein.
Figure 3.
 
Stimulation of hRPE caspase-4 mRNA synthesis (A), protein activation (B, C), and caspase-3 activity (D) by tunicamycin (Tu) or thapsigargin (Tha). The hRPE cells were cultured either without (untreated; Ctl) or with tunicamycin (3 or 10 μM) or thapsigargin (25 ng/mL) for 6 (A), 24 (B, D), 48 (B, C), or 72 (B) hours. (C, D) In hRPE, cells were pretreated with or without caspase-4 inhibitor Z-LEVD and caspase-1 and -4 inhibitor Z-YVAD. (A) To determine the steady state levels of caspase-4 mRNA, total RNA was isolated and subjected to semiquantitative RT-PCR. The fold changes were expressed as ratios between treated and untreated samples after normalization by β-actin. (B, C) Caspase-4 protein production and activation. Western blot analysis of proteins from the whole cell lysates treated or untreated were detected by anti–caspase-4 antibody. (D) Caspase-3 activity was determined by cleavage of substrate Ac-DEVD-pNA, and the absorbance was read at 405 nm. Protein content of each sample was determined with a BCA assay. Values represent mean ± SEM; n = 4; ***P < 0.001.
Figure 3.
 
Stimulation of hRPE caspase-4 mRNA synthesis (A), protein activation (B, C), and caspase-3 activity (D) by tunicamycin (Tu) or thapsigargin (Tha). The hRPE cells were cultured either without (untreated; Ctl) or with tunicamycin (3 or 10 μM) or thapsigargin (25 ng/mL) for 6 (A), 24 (B, D), 48 (B, C), or 72 (B) hours. (C, D) In hRPE, cells were pretreated with or without caspase-4 inhibitor Z-LEVD and caspase-1 and -4 inhibitor Z-YVAD. (A) To determine the steady state levels of caspase-4 mRNA, total RNA was isolated and subjected to semiquantitative RT-PCR. The fold changes were expressed as ratios between treated and untreated samples after normalization by β-actin. (B, C) Caspase-4 protein production and activation. Western blot analysis of proteins from the whole cell lysates treated or untreated were detected by anti–caspase-4 antibody. (D) Caspase-3 activity was determined by cleavage of substrate Ac-DEVD-pNA, and the absorbance was read at 405 nm. Protein content of each sample was determined with a BCA assay. Values represent mean ± SEM; n = 4; ***P < 0.001.
Figure 4.
 
Time-dependent effects of tunicamycin (Tu) on GRP78 (A), Bcl-2, and Bax (B) protein expression. The hRPE cells were cultured either without or with tunicamycin (10 μM) for 24 and 48 hours. The hRPE whole cell lysates were subjected to Western blot analysis for GRP-78, Bcl-2, and Bax expression. The data shown represent results from a typical experiment. The fold changes were calculated by normalization of band density with actin and assigned control value as 1.
Figure 4.
 
Time-dependent effects of tunicamycin (Tu) on GRP78 (A), Bcl-2, and Bax (B) protein expression. The hRPE cells were cultured either without or with tunicamycin (10 μM) for 24 and 48 hours. The hRPE whole cell lysates were subjected to Western blot analysis for GRP-78, Bcl-2, and Bax expression. The data shown represent results from a typical experiment. The fold changes were calculated by normalization of band density with actin and assigned control value as 1.
Figure 5.
 
Expression of GRP78 protein in hRPE cells that were stimulated by proinflammatory agents (A) or in combination with anti-inflammatory agents (B). The hRPE cells were cultured either without (Ctl, control) or with IL-1β (IL-1, 2 ng/mL), TNF-α (TNF, 20 ng/mL), LPS (1000 ng/mL), IFN-γ (500 U/mL), or overlaid monocytes (RM) for 2 to 24 hours (A, middle and bottom) or 24 hours (A, top; B). (B, C) Cultures were pretreated with or without dexamethasone (Dex, 1 μM), cyclosporine (CsA, 3 or 30 ng/mL), or IL-10 (100 U/mL) for 30 minutes and then were coincubated with IL-1β and IFN-γ for an additional 24 hours. Whole hRPE cell lysates were subjected to Western blot analysis for GRP-78 expression. The fold changes of cleaved caspase-4 were determined by the ratios between treated and untreated samples of the band densities, which were quantified by densitometry and normalized by actin.
Figure 5.
 
Expression of GRP78 protein in hRPE cells that were stimulated by proinflammatory agents (A) or in combination with anti-inflammatory agents (B). The hRPE cells were cultured either without (Ctl, control) or with IL-1β (IL-1, 2 ng/mL), TNF-α (TNF, 20 ng/mL), LPS (1000 ng/mL), IFN-γ (500 U/mL), or overlaid monocytes (RM) for 2 to 24 hours (A, middle and bottom) or 24 hours (A, top; B). (B, C) Cultures were pretreated with or without dexamethasone (Dex, 1 μM), cyclosporine (CsA, 3 or 30 ng/mL), or IL-10 (100 U/mL) for 30 minutes and then were coincubated with IL-1β and IFN-γ for an additional 24 hours. Whole hRPE cell lysates were subjected to Western blot analysis for GRP-78 expression. The fold changes of cleaved caspase-4 were determined by the ratios between treated and untreated samples of the band densities, which were quantified by densitometry and normalized by actin.
Figure 6.
 
ER stress-induced hRPE apoptotic cell death. (A) HRPE cells were cultured either without or with 10 μM tunicamycin in the presence or absence of caspase-4 inhibitor Z-LEVD, caspase-1 and -4 inhibitor Z-YVAD, or pan-caspase inhibitor Z-VAD for 72 hours. (B) The hRPE cells were cultured either without (Ctl, control) or with IL-1β (IL-1, 2 ng/mL), TNF-α (TNF, 20 ng/mL), LPS (1000 ng/mL), and tunicamycin (3 μM) for 24 hours (left) or with cyclosporine (CsA, 3 ng/mL), IL-10 (100 U/mL), dexamethasone (Dex, 1 μM), and tunicamycin (10 μM) for 72 hours (right). Apoptosis was determined by the absorbance difference between A405nm and A490nM using an ELISA cell death detection kit. For comparisons, the stress inducer-treated cells were arbitrarily assigned as 100% apoptotic cell death by tunicamycin. ***P<0.001; **P < 0.01; *P < 0.05 compared with tunicamycin treatment without inhibitors.
Figure 6.
 
ER stress-induced hRPE apoptotic cell death. (A) HRPE cells were cultured either without or with 10 μM tunicamycin in the presence or absence of caspase-4 inhibitor Z-LEVD, caspase-1 and -4 inhibitor Z-YVAD, or pan-caspase inhibitor Z-VAD for 72 hours. (B) The hRPE cells were cultured either without (Ctl, control) or with IL-1β (IL-1, 2 ng/mL), TNF-α (TNF, 20 ng/mL), LPS (1000 ng/mL), and tunicamycin (3 μM) for 24 hours (left) or with cyclosporine (CsA, 3 ng/mL), IL-10 (100 U/mL), dexamethasone (Dex, 1 μM), and tunicamycin (10 μM) for 72 hours (right). Apoptosis was determined by the absorbance difference between A405nm and A490nM using an ELISA cell death detection kit. For comparisons, the stress inducer-treated cells were arbitrarily assigned as 100% apoptotic cell death by tunicamycin. ***P<0.001; **P < 0.01; *P < 0.05 compared with tunicamycin treatment without inhibitors.
Figure 7.
 
Quantification of the effects of ER stress-induced hRPE cell death by TUNEL assays. HRPE cells were cultured either without or with 10 μM tunicamycin in the presence or absence of caspase-4 inhibitor Z-LEVD or pan-caspase inhibitor Z-VAD for 24 or 48 hours. (A) Dark TUNEL staining, 400×. The hRPE cells stained darkly for vimentin. Top: unstimulated hRPE cells (cultures and in TUNEL assay). Bottom: hRPE cells treated with tunicamycin showing nuclear condensation and cell shrinkage. (B) Data are expressed as percentage of TUNEL-positive hRPE cells. Values represent mean ± SEM. ***P < 0.001; **P < 0.01 compared with tunicamycin treatment without inhibitors.
Figure 7.
 
Quantification of the effects of ER stress-induced hRPE cell death by TUNEL assays. HRPE cells were cultured either without or with 10 μM tunicamycin in the presence or absence of caspase-4 inhibitor Z-LEVD or pan-caspase inhibitor Z-VAD for 24 or 48 hours. (A) Dark TUNEL staining, 400×. The hRPE cells stained darkly for vimentin. Top: unstimulated hRPE cells (cultures and in TUNEL assay). Bottom: hRPE cells treated with tunicamycin showing nuclear condensation and cell shrinkage. (B) Data are expressed as percentage of TUNEL-positive hRPE cells. Values represent mean ± SEM. ***P < 0.001; **P < 0.01 compared with tunicamycin treatment without inhibitors.
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