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(Investigative Ophthalmology and Visual Science. 2001;42:1356-1362.)
© 2001 by The Association for Research in Vision and Ophthalmology, Inc.

Blue Light–Induced Apoptosis of A2E-Containing RPE: Involvement of Caspase-3 and Protection by Bcl-2

Janet R. Sparrow and Bolin Cai

From the Department of Ophthalmology, Columbia University, New York.


   Abstract
Top
 Abstract
Introduction
 Methods
 Results
 Discussion
 References
 
PURPOSE. The lipofuscin fluorophore A2E has been shown to mediate blue light–induced damage to retinal pigment epithelial (RPE) cells. The purpose of this study was to evaluate caspase-3 and Bcl-2 as executor and modulator, respectively, of the cell death program that is initiated in A2E-containing cells in response to blue light.

METHODS. Human RPE cells (ARPE-19) that had accumulated A2E were exposed to blue light. Caspase-3 activity was assayed by observing cleavage of a fluorogenic peptide substrate, and the effect of a peptide inhibitor of caspase-3 (Z-DEVD-fmk) on the quantity of apoptotic nuclei was determined. ARPE-19 cells were transfected with either a neomycin-selectable expression vector containing Bcl-2 cDNA or a control neomycin-selectable expression vector without Bcl-2 cDNA. Expression of Bcl-2 transcripts by independently derived clones was established by in situ hybridization, and Bcl-2 protein expression was confirmed by Western blot analysis. Cell viability was assayed by TdT-dUTP terminal nick-end labeling (TUNEL) in conjunction with 4'6'-diamidino-2-phenylindole (DAPI) staining and by fluorescence staining of the nuclei of membrane-compromised cells.

RESULTS. In RPE cells that had previously accumulated A2E, caspase-3 activity was detected within 5 hours of blue light exposure. The incidence of apoptotic nuclei was attenuated when A2E-containing RPE cells were exposed to blue light in the presence of caspase-3 inhibitor and in A2E-loaded RPE cells that had been stably transfected with Bcl-2.

CONCLUSIONS. Blue light illumination of RPE in the setting of intracellular A2E initiates a cell death program that is executed by a proteolytic caspase cascade and that is regulated by Bcl-2.


   Introduction
Top
 Abstract
 Introduction
Methods
 Results
 Discussion
 References
 
The death of retinal pigmented epithelial (RPE) cells in a number of retinal disorders, including Stargardt’s disease, Best’s disease, some forms of retinitis pigmentosa, and nonexudative age-related macular degeneration (AMD) with geographic atrophy, is a crucial event in the disease process. Areas of RPE cell atrophy in these disorders are readily visualized by in vivo laser scanning ophthalmoscopy or fundus spectrophotometry as regions of decreased fundus autofluorescence with the margins of the atrophic areas exhibiting pronounced fluorescence.1 2 3 4 5 It is generally accepted that fundus autofluorescence is attributable to the lipofuscin3 5 6 that accumulates in RPE cells with age and that is also amassed in excessive amounts in a number of inherited retinal disorders.7 8 9 10 11 12

Analysis of extracts of human RPE has revealed that the major hydrophobic fluorophore of RPE lipofuscin is A2E,13 14 a quaternary pyridinium salt14 15 16 that is generated after hydrolytic cleavage of the all-trans-retinal-phosphatidylethanolamine conjugate, A2PE (phosphatidyl-pyridinium bisretinoid).17 It is also now clear that the accumulation of A2E has adverse consequences for the cell. Thus, as an amphiphilic detergent, A2E has been shown to exert a detergent-like perturbation of cell membranes,18 an effect that may explain the propensity for A2E to interfere with the adenosine triphosphatase (ATPase)–dependent acidification of lysosomes.19 A2E has also been shown to confer a susceptibility to photo-induced damage.20 21 In particular, the blue (480 nm) region of the spectrum was found to induce the death of A2E-containing cultured RPE cells in a manner that was directly dependent on the A2E content of the cells.20 Conversely, green light (540 nm) was considerably less effective. This wavelength dependence was consistent with the absorbance and excitation spectra of A2E.20

Although the photochemical events triggering apoptosis under conditions of blue light exposure are not fully understood, the cell death program is probably executed by caspases, a family of cysteine-dependent proteases located in the cytoplasm.22 23 24 25 26 27 Several lines of evidence indicate that the caspase-mediated cleavage of manifold cellular substrates, including enzymes involved in DNA repair, structural components of the cytoplasm and nucleus, and various protein kinases, is directly responsible for the demise of the cell. Caspases are synthesized as inactive zymogens (procaspases) whose activation requires cleavage on the carboxyl side of aspartate residues to liberate one large (~20 kDa) and one small (~10 kDa) subunit. The active enzyme is then formed as a tetramer consisting of two of each of these subunits. Caspases cleave substrate proteins exclusively after aspartate residues, and the sequence of the four amino acid NH2-terminals to the cleavage site determines the substrate specificity of the different caspases. Distinct members of the caspase family are involved in both the initiation and execution phases of apoptosis, with the initiator caspases coupling cellular signaling pathways to caspase activation and the downstream effector caspases being responsible for the cleavage of cellular substrates.

Although several human caspases have been identified, it is becoming increasingly clear that not only does the specific subset of caspases recruited and the sequence in which they are activated vary with the particular cell death paradigm, but the cascade may also exhibit cell-type specificity.24 28 29 30 31 Furthermore, the inhibition of caspases does not always prevent cell death elicited by proapoptotic signals.32 Perhaps the best studied of the cell death pathways are those that are triggered by binding of cognate ligand to one of a number of cell surface death receptors. Subsequent clustering of the receptor leads to physical association with an adaptor protein at the cytoplasmic face and ultimately to the clustering and activation of initiator caspases. Conversely, other cell death pathways are initiated by mitochondria, with cytochrome c and probably other proteins being released from the mitochondrial intermembrane space. On entering the cytosol, cytochrome c forms a complex with an adaptor protein (APAF-2), thereby recruiting and activating the initiator caspase. Upstream of this process, an additional level of regulation is provided by the Bcl-2 family of proteins, many but not all of which reside in the mitochondrial outer membrane and among other actions, control the release of mitochondrial apoptogenic factors, such as cytochrome c.30 33 34 35 36 37 Nevertheless, because one antiapoptotic member of this family, Bcl-2 protein, can inhibit some apoptotic paradigms but not others, it is possible that certain death stimuli can either circumvent or operate downstream of Bcl-2.

Although a number of exogenous photosensitizers—for instance, those used in photodynamic therapy38 39 —have been studied for their ability to initiate apoptosis after their activation by specific wavelengths of light, far less is understood of the mechanism by which an identified, naturally occurring fluorophore, such as A2E, induces apoptosis after exposure to visible light. By studying caspase-3 as an effector of the death process and Bcl-2 as a potential negative regulator, we addressed the molecular pathways involved in executing RPE cell death in the context of A2E and blue light.


   Methods
Top
 Abstract
 Introduction
 Methods
Results
 Discussion
 References
 
RPE Cultures
A human adult RPE cell line ARPE-19 (American Type Culture Collection, Manassas, VA), which is devoid of endogenous A2E,18 was grown as previously described,18 and all experiments were performed at confluence.

A2E Synthesis and Loading
A2E was synthesized from all-trans-retinal and ethanolamine14 and stored as a stock solution in dimethyl sulfoxide (DMSO). For loading of RPE cells, A2E was delivered in 100-µM concentrations in culture media, as previously described.18 The autofluorescence of cell-associated A2E was detected by epifluorescent illumination under a microscope (Axiovert S100; Carl Zeiss, Thornwood, NY) and standard fluorescein isothiocyanate (FITC) filters (460–500-nm excitation, 510–560-nm emission).

Blue Light Illumination
Cells growing in eight-well plastic chamber slides were exposed, either to a single spot of blue light delivered from a 100-W mercury lamp (480 ± 20 nm; 35 mW/mm2, 60 seconds)20 or to a light line delivered from a tungsten halogen source (470 ± 20 nm; 0.4 mW/mm2; 20-minute exposure), as indicated. These wavelengths are consistent with the excitation spectrum of A2E.20 .

Caspase-3 Cleavage Activity
Caspase-3–like protease activity was studied by detecting the cleavage of the cell-permeable fluorogenic peptide substrate GDEVDGI (Gly-Asp-Glu-Val-Asp-Gly-Ile; PhiPhiLux-G2D2; Alexis, Laufelfingen, Switzerland).40 41 42 43 Briefly, 5 hours after light exposure, the cells were incubated for 1 hour at 37°C in the dark with 10 µM substrate prepared in RPMI-1640 supplemented with 10% fetal calf serum (FCS). Immediately after washing, the cleaved substrate was detected by fluorescence microscopy using rhodamine-appropriate filters.

Inhibition of Caspase-3
Before exposure to blue light, A2E containing ARPE-19 cells was preincubated at 37°C for 1 hour with 20 µM Z-Asp-Glu-Val-Asp-fluoromethylketone (Z-DEVD-fmk; Alexis), a cell-permeable peptide inhibitor whose tetrapeptide structure is based on the optimal sequence recognized by caspase-3.44

Transfection of Bcl-2
ARPE-19 cells grown in 100-mm culture dishes to 70% to 80% confluence were transfected with the neomycin-resistant pSFFV/Bcl-2 plasmid (a generous gift from Ralph Buttyan, Columbia University, the College of Physicians and Surgeons, New York, NY).45 Control cells were transfected with the neomycin-resistant expression vector (pCMV-Script; Stratagene, La Jolla, CA) without insertion of the Bcl-2 cDNA. Transfection complexes were prepared by preincubation of plasmid (4 µg) with reagents (Plus-Lipofectamine; Gibco–Life Technologies, Grand Island, NY) in serum-free/antibiotic-free Dulbecco’s modified Eagle’s medium (DMEM; 0.75 ml), according to the manufacturer’s instructions. Subsequently, 1.5 ml of the reagent complex was gently mixed with the 5 ml of DMEM in each culture plate. After 3 hours of incubation at 37°C and 8.5% CO2, 6.5 ml of antibiotic-free medium containing 20% FCS was added to each dish. Twenty-four hours after the start of transfection, the cells were replated in DMEM with 10% FCS at a density of 105 cells/100-mm dish, and after an additional 24 hours, G418 sulfate (700 µg/ml; Gibco–Life Technologies) was added to begin selection. Medium containing G418 was renewed weekly, and after 3 weeks, individual colonies were isolated with cloning rings and were subcultured and eventually expanded. Seventeen pSFFV Bcl-2 and eight pCVM-Script-neo clones were screened for Bcl-2 expression.

Probe Generation and In Situ Hybridization
To generate RNA probes for Bcl-2 in situ hybridization, a fragment of a 630-bp Bcl-2 cDNA was subcloned into the pBluescript II KS(±) vector. BssHII/EcoRI (antisense) and BssHII/BamHI (sense) linearized DNA templates were purified by electrophoresis on agarose gel and transcribed using the digoxigenin RNA labeling system (Boehringer–Mannheim, Indianapolis, IN). Forward transcription from the T7 promoter generated the antisense probe. Before hybridization, probe size was determined by electrophoresis on a 1% agarose gel, and labeling efficiency was detected by dot blot hybridization.

Cells grown in eight-well chambers were fixed with 4% paraformaldehyde for 20 minutes, washed in phosphate-buffered saline (PBS), and digested with 7.5 µg/ml proteinase K in 50 mM EDTA and 0.1 M Tris-HCl (pH 8.0) for 20 minutes at 37°C. After rinsing in 0.2% glycine to arrest digestion, the sections were acetylated in 0.25% acetic anhydride containing 0.1 M triethanolamine for 10 minutes. The slides were prehybridized for 2 hours at 37°C in a solution containing 50% formamide, 2x SSC, 1x Denhardt’s solution, 10% dextran sulfate, 0.1% sodium dodecyl sulfate (SDS), 4 mM EDTA, 250 µg/ml yeast t-RNA, and denatured salmon testis DNA. Hybridization of Bcl-2 antisense digoxigenin UTP-labeled RNA probe (2.5 ng/µl) was then performed overnight at 42°C. Sense probe served as a negative control. RNase A was subsequently applied to the sections for 30 minutes to digest any unbound probe. The slides were then washed repeatedly with gentle agitation in declining concentrations of SSC (2x to 0.1x SSC) followed by digoxigenin buffer (0.1 M Tris-HCl [pH 7.5], 0.15 M NaCl). After blocking with 10% normal serum, the sections were incubated for 2 hours with alkaline phosphatase–conjugated anti-digoxigenin polyclonal sera (Boehringer–Mannheim), diluted 1:750 in 100 mM Tris-HCl (pH 7.5) containing 150 mM NaCl and 10% normal serum. The bound antibody was detected using nitroblue tetrazolium chloride/5-bromo-4-chloro-3-indolyl-phosphate as substrate (Boehringer–Mannheim).

Detection of Bcl-2 by Western Blot Analysis
Cells were washed with PBS and lysed in 50 mM Tris-HCl (pH 7.5) containing 150 mM NaCl, 1% Nonidet P40, 0.5% sodium deoxycholate, and protease inhibitors. Lysates were centrifuged at 12,000g for 10 minutes, and the protein concentration of the supernatant was determined using a protein assay system (Bio-Rad, Richmond, CA). Bcl-2 proteins were immunoprecipitated using monoclonal antibody to human Bcl-2 (Dako, Glostrup, Denmark), and samples containing equal amounts of immunoprecipitated protein (20 µg) were separated by 10% sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to nitrocellulose membranes. Bcl-2 protein was detected using the antibody to human Bcl-2, and binding of secondary antibody was detected using the blot detection system (Hybond ECL; Amersham Pharmacia Biotech, Piscataway, NJ).

Assays of Cell Viability
To count nonviable cells using assays based on the detection of nuclear condensation and DNA fragmentation, terminal deoxynucleotidyl transferase (TdT)-mediated dUTP nick-end labeling (TUNEL) of apoptotic nuclei was performed, together with 4'6'-diamidino-2-phenylindole (DAPI) labeling of all nuclei, 6 hours after blue light exposure. For staining, cultures were fixed in 2% paraformaldehyde for 30 minutes, permeabilized with 0.1% Triton X-100 in 0.1% sodium citrate (2 minutes, 4°C), incubated in TdT together with dUTP-rhodamine (37°C, 60 minutes; Boehringer–Mannheim), and stained with DAPI (0.3 µM). To determine the percentage of apoptotic nuclei, TUNEL- (rhodamine) and DAPI-stained nuclei were visualized by fluorescence microscopy (x40 objective), and counting was performed from digital images after contrast enhancement by computer (PhotoShop ver. 5; Adobe, San Jose, CA).

Nonviable cells were also labeled by fluorescence-exclusion assays that allow for the labeling of apoptotic nuclei because of a loss of plasma membrane integrity during the latter stages of apoptosis. Accordingly, 12 and 18 hours after blue light exposure, the nuclei of dead cells were stained with the membrane-impermeable dyes propidium iodide (15 µM in medium, 15-minute incubation; Molecular Probes, Eugene, OR) or Dead Red (1:500 dilution, 15-minute incubation; Molecular Probes) alone or in combination with Hoechst 33342 (5 µg/ml) to stain all nuclei. In all experiments, replicates were assayed as indicated in the figure legends.


   Results
Top
 Abstract
 Introduction
 Methods
 Results
Discussion
 References
 
Blue Light–Induced Apoptosis of A2E-Containing RPE and Caspase-3 Activation
To determine whether caspase-3 is activated after blue light irradiation of A2E-containing RPE cells, we used a cell-permeable rhodamine-conjugated caspase-3 substrate that exhibits red fluorescence only after enzyme cleavage. Accordingly, fluorescence microscopic examination 5 hours after illumination revealed that A2E-containing cells exposed to blue light (spot illumination) for 60 seconds exhibited red fluorescence, a finding indicative of proteolytic cleavage of the fluorogenic substrate by activated caspase-3 (Figs. 1A , 2 A ). The absence of red fluorescence in A2E-loaded cultures that were not illuminated before incubation with the caspase-3–specific substrate, demonstrated that the red fluorescence was not attributable to cross-contamination of signal (Fig. 1B) . To confirm that the cleavage of the fluorogenic substrate PhiPhilux-G6D2 was caspase-3 dependent, the cells were also treated with the caspase-3 inhibitor Z-DEVD-fmk before light exposure. Suppression of the fluorescence emission of the substrate PhiPhilux-G6D2 occurred (Figs. 1C 2D) .



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  		Figure 1. Activation of caspase-3 in ARPE-19 cells that had accumulated A2E and were exposed to 480-nm light. (A) Rhodamine fluorescence, indicative of caspase-3–mediated substrate cleavage, was exhibited by A2E-loaded RPE cells illuminated with 480-nm light and incubated with the fluorogenic substrate PhiPhilux-G6D2. (B) Rhodamine fluorescence was not detected in A2E-loaded RPE cells that were incubated with the fluorogenic caspase-3–specific substrate but not exposed to 480-nm light. (C) Treatment of A2E-loaded RPE cells with the caspase-3 inhibitor Z-DEVD-fmk before 480-nm illumination suppressed the fluorescence emission of the substrate PhiPhilux-G6D2. Representative of three experiments. Scale bar, 30 µM.

 


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  		Figure 2. Blue light–induced caspase-3 activity colocalized with intracellular A2E autofluorescence. (A) Epifluorescence detection of the autofluorescent A2E that accumulated in cultures of ARPE-19 cells. (B) Caspase-3 activity was visualized in A2E-containing cells after exposure to 480-nm light and incubation with the fluorogenic caspase substrate PhiPhilux-G6D2. Same field as in (A). Several of the cells that were heavily loaded with A2E (A) simultaneously exhibited caspase-3 activity (B; arrows). (C) Phase-contrast micrograph illustrates confluence of the field of cells shown in (A) and (B). (D) Caspase-3 activity is not detected in unexposed A2E-containing cells after incubation with the fluorogenic caspase substrate. Representative of three experiments. Scale bar, 15 µm.

 
When we examined the autofluorescence of A2E-accumulating cells by epifluorescence microscopy, together with the corresponding phase-contrast images indicating the confluence of the cultures (Figs. 2A 2C) , it was apparent that the RPE cells varied in the amounts of A2E that they had accumulated. Moreover, comparison of the A2E autofluorescence with the fluorescence emission of the caspase-3 substrate (Figs. 2A 2B) revealed that all the cells that exhibited caspase-3 activity had accumulated readily visible levels of A2E. This observation is consistent with the concept that blue light toxicity is dependent on the cells’ accumulating critical concentrations of A2E.

The role of caspase-3 in mediating the blue light–induced death of A2E-containing RPE cells was further assessed by exposing the cells to a spot of 480 nm illumination (60 seconds) in the presence of the caspase-3 inhibitor Z-DEVD-fmk. Using a fluorescence assay in which the nuclei of nonviable cells were labeled with the membrane-impermeable dye Dead Red 18 hours after light exposure, we observed that treatment with Z-DEVD-fmk reduced the numbers of nonviable cells in the 0.5-mm diameter zones corresponding to the areas of illumination (Figs. 3A 3B ). Counting of fluorescently labeled nuclei in the illuminated fields revealed that the addition of Z-DEVD-fmk decreased the numbers of apoptotic nuclei to an average of 55% of control numbers (three experiments with two-tailed P = 0.0006, 0.02, and 0.05, by Student’s t-test; Fig. 3C ). As previously reported,20 the frequency of apoptotic nuclei among cells that had not been loaded with A2E but that were exposed to blue light was not greater than background levels observed in nonilluminated regions of the cultures (not shown).



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  		Figure 3. The death of A2E-loaded RPE cells that are exposed to 480 nm light is inhibited in the presence of the cell-permeable caspase-3 inhibitor Z-DEVD-fmk. (A) ARPE-19 cells accumulated A2E in culture 7 days before 480-nm illumination for 60 seconds. The nuclei of nonviable cells were labeled by a membrane-impermeable dye, 18 hours after exposure. The zone of nonviable cells (0.5 mm in diameter) corresponded to the area of illumination. (B) A2E-containing RPE cells were incubated with Z-DEVD-fmk 1 hour before 480-nm exposure, and 18 hours after exposure nuclei of nonviable cells were labeled. Scale bar, 80 µm. (C) Quantification of the effect of the caspase-3 inhibitor Z-DEVD-fmk on the numbers of nonviable nuclei located in zones of illumination after blue light exposure of A2E-loaded RPE cells. Values are the mean ± SEM of three experiments, 3 to 10 replicates per experiment.

 
Blue Light-Induced Apoptosis of A2E-Containing RPE: Protection by Bcl-2
To determine whether enhanced Bcl-2 expression could alter the response of A2E-loaded RPE to blue light, human ARPE-19 cells were transfected with either a neomycin-selectable expression plasmid containing cDNA for human Bcl-2 (pSFFV/Bcl-2) or a control neomycin-selectable expression vector containing no cDNA for Bcl-2 (pCMV-Script). Seventeen clones independently derived from ARPE-19 cells transfected with pSFFV/Bcl-2 and eight clones transfected with the pCMV-Script-neo vector (control transfectants) were screened by Western blot analysis for Bcl-2 protein expression. Of the Bcl-2 transfectants, six demonstrated greatly enhanced expression of Bcl-2 protein when compared with the parental ARPE-19 cell line and the pCMV-Script-neo control-transfected lines. The immunoblot analysis of two of these clones is presented in Figure 4 . As evidenced by hybridization of digoxigenin-labeled Bcl-2 RNA probe (antisense), these Bcl-2–transfected clones also exhibited enhanced expression of Bcl-2 transcripts (Fig. 5A ). Labeled sense probe and control transfectants hybridized with Bcl-2 antisense probe served to control for nonspecific hybridization (Figs. 5B 5C) . Two Bcl-2–transfected clones, Bcl-2/1 and Bcl-2/6, together with a control-transfected clone were selected for further study and were grown continuously in medium containing G-418.



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  		Figure 4. Immunoblot analysis of Bcl-2 expression in wild-type (WT) nontransfected ARPE-19 cells, the same cell line transfected with the pCMV control vector alone (control transfected) and two independently derived clones transfected with pSFFV/Bcl-2 (Bcl-2/1, Bcl-2/6). Each lane was loaded with 20 µg of protein after immunoprecipitation. The blot was probed with monoclonal antibody to human Bcl-2.

 


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  		Figure 5. Detection of Bcl-2 mRNA in Bcl-2–transfected ARPE-19 cells by in situ hybridization. Bcl-2–transfected cells were hybridized with digoxigenin-labeled Bcl-2 antisense (A) and sense (B) probe. Control-transfected cells were hybridized with Bcl-2 antisense probe (C). Scale bar, 20 µM.

 
Clonal derivatives of ARPE-19 cells stably transfected with Bcl-2 (Bcl-2/1 and Bcl-2/6) and subsequently loaded with A2E demonstrated a resistance to blue light damage. Thus, when the viability of A2E-containing cells was tested after blue light irradiation by labeling all nuclei with DAPI and apoptotic nuclei by TUNEL, the Bcl-2–transfected lines displayed a 50% to 60% reduction in the proportion of apoptotic nuclei compared with control-transfected cells (P < 0.05; Fig. 6A ). Combined propidium iodide and Hoechst 33342 staining 12 hours after blue light exposure revealed a similar decrease in the percentage of apoptotic nuclei (Fig. 6B) . Moreover, the incidence of apoptotic nuclei in a Bcl-2-overexpressing line (Bcl-2/6) was decreased 58% and 52% compared with control-transfected and wild-type cells, respectively, when cell death was assayed by the exclusion of propidium iodide at 18 hours after blue light exposure (Fig. 6C) .



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  		Figure 6. Overexpression of Bcl-2 protected A2E-loaded RPE cells from death after blue light exposure. Two independently derived Bcl-2–transfected clones (Bcl-2/1, Bcl-2/6), a control-transfected clone and wild-type (WT) cells accumulated A2E 7 days before blue light exposure (470 nm; band illumination). (A) The percentage of apoptotic nuclei was determined by labeling all nuclei with DAPI and apoptotic nuclei by the TUNEL (rhodamine) method 6 hours after blue light illumination. Replicates were assayed by counting cells within 10 microscopic fields (x40 objective) situated along the band of illumination in each culture. Data are the means ± SEM of three experiments. Bcl-2/1- and Bcl-2/2-transfected cells were significantly different from control-transfected (P < 0.05; one-way analysis of variance). (B) The percentage of apoptotic cells was determined 12 hours after blue light exposure by staining all nuclei with Hoechst 33342 and dead cells with propidium iodide. Replicates were assayed by counting cells within 10 microscopic fields (x40 objective) situated along the band of illumination in each culture. Data are the mean ± SEM of one experiment. (C) The numbers of apoptotic nuclei were determined by labeling with membrane-impermeable propidium iodide, 18 hours after exposure to blue light. Data (mean ± SEM) are expressed as number of apoptotic nuclei per field of confluent cells (1.34 mm2) and are based on the sampling of five fields of illumination for each condition in one experiment.

 

   Discussion
Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
References
 
The evidence for caspase-3 activation, together with the observation that inhibition of caspase-3 and overexpression of Bcl-2 attenuate the frequency of apoptosis in cells, indicates that exposure of RPE to blue light in the setting of intracellular A2E initiates a cell death program that is executed by a proteolytic caspase cascade and that is regulated by Bcl-2. These observations build on our previous work implicating A2E as an initiator of blue light–induced damage to the RPE20 and are consistent with the known susceptibility of RPE cells to blue light toxicity in vivo.46 47 48 49 50

Apoptosis induced in A2E-loaded RPE cells by blue light was inhibited by approximately 50% in the presence of the caspase-3 inhibitor, Z-DEVD-fmk. This level of inhibition is consistent with that achieved in other studies using Z-DEVD-fmk to block apoptosis in whole cells induced by a variety of agents.51 52 53 The failure of Z-DEVD-fmk to completely prevent apoptosis may be interpreted as evidence for a redundant caspase-3–independent pathway. On the other hand, because signal augmentation occurs along the caspase cascade, blockage of a single protease may reflect the kinetics of amplification along this enzyme pathway.51 If even a small amount of caspase is activated, it could be sufficient to induce the death program. In addition, although Z-DEVD-fmk is membrane permeable, some restriction on the penetrability of the tetrapeptide inhibitor is suggested by the observation that considerably higher concentrations of inhibitor are required to inhibit the death of intact cells than for inhibition of caspase-3 in cell-free systems.54 55

The Bcl-2 protein resides on the outer of the two mitochondrial membranes and is one member in a family of proteins that play a pivotal role in the regulation of cell death. Some of the members of the Bcl-2 family, such as Bcl-2, Bcl-XL, and Bcl-w, inhibit apoptosis, whereas others, for instance Bax, Bak, and Bad, are promoters.27 33 Ectopic expression of Bcl-2 in a transgenic approach has been shown to rescue photoreceptor cells in retinal degeneration slow (rds) mice,56 but has little or only temporary effects in other forms of retinal degeneration in mice, including a model of light damage.57 58 59 Similarly, enforced overexpression of Bcl-2 in cultured cells has been shown to confer a resistance to apoptosis induced by many,45 60 61 62 63 64 65 but not all,60 66 67 68 cell death stimuli. Under some circumstances, the ratio between the pro- and anti-apoptotic molecules is considered to be at least one of the determinants of the susceptibility of a cell to a death stimulus.69 In keeping with this, the extent to which overexpression of Bcl-2, after transfection into cultured cells, inhibits apoptosis has been shown to vary with the level of Bcl-2 protein expression.61

A number of mechanisms have been proposed to explain the ability of Bcl-2 to suppress apoptosis. For instance, the formation of heterodimers between antiapoptotic and proapoptotic proteins is thought to lead to the neutralization of activity.69 Apart from heterodimerization, Bcl-2 also can protect against release of the apoptogenic protein cytochrome c35 70 and avert a loss of mitochondrial membrane potential by inducing an H+ efflux from the mitochondria71 —measures that guard against downstream caspase activation.

Because the accumulation of lipofuscin by aging RPE cells is greatest in the macula,11 12 A2E-mediated blue light damage may contribute to the development of areas of RPE atrophy within the parafovea. In fundus photographs, RPE atrophy can initially appear as multiple small (150–200 µm) lesions that slowly enlarge and coalesce to form the large geographic areas of atrophy typical of non-neovascular AMD.72 73 It is interesting that laser scanning ophthalmoscopy reveals focal areas of increased autofluorescence at locations on the fundus that are otherwise unremarkable ophthalmoscopically.3 74 It has been suggested that these areas of increased autofluorescence may correspond to groups of RPE cells that contain higher quantities of lipofuscin than surrounding cells and that may be at risk for cell loss.3 74

Although epidemiologic studies concerned with a potential causal relationship between light exposure and AMD have been inconclusive,75 76 it is potentially relevant that the Chesapeake Bay Waterman Study found that individuals with advanced AMD, including geographic atrophy, reported the highest estimates of blue light exposure during the 20-year period leading up to the study.76 The propensity for blue light damage to the RPE may be particularly significant in the elderly aphakic or pseudophakic eye, wherein lipofuscin accumulation is substantial and the crystalline lens, which yellows with age and thus provides some protection from blue light, has been removed. Indeed, in an investigation analyzing associations between lens opacities and AMD, it was concluded that cataract extraction, without implantation of a UV-blue light–absorbing intraocular lens, leads to an increased risk of AMD.77 In another study, progression to AMD also occurred more frequently in eyes undergoing cataract extraction with intraocular lens implantation than in fellow eyes.78 The contribution of A2E to the pathogenesis of AMD under these and other conditions, deserves further study.


   Acknowledgements
 
The authors thank Koji Nakanishi for providing A2E.


   Footnotes
 
Supported by National Institutes of Health Grant EY-12951, Fight for Sight, and unrestricted funds from Research to Prevent Blindness.

Submitted for publication October 30, 2000; revised January 25, 2001; accepted February 7, 2001.

Commercial relationships policy: N.

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.

Corresponding author: Janet R. Sparrow, Department of Ophthalmology, Columbia University, 630 W. 168th Street, New York, NY 10032. jrs88{at}columbia.edu


   References
Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

  1. Holz, FG, Bellmann, C, Margaritidis, M, Schutt, F, Otto, TP, Volcker, HE (1999) Patterns of increased in vivo fundus autofluorescence in the junctional zone of geographic atrophy of the retinal pigment epithelium associated with age-related macular degeneration Graefes Arch Clin Exp Ophthalmol 237,145-152[Medline][Order article via Infotrieve]
  2. Delori, FC, Dorey, CK (1998) In vivo technique for autofluorescent lipopigments Methods Mol Biol 108,229-243[Medline][Order article via Infotrieve]
  3. von Ruckmann, A, Fitzke, FW, Bird, AC (1997) Fundus autofluorescence in age-related macular disease imaged with a laser scanning ophthalmoscope Invest Ophthalmol Vis Sci 38,478-486[Abstract/Free Full Text]
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