To investigate the role of extracellular zinc on the death process of cultured human retinal pigment epithelial (RPE) cells.
Confluent cells on borosilicate glass coverslips were treated with substances in serum-free growth medium for various times and were analyzed for death by means of changes in morphologic features, numbers of attached cells, and terminal deoxynucleotidyl transferase–mediated deoxyuridine triphosphate nick-end labeling (TUNEL) procedure. Some cultures were also exposed to experimental ischemia (defined as a lack of oxygen, glucose, and serum). Electrophoresis and Western blotting and enzyme assays were used to investigate changes in expression of the protease enzyme, caspase-3.
Experimental ischemia caused death of RPE cells. Zinc sulfate had no effect on these cells at low concentrations (100 pmol/L to 10 nmol/L), but protected them at higher concentrations (≤10 µmol/L) and appeared to exacerbate cell death at still greater concentrations. Moreover, zinc compounds(>10 µmol/L) also induced death of cells in control cultures that could be blocked by zinc chelators and partially by the caspase-3 inhibitor, DEVD-FMK. Zinc also increased the amount of the active form of caspase-3 in RPE cells.
Zinc salts protect RPE cells from experimental ischemia–induced death at low concentrations (100 pmol/L-10 nmol/L). However, at higher concentrations, zinc causes cell death and alters the cellular level of caspase-3. These observations are consistent with the death process being apoptosis.
Zinc supplements are taken by many individuals. Low doses of zinc can protect RPE cells against ischemic-type insults as may occur in certain ocular complaints. Furthermore, high concentrations of zinc can damage RPE cells. Because zinc ions are known to be taken up by RPE cells from the choroidal circulation, the actual therapeutic dose taken by patients is critical.
ZINC IS one of the most abundant trace elements in nervous tissues, including the neural retina and the associated retinal pigment epithelium(RPE).1,2 Zinc ions bind to and contribute to the actions of more than 400 metalloproteins that play a part in nucleic acid and protein synthesis, energy metabolism, and intracellular signaling.3 By far the most studied roles for zinc in mammalian cells, however, are those involving maintenance of cellular antioxidation states, by acting on catalase, copper-zinc superoxide dismutase, metallothionein, and some nicotinamide adenine dinucleotide phosphate oxidases.3 Deficiency of zinc ions causes night blindness, impaired dark adaptation, and reduced visual acuity that can be reversed by dietary supplementation of this ion.4
Contradictory roles for zinc have been described in the induction, maintenance, and inhibition of apoptotic cell death.1,5,6 Apoptosis is a form of programmed death that is under tight genetic control and that involves removal of damaged or superfluous cells without inflammation, eg, during aging and development.7-9 Apoptosis is characterized by cytoplasmic and nuclear shrinkage and precise internucleosomal chromatin cleavage.10,11 It has been shown in different cells that zinc supplementation prevents apoptosis induced by a wide variety of agents, and that cells grown under conditions of zinc deficiency or in the presence of zinc chelators can undergo this death process spontaneously.1,5 It is thought that zinc acts as an inhibitor of the endonuclease that is responsible for apoptotic DNA degradation.12 Furthermore, it has been reported recently that zinc acts as a potent inhibitor of the protease caspase-3 (CPP32/yama/apopain).13 This protease is frequently activated in mammalian cell apoptosis to cleave key cellular proteins, leading to completion of the apoptotic process once the cell has been committed to die.14-16 Evidence exists, however, that zinc may actually induce apoptosis at physiologic concentrations, and this has been demonstrated in pancreatic acinar cells17 and mouse thymocytes.18-20 It is now believed, therefore, that the concentration of this ion is important in determining whether it acts as an activator or inhibitor of the death process.
The RPE forms a cellular monolayer that lies immediately posterior and in juxtaposition to the retina, playing a vital role in the maintenance of the normal functioning of this tissue. Damage to the RPE will necessarily affect the retina and may lead to its degeneration secondary to an initial insult. It has been suggested that RPE cells die by apoptosis in conditions such as age-related macular degeneration, retinal-choroidal ischemia, or proliferative vitreoretinopathy and as a result of intense light damage.21-24 Moreover, in vitro, RPE cell apoptosis can be induced by nutrient deprivation(experimental ischemia),25,26 inhibition of glutathione biosynthesis,27 or hypoxia.28 Therefore, any compounds that prevent or reduce incidences of RPE apoptosis, or indeed any other modes of RPE cell death, should be identified to preserve retinal function in relevant ocular disease states.
Zinc is known to be involved in the maintenance of RPE cell antioxidant levels,29,30 and it is also suspected that oxidative insults may contribute to the development of diseases such as age-related macular degeneration.31-33 Dietary zinc supplementation has therefore been suggested as a measure to counteract ocular damage resulting from this and other similar conditions.33-35 Because raised intracellular levels of oxidative agents (reactive oxygen species) are also involved in death processes,36 zinc may have a protective effect on death of RPE cells. Our aims, therefore, were to investigate the effects of zinc on experimental ischemia–induced death of cultured human RPE cells. Furthermore, a range of zinc concentrations were investigated to see whether zinc had any detrimental effects on RPE cells.
Deoxyribonuclease 1, proteinase K, and biotin-16-2′-deoxyuridine-5′-triphosphate(biotin-16-dUTP) were obtained from Boehringer Mannheim (Lewes, England). Terminal deoxynucleotidyl transferase was obtained from Promega (Southampton, England) and avidin-biotin complex kits from Vector Laboratories (Peterborough, England). Fetal bovine serum (approved by the European Community), Hams F-10, amphotericin B (Fungizone), glutamine, 0.25% (weight to volume) trypsin solution, and 24-multiwell plates (NUNC) were from GIBCO (Paisley, Scotland); 25- and 75-cm2 tissue culture flasks were obtained from Falcon (Oxford, England). Monoclonal anti–caspase-3 antibody was obtained from Signal Transduction Labs (distributed by Affinity, Exeter, England). The caspase-3 activity assay kit was obtained from Clontech Labs Inc (Palo Alto, Calif), including the caspase-3 inhibitor DEVD-FMK. All other chemicals were obtained from Sigma-Aldrich (St Louis, Mo).
Postmortem donor human eyes (donors aged 7, 26, 48, 54, and 58 years) were obtained without their cornea (for transplantation purposes) from Bristol Eye Bank, Bristol, England, up to 48 hours after enucleation. These were used immediately or stored for up to 24 hours at 4°C. All culture work was undertaken in a sterile laminar flow hood (ICN Flow, Thame, England). Cultures of RPE cells were prepared and characterized by labeling for cytokeratin (KG 8.13) as described previously.37 Culture medium consisted of Hams-F10 supplemented with 10% (volume-volume) fetal bovine serum, 0.4% glucose, 2-mmol/L glutamine, amphotericin B (25 µg/mL), and gentamicin (100 µg/mL). Primary cultures were grown in 25-cm2 culture flasks and passaged in a ratio of 1:3, and thereafter, in 75-cm2 flasks. While growing, cultures were kept in an incubator at 35.5°C, with saturating humidity and an atmosphere of 5% carbon dioxide to 95% air. After reaching the third passage, some cells were transferred to 13-mm glass coverslips in 24-well plates at an approximate seeding density of 2.0 × 104 cells per well.
Confluent RPE cells on coverslips were transferred to serum-free culture medium for 4 hours before the experiments. Cultures were then transferred in fresh serum-free medium to a sealed glass chamber with saturating humidity, within the normal incubator, which was linked to a reservoir of oxygen-free gas mixture (95% nitrogen to 5% carbon dioxide). This delivered anoxic conditions to the cells. Once the cultures were inside, the chamber was completely sealed, and the anoxic gas mixture passed through for 3 hours to ensure that all oxygen was removed. Hypoglycemia was achieved by incubating the cells with serum-free medium lacking additional glucose. The combination of anoxic and hypoglycemic conditions was defined as experimental ischemia.
To assess whether zinc could protect against apoptotic death that was induced by experimental ischemia, increasing concentrations of zinc sulfate were added to cells that were to be deprived of nutrients for 48 or 72 hours. To see the effect of zinc on untreated cultures, zinc sulfate or other zinc salts also were added at different concentrations to control cells for varying periods. In other instances, different compounds were coincubated with zinc(fetal calf serum, melatonin, flupirtine gluconate, EDTA, diethyldithiocarbamate[DDCA], and cycloheximide) in serum-free medium at the concentrations described. To determine the effectiveness of caspase-3 inhibition in the prevention of cell death, cultures were preincubated with DEVD-FMK for 24 hours before insult(10 or 100 µmol/L).
At the appropriate times, coverslips were removed and cells were fixed with 4% paraformaldehyde in phosphate-buffered saline solution (137-mmol/L sodium chloride, 5.4-mmol/L potassium chloride, 1.28-mmol/L sodium dihydrogen phosphate, and 7-mmol/L disodium hydrogen phosphate [pH, 7.4]) for 30 minutes. Some cells were visualized with a solution of toluidine blue (0.5% toluidine blue, 0.5% thionine, and 1% sodium tetraborate) to enable culture density(total number of cells per square millimeter) to be determined. Other cells were analyzed for DNA degradation by staining with the terminal deoxynucleotidyl transferase–mediated deoxyuridine triphosphate nick-end labeling (TUNEL) method. A single culture density determination was obtained by using a standard hemocytometer to count and then average the amount of cells from 4 different fields. Statistical analyses were performed by using the paired t test to compare experimental counts with those obtained from parallel control cultures. A value of P<.05 was considered significant.
Other cultures were assessed for activity and expression of the enzyme caspase-3 using a spectrophotometric assay and using electrophoresis and Western blotting analysis, respectively. The cells that were analyzed for caspase-3 expression and activity were grown to confluence in 75-cm2 culture flasks for use.
For each set of analyses, cultures derived from different donors were used in a randomized way, ensuring that at least 3 different cell lines were represented in each case. Values presented for n in each case refer to replicates from different experiments.
Assessment of dna breakdown using the tunel method
This method was performed as described previously.37 Briefly, cells were treated with 1% hydrogen peroxide, and then the free DNA ends were labeled by incubating in buffer (30-mmol/L Tris hydrochloride [pH, 7.2] containing 140-mmol/L sodium cacodylate and 1-mmol/L cobalt chloride) along with 0.25 U/µL of terminal deoxynucleotidyl transferase and 10-µmol/L biotin-16-dUTP. The reaction was stopped by washing (2 × 15 minutes) in saline sodium citrate buffer (30-mmol/L sodium citrate with 300-mmol/L sodium chloride). After an additional wash in 2% bovine serum albumin (in phosphate-buffered saline solution) for 15 minutes, stained nuclei were visualized using an avidin-biotin-peroxidase complex kit with 3-3′-diaminobenzidine and 0.1% hydrogen peroxide as substrates. The number of cells with positive TUNEL findings was determined as for the total counts with a hemocytometer.
After the appropriate drug incubation times, adherent cells were washed and harvested in phosphate-buffered saline solution (35.5°C) using a cell scraper. Cell counting was used to determine that aliquots of 2 × 106 cells were assayed in each case. These were collected by centrifugation(80g for 8 minutes at 4°C). Cells were then lysed and incubated with the reaction substrate (peptide with the sequence DEVD, conjugated to p-nitroanilide) for 1 hour at 37°C in a reaction buffer as described in the manufacturer's instructions and by Gurtu et al.38 Samples were read on a spectrophotometer at a wavelength of 405 nm. The change in optical density values in the apoptotic samples relative to control samples indicates the increase in DEVD-dependent caspase-3 activity in these samples.
Electrophoresis and western blotting
Confluent RPE cells grown in 75-cm2 flasks were harvested as per the cells assayed for caspase-3 activity. Cells were sonicated in buffer(20-mmol/L Tris hydrochloride [pH, 7.4] containing 2-mmol/L EDTA, 0.5-mmol/L ethyleneglycoltetracetic acid, and 0.1-mmol/L phenylmethylsulfonyl fluoride). An equal volume of sample buffer 62.5-mmol/L Tris hydrochloride [pH, 7.4] containing 4% sodium dodecyl sulfate, 10% glycerol, 10% β-mercaptoethanol, and 0.002% bromophenol blue) was added, and samples were boiled for approximately 3 minutes. An aliquot was taken for the determination of protein concentration using the method of Bradford.39 Electrophoresis of samples was performed using the method of Laemmli40 with 10% polyacrylamide gels containing 0.1% sodium dodecyl sulfate. Proteins were transferred to nitrocellulose41 and blots were stained as previously described37 for the presence of caspase-3.
The RPE cells died when subjected to a deprivation of oxygen, glucose, and serum from their growth medium (experimental ischemia) (Table 1). As reported previously,26 after 72 hours of experimental ischemia, culture density was reduced and nuclear labeling by means of the TUNEL procedure increased when compared with untreated cells.
Zinc had no obvious effect on cultures at low concentrations (100 pmol/L to 10 nmol/L), but protected cells from experimental ischemia (48 or 72 hours) at higher concentrations (≤10 µmol/L) as indicated by a greater density of cells associated with the coverslips (Figure 1A) and a reduced amount of TUNEL-positive nuclei (Figure 1B). At concentrations higher than 10 µmol/L, zinc exacerbated cell death caused by experimental ischemia.
When control cultures were treated with concentrations of zinc sulfate higher than 10 µmol/L, large numbers of cells died (Figure 2 and Figure 3). Cell shrinkage and TUNEL assessment suggested this death process to be apoptosis(Figure 2). As can be seen in Table 1, the effects of other zinc salts(zinc acetate and zinc chloride) did not vary significantly from those of zinc sulfate in this regard.
The mechanism of cell death induced by zinc sulfate appeared to differ from that of experimental ischemia, as substances that have been shown previously to attenuate the latter process had no effect on zinc-induced cell death (Table 1) (100-µmol/L flupirtine, 100-µmol/L melatonin, and 10% [volume-volume] fetal calf serum). The cation chelator, EDTA, and to a lesser extent, the zinc chelator, DDCA, significantly attenuated RPE cell death induced by zinc, however (Table 1).
Caspase-3 activity (Figure 4)was detectable in control cells that had been incubated in serum-free medium for 24 hours. The amount of activity increased optimally by approximately 10-fold in RPE cells treated with 10-µmol/L zinc sulfate for 24 hours(Table 2). Greater concentrations of zinc sulfate (≤10 mmol/L) produced less of a measurable amount of caspase-3. Significantly, experimental ischemia had no effect on caspase-3 (Table 2). Western blot analysis revealed an increased presence of the 17-kd active protein for caspase-3 after incubation of cultures with 100-µmmol/L zinc sulfate for 6 and then 24 hours, compared with cells incubated in just serum-free medium or medium containing 10% (volume-volume) fetal bovine serum (Figure 5). Furthermore, there was no appearance of the 17-kd immunoreactive protein for caspase-3 in cells subjected to experimental ischemia for 24 hours (Figure 5). Coincubation of 100-µmol/L cycloheximide also had no effect on the death of RPE cells induced by 100-µmol/L zinc sulfate(Table 1). Zinc-induced death but not experimental ischemia–induced cell death could be partially ameliorated by preincubation of the cells with DEVD-FMK, a specific and irreversible caspase-3 inhibitor, at concentrations of 100 µmol/L and, to a lesser extent, 10 µmol/L (Table 1).
Previous studies have shown that human RPE cells subjected to experimental ischemia die by means of a process that is characteristic of apoptosis.25,26,37 This is because cells treated in this way exhibit a shrunken appearance and have nuclei that stain positively for the TUNEL reaction. Furthermore, apoptosis-related genes have also been reported to be affected, consistent with this idea.25 It is believed, however, that the differences between distinct types of cell death are not so clear.42 Indeed, many of the events that have been ascribed previously to a particular mode of death have now been shown to be less specific. Thus, although the data described herein suggest that the death process involved is apoptosis, this cannot be stated definitively.
The present data show that zinc sulfate has contrasting effects on cultured human RPE cells, depending on the concentrations used. Previous reports have demonstrated that exogenously applied zinc can have biphasic effects on nonneuronal cells by either initiating or counteracting apoptosis.5,6,18,43 The concentrations of zinc required to cause or prevent apoptosis, however, seem to be variable and may depend on the cell type under investigation. In mouse thymocytes, for example, high zinc concentrations (10 µmol/L to 1 mmol/L) inhibit serum-free medium- and dexamethasone-induced apoptosis, whereas zinc at concentrations of less than 10 µmol/L induces apoptosis.18,20,43
The mechanism by which zinc induces death of RPE cells cannot be inferred from the present data. Each of the zinc salts tested had the same ability to initiate death of RPE cells in culture, however, confirming that it was zinc and not the respective anion in each case that was responsible. It is also likely that zinc is equally effective at entering RPE cells in each case. This has been described in intact erythrocytes, which accumulated zinc sulfate, zinc acetate, or zinc chloride from an incubation medium at the same rate.44 It is known that intracellular zinc can modulate expression and activity of immediate early gene products1 such as c-fos or c-jun and that these have been correlated with apoptosis, particularly in neurons.45 Furthermore, protein kinases such as protein kinase C have high-affinity binding sites for zinc.46 This cation has a biphasic effect on protein kinase C, acting as a stimulator at concentrations lower than 10 µmol/L and an inhibitor at higher amounts.1 The latter effect may well explain the toxic effects of zinc that are reported in the present study, as it has already been shown that inhibition of protein kinase C with staurosporine or polymyxin B sulfate can cause apoptosis of human RPE cells,37 and with hypericin can cause apoptosis of bovine RPE cells.47
It has been thought that the primary mode of action for zinc as an attenuator of apoptosis is to inhibit the endonuclease that is responsible for internucleosomal DNA degradation5,6,12 usually but not always accompanying this death process.48 A number of recent observations have suggested, however, that zinc may have other cellular targets during apoptosis.49,50 One such enzyme that is activated by zinc at submicromolar concentrations is the DNA repair enzyme poly-(adenosine diphosphate)-ribose polymerase (PARP).51 When PARP is degraded during the early stages of apoptosis, the cell loses its capacity to repair DNA strand damage. Increases in intracellular zinc concentrations, such as in the present study, will therefore increase the antiapoptotic activity of PARP. Zinc is also responsible for the activity of many antioxidant or metabolic enzymes in the RPE such as catalase, alkaline phosphatase, α-mannosidase, and metallothionein.30 Because increases in intracellular levels of pro-oxidants are known to be associated with apoptosis, raising the cellular supply of zinc to the level where its action on the antioxidant status becomes optimal could counteract cell-death processes. Furthermore, zinc has also been reported to potently inhibit the protease enzyme, caspase-3,13 which is involved in processing of the apoptotic signals in the regulation stages of this type of cell death.15 It is unlikely that this action accounts for the counteraction of experimental ischemia–induced apoptosis in the present study, however, as there is no increase in the active form of caspase-3 during this process (Table 2), and this cell death cannot be blocked by the caspase-3 inhibitor (Table 1).
Increases in the amount of caspase-3 present in cultures after zinc treatment (Figure 4 and Table 2) confirm that this enzyme can be induced in an active form during RPE cell death. A small quantity of caspase-3 is also present in control cultures incubated in serum-free medium for 24 hours, and this may reflect the low incidence of death of these cells when incubated without serum, as shown previously.26 Because experimental ischemia did not give rise to any changes in caspase-3, and because the irreversible caspase-3 inhibitor, DEVD-FMK, partially attenuated zinc- but not ischemia-induced death, then it is clear that both death processes involve distinct pathways. It is obvious, therefore, that distinct pathways are involved in apoptotic death of certain cells and that caspase-3 does not participate in all of these. This has been suggested previously.52
The data shown in Figure 5indicate that detection of the active 17-kd caspase-3 enzyme is possible after 6 and 24 hours of treatment of the cultures with zinc sulfate at concentrations of 100 µmol/L. However, the toxic effects of zinc sulfate are not blocked in the presence of the protein synthesis inhibitor cycloheximide. Caspase-3 is therefore not produced, de novo, during this mode of death. These results are explained by the cellular presence of procaspase-3, an inactive precursor enzyme, which is cleaved in the early stages of an apoptotic insult by other proteases (eg, caspase-9) to release an active 17-kd form.15 It is suspected that without a signal telling the cells to enter a death pathway, procaspase-3 is present but inactive. In agreement with the assay data for caspase-3, the active form of this enzyme could not be detected after 24 hours of experimental ischemia. These data also confirm that nutrient deprivation–induced death of RPE cells does not involve this protease. The fact that substances that have been shown previously to block experimental ischemia–induced death in human RPE cultures (eg, serum, melatonin, and flupirtine)25,26 have no effect on death induced by zinc also confirms that both insults must involve distinct mechanisms.
In the present investigation, coincubation of cultures with zinc sulfate and the metal-chelating agent EDTA led to a complete counteraction of toxic effects. The marked decrease in RPE cell death in such instances indicates that zinc uptake is critical for toxic events to occur, as EDTA will not pass the plasma membrane and so must have exerted its effects extracellularly. The more readily membrane-permeable chelator, DDCA, was not quite so effective in counteracting cell death as EDTA. In this instance it is possible that the chelator exerted its protective effect on RPE cells by binding extracellular zinc, as did EDTA. Moreover, because it is known that intracellular entry of zinc chelators will promote cell death,6,20,53 then it is also possible that in the present study the effect of DDCA was manifested as an intracellular toxic effect. However, previous data (J.W., unpublished data, January 1998) have indicated that DDCA is not toxic to RPE cells alone, and so the difference in counteracting zinc-induced RPE cell death in the present study was probably caused by a greater zinc-chelating property for EDTA.
It is obvious that the cellular effects of zinc, nutrient deprivation, and pathways leading to death are extremely complex events that are likely specific for cell type and paradigm. It must be borne in mind, therefore, that the effects of zinc reported herein are relevant only for RPE cells that have been cultured and treated under the conditions described. This means that the effects may differ in different strains of RPE cells, in cells cultured in different ways, under different conditions of assay, or for RPE cells in situ. Generally, RPE cells in culture have been shown to perform a number of in vivo functions readily, such as ingestion and degradation of photoreceptor outer segments54,55 and uptake of important ions such as zinc,56 and so studies on these cells provide useful data for the understanding of their in situ functioning. Furthermore, the use of donors of different ages for establishment of the RPE cells used herein reveals consistency in the data, suggesting further relevance.
With these points in mind, then, it can be stated that the present studies show zinc to protect against or induce RPE cell death that is characteristic of apoptosis, according to its concentration. Further studies will determine the effects of zinc on RPE cells in vivo. It has also been reported previously that zinc has a role in ischemic, necrotic-type cell death in the brain57,58 and the retina.59 All of these results implicate zinc involvement in cellular death processes. Further experiments will have to be undertaken to describe the mechanism of damage and the role of caspase-3. Indeed, a more detailed analysis of the interplay between cultured RPE cells and zinc will need to be performed, and this will have to be correlated with in situ data. The present data should, however, be taken into account by individuals taking high-dose dietary supplements of zinc, because this ion obviously can be specifically harmful to certain cells, under certain conditions, at elevated concentrations.
Accepted for publication June 22, 2000.
The financial assistance of The National Eye Research Centre, Bristol, England, is gratefully acknowledged.
Reprints: Neville N. Osborne, PhD, DSc, The Nuffield Laboratory of Ophthalmology, University of Oxford, Walton Street Oxford, OX2 6AW, England(e-mail: firstname.lastname@example.org or email@example.com).
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