Phototoxicity and cytotoxicity of fullerol in human retinal pigment epithelial cells

Toxicology and Applied Pharmacology 242 (2010) 79–90

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Toxicology and Applied Pharmacology j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / y t a a p

Phototoxicity and cytotoxicity of fullerol in human retinal pigment epithelial cells Albert R. Wielgus a, Baozhong Zhao a, Colin F. Chignell a, Dan-Ning Hu b, Joan E. Roberts c,⁎ a b c

Laboratory of Pharmacology, National Institute of Environmental Health Sciences, Research Triangle Park, NC 27709, USA Tissue Culture Center, New York Eye and Ear Infirmary, New York, NY 10002, USA Department of Natural Sciences, Fordham University, Lincoln Center, 113 West 60th Street, New York City, NY 10023, USA

a r t i c l e

i n f o

Article history: Received 28 April 2009 Revised 27 August 2009 Accepted 28 September 2009 Available online 2 October 2009 Keywords: Nanoparticles Fullerenes Fullerol Ocular toxicology Ocular phototoxicity Human retinal pigment epithelial cells Singlet oxygen Superoxide Lipid peroxidation

a b s t r a c t The water-soluble nanoparticle hydroxylated fullerene [fullerol, nano-C60(OH)22–26] has several clinical applications including use as a drug carrier to bypass the blood ocular barriers. We have previously found that fullerol is both cytotoxic and phototoxic to human lens epithelial cells (HLE B-3) and that the endogenous antioxidant lutein blocked some of this phototoxicity. In the present study we have found that fullerol induces cytotoxic and phototoxic damage to human retinal pigment epithelial cells. Accumulation of nano-C60(OH)22–26 in the cells was confirmed spectrophotometrically at 405 nm, and cell viability, cell metabolism and membrane permeability were estimated using trypan blue, MTS and LDH assays, respectively. Fullerol was cytotoxic toward hRPE cells maintained in the dark at concentrations higher than 10 μM. Exposure to an 8.5 J·cm− 2 dose of visible light in the presence of N 5 μM fullerol induced TBARS formation and early apoptosis, indicating phototoxic damage in the form of lipid peroxidation. Pretreatment with 10 and 20 μM lutein offered some protection against fullerol photodamage. Using time resolved photophysical techniques, we have now confirmed that fullerol produces singlet oxygen with a quantum yield of Φ = 0.05 in D2O and with a range of 0.002–0.139 in various solvents. As our previous studies have shown that fullerol also produces superoxide in the presence of light, retinal phototoxic damage may occur through both type I (free radical) and type II (singlet oxygen) mechanisms. In conclusion, ocular exposure to fullerol, particularly in the presence of sunlight, may lead to retinal damage. Published by Elsevier Inc.

Introduction Water-soluble fullerene nanoparticles can bypass the blood–brain and blood–retina barriers (Ji et al., 2006). Nanoparticle delivery of photosensitizers to ocular (and dermal) cells improves photodynamic therapy of tumors (Zhao et al., 2009). Nanoparticles also show promise as drug carriers to the retina (Yamago et al., 1995; Calvo et al., 1996; Dugan et al., 1997; Da Ros and Prato, 1999; Nakamura and Isobe, 2003) and are currently being developed as carriers for antiangiogenesis drugs and vectors for gene delivery to retinal pigment epithelial cells (RPE) (Prow et al., 2008; Bejjani et al., 2005) for treatment of wet macular degeneration. It has been shown that a single intravitreous injection of aliphatic nanoparticles preferentially localizes in retinal pigment epithelial cells (Bourges et al., 2003; Bejjani et al., 2005), and that these nanoparticles are retained by RPE cells for a significant period of time (Bourges et al., 2003; Bejjani et al., 2005). Water-soluble forms of fullerene exhibit antitumor, antibacterial and antiviral activity, including inhibition of HIV protease (Bogdanović et al., 2004; Lyon et al., 2006; Badireddy et al., 2007; Friedman et al., 1998; Nakamura and Isobe, 2003; Schinazi et al., 1993). Radiolabeled

⁎ Corresponding author. Fax: +1 212 636 7217. E-mail address: [email protected] (J.E. Roberts). 0041-008X/$ – see front matter. Published by Elsevier Inc. doi:10.1016/j.taap.2009.09.021

fullerol has been used to determine the biodistribution and tumor uptake in five mouse tumor-bearing models (Ji et al., 2006). The pristine form of fullerol showed a tumor-inhibitory effect in the murine H22 hepatocarcinoma model (Zhu et al., 2008) in vivo and is cytotoxic to human liver carcinoma cells (HepG2) in vitro (Sayes et al., 2004, 2005). Fullerol exhibited a protective influence on the heart and liver tissue against chronic toxicity induced by doxorubicin (Injac et al., 2009) and radioprotective activity against (8 Gy) X-rays in animal models (Dordević and Bogdanović, 2008). Beyond their use as pharmaceutical agents (Dordević and Bogdanović, 2008; Markovic and Trajkovic, 2008), fullerols have been recommended for use in waste treatment of contaminated waterways (Anderson and Barron, 2005; Brant et al., 2007). Water-soluble fullerenes are not genotoxic (Zakharenko et al., 1997) and have low cytotoxicity, although they have been found to show some cytotoxicity to human dermal fibroblasts (Sayes et al., 2004, 2005, 2007), keratinocytes (Zhao et al., 2008b) and human lens epithelial cells (Roberts et al., 2008; Zhao et al., 2009). These nanoparticles can be stored in the body for months (Yamago et al., 1995), thus offering the potential for chronic side effects. Although many nanoparticles absorb in both the UV and visible light range, few nanomaterials have been assessed for their potential phototoxicity. The human eye is constantly subjected to environmental light. Only visible light (wavelengths longer than 400 nm) is transmitted to the adult human retina, as ultraviolet radiation is

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filtered by the human lens (Roberts, 2001). When any substance accumulates in the retina, absorbs visible light (λ N 400 nm), and produces reactive oxygen species, it has the potential to induce phototoxic damage in the human retina (Roberts, 2002). Such damage is seen in patients taking phototoxic prescription drugs, diagnostic dyes, or over-the-counter herbal medications (Wielgus et al., 2007). As human RPE cells are postmitotic after birth (Hjelmeland et al., 1999), damage to these cells can lead to transient or permanent loss of vision due to early retinal degeneration. Water-soluble hydroxylated fullerene [fullerol, C60(OH)22–26] (Fig. 1A) and its aggregated form (Fig. 1B) absorb light in both the UVA and visible light regions and therefore have the potential to be phototoxic to the skin and the eye. We have previously found that fullerol exposed to UV radiation or visible light is phototoxic to human keratinocytes in vitro (Zhao et al., 2008a). We have just reported that

fullerol induces phototoxic damage to human lens epithelial cells (Roberts et al., 2008) when irradiated with either UV or visible light. We have also found that water-soluble cyclodextrin-complexed fullerene also induces phototoxic damage to human lens epithelial cells (Zhao et al., 2009). Phototoxic reactions occur when light activates a substance, forms a long lived triplet, and from there produces reactive oxygen species. Photoexcitation of fullerene derivatives efficiently produces an excited triplet state (Arbogast et al., 1991; Guldi and Prato, 2000) when irradiated with visible light. Energy and electron transfer from this triplet state to molecular oxygen produces superoxide (Zhao et al., 2008a) and other reactive oxygen species (Prat et al., 1999; Yamakoshi et al., 2003; Chin et al., 2008). Our previous studies (Zhao et al., 2008b) and others (Pickering and Wiesner, 2005) have shown that fullerol produces superoxide in the presence of light.

Fig. 1. (A) The structure of fullerol [C60(OH)22–26] and (B) possible structure of fullerol aggregates in aqueous solution. (C–F) TEM images of 50 μM fullerol in different solutions containing 1% DMSO. (C) Cell culture medium HAM F-12. (D) 5 mM PBS. (E) Deionized H2O; (F) 50 mM NaCl.

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In this study we have assessed the potential phototoxicity and cytotoxicity of fullerol in vitro in human retinal pigment epithelial cells and determined whether non-toxic quenchers might mitigate some of the phototoxic damage. We have also measured fullerol's production of singlet oxygen, another reactive oxygen species. Materials and methods Reagents. Perinaphthenone, potassium iodide, Rose Bengal, sodium deuteroxide, and 85% phosphoric acid–d3 in D2O were purchased from Aldrich (Milwaukee, WI); fetal bovine serum (FBS) was from Biofluids (Rockville, MD); deuterium oxide was from Cambridge Isotope Laboratories (Andover, MA); N-acetyl-L-cysteine was from Fluka (Milwaukee, WI); cell culture medium HAM F-12 with phenol red, gentamicin, trypan blue, and trypsin–EDTA were from GIBCO Invitrogen Corporation (Carlsbad, CA); sodium hydroxide and HPLC grade water from J.T. Baker (Phillipsburg, NJ); acetone, 38% hydrochloric acid, and trichloroacetic acid were from Mallinckrodt (Paris, KY); and acetonitrile, chloroform, dichloromethane (CH2Cl2), 2,6-di-tert-butyl-4-methylphenol (butylated hydroxytoluene), sterile dimethyl sulfoxide (DMSO), 1,4-dioxane, ethanol, L-glutamine, lutein, bovine albumin (fraction V), magnesium perchlorate, 5-sulfosalicylic acid, tetrahydrofuran (THF), 2-thiobarbituric acid, and toluene were from Sigma (St. Louis, MO). Phosphate buffered saline without Ca2+ and Mg2+ (PBS) and Hank's balanced salt solution containing Ca2+ and Mg2+ (HBSS), both sterile, were prepared at Media and Glassware Units at NIEHS (RTP, NC). All reagents used for experiments were at least analytical grade. Fullerene derivatives. Hydroxy fullerene [C60(OH)22–26], fullerene (C60), and fullerene hydride (C60Hx, where x ≈ 36) used during this study were synthesized by Material and Electrochemical Research (MER) Corporation (Tucson, AZ) for the National Toxicology Program (NTP) at NIEHS. The materials presented in this paper are quality controlled by the National Toxicology Program and have been supplied under contract (TOXBC) by Battelle, Ohio. In order to have consistent and reproducible results, these specific fullerene materials are used for all of the approved in vitro and in vivo experiments for NTP. Based on the Material Safety Data Sheet provided by the company, purity of the fullerene and its derivatives was 99.5%. The producer included the synthetic route for fullerol in the product description. Fullerol was produced by hydrolysis of C60Br24, and the only potential additional materials present are sodium and water. IR and mass spectrum data supplied by MER are consistent with previous published data (Rodríguez-Zavala and Guirado-López, 2004). According to the manufacturer, these samples did not contain transition metals or other catalysts. Physicochemical characterization. Absorption spectra of fullerene derivatives in various solvents were recorded with a Hewlett Packard diode array 8453 spectrophotometer (Hewlett Packard GmbH, Waldbronn, Germany). Transmission electron microscopy (TEM) images were taken on a Tecnai-12 Bio-Twin transmission electron microscope (FEI, The Netherlands) operating at 80 kV. TEM grids were prepared by evaporating approximately 20 μl of water-soluble fullerol in either deionized H2O, cell culture medium (HAM F-12 nutrient mixture supplemented with 10% fetal bovine serum, 2 mM glutamine, and 50 μg/ml gentamicin), 5 mM PBS, or 50 mM NaCl solution onto a 300 mesh carbon-coated copper grid. Dynamic light scattering (DLS) measurements of particle size and surface charge were carried out using a light scattering Zetasizer Nano-S90 instrument (Malvern Instruments, Enigma Business Park, UK). Fullerol was measured in deionized H2O, cell culture medium, 5 mM PBS, or 50 mM NaCl. These values are the average of four runs. Dynamic light scattering (DLS) was also used to measure the particle size of fullerene and fullerene hydride.

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Cell culture. Human RPE (hRPE) cells were isolated from donor eyes using the method of Hu et al. (2002) and cultured in Falcon flasks (75 cm2) with F12 nutrient mixture supplemented with 10% fetal bovine serum, 2 mM glutamine, and 50 μg/ml gentamicin. The hRPE cells were detached with trypsin–EDTA solution, diluted 1:3 to 1:4, and plated for subculture. The cells used in the present studies consisted of a pure culture of one cell line of hRPE cells in active growth status. The purity of the cell line was demonstrated by immunocytochemical methods: hRPE cells display S-100 and cytokeratin, while uveal melanocytes display S-100 antigen but not cytokeratin, and fibroblasts display neither of these proteins (Hu et al., 1993). The initial viability of the hRPE cells was 97.0 ± 0.4% as estimated by trypan blue exclusion. Preparation of solutions of fullerol for in vitro experiments. Fullerol was first dissolved in dimethyl sulfoxide (DMSO) and then diluted with HBSS in a ratio of HBSS/DMSO (99:1). The DMSO concentration was 1% in all cell culture plates including control samples. The solutions of DMSO were freshly made immediately before addition to the cell culture. Incubation of cells with fullerol. Cells were incubated in the dark at 37 °C in a 5% CO2/95% air atmosphere with 1–50 μM fullerol and without (control cells) in HBSS/DMSO (99:1). Cell viability tests were performed after incubation with fullerol for 24 h. Cells were trypsinized, stained with 0.2% trypan blue, and counted. In vitro uptake studies of fullerol in hRPE cells. For light microscope analysis, hRPE cells were incubated in 60-mm Petri dishes in 3 ml of 0–50 μM of fullerol solution in HBSS/DMSO (99:1). After a 17-h incubation in the dark in an atmosphere of 5% CO2/95% air at 37 °C, the cells were washed with HBSS. They were then overlaid with 3 ml of the cell culture medium and incubated for 2 h. Directly before microscopic observation the cells were again washed with HBSS and overlaid with 3 ml of the buffer. The cells were observed using phase contrast illumination with an Olympus IX70 microscope equipped with a Zeiss 10× UPlanFl objective (N.A. 0.3 and phase 1). A Zeiss AxioCam XEL color camera was used to acquire the images using an exposure time of 6 ms with AxioVision 4.4 software. For determination of fullerol uptake, the hRPE cells were incubated in 96-well plates in the dark in 50 μM fullerol solution in HBSS/DMSO (99:1) for up to 24 h. At a specific time of incubation, the supernatant was transferred quantitatively to an empty plate and saved for measurement. The cells in each well were washed twice with HBSS and overlaid with 50 μl of the buffer. The uptake of fullerol by hRPE cells was confirmed by fullerol absorbance at 405 nm using a GENios plate reader (Tecan US SPECTRAFluor Plus, Research Triangle Park, NC). Phototoxicity assays. The methods and assays used for detecting phototoxicity of fullerol nanoparticles were previously applied (Murdock et al., 2008; Kroll et al., 2009) and in our studies with human lens epithelial cells (HLE) (Roberts et al., 2008; Zhao et al., 2009) and with human retinal pigment epithelial cells with another photosensitizer, hypericin (Wielgus et al., 2007). These and other methods used to determine ocular phototoxicity of drugs have been reviewed (Roberts, 2002). Samples containing no fullerol (represented as zero concentration) in the dark and in the light served as the negative controls. Concentrations of fullerol RPE cells kept in the dark (cytotoxicity) served as additional negative controls for the phototoxicity studies (fullerol with light). There are clear differences in these data indicating that the greatest damage induced by fullerol in RPE cells occurs in the presence of light. Hypericin, which is a very efficient photosensitizer used in photodynamic therapy of

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the retina (Busch, 2009), was used as a positive control for the assays (data not shown). Light exposure. Human RPE cells were incubated in the dark for 17 h with 1–50 μM fullerol in HBSS/DMSO (99:1). The cells were washed and overlaid with plain HBSS, then irradiated with two fluorescent bulbs (Philips F40AX50, 40 W, 5000 K, Advantage X) through a liquid filter transmitting light of wavelengths longer than 400 nm (Wielgus et al., 2007) directly in the covered dishes or plates in which they were grown. Cells were irradiated for 60 min at an irradiance of 23.5 W/m2 measured with a spectroradiometer (LuzChem Research, Inc., Ottawa, ON, Canada) for a total dose of 8.5 J·cm− 2. The visible light exposures used in the experiments were similar to a daily dose of sunlight (Sliney, 2002; Turner and Mainster, 2008). Phototoxicity inhibition studies. In selected studies, the hRPE cells were incubated with 10 and 20 μM lutein or 0.5 and 1.0 mM N-acetylL-cysteine in HBSS/DMSO (99:1) for 2 h prior to incubation with fullerol. After incubation with a specific phototoxicity inhibitor, the supernatant was removed, and the cells were washed with HBSS and treated with 20 μM fullerol in HBSS/DMSO (99:1) for 17 h as described earlier. Then the cells were washed with HBSS, exposed to visible light as previously described, and incubated in medium/FBS (9:1) for 24 h. Cell metabolic activity and cell membrane damage tests. After light exposure, HBSS/FBS (9:1) supernatant was replaced with medium and the cells were incubated in the dark at 37 °C in a 5% CO2/95% air atmosphere for 24 h, after which the MTS (3-(4,5-dimethylthiazol-2-yl)5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium, inner salt) and LDH assays were carried out. Metabolic activity and

lactate dehydrogenase release of the cells were determined as described previously (Wielgus et al., 2007). Cell metabolic activity was determined using MTS (Cell Titer 96® AQueous Non-Radioactive Cell Proliferation Assay, Promega Corp., Madison, WI). Membrane damage was determined by LDH leakage, which was measured using the CytoTox 96® Non-Radioactive Cytotoxicity Assay (Promega Corp., Madison, WI). Measurement of apoptotic and necrotic cells. Apoptotic and necrotic cells were quantitatively evaluated by flow cytometry (Martin et al., 1995; Reno et al., 1998). After irradiation, the cells were harvested by trypsinization and collected by centrifugation at 140g for 6 min at 22 °C. Cells were washed with cold PBS and stained with Annexin V– FITC and propidium iodide (PI) using a TACS™ Apoptosis Detection Kit according to the manufacturer's instructions (Trevigen, Gaithersburg, MD). Cells positive for PI, for Annexin V–FITC, or for both were quantified by flow cytometry using a Becton Dickinson FACSort (Becton Dickinson, Mountain View, CA). Caspase-3 activity. Caspase-3 activity was determined using an ApoAlert Caspase Fluorescent Assay Kit (Clontech Laboratories, Palo Alto, CA). After irradiation, the cells were incubated in the cell culture medium/FBS (9:1) at 37 °C in a 5% CO2 atmosphere for 5 h, then removed with a cell lifter, washed with PBS by centrifugation at 400g for 10 min at 4 °C, and lysed. The cell lysates were centrifuged at 12,000g for 10 min at 4 °C to precipitate cellular debris. Supernatants were used for determination of caspase-3 activity by incubation at 37 °C for 1 h with a fluorescent substrate, DEVD-7-amino-4trifluoromethyl coumarin (DEVD-AFC). Fluorescence of cleaved AFC was measured in the GENios plate reader at excitation of wavelengths 360 and 405 nm and emission at 535 nm. A stock solution of free AFC

Fig. 2. Accumulation of fullerol in hRPE cells. The cells were incubated in the dark in HBSS/DMSO (99:1) containing (A) 0 μM, (B) 20 μM, and (C and D) 50 μM fullerol. (A–C) Microscope images taken in white light. (D) Time course of fullerol accumulation in the hRPE cells. The fullerene derivative amount taken up by the cells was determined spectrophotometrically and normalized to number of cells. Values are the means ± SD (n = 6).

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was used for preparation of a calibration curve. The results were expressed as a ratio of AFC released per hour and normalized to the amount of protein in the cell lysate. Preparation of cell samples for TBARS assay. After incubation of the cells with fullerol for 17 h in the dark, the supernatant containing fullerol was removed. The cells in each plate were flushed twice with HBSS and suspended in 5 ml of the buffer. Directly after visible light exposure the cells were scraped out of each plate, transferred to separate tubes, and then diluted with 10 mM HCl solution to 1 ml. A 700 μl portion of each cell suspension was transferred to a separate empty tube for the TBARS assay (Janero, 1990), mixed at 25 °C for 2 h, and placed on dry ice. The remaining cell suspension was sonicated for 10 s and centrifuged at 10,000g for 5 min at 4 °C. Each supernatant was transferred to a separate empty tube, frozen, and stored in a −80 °C freezer until the protein assay was performed.

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concentration was determined spectrophotometrically using an absorption coefficient of C60 in toluene of ε336 =5.04×104 M− 1 cm− 1. Singlet oxygen measurements. Photogeneration of singlet oxygen [1O2] by fullerene derivatives and reference photosensitizers was measured by direct detection of its steady-state phosphorescence in the range 1200–1350 nm in a 1O2 spectrometer described previously

TBARS assay. The modified thiobarbituric acid reactive substance (TBARS) assay was used for lipid peroxidation monitoring (Hong et al., 2000). Samples assigned to the TBARS assay were thawed. Twenty microliters of 4.4 mM butylated hydroxytoluene was added to each cell suspension, followed by 10.2 μl of 10 M NaOH. The samples were shaken at 60 °C for 30 min. After cooling down, 170 μl of a mixture containing 38% trichloroacetic acid and 2% KI was added to each sample, which was then mixed and placed on ice for 10 min. Then the samples were centrifuged at 7000g for 10 min at 4 °C. A 350 μl aliquot of each supernatant was transferred to a new tube containing the same volume of 0.66% 2-thiobarbituric acid. The samples were mixed and incubated at 95 °C for 30 min, then allowed to cool to room temperature. Absorbance was then registered at 534 nm (εMDA = 157,000 M− 1 cm− 1) in the HP spectrophotometer using water as a reference. Protein determination. Protein concentration was determined using a bicinchoninic acid (BCA) protein assay kit (Pierce, Rockford, IL). The assays were carried out in 96-well plates. Absorbance of the samples was measured at 560 nm in the GENios plate reader. Bovine albumin was used as a standard. Preparation of solutions and suspensions of fullerene derivatives for photophysical studies. Fullerene and fullerene hydride were stirred intensively for 3 days at room temperature in 3 ml of acetonitrile or D2O placed in a glass vial. Then they were stored at room temperature for 2 days to precipitate. Decanted supernatants were used for further measurements. Fullerol samples were prepared in various organic and aqueous solvents in the same way as the other two fullerene derivatives. Additionally, a fullerene (C60) suspension in H2O was prepared by sonication according to a modified procedure described previously (Andrievsky et al., 1995; Zhao et al., 2008b). Briefly, one mg/ml C60 solution in toluene was added to a 10-times larger volume of H2O. The mixture was sonicated (Ultrasonic Homogenizer 4710 Series, Cole-Parmer instruments Co., Chicago, IL) at room temperature for 15-min intervals to avoid rapid evaporation of the organic phase. The mixture was sonicated until all the toluene evaporated and an almost clear brown water solution was formed. Then the fullerene suspension was additionally evaporated under vacuum at 37 °C to remove all traces of toluene. The fullerene suspension was filtered under vacuum through a flask filter containing 0.2 μm cellulose acetate membrane. A clear yellow solution of crude C60 was concentrated in 15-ml portions in a centrifugal filter device (Amicon Ultra 15, 30,000 MWCO) by centrifugation at 754g and 23 °C until there was ∼ 1 ml of concentrated solution left. To determine fullerene concentration, 50 μl of the C60 suspension was diluted with 950 μl H2O and 400 μl of 0.1 M Mg(ClO4)2 and extracted into 1 ml toluene. The fullerene

Fig. 3. Dark (A–C)- and visible light (A and B)-induced cytotoxicity of fullerol in hRPE cells. The hRPE cells were incubated in the dark in 0–50 μM fullerol solution in HBSS/ DMSO (99:1) and exposed to visible light (A and B) for 60 min. Metabolic activity, lactate dehydrogenase release from the cells, and cell viability were determined with MTS, LDH, and trypan blue exclusion assays, respectively. Percentage of (A) metabolically active cells, (B) LDH release from the cells, and (C) cell viability were normalized to the control cells: (A) incubated in HBSS/DMSO (99:1) without fullerol and stored in the dark, (B) treated in the same way as measured cells but lysed directly before the assay, and (C) incubated in medium/FBS (9:1). Values are expressed as mean ± SD (n = 6). ⁎p b 0.001, ⁎⁎p b 0.01 comparing corresponding values in each pair of samples (irradiated vs. dark). †p b 0.001, ††p b 0.002, †††p b 0.005 comparing results obtained for cells treated with fullerol vs. controls (0 μM fullerol) within each group of cells.

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(Hall and Chignell, 1987; Bilski and Chignell, 1996). Samples were irradiated with an Oriel 500 W Hg lamp operating at 300 W. For general detection of 1O2 production the light was passed through a broad-band filter transmitting UV at 260–390 nm and for determination of quantum yield of its photogeneration it was passed through an interference filter with maximal transmittance at 365 nm. Recorded 1O2 phosphorescence spectra were normalized to the same number of absorbed photons (Bilski et al., 1996) and the quantum yield was determined using perinaphthenone as a reference photosensitizer (Schmidt et al., 1994; Marti et al., 1996; Oliveros et al., 1999; Beeby and Jones, 2000; Mitzel et al., 2004). The quantum yield of singlet oxygen photoproduction of perinaphthenone in 5 mM deuterated phosphate buffer in D2O (pD 7.5) was determined as Φ = 0.102 ± 0.004 using Rose Bengal as a reference (Redmond and Gamlin, 1999). Statistical analysis. Data in graphs in Figs. 2–3 and 5–7 are presented as the mean ± SD of three to six experiments. All p values were calculated using the ANOVA test.

Results Physiochemical properties Fullerols are capable of hydrogen bonding to form large aggregates (Figs. 1A and B). We found that the average size of fullerol particles (Table 1, Figs. 1C–F) was greatest in water, suggesting the highest aggregation, while there was less aggregation in cell culture medium containing serum proteins and in buffer and NaCl solutions. Water supports the most hydrogen bonding (Fig. 1B), while buffer and the NaCl solution would prevent some of the hydrogen bonding. Serum in the medium has been shown to decrease aggregation. There was a net zero charge on the fullerol in all solvents, indicating that there is little or no surface charge at neutral pH for fullerol; this result is in agreement with Sayes et al. (2007). The exception is medium with serum. Serum has been shown to interact with nanoparticles to change the size and surface charge (Murdock et al., 2008). We determined the particle sizes of three fullerene derivatives in acetonitrile and D2O (Table 2). In acetonitrile, the sizes of all three

Fig. 4. Apoptosis and necrosis in hRPE cells induced by fullerol. The cells were pretreated in the dark with 0–10 μM fullerol solution in HBSS/DMSO (99:1) for 17 h and stained with propidium iodide (PI) and Annexin V–FITC. Cell viability, apoptosis, or necrosis of each cell was monitored using flow cytometry.

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incubation in the dark, fullerol was observed to accumulate in hRPE cells (Figs. 2A–C). Uptake from a 10 μM solution of fullerol was indicated by a distinct yellow color appearing in the microscopic image. At this concentration the cells became slightly rounded. There was an increase in intensity of this yellow color with uptake from a 50 μM solution of fullerol, accompanied by increased damage to the cells. We determined quantitative uptake of fullerol in hRPE cells by its absorbance at 405 nm (Fig. 2D). When the cells were incubated in 50 μM solution, the amount of fullerol accumulated in the cells increased with time, reaching a plateau after ∼16 h. Cell viability

Fig. 5. Caspase-3 activation in hRPE cells pretreated with fullerol and exposed to visible light. Caspase-3 activity was determined in cytosolic extracts with an ApoAlert Caspase Fluorescent Assay Kit, where DEVD-AFC fluorescent substrate was used and normalized to protein content in the cells. Values are expressed as mean ± SD (n = 4). ⁎p b 0.01, ⁎⁎p b 0.1 comparing results obtained in cells incubated with fullerol and irradiated vs. the fullerol-treated cells incubated in the dark. The differences in caspase-3 activity between fullerol-treated cells vs. control cells were statistically significant at p b 0.01 (†) and p b 0.1 (††) for 20 and 50 μM fullerol, respectively, within the group of cells exposed to light.

compounds were in the nanoscale range. Fullerene and fullerol were of similar sizes – 291 and 282 nm in diameter, respectively – while fullerene hydride particles had a mean diameter of 364 nm. A more significant difference was observed in D2O. The size of fullerene particles obtained by sonication of C60 was 173 nm. We could not detect fullerene hydride particles in D2O. A fullerol suspension in D2O was composed of particles of three different diameters of 56, 347, and 5098 nm, with the medium-sized ones the most abundant. Accumulation in cells Overnight incubation of hRPE cells in HBSS containing 1% DMSO had no significant effect on cell density and morphology. During

We measured the metabolic activity and lactate dehydrogenase release of hRPE cells in the presence of 0–50 μM fullerol, either maintained in the dark or exposed to visible light. After a 17-h dark exposure, fullerol at concentrations of up to 5 μM had little effect on cells, as measured by the MTS, LDH, and trypan blue assays (Figs. 3A–C). However, at concentrations of 10–50 μM, fullerol was cytotoxic toward hRPE cells. When cells preincubated with fullerol for 17 h were exposed to visible light (N400 nm), phototoxicity, as measured by the MTS and LDH assays, was evident at concentrations of 10–50 μM (Figs. 3A and B). The clear differences in these data indicate that fullerol induces the greatest damage in RPE cells in the presence of light. Apoptosis Fullerol was seen to specifically bind to hRPE cells and cause their apoptosis and necrosis (Fig. 4). In the fluorescence dot plot histogram of Annexin V/PI stained cells, the lower left quadrant shows normal viable cells, which are negative for both Annexin V and PI; the lower right quadrant shows early-apoptotic cells, which are positive for Annexin V; the upper left quadrant shows necrotic cells, which are positive for PI; while the upper right quadrant shows late-apoptotic cells, which are positive for both Annexin V and PI (Martin et al., 1995; Reno et al., 1998). Fullerol was seen to induce apoptosis and necrosis in the hRPE cells when they were pretreated with the fullerene derivative at concentrations of 2–10 μM (Fig. 4). Incubation in the dark with 50 μM fullerol caused activation of caspase-3 in hRPE cells (Fig. 5). The protease activity was enhanced by visible light exposure. A significant increase in the enzyme activity was observed in irradiated cells preincubated with fullerol at concentrations higher than 20 μM. TBARS formation Dark incubation of hRPE cells with 50 μM fullerol led to TBARS formation (Fig. 6). Additionally, visible light exposure of the cells enhanced TBARS generation by a factor of 2–3 when hRPE cells were preincubated in 10–50 μM fullerol solution. Increased TBARS production induced by light was correlated with fullerol concentration in hRPE cells. Inhibition of fullerol phototoxicity to hRPE cells

Fig. 6. TBARS production in hRPE cells induced by fullerol. The cells were incubated for 17 h in the dark in 2–50 μM fullerol solutions in HBSS/DMSO (99:1, v/v). Control cells were incubated in the same solvent mixture without fullerol. One set of cells was exposed to light (λ N 400 nm) for 60 min while the other was stored in the dark. After irradiation the cells were incubated for 2 h at room temperature before the TBARS assay. Values are expressed as mean ± SD (n = 3). ⁎p b 0.001, ⁎⁎p b 0.05 comparing results obtained in cells incubated with fullerol and irradiated vs. the fullerol-treated cells incubated in the dark. The differences in TBARS concentration between fulleroltreated cells vs. control cells were statistically significant at p b 0.005 (††) for 10 μM fullerol and p b 0.001 (†) for 20 and 50 μM fullerene derivatives within the group of cells exposed to light and p b 0.01 (†††) for 50 μM fullerol within the group of cells kept in the dark.

To identify potential non-toxic antioxidants that might prevent phototoxic damage to hRPE cells, we preincubated the cells with lutein and the glutathione mimic N-acetyl-L-cysteine (NAC) – the quenchers endogenous to the human retina. When 10 or 20 μM lutein was added 2 h prior to incubation of the cells with fullerol and the cells were irradiated, phototoxic damage decreased (Fig. 7). However, NAC at concentrations of 0.5–1.0 mM did not show a significant protective effect on metabolic activity or cell membrane damage in the hRPE cells.

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Fig. 7. The effect of lutein and N-acetylcysteine (NAC) on fullerol phototoxicity in hRPE cells. The cells were incubated with lutein or NAC in HBSS/DMSO (99:1) for 2 h and with 20 μM fullerol in HBSS/DMSO (99:1) for the next 17 h. Then the cells were washed and overlaid with HBSS and exposed to visible light for 60 min. After irradiation, the cells were incubated for 24 h in medium/serum (9:1). Metabolic activity (A and C) and lactate dehydrogenase release (B and D) from the cells were determined with MTS and LDH assays, respectively. Results are expressed as mean ± SD (n = 4). ⁎p b 0.001, ⁎⁎p b 0.002 comparing metabolic activity or LDH release in cells incubated with an antioxidant and fullerol vs. cells incubated only with fullerol in both the “dark” and “light exposed” groups.

Singlet oxygen photogeneration All three fullerene derivatives photogenerated singlet oxygen in acetonitrile (Table 2), while only fullerene and fullerol did so in D2O. In acetonitrile, the 1O2 phosphorescence spectrum of the fullerene hydride and fullerene solutions was characteristically red-shifted (Zhao et al., 2008b) compared to fullerol. A similar effect was observed in D2O. The quantum yield of 1O2 photoproduction by fullerene was highest (0.9) in acetonitrile, with a much lower value for fullerol (0.02) and negligible production by fullerene hydride (0.004). In D2O it was less efficient for fullerene (0.7), for fullerol similar to the value in acetonitrile within experimental error, and below detectable levels for fullerene hydride. We also determined the singlet oxygen quantum yield of fullerol in both organic and aqueous solvents (Table 3). The highest value of the quantum yield was detected in tetrahydrofuran (0.139) while the

Table 1 DLS determination of particle size (Z-average) and surface charge (Zeta potential) of 50 μM fullerol in different solutions (DMSO: 1%). Sample Fullerol Fullerol Fullerol Fullerol

in in in in

cell culture medium 5 mM PBS deionized H2O 50 mM NaCl

Z-Average (d, nm)

Zeta potential (mV)

224.9 248.8 552.4 219.6

2.91 0.14 1.37 0.04

lowest was in 5 mM deuterated phosphate buffer (pD 7.6) in D2O (0.002). There was an observed relationship between particle size and 1 O2 photoproduction. The 1O2 quantum yield rose from 0.014 to 0.139 and fullerol particle size dropped simultaneously from 516 to 1.85 nm in CH2Cl2 and THF, respectively. The highest yields were observed in solvents with the highest fullerol solubility. In solvents that allow for the formation of larger fullerol particles, ethanol, D2O, and buffer, the singlet oxygen production decreased dramatically. Interestingly, the 1 O2 quantum yield in D2O was more than 20 times higher than in phosphate buffer (pD 7.6) prepared in D2O. Although singlet oxygen photoproduction of fullerol in acetonitrile (0.032) was much lower than that of fullerene (0.934), the former rate is of the same order of magnitude as that of lipofuscin (0.09) (Różanowska et al., 1998), the endogenous photooxidizing agent in the human retina (Roberts et al., 2002). Discussion In order to assess the potential in vitro toxicity of a particular nanoparticle, the particles must be accurately defined (Murdock et al., 2008; Kroll et al., 2009). In this study we determined the size, aggregated state and zeta potential of fullerol in physiologically relevant solutions. As seen in Figs. 1A and B, fullerol can form aggregates through hydrogen bonding of its hydroxyl group in polar solvents. TEM images indicate that all samples of fullerol (Figs. 1C–F) are aggregated. However, DLS (Table 1) shows that the highest aggregation is found when these nanoparticles are dispersed in water,

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Table 2 Particle size determined with DLS and quantum yield of singlet oxygen production by fullerene and its derivatives irradiated with UVA at λ = 365 nm in acetonitrile and D2O. Solvent

Acetonitrile

D2O

Fullerene derivative

A365

Fullerene Fullerol Fullerene hydride Fullerene (5 μM) Fullerol

0.300 0.301 0.300 0.109 0.105

Fullerene hydride

0.000

Particle size (nm)

291 ± 29 282 ± 25 364 ± 18 173 ± 78 56.3 ± 8.3 347 ± 21 5098 ± 102 b.d.l.

λImax (nm)

Integration value of 1O2 phosphorescence spectrum (a.u.) Fullerene derivative

Perinaphthenone

1284 1274 1278 1283 1276

29.2 ± 1.6 0.64 ± 0.04 0.12 ± 0.01 3.17 ± 0.25 0.33 ± 0.09

31.3 ± 1.8 32.4 ± 2.1 29.2 ± 2.4 4.44 ± 0.34 4.36 ± 1.10

b.d.l.

b.d.l.

Quantum yield of 1O2 photoproduction

0.934 ± 0.075 0.020 ± 0.002 0.004 ± 0.001 0.700 ± 0.077 0.073 ± 0.028

Quantum yield was determined using perinaphthenone as a reference photosensitizer. Fullerene solution in D2O was prepared by sonication. λImax, wavelength of maximal intensity of 1O2 phosphorescence spectrum; b.d.l., below detection level.

and this aggregation was decreased in buffer and medium with serum. In our previous studies (Roberts et al., 2008) we found that particle size decreases when fullerol binds to lens protein. Murdock et al. (2008) found that a number of nanoparticles showed decreased aggregation in the presence of serum proteins. In previous studies we have noted that the state of aggregation of fullerene derivatives affects their toxicity. Monomeric nanoparticles were more phototoxic than aggregated nanoparticles (Zhao et al., 2008a, 2009). As seen in Table 1, the zeta (surface charge) measurements showed that the fullerols dissolved in medium with serum had a small but significant net positive charge. There was a net zero charge on the fullerol in all other solvents, indicating that there is little or no surface charge at neutral pH for fullerol; this result is in agreement with Sayes et al. (2007). The hydrogen atom in the hydroxyl group is not labile at physiological pH, but there may be some interactions with the serum proteins in the media. Human RPE cells incubated with 10–50 μM fullerol but kept in the dark showed decreased metabolic activity and increased lactate dehydrogenase release, indicating a cytotoxic response. However when the RPE cells were irradiated with a dose of 8.5 J·cm− 2 visible light, metabolic activity was decreased and lactate dehydrogenase release was apparent in the presence of a much lower concentration of fullerol (b10 μM). Fullerol, both with and without irradiation by visible light, induced early apoptosis, indicating membrane damage. Photoactivated fullerol increased the production of TBARS, a measure of lipid peroxidation. Therefore the cytotoxic and phototoxic damage of fullerol to hRPE cells is due at least in part to an increase in damage to the cell cytoplasmic membrane and induction of lipid peroxidation. Enhanced lipid peroxidation has been seen with fullerol in other cell lines (Sayes

et al., 2004, 2005; Isakovic et al., 2006) and in vivo (Oberdőster et al., 2005; Sayes et al., 2007). We did not examine the cyto- and phototoxicity of fullerene and fullerene hydride because they are not water-soluble and thus would not be incorporated into the same compartments of the retinal pigment epithelial cells. Furthermore, they are not being considered for drug delivery to the retina. Fullerol and fullerene phototoxicity were compared in previous studies performed in our lab on dermal cells (keratinocytes) (Zhao et al., 2008a) since there is good evidence of in vivo dermal phototoxicity of all fullerenes. The level of oxygen in the retina is sufficiently high for photooxidation to induce the phototoxic effects seen in the cells. The partial pressure in the vitreous humor is much lower than in the blood vessels and is at the level of 7.5 mm Hg, which equals ∼ 1% pO2 (Shui et al., 2009). However, RPE cells are more saturated with oxygen than is the vitreous humor. Human RPE cells form a single-cell layer that is the outermost part of the retina. The RPE layer is located nearest the choroid, which is rich in blood vessels. The choroid supplies RPE cells with nutrients and oxygen. Measurements of oxygen diffusion through the RPE and the retina showed that pO2 level is relatively high, ∼ 70 mm Hg in a rabbit (Yu et al., 2005), and decreases linearly with distance from the choriocapillaris to the inner portion of the photoreceptors (Spaide, 2005). Moreover, there is an additional layer of blood vessels between the vitreous humor and the upper level of the retina, which supplies the neural retina. Thus the oxygen saturation is high enough in the retina and particularly in RPE to allow fullerol, when it is present there, to produce reactive oxygen species efficiently. Photooxidative damage to cells can occur through a type I (free radical and superoxide) and/or type II (singlet oxygen) mechanism. The exact reactive oxygen species and/or free radical intermediate for

Table 3 Fullerol particle size and quantum yield of singlet oxygen production by the fullerene derivative irradiated with UVA at λ = 365 nm in selected solvents. Solvent

Particle size (nm)

Dichloromethane Chloroform Acetone Acetonitrile DMSO (5 μM) Tetrahydrofuran 1,4-Dioxane Ethanol D2O

516 ± 63 410 ± 85 268 ± 76 173 ± 36 2.42 ± 0.41 1.85 ± 0.24 2.95 ± 0.55 57 ± 17 412 ± 120

5 mM NaPi in D2O (pD 7.6)

37.0 ± 6.5 359 ± 125

Integration value of 1O2 phosphorescence spectrum (a.u.) Fullerol

Reference photosensitizer

1.04 ± 0.08 1.82 ± 0.22 1.83 ± 0.16 0.43 ± 0.03

PN PN PN PN

71.0 ± 5.4 49.8 ± 5.9 23.1 ± 2.0 13.2 ± 0.9

1.92 ± 0.16 2.65 ± 0.15 1.03 ± 0.06 1.04 ± 0.08

PN PN PN PN RB PN

13.3 ± 1.2 19.9 ± 1.2 7.8 ± 0.5 3.1 ± 0.8 3.4 ± 0.7 15.8 ± 1.9

0.26 ± 0.03

Perinaphthenone (PN) and Rose Bengal (RB) were used as reference photosensitizers for determination of quantum yield of 1O2 photogeneration. PN, perinaphthenone; RB, Rose Bengal; n.d., not determined; NaPi, deuterated sodium phosphate buffer.

Quantum yield of 1O2 photoproduction 0.014 ± 0.001 0.036 ± 0.006 0.079 ± 0.010 0.032 ± 0.003 n.d. 0.139 ± 0.017 0.133 ± 0.011 0.128 ± 0.034 0.024 ± 0.012 0.023 ± 0.011 0.002 ± 0.001

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the phototoxicity of fullerol on RPE cells is not clear. Previous reports have shown that fullerol produces superoxide (Zhao et al., 2008a), and these present studies indicate that fullerol is a poor generator of singlet oxygen with a quantum yield of Φ = 0.05 in D2O and a range of 0.002– 0.139 in various solvents. On the other hand, fullerol phototoxicity induced in human lens epithelial cells and keratinocytes was prevented by quenchers of singlet oxygen (azide, lutein) (Zhao et al., 2008b; Roberts et al., 2008). Lutein is a moderate quencher of singlet oxygen and also quenches free radicals efficiently (Cantrell et al., 2003). We did not see quenching of photo- or cytotoxicity with the reductant and glutathione mimic N-acetylcysteine. However, photochemical properties of fullerol are mediated by another reductant, NADH (Zhao et al., 2008a). These ESR studies showed that NADH enhances superoxide anion production by fullerol when irradiated at wavelengths longer than 300 nm. NADH also increased the rate of oxygen photoconsumption by irradiated fullerol. Addition of NaN3, which is a specific quencher of singlet oxygen, to the fullerol and the reductant mixture only slightly decreased the rate of oxygen uptake by fullerol. However when catalase was added instead of azide, slower oxygen photoconsumption was observed, suggesting that hydrogen peroxide was present. Previous studies have determined that damage from some fullerene derivatives was prevented by ascorbate in human dermal fibroblasts (Sayes et al., 2005). However, we did not see a protective effect against fullerol photodamage by ascorbate either in human lens epithelial cells (Roberts et al., 2008) or in the present study with human retinal pigment epithelial cells. Although the ascorbate (AscH) concentration in the vitreous body (∼ 2 mM) (Varma, 1987; Hanashima and Namiki, 1999) is much higher than in blood plasma (22–57 μM) (VanderJagt et al., 1987; Block et al., 1999; Ayaori et al., 2000; Varma, 1987), its concentration decreases significantly with distance within the retina from 20.6 mg/dl (1.17 mM) in retinal cytosol to 5.8 mg/dl (0.33 mM) in RPE cytosol (Lai et al., 1986). Since the RPE is the most remote layer of the retina, ascorbate concentration is much lower in the RPE than in the vitreous body. Thus the ascorbate quenching effect of 1O2 in RPE cells would be significantly lower than in the vitreous body. We conducted further photophysical studies in order to define the effect of various environmental factors on the production of singlet oxygen of fullerene nanoparticles and its derivatives. All cells, including ocular cells, have compartments of different pH and different polarities, with the cytoplasm being the most polar and the membrane being the least. As seen in Table 2, in acetonitrile the quantum yield of singlet oxygen of fullerene derivatives changes with the chemical structure; the greater number of conjugated double bonds found in fullerene is reflected in the higher singlet oxygen yield. In comparison, the lower number of conjugated double bonds in fullerol significantly decreased the singlet oxygen yield, and the nearly complete loss of conjugation with fullerene hydride resulted in little or no singlet oxygen production. Here acetonitrile was chosen as the solvent because all three fullerene derivatives were soluble in it, allowing for a direct comparison. In the more polar D2O, fullerene solubility is decreased and its singlet oxygen production is increased. For fullerol, there is a distribution of different size particles in D2O with little significant change in singlet oxygen production. Fullerene hydride is not soluble in D2O. We next examined the effect of solvent polarity on fullerol's singlet oxygen quantum yield (Table 3). The solvents are listed from most hydrophobic (dichloromethane) to most hydrophilic (D2O and D2O buffer). Initially, as hydrophilicity increased, particle size decreased, with a size corresponding to monomers in THF and dioxane. This decrease indicated better solubility and lower fullerol aggregation in the more hydrophilic solvents, with a concomitant increase in singlet oxygen quantum yield. However, highly hydrophilic solvents (e.g., D2O) allow hydrogen bonding to the OH groups in fullerol, leading to aggregation with a concomitant decrease in singlet oxygen produc-

tion. We could not determine the singlet oxygen quantum yield of fullerol in DMSO because of high solvent viscosity. (At high viscosity, the oxygen molecule cannot move away quickly enough to remove energy from the excited fullerol particle.) We also examined the effect of pH on singlet oxygen production. D2O has a pD of 5; the phosphate buffer was at pD 7.6. At the higher pD, there was better solubilization of fullerol (Vileno et al., 2006) with a smaller particle size, but there was a dramatic decrease in singlet oxygen production. At the higher pD, the H on the fullerol OH groups is labile, leading to easier electron transfer and increased production of superoxide anion (Pickering and Wiesner, 2005; Badireddy et al., 2007; Zhao et al., 2008a). We determined the singlet oxygen production quantum yield of fullerol in several solvent systems. Fullerol is not only water soluble but can be solubilized in a wide spectrum of solvents (Alvez et al., 2006; Bogdanović et al., 2004; Da Ros and Prato, 1999; GuiradoLoópez and Rincón, 2006; Pickering and Wiesner, 2005; Zhao et al., 2008a). We found that although the production of singlet oxygen by fullerol is low with a range of 0.002–0.139 we could detect it in several solvent systems of different polarities. This indicates that fullerol can produce singlet oxygen not only in hydrophilic environments of the cytoplasm but also in hydrophobic environments such as the cell membrane (Vileno et al., 2006). The production of singlet oxygen by fullerol is not in the range of an efficient photosensitizer used in photodynamic therapy of the retina, for instance, hypericin (quantum yield = 0.73) (Racinet et al., 1988), but it is sufficient to do phototoxic damage to the retina. As has been seen with the endogenous agent for blue light damage to the retina, lipofuscin (quantum yield = 0.09) (Różanowska et al., 1998) and indocyanine green (quantum yield = 0.08) (Wu et al., 2005; Kassab, 2002), a dye used with vitreoretinal surgery, substances with very low quantum yields of singlet oxygen can permanently damage the human retina. Human health implications We have determined that fullerol is only mildly cytotoxic in the dark but significantly phototoxic to human retinal pigment epithelial cells in the presence of visible light. This phototoxic damage induces oxidative stress and lipid peroxidation in hRPE cells in vitro. Lipid peroxidation is an important risk factor in the induction of retinal and choroidal neovascularization (wet macular degeneration) and diabetic retinopathy (Hardy et al., 2005). The products of lipid peroxidation (Ayalasomayajula and Kompella, 2002) increase expression and secretion (Lu et al., 1998; Treins et al., 2001; Lee et al., 2004) of the angiogenesis growth factor VEGF by RPE cells. Increased expression of angiogenic growth factors has been shown to lead to age-related maculopathy (Kliffen et al., 1997). Fullerol produces both singlet oxygen and superoxide. Human retinal tissues are damaged by photooxidation through both type I (superoxide and other free radicals) and type II (singlet oxygen) mechanisms. We found that the phototoxic damage to hRPE cells could be partially quenched by lutein, and this compound can be supplemented in vivo to the human retina (Bernstein et al, 2001; Richer et al., 2004; van de Kraats et al., 2008) for protection against light damage. Conclusion Exposure to fullerols, particularly in the presence of sunlight, may lead to retinal damage. Although the acute toxicity of water-soluble nano-C60 is low, these compounds are retained in the body for long periods, raising concern for their chronic toxic effect. Conflict of interest statement The authors declare that there are no conflicts of interest.

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Acknowledgments This research was supported by the Intramural Research Program of the NIH, National Institute of Environmental Health Sciences (NIEHS). We wish to thank the following persons from NIEHS: Dr. Ann Motten for help in preparation of this manuscript, Dr. Carl Bortner and Maria Sifre for their assistance with flow cytometry measurements, Ms. Deloris Sutton for the TEM images, and Dr. Piotr Bilski for his help in setting up the 1O2 detection system. Images of the cells were taken by C. Jeff Tucker at the NIEHS Fluorescence Microscopy and Imaging Center. We also wish to thank Dr. Cynthia Smith of National Toxicology Program, NIEHS for supplying the fullerol, fullerene and fullerene hydride used in these experiments and Drs. William Boyes and Laura Degn of the US Environmental Protection Agency Research (USEPA) for assistance with the Zeta measurements. This manuscript has been reviewed by the National Institutes of Environmental Health Sciences and approved for publication. 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