Toxicology and Applied Pharmacology 258 (2012) 226–236

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Toxicology and Applied Pharmacology

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In Vitro Phototoxicity and Hazard Identification of Nano-scale Titanium Dioxide☆

Kristen Sanders a,b,1, Laura L. Degn a, William R. Mundy c, Robert M. Zucker a, Kevin Dreher b,Baozhong Zhao d,f, Joan E. Roberts d,e, William K. Boyes a,⁎a Toxicity Assessment Division, National Health and Environmental Effects Research Laboratory, Office of Research and Development, U.S. Environmental Protection Agency,Research Triangle Park, North Carolina, USAb Environmental Public Health Division, National Health and Environmental Effects Research Laboratory, Office of Research and Development, U.S. Environmental Protection Agency,Research Triangle Park, North Carolina, USAc Integrated Systems Toxicology Division, National Health and Environmental Effects Research Laboratory, Office of Research and Development, U.S. Environmental Protection Agency,Research Triangle Park, North Carolina, USAd Laboratory of Toxicology and Pharmacology, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, North Carolina, USAe Fordham University, New York, New York, USAf National Center for Nanoscience and Technology, Beijing, China

☆ This manuscript has been reviewed by the Nationalfects Research Laboratory, U.S.E.P.A. and approved fornames and commercial products does not constitute endfor use.⁎ Corresponding author at: B105-05, Toxicity Assessm

tal Protection Agency, Research Triangle Park, NC 27711E-mail address: boyes.william@epa.gov (W.K. Boyes

1 Present address.

0041-008X/$ – see front matter © 2011 Elsevier Inc. Alldoi:10.1016/j.taap.2011.10.023

a b s t r a c t

a r t i c l e i n f o

Article history:Received 8 April 2011Revised 21 October 2011Accepted 24 October 2011Available online 15 November 2011

Keywords:NanoparticlesTitanium dioxidePhototoxicityRetinal pigment epithelium

Titanium dioxide nanoparticles (nano-TiO2) catalyze reactions under UV radiation and are hypothesized tocause phototoxicity. A human-derived line of retinal pigment epithelial cells (ARPE-19) was treated withsix samples of nano-TiO2 and exposed to UVA radiation. The TiO2 nanoparticles were independently charac-terized to have mean primary particle sizes and crystal structures of 22 nm anatase/rutile, 25 nm anatase,31 nm anatase/rutile, 59 nm anatase/rutile, 142 nm anatase, and 214 nm rutile. Particles were suspendedin cell culture media, sonicated, and assessed for stability and aggregation by dynamic light scattering.Cells were treated with 0, 0.3, 1, 3, 10, 30, or 100 μg/ml nano-TiO2 in media for 24 hrs and then exposed toUVA (2 hrs, 7.53 J/cm2) or kept in the dark. Viability was assessed 24 hrs after the end of UVA exposure bymicroscopy with a live/dead assay (calcein-AM/propidium iodide). Exposure to higher concentrations ofnano-TiO2 with UVA lowered cell viability. The 25 nm anatase and 31 nm anatase/rutile were the most pho-totoxic (LC50 with UVAb5 μg/ml), while the 142 nm anatase and 214 nm rutile were the least phototoxic. Anacellular assay ranked TiO2 nanoparticles for their UVA photocatalytic reactivities. The particles were foundto be capable of generating thiobarbituric acid reactive substances (TBARS) under UVA. Flow cytometryshowed that nano-TiO2 combined with UVA decreased cell viability and increased the generation of reactiveoxygen species (ROS, measured by Mitosox). LC50 values under UVA were correlated with TBARS reactivity,particle size, and surface area.

© 2011 Elsevier Inc. All rights reserved.

Introduction

Engineered nanomaterials, as defined by the National Nanotechnolo-gy Initiative (National Science and Technology Council, 2007), are rapidlybeing developed and deployed in a variety of commercial applications.For the safe use and application of titanium dioxide nanomaterials(nano-TiO2), it is important to define the critical parameters that mightinfluence their impact on health and the environment. Among themore commonly considered properties evaluated in nanotoxicity studiesare particle size, surface area, composition, reactivity, coatings, stability,

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and propensity to agglomerate or aggregate in different media (Balbuset al., 2007; Oberdorster et al., 2005). An additional property of concernfor nano-TiO2 is photoreactivity.

Titanium dioxide is one of the most frequently used engineerednanomaterials because it can serve two distinct functions: absorbingand scattering ultraviolet (UV) radiation and/or semiconductorphotocatalysis activated by UV radiation (Fujishima et al., 2000;Kwon et al., 2008; Mills and Le Hunte, 1997). As a UV reflectingagent, it is found in a variety of applications, including sunscreens, paints,and coatings, where it serves in a protective role (Klaine et al., 2008). Thephotocatalytic ability of nano-TiO2 is used in photovoltaic devices andself-cleaning or self-sterilizing product coatings (Kwon et al., 2008;WoodrowWilson International Center for Scholars, 2011). Additionally,as a photocatalytic agent, nano-TiO2 could be useful for degrading envi-ronmental contaminants such as organic compounds (Muneer et al.,2002; Yeber et al., 2000; Yeo and Kang, 2006) and pesticides (Muneerand Bahnema, 2001; Zoh et al., 2005; Zoh et al., 2006), decontaminatingindoor air (Salthammer and Fuhrmann, 2007), or sterilizing surfaces or

227K. Sanders et al. / Toxicology and Applied Pharmacology 258 (2012) 226–236

pathogen-contaminated drinking water (Amezaga-Madrid et al., 2002;Paul et al., 2007; Prasad et al., 2009). The two common crystalline formsof nano-TiO2, rutile and anatase, are found in commercially availablenano-TiO2 and are considered photocatalytic, with anatase being themore photoreactive (Kwon et al., 2008).

However, the same properties that make nano-TiO2 a photocatalyticagent may also lead to phototoxic reactions in humans. Absorption ofUVA radiation activates a surface electron of the TiO2 molecule to theconduction band, leaving a valence band hole that extracts electronsfromwater or hydroxyl ions, generating hydroxyl radicals. The electronsof the hydroxyl radicals then reduce O2 to produce superoxide anion,which leads to a cascade of reactive oxygen species (ROS) (Brunet etal., 2009; Kwon et al., 2008). UVA radiation can also induce the produc-tion of highly reactive singlet oxygen (Konaka et al., 2001). The genera-tion of an ROS cascade can damage tissues through oxidation of nearbylipids, proteins, nucleic acids, or other biomolecules (Straight andSpikes, 1985).

Nano-TiO2 has been shown to be phototoxic to isolated DNA(Hirakawa et al., 2004), bacteria (Adams et al., 2006), and several aquat-ic species (Hund-Rinke and Simon, 2006;Matsuo et al., 2001), aswell asmammalian lymphoma (Nakagawa et al., 1997), ovary (Uchino et al.,2002), dermal fibroblast, and pulmonary epithelial carcinoma celllines (Sayes et al., 2006). Because skin and eye tissues are exposed tolight, they are typically of greatest concern for phototoxic damage. Inthe skin, phototoxicity can lead to sunburn or an increased risk of cancer(Johnson and Ferguson, 1990). In the eye, phototoxic reactions candamage the lens, retina, or retinal pigment epithelium (RPE), leadingover time to cataracts in the lens or photoreceptor loss in the retina(Roberts, 2002).

To our knowledge, the potential for nano-TiO2 phototoxicity in ocularcells has not been previously reported. However, models of human lens(Roberts et al., 2008) and RPE cells have been sensitive to phototoxic ef-fects of other nanomaterials, specifically to hydroxylated fullerene(Nohynek et al., 2008; Wielgus et al., 2010). In addition, cultured RPEcells internalize TiO2 nanoparticles and combine them into perinuclearaggregates in a dose-related fashion (Zucker et al., 2010). In the studiesreported in this communication, the potential phototoxicity of severaltypes of nano-TiO2 of various particle sizes and crystalline formswas eval-uated in a culture of human-derived RPE cells. The hypotheses tested in-cluded: 1) that nano-TiO2 toxicity would be enhanced by UVA radiation,2) that smaller particleswould bemore phototoxic than larger particles,and 3) that the anatase crystal formwould be themost phototoxic,withmixed anatase/rutile being intermediate and rutile samples being leastphototoxic. It was found that: 1) nano-TiO2 was more toxic followingexposure to UVA radiation, 2) smaller particles tended to be moretoxic than larger ones, and 3) the effect of crystal structure on phototox-icity could not be determined in these experiments. In addition, it wasobserved that the generation of reactive oxygen species with nano-TiO2 under UVA radiation correlated with cytotoxicity.

Materials and Methods

ARPE-19 cells. A human-derived RPE cell line (ARPE-19) was obtainedfrom ATCC (Manassas, Virginia) and grown in a 1:1 combination ofDulbecco's Modified Eagle's Medium and Ham's F-12 Nutrient Mixture(DMEM/F-12; cat # 11039047, Invitrogen, Carlsbad, CA) with 10% fetalbovine serum (FBS; cat # 511150, Atlanta Biologicals, Lawrenceville,GA) and 1% penicillin/streptomycin (cat # 17-602E, Lonza, Walkersville,MD). Phenol red was omitted from the media due to its potential inter-action with UV radiation. All testing was done with cells at passage 28.

TiO2 nanoparticles. Six samples of nano-TiO2, which varied in crystalstructure and primary particle size, were evaluated. None of the particleswere coated. Dry TiO2 nanoparticles were characterized under a contractwith theUniversity of Kentucky (EPA contract #EP-D-08-074). Elementalanalysis was conducted by transmission electron microscopy, electron

dispersive X-rays, and inductively coupled plasma-atomic emission spec-trometry; surface area by the Brunauer-Emmett-Teller method; primaryand aggregate particle sizes by transmission electron microscopy(TEM); crystal structure by X-ray diffraction; and particle shapeand morphology by TEM and scanning electron microscopy. The pri-mary results of the characterization of the six nano-TiO2 samples arepresented in Table 1. An evaluation of free radical oxidative damageto DNA caused by these six samples has been published (Kitchin etal., 2010).

In thismanuscript, the particleswill be designated by theirmean par-ticle size and crystal structures as determined by the independent char-acterization: 22 nm anatase (catalog # 39953, Alfa Aesar, Wardfill, MA),25 nm anatase (catalog # 44690, Alfa Aesar, Wardfill, MA), 31 nm ana-tase/rutile (catalog # Aeroxide® TiO2 P25, Degussa, Alpharetta, GA),59 nm anatase/rutile (catalog # 5485HT, NanoAmor, Houston, TX),142 nm anatase (catalog # 21358; Acros, Morris Plains, NJ), and214 nmrutile (catalog#MKN-TiO2-R250,Mknano,Mississauga, Ontario,Canada).

For in vitro treatment, particles were suspended in cell culturemedia containing 10% FBS (Allouni et al., 2009). A stock suspensionof 1 mg/ml was prepared and then dispersed using a probe sonicator(Misonix Microson Ultrasonic Cell Disrupter XL, Farmingdale, NY)operating at 4.5 watts for 3 pulses, 2 seconds/pulse. This sonicationprotocol consistently dispersed the nanoparticles until the suspen-sion appeared homogenous to the naked eye. Other concentrationswere prepared by serially diluting the stock suspension in the samemedia. A new stock suspension wasmade from dry nano-TiO2 imme-diately before treating cells in each experiment. The suspensionswere relatively stable over time (for at least 20 minutes accordingto dynamic light scattering) and could readily be evaluated and appliedto the cells to be treated, as reported by others (Allouni et al., 2009).Samples of 25 nm anatase and 59 nm anatase/rutile were evaluatedusing dynamic light scattering (Malvern Zetasizer Nano, Westborough,MA) (Murdock et al., 2008). The following parameters were used:refractive index of anatase, 2.493; refractive index of rutile, 2.903(Weast et al., 1984); refractive index of dispersant, 2.3; viscosityof dispersant, 0.75 cP at 35 °C (Gallik et al., 1993). Measurementswere taken three times and checked for quality of data using thesoftware's correlograms and error reports.

Uptake and accumulation of nanoparticles in ARPE-19 cells. Two experi-ments were conducted to determine if nanoparticles would entercells and be retained over time. In a short-term experiment, ARPE-19cells were grown to confluence on a glass culture slide. The cellswere treated with 15 μg/ml 59 nm anatase/rutile TiO2 (2 ml) and ob-served under modulation contrast microscopy (Nikon Eclipse TE200inverted epifluorescent microscope). Images were acquired at 100xmagnification over the course of three hours after the start of exposureand again three days later.

In a second, long-term experiment, ARPE-19 cells were grown toconfluence in three T75 culture flasks. Flasks were treated with 0,10, or 30 μg/ml 59 nm anatase/rutile TiO2 (19 ml). All flasks were ob-served under modulation contrast microscopy intermittently from 1to 25 days post-exposure. Media was removed and replaced 2–3times weekly. Images were acquired as described for the short-term experiment at 100x and 200x magnification using a 10x or 20xobjective.

Dose-dependent uptake of nano-TiO2 by ARPE-19 cells was alsoobserved using dark field microscopy (Nikon E-800 with 60x PlanFluor iris adjustable lens, NA 0.5-1.25) and flow cytometry (Zuckeret al., 2010). Data from these experiments suggested that particleswere incorporated within cells by 24 hours, so this time point wasused in further studies assessing phototoxicity.

Phototoxicity. ARPE-19 cells were plated in T-75 culture flasks (ap-proximately 1×106 cells in 20 ml media) and incubated in the

Table 1TiO2 Nanoparticle Sample Designation and Physicochemical Properties.

Vendor Vendora CrystalStructure

Vendor Size(nm)

VendorPurity

Vendor SurfaceArea (m2/g)

Analyzedb CrystalStructure

Analyzed TEM Avg Sizeand (Size Range) (nm)

Analyzed Purity(EDS and ICPMS)

Analyzed SurfaceArea (BET) (m2/g)

Alfa Aesar Stock # 39953Lot # C27R043

anatase 32 99.9% 45 94% anatase/6% rutile

22 (9–61) 97.7% 49.8

Alfa Aesar Stock # 44690Lot # D22T034

anatase 10 notprovided

100 - 130 anatase 25 (6–60) 98.8% 118

Degussa Aeroxide P25Lot # 4165012298

86% anatase/14% rutitle

27.5 >99% 49 86% anatase/14% rutile

31 (12–88) 99.9% 52.9

Nanostructured andAmorphous Materials Inc.Stock # 5485HTLot # 5485-030007

rutile 30–40 95.1% >30 85% anatase/15% rutile

59 (36–97) 99.9% 22.2

Acros Organics Cat. # 21358Lot # A0075656

anatase 200 notprovided

not provided anatase 142 (67–322) 99.9% 6.99

Mknano MKN-TiO2-R250Lot # 459/2007

rutile 250 100.0% 6.8 rutile 214 (37–410) 98.7% 11.6

a TiO2 nanoparticle physicochemical properties provided by specific vendor.b TiO2 nanoparticle physicochemical properties determined by an independent analysis performed by the University of Kentucky.

228 K. Sanders et al. / Toxicology and Applied Pharmacology 258 (2012) 226–236

dark with an atmosphere of 5% CO2 and 95% air at 37 °C. After cellsreached confluence (3–4 days), they were trypsinized and split into24-well plates (1×105 cells/ml, 1 ml/well, 1.9 cm2 surface area).These plates were incubated for 24 hours.

Cells on each plate were treated by removal ofmedia from eachwelland replacementwith a 1 ml suspension of 0.3, 1, 3, 10, 30, or 100 μg/mlnano-TiO2. A 0 μg/ml condition (media only) was included on eachplate as a negative control. Over the course of the experiments, threereplicates of each concentration (including controls) for each samplewere treated. The plates were incubated for 24 hours post-treatmentto allow cells to incorporate TiO2 nanoparticles.

After incubation, without changing media, plates were exposed tovisible light, UVA radiation, or a dark condition. During the exposure,the cells were placed on heating blocks under the bulbs to stabilizetheir temperature at 37 °C. The spectra and irradiance of the visibleand UVA bulbs were recorded using SpectrILight™ III software and anILT900 spectroradiometer (International Light Technologies, Peabody,MA). “Visible” plates were exposed to visible light (Daylight (6500 K)Model S, LED Supply, Randolph, VT) for 4 hours for a total irradianceof 17.9 J/cm2. The spectrum of visible light ranged from approximately434 to 650 nm and was bimodal with peaks at 452 and 540 nm.“UVA” plates were placed under UVA bulbs (catalog # F20T12/BL/HO(UVA), National Biological Corporation, Beachwood, OH) for 2 hoursfor a total UVA irradiance of 7.5 J/cm2. The levels of visible and UVA ra-diation employed were based on prior phototoxicity experiments (Heet al., 2004; Roberts et al., 2008) and were consistent with Organizationfor Economic Co-operation and Development (OECD) guidance on pho-totoxicity. This guidance states that the highest levels of radiation thatalone do not cause cytotoxicity in the test system should be employed(OECD, 2004). The UVA spectrum extended from approximately 330to383 nmwith a peak at approximately 350 nm. The spectraweremea-sured through a culture plate lid but did not account for any attenuationin radiation due tomedia or nanoparticles between the light source andthe cells. The remaining “dark” plates served as negative controls. Theywere wrapped in aluminum foil and placed beside the visible or UVAplates during exposure. After the visible light, UVA radiation, or darkcondition, cells were returned to the incubator for 24 hours.

Cell viability was assessed with a live/dead assay using calcein-AMand propidium iodide (PI) (catalog # C-3099 and P-3566, Invitrogen,Carlsbad, CA). Both stains (100 μl, 5% calcein-AM and 10% PI inDMEM/F-12) were applied to each well of the 24-well plates. Calcein-AM is hydrolyzed by an esterase in live cells with intact cell membranesto form fluorescent calcein, while PI stains cells with compromised cellmembranes. Approximately 40 minutes after the stains were added,the cells were examined using epifluorescent microscopy at 100xmagnification. Digital images were taken at three locations in each

well with filters yielding either green fluorescence (live cells) or redfluorescence (dead cells). Images were divided into 48 squares, andboth live and dead cells in the middle 8 squares of each photographwere counted. Cell viability was calculated as the number of live cellsdivided by the total number of cells. Average viability was calculat-ed and graphed in Microsoft Office Excel 2007. Cell viability data afterUVA exposure were fit with a four-parameter logistic function thatwas restricted to have a maximum value of 1.0 and a minimum valueof 0 (SigmaPlot 11, Systat Software Inc., San Jose, CA). For logistic func-tions having an R2 value of 0.8 or greater, LC50 valueswere derived fromthe curves. Data from one sample (25 nm anatase) could not be fit witha minimum value of 0, but was well fit with an unrestricted minimumvalue of approximately 0.13. Two of the samples tested (142 nm ana-tase and 250 nm rutile) could not be fit suitably with a four parameterlogistic function, so LC50 valueswere determined by linear interpolationbetween adjacent data points (30 and 100 μg/ml).

Flow cytometry. ARPE-19 cells were plated in twenty T-25 flasks (5 ml/flask, 1×105 cells/ml, 25 cm2 surface area) and incubated for 48 hrs.After incubation, 3 and 10 μg/ml suspensions of 25 nm anatase, 31 nmanatase/rutile, and 214 nm rutile TiO2 were prepared as describedabove for thephototoxicity studies. Themedia in eachflaskwas replacedwith 5 ml of nano-TiO2 suspension. Two flasks received each treatmentof nano-TiO2, and eight control flasks received only fresh media. After24 hrs of incubation, flasks were placed under UVA lights or kept in thedark for 2 hrs. Mitosox (Invitrogen, Carlsbad, CA) in DMSO was dilut-ed into 37 °C DMEM/F-12 for a final concentration of 500 nM andadded to cells (1 ml per flask) during the last 30 min of UVA or dark ex-posure. Immediately following exposure, cells were trypsinized, cen-trifuged, resuspended in 0.5 ml media, and directly put on ice. Between50–100 μl of the cells were fixed in 4% paraformaldehyde for dark fieldmorphological analysis (data not shown). For viability studies, the cellswere incubatedwith 10 μg/ml PI in awater bath for 15 min and then ret-urned to ice prior to beingmeasured on the flow cytometer. This viabilitytest measured PI uptake vs. side scatter (SSC) in a cytogram and wasequivalent to the calcein-AM/PI assay in predicting the percentage ofdead cells, with greater accuracy due to the statistical nature of the flowcytometer.

A BD FACSCalibur™ (BD Biosciences, San Jose, CA) flow cytometercontaining a 488-nm laser, forward scatter (FSC) diode detector, andphotomultiplier tube SSC detector was used in this study. Prior toeach experiment, the instrument was checked for functionality andperformance using Duke 3.0-μm alignment beads (Shapiro, 2001;Zucker, 2008). The instrument yielded coefficient of variation valuesbelow 2% on all the fluorescence channels at low flow rates withMolecular Probes 2.5-μm and below 2.5% with Duke 3.0-μm

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alignment beads. The cytometer was set up to measure SSC logarithmi-cally and FSC linearly. Mitosox fluorescence was measured using log am-plifiers in the FL1 (525/25 nm) and FL2 (585/25 nm) channels. Most ofthe scales’ dynamic ranges were used to optimize the differentiationamong the different doses. The highest dose of nanoparticles was runfirst to set the range for the maximum SSC signal and the minimum FSCsignal. The fluorescence and scatter values derived from treated sampleswere normalized to control values and expressed as a percent increase.The data were evaluated using FCS Express 4.0 (DeNovo Software, LosAngeles, California).

Nanoparticle reactivity. The thiobarbituric acid reactive substance(TBARS) assay was chosen for this experiment due to its successful usein air particulate pollution health effects research,where it linked the re-activity of nano-scale particulate matter with its acute pulmonary toxic-ity (Ghio et al., 1996; Molinelli et al., 2002; Pritchard et al., 1996). Inaddition, TBARS assay substrates are not highly sensitive to UVA irradi-ation, which can inactivate substrates employed in similar assays. Inthis procedure, metals associated with the nanoparticles were reducedby ascorbate. The particles then reacted with hydrogen peroxide to pro-duce hydroxide and hydroxyl radicals. Next, the hydroxyl radicalsreactedwith 2-deoxy-2-ribose to formmalondialdehyde (MDA). Finally,at 100 °C, MDA and thiobarbituric acid formed a pink-colored adduct,which was detected by absorbance at 532 nm. Therefore, absorbancevalues reflect each sample's inherent capacity for participating inredox reactions and potentially causing oxidative stress in cells.

The TBARS assay was carried out as previously described byPritchard et al. (1996) and Molinelli et al. (2002) with modificationsfor UVA exposure. Briefly, TiO2 nanoparticles at 4 mg/ml in salinewere sonicated at a power output of 6 watts 2 times for 1–2 minusing a model TM 100 sonicator (TekMar, Cincinnati, OH) with micro-probe tip and were subsequently placed in a Model ME4.6 ultrasonicbath (Mettler Electronic Corp. Anaheim, CA) until analyzed. A reac-tion mixture consisted of 0.4 ml of 4 mg/ml sonicated nanoparticlesuspension, 0.4 ml of 4 mM H2O2 in normal saline, 0.4 ml of 4 mM so-dium ascorbate in normal saline, and 0.4 ml of 4 mM deoxyribose innormal saline, for a final concentration of 0.1 mg/ml nano-TiO2. Con-trol reactions employed 0.4 ml of saline instead of nanoparticles. Eachreaction mixture was placed into a single well of a 6-well tissue cul-ture plate (Costar 3516, Corning Incorporated Life Sciences, Lowell,MA), which did not contain cells.

Plates containing samples not exposed to UVA light were coveredwith a lid and tin foil, while plates exposed to UVA radiation were leftuncovered. All plates were mixed using an orbital shaker placed in adry heat incubator maintained at 37 °C for one hour. During thishour, UVA plates were exposed to 364 nm radiation using a 8 WModel UVLMS-38, 30 V UVA lamp (UVP, Upland, CA) at an energy of1490 μW/cm2 (5.4 J/cm2) as measured using a Model J-221 LongWave UV monitor (UVP, Upland, CA). After incubation, samples weretransferred to Eppendorf tubes and centrifuged at 1300×g for 10 min.One ml of reaction mixture supernatant was mixed with 1 ml each of1% thiobarbituric acid and 2.8% trichloroacetic acid. The mixture washeated at 100 °C for 10 min and then cooled on ice. The absorbancewas read at 532 nm using a Spectronic Model 601 UV/VIS spectropho-tometer (Milton Roy, Rochester, NY).

The data from the nanoparticle reactivity assay were analyzed sta-tistically using a 2-way analysis of variance (proc GLM, SAS 9.1, SASInstitute Inc, Cary, NC) with the independent variables being thetype of TiO2 nanoparticles and whether the sample was exposed toUVA radiation. The dependent measure was the absorbance at532 nm, which was indicative of the amount of TBARS formed. Thecriterion for a statistically significant main effect or interaction wasα=0.05. Statistically significant main effects were followed withTukey's tests to evaluate differences between each particle typewith or without UVA exposure while maintaining an overall level ofα=0.05.

Results

Characterization of TiO2 nanoparticles

DLS measurements of aggregate size varied based on dispersant,concentration, and primary particle size. Aggregates had smallersizes and were more monodispersed in media with serum than inmedia without serum, Hank's Buffered Salt Solution, or water. The sta-bilizing effect of serumwas likely due to the association of nanoparticleswith serumproteins (Deguchi et al., 2007; Ji et al., 2006). Aggregate sizealso increased with greater concentrations and primary particle sizes.Aggregates were between 700–1000 nm at the concentrations testedin the phototoxicity experiment, similar to those reported by others(Allouni et al., 2009).

Uptake and accumulation of nanoparticles in ARPE-19 cells

In the short-term experiment, TiO2 aggregates appeared to beginsettling within minutes of treatment. The largest particles settled first,according to Stokes’ law. Within hours, particles could be observedinside cells by flow cytometry or dark field microscopy (Zuckeret al., 2010). After 1–3 days, particles were incorporated into cellsand appeared to surround the nucleus. The agglomerates werelarge enough to also be seen through Hoffman modulation contrastmicroscopy.

In the long-term experiment, perinuclear rings of TiO2 aggregateswere seen after 24 hours. The amount of TiO2 observed within cellswas dose-dependent, with more particles in the 30 μg/ml conditionthan in the 10 μg/ml condition (Fig. 1). No gross effects on viability ormorphology were observed. The rings of dark TiO2 material could be ob-served inside cells through 25 days post-treatment under Hoffman inter-ference microscopy. Over time, the subcellular localization of theparticleswas consistent, but the concentration of particles in each cultureseemed to decrease.

Phototoxicity

Images typical of those scored for the live dead/assay are shown inFig. 2. Live cells stained with calcein-AM and dead cells stained withpropidium iodide were readily observed by fluorescence microscopy.Cell viability data from these experiments are presented in Fig. 3. Cellstreated with any of the nano-TiO2 samples and kept in the dark wereessentially 100% viable up to 30 μg/ml or higher, except for the 31 nmanatase/rutile, which showed a dose-related decrease in viability be-ginning at 10 μg/ml. Preliminary experiments with this sample undervisible light showed that viability did not differ from dark conditions,so no other samples were tested with visible light. Cells exposed toUVA radiation without nano-TiO2 were almost 100% viable (similarto 0.3 μg/ml TiO2 treatments, Fig. 3), but for each nano-TiO2 sam-ple, there was a dose-related decrease in viability after UVA expo-sure (Fig. 3, UVA curves). The six samples differed in UVA-inducedphototoxic potency. The LC50 values were lowest for 25 nm anataseand 31 nm anatase/rutile, intermediate for 22 nm anatase/rutile and59 nm anatase/rutile, and highest for 142 nm anatase and 214 nmrutile (Table 2).

Flow cytometry

The three TiO2 samples tested (25 nm anatase, 31 nm anatase/rutile,and 214 nmrutile) showed an increase in SSCwith higher concentrationsof TiO2 relative to control (Table 3A, Fig. 4). UVA treatment did not greatlyaffect SSC, as the values were equivalent to those of the dark controls(Table 3A, Fig. 5). UVA increased the FSC for 3 μg/ml and 10 μg/ml treat-ments of 25 nm anatase and 31 nm anatase/rutile in a dose-dependentmanner (Table 3B, Fig. 4).

Fig. 1. Long-term (25-day) experiment. ARPE-19 cells were grown to confluence in culture flasks, then treated with 59 nm anatase/rutile TiO2. Each flask was periodically observedunder modulation contrast microscopy from 1 to 25 days post-treatment, with media changed twice weekly. Aggregates of TiO2 particles were observed after 24 hours in ring-likeformations surrounding the cell nuclei. The amount of TiO2 visible within cells was dose-dependent. Some cells maintained visible rings of TiO2 through 25 days post-treatment.Magnification 200x.

230 K. Sanders et al. / Toxicology and Applied Pharmacology 258 (2012) 226–236

Mitosox fluorescence (related to ROS production) in FL2 (585/25 nm) showed an increasewith increasing doses of nano-TiO2. UVA in-duced greater Mitosox fluorescence than dark controls for both 3 μg/mland 10 μg/ml nano-TiO2 (Table 3C, Fig. 4). Though Mitosox is reportedto emit primarily in the FL2 range, this response was also observed tothe same magnitude in the FL1 (525/25 nm) detector (data notshown). 25 nm anatase and 31 nm anatase/rutile had the highest levelsof Mitosox fluorescence with UVA. All three particles produced compa-rable values in the dark (Table 3C, Fig. 5).

The25 nmanatase and31 nmanatase/rutile showed almost an iden-tical pattern for all the parameters measured (SSC, FSC, Mitosox, and vi-ability; Table 3). In contrast, the 214 nm rutile showed reducedMitosoxfluorescence, no decrease in viability, and no FSC changes relative tocontrol (Table 3). In the 10 μg/ml treatments of 25 nm anatase and31 nm TiO2 anatase/rutile with UVA, there were small subpopulationsof cells where Mitosox and scatter patterns were lower, most likelydue to some of these cells thatmay have been dead or dying (Fig. 4). Vi-ability data from flow cytometry PI staining (Table 3D) supported this

Fig. 2. Live/dead cells after UVA radiation. Results of a typical experiment showing ARPE-19pidium iodide stain) cells after treatment with a range of concentrations of anatase TiO2 (2creased and the number of dead cells increased with higher TiO2 concentrations. Cells exposeTiO2. This pattern of results is indicative of a UVA-induced phototoxic reaction.

observation and showed a similar pattern as cell viability determinedby the calcein-AM/PI assay in the phototoxicity experiments.

Nanoparticle reactivity

In the dark, the TiO2 nanoparticles caused minimal TBARS reactiv-ity relative to assay control levels (Fig. 6), indicating they possessedlittle or no inherent reactivity. The 22 nm anatase/rutile sample wasan exception, since it showed a small but statistically significant in-crease of TBARS reactivity over assay control without UVA stimulation.In contrast, five of the TiO2 nanoparticles displayed photocatalyticactivity when exposed to UVA radiation. The statistical analysisshowed significant main effects of particle type (F(6,40)=33.21,pb0.0001) and UVA radiation (F(1,40)=429.98, pb0.0001) withan interaction between particle type and UVA radiation (F(6,40)=28.87, pb0.0001). Tukey's tests revealed that under UVA radiationall particle types except the 25 nm anatase caused greater TBARSreactivity than controls and that the 25 nm anatase sample caused

cell cultures stained for living (top row, calcein-AM stain) or dead (bottom row, pro-5 nm) combined with UVA radiation. After UVA exposure, the number of live cells de-d to darkness or visible light showed little cytotoxicity at these concentrations of nano-

Fig. 3. Phototoxicity experiments. The percentage of viable cells was counted from live/dead cell micrographs as depicted in Fig. 2. The six nano-TiO2 samples tested are depictedseparately in each panel. For each sample, the mean (± standard error) percent of viable cells (from three replicates of each condition) is presented for cells exposed to darknessand for cells exposed to UVA radiation. Standard error bars are included on all points but may not be visible due to small magnitude. A four-parameter Hill function was fit to theUVA data for four of the particles, and LC50 values were obtained from these functions. The Hill function would not fit sufficiently for the two larger size particles, so these LC50

values were identified by linear interpolation between the two consecutive doses flanking 50% viability (the two highest dose points). Control (0 μg/ml) values are not showndue to log scaling; control viability averaged across all particles was 99.9% in the dark and 99.2% with UVA radiation.

Table 2LC50 values for TiO2 samples exposed to UVA radiation.

Sample Regression - R2 UVA LC50 (μg/ml)

22 nm anatase/rutile 0.878 41.225 nm anatase 0.992 4.931 nm anatase/rutile 0.942 4.559 nm anatase/rutile 0.856 47.1142 nm anatase N/A 62.8*214 nm rutile N/A 77.4*

* determined by linear interpolation between 30 and 100 μg/ml data points.

231K. Sanders et al. / Toxicology and Applied Pharmacology 258 (2012) 226–236

significantly less TBARS formation than the other samples. Asidefrom the 25 nm anatase, the differences in reactivity under UVAradiation between the other types of TiO2 nanoparticles were notstatistically significant.

Correlations among parameters

In Fig. 7, the LC50 values (Table 2) were plotted against the followingparameters: reactivity as determined under UVA radiation in the TBARS

Table 3SSC, FSC, Mitosox Fluorescence, and Viability Data Obtained from Flow Cytometry.

Dark UVA

Sample 3 μg/ml 10 μg/ml 3 μg/ml 10 μg/ml

A. Side Scatter (Relative to Control):Control 1 1 1 125 nm 3.16 7.07 3.14 6.7731 nm 2.79 5.97 2.63 4.88214 nm 2.82 4.51 2.82 4.91

B. Forward Scatter (Relative to Control):Control 1 1 1 125 nm 0.97 0.83 1.03 1.2331 nm 0.98 0.82 1.17 1.19214 nm 1.04 0.91 1.05 0.96

C. Mitosox Fluorescence (Relative to Control):Control 1 1 1 125 nm 1.13 1.50 1.34 3.2231 nm 1.16 1.47 1.65 3.10214 nm 1.22 1.51 1.11 1.47

D. Viability (% live out of total)Control 95.6 95.9 95.3 92.825 nm 95.8 97.6 92.4 45.331 nm 97.8 96.5 80.9 10.5214 nm 95.0 95.8 95.9 93.2

232 K. Sanders et al. / Toxicology and Applied Pharmacology 258 (2012) 226–236

assay (Fig. 6), primary particle size, and primary particle surface area(Table 1). For the TBARS assay, the correlation is shown with and with-out the 25 nmanatase sample because it appeared to provide a discrep-ant value in relation to the other data due to being much less reactivethan the other particles. Without data for the 25 nm anatase, theTBARS assessment showed a higher correlation coefficient than theothermeasures. Particle sizewas directly correlated and particle surface

Fig. 4. Flow cytometry histograms of 31 nm anatase/rutile. ARPE-19 cells were treated withdark or exposed to UVA radiation. Nano-TiO2 treatment caused a dose-dependent increasedark and UVA cells, indicating that the experimental conditions of radiation did not introddependent manner. Mitosox fluorescence in cells exposed to TiO2 increased more after Uwhile FSC is graphed on a linear scale.

area was inversely correlated to LC50 values, as expected. With one ex-ception, the smaller primary particles had greater surface area, weremore reactive, and were more potent UVA phototoxic substances.

Discussion

The data reported in this manuscript show that nano-TiO2 cancause phototoxicity in ocular cells in culture. As hypothesized, thedose-dependent toxicity of nano-TiO2 was much greater after UVA ra-diation than in the dark. Additionally, smaller particles resulted inhigher phototoxicity than larger particles, but the effects of crystalstructure on phototoxic potency were not clear. UVA-catalyzed ROSproduction, as measured by two assays (TBARS and Mitosox), corre-lated with phototoxicity, suggesting that ROS production was in-volved in the phototoxic mechanism of action.

TiO2 nanoparticles entered ARPE-19 cells in vitro and remainedthere for an extended period of time. The increased SSC observedwith flow cytometry indicated that particles of nano-TiO2 were asso-ciated with the ARPE-19 cells. Dark field microscopy (Zucker et al.,2010) confirmed the association of nanoparticles with the cells andrevealed that the particles were inside the cells in clumps around thenucleus. SSC increased in a dose-dependent manner with nanoparticletreatment regardless of dark or UVA exposure, suggesting that UVA didnot affect the amount of nano-TiO2 in the cells. However, SSC intensitydid vary greatly among the three samples tested, with smaller particles(25 nm anatase/rutile and 31 nm anatase/rutile) showing a higher SSCvalue than the larger particle (214 nm rutile). This result suggests thatTiO2 nanoparticles may be absorbed by the cells at different rates, possi-bly affected by particle size.

Once inside the cells, all six nano-TiO2 samples produced a photo-toxic response, but smaller particles tended to be more phototoxicthan larger ones. The most phototoxic samples were 25 nm anataseand 31 nm anatase/rutile, which had LC50 values ofb5 μg/ml with

0 (black), 3 (red), or 10 μg/ml (blue) 31 nm anatase/rutile TiO2 and either kept in thein SSC and Mitosox fluorescence (related to ROS production). SSC was similar betweenuce cellular changes. However, FSC increased with UVA exposure and TiO2, in a dose-VA exposure than after dark exposure. SSC and Mitosox are graphed on a log scale,

Fig. 5. Flow cytometry histogram comparison of SSC andMitosox fluorescence using three types of TiO2 nanoparticles (10 μg/ml). The cells were treatedwith UVA or kept in the dark fortwo hours. ARPE-19 cells (control, black) were treated with 10 μg/ml of 25 nm anatase (red), 31 nm anatase/rutile (green), or 214 nm rutile (blue). Cellular uptake of the nano-TiO2 (asmeasured by SSC) and ROS production (as measured by Mitosox fluorescence) varied among the three particles. The two smallest particles, 25 nm anatase and 31 nm anatase/rutile,showed more Mitosox activity than the largest particle (214 nm), with all three particles showing greater fluorescence than the control. There was an equivalent Mitosox response inthe three types of nano-TiO2 in the dark, which was less than that of the cells exposed to UVA. Both SSC and Mitosox are graphed on a log scale.

233K. Sanders et al. / Toxicology and Applied Pharmacology 258 (2012) 226–236

UVA. 22 nm anatase/rutile and 59 nm anatase/rutile had moderatephotoxicity, while the largest samples tested (142 nm anatase and214 nm rutile) showed the least phototoxicity. Differences in toxicityamong similarly sized particles may have been due to their unknownsurface properties. Likewise, flow cytometry with PI staining indicatedthat 10 μg/ml treatments of 25 nm anatase and 31 nm anatase/rutilewith UVA radiation caused a decrease in viability, while viability afterthe 214 nm rutile treatment was roughly equal to control.

In addition to size, other physical characteristics of nano-TiO2 havebeen shown to affect its cytotoxic potency. Different crystal formswere expected to impact phototoxicity, with anatase hypothesizedto be more active than rutile. However, it was observed that the

Fig. 6. Nanoparticle reactivity. TiO2 nanoparticles were examined for their inherentphotocatalytic reactivity using the TBARS assay. UVA: 365 nm UV radiation with an in-tensity of 5.4 J/cm2. Assay control contains saline instead of nano-TiO2. N=3–5.

samples containing a mixture of anatase/rutile were also highly photo-toxic. Contrary to initial expectations, most of the samples tested weremixed rutile/anatase structures (Table 1). There was only one sampleof pure rutile, which also contained the largest particles tested. There-fore, it was difficult to test the hypothesis regarding activity of differentcrystal forms. Other studies have reported that mixed anatase/rutilesamples were more photoreactive than smaller pure anatase particles(Coleman et al., 2005; Uchino et al., 2002).

The data from the TBARS andMitosox fluorescence assays suggestedthat UVA radiation prompted TiO2-catalyzed production of ROS as apossible mechanism for cytotoxicity. Aside from the 25 nm anatasesample (which had unusually low reactivity), the TBARS reactivitywas highly correlatedwith the in vitro cytotoxicity under UVA radiation.The 25 nm sample's low reactivity may have been due to its crystalstructure; of the four smaller particles, it was the only pure anatasesample.

The highest Mitosox fluorescence was observed in cells that wereincubated with small particles (25 nm anatase and 31 nm anatase/rutile) and exposed to UVA radiation. In the dark,Mitosoxfluorescence(averaged across all three nano-TiO2 samples tested) increased 17%with 3 μg/ml treatments and 49% with 10 μg/ml treatments (rela-tive to control samples). With UVA radiation, the average Mitosoxfluorescence for these samples increased to 37% at 3 μg/ml and159% at 10 μg/ml. These results complement the phototoxicity dataand support ROS production as a possible pathway for toxicity.Similarly, Freyre-Fonseca et al. observed that TiO2 nanoparticles canimpair mitochondrial function through ROS production in the absenceof radiation exposure, although the dose levels in that study (expressedin μg TiO2/mg mitochondrial protein) are difficult to relate to those ofthe current study (Freyre-Fonseca et al., 2011).

The flow cytometry histograms of the 10 μg/ml treatments of25 nm anatase and 31 nm anatase/rutile with UVA radiation showeda subpopulation of cells with low Mitosox fluorescence and SSC.Cells with minor damage from the treatment appear to produce

Fig. 7. Correlation between phototoxic potency and particle parameters. The figure shows the relationship between the LC50 values for each of the six samples as a function of re-activity (TBARS), primary particle size, and primary particle surface area. The TBARS data are presented with (upper left panel) and without (upper right panel) the data for 25 nmanatase particle. Linear regressions were fit to the data in each panel and the R2 values are presented.

234 K. Sanders et al. / Toxicology and Applied Pharmacology 258 (2012) 226–236

ROS, but cells severely damaged or killed by the treatment will gener-ate little or no ROS, as functional mitochondria are necessary for acti-vating Mitosox. Thus, the second subpopulation of cells was evidentlycomposed of dead, dying, or apoptotic cells that were not convertingthe Mitosox probe to its active, fluorescent form. In addition, the twosmaller nanoparticles showed an increase in FSC at 10 μg/ml nano-TiO2 with UVA, indicating that the cells were affected by the treatment.Together with the viability data (Table 3D), these results suggest thatdead or dying cells may have changed their biophysical characteristics(increased their size, causing increased FSC).

The phototoxicity of TiO2 has been described in other systems. Co-exposure to TiO2 and sunlight or UV radiation caused greater toxicityto bacteria, algae, daphnia, and marine plankton than did exposure toTiO2 alone (Adams et al., 2006; Hund-Rinke and Simon, 2006; Matsuoet al., 2001). Anatase TiO2 in the presence of UV radiation catalyzedgeneration of hydroxyl free radicals, hydrogen peroxide-catalyzedDNA damage, and genotoxicity, while rutile crystals were less activeor inactive (Hirakawa et al., 2004; Nakagawa et al., 1997; Uchino etal., 2002). In cultures of human-derived dermal fibroblasts and lungepithelial carcinoma cells, anatase TiO2 with UV radiation was moretoxic than other forms of TiO2 (Sayes et al., 2006). Recently, it wasreported that the Degussa P25 form of TiO2 (designated 21 nm ana-tase/rutile in their experiments, 31 nm anatase/rutile in ours) underUVA radiation was more phototoxic to human keratinocytes thanwere other forms of TiO2; however, 10 nm anatase was similar toDegussa P25 in assessments of superoxide dismutase activity andmalondialdehyde concentration (Xue et al., 2010). Together withthe data from the current experiments, these results demonstrate

the necessity of considering phototoxic mechanisms when evaluatingpotential risks of exposure to nano-TiO2.

If TiO2 nanoparticles reach the retina, they may accumulate in RPEcells over time. TiO2 nanoparticles have been found to enter keratinocytesin culture and to co-localize in the Golgi apparatus (Simon et al., 2011). Itis possible that nano-TiO2 entered the cells due to phagocytosis, as RPEcells in vivo have a phagocytic role (Besharse and Defoe, 1998) and tis-sue cultures of brain microglia have appeared to internalize TiO2 aggre-gates this way (Long et al., 2006). However, it is unclear in the currentexperiments whether TiO2 entered the ARPE-19 cells by passive diffu-sion or active uptake. Nano-TiO2 was observed to be retained in a ringpattern around the nucleus within some cells for up to 25 days in cul-ture after a single exposure. This observation raises the possibility of ti-tanium accumulation and retention in the RPE cells after long-termlow-level environmental exposures.

Phototoxic damage to ocular cells accumulates over time as a con-sequence of normal human aging. The presence of phototoxic com-pounds in the eye could accelerate this damage and promote earlieronset or more severe expression of conditions such as cataracts orage-related macular degeneration. The current experiments demon-strated that nano-TiO2 is phototoxic following exposure to UVA radi-ation, but not visible wavelengths of light. In adult humans, thecornea filters short UVB wavelengths (b 295 nm), and the lens filtersthe long UVB wavelengths (295–315 nm) and UVA (315–400 nm), sothat only visible light reaches the retina. Therefore, if TiO2 nanoparti-cles reached the lens, they could be exposed to UVA and undergophototoxic reactions. However, in children and adolescents, the UV-filtering capacity of the lens has not fully developed, so UVA radiation

235K. Sanders et al. / Toxicology and Applied Pharmacology 258 (2012) 226–236

is also able to reach the retina (Gaillard et al., 2000; Lerman, 1983).Because of this, UVA-aggravated phototoxicity of nano-TiO2 in theRPE cells may be more of a potential problem in younger eyes.

In summary, phototoxic damage was produced in retinal pigmentepithelial cells in vitro when there was co-exposure to nano-TiO2 andUVA radiation. This damage was apparently mediated by the produc-tion of ROS after UVA activation of the TiO2 nanoparticles. The relativepotency of the six samples of nano-TiO2 was related to particle size,with smaller particles being more toxic; particle surface area, withlarger surface areas being more toxic; and reactivity, with particlesgenerating more ROS being more toxic. These results illustrate theimportance of considering potential phototoxic mechanisms as a com-ponent of the hazard assessment of photo-catalytic nanomaterials.

Conflict of interest statement

There are no conflicts of interest.

Acknowledgments

K. Sanders was supported by a contract to EPA (EP09D000042).Thanks are extended to Theresa Freudenrich for her excellent adviceand guidance with cell culture, to Kaitlin Daniel for her evaluationof the flow cytometry data and preparation of Figs. 4 and 5, and toSarah Karafas for her help with the Mitosox experiments. The authorsare indebted to Amy Wang, Christy Powers, Steve Diamond, TomLong, and Carl Blackman for editorial comments on a previous versionof the manuscript. This manuscript has been reviewed by the NationalHealth and Environmental Effects Research Laboratory, U.S.E.P.A. andapproved for publication. Mention of trade names and commercialproducts does not constitute endorsement or recommendation foruse.

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