Evaluating Cytotoxicity and Cellular Uptake from the Presence of

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Chem. Res. Toxicol. 2010, 23, 871–879

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Evaluating Cytotoxicity and Cellular Uptake from the Presence of Variously Processed TiO2 Nanostructured Morphologies Jingyi Chen,†,§ Hongjun Zhou,‡,| Alexander C. Santulli,‡ and Stanislaus S. Wong*,†,‡ Condensed Matter Physics and Materials Sciences Department, BrookhaVen National Laboratory, Building 480, Upton, New York 11973, and Department of Chemistry, State UniVersity of New York at Stony Brook, Stony Brook, New York 11794-3400 ReceiVed NoVember 22, 2009

We evaluated the cytotoxicity of various morphological classes of TiO2 nanostructures (including 0-D nanoparticles, 1-D nanorods, and 3-D assemblies) toward living cells. These TiO2 nanostructures were modified with fluorescent dye molecules, mediated via a dopamine linkage, in order to facilitate a confocal study of their internalization. Specifically, we noted that both TiO2 1-D nanorods and 0-D nanoparticles could internalize into cells after 24 h of incubation time. However, only incubation with TiO2 1-D nanorods and 3-D micrometer-scale sea urchin-like assemblies at concentrations of up to 125 µg/mL yielded data suggestive of cell viabilities of close to 100%. Moreover, upon irradiation with UV light for periods of a few minutes at energy densities of up to 1 J/cm2, we observed up to 60% mortality rates, indicative of the cytotoxic potential of photoirradiated TiO2 nanostructures due to the generation of reactive oxygen species. 1. Introduction In the past decade, structures at the nanometer scale have been extensively produced and studied in academic, industrial, and government research settings. Recently, concerns have been raised over the potentially deleterious effects of these nanomaterials on human health and the environment (1). For a given nanomaterial, morphology (e.g., in terms of its size and shape) is thought to be one of the key factors that can decisively determine the degree of its cytotoxicity and cellular uptake. As a model system to demonstrate this idea, nanocrystallites of titanium dioxide (TiO2) are of great interest for photocatalysts, gas sensors, pigments, photovoltaic applications, as well as additives in cosmetics, sun tan lotions, pharmaceuticals, and food colorants because of their electronic, optoelectronic, and catalytic properties, which are intrinsically coupled to their high surface area, porosity, low cost, and chemical stability (2). Titanium dioxide is a wide band gap semiconductor (3.2 eV) with an efficient photocatalytic ability (2b). Upon irradiation by a photon possessing an energy larger than its band gap, an electron associated with the TiO2 nanoparticle can be excited to the conduction band, thereby creating a positively charged hole. This hole can then remove an electron from a molecule (such as water) that is in contact with the surface of the TiO2, thereby resulting in the formation of reactive oxygen species (3). Ultrafine TiO2 nanoparticles (4 mg/kg (7); specifically, 10-20 nm-sized TiO2 particles in the absence of photoactivation have been found to induce oxidative DNA damage, lipid peroxidation, and micronuclei formation as well as increased hydrogen peroxide and nitric oxide production in human bronchial epithelial cell lines (8). Additional pulmonary instillation studies with a number of TiO2 nanostructures in rats have deduced that toxicity is not necessarily dependent upon particle size and surface area (9) but rather is more associated with inherent distinctions based on crystal structure (e.g., rutile versus anatase polymorphs of TiO2 which possess the same crystal symmetry but maintain distinctive structures), inherent surface pH of the particles, or surface chemical reactivity (10). Intratracheal instillation of rutile TiO2 nanorods caused upregulation of lung inflammation, pulmonary and cardiac edema, and systemic inflammation and triggered platelet aggregation both in ViVo and in Vitro at concentrations of 0.4 to 10 µg/mL (11). A number of studies of TiO2 nanoparticles have been conducted in Vitro on different types of cell lines (1b). Some results have suggested that there is no cytotoxic effect with relatively large particles at low concentrations (12). However, particles at the nanometer scale could clearly cause pulmonary damage, inflammation, and even DNA damage with classic dose-response behavior (13). It was recently found that 100% anatase TiO2 nanoparticles, regardless of size, could induce cell necrosis, whereas rutile TiO2 nanoparticles could initiate apoptosis through the formation of reactive oxygen species (14). Moreover, it has also been demonstrated that catalytically inactive materials (rutile) were 2 orders of magnitude less

10.1021/tx900418b  2010 American Chemical Society Published on Web 04/21/2010

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cytotoxic than similarly sized anatase counterparts, possessing high catalytic activity (13c). In addition, upon exposure to UV light, TiO2 particles can induce the formation of hydroxyl radicals and hydrogen peroxide in water that can cause the loss of cell viability (15). Not surprisingly, photoactivated TiO2 nanostructures have already been successfully applied to treat cancer cells (15b, 16). However, as mentioned, most studies were generally focused on size and concentration effects upon cytotoxicity. Other factors, including morphology, also play an important role with respect to cell viability (17). Specifically, few studies have reported on how reproducible variations in morphology, especially unconventional, nonspherical shapes, can affect TiO2 cytotoxicity. In our laboratory, we have previously synthesized a number of zero-, one-, and three-dimensional (i.e., 0-D, 1-D, and 3-D) anatase motifs of TiO2 with well-controlled morphology. Specifically, in our system, 0-D (e.g., reasonably monodisperse nanoparticles measuring less than 100 nm in diameter), 1-D (e.g., anisotropic, high aspect ratio materials with two dimensions, measuring less than 100 nm in diameter), and 3-D (defined as sterically bulky aggregates of constituent nanoparticles or nanowires) manifestations of TiO2 were represented by nanoparticles, nanowires, and micrometer-scale, sea urchin-like assemblies, respectively. In this work, we specifically (i) compared the cytotoxicity of these various anatase TiO2 morphologies in human epithelial cells, (ii) evaluated the cell viability of these nanostructures upon UV light exposure, and finally (iii) examined their cellular uptake using confocal microscopy.

2. Experimental Procedures 2.1. Synthesis of TiO2 Nanostructures. TiO2 nanorods were synthesized using a modified hydrothermal method, as previously described in detail (18). Briefly, a commercial anatase TiO2 powder (Alfa Aesar, 0.1-1 g) was dispersed in an 18 mL aqueous solution of NaOH (5-10 M) and placed into a Teflon-lined autoclave with an 80% filling factor. The autoclave was then oven-heated at 180 °C for 24 h. A white precipitate was eventually isolated upon filtration and washed repeatedly with copious amounts (100 to 200 mL) of distilled, deionized water until the pH value of the supernatant had attained a reading close to 7. After collection by centrifugation and oven drying at 120 °C overnight, as-produced 1-D sodium hydrogen titanate nanomaterials were neutralized using a 0.1 M HCl solution. These were then washed with distilled, deionized water (as much as 100 to 200 mL) until the pH of the supernatant had attained a value of ∼7, in order to prepare their hydrogen titanate analogues, which were subsequently oven-dried overnight at 120 °C. Prior to conversion to TiO2, the resultant hydrogen titanate samples were then dispersed in distilled water, transferred to an autoclave, and heated at 170 °C for 24 h. It is worth pointing out that prior to our biological experiments, the TiO2 nanorods were shortened using a sonication bath (Branson, CT) for 5 min to lengths of 1-2 µm in order to facilitate cell internalization. As for the synthesis of TiO2 nanoparticles, hydrogen titanate nanotubes (∼7 to 10 nm) were transformed into high-purity, singlecrystalline anatase nanoparticles, using a previously reported hydrothermal process, run at 120 °C (18a). Regarding the production of 3-D TiO2 aggregates, as-prepared sodium or potassium hydrogen titanate 1-D nanostructures were neutralized using approximately 15 mL of 0.1 M HCl solution and subsequently washed with distilled, deionized water in order to yield assemblies of hydrogen titanate 1-D nanostructures. Three-dimensional assemblies of anatase TiO2 1-D nanostructures were then synthesized by annealing aggregates of hydrogen titanate 1-D nanostructures in air at 350-500 °C for 1-10 h, as previously noted (18b).

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2.2. Modification of TiO2 Nanostructures. To facilitate the conjugation of a fluorescent label, TiO2 nanostructures were then incubated with an excess of dopamine (DA, 1 mM) in aqueous solution. After the removal of free dopamine by centrifugation, the resulting DA-modified TiO2 nanostructures were further reacted with fluorescein isothiocyanate (FITC) so as to yield FITC-labeled TiO2 nanostructures. Free FITC molecules were subsequently removed by centrifugation, and the sample was resuspended in phosphate buffered saline (PBS) for further processing. 2.3. Structural Characterization of TiO2 Nanostructures. The diameters and lengths of as-prepared 1-D nanorods and 3-D nanostructures were initially characterized using a field emission scanning electron microscopy instrument (FE-SEM Leo 1550), operating at an accelerating voltage of 15 kV and equipped with energy-dispersive X-ray spectroscopy (EDS) capabilities. Samples for scanning electron microscopy (SEM) were prepared by dispersing as-prepared nanostructures in ethanol, sonicating for about 2 min, and then depositing the sample onto either a conductive tape or a silicon wafer, attached to a SEM brass stub. All of these samples were then conductively coated with gold by sputtering for 15 s to minimize charging effects under SEM imaging conditions. Low-magnification transmission electron microscopy (TEM) images of 0-D nanoparticles were obtained at an accelerating voltage of 80 kV on an FEI Tecnai12 BioTwinG2 instrument, equipped with an AMT XR-60 CCD Digital Camera System. Specimens for these TEM experiments were prepared by sonicating the as-prepared product for 2 min in ethanol to ensure adequate dispersion of the nanostructures and placing one drop of the solution onto a 300 mesh Cu grid, coated with a lacey carbon film. Zeta potential measurements were obtained using a Zetasizer Nano ZS (Malvern) instrument. All of the data were collected at 25 °C on samples prepared by dispersing TiO2 nanostructures in PBS solution at pH 7.4. Nitrogen isotherms were measured using a BET Micromeritics Sorption Analyzer Model 2010 at 77 K. Before the experiments, samples were outgassed at 120 °C for a period of 1.5 to 2 h. From the isotherms, we were able to compute the total surface area. Mid-infrared spectra were obtained on a Nexus 670 (Thermo Nicolet) equipped with a single reflectance ZnSe ATR accessory, a KBr beam splitter, and a DTGS KBr detector. As-prepared solid powder samples were placed onto a ZnSe crystal where data were taken with a reproducible pressure. A background correction was performed using the ZnSe crystal in the spectral range studied. Thermogravimetric Analysis (TGA) was performed in an argon environment using a TA Instruments Q500 TGA model. Experiments were carried out from room temperature to 1000 °C with a heating rate of 10 °C/min. Samples of 3-4 mg were placed in a platinum pan for individual runs. 2.4. Cell Culture. HeLa cells were grown in a minimal essential medium (MEM, Giboco) solution. This treatment was supplemented, however, with 10% (v/v) heat-inactivated fetal bovine serum (FBS) as well as with 1% (v/v) penicillin and streptomycin (P/S) at 37 °C in a humidified atmosphere in the presence of 5% CO2. Cells were subsequently trypsinized and reseeded onto a lysine-coated 35-mm bottom glass dish (MatTek Corp.) at a concentration of 1 × 105 cells/mL overnight, prior to additional use. 2.5. 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium Bromide (MTT) Cell Viability Assay. HeLa cells were plated at a density of 2 × 104 per well in 96-well plates and cultured for 24 h in the presence of serially diluted nanostructures. At the end of this period, the number of viable cells was determined by a quantitative colorimetric staining assay using a tetrazolium salt (MTT, Sigma Chemical Co.). To probe the effect of UV irradiated TiO2 nanostructures on cell viability, HeLa cells were exposed to medium-wave (302 nm, UV lamp Model UVM-57) UV light irradiation at 8 mW/cm2 for time periods of 1 and 2 min, corresponding to energy fluxes of 0.50 and 1.0 J/cm2, respectively, in the presence of different concentrations of nanostructures, prior to the MTT assay itself. The distance between the UV lamp and the probed cells was ∼1.5 cm.

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2.6. Reactive Oxygen Species (ROS) Assay. HeLa cells were cultured in 100 mm dishes in the presence of culture media containing various TiO2 nanostructures at a final concentration of 100 µg/mL for 24 h. These cells were subsequently collected and incubated in 2 mL of a working solution of 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA, Invitrogen), a fluorogenic probe commonly used to detect intracellular generation of reactive oxygen species, at 37 °C for 40 min. Fluorescence data of oxidized H2DCFDA were recorded by using a filter based microplate reader (Tecan Infinite F200) with the excitation and emission wavelengths set at 485 ( 20 nm and 535 ( 25 nm, respectively. For the UV experiments, cells were exposed to UV light at 8 mW/cm2 for periods of 1 min after the addition of various TiO2 nanostructures. 2.7. Fluorescence Imaging. Alterations in the morphology of HeLa cells were investigated using epifluorescence microscopy after selective staining of their F-actin cytoskeleton network and nuclei using rhodamine phalloidin (lyophilized powder, Molecular Probes, Invitrogen) and DAPI (4′, 6-diamidino-2-phenylindol, Sigma), respectively. Specifically, cells were initially fixed with 4% paraformaldehyde in PBS for 10 min at room temperature, and permeabilized with 0.1% Triton X-100 in PBS for 20 min. Subsequently, cells were incubated with rhodamine phalloidin at a concentration of 1 µg/mL in PBS for 40 min. Thereafter, DAPI was applied for 15 min at room temperature in order to label cell nuclei. The extent of staining was visualized using an Olympus IX71 fluorescence microscope. Digital images were acquired with a QICAM Fast Cooled Mono 12-bit camera (Q Imaging), working with the Olympus capture software version 2.90.1. Fluorescence signals were detected using the following filters: rhodamine (excitation, 543 nm; emission, 565 nm) and DAPI (excitation, 365 nm; emission, 420 nm). 2.8. Confocal Fluorescence Microscopy (CFM) Imaging. For nanomaterials only, a diluted sample was drop cast onto a 36-mm uncoated bottom glass dish (MatTek Corp.), prior to imaging. For cell experiments, the nanomaterial sample was additionally incubated with cells in a lysine-coated glass dish, overnight. Cells were then twice washed with PBS so as to remove excess nanomaterials. Cell membranes were subsequently stained with the Texas Red-X conjugate of wheat germ agglutinin (TR-WGA, Invitrogen). Confocal microscopy experiments were performed using a Zeiss LSM 510 META NLO two-photon laser scanning confocal microscope system. For green fluorescence, the system was operated using a 488 nm excitation wavelength; we detected emission wavelengths of 527 ( 23 nm using a 500-550 nm bandpass filter. For red fluorescence, the system was operated using a 543 nm excitation wavelength; we detected emission at 615 ( 20 nm using a 560 nm long-pass filter. Images were captured using either a C-Apochromat 63×/1.2 Water (corr.) objective or a Plan-Apochromat 100×/1.45 oil objective. Acquired data were then analyzed using LSM 510 META software.

3. Result and Discussion 3.1. Nanostructure Characterization. We have prepared TiO2 possessing three different morphologies: (a) reasonably monodisperse nanoparticles (mainly cubes and rhombohedra) with diameters of ∼14 nm, (b) nanorods with diameters of 50 nm and lengths of up to several micrometers (e.g., 2 µm), and (c) 3-D nanostructures with diameters on the micrometer scale, as shown in Figure 1A-C. The crystal structures of all three of these morphologies were determined to be anatase in nature by means of powder X-ray diffraction (Figure 1D). The X-ray diffraction pattern itself was indexed to a pure tetragonal anatase phase of TiO2, with calculated cell constants of a ) b ) 3.785 Å and c ) 9.514 Å (18). Measured peak intensities and positions are in good agreement with standard database values (JCPDS File No. 21-1272). Table 1 lists relevant physical parameters collected for our as-prepared TiO2 nanostructures. Brunauer-Emmett-Teller

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Figure 1. TEM image (A), SEM images (B and C), and (D) corresponding XRD patterns of anatase TiO2 0-D nanoparticles, 1-D nanorods, and 3-D aggregate motifs.

Table 1. Selected Physical Characterization of Classes of TiO2 Nanostructures nanostructure type

zeta potential (mV)

BET measurements (m2/g)

0-D nanoparticles 1-D nanorods 3-D sea urchin-like aggregates

-21.3 ( 1.3 -24.3 ( 0.9 -22.8 ( 1.0

84.13 ( 1.10 66.55 ( 0.19 109.91 ( 1.07

(BET) measurements of surface area were computed to be 84.13 ( 1.10, 66.55 ( 0.19, and 109.91 ( 1.07 m2/g for 0-D nanoparticles, 1-D nanorods, and 3-D nanostructures, respectively. Zeta potential data suggested that the surfaces of all of these nanostructures were negatively charged. 3.2. Cell Viability. To verify whether the TiO2 nanostructures had the potential for cellular toxicity, cell viability assays were performed, on the basis of the reduction activity of methyl thiazolyl tetrazolium (MTT) (19), and the resulting data are shown in Figure 2. The viability of untreated cells was assumed to be 100%. Figure 2A shows the cell viability after incubation with different concentrations of TiO2 at 37 °C for 24 h. For the 3-D and 1-D anatase motifs, cell viability only slightly lowered as a function of concentration and remained at over 90% up to a final concentration of 125 µg/mL. In the case of 0-D anatase nanoparticles, however, cell viability gradually decreased to a level of 80% at a final concentration of 125 µg/mL. This observation may be attributed not only to the inherently higher surface-area-to-volume ratio of nanoparticles as compared with larger morphological motifs but also, more importantly, to endocytosis as a more facile mechanism of 0-D nanoparticle uptake (13c, 14). Hence, not surprisingly, the cytotoxicity of 3-D structures exhibited a trend similar to that of 1-D nanorods. That is, both 1-D and 3-D motifs were relatively nontoxic in the concentration range of study, as these unmodified, bulkier, and geometrically more exotic structures were less likely to enter into and diffuse across HeLa cell membranes as opposed to their smaller, less bulky counterparts. Moreover, with respect to our observations, we can rule out the possibility of the differential solubility of the various as-prepared nanostructured motifs as all of these were sufficiently well suspended (inset to Figure 2A) over the course of the incubation period. Although it has been previously reported that dendritic TiO2 particles are more toxic than analogous particles with a rod-like shape, in those particular experiments, the large micrometer-scale particle size was primarily responsible for the observed cytotoxicity (12a, 20). We then exposed the cells to UV irradiation in order to explore possible cell death induced by the presence of TiO2 0-D nanoparticles, 1-D nanorods, and 3-D nanostructures, respectively (Figure 2B-D). Prior to exposure to UV light with

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Figure 2. MTT assay associated with cell viability: (A) after a 24 h incubation time period with varying concentrations (normalized on the basis of surface area data collected from BET measurements) of TiO2 nanostructures as well as after treatment with UV light irradiation in the presence of varying concentrations of (B) 0-D nanoparticles; (C) 1-D nanorods; and (D) 3-D sea urchin-like assemblies of TiO2 nanostructures. All measurements are averages of three replicas. Error bars represent standard deviations of three replicates of data taken. Inset in Figure 2A presents three different morphologies of TiO2 nanostructures, well suspended in culture medium up to 24 h at a final concentration of 125 µg/mL.

wavelength at 302 nm, a series of HeLa cells was treated with variable concentrations of TiO2 nanostructures. Cell viability was calculated as the percentage of living, treated cells as a fraction of the initial total of untreated cells. As expected, in the absence of TiO2 nanostructures, cell viability remained close to 100% even at a power density of 0.5 J/cm2, which is below the threshold of UV irradiation-induced cell death (21). By contrast, in the presence of TiO2 nanostructures, at an identical power density of 0.5 J/cm2, the amount of cell death increased with increasing concentrations of TiO2 nanostructures present, suggestive of the photoinduced cytotoxic potential of TiO2. Specifically, relative to nonirradiated cells, cell viability dropped by amounts of 37%, 40%, and 30% for 0-D nanoparticles, 1-D nanorods, and 3-D nanostructures, respectively, at a concentration of 125 µg/mL. Not surprisingly, when the power density was increased to 1 J/cm2, relative to nonirradiated cells, cell viability decreased by values of 50%, 40%, and 52% for 0-D nanoparticles, 1-D nanorods, and 3-D nanostructures, respectively, with increasing TiO2 concentrations (up to 125 µg/mL) due to the photoinduced cytotoxicity incurred by UV irradiation. Furthermore, our results are consistent with previous data reported on HeLa cell line targeting through photoexcitation of folic acid-modified TiO2 nanoparticles (16). To investigate potential changes in cellular morphology upon UV irradiation, we selectively red stained the F-actin skeletal network with rhodamine phalloidin and blue stained the cell nuclei with DAPI. In absence of TiO2, irradiated cells appeared to possess a shape similar to that of untreated cells after 1 min of UV light exposure (Figure 3A and B). After 2 min of UV light exposure alone, treated cells appeared slightly more distorted (Figure 3C). By contrast, in the presence of TiO2

nanostructures, a number of cells appeared to have swelled and elongated (Figure 3D and E). Other cells seemed slightly more distorted, while a few ostensibly lost their structural membrane integrity altogether (Figure 3F). To investigate the possibility of oxidative stress as a factor in incurring the observed cell damage (e.g., membrane damage noted in Figure 3F for instance) (22) under UV irradiation, we measured possible reactive oxygen species formation using a ROS detection reagent, namely, H2DCFDA (Figure 4). Specifically, neither chemically reduced nor acetylated forms of 2′,7′dichlorofluorescein (DCF) are inherently fluorescent. However, there is measurable fluorescence if the acetate group is removed by enzymes such as intracellular esterases and when the compound is oxidized within the cells (23). By comparison with untreated controls, cells cultured in media containing TiO2 0-D nanoparticles for 24 h showed a marked increase in the total DCF fluorescence intensity up to 174%, whereas 1-D nanorods and 3-D nanostructures evinced little increase in the fluorescence intensity with data corresponding to 103% and 109% of control values, respectively. Interestingly, upon exposure to UV light irradiation for 1 min, the measured fluorescence intensities rose to 209%, 183%, and 167%, relative to analogous intensity values for controls, for 0-D nanoparticles, 1-D nanorods, and 3-D nanostructures, respectively. This result is consistent with a previous study in which a 500 W Hg lamp was used to photoirradiate TiO2 nanoparticles (15b). Hence, the observed cell death, manifesting itself in part by a marked drop in cellular stiffness (24), is possibly caused by the presence (and accompanying stress) of reactive oxygenated species (e.g., hydroxyl and peroxy radicals, superoxide anions, singlet oxygen,

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Figure 3. HeLa cells subjected to different treatment conditions: (A) without any treatment; (B) UV light irradiation for 1 min; (C) UV light irradiation for 2 min; (D) treatment with 0-D nanoparticles for 1 min of UV light irradiation; (E) treatment with 1-D nanorods for 1 min of UV light irradiation; and (F) treatment with 3-D sea urchin-like micrometer-scale aggregates of TiO2 nanostructures under UV light irradiation. Cytoskeletal filaments have been stained red while the nuclei are stained blue.

Figure 4. DCF fluorescence measurements of HeLa cells associated with ROS formation. The fluorescence intensity of each nanoparticle (NP), nanorod (NR), and 3-D micrometer-scale sea urchin-like aggregate (3-D) sample was compared with and computed relative to that of untreated, control cells. All measurements are averages of three replicates. In addition, error bars represent standard deviations of three replicates of data taken.

and hydrogen peroxide) generated during the photoexcitation process (13c, 13d, 15b). 3.3. Case Study: Cellular Uptake of 1-D TiO2 Nanorods. To demonstrate the potential capacity for TiO2 nanorod uptake into the cells, we functionalized the nanorods with a dye molecule, namely, fluorescein isothiocyanate (FITC), through the mediation of a bridge linker dopamine (DA), as shown in Figure 5A. The OH groups on the DA molecules can be readily coupled to the surface of TiO2 via uncoordinated surface defect sites (25). Subsequently, the isothiocyanate groups (-NdCdS) of the FITC molecules reacted with the dangling NH2 groups

Figure 5. (A) Schematic illustrating the conjugation of fluorescein isothiocyanate (FITC) molecules onto the surface of TiO2 nanorods through dopamine (DA) bridges. (B) Thermogravimetric analysis profile (TGA) of TiO2 nanorods. (C) FT-IR spectra of variously functionalized TiO2 nanorods. In all cases, NR represents a naked nanorod, DA symbolizes dopamine, NR-DA is associated with DA-modified nanorods, and NR-DA-FITC can be attributed to FITC-labeled nanorods.

of DA molecules that had been initially immobilized onto the TiO2 nanostructure so as to yield FITC-TiO2 nanorods. The amount of attached and conjugated DA molecules can be quantified using thermogravimetric analysis (TGA). Figure 5B shows a 22% weight loss, after heating DA-modified TiO2 nanorods (red curve) under nitrogen at temperatures up to 900 °C. It was evident that the DA molecules started to decompose at 200 °C and had completed their decomposition by 500 °C.

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Figure 7. Confocal images of HeLa cells after incubation with FITClabeled TiO2 nanorods (denoted by white arrows): (A) phase contrast image; and (B) superimposition of both phase contrast and florescence images. Note that the cell membrane has been stained with the Texas Red-X conjugate of wheat germ agglutinin.

Figure 6. Confocal images of green FITC-labeled TiO2 nanorods: (A) fluorescence image; (B) phase contrast image; and (C) superimposition of both fluorescence and phase contrast images.

Moreover, the sample had reattained a white coloration suggestive of pristine TiO2. If we assume that moisture can account for ∼2% of the weight loss at temperatures below 100 °C, the percentage weight of attached DA molecules in the samples therefore measured ∼20%. By contrast, in the control experiment (black curve), no obvious weight loss of naked TiO2 nanorods was observed. The FTIR spectra also confirmed the formation of both DAmodified TiO2 and FITC-labeled TiO2 nanorods (Figure 5C). In both samples, the broad band at 3410 cm-1 assigned to OH stretching vibrations can be ascribed to a water residue, consistent with the TGA data. For DA-modified TiO2 nanorods, bands at 1600 and 1496 cm-1 could be assigned to the benzene ring stretch of DA molecules, whereas the band at 1286 cm-1

Figure 8. Z-stacked confocal images, representing a superposition of green and red fluorescence, of a single HeLa cell incorporating green FITC-labeled TiO2 nanorods. Slices were obtained from top to bottom in regular one-micrometer-sized intervals. Note that the cell membrane has been red stained with the Texas Red-X conjugate of wheat germ agglutinin. Each image measures 20 µm × 15 µm.

could be attributed to a C-O stretch (26). In the case of FITClabeled TiO2 nanorods, most of the observed bands were similar to those of DA-modified TiO2 nanorods. However, there were

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Figure 9. Representative single orthogonal slice (slice 5, 20 µm × 20 µm) from a set of 3-D images (Z-stacks) as shown in Figure 8, indicating that the nanorods were incorporated within the cells.

additional peaks, associated with the inherent structure of FITC, which could be assigned to the benzene ring stretch and which were located at 1593, 1500, and 1442 cm-1, respectively. To further confirm the successful surface modification of TiO2 nanorods with FITC molecules, confocal fluorescence microscopy images were taken, as shown in Figure 6. Figure 6A shows a typical phase contrast image of a FITC-labeled TiO2 nanorod suspension, highlighting its morphological profile within the field of view. We then excited our sample at 488 nm and noted an expected emission at ∼520 nm for FITC in aqueous solution, thereby yielding a readily detectable green fluorescence (Figure 6B). The superimposition of both phase contrast and the corresponding fluorescent images in Figure 6C is indicative of a very strong degree of overlap and a correlation in signal, thereby suggesting that the vast majority of as-prepared TiO2 nanorods were labeled with fluorescent dye molecules. We subsequently incubated FITC-labeled TiO2 nanorods (∼25 µg/mL) with HeLa cancer cells and studied their uptake using confocal microscopy. Cell membranes were stained with the red Texas Red-X conjugate of wheat germ agglutinin (TR-WGA) for enhanced visualization. CFM images of the cells were taken after removal of the excess TiO2 by washing with PBS three times. As seen in Figure 7A, some rod-like particles, delineated by white arrows, appeared to be colocalized with the cells themselves. A superimposition of the phase-contrast image with the fluorescence image (Figure 7B) suggested that these rod-like particles could be ascribed to FITC-labeled TiO2 nanorods. The fact that some of the rod-like particles may have

appeared brighter than others was due to the likelihood that these were situated within the focal plane itself. To confirm the spatial localization of nanorods within a typical cell, we obtained a series of Z-stacked images of an individual cell at 1 µm slice intervals (Figure 8) from top to bottom. FITC-labeled TiO2 nanorod composites evinced a green fluorescence that became apparent by slice 3 but which had clearly faded in intensity as one proceeded vertically to slice 9. We focused on slice 5 situated in the cellular interior. Data corresponding to the orthogonal X, Y, and Z planes, respectively, within the cell interior are shown in Figure 9. Specifically, the main, central portion of Figure 9 corresponds to fluorescence data measured in a plane in the Z direction, whereas the rightmost and uppermost edge regions of the image present data at planes in the X and Y directions, respectively. Because these three planes share a common focal center within the cell itself and, moreover, as these intimately interconnected planes evidently all demonstrate green fluorescence simultaneously, we can reasonably conclude that the FITC-labeled TiO2 nanorods are localized (and can therefore theoretically transport cargo) within the interior of the HeLa cells themselves. We cannot discount the possibility that some of the nanorods did still remain on the surface of the cells themselves. 3.4. Additional Studies. In parallel, complementary experiments using identical protocols, we noted that FITC-labeled TiO2 nanoparticles can also be taken up into cells by means of endocytosis (27) (Figure 10), although we have previously noted their inherently larger toxicity in Figure 2A. In fact, both TiO2

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4. Conclusions

Figure 10. Confocal images of HeLa cells after incubation with FITClabeled TiO2 nanoparticles, as shown in green: (A) fluorescence image (100 µm × 100 µm) and (B) a representative single orthogonal slice (middle slide, 40 × 40 µm) from a set of 3-D images (Z-stacks, 9 slices), indicating that the nanoparticles were internalized within the cells. Note that the cell membrane has been red stained with the Texas Red-X conjugate of wheat germ agglutinin.

Figure 11. Z-stacked confocal images, representing a superposition of green and red fluorescence, of a single HeLa cell incorporating FITClabeled 3-D micrometer-scale sea urchin-like assemblies of 1-D TiO2 nanostructures, as shown in green. Slices were obtained from A to D in regular two-micrometer-sized intervals. Note that the cell membrane has been red stained with the Texas Red-X conjugate of wheat germ agglutinin. Each image measures 100 µm × 100 µm.

1-D nanorods and 0-D nanoparticles evinced very similar cellular incorporation behavior with better uptake performance noted with 0-D nanoparticles. Moreover, Figure 11 highlights our observation that there appeared to be a relatively smaller degree of cellular incorporation of 3-D micron-scale sea urchinlike assemblies of TiO2 nanostructures as compared with other nanostructures tested under similar incubation conditions. From our overall analysis, a significant proportion of these 3-D nanostructures seemed to remain on the external surfaces of the cells, presumably due to the larger size and steric bulk of these nanomaterials, a finding consistent with their lower observed tendency towards cytotoxicity (Figure 2A). Hence, in summary, (a) the relatively facile internalization and (b) associated low cytotoxicity, thereby suggesting the potential for an inherent biocompatibility, of TiO2 nanorods in particular imply that these anisotropic nanostructures with cylindrical motifs may find useful applicability as plausible delivery vehicles.

We tested the cytotoxicity of various anatase nanostructures, possessing three different morphological motifs. In the case of 0-D anatase nanoparticles, cell viability gradually decreased to a level of 80% at a final concentration of 125 µg/mL. By contrast, the observed cell viability of 1-D (similarly sized to the 0-D nanomaterials) and 3-D structures remained close to 100%, even after treatment with TiO2 at concentrations of up to 125 µg/mL. This behavioral difference may be related to factors including the differential uptake ability into cells of unmodified structures of varying geometries. Upon irradiation with UV light for 1 min at a power of 8 mW/cm2, we observed a 37%, 40%, and 30% cell mortality rate, relative to nonirradiated cells, for 0-D nanoparticles, 1-D nanorods, and 3-D sea urchin-like aggregates of nanostructures, respectively, induced by the presence of photoexcited TiO2 nanostructures at concentrations of up to 125 µg/mL. Additional experiments attributed the observed cell death under UV irradiation to the generation of reactive oxygen species for all of the TiO2 morphologies tested. To probe cellular uptake potential, fluorescently labeled 1-D TiO2 nanostructures were prepared by chemical modification with dye molecules, mediated using a dopamine linkage. Confocal studies confirmed that the size and shape of nanomaterials influenced their degree of cellular uptake. Whereas TiO2 1-D nanorods and 0-D TiO2 nanoparticles could be readily internalized into the cells after 24 h of incubation time, the more sterically unwieldy, highest surface area 3-D aggregates of TiO2 were less likely to be incorporated into HeLa cells. Our overall results are consistent with the modality of the localized administration of photoexcited surface-functionalized TiO2 nanostructures, which can generate reactive oxygenated species that can subsequently permeate into tumor tissue, as a powerful and flexible anticancer treatment methodology. In particular, the reasonably low cytotoxicity of 1-D TiO2 nanorods to cells coupled with their photocatalytic ability to induce apoptosis only upon UV irradiation are reminiscent of the prior use of titanium dioxide whiskers (16) derivatized with daunorubicin and magnetically guided titania nanotubes (28) as plausible anticancer agents. We have gone one step further in this work, in that we have shown direct confocal evidence for the internalization of TiO2 nanostructures into our cells. Acknowledgment. Synthesis work on TiO2 morphologies was performed at Brookhaven National Laboratory under contract number DE-AC02-98CH10886. We also thank the National Science Foundation (CAREER Award DMR-0348239) and the Alfred P. Sloan Foundation (2006-2008) for PI support as well as for support of the biological work and characterization studies. The authors also acknowledge the technical service and advice provided by Susan Van Horn for TEM and Guo-Wei Tian for CFM, performed at the Central Microscopy Imaging Center at Stony Brook. They also thank Rebecca Rowehl for her invaluable help with cell culture preparations at the Cell Culture and Hybridoma facility at Stony Brook. Amanda L. Tiano and Fulya Dogan are also thanked for their assistance with the BET measurements.

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