Molecular mechanism of DNA damage induced by titanium dioxide nanoparticles in toll-like receptor 3 or 4 expressing human hepatocarcinoma cell lines

Background Titanium dioxide nanoparticles (TiO2 NPs) are widely used in the biological sciences. The increasing use of TiO2 NPs increases the risk of humans and the environment being exposed to NPs. We previously showed that toll-like receptors (TLRs) play an important role in the interactions between NPs and cells. Our previous results indicated that TLR4 increased the DNA damage response induced by TiO2 NPs, due to enhanced NP uptake into the cytoplasm, whereas TLR3 expression decreased the DNA damage response induced by TiO2 NPs because of NP retention in the endosome. In this study, we explored the molecular mechanism of the DNA damage response induced by TiO2 NPs using TLR3 or TLR4 transfected cells. We examined the effect of TLR3 or TLR4 over-expression on oxidative stress and the effect of DNA damage induced by TiO2 NPs on gene expression levels. Results Our results showed evidence for elevated oxidative stress, including the generation of reactive oxygen species (ROS), with increased hydrogen peroxide levels, decreased glutathione peroxidase, and reduced glutathione and activated caspase-3 levels in cells exposed for 48 h to 10 μg/ml TiO2 NPs. These effects were enhanced by TLR4 and reduced by TLR3 over-expression. Seventeen genes related to DNA double-strand breaks and apoptosis were induced, particularly IP6K3 and ATM. Conclusion Our results indicated that TiO2 NPs induced ROS, and the above molecules are implicated in the genotoxicity induced by TiO2 NPs.


Background
Nanotechnology is one of the fastest growing sectors of the high-tech economy. Several consumer products currently use nano-materials; these products have personal, commercial, medical, and military uses [1,2]. Engineered nano-materials are suited to a wide range of novel applications in the electronics, healthcare, cosmetics, technologies and engineering industries. The dearth of toxicological data on nano-materials makes it difficult to determine if there is a risk associated with exposure to nano-materials. Thus, there is an urgent need to develop rapid, accurate and efficient testing strategies to assess the health effects of these emerging materials [3].
Titanium dioxide nanoparticles (TiO 2 NPs) possess dramatically different physicochemical properties compared to TiO 2 fine particles (FPs). TiO 2 NPs are widely used in the biomedical and bioengineering fields due to their strong oxidizing properties and chemical inertness [4]. Moreover, TiO 2 nano-materials are widely used in industrial and consumer products due to their strong catalytic activity attributed to their small size, which provides a larger surface area per unit mass. These properties of TiO 2 NPs may present both unique bioactivity properties and challenges to human health [5][6][7]. Indeed, the physicochemical properties of TiO 2 NPs have been demonstrated to correlate with their toxicological effects [8]. TiO 2 nano-materials have attracted much interest in medical fields due to their photo-reactivity [9]. It was previously reported that a TiO 2 photo-catalyst can kill bacterial cells in water due to the generation of compounds such as reactive oxygen species (ROS) [10,11]. Furthermore, photo-excited anatase TiO 2 nanoparticles effectively induce cytotoxicity in HeLa cancer cells [12]. TiO 2 NPs are used increasingly in industrial products, such as toothpastes, sunscreens, cosmetics, pharmaceuticals, and food products [13]. Human exposure may occur during both the manufacturing process and use. The widespread use of TiO 2 NPs, and their potential entry into the body through dermal, ingestion, and inhalation routes, suggests that TiO 2 NPs pose a potential exposure risk to humans, livestock, and the ecosystem [14][15][16][17][18]. TiO 2 NP toxicity may be due to the ease with which these NPs can pass through the cellular membranes and disrupt biological systems [19]. It has been suggested that the small size and corresponding large specific surface area are the major determinants of NP toxicity [20]. It has also been proposed that the large surface area of NPs greatly increases their ability to produce potentially toxic ROS [21].
ROS are reactive molecules and free radicals derived from molecular oxygen. These molecules are produced as byproducts during the mitochondrial electron transport of aerobic respiration or by oxidoreductase enzymes and metal catalyzed oxidation, and have the potential to cause a number of deleterious events. ROS play a role in cellular signaling, including apoptosis, gene expression, and the activation of cell signaling cascades [22]. Oxidative DNA damage induced by ROS and free radicals is important in the pathogenesis of many human diseases, including cancer, muscle degeneration, coronary heart disease and ageing [23]. Moreover, studies have indicated that TiO 2 NPs induce photo-damage to DNA in human cells, mouse lymphoma cells, and phage [24][25][26].
Toll-like receptors (TLRs) play an essential role in the activation of innate immunity by recognizing specific molecular patterns of microbial components. TLRs are transmembrane proteins composed of both an extracellular domain (responsible for ligand recognition) and a cytoplasmic domain (required for initiating signaling) [27]. As suggested by their range of ligands and subcellular locations, TLRs recognize a wide range of foreign materials [28,29]. For example, TLRs that localize to the cell surface, such as TLR4, primarily recognize bacterial components. In contrast, TLRs that localize to the endocytic compartments, such as TLR3, mainly recognize viruses. We have previously shown that TLRs are also involved in the cellular response and cellular uptake of TiO 2 NPs [30,31]. We have also shown that the exposure of HepG2 cells to TiO 2 NPs induces DNA damage responses; this damage was increased by TLR4 overexpression, and decreased by TLR3 over-expression [32]. TLR3, which has a subcellular location distinct from TLR4, reduced the DNA damage response caused by TiO 2 NPs. These results suggested that TLRs could be involved in many cellular responses, including genotoxicology. However, the molecular mechanism of DNA damage induced by TiO 2 NPs is unknown.
In the present study, we aimed to elucidate the molecular mechanism of DNA damage induced by TiO 2 NPs by using PCR array and real-time PCR (RT-PCR). Specifically, we examined the effect of TiO 2 NP exposure on gene expression changes in DNA damage signaling pathways involving apoptosis, cell-cycle arrest, and DNA repair. Furthermore, we also confirmed elevated ROS generation by demonstrating increased hydrogen peroxide (H 2 O 2 ) levels, decreased glutathione peroxidase (GPX) and glutathione (GSH) levels, as well as caspase-3 activation, in cells exposed to TiO 2 NPs with or without TLR3 and TLR4 over-expression. We expect our work to advance the understanding of the molecular mechanism of DNA damage induced by TiO 2 NPs.

Results
In general, ROS generation results in DNA damage. Our aim was to examine TiO 2 NP-induced ROS generation and its association with DNA damage responses. ROS generation was measured in HepG2 cells exposed to TiO 2 NPs with and without TLR3 or TLR4 overexpression ( Figure 1). The results showed that in TiO 2 Figure 1 Reactive oxygen species (ROS) generation in TiO 2 NP-exposed HepG2 cells with and without TLR3 or TLR4 transfection. The transfected cells were exposed to 10 μg/ml TiO 2 NPs for 48 h. Each plot was produced from at least 3 replicate measurements. All values are presented as mean ± S.D. (n ≥3), (*P <0.05). NP-exposed cells, ROS levels were significantly increased (approximately 1.9 fold) compared with control cells (untreated, untransfected HepG2 cells). Cells exposed to TiO 2 NPs and over-expressing TLR4 showed a significant increase in ROS levels compared to untransfected cells exposed to TiO 2 NPs, reaching an approximately 2.6 fold increase compared to control. In comparison, ROS levels in HepG2 cells exposed to TiO 2 NPs with TLR3 over-expression were slightly (≈1.3 fold) increased compared to control cells, as shown in Figure 1.
These results indicate that ROS generation is a factor in the DNA damage response induced by TiO 2 NPs.
Oxidative stress reflects an imbalance between the systemic manifestation of reactive oxygen species and a biological system's ability to readily detoxify the reactive intermediates. Certain oxidant and antioxidant parameters were evaluated in order to obtain information on the cellular mechanism of oxidative stress in response to TiO 2 NP exposure, as well as the effect of TLR3 or TLR4 over-expression. H 2 O 2 is a reactive oxygen metabolic byproduct that serves as a key regulator of a number of oxidative stress-related states. Measurement of this reactive species provides an indication of the modulation of intracellular pathways by oxidative stress during TiO 2 NP exposure and TLR3 and TLR4 overproduction. In the present study, H 2 O 2 concentrations were elevated 1.9 fold in cells exposed to TiO 2 NPs compared to control cells. H 2 O 2 concentration was further increased in cells expressing TLR4 (3.2 fold), whereas cells expressing TLR3 exhibited only a 1.4 fold increase ( Figure 2). Our data show that H 2 O 2 is an intermediate in TiO 2 NP-induced oxidative stress, regardless of TLR3 or TLR4 over-expression.
GPX catalyzes the reduction of hydroperoxides, including H 2 O 2 , using GSH tripeptide as a hydrogen donor. In order to confirm the elevated H 2 O 2 levels, we measured GPX activity and GSH levels. The results showed that TiO 2 NP treatment inhibited GPX enzyme activity in HepG2 cells by 1.6 and 3.6 fold in the absence and presence of TLR4 transfection, respectively, when compared to control cells. Over-expression of TLR3 with TiO 2 NP exposure resulted in a 1.3 fold decrease in GPX activity compared to control cells ( Figure 3). Similarly, TLR4 overexpression exacerbated TiO 2 NP-induced reductions in GSH levels, while TLR3 over-expression appeared to reduce the effects of TiO 2 NP exposure ( Figure 4). These results showed that reduced GPX detoxification of H 2 O 2 is involved in the oxidative stress response stimulated by TiO 2 NPs, regardless of TLR3 or TLR4 over-expression, and confirmed the accumulation of H 2 O 2 due to inhibition of the antioxidants GPX and GSH.
Caspase-3, an effector cysteine protease involved in apoptosis and necrosis, is activated by H 2 O 2 [33]. Therefore, monitoring caspase-3 activation is important for evaluating apoptotic responses to oxidative stress occurring in HepG2 cells exposed to TiO 2 NPs. Our results showed that caspase-3 activity significantly increased in cells exposed to TiO 2 NPs, and that TLR4 expression further increased caspase-3 activation. Conversely, TLR3 expression resulted in an almost complete reversal of TiO 2 NP-induced caspase-3 activation ( Figure 5). Therefore, TiO 2 NP-treatment induces apoptosis involving Figure 2 Hydrogen peroxide (H 2 O 2 ) levels in TiO 2 NP-exposed HepG2 cells with and without TLR3 or TLR4 transfection. The transfected cells were exposed to 10 μg/ml TiO 2 NPs for 48 h. Each plot was produced from at least 3 replicate measurements. All values are presented as mean ± S.D. (n ≥3), (*P <0.05).

Figure 3
Glutathione peroxidase (GPx) activities in TiO 2 NP-exposed HepG2 cells with and without TLR3 or TLR4 transfection. The transfected cells were exposed to 10 μg/ml TiO 2 NPs for 48 h. Each plot was produced from at least 3 replicate measurements. All values are presented as mean ± S.D. (n ≥3), (*P <0.05). elevated caspase-3 activity in TLR4-expressing and normal HepG2 cells.
We used PCR array and RT-PCR to assess the cellular mechanisms operating in response to TiO 2 NP exposure in the presence of TLR4 over-expression. The genes upregulated by greater than 1-fold are listed in Table 1, and consist of genes in the human DNA damage signaling pathways (Table 1). In particular, the expression of the genes for apurinic/apyrimidinic exonuclease 1 (APEX1), ataxia telangiectasia mutated (ATM), growth arrest and DNA-damage-inducible, alpha (GADD45A), inositol hexakisphosphate kinase 3 (IP6K3), methyl-CpG binding domain protein 4 (MBD4), and structural maintenance of chromosomes 1A (SMC1A) were increased by >1.5 fold, with the remainder of the genes in the PCR array exhibiting <1.5 fold changes. In order to confirm the induction of the above-mentioned genes, the mRNA induction levels were determined by real-time PCR. The real-time PCR results confirmed the induction of mRNA expression observed for each of the genes ( Figure 6). Indeed, the real-time PCR results indicated that the genes induced to the greatest extent were ATM and IP6K3, which is consistent with double-strand breaks in the DNA that result in DNA fragmentation and apoptosis [34].
To confirm the enhancement of apoptosis in HepG2 cells, we used Hoechst DNA staining to observe nuclear fragmentation as an indication of apoptosis. As shown in Figure 7, morphological changes consistent with cellular apoptosis, including condensation of chromatin and nuclear fragmentation, were observed in the cells exposed to TiO 2 NPs. Again, expression of TLR4 ( Figure 7C) appeared to enhance the effects of TiO 2 NPs ( Figure 7B) on apoptosis. Microscopy analysis confirmed that cells exposed to TiO 2 NPs and transfected with TLR4 undergo programmed cell death (apoptosis) because of DNA damage. HepG2 cells that did not express TLRs and that were not exposed to TiO 2 NPs also underwent apoptosis: by counting the number of apoptotic cells, we determined that 28% of untreated, untransfected cells had fragmented nuclei ( Figure 7A), 55% of HepG2 cells exposed to TiO 2 NPs were apoptotic ( Figure 7B), and 75% of cells over-expressing TLR4 and exposed to TiO 2 NPs were apoptotic ( Figure 7C).

Discussion
The purpose of this study was to examine the molecular mechanism of DNA damage caused by exposure to TiO 2 NPs (10 μg/ml). A high concentration of TiO 2 NPs should amplify the effects of the NPs and thus aid examination of their mechanism of action. The interactions of NPs with cells resulted in the generation of ROS, and the resultant oxidative stress may cause DNA fragmentation [35,36]. We found a significant increase in ROS generation in cells exposed to TiO 2 NPs, which is consistent with our previous report of DNA damage responses in TiO 2 NP-exposed cells [32]. In this paper, the results indicated that TiO 2 NPs induced oxidative stress in cells, which can cause oxidative DNA damage and lead to the activation of p53 tumor suppressors and bcl-2 apoptotic factors. Additionally, oxidative stress can affect the mitochondria, the richest source of ROS, in which oxygen is metabolized and converted to O 2 − by several Reduced glutathione (GSH) levels in TiO 2 NP-exposed HepG2 cells with and without TLR3 or TLR4 transfection. The transfected cells were exposed to 10 μg/ml TiO 2 NPs for 48 h. Each plot was produced from at least 3 replicate measurements. All values are presented as mean ± S.D. (n ≥3), (*P <0.05).

Figure 5
Caspase-3 activities in TiO 2 NP-exposed HepG2 cells with and without TLR3 or TLR4 transfection. The transfected cells were exposed to 10 μg/ml TiO 2 NPs for 48 h. Each plot was produced from at least 3 replicate measurements. All values are presented as mean ± S.D. (n ≥3), (*P <0.05).
components of the mitochondrial respiratory chain (Figure 8). TLRs recognize and respond to exogenous and endogenous ligands through signaling pathways, leading to inflammatory cascade mediator production, which directs the innate and adaptive immune responses. TLRs interact with microbial components, such as lipopeptides, and non-self nucleic acids [37]. TLR4 localizes to the cell surface and TLR3 localizes in the endosome. We have shown that TLR4 is involved in TiO 2 NP-induced inflammatory responses and TiO 2 NP-incorporation [38,39]. We also have shown that TLR3 and TLR4 are involved in DNA damage induced by TiO 2 NPs, indicating that TLR3 reduces DNA damage while TLR4 exacerbates it [32]. In this paper, our results showed more significant effects in HepG2 cells exposed to TiO 2 NPs with TLR4 overexpression due to increased TiO 2 NP uptake into the cytoplasm and increased signal transduction involving ROS in TLR4-dependent activation of NF-kB [40], while TLR3 reduced the effects caused by TiO 2 NPs.
Various endogenous and exogenous genotoxic insults induce DNA-damage checkpoint signaling. The biological outcomes of checkpoint signaling include the control and coordination of cell-cycle progression, transcription, DNA replication, DNA repair, and apoptosis. DNA lesions trigger the activation of various kinases, which constitute the primary transducers in the signaling cascade. Of utmost importance are the phosphoinositide-3-kinase-related protein kinase (PIKK) family members, ATM, ATR, and DNA-dependent protein kinase. While ATR activation is associated with single-stranded DNA and stalled DNA replication forks, ATM and DNA-dependent protein kinase respond mainly to DNA double-strand breaks (DSBs) [34]. To identify marker genes of DNA damage-related cytotoxic stimulation, PCR array and RT-PCR analysis were performed using a commercial array system. Our results showed the induction of six genes, as follows: (1) APEX1 gene: a multifunctional DNA repair enzyme that plays a central role in the cellular response to oxidative stress. The major function of APEX1 in DNA repair

Figure 6
Expression of DNA damage marker mRNAs in TiO 2 NP-exposed HepG2 cells. Cells were exposed to 10 μg/ml TiO 2 NPs for 48 h. Results are shown as the mean ± SD, n ≥3 for each marker, (*P <0.05).  N-glycosylase activity involved in DNA repair. They can also remove uracil or 5-fluorouracil in G:U mismatches. (6) Finally, the sixth gene is SMC1A; the encoded protein is thought to be an important part of functional kinetochores. This protein interacts with BRCA1 and is phosphorylated by ATM, indicating a potential role for this protein in DNA repair. These data suggest that these six genes are useful markers for DNA damage signaling pathways in response to TiO 2 NP exposure, with the highest induction observed with ATM and IP6K3, as illustrated in Figure 6. Given that ATM and IP6K3 gene induction are involved in DSBs [34], the type of DNA damage induced by TiO 2 NP exposure is likely DNA DSBs that cause eventual DNA fragmentation and apoptosis.

Conclusions
Our results showed that exposing HepG2 cells to TiO 2 NPs enhances ROS generation and activates caspase-3 and oxidative stress-induced apoptosis. These effects were increased by TLR4 over-expression and decreased by TLR3 over-expression. We conclude that exposure to TiO 2 NPs causes oxidative stress, with increased H 2 O 2 and · OH levels leading to DNA damage and p53 activation, and induces apoptosis by releasing cytochrome c into the cytoplasm and activating caspase-3. Overexpression of TLR3 protects against oxidative stressinduced damage in response to TiO 2 NP exposure, but over-production of TLR4 enhances the oxidative stress mediated by TiO 2 NPs. TiO 2 NPs induce the expression of 17 DNA damage marker genes, especially the ATM and IP6K3 genes. This indicates that the type of DNA damage caused in HepG2 cells is double strand breaks, as well as chromatin condensation, nuclear fragmentation, and apoptosis.

Plasmids employed
TLR-encoding genes were purchased from InvivoGen (San Diego, CA, USA). The pUNO1-mcs expression vector was used as an "empty" control vector. Since pUNO1-mcs does not contain a therapeutic gene, it can be used in conjunction with other vectors of the pUNO1 family to serve as an experimental control. Overproduction of TLR3 and TLR4 was provided by transfection with pUNO-hTLR3 (which encodes the human TLR3 protein) and pUNO1-hTLR04a (CD284a) (which harbors the human TLR04a (CD284a) encoding open reading frame), respectively. HepG2 cells were seeded in 6-well plates. After overnight incubation, the cells were cotransfected with TLR3 or TLR4 expression vectors and control plasmid (pUNO1-mcs) using Lipofectamine™ LTX Reagent (Invitrogen, Carlsbad, CA, USA) according to the supplier's protocol. Transfection efficiency of at least 50% was obtained.

Preparation and exposure to TiO 2 NPs
The preparation and characterization of TiO 2 NPs were described in previous studies [26]. Briefly, nano-TiO 2 (AeroxideR P25; Sigma-Aldrich, St Louis, MO, USA) was dispersed in distilled water and autoclaved at 120°C for 20 min. The suspension was cooled to room temperature and then sonicated for 10 min at 200 kHz using a high-frequency ultrasonic sonicator (MidSonic 600, Kaijo Corp., Tokyo, Japan). The resulting nano-TiO 2 suspension was designated "TiO 2 NPs". The concentration of TiO 2 NPs was determined using a UV-vis spectrophotometer at 370 nm (UV-1600, Shimadzu, Kyoto, Japan). The suspension was adjusted to the desired concentration by the addition of distilled water and stored at 4°C until use. The particle size distribution was measured by dynamic light scattering (Zetasizer Nano-ZS, Malvern Instruments, Malvern, UK). The aggregated particle size of the TiO 2 NPs was determined to be 216 ± 70 nm. The size of the aggregated TiO 2 NPs remained stable for several weeks under the indicated storage conditions. Prior to addition to the cell cultures, the suspension of TiO 2 NPs was diluted with supplemented medium and used as described above. For the reporter gene (transfected cell) assays, the culture medium was replaced (1 day after transfection) with medium containing the TiO 2 NPs at the indicated concentration. Specifically, TiO 2 NPs were added to the culture medium immediately before the medium was applied to the cells. After 48 h, the cells were harvested and assayed.

DCF assay for oxidative stress determination
The accumulation of intracellular free radicals was quantified using a ROS assay kit (OxiSelect, Cell Biolabs, Inc., San Diego, CA, USA), which employs the cell-permeable fluorogenic probe 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA). DCFH-DA can cross cell membranes and be deacetylated by intracellular esterases to non-fluorescent 2′, 7′-dichlorodihydrofluorescein (DCFH). In the presence of ROS, DCFH is rapidly oxidized to the highly fluorescent DCF, which is readily detectable. The fluorescence intensity is proportional to the ROS levels in the cell cytosol. HepG2 cells were cultured in 96-well black plates; after overnight incubation, the cells were co-transfected with TLR3, TLR4 or control plasmid (pUNO1-mcs). After 24 h, the cells were exposed to TiO 2 NPs for 48h, and were then incubated with DCHF-DA for 30 min in the dark. Parallel sets of wells containing freshly cultured cells, which were not treated with NPs or plasmids, and were suspended in the same concentration ratio of DPBS and DMEM, were regarded as negative controls. The fluorescence emission of DCF was monitored at regular intervals at an excitation wavelength of 480 nm and an emission wavelength of 530 nm using a fluorescence plate reader (Twinkle LB 970 Microplate Fluorometer, BERTHOLD TECHNOLOGIES GmbH & Co. KG, Calmbacher, Bad Wildbad Germany). The amount of DCF formed was calculated from a calibration curve constructed using an authentic DCF standard.

Measurement of H 2 O 2
The levels of hydrogen peroxide (H 2 O 2 ) were measured using a hydrogen peroxide assay kit (ab102500, Abcam, Tokyo, Japan

Measurement of GPX
Glutathione peroxidase activity was measured using a glutathione peroxidase assay kit provided by Cayman Chemical Company (Ann Arbor, MI, USA). Cells were washed in phosphate buffer, pH 7.4, collected by centrifugation (2000 × g for 10 min at 4°C), then homogenized in cold assay buffer (50 mM Tris-HCL, pH 7.5, 5 mM EDTA, 1 mM DTT). Following centrifugation at 10,000 × g for 15 min at 4°C, the supernatant was removed for assay. Sample (20 μl of supernatant) was added to the desired well of a 96-well plate, then 100 μL of assay buffer and 50 μl of co-substrate mixture was added. The reaction was initiated by adding 20 μl of cumene hydroperoxide to each reaction well, then mixed by shaking for second. The absorbance was read at 340 nm using a plate reader. At least 5 time points were obtained.

Measurement of GSH
The total glutathione concentration (reduced and oxidized forms) was determined in a microtitre plate assay using a glutathione assay kit (Sigma-Aldrich). After TLR transfection and nanoparticle exposure, HepG2 cells were washed twice with phosphate-buffered saline (PBS), resuspended in a 5% 5-sulfosalicylic acid solution, then centrifuged at 10,000 × g for 10 min. Supernatant (10 μl) was mixed with 150 μl of working solution, incubated for 5 min at room temperature, then 50 μl of the diluted NADPH solution was added. The absorbance of each sample was measured at 412 nm using the plate reader, as was the absorbance of the reagent blank (10 μl of 5% 5-sulfosalicylic acid); the absorbance of the blank was then subtracted from the absorbance of each sample. The final concentration of the components in the reaction mixture was 95 mM potassium phosphate buffer, pH 7.0, containing 0.95 mM ethylenediamine tetra-acetic acid (EDTA), 0.038 mg/ml (48 μM) NADPH, 0.031 mg/ml DTNB, 0.115 units/ml glutathione reductase, and 0.24% 5-sulfosalicylic acid. All measurements were performed in triplicate; the concentration (nmoles) of GSH in the samples was calculated.

Measurement of caspase-3 activity
All reagents for assessing caspase-3 activity were provided in a caspase-3 colormetric assay kit, (Sigma Aldrich). HepG2 cells (1 × 10 6 ) were cultured in 6-well plates and treated as described above. At the end of the experiment, the cells were washed and lysed in 100 μl of lysis buffer provide in the kit, then 80 μl of the sample was added to 10 μl of the 10× assay buffer and 10 μl of in a well of a 96-well plate. The reaction mix was incubated for 10 hours at 37°C, then the absorbance was read at 405 nm.

Gene expression analysis: PCR array
For polymerase chain reaction (PCR) array analysis, HepG2 cells (at 6 × 10 5 cells/ml) with or without TLR4 transfection were seeded in a culture dish containing culture medium with or without TiO 2 NPs (suspended at 10 μg/ml). After 48 h exposure to the TiO 2 NPs, the cells were detached by mechanical dissociation and total cellular RNA was extracted using an RNeasy kit (Qiagen, MD, USA). An aliquot (1 μg) of the extracted total RNA was reverse transcribed into cDNA with random hexamer primers using a RT 2 First Strand kit (SABiosciences/ Quiagen MD, USA) and the expression of 89 human DNA damage-related genes involved in signaling pathways were examined using a RT 2 Profiler PCR array kit (SABiosciences/Quiagen) according to the manufacturer's instructions. PCR array analysis was performed using an ABI PRISM 7000 sequence detection system (Applied Biosystems, Singapore).

Real-Time (RT) PCR
For mRNA expression analysis, 6 × 10 5 HepG2 cells/ml were seeded in cell culture dishes, the cells were transfected with TLR4 expression vector and exposed to a suspension of TiO 2 NPs at a final concentration of 10 μg/ml for 48 h, then the cells were detached and subjected to gene expression analysis. The expression of marker genes was determined using quantitative real-time PCR (RT-PCR) as follows. Total RNA and cDNA were synthesized as described for the PCR array. The PCR primers for human APEX1, ATM, GADD45A, IP6K3, MBD4, SMC1A were purchased from SABiosciences/ Qiagen. The data were normalized using the housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as an endogenous control in the same reaction as the gene of interest [42]. The reaction mixture was composed of 12.5 μl of RT 2 SYBR Green qPCR Master Mix (SABiosciences;/Qiagen), 1 μl of 10 μM gene-specific RT 2 qPCR forward and reverse primers, 2 μl of cDNA, and nuclease-free water to a final volume of 25 μl. The thermocycling conditions were 95°C for 10 min, followed by 40 cycles of 95°C for 15 s and 60°C for 1 min.

Confocal microscopy observation
Confocal laser scanning microscopy was performed using a Zeiss LSM510 microscope (Carl Zeiss, Oberkochen, Germany). HepG2 cells were cultured on cover-slips (13 mm diameter; Matsunami Glass Ind., Ltd., Osaka, Japan). The following day, cultures were transfected (using the Lipofectamine™ LTX Reagent, as described above) with the expression vector encoding TLR4. At 24 h after transfection, the culture medium was replaced with medium containing 10 μg/ml TiO 2 NPs. Untransfected cells and cells without NP exposure were used as controls. After 48 h incubation, the cells were washed with PBS and fixed with 4% paraformaldehyde for 5 min. Fixed cells were then stained for nuclei using 1 μg/ml Hoechst33342 (Dojin Chemical, Japan) for 30 min in a 5% CO 2 environment. Figures were created using NIH ImageJ software.

Statistical analysis
Data were expressed as the mean ± SD, (n ≥3). All experiments were carried out independently. The data were analyzed using Student's t test to evaluate the significance of differences between the treated groups and control groups. Statistical significance was accepted at P < 0.05.