http://informahealthcare.com/txm ISSN: 1537-6516 (print), 1537-6524 (electronic) Toxicol Mech Methods, 2014; 24(3): 196–203 ! 2014 Informa Healthcare USA, Inc. DOI: 10.3109/15376516.2013.879505

RESEARCH ARTICLE

Evaluation of cellular effects of silicon dioxide nanoparticles

1

Institute of Industrial Ecological Sciences, University of Occupational and Environmental Health, Fukuoka, Japan, 2Health Research Institute (HRI), National Institute of Advanced Industrial Science and Technology (AIST), Ikeda, Osaka, Japan, 3National Metrology Institute of Japan, Higashi, Tsukuba, Ibaraki, Japan, 4Technology Research Association for Single Wall Carbon Nanotubes (TASC), Tsukuba, Ibaraki, Japan, 5Research Institute of Science for Safety and Sustainability (RISS), AIST, Tsukuba, Ibaraki, Japan, and 6Faculty of Applied Biological Sciences, Gifu University, Gifu, Japan Abstract

Keywords

Silica nanoparticles (nSiO2s) are an important type of manufactured nanoparticles. Although there are some reports about the cytotoxicity of nSiO2, the association between physical and chemical properties of nSiO2s and their cellular effects is still unclear. In this study, we examined the correlation between the physiochemical properties and cellular effects of three kinds of amorphous nSiO2s; sub-micro-scale amorphous SiO2, and micro-scale amorphous and crystalline SiO2 particles. The SiO2 particles were dispersed in culture medium and applied to HaCaT human keratinocytes and A549 human lung carcinoma cells. nSiO2s showed stronger protein adsorption than larger SiO2 particles. Moreover, the cellular effects of SiO2 particles were independent of the particle size and crystalline phase. The extent of cell membrane damage and intracellular ROS levels were different among nSiO2s. Upon exposure to nSiO2s, some cells released lactate dehydrogenase (LDH), whereas another nSiO2 did not induce LDH release. nSiO2s caused a slight increase in intracellular ROS levels. These cellular effects were independent of the specific surface area and primary particle size of the nSiO2s. Additionally, association of solubility and protein adsorption ability of nSiO2 to its cellular effects seemed to be small. Taken together, our data suggest that nSiO2s do not exert potent cytotoxic effects on cells in culture, especially compared to the effects of micro-scale SiO2 particles. Further studies are needed to address the role of surface properties of nSiO2s on cellular processes and cytotoxicity.

Cytotoxicity, lactate dehydrogenase, nanoparticle, oxidative stress, silica

Introduction Silicon dioxide nanoparticles (nanosilica; hereafter abbreviated as ‘‘nSiO2s’’) are one of the important manufactured nanoparticles with applications in various industrial products such as paint, ink, toner, adhesive, sealer, water-absorptive polymers, and cosmetics. There are two types of crystalline phases in manufactured SiO2; crystalline silica and amorphous silica. It is well known that crystalline SiO2 has hazardous properties. Inhalation of crystalline SiO2 leads to silicosis and lung cancer via induction of oxidative stress and inflammation (Donaldson & Borm, 1998; Peretz et al., 2006). SiO2 also exerts cytotoxic effects (Lesur et al., 1992). Pulmonary toxicity induced by amorphous SiO2 induces pulmonary toxicity to a lower extent than does crystalline SiO2. Although inhalation of amorphous SiO2 at concentrations of 25 mg/m3 induces pulmonary injury, the toxicity is transient (Arts et al., 2007). In the past decade, production of nSiO2s has been Address for correspondence: Masanori Horie, Institute of Industrial Ecological Sciences, University of Occupational and Environmental Health, 1-1 Iseigaoka, Yahata-Nishi, Kitakyushu, Fukuoka 807-8555, Japan. Tel: 81936917466. E-mail: [email protected]

History Received 26 September 2013 Revised 26 December 2013 Accepted 26 December 2013 Published online 27 January 2014

increasing. In most cases, the crystalline phase of the manufactured nSiO2s is amorphous. Although the toxicity of amorphous silica is less potent than that of crystalline silica, many reports have described the cytotoxicity of nSiO2s. In certain instances, nSiO2s with a particle size of 15 nm had stronger cytotoxicity than crystalline SiO2 (Min-U-Sil 5). In addition, the nSiO2s also induces oxidative stress in A549 cells (Lin et al., 2006). Amorphous nSiO2 causes the release of interleukin (IL)-6 and IL-8, reactive oxygen species (ROS) production, and induction of apoptosis in submucosal cells (McCarthy et al., 2012). Well-dispersed and bovine serum albumin (BSA)-stabilized amorphous nSiO2s induce ROS generation in A549 cells (Foldbjerg et al., 2013). Additionally, some studies have also indicated that amorphous nSiO2s induce oxidative stress in cells (Berg et al., 2013; Nabeshi et al., 2011; Napierska et al., 2012). Oxidative stress-independent cellular effects of amorphous nSiO2 have also been reported (Gehrke et al., 2013). Induction of oxidative stress is one of the most important cellular effects of nanoparticles. Oxidative stress promotes other cellular processes such as apoptosis and damage to cell membranes.

20 14

Toxicology Mechanisms and Methods Downloaded from informahealthcare.com by University of Laval on 06/18/14 For personal use only.

Masanori Horie1,2, Keiko Nishio2, Haruhisa Kato3, Shigehisa Endoh4, Katsuhide Fujita5, Ayako Nakamura3, Yoshihisa Hagihara2, Yasukazu Yoshida2, and Hitoshi Iwahashi2,6

Cellular influences of silica nanoparticles

Toxicology Mechanisms and Methods Downloaded from informahealthcare.com by University of Laval on 06/18/14 For personal use only.

DOI: 10.3109/15376516.2013.879505

Some metal oxide nanoparticles are also cytotoxic in that they induce oxidative stress. Metal ion release, adsorption ability, cellular uptake, and surface activity are important factors that affect the cytotoxic potential of nanoparticles (Horie et al., 2012a). Among these factors, metal ion release is considered the most important. Generally, cytotoxicity of soluble metal oxide nanoparticles such as ZnO, CuO, NiO, and Cr2O3 is impotent (Cronholm et al., 2013; Horie et al., 2009, 2012a, 2013). These soluble metal oxide nanoparticles induce oxidative stress and subsequently lead to cell death, including apoptosis (Horie et al., 2013). The soluble nanoparticles are taken up into cells by endocytosis, and the internalized particles release metal ions inside the cell, resulting in many cytotoxic effects (Horie et al., 2009, 2013). Cellular effects of insoluble nanoparticles are less potent than those of soluble nanoparticles, which are not insignificant. Another important factor that impacts the cytotoxic potential of nanoparticles in their capacity for adsorbing macromolecules, particularly, is their protein adsorption capacity. Protein adsorption affects the cellular uptake of the nanoparticles (Lesniak et al., 2012). Therefore, understanding the factors that modulate the cellular effects of nanoparticles is important for evaluation of their toxic activity, from the perspective of both the industrial applications of nSiO2s and their effects on the environment and human health. The purpose of the present study is to compare the cell influence by some kinds of nSiO2s. Especially, induction of oxidative stress which is important cellular response caused by nanoparticles exposure was examined. And we investigated the association between these effects and the physical and chemical properties of the nSiO2s, particularly, solubility and protein adsorption ability.

Materials and methods Cell culture HaCaT human keratinocytes were purchased from the German Cancer Research Center (DKFZ, Heidelberg, Germany). A549 human lung carcinoma cells were purchased from the RIKEN BioResource Center (Tsukuba, Ibaraki, Japan). The cells were cultured in Dulbecco’s modified Eagle medium (DMEM; Gibco, Invitrogen Corporation, Gland Island, NY) supplemented with 10% heat-inactivated fetal bovine serum (FBS; CELLect GOLD; MP Biomedicals Inc., Solon, OH), 100 units/mL of penicillin, 100 mg/mL of streptomycin, and 250 ng/mL of amphotericin B (Nacalai Tesque Inc., Kyoto, Japan). In this study, this DMEM cocktail

197

is referred to as ‘‘DMEM-FBS’’. The cultures were placed in a 75-cm2 flask (Corning Incorporated, Corning, NY) and incubated at 37  C in an atmosphere of 5% CO2. For cell viability assays, the cells were seeded in 96-well or six-well plates (Corning Incorporated) at 1  105 cells/mL and incubated for 24 h; subsequently, they were removed from the culture medium, subjected to metal oxide dispersion, and incubated for another 24 h. SiO2 particles and preparation of SiO2 medium dispersions The SiO2 particles used in this study are listed in Table 1. Three kinds of nSiO2s were purchased from Denki Kagaku Kogyo Kabushiki Kaisha (Tokyo, Japan), Tokuyama Corporation (Tokyo, Japan), and CIK NanoTek Corporation (Tokyo, Japan). Fine SiO2 particles were purchased from Denki Kagaku Kogyo Kabushiki Kaisha. Micro-scale amorphous SiO2 particles were purchased from Nippon Steel & Sumikin Materials Co., Ltd. (Micron; Himeji, Japan) and micro-scale crystalline SiO2 particles were purchased from U.S. Silica Company (Berkeley Springs, WV). For the cytotoxicity assays, SiO2 powder was dispersed directly in DMEM-FBS at concentrations of 10, 100, and 1000 mg/mL without a dispersant. For some experiments, stable and uniform dispersions were prepared using a previously described pre-adsorption and centrifugation method (Horie et al., 2010a). Nanoparticles can induce nonspecific effects on cells by medium depletion, because of their protein adsorption ability (Horie et al., 2009). To prevent cell starvation caused by adsorption of the medium components onto the particle surface, SiO2 particles were initially dispersed in FBS at a concentration of 40 mg/mL. Subsequently, the dispersion was centrifuged at 16 000  g for 20 min. The precipitated SiO2 particles were washed once with FBS-free DMEM and re-dispersed in an equivalent volume of fresh DMEM-FBS. This dispersion of ultrafine particles in DMEM-FBS was centrifuged at 8000  g for 20 min. After discarding the supernatant, the precipitate was re-suspended in an equal volume of fresh DMEM-FBS, and the resulting SiO2 dispersion was centrifuged at 4000  g for 20 min. The supernatant was collected as the stable SiO2 dispersion. Characterization of SiO2-DMEM-FBS dispersions In this study, we defined ‘‘secondary particles’’ as complex aggregates of primary particles, proteins from FBS, and other medium components. In addition, the ‘‘average particle size’’

Table 1. Properties of the SiO2 particles used in this study.

Matrial Silicon dioxide

a

Product name

Average primary perticle size (nm)a

Specific surface area (m2/g)a

Amorphous

UFP-80 REOLOSIL; QS-30 NanoTek SFP-20 M HS-301

34 7 25 300 2400

80.0 300 86.0 11.3 8

Crystalline

MIN-U-SIL 5

1800

Code in thes study

Crystalline structure

nSiO2-1 nSiO2-2 nSiO2-3 fSiO2 mSiO2 mcSiO2

These data are according to the manufacturer’s data sheet.

Vendor Denki Kagaku Kogyo Kabushiki Kaisha Tokuyama Corporation CIK NanoTek Corporation Denki Kagaku Kogyo Kabushiki Kaisha Nippon Steel & Sumikin Materials Co., Ltd. (Micron) U.S. Silica Company

Toxicology Mechanisms and Methods Downloaded from informahealthcare.com by University of Laval on 06/18/14 For personal use only.

198

M. Horie et al.

was defined as the size of the secondary particles estimated from light scattering intensity measurements made under the assumption that the aggregates were globular. SiO2 DMEM-FBS dispersions prepared by the above-mentioned methods were divided into three aliquots that were used for simultaneous biological studies, and measurement of calcium concentration and particle size. The secondary particle sizes in the SiO2 DMEM-FBS dispersions were measured by dynamic light scattering (DLS), as described previously (Kato et al., 2010). The estimated diameters of the secondary particles were calculated as the average of three measurements at different wavelengths obtained with the following devices: a DLS-7000 spectrophotometer (633 nm; Otsuka Electronics Co. Ltd., Hirakata, Japan); an FPAR-1000 fiberoptics particle analyzer (660 nm; Otsuka Electronics Co. Ltd.); and Nanotrac (780 nm; Nikkiso Co. Ltd., Tokyo, Japan). Undiluted dispersions were used for the above measurements. SiO2 concentration was measured by X-ray fluorescence (XRF) analysis. For this analysis, 13 mL of the SiO2 DMEMFBS dispersion was added to 13 mL of a standard solution containing 0.1 mg/mL of iron as an internal standard element and mixed thoroughly. Next, 5 mL of the mixture was dried in an oven at 200  C for 24 h. The dried sample was ground in an agate mortar and subjected to XRF analysis using a dispersive XRF spectrometer (JSX-3201; JEOL Ltd., Tokyo, Japan). The amount of calcium was estimated from the molar ratio of SiO2 and the internal standard. The values of SiO2 were corrected by background concentration for the SiO2 in the medium. Although DMEM does not include Si and its compounds, background Si concentration in the DMEM-FBS was estimated as 0.31 mM. Source of the background Si is unknown. Determination of protein and calcium adsorption abilities of the SiO2 particles SiO2 powder was dispersed directly in DMEM-FBS at concentrations of 10 and 100 mg/mL and then the dispersion was centrifuged at 16 000  g for 20 min. The supernatant was collected carefully for the determination of protein and Ca2+ concentrations. The protein content of the supernatant was determined using a BCA protein assay kit (Thermo Fisher Scientific Inc., Rockford, IL) with BSA as the standard. The Ca2+ content of the culture medium was determined using the methyl xylenol blue (MXB) method using a Calcium-E test Wako system (Wako Pure Chemical Industries, Osaka, Japan). DMEM without serum and Ca2+ were used as the blank. Measurement of mitochondrial activity and cell membrane damage Cells were seeded in a 96-well culture dishes (Corning) at 2  105 cells/well, incubated for 24 h, and the culture medium was replaced with the SiO2 DMEM-FBS dispersion. The cells were then incubated for 6 and 24 h. For determining the mitochondrial activity, a 3-(4,5-dimethyl-2-thiazolyl)-2,5diphenyltetrazolium bromide (MTT) assay was performed. Cells were incubated with 0.5 mg/mL MTT (Nacalai Tesque, Inc.) at 37  C for 2 h. Isopropyl alcohol containing 40 mM HCl was added to the culture medium (3:2, by volume), and

Toxicol Mech Methods, 2014; 24(3): 196–203

the samples were mixed by pipetting until the formazan was completely dissolved. The optical density of formazan was measured at 570 nm by using a Multiskan Ascent plate reader (Thermo LabSystems, Helsinki, Finland). For determining the extent of damage to the cell membrane, the amount of lactate dehydrogenase (LDH) released into the culture supernatant was determined. The LDH release was measured with a tetrazolium salt using the Cytotoxicity Detection KitPLUS for LDH (Roche Diagnostics GmbH, Mannheim, Germany) according to the manufacturer’s protocol. The amount of formazan salt formed was estimated by measuring the absorbance at 492 nm using a Multiskan Ascent plate reader. The maximum amount of LDH released was determined by incubating the cells with a lysis solution provided in the kit. Cytotoxicity was calculated as follows: cytotoxicity (%) ¼ [(experimental value  low control)/(high control  low control)]  100. The low control, which refers to spontaneous LDH release, was determined as the LDH released from untreated normal cells. The high control, which refers to the maximum LDH release, was determined as the LDH released from cells lysed by treatment with a surfactant. Measurement of intracellular oxidative stress Intracellular ROS levels were detected using dichlorofluorescein diacetate (DCFH-DA; Sigma-Aldrich, St. Louis, MO). DCFH-DA was dissolved in dimethyl sulfoxide at a concentration of 5 mM as a stock solution and stored at 20  C; for use in further experiments, the stock solution was diluted 500 times with serum-free medium. The cells were exposed to the SiO2 dispersion for 2, 6, 12, and 24 h. The dispersion was then exchanged with serum-free DMEM that included 10 mM DCFH-DA, and the cells were incubated for 30 min at 37  C. Cells were then washed once with phosphate-buffered saline (PBS), treated with 0.25% trypsin, washed once with PBS, and resuspended in 500 mL of PBS. The cell samples in PBS were excited with a 488-nm argon ion laser in a Cytomics FC500 flow cytometry system, and the emission of 20 ,70 dichlorofluorescein (DCF) was recorded at 525 nm. Data were collected from at least 5000 gated events. Intracellular hydroperoxide was detected using diphenyl1-pyrenylphosphine (DPPP; Dojindo Laboratories, Kumamoto, Japan). DPPP was dissolved in DMSO at 5 mM as a stock solution and stored at 20  C. After incubation of cells for another 24 h with the nSiO2 dispersion, the medium was changed to serum-free DMEM containing 50 mM of DPPP and incubated for 30 min at 37  C. Cells were then washed with PBS, harvested by trypsinization, washed again with PBS and resuspended in 3 ml of PBS. Cell samples in PBS were excited with a 351-nm argon laser. The emission of DPPP oxide was measured at 380 nm using an RF-5300PC spectrofluorophotometer (Shimadzu Corporation, Kyoto, Japan). After measurement, cells were collected and measured for protein concentration. DPPP oxide fluorescence was normalized by cellular protein concentration. Determination of caspase-3 activity To obtain total cell extracts, SiO2-treated cells were collected by treatment with 0.25% trypsin, washed with cold PBS, and

Cellular influences of silica nanoparticles

Statistical analysis Data are expressed as the mean ± SD of at least three independent repetitions. Analysis of variance (ANOVA) using Dunnett’s test was performed to compare the differences between data sets.

Results Protein and calcium adsorption abilities of SiO2 particles Protein and calcium adsorption abilities of nanoparticles are key factors that determine their cellular effects (Horie et al., 2012b). We examined the protein and calcium adsorption abilities of SiO2 particles that were dispersed in DMEM-FBS at concentrations of 10 and 100 mg/mL. The protein and calcium concentration in the supernatant was measured after the removal of particles from the dispersion by centrifugation (Figure 1). Neither protein nor calcium was adsorbed onto the micro-scale SiO2 particles (mSiO2s and mcSiO2s). The protein and calcium adsorption ability of fine SiO2 particles was not significant. Protein concentration of the DMEM-FBS decreased slightly with the 100 mg/mL SiO2 particle dispersion, whereas nSiO2s showed significant protein adsorption abilities (Figure 1A). Among the examined SiO2 particles, nSiO2-2 had the strongest protein adsorption. nSiO2s also adsorbed calcium to a slight extent (1B), while calcium adsorption abilities of SiO2 particles were not significant. Cytotoxicity of SiO2 particles: unstable dispersion SiO2 particles were dispersed in DMEM-FBS directly at concentrations of 10, 100, and 1000 mg/mL, and the dispersions were applied to cells. After 24 h, mitochondrial activity and cell membrane damage was determined. SiO2 particles decreased mitochondrial activity in a dose-dependent manner (Figure 2A). There was no significant difference in the changes in mitochondrial activity induced by different kinds of particles. The influence of SiO2 particles on the mitochondrial activity was not potent; even at 1000 mg/mL SiO2, mitochondrial activity was approximately 70% that of unexposed cells. Moreover, different SiO2 particles

(A) Protein concentration in DMEM-FBS (mg/ml)

resuspended in lysis buffer (150 mM NaCl, 50 mM Tris-HCl, pH 7.4), 50 mM NaF, 5 mM EDTA, 0.5% Triton X-100, and 1 mM Na3VO4, along with a protease inhibitor cocktail tablet [Roche Diagnostics GmbH], on ice for 10 min. Nuclei and unlysed cellular debris were removed by centrifugation at 10 000  g for 1 min. The protein concentration was determined using a BCA protein assay kit (Thermo Fisher Scientific Inc., Rockford, IL) with BSA as the standard. Caspase-3 activity was measured by cleavage of the AspGlu-Val-Asp (DEVD) peptide conjugated to 7-amino-4trifluoromethyl coumarin (AFC) according to the manufacturer’s instructions for the caspase-3 fluorimetric protease assay kit (Medical & Biological Laboratories Co., Ltd., Nagoya, Japan). Substrate cleavage, which resulted in the release of AFC fluorescence (excitation at 400 nm; emission at 505 nm), was measured using a Fluoroskan Ascent CF plate reader (Thermo Fisher Scientific Inc.).

4 3.5

Proteinconcentration of DMEM-FBS *

199

10 100 mg/ml

**

3 **

2.5

**

2 1.5

**

**

1 0.5

**

0 nSiO2-1 nSiO2-2 nSiO2-3 fSiO2

mSiO2 mcSiO2

(B) Calcium concentration in DMEM-FB (mM)

Toxicology Mechanisms and Methods Downloaded from informahealthcare.com by University of Laval on 06/18/14 For personal use only.

DOI: 10.3109/15376516.2013.879505

2.5 2

Calcium concentration of DMEM-FBS ** **

1.5

** **

** **

**

10 100 mg/ml

* **

**

1 0.5 0 nSiO2-1 nSiO2-2 nSiO2-3 fSiO2

mSiO2 mcSiO2

Figure 1. Protein and calcium adsorption ability of SiO2 particles. SiO2 particles were dispersed in DMEM-FBS at 10 and 100 mg/mL. After removing the particles by centrifugation, the protein (A) and calcium (B) concentrations in the DMEM-FBS were measured by BCA method and MXB method, respectively. The protein (3.2 mg/mL) and calcium (2.0 mM) concentrations in the DMEM-FBS are indicated by the broken lines. *p50.05 and **p50.01, versus DMEM-FBS, calculated using Dunnett’s one-way ANOVA post-hoc test.

induced varying extents of cell membrane damage, with some particles inducing significant extents of damage (Figure 2B). However, the cell membrane damage induced by nanoparticles was not particularly significant compared with the fine- and micro-scale particles. Among the nSiO2s, nSiO2-3 induced the largest amount of cell membrane damage, while the effect of nSiO2s overall was not as significant. Particularly, even at 1000 mg/mL nSiO2, nSiO2-2 did not damage the cell membrane in A549 cells. Next, the influence of SiO2 particles on intracellular ROS levels was examined. Exposure to 1000 mg/mL nSiO2-1, nSiO2-2, and fSiO2 particles caused an increase in intracellular ROS levels (Figure 3). When cells were exposed to 100 mg/mL SiO2 particles, intracellular ROS levels were unchanged. Cytotoxicity of SiO2 particles: stable dispersions When nSiO2s were directly dispersed in DMEM-FBS, the nanoparticles formed large secondary particles and accumulated onto cells by gravity sedimentation. Therefore, the concentrations of the nanoparticles in dispersion did not correctly reflect the concentration to which the cells were exposed. Therefore, we prepared stable nSiO2 dispersions in culture medium. The physiochemical properties of the nSiO2 dispersions are listed in Table 2. nSiO2-1 was stably dispersed in the DMEM-FBS, and the changes in secondary particle

200

Toxicol Mech Methods, 2014; 24(3): 196–203

HaCaT

A549

120 ** **

100 **

80

** ** ** **

**

**

* **

**

20

HaCaT

**

** **

40

mcSiO2

mSiO2

A549

** **

1000 µg/ml

** **

**

**

30 **

**

20

fSiO2

mSiO2

nSiO2-3

nSiO2-2

mcSiO2

nSiO2-1

mSiO2

fSiO2

nSiO2-3

nSiO2-2

nSiO2-1

0

mcSiO2

**

** **

10 **

Figure 2. Effect of SiO2 particles on mitochondrial activity and cell membrane. (A) Effect of the SiO2 DMEM-FBS dispersions on mitochondrial activity. SiO2 dispersions in DMEM-FBS were applied to HaCaT and A549 cells at concentrations of 10, 100, and 1000 mg/mL. After incubation for 24 h, mitochondrial activity was measured with the MTT assay and represented by the percentage of MTT conversion. The percentage of MTT conversion of the unexposed cells was set as 100%. (B) Evaluation of the cell membrane damage caused by SiO2 dispersions in DMEM-FBS. Cells were treated with SiO2 dispersions in DMEM-FBS for 24 h at concentrations of 100 and 1000 mg/mL and the extent of cell membrane damage was determined by measuring the release of intracellular LDH. *p50.05 and **p50.01, versus unexposed cells, calculated using Dunnett’s one-way ANOVA post-hoc test. HaCaT RelativeIntracellular ROS level (vs control)

3.5

A549

100

**

1000 µg/ml

3 2.5 2 1.5

**

**

**

nSiO2-2 and nSiO2-2 dispersions contained 0.89, 1.68 and 1.49 of soluble Si, respectively. Subsequently, their cellular effects were examined. Exposure of cells to the stable nSiO2 dispersions decreased mitochondrial activities, as measured by the MTT assay (Figure 4A). However, cell membrane damage caused by nSiO2-1 and nSiO2-2 exposures was not significant (Figure 4B). Both HaCaT and A549 cells exposed to nSiO2-3 exhibited significant amounts of cell membrane damage. Induction of oxidative stress by nSiO2s

100

** **

** **

50

fSiO2

nSiO2-3

nSiO2-2

nSiO2-1

mcSiO2

mSiO2

fSiO2

nSiO2-3

nSiO2-2

0

60 Cytotoxicity (%)

** **

40

(B) 70

**

1 0.5

The effect of stable nSiO2 dispersions on the intracellular ROS levels and lipid peroxidation levels were examined. Exposure of cells to nSiO2s for 24 h caused an increase in intracellular ROS levels (Figure 5A). In A549 cells, nSiO2-3 did not induce any changes in the intracellular ROS levels (Figure 5A). After 24 h, the intracellular ROS levels in the nSiO2-exposed cells were approximately 1.6-times that of unexposed cells. This result indicates that the nSiO2 induced slightly cellular oxidative stress. Additionally, exposure of cells to nSiO2s for 24 h also caused an increase in intracellular lipid peroxidation levels (Figure 5B). In HaCaT cells, the lipid peroxidation in the nSiO2-1 and nSiO2-2 exposed cells were approximately 2-times that of unexposed cells. The intracellular ROS levels in the nSiO2-3 exposed cells were approximately 5-times that of unexposed cells. In A549 cells, increase of lipid peroxidation level was observed only in the nSiO2-2 exposed cells. Induction of caspase-3 activity by nSiO2s To evaluate apoptosis-inducing ability of nSiO2s, caspase-3 activity, a marker for apoptosis, was determined (Figure 6). Exposure of cells to nSiO2 dispersions for 24 h increased caspase-3 activity in both HaCaT and A549 cells (Figure 6A). Particularly, the nSiO2-3 dispersion caused a significant induction of caspase-3 activity. Caspase-3 activities in nSiO2-3-exposed cells (exposed to 70.8 mg/mL nSiO2-3) were 10-times or even greater than that in unexposed cells (Figure 6B). When the concentration of nSiO2-3 was 35.4 mg/ mL or less, caspase-3 activity was not affected by exposure to nSiO2-3.

Discussion mcSiO2

mSiO2

fSiO2

nSiO2-3

nSiO2-2

nSiO2-1

mcSiO2

mSiO2

fSiO2

nSiO2-3

nSiO2-2

0 nSiO2-1

Toxicology Mechanisms and Methods Downloaded from informahealthcare.com by University of Laval on 06/18/14 For personal use only.

*

** **

10 100 1000 µg/ml

60

nSiO2-1

MTT conversion (% of control)

(A)

M. Horie et al.

Figure 3. Effect of nSiO2 medium dispersions on intracellular ROS levels. Cells were exposed to nSiO2 dispersions in DMEM-FBS for 24 h. Then, the medium was exchanged with fresh FBS-free DMEM containing 10 mM of DCFH-DA. After incubation for 30 min, cells were collected and washed. DCFH fluorescence in cells was measured by flow cytometry. The value of DCFH fluorescence in untreated control cells was set to 1 and the intensities in all other samples are represented as the relative fluorescence intensity. **p50.01, versus unexposed cells, calculated using Dunnett’s one-way ANOVA post-hoc test.

size and light intensity of the nSiO2-1 dispersion were small. On the other hand, the secondary particle size in the nSiO2-2 and nSiO2-3 dispersions could not be estimated because their size distribution curves had multiple peaks. The nSiO2-1,

In the present study, the cellular effects of five kinds of SiO2 particles, including three kinds of nanoparticles, were compared. The cellular effects of thee kinds of the nSiO2 were different. The primary particle size did not seem to relate directly to the cellular effects; for instance, similar results were obtained with the MTT assay regardless of the size of the particle. In addition, even if the primary particle size was nano-scale, different degrees of effects were observed with different nano-scale particles. nSiO2-1 and nSiO2-3 induced cell membrane damage, whereas nSiO2-2 did not. Moreover, nSiO2-3 induced potent damage to the cell membrane. The cytotoxic effects of nSiO2-3 were greater than that of micro-scale crystalline SiO2 particles, consistent with the results from another study (Lin et al., 2006). nSiO2s, with particle sizes of 46 and 15 nm, exerted stronger cytotoxic effects than crystalline SiO2 (Min-U-Sil 5) particles in A549

Cellular influences of silica nanoparticles

DOI: 10.3109/15376516.2013.879505

201

Table 2. Properties of nSiO2 dispersion. Inorganic salt concentration in the dispersion (mM) Concentration of SiO2 particles (mg/mL)

Na

P

K

Sia

nSiO2-1

53.3 ± 11.2

35.85 ± 5.07

0.77 ± 0.08

2.22 ± 0.12

0.89 ± 0.19

nSiO2-2 nSiO2-3

101.2 ± 12.0 70.8 ± 7.41

34.75 ± 4.24 31.99 ± 4.77

0.77 ± 0.11 0.69 ± 0.06

2.20 ± 0.25 2.21 ± 0.14

1.68 ± 0.20 1.49 ± 0.12

Size (nm) dl dn

u (nm)

utime (nm)

uapp (nm)

umethod (nm)

14.7 21.8

5.7 5.7

13.2 11.2

3.0 17.7

304.8 104.3

ND ND

a

Background Si concentration in the DMEM-FBS was 0.31 mM.

**

** **

**

**

**

nSiO2-2

nSiO2-2

nSiO2-3

53.3

101.2

50.6

70.8

53.3

101.2

50.6

70.8

**

35

**

15 *

*

0 SiO2 (µg/ml)

nSiO2-1

nSiO2-2

nSiO2-2

nSiO2-3

53.3

101.2

50.6

70.8

Figure 4. Effect of stable nSiO2 medium dispersions on mitochondrial activity and cell membrane. (A) Effect of the stable nSiO2 dispersions in DMEM-FBS on mitochondrial activity. nSiO2 dispersions in DMEMFBS were applied to HaCaT and A549 cells. After incubation for 6 and 24 h, mitochondrial activity was measured with the MTT assay. (B) Effect of the nSiO2 dispersions in DMEM-FBS on cell membrane damage. Cells were treated with nSiO2 dispersions in DMEM-FBS for 24 h and cell membrane damage was determined by measuring the release of intracellular LDH. *p50.05 and **p50.01, versus unexposed cells, calculated using Dunnett’s one-way ANOVA post-hoc test.

cells. Overall, the cellular effects of the SiO2 particles were not remarkable. When the effects of SiO2 were compared with those of metal oxide nanoparticles, according to the MTT assay, the cellular toxicities corresponded to those of SnO2 and ZrO2 nanoparticles (Horie et al., 2012a), and they were moderately greater than those of TiO2 and CeO2 (Horie et al., 2010a, 2011). The cellular effects of nSiO2s were less potent than those of soluble nanoparticles such as ZnO, CuO, and NiO nanoparticles (Horie et al., 2012a; Karlsson et al., 2008). Some cellular factors that determine the response to nanoparticles have been suggested in previous studies (Horie et al., 2012a,b); these include metal ion release, adsorption ability, cellular uptake, and surface activity. In the present study, nSiO2-2 and nSiO2-3 dissolved to some extent in DMEM-FBS. However, the influence of soluble Si on the

A549 **

**

6h 24h

**

nSiO2-1 nSiO2-2 nSiO2-3 nSiO2-1 nSiO2-2 nSiO2-3 SiO2 (µg/ml)

53.3 (B)

20

**

**

A549

25

5

HaCaT

2 1.8 1.6 1.4 1.2 1 0.8 0.6 0.4 0.2 0

HaCaT

30

10

(A)

Relativelipid peroxidation level (vs control)

nSiO2-1

**

(B) 40 Cytotoxicity (%)

** **

nSiO2-3

**

** **

24h

nSiO2-2

**

**

**

6h

nSiO2-2

SiO2 (µg/ml)

**

A549

RelativeIntracellular ROS level (vs control)

HaCaT 100 90 80 70 60 50 40 30 20 10 0

nSiO2-1

MTT conversion (% of control)

Toxicology Mechanisms and Methods Downloaded from informahealthcare.com by University of Laval on 06/18/14 For personal use only.

(A)

101.2

70.8

53.3

6

101.2

**

70.8

HaCaT A549

5 4 3

**

**

**

2 1 0

SiO2 (µg/ml)

nSiO2-1 53.3

nSiO2-2 101.2

nSiO2-3 70.8

Figure 5. Effect of stable nSiO2 medium dispersions on cellular oxidative stress. (A) Intracellular ROS level. Cells were exposed to nSiO2 dispersions in DMEM-FBS for 6 and 24 h. Then, the medium was exchanged with fresh FBS-free DMEM containing 10 mM of DCFH-DA. After incubation for 30 min, cells were collected and washed. DCFH fluorescence in cells was measured by flow cytometry. (B) Intracellular lipid peroxidation level. Cells were exposed to nSiO2 dispersions for 24 h. After incubation, the intracellular lipid peroxidation level was measured using DPPP. The value of DCFH and DPPP fluorescence in untreated control cells was set to 1 and the intensities in all other samples are represented as the relative fluorescence intensity. **p50.01, versus unexposed cells, calculated using Dunnett’s one-way ANOVA post-hoc test.

cells seemed small. Although Si concentrations in the nSiO2-2 and nSiO2-3 dispersion were similar, the LDH release of the cells exposed by nSiO2-2 and nSiO2-3 dispersion was significantly different. nSiO2s had a higher ability for protein adsorption than fine and micro-scale SiO2 particles. Although the nSiO2s had the largest protein adsorption ability, the induction of cell membrane damage was the lowest in nSiO2exposed cells, among the three kinds of SiO2 particles. These results suggest that protein adsorption is not the main factor that determines the cellular cytotoxicity of nSiO2s. In this study, we did not observe cellular uptake of nSiO2s. Many studies have shown that manufactured nanoparticles

202

Relative caspase-3 activity (vs control)

(A)

M. Horie et al.

Toxicol Mech Methods, 2014; 24(3): 196–203

HaCaT 18 16 14 12 10 8 6 4 2 0

A549

6h

**

**

24h

**

**

**

**

**

**

**

nSiO2-1 nSiO2-2 nSiO2-3 nSiO2-1 nSiO2-2 nSiO2-3 SiO2 (µg/ml)

Relative caspase-3 activity (vs control)

Toxicology Mechanisms and Methods Downloaded from informahealthcare.com by University of Laval on 06/18/14 For personal use only.

(B)

53.3 18 16 14 12 10 8 6 4 2 0

101.2

70.8

53.3

101.2

**

HaCaT

**

70.8

70.8

A549

35.4

14.2

7.1

Concentration of SiO2 (mg/ml)

Figure 6. Effect of stable nSiO2 medium dispersions on caspase-3 activity. (A) Cell lysate samples were prepared after treating cells with the nSiO2 dispersions in DMEM-FBS for 6 and 24 h. (B) Cell lysate samples were prepared after treating cells with serial dilutions of nSiO23 dispersion in DMEM-FBS for 24 h. Caspase-3 activity in the cell lysates was measured using DEVD-AFC as substrate as described in Materials and methods section. **p50.01, versus unexposed control cells, calculated using Dunnett’s one-way ANOVA post-hoc test.

and micro-scale particles are taken up into cells by endocytosis (Gehrke et al., 2013). The nSiO2s examined in the present study might have also been taken up into the cell. Therefore, differences in the cellular effects of nSiO2s, especially the cell membrane damage, are likely to be due to differences in their surface conditions. Furthermore, the extent of cell membrane damage was different depending on the kind of nSiO2. Among the tested nSiO2s, nSiO2-3 elicited the most potent cell membrane damage, likely an effect of the differences in the surface properties of the nSiO2s. nSiO2 is adhered to the cell membrane during the earlier stages of endocytosis. The adhesion of nanoparticles to cells affects to their cytotoxicity effects (Lesniak et al., 2013). The surface characteristics and adsorption of proteins are important to the adhesion of nanoparticles to cells (Lesniak et al., 2012; Nazli et al., 2012; Shinto et al., 2013). Large protein adsorption abilities of nSiO2s may impinge on their cellular uptake. Surface modifications of nSiO2s also affect their cellular effects (Yoshida et al., 2012). The manufacturing methods of the nSiO2 in the present study were different. According to the descriptions provided by the manufacturer, nSiO2-1 was fused silica that was produced by a fusion method, while nSiO2-2 was fumed silica that was produced by hydrolysis of SiCl4, and nSiO2-3 was produced by a physical vapor synthesis (PVS) method. However, the details of the manufacturing method are unclear. Additionally, surface properties of these nSiO2s are also unknown. Therefore, the identity of the

specific properties of nSiO2s that correlate to the cellular effects and their potency are unclear. Another important cellular effect of nanoparticles is induction of oxidative stress. In many cases, cytotoxic nanoparticles induce oxidative stress in the cell. First, intracellular ROS levels increase, which then leads to various cellular effects such as apoptosis (Saito et al., 2006). In the present study, nSiO2s also caused a decrease of mitochondrial activity, an increase in intracellular ROS levels, and induction of caspase-3 activity. An increase of the caspase-3 activity suggested the possibility of the induction of apoptosis. In the case of an induction of cellular oxidative stress by metal oxide nanoparticles, an intracellular ROS level increases first. The increase of intracellular ROS drives cellular anti-oxidative systems such as an expression of heme oxigenase-1 (HO-1) and weak oxidative stress is suppressed. However, strong oxidative stress leads to oxidation of biomolecule such as lipids, proteins and nucleic acids. Particularly, polyunsaturated fatty acid is easily oxidized. Intracellular lipid peroxidation level was increased by exposure of nSiO2s as well as intracellular ROS level. These results suggest that exposure of nSiO2 induced oxidative stress to cells. However, compared with some cytotoxic metal oxide nanoparticles such as ZnO and CuO (Fukui et al., 2012; Karlsson et al., 2008), the cellular oxidative stress derived by nSiO2 was not severe. The cell was not annihilated by the nSiO2 exposure. A mild oxidative stress causes apoptosis while a strong oxidative stress causes the necrosis (Saito et al., 2006). Therefore, there is possibility that nSiO2 exposure induced mild oxidative stress to cells and then the oxidative stress led to activation of caspase-3. It is likely that intracellular nSiO2 uptake caused mitochondrial dysfunction, which subsequently induced intracellular ROS production, which, in turn, activated caspase-3. The induction of intracellular ROS levels and caspase-3 activity were not nSiO2-specific cellular responses. They have also been observed in cells exposed to other nanoparticles (Horie et al., 2013). The cellular responses caused by internalized nSiO2s may be similar to those caused by other insoluble metal oxide nanoparticles. Results of the MTT assay suggested that the nSiO2s induced a modest degree of mitochondrial dysfunction, but that the degree of dysfunction was independent of the kind of nSiO2. However, the result of the MTT assay should be noted. It has been suggested that in some cases, the MTT assay is not suitable for evaluation of the cellular influences of nanomaterials (Horie et al., 2010b; Wo¨rle-Knirsch et al., 2006). Additionally, it is also reported that porous SiO2 particles affect the key reaction in the MTT assay (Laaksonen et al., 2007). Therefore, additional assessments of cell viability that do not rely on the same chemical reaction as the MTT assay need to be performed to further evaluate the cytotoxic effects of nSiO2s. Long-term in vivo toxicity of inhaled nSiO2s such as fibrosis and carcinogenesis cannot be evaluated by the methods used in the present study. Indeed, although it is known that crystalline silica caused silicosis in vivo, the cellular effects of mcSiO2 measured in this study were not as significant. These data support the conclusion that the long-term in vivo effects of nano-scale and micro-scale

Toxicology Mechanisms and Methods Downloaded from informahealthcare.com by University of Laval on 06/18/14 For personal use only.

DOI: 10.3109/15376516.2013.879505

SiO2 particle need to be addressed further, perhaps in animal models. In conclusion, cellular effects of nSiO2 were different by manufacturer. Unfortunately, in this study, a physical and/or chemical factor of cellular effect caused by nSiO2 was not able to determine clearly. Association of solubility and protein adsorption ability of nSiO2 to its cellular effects seemed to be small. One of the causes of different cellular effects of nSiO2s may be different surface property. Although the cellular effects of nSiO2s are similar to that of metal oxide nanoparticles, the mechanism of the cellular effect may be different. It should be noted that these effects may depend on their surface property and that they were not especially potent, compared with the cellular effects of micro-scale SiO2 particles. According to our previous investigations, cytotoxic activity of nSiO2s can be summarized as the effects of ZnO, CuO4nSiO24TiO2, CeO2 (Horie et al., 2010a, 2011, 2012a).

Acknowledgements We appreciate technical assistance provided by Ms. Arisa Miyauchi.

Declaration of interest This work was funded by a grant from the New Energy and Industrial Technology Development Organization (NEDO) entitled ‘‘Evaluating risks associated with manufactured nanomaterials (P06041)’’.

References Arts JH, Muijser H, Duistermaat E, et al. (2007). Five-day inhalation toxicity study of three types of synthetic amorphous silicas in Wistar rats and post-exposure evaluations for up to 3 months. Food Chem Toxicol 45:1856–67. Berg JM, Romoser AA, Figueroa DE, et al. (2013). Comparative cytological responses of lung epithelial and pleural mesothelial cells following in vitro exposure to nanoscale SiO2. Toxicol In Vitro 27: 24–33. Cronholm P, Karlsson HL, Hedberg J, et al. (2013). Intracellular uptake and toxicity of Ag and CuO nanoparticles: a comparison between nanoparticles and their corresponding metal ions. Small 9:70–82. Donaldson K, Borm PJ. (1998). The quartz hazard: a variable entity. Ann Occup Hyg 42:287–94. Foldbjerg R, Wang J, Beer C, et al. (2013). Biological effects induced by BSA-stabilized silica nanoparticles in mammalian cell lines. Chem Biol Interact 204:28–38. Fukui H, Horie M, Endoh S, et al. (2012). Association of zinc ion release and oxidative stress induced by intratracheal instillation of ZnO nanoparticles to rat lung. Chem Biol Interact 198:29–37. Gehrke H, Fru¨hmesser A, Pelka J, et al. (2013). In vitro toxicity of amorphous silica nanoparticles in human colon carcinoma cells. Nanotoxicology 7:274–93. Horie M, Nishio K, Fujita K, et al. (2009). Ultrafine NiO particles induce cytotoxicity in vitro by cellular uptake and subsequent Ni(II) release. Chem Res Toxicol 22:1415–26. Horie M, Nishio K, Fujita K, et al. (2010a). Cellular responses by stable and uniform ultrafine titanium dioxide particles in culture-medium dispersions when secondary particle size was 100 nm or less. Toxicol In Vitro 24:1629–38.

Cellular influences of silica nanoparticles

203

Horie M, Komaba LK, Kato H, et al. (2010b). In vitro evaluation of cellular responses induced by Al(OH)3-treated rutile TiO2 nanoparticles. Nano Biomed 2:182–93. Horie M, Nishio K, Kato H, et al. (2011). Cellular responses induced by cerium oxide nanoparticles: induction of intracellular calcium level and oxidative stress on culture cells. J Biochem 150:461–71. Horie M, Fujita K, Kato H, et al. (2012a). Association of the physical and chemical properties and the cytotoxicity of metal oxide nanoparticles: metal ion release, adsorption ability and specific surface area. Metallomics 4:350–60. Horie M, Kato H, Fujita K, et al. (2012b). In vitro evaluation of cellular response induced by manufactured nanoparticles. Chem Res Toxicol 25:605–19. Horie M, Nishio K, Endoh S, et al. (2013). Chromium(III) oxide nanoparticles induced remarkable oxidative stress and apoptosis on culture cells. Environ Toxicol 28:61–75. Karlsson HL, Cronholm P, Gustafsson J, Mo¨ller L. (2008). Copper oxide nanoparticles are highly toxic: a comparison between metal oxide nanoparticles and carbon nanotubes. Chem Res Toxicol 21: 1726–32. Kato H, Fujita K, Horie M, et al. (2010). Dispersion characteristics of various metal oxide secondary nanoparticles in culture medium for in vitro toxicology assessment. Toxicol In Vitro 24:1009–18. Laaksonen T, Santos H, Vihola H, et al. (2007). Failure of MTT as a toxicity testing agent for mesoporous silicon microparticles. Chem Res Toxicol 20:1913–18. Lesniak A, Fenaroli F, Monopoli MP, et al. (2012). Effects of the presence or absence of a protein corona on silica nanoparticle uptake and impact on cells. ACS Nano 6:5845–57. Lesniak A, Salvati A, Santos-Martinez MJ, et al. (2013). Nanoparticle adhesion to the cell membrane and its effect on nanoparticle uptake efficiency. J Am Chem Soc 135:1438–44. Lesur O, Cantin AM, Tanswell AK, et al. (1992). Silica exposure induces cytotoxicity and proliferative activity of type II pneumocytes. Exp Lung Res 18:173–90. Lin W, Huang YW, Zhou XD, Ma Y. (2006). In vitro toxicity of silica nanoparticles in human lung cancer cells. Toxicol Appl Pharmacol 217:252–9. McCarthy J, Inkielewicz-Ste˛pniak I, Corbalan JJ, Radomski MW. (2012). Mechanisms of toxicity of amorphous silica nanoparticles on human lung submucosal cells in vitro: protective effects of fisetin. Chem Res Toxicol 25:2227–35. Nabeshi H, Yoshikawa T, Matsuyama K, et al. (2011). Amorphous nanosilica induce endocytosis-dependent ROS generation and DNA damage in human keratinocytes. Part Fibre Toxicol 8:1. doi: 10.1186/ 1743-8977-8-1. Napierska D, Rabolli V, Thomassen LC, et al. (2012). Oxidative stress induced by pure and iron-doped amorphous silica nanoparticles in subtoxic conditions. Chem Res Toxicol 25:828–37. Nazli C, Ergenc TI, Yar Y, et al. (2012). RGDS-functionalized polyethylene glycol hydrogel-coated magnetic iron oxide nanoparticles enhance specific intracellular uptake by HeLa cells. Int J Nanomed 7:1903–20. Peretz A, Checkoway H, Kaufman JD, et al. (2006). Silica, silicosis, and lung cancer. Isr Med Assoc J 8:114–18. Saito Y, Nishio K, Ogawa Y, et al. (2006). Turning point in apoptosis/ necrosis induced by hydrogen peroxide. Free Radic Res 40:619–30. Shinto H, Hirata T, Fukasawa T, et al. (2013). Effect of interfacial serum proteins on melanoma cell adhesion to biodegradable poly(l-lactic acid) microspheres coated with hydroxyapatite. Colloids Surf B Biointerfaces 108C:8–15. Wo¨rle-Knirsch JM, Pulskamp K, Krug HF. (2006). Oops they did it again! Carbon nanotubes hoax scientists in viability assays. Nano Lett 6:1261–8. Yoshida T, Yoshioka Y, Matsuyama K, et al. (2012). Surface modification of amorphous nanosilica particles suppresses nanosilica-induced cytotoxicity, ROS generation, and DNA damage in various mammalian cells. Biochem Biophys Res Commun 427: 748–52.