Genotoxicity evaluation of stearic acid grafted chitosan oligosaccharide nanomicelles

Mutation Research 751 (2013) 116–126

Contents lists available at SciVerse ScienceDirect

Mutation Research/Genetic Toxicology and Environmental Mutagenesis journal homepage: www.elsevier.com/locate/gentox Community address: www.elsevier.com/locate/mutres

Genotoxicity evaluation of stearic acid grafted chitosan oligosaccharide nanomicelles Peng Hu a , Tingting Wang a , Qin Xu a , Yan Chang b , Honggang Tu b , Yifan Zheng a , Jun Zhang a , Yuying Xu a , Jun Yang c , Hong Yuan d , Fuqiang Hu d,∗ , Xinqiang Zhu a,∗∗ a

Department of Toxicology, Zhejiang University School of Public Health, 866 Yu-Hangtang Road, Hangzhou, Zhejiang 310058, China National Shanghai Center for New Drug Safety Evaluation & Research, 199 Guoshoujing Rd., Zhangjiang Hi-tech Park, Pudong, Shanghai 201203, China c Department of Toxicology, Hangzhou Normal University School of Public Health, Hangzhou, Zhejiang 310036, China d College of Pharmaceutical Science, Zhejiang University, 866 Yu-Hang-Tang Road, Hangzhou, Zhejiang 310058, China b

a r t i c l e

i n f o

Article history: Received 14 August 2012 Received in revised form 9 November 2012 Accepted 19 December 2012 Available online 27 December 2012 Keywords: Chitosan oligosaccharide nanomicelles Genotoxicity Apoptosis Oxidative stress

a b s t r a c t Stearic acid grafted chitosan oligosaccharide (CSO-SA) nanomicelles could be promptly internalized into cancer cells; therefore, it is regarded as a promising drug carrier for cancer therapy. However, the toxicity of CSO-SA is not clear. In the present study, the genotoxic effects of CSO-SA nanomicelles (with high substitution degree of SA, 42.6 ± 3.8%) were evaluated with a battery of genotoxicity assays. Mutagenicity was not found in Ames Salmonella/microsome mutagenicity assay (Ames test), while mild but definite positive results were observed in mouse bone marrow micronucleus assay and single cell gel electrophoresis (SCGE or comet assay) in A549 cells. CSO-SA was also found to induce apoptosis and oxidative stress through the induction of reactive oxygen species (ROS) in a dose-dependent manner in A549 cells. Preincubation with the free radical scavenger N-acetyl-l-cysteine (NAC) decreased the intracellular ROS level and alleviated the DNA damage in A549 cells. Expression levels of cleaved poly ADP-ribose polymerase (PARP), caspase-9 and caspase-3, markers of apoptosis, were significantly higher in CSO-SA treated cells. In conclusion, these results suggested significant genotoxicity of high doses of CSO-SA nanomicelles in vivo and in vitro. Oxidative stress was, at least partially, the possible mechanism underlying the genotoxicity induced by CSO-SA. © 2012 Elsevier B.V. All rights reserved.

1. Introduction With the advancement of nanotechnology, more and more nanoparticles (NPs) are being generated by human. Due to their unique size and structure, they have many new physical and chemical properties, such as small size effect, large specific surface, high reactivity, quantum effects, etc. Compared to regular size materials, they are superior in strength, toughness, specific heat capacity, as well as catalytic ability, conductivity, diffusivity, magnetic susceptibility, optical property, and electromagnetic wave absorption. Therefore, NPs are widely used in many fields including medical and pharmaceutical research, material science, and other industrial applications [1]. Developing efficient drug carrier is a key step in the pharmaceutical industry. The application of nano-carrier has made rapid progress, such as chitosan and its derivatives. Chitosan is a natural cationic polysaccharide, which is a biodegradable,

∗ Corresponding author. Tel.: +86 571 8820 8441; fax: +86 571 8820 8441. ∗∗ Corresponding author. Tel.: +86 571 8820 8143; fax: +86 571 8820 8143. E-mail addresses: [email protected] (F. Hu), [email protected] (X. Zhu). 1383-5718/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.mrgentox.2012.12.004

low toxic, biocompatible, anti-bacterial and anti-tumor biopolymer with diverse applications [2]. For example, it has been used to develop the optimal drug sustained release and controlled release formula [3]. Also it could enhance the absorption of insoluble drugs by influencing the mucus [4]. In addition, it could improve the penetration of macromolecular drugs by opening the tight junctions, promoting transport function across epithelial cells [4]. However, the problems of low transfection efficiency, blood compatibility and effects on cell viability of chitosan remain to be solved [5]. To solve the problem, chitosan oligosaccharide (CSO), the depolymerized product of chitosan, has received considerable attention [6]. Recent reports indicated that CSO itself exhibits some beneficial effects. CSO could reduce the expression of obesity-related gene expression in ob/ob mice, overcome metabolic disorders of obesity [7], and suppress metastasis of human breast cancer cells [8]. Stearic acid (SA), an endogenous long-chain saturated fatty acid with biocompatibility and low toxicity, can be used to modify the structure of CSO to generate SA grafted CSO (CSO-SA) [9]. Nanomicelles can be formed in the aqueous medium when CSO-SA dissolves, and the nanomicelles would be promptly internalized into cancer cells [9–12]. As vectors, the optimal transfection efficiency of CSO-SA nanomicelles (about

P. Hu et al. / Mutation Research 751 (2013) 116–126

15%) is approximately the same as that of LipofectamineTM 2000 (about 20%). Moreover, the cytotoxicity of CSO-SA nanomicelles was lower in A549 cells compared to that of LipofectamineTM 2000 [13]. In addition, the CSO-SA nanomicelles encapsulated drugs were able to reverse the drug resistance and improve the cytotoxicity [12,14]. However, it has been demonstrated that NPs could traverse the lung blood barrier [15], the blood brain barrier [16] and the placental barrier [17], to reach the cardiovascular system and a variety of tissues and organs. And many groups have reported that NPs have impacts on genes, proteins, cells and whole animals [18–22]. For example, nickel ferrite NPs could induce apoptosis through oxidative stress in A549 cells [18], and multi-wall carbon nanotubes exposure resulted in cytotoxic and genotoxic effect [19]. Current research on chitosan and its derivatives almost all focus on their synthesis, physicochemical property and therapeutic effect, and there are few studies evaluating the possible toxicity of CSO–SA nanomicelles. In particular, the potential genotoxicity of CSO-SA is of concern, as DNA damage is associated with tumorigenesis. It has been shown that some NPs can induce DNA damage through increasing ROS and promoting inflammatory responses [23,24], which eventually can lead to necrotic, apoptotic or autophagic cell death [18,19,25–27]. Therefore, a better understanding of the toxicity of CSO-SA nanomicelles is essential for the development of safer drug carriers. Previously, in acute toxicity testing, our group had found that CSO-SA could cause animal harm in high dose, and the harm degree enhanced with increasing the substitution degree of CSO-SA (unpublished data), suggesting that CSO-SA may be some toxic. For the good cellular uptake properties of CSO-SA, we used the CSO-SA with high substitution degree of SA which could be quickly internalized into cancer cell for toxicity studies. In this context, the aim of this study was to investigate the adverse effect (including cytotoxicity and genotoxicity) and possible mechanism of CSO-SA nanomicelles on bacterium, mouse and human cells, in an effort to obtain more detailed information concerning the safety of this type of nanomicelles for its potential application in the medical field. 2. Material and methods 2.1. Stearic acid-grafted chitosan oligosaccharide (CSO-SA) micelles preparation CSO-SA was prepared as described before by our group [9–12]. The CSO-SA was synthesized by the coupling reaction of the amino groups of CSO (obtained by enzymatic degradation of 95% deacetylated chitosan) with carboxyl group of SA. The CSO-SA could be easily dissolved in DI water and self-aggregated in aqueous environment. The CSO-SA micelles were prepared by ultrasonic treatment (JY92-IIN, Ningbo Xinzhi Scientific Instrument Institute, Zhejiang, China) at the strength of 400 w for 30 cycles (active every 2 s for a 3 s duration) on ice bath. 2.2. Cell culture A549 human lung carcinoma cells (ATCC No. CCL-185TM ) were cultured in RPMI 1640 medium containing 100 mg/ml penicillin/streptomycin and 10% (v/v) fetal bovine serum (FBS) in a 37 ◦ C, 5% CO2 humidified incubator. 2.3. Animals Sixty ICR male mice (4 weeks, 20 ± 2.0 g body weight) were obtained from Zhejiang University Laboratory Animal Center. They were adapted in the conditional environment (room temperature,

117

23◦ ±2 ◦ C; controlled relative humidity, 50 ± 20%; 12 h light/dark cycle) for a period of 1 week prior to the experiments. Mice were given sterilized water and diet ad libitum.

2.4. CSO-SA characterization The degree of substitution (DS), defined as the number of stearic acid groups per 100 anhydroglucose units of chitosan, was examined by the TNBS method [28]. Size and zeta potential of the micelles with 1 mg/ml of CSO–SA concentration were measured by dynamic light scattering using a Zetasizer (3000HS, Malvern Instruments Ltd., UK). Morphology of the CSO-SA with the concentration of 1.0 mg/mL in DI water was examined by transmission electron microscopy (TEM) (JEM-1230, JEOL, Japan). The samples were stained by phosphotungstic acid, placed onto the carbon-coated copper TEM grids, and observed with TEM.

2.5. Bacterial reverse mutation test The Ames test was conducted following the standard procedures of the OECD guideline No. 471 (1997). The tester strains used in this study were Salmonella typhimurium strains TA97a, TA98, TA100, TA102, and TA1535. The test was performed by the stand plate method [29,30], with or without the exogenous metabolic activation (S9 mix). The concentrations in the study were 20.6, 61.7, 185.2, 555.6, 1666.7, and 5000.0 ␮g per plate. These doses were determined after toxicity test had been carried out. The positive control used in the absence of S9 mix was NaN3 for the TA100 and TA1535 strains, methyl methanesulfonate (MMS) for the TA102 strain, daunorubicin (DRN) for the TA98 strain, and ICR-191 for the TA97a strain; in the presence of S9 mix 2-aminofluorene (2-AF) was used for all tester strains. Dimethylsulfoxide (DMSO) was used to dissolve the positive controls in the Ames test. So DMSO was used as negative control. To determine the mutagenicity without S9 mix, CSO-SA was mixed with 0.1 ml bacterial suspension and 0.5 ml 100 mM sodium phosphate buffer (pH 7.4). For testing in the presence of S9 mix, 0.5 ml of S9 instead of the sodium phosphate buffer was used. Overlay agar (2 ml) was added to the mixture. The mixtures were then poured onto agar plates and incubated for 48 h at 37 ◦ C. After the incubation, the revertant colonies on each plate were counted. There was a mutagenic potential in the test compound if the mutant frequency was 2.0 or higher, or revertant colonies were induced in a dose-dependent manner. Each dose was tested on triple plates.

2.6. Cytotoxicity assay Cytotoxicity was assessed by using the 3-[4,5-dimethylthiazol2-yl] -2,5-diphenyltetrazolium bromide (MTT) assay. A549 cells were grown in 96-well microtitre plates in 200 ␮l medium and incubated for 24 h. Then the cells were treated with 200 ␮l of CSO-SA micelles solution (12.5–200 ␮g/ml) for 12, 24 and 48 h. Cells incubated with medium without CSO-SA were used as control (0 ␮g/ml). After various treatments, 20 ␮l of MTT solution (5 mg/ml) was added. Following 4 h of incubation at 37 ◦ C, the medium was removed, 150 ␮l of DMSO was added to dissolve the MTT formazan crystal which formed inside the cells. Finally, the plates were shaken for 10 min, and the absorbance of formazan product was measured at 570 nm in an Infinite 200 microplate reader (Tecan Group Ltd., Männedorf, Switzerland). The cytotoxicity was calculated as a percentage of the absorbance of the negative control.

118

P. Hu et al. / Mutation Research 751 (2013) 116–126

2.7. Alkaline single-cell microgel electrophoresis assay The alkaline comet assay was used to assess CSO-SA-induced DNA damage as described by Singh et al. [31] with modifications. The A549 cells were grown in a six-well plate (about 1 × 106 cells/well). Cells were incubated with different concentrations (25–200 ␮g/ml) of CSO-SA for 12, 24 and 48 h. The negative control cells were incubated with medium without CSO-SA. The positive control cells were treated with 500 ␮M H2 O2 for 10 min. After treatment, the cells were harvested and washed in PBS and resuspended in Ca2+ and Mg2+ free PBS solution. The cell suspension (10,000 cells in 75 ␮l of suspension) was mixed with 0.75% low melting point agarose (LMA), which was spread over a pre-coated (0.3% normal melting point agarose (NMA) slide. The slides were allowed to solidify on ice and lysed in pre-chilled lysis solution (2.5 M NaCl, 100 mM EDTA, 10 mM Tris base, 10% DMSO, 1% Triton X-100, pH 10) for 1 h at 4 ◦ C. After lysis, DNA was subjected to unwind for 20 min in alkaline electrophoresis solution (0.3 M NaOH and 1 mM EDTA, pH > 13) in the dark at room temperature. Electrophoresis was performed at 20 V and 300 mA for 20 min. Subsequently the slides were neutralized with neutralization buffer (0.4 M Tris-HCl, pH 7.5) for 15 min and fixed in methanol for 10 min. After staining with GelRed (40 ␮M, 20 min), at least 200 comets per slides were examined with a fluorescence microscope (Olympus Corporation, Tokyo, Japan). The Olive tail moment and Tail DNA were measured as endpoints of DNA damage and analyzed by using CASP v1.2.2 software. Experiments were done in duplicates. 2.8. Micronuclei assay The micronuclei assay was performed according to the OECD Guideline 474 (1997). Mice were divided into six groups of 10 animals each. Mice received CSO-SA treatment (5, 10, 25 and 50 mg/kg/day, 0.5 ml by tail vein injection) for 5 days. The negative control group was injected with saline. The positive control group was treated with CTX (40 ml/kg) once by intraperitoneal injection. The animals were sacrificed by cervical dislocation 24 h after the final administration. Bone marrow cells were obtained from sternum and suspended in fetal calf serum and smears were spread on a glass slide. The smears were air-dried, fixed in methanol for 5 min, and stained with Giemsa stain (3%) for 25 min. To detect the mutagenic property, 2000 polychromatic erythrocytes (PCEs) from each animal were scored and the frequency of micronucleated polychromatic erythrocytes (MNPCEs) was estimated. The PCE/NCE (normochromatic erythrocytes) ratio in 200 erythrocytes for each animal was calculated to determine the genotoxicity. The slides were examined at ×100 magnification under oil lens. 2.9. Measurement of reactive oxygen species (ROS) CSO-SA-induced ROS generation was determined in A549 cells exposed to different concentrations (25–200 ␮g/ml) of CSO-SA for 12, 24 and 48 h. Cells treated with 1 ␮l ROSup (Beyotime Institute of Biotechnology, Haimen, China) for 30 min served as positive control. The negative control cells were incubated with medium without CSO-SA. The level of intracellular ROS was measured using 2,7-dichlorofluorescin diacetate (DCFH-DA) as fluorogenic probe[32,33]. Briefly, the cells were seeded in 96-well plates. After exposure, the cell were washed and incubated with fresh culture medium without serum. DCFH-DA dissolve in methanol (10 mM) was diluted in culture to a final concentration of 20 ␮M. Then cells were incubated at 37 ◦ C for 30 min and protected from light. Subsequently, cells were washed with PBS and fluorescence was read at 485 nm excitation and 520 nm emission wavelength using a Hybrid Multi-Mode Microplate Readers (Bio-Tek Instruments, Inc.,

Winoosk, Vermont, USA). Fluorescence from untreated cells was set as 100%. 2.10. N-Acetylcysteine (NAC) pretreatment Cells were treated with antioxidant NAC (6 mM) for 2 h prior to CSO-SA treatment to delineate the potential involvement of oxidative stress in cells damage. 2.11. Apoptosis analysis The Annexin V/Propidium Iodide(PI) assay was applied to assess apoptosis and necrosis of cells exposed to CSO-SA for 24 h as described by Van Engeland et al. [34]. Phosphatidylserine (PS) is only distributed in the inner membrane lipid bilayer in normal cells. The externalization of PS could be regarded as a marker of early-stage apoptosis. Annexin V binds to PS, and Annexin V protein conjugated to FITC could be used as an indicator for PS. Propidium iodide (PI) is a nucleic acid dye that could not permeate the cell membrane. But after membrane damage due to late-stage apoptosis/necrosis, PI could bind to nuclear DNA. In short, A549 cells were seeded in six-well plates and cultured for 24 h. Cells were then exposed to various concentrations of CSO-SA (25–200 ␮g/ml) and incubated for another 24 h. The positive control cells were treated with 500 ␮M H2 O2 for 24 h. The negative control cells were incubated with medium without CSO-SA. After exposure, cells were harvested and washed with 1 ml of PBS twice, resuspended in binding buffer (KeyGEN Biotech, Nanjing, China). Cells were stained for 15 min in binding buffer containing Annexin V-FITC and PI. Apoptosis was analyzed by FC500 flow cytometry (Beckman Coulter, Inc., CA, USA). The 488-nm laser was used for excitation, and FITC was detected in FL-1, while PI was detected in FL-3. Single-stained and unstained cells were used as standard compensation. Early apoptosis (Annexin V+, PI−), late apoptosis/necrosis (Annexin V+, PI−) and live (Annexin V−, PI−) cells were showed as percentages of the all analyzed cells. 2.12. Protein extraction Cells were treated with different concentrations (25–200 ␮g/ml) of CSO-SA for 24 h. Cells treated with 500 ␮M H2 O2 for 24 h served as positive control. The negative control cells were incubated with medium without CSO-SA. After treatment, cells were harvested with a cell scrapper. The cell pellets were collected after centrifugation at 1000 rpm for 10 min at 4 ◦ C and washed twice with cold PBS. The cell pellets were subsequently resuspended in RIPA lysis Buffer (Beyotime Institute of Biotechnology, Haimen, China) containing 1 mM Phenylmethanesulfonyl fluoride (PMSF) and incubated on ice for 60 min. The lysate was harvested into a 1.5 ml centrifuge tube and centrifuged at 12,000× g at 4 ◦ C for 10 min. The supernatant was transferred into a new tube which was used for Western blotting. 2.13. Protein concentration determination A BCA protein assay kit (Beyotime Institute of Biotechnology) was used to measure the concentration of protein. The BCA protein assay kit was based on bicinchoninic acid (BCA) for the colorimetric detection and quantification of protein. Bovine serum albumin was used as standard. 2.14. Western blotting Equal amounts of protein were resolved on 12% SDS-PAGE gel under denaturing condition and then transferred to polyvinylidene difluoride membranes (PVDF, EMD Millipore Corporation,

P. Hu et al. / Mutation Research 751 (2013) 116–126

119

Fig. 1. CSO-SA characterization. (A) Size distribution; (B) Zeta potential; (C) TEM image, a bar is 50 nm.

Billerica, Massachusetts, USA) using a Protein Electrophoresis and Blotting System (Bio-Rad Laboratories, Inc., Hercules, CA, USA). The membranes were blocked in Blocking Buffer (Beyotime Institute of Biotechnology) for 2 h at 25 ◦ C and then incubated with primary antibodies against poly ADP-ribose polymerase (PARP) (Cell Signaling Technology, Inc., Danvers, Massachusetts, USA), caspase-3 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA. USA), caspase-9 (Sigma-Aldrich Co. LLC, St. Louis, Missouri, USA), and ␤-actin (MultiSciences Biotech Co., Ltd., Hangzhou, China). Then the membranes were washed in Tris-buffered saline with 0.1% Tween and incubated with FITC-conjugated secondary antibody (LI-COR Biotechnology, Lincoln, Nebraska, USA). Protein bands were visualized by Odyssey Infrared Imaging System (LI-COR Biotechnology). Densities of the bands were analyzed with Quantity One image analysis software (Bio-Rad). ␤-actin was used as an internal control. 2.15. Capase-3 activity assay Activity of caspase-3 was measured in A549 cells using the Caspase-3 Colorimetric Assay Kit (KeyGEN Biotech). The positive control cells were treated with 500 ␮M H2 O2 for 24 h. The negative control cells were incubated with medium without CSO-SA. The assay was based on the principle that caspase-3 in apoptotic cells could cleave the synthetic Caspase-3 Substrate (tertrapeptide substrates DEVD-pNA) and release chromophore p-nitroanilide (pNA), which can be measured at absorbance (OD, optical density) of F405 nm[35,36]. Briefly, A549 cells were washed with cold PBS by centrifugation at 2000× g at 4 ◦ C for 5 min, and then resuspended in lysis buffer containing 0.5 mM PMSF and 1 mM Dithiothreitol

(DTT), and incubated on ice for 60 min. The lysate was centrifuged at 12,000× g at 4 ◦ C for 10 min. Assays were performed on 96well microtitre plates. The reaction mixture consisted of 50 ␮l equal amounts of protein, 50 ␮l of 2× reaction buffer (containing 1 mM DTT) and 5 ␮l Caspase-3 Substrate (DEVD-pNA). After incubation at 37 ◦ C for 4 h, the absorbance of pNA was determined using the Infinite 200 microplate reader (Tecan Group Ltd) at wavelength of 405 nm. The activity of caspase-3 was expressed as OD405 (test)/OD405 (negative control). Assays were performed in duplicate, and at least three independent experiments were performed. 2.16. Statistical analysis Statistical analyses were conducted using the PASW Statistics 18.0 package (SPSS Inc, Chicago, IL, USA). Data were expressed as mean ± SD. One-way ANOVA, paired-t test, Least-Significant Difference (LSD) test and Pearson’s Chi-Square test were used to determine statistical significance. p < 0.05 was considered statistically significant. 3. Results 3.1. CSO-SA characterization CSO-SA could be easily prepared by dispersing CSO–SA in DI water and self-assemble to form nanomicelles due to the hydrophobic SA modification of water-soluble CSO molecules. The DS was 42.6 ± 3.8%. The mean hydrodynamic diameter and zeta potential of the CSO-SA determined by the DLS measurement was 51.0 nm

120

P. Hu et al. / Mutation Research 751 (2013) 116–126

Fig. 2. CSO-SA decreases cell viabilities in a dose-dependent manner in A549 cells. Cytotoxicity of CSO-SA was measured by MTT. Data were presented as mean (n ≥ 3) ± SD. *, p < 0.05 (12 h); # , p < 0.05 (24 h); and &, p < 0.05 (48 h), compared to the control group.

and 39.3 ± 0.5 mV, respectively (Fig. 1A–B). It meant that CSO-SA had small size and narrow size distribution in DI water. The CSOSA micelles were stable in colloidal dispersion because the zeta potential of CSO-SA micelles was larger than 30 mV in DI water [37]. Fig. 1C showed TEM image of CSO-SA. The nanomicelles had near spherical shape. And the nanomicelles sizes observed by TEM were similar to the results from a Zetasizer.

3.2. CSO-SA could not increase the number of revertant colonies in tester strains Table 1 showed the result of the bacterial reverse mutation test. The number of revertant colonies in the positive control was significantly increased in all tester strains, indicating that the assay was conducted properly for testing mutagenicity. Precipitation of CSO-SA was not seen in any of the tester strains. Growth inhibition was observed at high-doses in all tester strains with or without S9 mix. Although CSO-SA significantly increased the number of colonies in strains TA97a (1666.7 ␮g/plate) and TA100 (185.2 and 555.6 ␮g/plate) with activation, the number of colonies was less than 2-fold of the negative control. Hence, according to the criteria, the mutagenicity of the CSO-SA was considered to be negative.

3.3. CSO-SA decreases cell viabilities in a dose-dependent manner in A549 cells Mitochondria function was used as a cell viability marker of A549 cells. MTT can be converted into formazan by a mitochondrial enzyme in living cells. Formazan has an absorption at 570 nm and the absorbance is proportional to the amount of living cells. The result of cytotoxicity assay was summarized in Fig. 2. In general, the result demonstrated a concentration- and time-dependent decrease in cell viability after CSO-SA treatment. The IC50 (50% of inhibition concentration) was 202.38 ␮g/ml (12 h), 157.13 ␮g/ml (24 h) and 70.84 ␮g/ml (48 h).

Fig. 3. CSO-SA induces DNA damage in A549 cells. Cells were treated with different concentrations of CSO-SA (25–200 ␮g/ml) for 12, 24, or 48 h and then subjected to comet assay. (A) Tail DNA. (B) Olive tail moment. H2 O2 was used as positive control. Data were presented as mean ± SD. *, p < 0.05 (12 h); # , p < 0.05 (24 h); and &, p < 0.05(48 h), compared to the negative control group.

3.4. CSO-SA induces DNA damage in A549 cells The alkaline single-cell microgel electrophoresis (comet) assay is sensitive in vitro assay used to detect the DNA damage effect of chemicals. DNA fragmentation served as a marker for DNA damage induced by CSO-SA using the alkaline single-cell microgel electrophoresis (comet) assay. The comet assay was able to detect DNA single- and double-strand breaks. Olive tail moment (OTM) and tail DNA were analyzed as the index of the assay. The results of the comet assay were presented in Fig. 3. A highly significant increase in DNA breakage after H2 O2 treatment (positive control) was observed (p < 0.05). Interestingly, all the OTM and tail DNA of the treatment groups were statistically significant increased (p < 0.05, compared to negative control), indicating that CSO-SA could induce DNA strand breakages. But there was no dose- or time-effect observed in the results. We also used the comet assay to assess the genotoxicity of SA or/and CSO alone, and the results were negative (data not shown).

P. Hu et al. / Mutation Research 751 (2013) 116–126

121

Table 1 The number of total colonies including spontaneous revertant colonies that appeared on a plate. Dose (␮g/plate)

0.0 20.6 61.7 185.2 555.6 1666.7 5000.0 Positive

Base-pair substitution type TA100 −S9mix

+S9mix

179 ± 197 ± 197 ± 206 ± 194 ± 142 ± 0± 957 ± (2.0)

173 ± 118 ± 171 ± 227 ± 226 ± 164 ± 32 ± 849 ± (20.0)

10 17 13 9 24 9a 0a 85

8 14 17 12 9 16 1a 21

Frameshift type TA1535 −S9mix

+S9mix

21 ± 17 ± 15 ± 15 ± 14 ± 15 ± 20 ± 341 ± (2.0)

14 ± 20 ± 14 ± 18 ± 17 ± 17 ± 5± 203 ± (3.0)

4 3 2 1 3 3 4a 15

2 1 4 3 3 1 1a 19

TA102 −S9mix

+S9mix

238 ± 246 ± 237 ± 227 ± 215 ± 170 ± 0± 871 ± (1.0)

283 ± 233 ± 230 ± 227 ± 227 ± 287 ± 257 ± 1224 ± (200.0)

12 13 14 10 6 7a 0a 27

10 7 15 8 13 7 13a 47

TA98 −S9mix

+S9mix

21 ± 19 ± 19 ± 19 ± 24 ± 15 ± 8± 964 ± (6.0)

29 ± 25 ± 25 ± 18 ± 22 ± 21 ± 6± 1190 ± (20.0)

3 4 4 6 1 3 3a 9

2 4 6 3 2 1 2a 27

TA97a −S9mix

+S9mix

94 ± 109 ± 102 ± 102 ± 99 ± 81 ± 0± 857 ± (1.0)

124 ± 131 ± 127 ± 108 ± 142 ± 183 ± 108 ± 943 ± (20.0)

3 7 7 7 4 3a 0a 27

7 6 10 10 12 11 4a 18

DMSO was used as negative control. The positive control used in the absence of S9 mix was NaN3 for the TA100 and TA1535 strains, methylmethanesulfonate (MMS) for the TA102 strain, daunorubicin (DRN) for the TA98 strain, and ICR-191 for the TA97a strain; in the presence of S9 mix 2-aminofluorene (2-AF) was used for all tester strains. a indicated that growth inhibition was observed. Values in parentheses represented the doses of positive control chemicals (␮g/plate).

3.5. CSO-SA induces increase of the micronuclei frequency in vivo Since the genotoxicity of CSO-SA in comet assay was observed, their genotoxicity in vivo was examined. Rodent erythrocyte micronuclei assay is the most common in vivo assay used to detect the genotoxicity of chemicals. This assay has been used to detect chromosomal damage or the mitotic apparatus damage. The incidence of micronuclei was analyzed as an index of cytogenetic damage. If there was a dose-related increase in frequency of MNPCEs or a significantly increase in the frequency of MNPCEs in at least one dose group, the result was judged as positive. Based on preliminary toxicity test, animals were administrated with different doses of CSO-SA. The results were shown in Table 2. There was a significant difference between negative control and positive control. Also, micronuclei frequency was increased significantly at all dose group of CSO-SA when compared to negative control. But a dose-dependent relationship was not observed. The PCE/NCE ratio of the CSO-SA group showed no significant decrease, indicating no cytotoxic effect. This result indicated that CSO-SA could induce detectable genotoxicity in mice.

antioxidant NAC (6 mM) for 2 h before CSO-SA treatment. It was noted that NAC pretreatment could significantly prevent the ROS generation (p < 0.05) and also significantly decrease the tail DNA and OTM (p < 0.05) compared to the group treated with CSO-SA only (Fig. 5). 3.8. CSO-SA induces apoptosis in A549 cells To further analyze the feature of cytotoxicity induced by CSOSA, flow cytometry was used to examine the ratio of apoptotic cells (Fig. 6). After 24 h exposure, as shown in Fig. 6B and C, CSO-SA induced apoptosis in a dose-dependent manner. The percentage of early apoptotic cells was increased to 1.39%, 5.60% and 8.26% by exposure to 50, 100 and 200 ␮g/ml of CSO-SA, respectively, compared to 0.54% in control cells (p < 0.05). And the ratio of necrotic/late apoptotic cells was also significantly increased in 100 and 200 ␮g/ml CSO-SA treated groups (p < 0.05). These results indicated that apoptosis is involved in CSO-SA-induced cell death. 3.9. CSO-SA increases the expression of proteins associated with apoptosis in A549 cells

3.6. CSO-SA increases ROS in A549 cells To determine the potential contribution of ROS to the observed positive results in micronuclei assay and comet assay after treatment with CSO-SA, intracellular ROS generation was measured by the fluorescence intensity of DCF. As presented in Fig. 4, after treatment with CSO-SA (25, 50, 100 and 200 ␮g/ml) for 12, 24 and 48 h, intracellular production of ROS in all dose groups was significantly increased (p < 0.05).

Caspase-3, caspase-9 and PARP play important roles in the process of apoptosis. Here, after CSO-SA exposure, we examined the protein expression of caspase-3, caspase-9 and PARP in A549 cells, and the data were shown in Fig. 7. The expression level of full length 116KD PARP was increased significantly only at 25 ␮g/ml group

3.7. Inhibition of ROS generation alleviates the genotoxicity of CSO-SA To determine whether ROS played a role in influencing CSOSA-induced genotoxic effects, A549 cells were pretreated with an Table 2 Results of the micronucleus assay of CSO-SA. Dose (mg/kg)

Number of MNPCEs

Negative control 5 10 25 50 Positive control

2.58 6.27 5.81 4.86 5.91 22.5

± ± ± ± ± ±

0.58 1.28* 1.49* 1.09* 1.59* 5.06*

All data were presented as mean ± SD. * p < 0.05, compared to negative control.

PCE/NCE 1.0 0.94 0.95 0.92 0.91 1.0

± ± ± ± ± ±

0.10 0.04 0.06 0.05 0.03 0.10

Number of mice 10 10 10 10 10 10

Fig. 4. Effect of CSO-SA on the intracellular ROS production of A549 cells. A549 cells were treated with CSO-SA of different concentrations (25–200 ␮g/ml) for 12, 24 and 48 h, and ROS was measured by Multi-Mode Microplate Readers. Data were expressed as mean ± SD, n = 3–4. *, p< 0.05 (12 h); #, p < 0.05 (24 h); and &, p < 0.05 (48 h), compared to the negative control group.

122

P. Hu et al. / Mutation Research 751 (2013) 116–126

Fig. 5. Inhibition of ROS generation alleviates the toxic effects of CSO-SA. (A) Effect of NAC pretreatment on ROS generation by different concentrations of CSO-SA at 24 h. The value at the negative control without NAC pretreatment was designated as 1. Paired-t test was used to test the difference between treatment ± NAC. The difference between CSO-SA treatment groups was tested by ANOVA. (B) Effect of NAC pretreatment on Tail DNA induced by CSO-SA at 24 h. (C) Effect of NAC pre-treatment on OTM induced by CSO-SA at 24 h. NAC− and NAC+, without or with NAC pretreatment, respectively. Data were represented as mean ± SD of three independent experiments. When appropriate, the difference in all groups was tested by ANOVA in (B) and (C). *p < 0.05 compared to the negative control (NAC−) in tail DNA and OTM.

(p < 0.05), while the expression level of the cleaved 89KD PARP was significantly increased in other groups except the 25 ␮g/ml group (p < 0.05). The expression level of the inactive 47 kD proenzyme (procaspase-9) was increased in all groups, but only reached statistical significance in high-dose groups (100 and 200 ␮g/ml) (p < 0.05). Unfortunately, no significant difference was found for the expression level of the cleaved 35KD caspase-9, even though there was a slight trend of increase after CSO-SA exposure. During apoptosis, procaspase-3 is activated by cleavage to generate the active caspase-3 which consists of the 17 and 12 KD subunit. As shown in Fig. 6, although no statistical significance was found in the expression of the inactive 32 kDa proenzyme (procaspase3), the expression of cleaved 17 KD caspase-3 was upregulated in three groups (50, 100 and 200 ␮g/ml, p < 0.05). Together, these data pointed to the involvement of caspase pathway in CSO-SA-induced A549 cells apoptosis.

3.10. CSO-SA induces the activation of caspase-3 Caspase-3 is a key enzyme for apoptosis. To support our hypothesis that CSO-SA induced apoptosis, we further tested the activity of caspase-3 using a colorimetric method. Results showed that CSO-SA could significantly induce the caspase-3 activity in a dosedependent manner in all treatment groups (Fig. 8, p < 0.05). 4. Discussion The hydrophilic CSO can be modified by the hydrophobic carboxyl group of SA to form the amphipathic compound CSOSA. Due to the amphipathic property, CSO-SA molecules could self-assemble to form nanomicelles in the aqueous phase [14]. The hydrophobic core could be described as container for poor water-soluble drugs, and the drugs could be incorporated into the

P. Hu et al. / Mutation Research 751 (2013) 116–126

123

Fig. 6. CSO-SA induces apoptosis in A549 cells. A549 cells were treated with different concentrations (25–200 ␮g/ml) of CSO-SA for 24 h; apoptotic cells were analyzed by flow cytometry. (A) Flow cytometric analysis of CSO-SA-induced apoptosis in A549 cells using annexinV-FITC/PI. Quadrants: B2, necrosis/late apoptotic cells; B3, live cells; B4, early apoptotic cells. (B) The percentage of early apoptotic cells in total A549 cells from (A). (C) The percentage of necrosis/late apoptotic cells in total A549 cells from (A). Data were presented as mean ± SD, n = 3, *p < 0.05, compared to negative control.

hydrophobic core, become stable and with increased bioavailability [38]. Due to the hydrophilic shells of CSO, there was a delayed release of the drugs [12]. Therefore, CSO-SA nanomicelles, as mentioned above, are ideal for drug carrier. However, before clinical application, the safety of such CSO-SA nanomicelles has to be thoroughly evaluated. Thus, in the present study, the potential toxicity, particularly, the genotoxicity of CSOSA, was examined. First, bacterial reverse mutation test was used to assess its mutagenic potential. Frameshift and base-pair substitution defects were represented in the different strains (TA97a and TA98 for frameshift, TA100, TA102 and TA1535 for base-pair substitution). According to the OECD guideline No. 471 (1997), the results of the bacterial reverse mutation test on CSO-SA were considered to be negative. Thus, it appears that CSO-SA could not induce gene mutation in Salmonella typhimurium strains.

Although numerous chemicals that are positive in the Ames test might act as carcinogens in animal experiments, a number of chemicals that are not mutagenic in Ames test could also cause cancer in animal experiments, indicating the false-positive and false-negative results in such system [39]. Therefore, further studies were needed to verify the mutagenic potential of CSO-SA. To date, very few genotoxicity studies in vitro and in vivo have been carried out for CSO-SA or related derivatives. Dube et al. reported that chitosan NPs could enhance the plasma exposure of (-)-epigallocatechin gallate in mice [40]. On the other hand, Yoon et al. reported the protective effects of CSO on paraquat-induced nephrotoxicity in rats [41]. The comet assay is a highly sensitive test to measure DNA damage. We demonstrated that CSO-SA could cause DNA damage (increased OTM and tail DNA) at all concentration after exposure in A549 cells. However, no dose-effect was

124

P. Hu et al. / Mutation Research 751 (2013) 116–126

Fig. 7. Effects of CSO-SA on the expression of proteins associated with apoptosis in A549 cells. (A) Representative images of immunoblot for PARP, caspase-9, caspase-3 and ␤-actin; (B) Densitometric analysis of PARP expression from (A); (C) Densitometric analysis of caspase-9 expression from (A); (D) Densitometric analysis of caspase-3 expression from (A). Data were expressed as mean ± S.D. of more than three independent experiments. *p < 0.05, compared to negative control. ␤-actin was used as internal standard.

observed from this assay for the genotoxicty of CSO-SA. In this study, micronuclei test was also used to evaluate the mutagenic potential of CSO-SA in vivo, in which ICR mice were treated with CSO-SA via tail vein injection for 5 days. Consistent with the results of the comet assay, the micronuclei test results showed that CSO-SA could affect the MNPCE generation (an indicator of DNA

damage), but not the PCE/NCE ratio (an indicator of the cytotoxicity of bone marrow cells). Again, dose-dependent genotoxicty was not observed. Taken together, we have applied three assays to evaluate the genotoxicity of CSO-SA. Two of the three assays revealed genotoxicity upon exposure to CSO-SA, though slight variation and no

P. Hu et al. / Mutation Research 751 (2013) 116–126

Fig. 8. Effect of CSO-SA on caspase-3 activity in A549 cells. Caspase-3 activity in lysates of A549 cells treated with different concentrations of CSO-SA (25–200 ␮g/ml) for 24 h was tested. Data were presented as mean ± S.D. (n = 6). *p < 0.05, compared to the negative control group.

dose-dependent relations were observed. Regarding the negative results of Ames test, there was concern that the Ames test might not be appropriate for evaluating nanoparticles. It has been reported that the Ames test was negative for TiO2 [42,43], SWCNT [44] and zinc oxide [45], which were shown to be genotoxic in other assays. In addition, not all particles could be able to penetrate the bacterial membrane [46]. Oxidative stress had been considered as one of the major toxic mechanism for NPs. Some reports had indicated that many types of NPs could induce intracellular generation of ROS [19,47–49]. Stress, inflammation, DNA damage and apoptosis could be elicited by oxidative stress [36]. Based on these reports, we hypothesized that CSO-SA could induce toxicity (including genotoxicity) in A549 cells through ROS generation. Therefore, ROS generation was measured by using DCFH-DA. Our data showed that ROS generation by CSO-SA was significantly increased compared to negative control. On the other hand, ROS generation was inhibited by pretreatment with the ROS scavenger NAC. Furthermore, CSO-SA-mediated DNA damage was inhibited by pretreatment with NAC. Thus, it was suggested that oxidative stress might play an important role in the toxicity of CSO-SA. For the underlying mechanism, one possible answer might be from the nano-scale of the CSO-SA structure. In many substances, when reach nano-scale, their physical/chemical properties may change. Indeed, nanomaterial cytotoxicity is size dependent [50]. For instance, bulk gold is inert and incapable of reacting, but the gold nanoparticles could induce oxidative damage [51,52]. The same thing happens with silver nanoparticles (Ag NPs) [53]. Therefore, it may also apply to the CSO-SA nanomicelles. Mitochondria appear to be a sensitive target for NPs toxicity [48,49]. Since mitochondria dysfunction, as well as the production of ROS, is associated with apoptosis [48,54,55], the Annexin V/Propidium Iodide (PI) assay was applied to assess apoptotic and necrotic cells. The results showed that there is a distinct dosedependent induction of apoptosis in A549 cells after treatment (Fig. 6). Apoptosis is a crucial way to maintain homeostasis in cells, and a large amount of genes acting as death switches play important roles in this process. PARP, a chromatin-associated polymerase that is related to the base excision repair (BER) pathway by modifying the poly(ADP-ribosyl)ation of various nuclear proteins. The cleaved PARP was considered as an important indicator of apoptosis. On the other hand, sequential activation of cysteine-aspartic

125

acid proteases (caspase) was a critical signaling pathway leading to apoptosis. They play central roles in the execution-phase of cell apoptosis [56,57]. Our results showed that CSO-SA treatment upregulated the expression of cleaved-PARP, procaspase-9, cleaved caspase-9, and cleaved caspase-3 protein, while the expression of procaspase-3 was downregulated. Furthermore, we used a colorimetric method to detect the activity of caspase-3 enzyme, and the result showed that there was a statistically significant increase in caspase-3 enzyme activity. Taken together, these data suggested that CSO-SA exposure induced a mild activation of procaspase-9, which in turn activated caspase-3 [18]. The active caspase-3 would then cleave many of the key proteins in the apoptotic process, including PARP. PARP would lose its enzyme activity after being cleaved by caspase-3, which might accelerate the instability of the cell. Of course, other types of cell death may also exist after treatment with CSO-SA, such as necrosis, necroptosis [58], or autophagy. And this is a direction we are interested in our future study. The toxic reactions of nanoparticles are closely related to its physical and chemical properties [53]. To decrease toxic reactions and overcome the shortcomings, we would introduce hydrophilic polyethylene glycol (PEG) or/and adjust DS to modify CSO-SA in our future study. 5. Conclusion In this study, we examined the toxic effects of CSO-SA (with high substitution degree of SA) using bacteria, mice and A549 cells as the model systems. It was shown that while mutagenicity was found to be negative in Ames test, positive results were observed in micronuclei assay and comet assay. We further demonstrated that CSO-SA could decrease cell survival and induce apoptosis in A549 cells. In addition, ROS generation, activation of caspase-9 and caspase-3, and inactivation of PARP were induced by CSO-SA. Furthermore, inhibition of ROS generation could alleviate the genotoxic effects of CSO-SA, suggesting that oxidative stress was, at least partially, the possible mechanism underlying the toxicity induced by CSO-SA. Taken together, it is clear that CSO-SA have toxic effects in different organic systems in high doses, and therefore, further studies are needed to evaluate the toxicity of CSO-SA in order to adequately assess the risks for its application as drug carrier, as well as the underlying mechanisms. It could provide necessary guide for better understanding and using of CSO-SA. Conflict of interest The authors declare that there are no conflicts of interest. Acknowledgements This work was supported by the National Basic Research Program of China (973 Program) [2009CB930300]. References [1] O. Salata, Applications of nanoparticles in biology and medicine, J. Neurobiol. 2 (2004) 3. [2] J. Zhang, W. Xia, P. Liu, Q. Cheng, T. Tahirou, W. Gu, B. Li, Chitosan modification and pharmaceutical/biomedical applications, Mar. Drugs 8 (2010) 1962–1987. [3] Y. Kato, H. Onishi, Y. Machida, Application of chitin and chitosan derivatives in the pharmaceutical field, Curr. Pharm. Biotechnol. 4 (2003) 303–309. [4] N.G.M. Schipper, K.M. Varum, P. Stenberg, G. Ocklind, H. Lennernas, P. Artursson, Chitosans as absorption enhancers of poorly absorbable drugs 3: Influence of mucus on absorption enhancement, Eur. J. Pharm. Sci. 8 (1999) 335–343. [5] K. Kojima, Y. Okamoto, K. Miyatake, Y. Tamai, Y. Shigemasa, S. Minami, Optimum dose of chitin and chitosan for organization of non-woven fabric in the subcutaneous tissue, Carbohydr. Polym. 46 (2001) 235–239. [6] S.C. Richardson, H.V. Kolbe, R. Duncan, Potential of low molecular mass chitosan as a DNA delivery system: biocompatibility, body distribution and ability to complex and protect DNA, Int. J. Pharm. 178 (1999) 231–243.

126

P. Hu et al. / Mutation Research 751 (2013) 116–126

[7] S.G. Kumar, M.A. Rahman, S.H. Lee, H.S. Hwang, H.A. Kim, J.W. Yun, Plasma proteome analysis for anti-obesity and anti-diabetic potentials of chitosan oligosaccharides in ob/ob mice, Proteomics 9 (2009) 2149–2162. [8] Y.-H. Shon, Suppression of metastasis of human breast cancer cells by chitosan oligosaccharides, J. Microbiol. Biotechnol. 19 (2009) 629–633. [9] F.Q. Hu, Y.H. Li, H. Yuan, S. Zeng, Novel self-aggregates of chitosan oligosaccharide grafted stearic acid: preparation, characterization and protein association, Pharmazie 61 (2006) 194–198. [10] F.Q. Hu, M.D. Zhao, H. Yuan, J. You, Y.Z. Du, S. Zeng, A novel chitosan oligosaccharide-stearic acid micelles for gene delivery: properties and in vitro transfection studies, Int. J. Pharm. 315 (2006) 158–166. [11] Y.Z. Du, P. Lu, J.P. Zhou, H. Yuan, F.Q. Hu, Stearic acid grafted chitosan oligosaccharide micelle as a promising vector for gene delivery system: factors affecting the complexation, Int. J. Pharm. 391 (2010) 260–266. [12] J. You, F.Q. Hu, Y.Z. Du, H. Yuan, B.F. Ye, High cytotoxicity and resistant-cell reversal of novel paclitaxel loaded micelles by enhancing the molecular-target delivery of the drug, Nanotechnology 18 (2007) 495101. [13] Y.H. Yun, H. Jiang, R. Chan, W. Chen, Sustained release of PEG-g-chitosan complexed DNA from poly(lactide-co-glycolide), J. Biomater. Sci., Polym. Ed. 16 (2005) 1359–1378. [14] Y.Y. Xu, Y.Z. Du, H. Yuan, L.N. Liu, Y.P. Niu, F.Q. Hu, Improved cytotoxicity and multidrug resistance reversal of chitosan based polymeric micelles encapsulating oxaliplatin, J. Drug Targeting 19 (2011) 344–353. [15] G. Oberdorster, Z. Sharp, V. Atudorei, A. Elder, R. Gelein, W. Kreyling, C. Cox, Translocation of inhaled ultrafine particles to the brain, Inhalation Toxicol. 16 (2004) 437–445. [16] J.A. Sarnat, J. Schwartz, H.H. Suh, Fine particulate air pollution and mortality in 20 US cities, N. Engl. J. Med. 344 (2001) 1253–1254. [17] C.A. Pope 3rd, R.T. Burnett, M.J. Thun, E.E. Calle, D. Krewski, K. Ito, G.D. Thurston, Lung cancer, cardiopulmonary mortality, and long-term exposure to fine particulate air pollution, JAMA 287 (2002) 1132–1141. [18] M. Ahamed, M.J. Akhtar, M.A. Siddiqui, J. Ahmad, J. Musarrat, A.A. Al-Khedhairy, M.S. AlSalhi, S.A. Alrokayan, Oxidative stress mediated apoptosis induced by nickel ferrite nanoparticles in cultured A549 cells, Toxicology 283 (2011) 101–108. [19] Y.Y. Guo, J. Zhang, Y.F. Zheng, J. Yang, X.Q. Zhu, Cytotoxic and genotoxic effects of multi-wall carbon nanotubes on human umbilical vein endothelial cells in vitro, Mutat. Res. 721 (2011) 184–191. [20] S. Maenosono, T. Suzuki, S. Saita, Mutagenicity of water-soluble FePt nanoparticles in Ames test, J. Toxicol. Sci. 32 (2007) 575–579. [21] X. Yang, J. Liu, H. He, L. Zhou, C. Gong, X. Wang, L. Yang, J. Yuan, H. Huang, L. He, B. Zhang, Z. Zhuang, SiO2 nanoparticles induce cytotoxicity and protein expression alteration in HaCaT cells, Part Fibre Toxicol. 7 (2010) 1. [22] B. Trouiller, R. Reliene, A. Westbrook, P. Solaimani, R.H. Schiestl, Titanium dioxide nanoparticles induce DNA damage and genetic instability in vivo in mice, Cancer Res. 69 (2009) 8784–8789. [23] M. Park, I. Lynch, S. Ramirez-Garcia, K.A. Dawson, L. de la Fonteyne, E. Gremmer, W. Slob, J.J. Briede, A. Elsaesser, C.V. Howard, H. van Loveren, W.H. de Jong, In vitro evaluation of cytotoxic and inflammatory properties of silica nanoparticles of different sizes in murine RAW 264.7 macrophages, J. Nanopart. Res. 13 (2011) 6775–6787. [24] I. Ahmad, M.I. Khan, G. Patil, L.K. Chauhan, Evaluation of cytotoxic, genotoxic and inflammatory responses of micro- and nano-particles of granite on human lung fibroblast cell IMR-90, Toxicol. Lett. 208 (2012) 300–307. [25] R. Foldbjerg, D.A. Dang, H. Autrup, Cytotoxicity and genotoxicity of silver nanoparticles in the human lung cancer cell line, A549, Arch. Toxicol. 85 (2011) 743–750. [26] H.L. Liu, Y.L. Zhang, N. Yang, Y.X. Zhang, X.Q. Liu, C.G. Li, Y. Zhao, Y.G. Wang, G.G. Zhang, P. Yang, F. Guo, Y. Sun, C.Y. Jiang, A functionalized single-walled carbon nanotube-induced autophagic cell death in human lung cells through Akt-TSC2-mTOR signaling, Cell Death Dis 2 (2011) e159. [27] C.G. Li, H.L. Liu, Y. Sun, H.L. Wang, F. Guo, S.A. Rao, J.J. Deng, Y.L. Zhang, Y.F. Miao, C.Y. Guo, J. Meng, X.P. Chen, L.M. Li, D.S. Li, H.Y. Xu, H. Wang, B. Li, C.Y. Jiang, PAMAM nanoparticles promote acute lung injury by inducing autophagic cell death through the Akt-TSC2-mTOR Signaling Pathway, J Mol Cell Biol 1 (2009) 37–45. [28] A. Bernkop-Schnurch, M.E. Krajicek, Mucoadhesive polymers as platforms for peroral peptide delivery and absorption: synthesis and evaluation of different chitosan-EDTA conjugates, J. Controlled Release 50 (1998) 215–223. [29] B.N. Ames, J. McCann, E. Yamasaki, Methods for detecting carcinogens and mutagens with the Salmonella/mammalian-microsome mutagenicity test, Mutat. Res. 31 (1975) 347–364. [30] D.M. Maron, B.N. Ames, Revised methods for the Salmonella mutagenicity test, Mutat. Res. 113 (1983) 173–215. [31] N.P. Singh, M.T. McCoy, R.R. Tice, E.L. Schneider, A simple technique for quantitation of low levels of DNA damage in individual cells, Exp. Cell Res. 175 (1988) 184–191. [32] H. Wang, J.A. Joseph, Quantifying cellular oxidative stress by dichlorofluorescein assay using microplate reader, Free Radical Biol. Med. 27 (1999) 612–616.

[33] O. Myhre, J.M. Andersen, H. Aarnes, F. Fonnum, Evaluation of the probes 2’,7’dichlorofluorescin diacetate, luminol, and lucigenin as indicators of reactive species formation, Biochem. Pharmacol. 65 (2003) 1575–1582. [34] M. van Engeland, F.C. Ramaekers, B. Schutte, C.P. Reutelingsperger, A novel assay to measure loss of plasma membrane asymmetry during apoptosis of adherent cells in culture, Cytometry 24 (1996) 131–139. [35] C. Berasain, E.R. Garcia-Trevijano, J. Castillo, E. Erroba, M. Santamaria, D.C. Lee, J. Prieto, M.A. Avila, Novel role for amphiregulin in protection from liver injury, J. Biol. Chem. 280 (2005) 19012–19020. [36] M. Ahamed, R. Posgai, T.J. Gorey, M. Nielsen, S.M. Hussain, J.J. Rowe, Silver nanoparticles induced heat shock protein 70, oxidative stress and apoptosis in Drosophila melanogaster, Toxicol. Appl. Pharmacol. 242 (2010) 263–269. [37] D. Cunningham, R.E. Littleford, W.E. Smith, P.J. Lundahl, I. Khan, D.W. McComb, D. Graham, N. Laforest, Practical control of SERRS enhancement, Faraday Discuss. 132 (2006) 135–145, discussion 147-158. [38] A.N. Lukyanov, V.P. Torchilin, Micelles from lipid derivatives of water-soluble polymers as delivery systems for poorly soluble drugs, Adv. Drug Delivery Rev. 56 (2004) 1273–1289. [39] K. Mortelmans, E. Zeiger, The Ames Salmonella/microsome mutagenicity assay, Mutat. Res. 455 (2000) 29–60. [40] A. Dube, J.A. Nicolazzo, I. Larson, Chitosan nanoparticles enhance the plasma exposure of (-)-epigallocatechin gallate in mice through an enhancement in intestinal stability, Eur. J. Pharm. Sci. 44 (2011) 422–426. [41] S.P. Yoon, M.S. Han, J.W. Kim, I.Y. Chang, H.L. Kim, J.H. Chung, B.C. Shin, Protective effects of chitosan oligosaccharide on paraquat-induced nephrotoxicity in rats, Food Chem. Toxicol. 49 (2011) 1828–1833. [42] Y. Nakagawa, S. Wakuri, K. Sakamoto, N. Tanaka, The photogenotoxicity of titanium dioxide particles, Mutat. Res. 394 (1997) 125–132. [43] D.B. Warheit, R.A. Hoke, C. Finlay, E.M. Donner, K.L. Reed, C.M. Sayes, Development of a base set of toxicity tests using ultrafine TiO2 particles as a component of nanoparticle risk management, Toxicol. Lett. 171 (2007) 99–110. [44] E.R. Kisin, A.R. Murray, M.J. Keane, X.C. Shi, D. Schwegler-Berry, O. Gorelik, S. Arepalli, V. Castranova, W.E. Wallace, V.E. Kagan, A.A. Shvedova, Single-walled carbon nanotubes: geno- and cytotoxic effects in lung fibroblast V79 cells, J. Toxicol. Environ. Health A 70 (2007) 2071–2079. [45] E.K. Dufour, T. Kumaravel, G.J. Nohynek, D. Kirkland, H. Toutain, Clastogenicity, photo-clastogenicity or pseudo-photo-clastogenicity: Genotoxic effects of zinc oxide in the dark, in pre-irradiated or simultaneously irradiated Chinese hamster ovary cells, Mutat. Res. 607 (2006) 215–224. [46] R. Landsiedel, M.D. Kapp, M. Schulz, K. Wiench, F. Oesch, Genotoxicity investigations on nanomaterials: methods, preparation and characterization of test material, potential artifacts and limitations–many questions, some answers, Mutat. Res. 681 (2009) 241–258. [47] Y.-J. Kim, M. Yu, H.-O. Park, S.I. Yang, Comparative study of cytotoxicity, oxidative stress and genotoxicity induced by silica nanomaterials in human neuronal cell line, Mol. Cell Toxicol 6 (2010) 336–343. [48] M. Ahamed, Toxic response of nickel nanoparticles in human lung epithelial A549 cells, Toxicol. in Vitro 25 (2011) 930–936. [49] S. Barillet, M.L. Jugan, M. Laye, Y. Leconte, N. Herlin-Boime, C. Reynaud, M. Carriere, In vitro evaluation of SiC nanoparticles impact on A549 pulmonary cells: cyto-, genotoxicity and oxidative stress, Toxicol. Lett. 198 (2010) 324–330. [50] S.K. Sohaebuddin, P.T. Thevenot, D. Baker, J.W. Eaton, L. Tang, Nanomaterial cytotoxicity is composition, size, and cell type dependent, Part Fibre Toxicol 7 (2010) 22. [51] J.J. Li, L. Zou, D. Hartono, C.N. Ong, B.H. Bay, L.Y.L. Yung, Gold nanoparticles induce oxidative damage in lung fibroblasts in vitro, Adv. Mater. 20 (2008) 138–142. [52] J.J. Li, D. Hartono, C.N. Ong, B.H. Bay, L.Y. Yung, Autophagy and oxidative stress associated with gold nanoparticles, Biomaterials 31 (2010) 5996–6003. [53] N. Singh, B. Manshian, G.J. Jenkins, S.M. Griffiths, P.M. Williams, T.G. Maffeis, C.J. Wright, S.H. Doak, NanoGenotoxicology: the DNA damaging potential of engineered nanomaterials, Biomaterials 30 (2009) 3891–3914. [54] X. Lu, J. Qian, H. Zhou, Q. Gan, W. Tang, J. Lu, Y. Yuan, C. Liu, In vitro cytotoxicity and induction of apoptosis by silica nanoparticles in human HepG2 hepatoma cells, Int. J. Nanomed. 6 (2011) 1889–1901. [55] H. Nakajima, J. Magae, M. Tsuruga, K. Sakaguchi, I. Fujiwara, M. Mizuta, K. Sawai, H. Yamagishi, N. Mizuta, Induction of mitochondria-dependent apoptosis through the inhibition of mevalonate pathway in human breast cancer cells by YM529, a new third generation bisphosphonate, Cancer Lett. 253 (2007) 89–96. [56] T. Takadera, T. Ohyashiki, Caspase-dependent apoptosis induced by calcineurin inhibitors was prevented by glycogen synthase kinase-3 inhibitors in cultured rat cortical cells, Brain Res. 1133 (2007) 20–26. [57] X. Tang, Y. Guo, K. Nakamura, H. Huang, M. Hamblin, L. Chang, L. Villacorta, K. Yin, H. Ouyang, J. Zhang, Nitroalkenes induce rat aortic smooth muscle cell apoptosis via activation of caspase-dependent pathways, Biochem. Biophys. Res. Commun. 397 (2010) 239–244. [58] J. Hitomi, D.E. Christofferson, A. Ng, J. Yao, A. Degterev, R.J. Xavier, J. Yuan, Identification of a molecular signaling network that regulates a cellular necrotic cell death pathway, Cell 135 (2008) 1311–1323.