Journal of Hazardous Materials 283 (2015) 519–528

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In depth analysis of apoptosis induced by silica coated manganese oxide nanoparticles in vitro Chao Yu a , Zhiguo Zhou a,∗ , Jun Wang a , Jin Sun a , Wei Liu a , Yanan Sun a , Bin Kong a , Hong Yang a , Shiping Yang a,b,∗ a The Education Ministry Key Lab of Resource Chemistry and Shanghai Key Laboratory of Rare Earth Functional Materials, Shanghai Normal University, Shanghai 200234, China b The Education Ministry Key Lab of Pesticide & Chemical Biology, South China Agricultural University, Guangzhou 510641, China

h i g h l i g h t s • L929 cell is more sensitive than HeLa cell under the same concentration of MnO@SiO2 NPs. • The forms of cell death are apoptosis for HeLa and L929 cells which induced by MnO@SiO2 NPs. • Internalized MnO@SiO2 NPs induced intracellular ROS overproduction, thus activates p53 pathways to promote cell apoptosis.

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Article history: Received 9 March 2014 Received in revised form 10 June 2014 Accepted 22 September 2014 Available online 7 October 2014 Keywords: MnO nanoparticle Silica Magnetic resonance imaging Toxicity In vitro

a b s t r a c t Manganese oxide nanoparticles (MnO NPs) have been regarded as a new class of T1 -positive contrast agents. The cytotoxicity of silica coated MnO NPs (MnO@SiO2 NPs) was investigated in human cervical carcinoma cells (HeLa) and mouse fibroblast cells (L929). The changes of cell viability, cell morphology, cellular oxidative stress, mitochondrial membrane potential and cell cycle induced by MnO@SiO2 NPs were evaluated. Compared to HeLa cells, L929 cells showed lower cell viability, more strongly response to oxidative stress and higher percentage in the G2/M phase of cell cycle. The appearance of sub-G1 peak, double staining with Annexin V-FITC/PI and the increase of Caspase-3 activity further confirmed apoptosis should be the major form of cell death. Moreover, the apoptotic pathway was clarified as follows. Firstly, reactive oxygen species (ROS) is generated induced by MnO@SiO2 NPs, then p53 is activated followed by an increase in the bax and a decrease in the bcl-2, ultimately leading to G2/M phase arrest, increasing the activity of caspase-3 and inducing apoptosis. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Magnetic resonance imaging (MRI) is one of the most powerful non-invasive imaging technologies. It is used to diagnose disease through the interaction of protons with molecules in tissues [1–3]. The current MRI contrast agents are in the form of T1 -positive agents of paramagnetic species and T2 -negative agents of superparamagnetic nanoparticles [4]. The recent development of T1 contrast agents has led to an increased interest due to their ability to create an enhanced imaging signal. Manganese oxide nanoparticles (MnO NPs), a new class of T1 -positive contrast agents, have

∗ Corresponding authors at: Shanghai Normal University, Department of Chemistry, 100 Guilin Road, Shanghai, China. Tel.: +86 21 64322343; fax: +86 21 64322511. E-mail addresses: [email protected] (Z. Zhou), [email protected] (S. Yang). http://dx.doi.org/10.1016/j.jhazmat.2014.09.060 0304-3894/© 2014 Elsevier B.V. All rights reserved.

recently attracted much attention [5,6]. MRI nanoparticulate contrast agents should be water-dispersible and biocompatible, and have functional groups for further bioconjugation. Silica coating has been widely developed as an effective strategy to meet this requirement due to its chemical/physical stable properties in the biological environment and as a general platform for further conjugation. For example, Krishnan et al. demonstrated the controllable T1 relaxivity of MnO@SiO2 NPs in response to the specific physiological environment [7]. Chou et al. developed trifunctional uniform NPs comprising a hollow MnO NPs core and a functionalized mesoporous silica coating for T1 -weighted MRI, phosphorescent imaging and photodynamic therapy [8]. Shi et al. developed MnObased multifunctionalized mesoporous silica nanoparticles for pH-responsive MRI, ultrasonography and circumvention of multidrug resistance in cancer cells [9]. Although MnO@SiO2 NPs have been widely used for T1 -weighted MRI and drug delivery, there are few reports on their risk assessment in biological applications.

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Seo et al. showed the in vitro cytotoxicity of PEG-ylated MnO NPs using a live/dead cell assay, lactate dehydrogenase assay, and reactive oxygen species detection in lung adenocarcinoma cells, breast cancer cells and glioblastoma cells, respectively [10]. In the present study, the cellular toxicity of MnO@SiO2 NPs was evaluated in human cervical carcinoma cells (HeLa) and mouse fibroblast cells (L929) using biochemical indicators, such as cell viability, cell morphology, cellular ROS, the impact on mitochondria and the cell cycle. Furthermore, the mechanism of apoptosis induced by MnO@SiO2 NPs was explored by measuring the activity of caspase-3 and the expression level of p53, bax and bcl-2. 2. Materials and methods 2.1. Materials and characterization Manganese chloride tetrahydrate (MnCl2 ·4H2 O, 99.0%), sodium oleate (C18 H33 NaO2 , 90.0%), n-hexane (97.0%), cyclohexane (99.5%), ethanol (C2 H5 OH, 99.7%), n-octanol (C8 H18 O, 99.0%), ammonia solution (NH4 OH, 25–28%) and tetraethoxysilane (TEOS) were purchased from Sinopharm Chemical Reagent Co. 1-octadecene (90.0%) and Triton-X100 were purchased from Acros. All chemicals were used without further purification. The water used in the experiments was purified using the Milli-Q Plus 185 water purification system (Millipore, Bedford, MA, USA) with a resistivity greater than 18 M cm. X-ray diffraction (XRD) was performed using a Rigaku DMAX 2000 diffractometer equipped with Cu/Ka not a, is Alpha radiation at a scanning rate of 4◦ /min in the 2 range of 20–80◦ ( = 0.15405 nm, 40 kV, 40 mA). The size and morphology of the synthesized nanoparticles were observed using a JEOL JEM-2100F transmission electron microscope (TEM) operating at 200 kV. The Fourier transform infrared (FT-IR) spectra were recorded on a Nicolet Avatar 370 FT-IR spectrophotometer using KBr pellets. The surface potential and hydrodynamic radius of nanoparticles were measured using a Malvern Zetasizer Nano ZS model ZEN3690 (Worcestershire, UK) equipped with a standard 633 nm laser. Absorption was performed on a microplate reader (Multiskan MK3, USA). Flow cytometry (Beckman Coulter, USA) was used to carry out related experiments. 2.2. Synthesis of MnO and MnO@SiO2 NPs MnO NPs were synthesized by high temperature pyrolysis as reported previously [6,11]. Briefly, 2 mmol of manganese oleate was dissolved in 20 mL of 1-octadecene and degassed at 70 ◦ C in vacuo (1 × 10−2 mbar) for 2 h, then intermittently backfilled with nitrogen to remove the moisture and oxygen. The reaction mixture was subsequently treated with a definitive temperature program. The temperature was held at 280 ◦ C for 1 h. The nanoparticles were washed with ethanol, and subsequently collected by centrifugation. MnO@SiO2 NPs were synthesized by the reverse microemulsion procedure according to the literature [12–14]. Briefly, 2 mg MnO NPs were dispersed in 60 mL cyclohexane and Triton-X100 (1.1 mL), NH4 OH (150 ␮L) and 1-octanol (400 ␮L) were added sequentially. Finally, tetraethyl orthosilicate (TEOS, 200 ␮L) was added to the mixed solution. The solution was stirred for 3 days at 500 rpm. The final product was precipitated with acetone and centrifuged with ethanol.

with 10% fetal bovine serum (FBS), and 1% Pen-Strep. Cells were incubated at 37 ◦ C in a 5% CO2 incubator and split with trypsin/EDTA solution (0.25%) as recommended by the manufacturer. 2.4. MTT assay The methyl thiazolyl tetrazolium (MTT) assay was used to detect viable cells and evaluate the metabolic activity of cells. All samples were analyzed in triplicate. HeLa and L929 cells were seeded in 96well plates at a density of 1 × 104 cells per well, then cultured in 5% CO2 at 37 ◦ C for 24 h. After the cells were incubated with MnO@SiO2 NPs at different concentrations for 0.5 h, 1 h, 4 h, 8 h, 12 h and 24 h, respectively, MTT (20 ␮L, 5 mg/mL) was added to each well for 4 h at 37 ◦ C. After dimethyl sulfoxide (DMSO, 200 mL/well) was added, the absorbance was measured at 490 nm using a microplate reader. 2.5. Optical microscopy of cells HeLa and L929 cells were seeded in 12-well culture plates at a density of 2 × 105 cells per well. Following incubation at 37 ◦ C overnight, the cells were treated with MnO@SiO2 NPs at 50 ␮g/mL in RPMI 1640 for 24 h. The remaining cells were fixed with paraformaldehyde (4%) for 15 min and stained with the standard hematoxylin & eosin (H&E) staining solution (HE, Ketgen, China). Cells were observed using an inverted biological microscope (Olympus, IX71, Japan). 2.6. TEM of cells HeLa and L929 cells were seeded in a culture dish (100 mm × 20 mm). After treatment with MnO@SiO2 NPs at 50 ␮g/mL for 24 h, the cells were washed in phosphate buffered saline (PBS) and fixed overnight with 2.5% glutaraldehyde in 0.01 M PBS at 4 ◦ C. The cells were then postfixed in 1% osmium tetroxide, dehydrated in ethanol, and embedded in Epon (Fluka). Ultrathin sections (60–80 nm) were stained with uranyl acetate and lead citrate and examined by TEM. 2.7. Reactive oxygen species (ROS) in cells The production of ROS in cells was measured using 2 ,7 dichlorofluorescein diacetate (DCFH-DA) (Ketgen, China) as an oxidation-sensitive probe. Briefly, 10 mM DCFH-DA stock solution was diluted 1000-fold with cell culture medium without serum or other additives to prepare a 10 mM working solution. HeLa and L929 cells were seeded in 12-well culture plates at a density of 2 × 105 cells per well. After exposure to MnO@SiO2 NPs at different concentrations for 24 h, the cells were washed with PBS and incubated in 1 mL of DCFH-DA at 37 ◦ C in the dark for 20 min. The DCFH-DA probe passively diffused into cells and was hydrolyzed by the intracellular esterase to yield DCFH, which was trapped inside the cells. ROS produced by cells oxidized DCFH to the highly fluorescent compound 2 ,7 -dichlorofluorescein (DCF). The cells were then washed twice with culture medium without serum and resuspended in RPMI 1640 for analysis. Rosup (50 mg/mL) was used as a positive control. The fluorescence intensity was measured using a flow cytometer at 510–540 nm after excitation at 488 nm. The intracellular ROS level was expressed as DCF relative fluorescence intensity from independent experiments. Data was analyzed using FlowJo software 7.6.5.

2.3. Cell culture 2.8. MDA assay The human cervical carcinoma cells (HeLa) and mouse fibroblast cells (L929) were provided by the Institute of Biochemistry and Cell Biology, SIBS, CAS (China). Cells were cultured in phenol-red-free Dulbecco’s modified essential medium (RPMI 1640) supplemented

HeLa and L929 cells were prepared as described in the lipid peroxidation MDA assay kit (Beyotime, China). MDA was required to react with thiobarbituric acid (TBA) to form a red substance. Cells

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were plated in 12-well plates at a density of 2.0 × 105 cells/well. After exposure to MnO@SiO2 NPs for 24 h, cells were washed with cold PBS and harvested by trypsinization. Cellular extracts were prepared using cell lysis buffer (Western and IP, Beyotime, China) and the lysed cells were centrifuged at 1600 rpm for 10 min to remove debris. MDA level and the protein content were measured in the supernatant. 100 ␮L cell homogenates, 50 ␮L 0.37% TBA and 150 ␮L diluent of TBA were added to each test tube. The mixture was incubated in a boiling water bath for 15 min. After cooling to room temperature, the reaction mixture was centrifuged at 1000 rpm for 10 min. Then, 200 ␮L supernatant was added to a 96-well plate, followed by measurement of the absorbance using a microplate reader at 540 nm. 490 nm was used as a control. An enhanced BCA protein assay kit (Beyotime, China) was used to quantify the protein concentration. 2.9. Mitochondrial membrane potential (MMP) of cells The change in MMP was evaluated using a flow cytometer and a JC-1 (5,5 ,6,6 -tetrachloro-1,1 ,3,3 -tetraethylbenzimidazolcarbocyanine iodide) mitochondrial membrane potential detection kit (Beyotime, China). Briefly, after treatment with MnO@SiO2 NPs at different concentrations (0, 10, 25, 50, 100 and 200 ␮g/mL in RPMI 1640, respectively) for 24 h, the cells were washed with PBS and incubated with JC-1 solution (5 ␮g/mL) at 37 ◦ C for 20 min. The staining solution was removed and cells were washed twice with JC-1 working buffer. Then, the cells were resuspended in 500 ␮L culture medium and analyzed using a flow cytometer. Normally polarized mitochondria (red J-aggregate) were recorded at Ex/Em 585/590 nm, while depolarized mitochondria (green monomer) were recorded at Ex/Em 514/529 nm. The value of MMP staining from each sample was expressed as the ratio of red fluorescence intensity to green fluorescence intensity. Cells treated with 10 ␮M carbonyl cyanide 3-chlorophenylhydrazone (CCCP), a protonophore which can cause dissipation of MMP, was used as a positive control. 2.10. Cell cycle HeLa cells and L929 cells were treated with MnO@SiO2 NPs at different concentrations (0, 10, 25, 50, 100 and 200 ␮g/mL in RPMI 1640, respectively) for 24 h. The cells were washed with PBS, trypsinized, and placed on ice as described above. Then the cells were treated with propidium iodide which was contained in DNA PREP stain (Beckman Coulter, USA). The flow cytometer was set to measure nuclei using DNA FL3-H as the detection trigger. The percentages of Sub-G1, G1, S, and G2/M phases for different concentrations in each treatment medium were calculated from histograms using the area parameter. Three independent experiments were performed and expressed as mean ± SD. The results of the cell cycle measurements were fitted using Flow Jo 7.6.5 software. 2.11. Annexin V-FITC and PI double staining HeLa and L929 cells were spread at a density of 2 × 105 cells per well in 12-well plates, then treated with MnO@SiO2 NPs at different concentrations (0, 10, 25, 50, 100 and 200 ␮g/mL in RPMI 1640, respectively) at 37 ◦ C. Following incubation for 24 h, Annexin V-FITC and PI double staining was performed according to the manufacturer’s instructions. Cells were collected by centrifugation, washed twice with cold PBS, and a cell suspension was prepared by adding 400 ␮L ×1 binding buffer and 10 ␮L Annexin V-FITC, which was incubated for 15 min at 4 ◦ C in the dark. Finally, 10 ␮L PI was added and incubated for 15 min under the same conditions. Flow cytometry experiments were carried out at an excitation

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wavelength of 488 nm for Annexin V-FITC and PI. Statistical data were obtained from the dot plots using Flow Jo software 7.6.5. 2.12. Caspase-3 activity assay Caspase-3 activity was determined using the caspase-3 activity assay kit (Beyotime, China) according to the manufacturer’s protocol. The assay is based on caspase-3 changing acetyl-AspGlu-Val-Asp p-nitroanilide (Ac-DEVD-pNA) into yellow formazan and product pNA. Briefly, after treatment with MnO@SiO2 NPs for 24 h, cells were collected by centrifugation, washed with cold PBS, then re-suspended in lysis buffer (100 ␮L per 2 × 106 cells) and maintained on ice for 15 min. The cells were centrifuged at 16,000 rpm and 4 ◦ C for 12 min. The assay was performed on a 96well plate by incubating the mixture composed of 10 ␮L protein in cell lysate, 80 ␮L reaction buffer and 10 ␮L caspase-3 substrate (Ac-DEVD-pNA, 2 mM) at 37 ◦ C for 6 h. The caspase-3 activity in cells was quantified using a microplate reader at an absorbance of 405 nm. The protein concentration in the cells was determined using the Bradford protein assay kit (Beyotime, China). The activity of caspase-3 was expressed as the percentage of enzyme activity compared to the control. 2.13. Western blot analysis After HeLa and L929 cells were treated with MnO@SiO2 NPs at different concentrations (0, 10, 25 and 50 ␮g/mL, respectively) for 24 h, the cells were incubated with lysis buffer (Beyotime, China). The protein concentration was determined by the BCA protein assay kit (Beyotime, China). The same amount of protein was electrophoresed in polyacrylamide gel, then transferred to nitrocellulose membrane and blocked with 10% skimmed milk in Tris-Buffered Saline Tween-20 (TBST) buffer for 1.5 h. Then the membranes were incubated with primary antibodies at 1:500–1:800 dilution in 10% skimmed milk overnight with continuous agitation at 4 ◦ C. The membranes were then incubated with secondary antibodies conjugated with horseradish peroxidase at a 1:3000 dilution at room temperature for 1 h. After washing three times with TBST, the protein bands were visualized on X-ray film using immobilon western chemilum hrp substrate (Millipore).

3. Results and discussion 3.1. Synthesis and characterization of MnO@SiO2 NPs MnO NPs were synthesized by thermolysis of the manganese oleate precursor in 1-octadecene according to the literature [6,11]. MnO NPs were well dispersed in hexane with a diameter of 19.5 ± 5.1 nm confirmed by TEM (Fig. 1a and Fig. S1a). The cubic structure with the characteristic peaks of (1 1 1), (2 0 0), (2 2 0), (3 1 1), and (222) was well-resolved from the X-ray diffraction (XRD) pattern. To ensure that the hydrophobic MnO NPs were dispersible in aqueous media, the reverse microemulsion method was used to coat the surface of the MnO NPs with silica [12–14]. After coating the silica shell, the average diameter of the as-prepared MnO@SiO2 NPs was measured to be 39.3 ± 3.7 nm (Fig. 1b and Fig. S1b). The peak located at 2 = 23◦ in the XRD pattern (Fig. 1c) and a strong broad band at ∼1109 cm−1 in the FT-IR spectrum of MnO@SiO2 NPs (Fig. S1c) both confirmed the existence of the silica shell. The hydrodynamic radius determined by DLS was ∼87 nm with a zeta potential of −25 mV (Fig. S1d and e) in aqueous media. To assess the colloidal stability of MnO@SiO2 NPs under physiological conditions, change in the hydrodynamic size as a function of time was monitored in RPMI 1640 plus 10% FBS. After 36 h, the size

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Fig. 1. Representative TEM images of as synthesized MnO (a), and MnO@SiO2 NPs (b). (c) X-ray diffraction pattern of MnO (black) and MnO@SiO2 NPs (red). (d) The change of the hydrodynamic size of MnO@SiO2 NPs incubated in RPMI 1640 plus 10% FBS. Values were the means ± SD from three independent experiments. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

varied only from ∼92 nm to ∼112 nm (Fig. 1d), which was stable enough for further biomedical application in vitro.

was lower than 30% at the concentration of 200 ␮g/mL after 24 h. Taken together, these findings showed that L929 cells were more sensitive than HeLa cells and had lower viability.

3.2. Cell viability 3.3. Cellular morphology The viability of HeLa and L929 cells incubated with concentrations of MnO@SiO2 NPs from 10 to 200 ␮g/mL for 0.5 h, 1 h, 4 h, 8 h, 12 h and 24 h were determined using the MTT assay [15,16], respectively. As shown in Fig. 2a and b, the toxicity of MnO@SiO2 NPs was highly time-dependent. The viability of HeLa cells maintained above 90% in 4 h. However, the viability decreased significantly after 8 h. After the incubation for 12 h and 24 h, respectively, the viability was below 80%, even less than 50% at the maximum concentration of 200 ␮g/mL. For L929 cells, the viability was less than 90% gradually after 4 h, and rapidly decreased after 8 h. The viability

In addition to testing the global effect of MnO@SiO2 NPs on cell viability, optical microscopy and TEM were used to observe changes in cell morphology. Under an optical microscope, the nuclear region and cytoplasm were seen in both HeLa and L929 cells in the control group (Fig. 3a1 and b1). After incubation with MnO@SiO2 NPs at 50 ␮g/mL for 24 h, morphological characteristics of cell death and cell damage, such as loss of cell/cell contact between neighboring cells, cytoplasm retraction in both cell types, shrunken nuclei, and multinucleated giant cells, were observed [17]. The cell nuclei were

Fig. 2. The viability of HeLa (a) and L929 cells (b) after the incubation of different concentrations of MnO@SiO2 NPs after 0.5, 1, 4, 8, 12, 24 h, respectively. Values were the means ± SD from three independent experiments, * p < 0.05 versus control cells.

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Fig. 3. The control of HeLa (a1) and L929 cells (b1) stained with H&E. HeLa (a2) and L929 cells (b2) incubated with MnO@SiO2 NPs (50 ␮g/mL) for 24 h, then stained with H&E. TEM images of HeLa cells (c1) and L929 cells (d1). TEM images of HeLa cells (c2) and L929 cells (d2) incubated with 50 ␮g/mL MnO@SiO2 NPs. N represents the nucleus. Red dots represent the border of the nucleus. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

shriveled (Fig. 3a2 and b2 with red arrows). Retracted cytoplasm is shown by black arrows. TEM analysis of HeLa and L929 cells incubated with 50 ␮g/mL of MnO@SiO2 NPs was also performed. As shown in the TEM images of whole cells (Fig. 3c1 and d1), many cilia and protrusions distributed around the cell membrane were clearly observed in both cell types, whereas the number of cilia and protrusions was reduced following incubation with MnO@SiO2 NPs (Fig. 3c2 and d2). Furthermore, the reduced number of cilia and protrusions was larger for L929 cells than for HeLa cells. In addition, the membrane structure of L929 cells became blurred and was difficult to identify, which

further indicated that L929 cells were more sensitive than HeLa cells. 3.4. Oxidative stress in cells In general, oxidative stress induced by nanoparticles causes mitochondrial dysfunction, and initiates cell apoptosis [18,19]. To obtain an insight into oxidative stress caused by MnO@SiO2 NPs, the production of ROS, which is generally considered a sign of oxidative stress [20], and malondialdehyde (MDA) was evaluated. It is known that nanoparticles increase the production of ROS and cause

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Fig. 4. (a) The ROS level of HeLa and L929 cells after incubation with different concentrations of MnO@SiO2 NPs for 24 h by DCFH-DA assay. Rosup (50 ␮g/mL) was used as a positive control. (b) The percentage of intracellular MDA level of HeLa and L929 cells after incubation with MnO@SiO2 NPs (50 ␮g/mL) for 24 h compared to the control group. Values were the means ± SD from three independent experiments, * p < 0.05 versus control cells.

Fig. 5. Representative plots of the change of MMP for HeLa cells (a1), and HeLa cells treated with MnO@SiO2 NPs of 10 ␮g/mL (a2), 25 ␮g/mL (a3), 50 ␮g/mL (a4). Representative plots of the change of MMP of L929 cells (b1), and L929 cells treated with MnO@SiO2 NPs of 10 ␮g/mL (b2), 25 ␮g/mL (b3), 50 ␮g/mL (b4). R represents the percentage of red fluorescence. G represents the percentage of green fluorescence. (c) The histogram of MMP for HeLa and L929 cells. Values were the means ± SD from three independent experiments, * p < 0.05 versus control cells. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

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Fig. 6. The percentage of HeLa (a) and L929 (b) cells in each phase of the cell cycle incubated with different concentrations of MnO@SiO2 NPs for 24 h. Values were the means ± SD from three independent experiments, * p < 0.05 versus control cells.

cell death in different types of cultured cells [21]. The extent of ROS production was determined by the fluorescence intensity of dichlorofluorescein (DCF). As shown in Fig. 4a, following incubation with MnO@SiO2 NPs for 24 h, intracellular ROS in HeLa and L929 cells increased gradually with increasing concentrations of MnO@SiO2 NPs. The generation of ROS increased slowly below 50 ␮g/mL, but increased rapidly when the concentration was higher than 50 ␮g/mL. The fluorescence intensity of DCF increased 4.5-fold in HeLa cells at 200 ␮g/mL and increased 5.5-fold in L929 cells. The reactive intermediates produced by oxidative stress can change membrane bilayers and lead to lipid peroxidation [22,23]. MDA, which is a lipid peroxidation product, was measured in this system [24]. As shown in Fig. 4b, the content of MDA increased as the concentration of MnO@SiO2 NPs increased. At the maximum concentration in our experimental range (200 ␮g/mL), the content of MDA in HeLa cells was 1.8-fold higher than that in the control. However, the content of MDA in L929 cells was 2.3-fold higher under similar incubation conditions, which again suggested that L929 cells were more sensitive than HeLa cells. The decrease in cell viability and increase in ROS and MDA suggested that cell death is mainly caused by membrane damage due to lipid peroxidation, which corresponds with a previous report that oxygen radicals are capable of directly oxidizing DNA, proteins, and lipids [25]. 3.5. Mitochondrial membrane potential (MMP) of cells Given that ROS has a significant association with mitochondria, and oxygen radicals are mainly enriched in mitochondria [26], it is necessary to assess the likely impact on mitochondria. Therefore, MMP, an indicator of mitochondrial activity [27], was measured in cells using flow cytometry. Normal MMP is expressed by the region of R, and reduced MMP is expressed by the region of G. Higher MMP was defined by the higher ratio of red fluorescence from J-aggregate

at the hyperpolarized membrane potential to green fluorescence from a monomer at depolarized membrane potential [28]. After incubation with different concentrations of MnO@SiO2 NPs for 24 h, the ratio of red fluorescence intensity to green fluorescence intensity decreased gradually, indicating increased depolarization of mitochondria induced by MnO@SiO2 NPs. As shown in Fig. 5c, the ratio was decreased in a concentration-dependent manner. These results indicated that intracellular ROS caused mitochondrial dysfunction, and induced apoptosis in both HeLa and L929 cells. 3.6. Effect on cell cycle The generation of ROS can cause DNA damage, and the early effect is seen in cell cycle progression [29]. Therefore, the influence of MnO@SiO2 NPs on the cell cycle was analyzed according to the results of HeLa and L929 cells flow cytometry. Both HeLa and L929 cells showed a concentration-dependent G2/M arrest, which was confirmed by an increase in the percentage of cells in the G2/M phase and a corresponding decrease in the G1 phase with increased concentrations of MnO@SiO2 NPs (the original data in Fig. S4). As shown in Fig. 6a, G2/M arrest was observed in HeLa cells only at concentrations greater than 50 ␮g/mL. Following incubation with MnO@SiO2 NPs at 200 ␮g/mL for 24 h, the percentage of cells in the G2/M phase was approximately 200% compared to the control group. However, for L929 cells, even when incubated at 10 ␮g/mL, a slight increase in the percentage of cells at G2/M was observed (Fig. 6b). At the highest concentration of 200 ␮g/mL in our experimental conditions, the percentage of cells in the G2/M phase was nearly 300% compared to the control group, which further confirmed that L929 cells were more sensitive than HeLa cells. G2/M arrest indicated that DNA repair was needed at the G2/M checkpoint, if repair occurred then the cell cycle would progress, or

Fig. 7. The percentages of apoptosis and necrosis of HeLa cells (a) and L929 cells (b) after the incubation with different concentrations of MnO@SiO2 NPs for 24 h at 37 ◦ C. Values were the means ± SD from three independent experiments, * p < 0.05 versus control cells.

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apoptosis would be activated [30,31], however, these two processes were not conducive for cell proliferation. In general, emergence of the Sub-G1 phase is often accompanied by apoptosis. As shown in Fig. 6, the percentage of both HeLa and L929 cells in the Sub-G1 phase increased with increased concentrations of MnO@SiO2 NPs, indicating apoptosis. 3.7. Apoptosis and necrosis To further verify the apoptosis process induced by MnO@SiO2 NPs, Annexin-V/PI double staining was carried out. Annexin V (aV) which is easily reacted with phosphatidylserine (PS) was used as a marker of apoptosis. Propidium iodide (PI), which was utilized to detect plasma membrane integrity, was employed to detect necrotic cell death [32]. Statistical data were extracted from the pseudo-color dot plot (Fig. S5) using FlowJo software 7.6.5, based on the percentage of aV−/PI− (viable cells), aV−/PI+ (necrotic cells),

aV+/PI− (apoptotic cells), and aV+/PI+ (late apoptotic cells). The data on HeLa and L929 cells from the annexin-V staining experiment are summarized in Fig. 7. Compared to the negative control group, the percentage of viable cells decreased significantly. Both HeLa and L929 cells underwent apoptosis following incubation with MnO@SiO2 NPs. Furthermore, apoptosis was dose-dependent. The proportion apoptosis of L929 cells was higher than that of HeLa cells. 3.8. The intracellular apoptotic signaling pathway by MnO@SiO2 NPs To explore the possible mechanism of MnO@SiO2 NPs-mediated apoptosis and cell cycle arrest, we measured the expression level of regulators involved in the apoptotic pathway and G2/M DNA damage checkpoint. It can be seen from Fig. 8b and c that p53 phosphorylation increased dose-dependently in both types of cells

Fig. 8. (a) Western blot analysis of HeLa and L929 cells. The expression level of p53, bax and bcl-2 for HeLa (b) and L929 cells (c) after the incubation with different concentrations of MnO@SiO2 NPs (0, 10, 25 and 50 ␮g/mL, respectively). (d) The caspase-3 activity of HeLa and L929 cells after the incubation with MnO@SiO2 NPs for 24 h. (e) The schematic mechanism of the signal pathway of apoptosis induced by MnO@SiO2 NPs. Values were the means ± SD from three independent experiments, * p < 0.05 versus control cells.

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when exposed to MnO@SiO2 NPs after 24 h. It is generally believed that ROS activates the p53 signal transduction pathway [19]. In response to cellular damage, p53 is of importance in genomic surveillance by monitoring genomic instability [33–36]. Bax and bcl-2 are also involved in mitochondrial-dependent apoptosis. Bcl2 family proteins and bax regulate outer mitochondrial membrane permeability and control the on/off intrinsic apoptotic pathway [37–40]. As shown in Fig. 8b and c, the expression of bcl-2 showed a decreasing trend, while the expression of bax was similarly increased in both HeLa and L929 cells. The change in the bax/bcl-2 ratio resulted in significant activation of caspases followed by induction of apoptosis [41,42]. It is believed that bax and bcl-2 are activated by ROS. Caspase-3 activity is unique to apoptosis, as it does not occur in other forms of cell death and provides strong evidence for the presence of apoptosis [43,44]. Caspase-3 activity was also assessed following incubation with MnO@SiO2 NPs for 24 h and compared to the control group. The activity of caspase-3 was expressed as a percentage compared to that of the control cells. As shown in Fig. 8d, there was no significant increase in caspase-3 activity at low concentrations of MnO@SiO2 NPs. After incubation with MnO@SiO2 NPs at 200 ␮g/mL, the activity of caspase-3 was more than 1.3-fold in HeLa and 1.6-fold in L929 cells, respectively, which suggested that apoptosis was the main form of cell death induced by MnO@SiO2 NPs for both HeLa and L929 cells. The apoptotic signal pathway by MnO@SiO2 is shown in Fig. 8e. After exposure to MnO@SiO2 NPs, ROS was induced in cells, then p53 activation followed by an increase in bax and a decrease in bcl-2, ultimately leading to G2/M phase arrest and apoptosis. 4. Conclusions In summary, HeLa and L929 cells were used as model cell lines to investigate the cytotoxicity of MnO@SiO2 NPs. L929 cells were affected more seriously by MnO@SiO2 NPs than HeLa cells. Internalized MnO@SiO2 NPs induced intracellular ROS generation, then p53 was activated followed by an increase in bax and a decrease in bcl-2, ultimately leading to G2/M arrest, increasing caspase-3 activity and induction of apoptosis. Our overall findings on the cytotoxicity of MnO@SiO2 NPs should be helpful for further application in vivo. Acknowledgements This work was partially supported by National Natural Science Foundation of China (Nos. 21271130 and 21371122), program for Changjiang Scholars and Innovative Research Team in University (No. IRT1269), Shanghai Science and Technology Fund Project (Nos. 12ZR1421800 and 13520502800), Pujiang Program of Shanghai Education Commission in China (13PJ1406600), Shanghai Municipal Education Commission (No. 13ZZ110) and Shanghai Normal University (Nos. DXL122 and SK201339).

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