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Copper(II) oxide nanoparticles penetrate into HepG2 cells, exert cytotoxicity via oxidative stress and induce pro-inflammatory response† Jean-Pascal Piret,*a Diane Jacques,a Jean-Nicolas Audinot,b Jorge Mejia,c Emmanuelle Boilan,a Florence No€el,a Maude Fransolet,a Catherine Demazy,a Stephane Lucas,c Christelle Saouta and Olivier Toussaint*a Received 9th July 2012, Accepted 24th September 2012 DOI: 10.1039/c2nr31785k The potential toxic effects of two types of copper(II) oxide (CuO) nanoparticles (NPs) with different specific surface areas, different shapes (rod or spheric), different sizes as raw materials and similar hydrodynamic diameter in suspension were studied on human hepatocarcinoma HepG2 cells. Both CuO NPs were shown to be able to enter into HepG2 cells and induce cellular toxicity by generating reactive oxygen species. CuO NPs increased the abundance of several transcripts coding for proinflammatory interleukins and chemokines. Transcriptomic data, siRNA knockdown and DNA binding activities suggested that Nrf2, NF-kB and AP-1 were implicated in the response of HepG2 cells to CuO NPs. CuO NP incubation also induced activation of MAPK pathways, ERKs and JNK/SAPK, playing a major role in the activation of AP-1. In addition, cytotoxicity, inflammatory and antioxidative responses and activation of intracellular transduction pathways induced by rod-shaped CuO NPs were more important than spherical CuO NPs. Measurement of Cu2+ released in cell culture medium suggested that Cu2+ cations released from CuO NPs were involved only to a small extent in the toxicity induced by these NPs on HepG2 cells.
1. Introduction Among the manufactured metal oxide nanoparticles (NPs), copper oxide (CuO) NPs are used in industrial catalysis and are components of gas sensors, batteries, solar energy converters and high-temperature superconductors.1 CuO NPs are also used for their antimicrobial activity with possible applications to disposable textiles or food containers.2–4 The increasing production of metal oxide NPs, particularly CuO NPs, has led to major concerns regarding the potential hazards for human health. Several in vitro studies investigated the potential toxic effects of CuO NPs, mainly on airway cells.5–8 CuO NPs were shown to trigger the production of Reactive Oxygen Species (ROS), to generate DNA damage5 and to induce
a URBC, Namur Nanosafety Center (NNC), Namur Research Institute for Life Sciences (NARILIS), University of Namur (FUNDP), 61 rue de Bruxelles, B-5000 Namur, Belgium. E-mail:
[email protected]; olivier.
[email protected]; Fax: +32 81 72 41 35; Tel: +32 81 72 41 32 b Department of Science and Analysis of Materials, Centre de Recherche Public – Gabriel Lippmann, 41 rue du Brill, L-4422 Belvaux, Luxembourg. Tel: +352 47 02 61 521 c Physics of Matter and Radiation (PMR-LARN), NNC-NARILIS, University of Namur (FUNDP), 61 rue de Bruxelles, B-5000 Namur, Belgium. Fax: +32 81 72 54 74; Tel: +32 81 72 54 81 † Electronic supplementary information (ESI) available: Additional tables and figures supporting the information presented in the manuscript. See DOI: 10.1039/c2nr31785k
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mitochondrial depolarisation in A549 cells.6 CuO NPs also induced a concentration- and time-dependent cytotoxicity and elicited the permeability and inflammation response in human cardiac microvascular endothelial cells9 and differentiated Caco2 intestinal cell monolayers.10 In vivo studies described lung toxicity of CuO NPs after intratracheal instillation in rats. Pulmonary edema was observed with macrophages and neutrophil infiltration in the alveoli after one day of exposure and lethality after 28 days of exposure.11 ROS can damage proteins, lipids and DNA and participate in the development of diseases like cancer, arteriosclerosis and arthritis.12 The mammalian oxidative stress response is mainly coordinated by the transcription factor, nuclear factor erythroid 2-related factor 2 (Nrf2).13,14 Under basal conditions, Nrf2 is targeted for proteasomal degradation by Keap1 and Nrf2-regulated genes are weakly expressed. Under oxidative stress, transducers such as mitogen-activated protein kinases (MAPKs), protein kinase C and phosphatidylinositol 3-kinase can phosphorylate Keap1 and Nrf2, disrupting the Keap1–Nrf2 complexes and allowing nuclear translocation of Nrf2. In addition, ROS also directly oxidize thiol groups of Keap1 protein15 which could act as a primary sensor for oxidative stress.16 Following partnering with small Maf proteins, activated Nrf2 binds to antioxidant response elements located within the promoter region of target genes and induces their transcription. NPs-induced activation of Nrf2 has recently been described: This journal is ª The Royal Society of Chemistry 2012
CeO2 or SiO2 NPs were shown to induce oxidative stress in Beas2B human bronchial epithelial cells, leading to the activation of p38MAPK–Nrf2 or ERKs–Nrf2 signalling pathways respectively.17,18 Multi-walled carbon nanotubes activate nuclear translocation of Nrf2 and induce the expression of the Nrf2 target gene, heme oxygenase 1 (HMOX1).19 Other redox-sensitive transcription factors like Activator Protein-1 (AP-1) or Nuclear factor kappa B (NF-kB) were also described to be activated after exposure to NPs.20,21 NF-kB consists of homodimers or heterodimers of Rel proteins that share a Rel homology domain in their N-termini. NF-kB regulates a large number of genes related to immune function, inflammation, apoptosis, cell proliferation, and synaptic plasticity.22 In
unstimulated cells, NF-kB dimers are sequestered in the cytoplasm by a family of inhibitors, IkBs, among which IkBa is the best known. Various stimuli like oxidative stress, cytokines, and UV irradiation induce phosphorylation of IkBa leading to its ubiquitination and proteasomal degradation. Free NF-kB dimers migrate to the nucleus and activate gene transcription.23 AP-1 is a homodimeric or heterodimeric transcription factor composed of basic region-leucine zipper (bZip) proteins that belong to the Fos, Jun, ATF, and Maf subfamilies. AP-1 regulates the induction of a variety of genes in response to a host of stimuli, such as oxidative stress, cytokines, growth factors, and infections, and thereby controls a number of cellular processes including differentiation, proliferation, and apoptosis.24
Fig. 1 TEM micrographs revealing the morphological variations of CuO NPs (A and B). Size distribution analysis of rod-shaped (R. CuO) and spherical (S. CuO) CuO NPs by centrifugal liquid sedimentation (C–F). Agglomeration state profiles were obtained for CuO NPs diluted at 100 mg ml1 into DMEM cell culture medium. Results are displayed as relative CuO NPs weight (C) or number (E) distribution against the hydrodynamic diameter. Maximal values in each sample were arbitrarily normalised to 100. Tables (D) and (F) give quantitative values of size distribution analysis of CuO NPs suspensions by centrifugal liquid sedimentation. Results are presented as relative weight (D) or number (F) percentages of CuO NPs detected between selected hydrodynamic diameter ranges, chosen in order to span the width of detection peaks.
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Fig. 2 (A) Amount of released Cu cations versus concentration of Cu from 4 mg ml1 to 80 mg ml1 of rod-shaped CuO NPs (i.e., 5–100 mg ml1 NPs; R. CuO), spherical CuO NPs (i.e., 5–100 mg ml1 NPs; S. CuO) or CuCl2 (i.e., 11–215 mg ml1 CuCl2) incubated for 24 h into DMEM cell culture medium. Results are expressed in fold increase compared to control. (B) Effect of CuO NPs or CuCl2 on HepG2 cell viability. HepG2 cells were incubated for 24 h in the presence or absence (control, CTL) of 4 mg ml1 to 80 mg ml1 of Cu from CuO NPs or CuCl2. The results are expressed as means s.d. (n ¼ 3), and presented as percentages of CTL. *: P < 0.05, **: P < 0.01 or ***: P < 0.001 vs. CTL; #: P < 0.05, ##: P < 0.01 or ###: P < 0.001 vs. R. CuO NPs at the same concentration; $$: P < 0.01 or $$$: P < 0.001 vs. S. CuO NPs at the same concentration.
In this study, we tested the effects of two categories of CuO NPs of different shapes and specific surface areas and with similar hydrodynamic diameter in solution on viability, ROS production, activation of redox-sensitive transcription factors and inflammatory response of the HepG2 hepatocarcinoma cell line, a classical hepatic model used to test compounds that are potentially cytotoxic, genotoxic or affect hepatocyte functions.25
2. Results 2.1 Characterization and dispersion of CuO NPs In this study we investigated two types of CuO NPs purchased from IoLiTec. According to the data sheet, the first type of CuO NPs has an average size of 12 nm (12 nm CuO NPs) and the second an average size between 50 and 80 nm (50 nm CuO NPs). The main physico-chemical properties of these NPs were described previously.10 Briefly, SEM and TEM analyses revealed that the raw powders 12 nm CuO NPs were rod-shaped with an average of 10 3 nm in thickness and 74 17 nm in length, whereas 50 nm CuO NPs appeared spherical (average of 40 16 nm in diameter). CuO NPs morphologies are illustrated in Fig. 1A and B. For the rest of the study, we decided to coin the CuO NPs based on their respective shape: rod-shaped CuO NPs (12 nm CuO NPs) and spherical CuO NPs (50 nm CuO NPs). EDX and XPS analyses revealed that both CuO NPs were composed of approximately half of Cu and O.10 The specific surface areas were 18.4 m2 g1 and 6.8 m2 g1 for the rod CuO NPs and the spherical CuO NPs, respectively. 7170 | Nanoscale, 2012, 4, 7168–7184
Before cell incubation, CuO NPs (from stock water suspensions) were diluted in DMEM cell culture medium containing 10% of fetal bovine serum necessary for the growth of HepG2 cells. The particle size distribution and the agglomeration state of both categories of CuO NPs in cell culture medium were characterized by centrifugal liquid sedimention.26–28 Results were expressed in relative weight to reveal the size distribution of CuO NP agglomerates (Fig. 1C) and in relative particle number (Fig. 1E) to distinguish the size distribution of individualized NPs. The maximal values in each sample were arbitrarily set to 100%. Interestingly, both categories of CuO NPs displayed a similar agglomeration state in the culture medium. Rod-shaped CuO NPs and spherical CuO NPs mainly formed agglomerates with respective maximum peaks (corresponding to the maximum weight measured) of 364 nm and 597 nm of hydrodynamic diameter values. A small proportion of CuO was present as individualized particles (24.5% and 11.9% respectively, Fig. 1C and Table D in Fig. 1). The size distribution of individualized rod-shaped CuO NPs or spherical CuO NPs appeared to be virtually identical with peaks spanning around 6.8 nm and 8.7 nm respectively (Fig. 1E and Table F in Fig. 1). Moreover, the size distribution of the majority of well-dispersed rod-shaped CuO NPs (97%) and spherical CuO NPs (95%) was included between 5 and 20 nm (Table F, Fig. 1). 2.2 Cytotoxicity of CuO NPs on HepG2 cells: a potential role of Cu cations We showed previously that the release of Cu cations partly mediates the toxicity of both rod-shaped and spherical CuO NPs This journal is ª The Royal Society of Chemistry 2012
Fig. 3 (A) Production of ROS in HepG2 cells after different incubation times with CuO NPs. Cells were incubated in the presence or absence (control, CTL) of 5 mg ml1 to 50 mg ml1 of CuO NPs for increasing incubation times. ROS were detected by fluorescence measurement of the oxidized 2,7-DCF probe. H2O2 (500 mM) was used as a positive control. Results in relative fluorescence units (RFU) per mg of proteins are expressed as means s.d. for n ¼ 3 and normalized to control. *: P < 0.05, **: P < 0.01 or ***: P < 0.001 vs. the corresponding control. (B) Effect of N-acetyl-cystein (NAC, 10 mM) on CuO NP-induced toxicity in HepG2 cells. HepG2 cells were preincubated for 1 h with NAC and then exposed for 24 h to 25 or 50 mg ml1 of rod-shaped (R. CuO) or spherical (S. CuO) CuO NPs in the presence or absence of NAC. The results are expressed as means s.d. (n ¼ 4), and presented as percentages of CTL. **: P < 0.01 or ***: P < 0.001 vs. the corresponding control; ###: P < 0.001 vs. the corresponding condition without NAC at the same concentration.
on intestinal Caco-2 cell monolayers.10 Here we compared the toxicity of both CuO NPs to CuCl2 on HepG2 cells. We expressed the concentration of Cu equivalent added to the culture wells in mg ml1 of Cu alone, regardless of the presence of O in CuO or Cl in CuCl2. Both CuO NPs and CuCl2 were shown to release Cu2+ in the DMEM culture medium in a dosedependent manner with rod-shaped CuO NPs releasing a stronger amount of Cu2+ than spherical CuO NPs (Fig. 2A). As expected, at all concentrations tested, more Cu2+ was released by CuCl2 than CuO NPs. The impact of CuCl2 and CuO NPs on the viability of HepG2 cells was investigated after 24 h of incubation (Fig. 2B). Both CuO NPs induced a dose-dependent decrease of cell viability, rod-shaped CuO NPs appearing to be more toxic than spherical CuO NPs at higher concentrations of 40 and 80 mg ml1 of Cu equivalent. CuCl2 was cytotoxic, albeit less than CuO NPs, at 80 mg ml1 of Cu equivalent. This suggested that Cu2+ released from CuO NPs was involved only to a small extent in the toxicity induced by these NPs on HepG2 cells, at the highest concentration tested. 2.3 Toxicity of CuO NPs is mediated by oxidative stress The pro-oxidant effect of CuO NPs was studied by following the oxidation of the 2,7-DCF probe after different incubation times (Fig. 3A). No probe oxidation was observed after 2 h of incubation with CuO NPs. Both CuO NPs increased doseThis journal is ª The Royal Society of Chemistry 2012
dependently the oxidation of 2,7-DCF from 4 h to 24 h of incubation. In order to investigate whether oxidative stress could play a role in the cytotoxicity of CuO NPs, HepG2 cells were exposed to CuO NPs in the presence of the anti-oxidant N-acetylcystein (NAC). NAC abolished almost fully the harmful effect of both CuO NPs at all concentrations studied (Fig. 3B). 2.4 Intracellular localization of CuO NPs In order to investigate whether CuO NPs could penetrate into the cells, the intracellular localization of CuO NPs was analysed by the Nano Secondary Ion Mass Spectrometry (NanoSIMS) technique. The subcellular distribution of Cu in individual cells was mapped by the measurement of the intensity of the two isotopes of copper (63Cu and 65Cu) in HepG2 cells exposed or not (CTL) to 25 mg ml1 of rod-shaped or spherical CuO NPs for increasing times. The isotopic ratio (2.22) between 63Cu and 65Cu was verified and was in agreement with the natural abundance. Only images of the isotope 63Cu are presented here. In order to provide cell background images, ion mapping of the main elements that occurs in biological tissues, cluster carbon– nitrogen (12C14N), phosphorus (31P) and sulphur (34S) was detected in parallel. The images of secondary ions of these elements were used to correlate the images of ions with the cellular structure. Indeed, the distribution of these elements gives images similar to that obtained by optical microscopy.29 Fig. 4A Nanoscale, 2012, 4, 7168–7184 | 7171
Fig. 4 (A) NanoSIMS50 analysis of HepG2 cells after different incubation times in the presence or absence (control, CTL) of rod-shaped (R. CuO) or spherical (S. CuO) CuO NPs (25 mg ml1) showing distribution of 31P (grey) and 63Cu (red). Increasing brightness indicates stronger secondary ion intensities within the image. Scale bar: 5 mm. (B) NanoSIMS50 analysis of HepG2 cells after different incubation times in the presence or absence (control, CTL) of rod-shaped (R. CuO) or spherical (S. CuO) CuO NPs (25 mg ml1) showing distribution of 34S (green) and 63Cu (red). Increasing brightness indicates stronger secondary ion intensities within the image. Arrows indicate the superposition of 34S and 63Cu signals. Scale bar: 5 mm.
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and B show signals arising from secondary ions of 63Cu and 31P or 63Cu and 34S respectively. The phosphorus and the sulfur signals allow us here to map the phospholipid membrane, nucleus (DNA), and imaging the global cell morphology.29 Intracellular localization of Cu was observed from 2 h to 24 h of incubation with both CuO NPs (Fig. 4A). No signal arising
from secondary ions of 63Cu was detected from HepG2 cells not exposed to CuO NPs (CTL). Superimposition of the 63Cu secondary ion signal with that of 31P allowed us to observe a cytoplasmic and perinucleus localization of Cu. Interestingly, an intracellular overlapping between 63Cu and 32S signals was observed in CuO NP-treated cells from 6 h to 24 h of incubation
Table 1 Effects of CuO NPs on the abundance of pro-inflammatory mRNAs in HepG2 cells. Results of the transcriptomic profiling (RT real time TaqMan PCR arrays), considering 96 genes involved in the immunity or pro-inflammatory response, of HepG2 cells treated for 24 h with 25 mg ml1 of CuO NPs. Only 42 transcript species were abundant enough to be detected in the control or CuO NPs-treated HepG2 cells. Results are expressed as the ratios of the transcript levels in treated cells vs. untreated cells (mean ratios s.d., n ¼ 3). All mean ratios are ranked in the decreasing order in cells treated with rod-shaped CuO NPs (R. CuO). Grey boxes indicate statistically significant effects with P < 0.05. N.D., not detected. Values in the spherical CuO NP (S. CuO) column underlined in red indicate similar values to those obtained after rod-shaped CuO NPs treatment
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(Fig. 4B), suggesting an accumulation of Cu in sulphur-rich areas, whatever that means. 2.5 Pro-inflammatory effects of CuO NPs The inflammatory response of HepG2 cells was investigated after 24 h of incubation with rod-shaped or spherical CuO NPs (25 mg ml1) using reverse transcription (RT) real time Taqman PCR low density arrays (TLDA). These arrays allow Taqman quantitative PCR (qPCR) amplification of 96 transcripts involved in human immunity and inflammation. 42 transcript species were abundant enough to be detected in the control or CuO NPstreated HepG2 cells (listed in Table S1 with gene names and functions, ESI†). Ratios between 0.7 and 1.3 were arbitrarily considered as biologically non-relevant. These 42 transcript species were sorted along the decreased mRNA abundance in cells treated with rod-shaped CuO NPs (Table 1). When 3 independent biological replicates could be considered, the mean ratios were calculated with standard deviations (s.d.). Among the 42 transcript species detected, only a few transcripts were statistically more abundant after treatment with both CuO NPs. This was not surprising since high variability is frequently obtained when profiling gene expression in triplicates with arrays.30,31 Other changes of abundance were not statistically
significant when large s.d. values were obtained or lacking (mean ratios calculated from only two values) despite all efforts. We observed the overexpression of transcripts coding for chemokines and proteins able to recruit and activate immune cells, such as various interleukins (IL-18-12A-8-7), as well as colony stimulating factor-1/macrophage-colony stimulating factor (CSF-1/M-CSF). An increase in the amount of tumor growth factor-b1 (TGF-b1) transcript was also measured. Moreover, overexpression of mRNAs coding for proteins usually associated to oxidative stress like HMOX1 was observed. These results are in accordance with those describing ROS production shown in Fig. 3A, confirming the pro-oxidant potential of CuO NPs. In addition, an increase of the abundance of mRNA coding for NFKB2 (NF-kB p100 subunit) was also detected. NF-kB has been identified as a transcription factor regulated by the intracellular redox status inducing the expression of a variety of proteins implicated in the immunological and cellular detoxifying defense systems. Moreover, some transcripts coding for chemokines or proinflammatory cytokines were specifically more or less abundant after exposure to one of the two CuO NPs. Chemokine receptors CCR7 and CCL5, cytokine Tumor Necrosis Factor-a (TNF-a) and colony stimulating factors-3 (CSF-3) transcripts were more abundant after exposure to rod-shaped CuO NPs while C3
Fig. 5 Effects of CuO NPs on expression levels of HMOX1 and IL-8 proteins. (A) Abundance of HMOX1 protein was evaluated by western blotting after increasing incubation times with 25 mg ml1 of rod-shaped (R. CuO) or spherical (S. CuO) CuO NPs. 3-Morpholinosydnonimine hydrocloride (Sin-1, 2.5 mM, 6 h) was used as a positive control of oxidative stress. b-Actin was used as a reference protein. (B) Abundance of IL-8 released in culture medium from HepG2 cells exposed to CuO NPs. 400 000 cells were incubated for increasing incubation times in the presence or absence (control, CTL) of rod-shaped (R. CuO) or spherical (S. CuO) CuO NPs at different concentrations or with IL-1b (0.01 ng ml1) used as a positive control. The amount of IL-8 present in cell culture medium was assayed by ELISA. Results are expressed as means s.d. for n ¼ 4, in pg of IL-8 per ml. *: P < 0.05, **: P < 0.01 or ***: P < 0.001 vs. the corresponding control. (C) 50 000 cells were incubated for 24 h with or without CuO NPs (25 mg ml1) in the presence or absence of NAC (10 mM) and the amount of released IL-8 was assayed by ELISA. Results are expressed as means s.d. for n ¼ 4, in pg of IL-8 per ml. ***: P < 0.001 vs. control; ##: P < 0.01 or ###: P < 0.001 vs. the corresponding condition without NAC. No IL-8 was detected after treatment with NAC.
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transcript was less abundant. ECE1, CD4 and VEGF were more abundant after exposure to spherical CuO NPs, BCL2L1, PRF1, NOS2A, CCR7 and SMAD7 being less abundant. Increased abundance of HMOX1 and TGF-b1 transcripts was confirmed by single RT-qPCR with SYBR green as intercalating agent. Due to the primer positions, different variants of RT-PCR techniques may have different dynamic ranges giving different ranges of ratios. Rod-shaped CuO NPs induced, after 24 h of incubation, a 6.4 1.09 (p < 0.01) and 1.56 1.1-fold increase of HMOX1 and TGF-b1 transcripts respectively. Spherical CuO NPs induced a 6.4 0.77 (p < 0.001) and 1.6 1.0-fold increase of HMOX1 and TGF-b1 transcripts respectively. Overexpression of HMOX1 protein was observed by western blotting (Fig. 5A) and immunofluorescence (ESI, Fig. S1†). Accumulation of HMOX1 protein could be detected from 4 to 24 h corroborating the results of 2,7-DCF shown in Fig. 3A. Interestingly, rod-shaped CuO NPs induced a higher production of HMOX1 protein in comparison with spherical CuO NPs. Overexpression of IL-8 was also confirmed at the protein level. A time- and dose-dependent accumulation of IL-8 protein was measured in culture medium after treatment with both CuO NPs (Fig. 5B). As observed for HMOX1 protein, accumulation of IL8 was more important after treatment with rod-shaped CuO NPs than with spherical CuO NPs. Moreover, IL-8 overproduction
depended on the ROS produced by CuO NPs, as antioxidant NAC reduced the amount of IL-8 detected in culture medium (Fig. 5C). Overexpression of other transcripts like CSF-1 was also confirmed at the protein level. Both CuO NPs increased the amount of CSF-1 released in the extracellular culture medium (ESI, Fig. S2†). 2.6 Increased Nrf2, NF-kB and AP-1 DNA binding after incubation with CuO NPs Oxidative stress can activate transcription factors like Nrf2, NFkB, AP-1 or p53.32 We evaluated the DNA binding of these transcription factors after increasing the incubation time of HepG2 cells in the presence of CuO NPs (Fig. 6). An increase of the DNA binding of Nrf2 occurred from 4 h of exposure to CuO NPs (25 mg ml1) onwards and was comparable to the oxidant positive control 3-morpholinosydnonimine hydrocloride (Sin-1) (Fig. 6A). Nuclear translocation of Nrf2 was revealed by immunofluorescence at 6 h of incubation in the presence of both CuO NPs (Fig. 6B). A slight increase in the DNA binding of NF-kB was also observed after 4 h of incubation with CuO NPs. However, the NF-kB DNA binding activity decreased after 6 h of incubation (Fig. 6C) suggesting that the potential role of NF-kB in the cell
Fig. 6 Effects of CuO NPs on activation of redox-sensitive transcription factors. Analysis of Nrf2 (A), NF-kB (C) and AP-1 (E) DNA binding to their respective consensus sequence using a colorimetric assay. HepG2 cells were incubated for increasing incubation times in the presence or absence (control, CTL) of rod-shaped (R. CuO) or spherical (S. CuO) CuO NPs (25 mg ml1). Sin-1 (2.5 mM, 6 h) or IL-1b (0.01 ng ml1, 30 min) was used as a respective positive control. Nuclear proteins from Jurkat cells incubated 6 h with phorbol-12-myristate-13-acetate (PMA, 0.1 ng ml1) and ionomycin (iono., 1 mM) were used as internal positive controls for AP-1 DNA binding activity. Results in optical density (O. D.) are expressed as means s.d. for n ¼ 3 and normalized to control. *: P < 0.05, **: P < 0.01 or ***: P < 0.001 vs. the corresponding control. (B) Nuclear translocation of Nrf2 (green) was studied by immunofluorescence after 6 h of incubation with CuO NPs. (D) Abundance of P-c-Jun was assayed by western blotting after increasing incubation times with or without (control, CTL) 25 mg ml1 of rod-shaped (R. CuO) or spherical (S. CuO) CuO NPs. b-Actin was used as a reference protein.
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response to CuO NPs is likely limited or transient. Activation of AP-1 was firstly evaluated by detecting phosphorylated c-Jun at increasing times of incubation with CuO NPs. The level of phospho-c-Jun increased between 4 h and 6 h but decreased after 24 h of incubation with CuO NPs, suggesting a transient activation of AP-1 (Fig. 6D). This was confirmed by evaluating the DNA binding activity of AP-1 (Fig. 6E). The DNA binding activity of AP-1 increased between 4 and 6 h of incubation with CuO NPs but decreased at 24 h. DNA binding of p53 and Hypoxia Inducible Factor-1 (HIF-1) whose activation can be modulated by ROS or copper33 was also evaluated. No modification of the DNA binding of these transcription factors was observed after CuO exposure (data not shown).
2.7 Inhibition of Nrf2, NF-kB or AP-1 modulates differently the expression of IL-8 and HMOX-1 The transcriptomic data indicated that CuO NPs increased the abundance of some transcripts like HMOX1 and IL-8, known to be regulated by redox-sensitive transcription factors.34–38 Implication of Nrf2, NF-kB and AP-1 in the regulation of IL-8 and HMOX-1 expression was investigated using specific siRNA antiNrf2, p65 (NF-kB) and c-Jun (AP-1). Effects of Nrf2 inhibition (more than 90% inhibition of Nrf2 mRNA) were evaluated at the transcriptional level after 24 h of incubation with CuO NPs (25 mg ml1). CuO NPs-induced overexpression of mRNA coding for HMOX1 (5.4 2.8-fold increase – p < 0.05 – and 5.3 1.9-fold increase – p < 0.05 – for rod-shaped and spherical CuO NPs
respectively) and IL-8 (3.8 2.2-fold increase and 2.8 0.3-fold increase – p < 0.01 – for rod-shaped and spherical CuO NPs respectively; 24 h of incubation) was reduced after transfection of specific Nrf2 siRNA (HMOX1: 2.3 0.4-fold increase and 2.5 0.9-fold increase for rod-shaped and spherical CuO NPs respectively; IL-8: 2.3 1.6-fold increase and 1.8 0.6-fold increase – p < 0.05 – for rod-shaped and spherical CuO NPs respectively) while no effect of non-targeting siRNA was observed (HMOX1: 4.7 1.2-fold increase and 6.2 2.1-fold increase for rod-shaped and spherical CuO NPs respectively; IL-8: 4.5 1.3-fold increase and 3.4 0.6-fold increase for rod-shaped and spherical CuO NPs respectively). These results were confirmed at the protein level. Inhibition of Nrf2 by siRNA reduced the amount of HMOX1 protein measured after CuO NPs treatment (Fig. 7A) and the quantity of IL-8 released into the culture medium (Fig. 7B). siRNA inhibition of p65 or c-Jun also reduced the amount of IL-8 (Fig. 7D and F) but has no effect on HMOX1 expression (Fig. 7C and E), suggesting a cooperative action of these different transcription factors in the expression of some CuO NPs-induced transcripts like IL-8. A potential regulation of CSF-1 by Nrf2 was evaluated by ELISA. Even in the absence of statistical significance, there was overall a tendency of a decrease of the amount of secreted CSF-1 after inhibition of Nfr2 (ESI, Fig. S3†).
2.8 Activation of MAPKs in CuO NP-treated HepG2 cells To determine whether CuO NPs activated mitogen-activated protein kinases (MAPKs) pathways, phosphorylated p38MAPK
Fig. 7 Effects of inhibition of Nrf2 (A and B), p65 (NF-kB, C and D) and c-Jun (AP-1, E and F) on the expression levels of HMOX1 and IL-8 after 24 h of incubation with CuO NPs. (A, C and E) Abundance of HMOX1 was evaluated by western blotting after 24 h of incubation with or without rodshaped (R. CuO) or spherical (S. CuO) CuO NPs (25 mg ml1) and transfection with or without non-targeting siRNA or specific siRNA. a-Tubulin was used as a reference protein. (B, D and F) Abundance of IL-8 released in culture medium from HepG2 cells exposed to CuO NPs (25 mg ml1) for 24 h after transfection with or without non-targeting siRNA or specific siRNA was assayed by ELISA. Results are expressed as means s.d. for n ¼ 3, in pg of IL-8 per ml. **: P < 0.01 or ***: P < 0.001 vs. control; #: P < 0.05 or ##: P < 0.01 vs. the corresponding condition without siRNA.
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(P-p38MAPK), phosphorylated extracellular-receptor kinases (P-ERKs) and phosphorylated c-Jun N-terminal kinase/stressactivated protein kinase (P-JNK/SAPK) were detected by western blotting. A strong increase of the level of the phosphorylated form of the three MAPKs was observed in CuO NPtreated cells compared to control cells at all incubation times (Fig. 8). In addition, the CuO NPs-induced phosphorylation of MAPKs decreased after 24 h of incubation suggesting a transient activation of MAPKs. Interestingly, a greater amount of phosphorylated form of the three MAPKs was observed after incubation with rod-shaped CuO NPs in comparison with spherical CuO NPs at each time of incubation.
2.9 CuO NPs-activated MAPKs regulate differently Nrf2, AP1 and NF-kB transcription factors Potential regulation of CuO NPs-activated transcription factors by MAPKs was examined using specific inhibitors for p38MAPK (SB203580), ERKs (U0126) and JNK/SAPK (SP600125). Effects of these inhibitors on the transcription factors DNA-binding activities are described in Fig. 9. None of the inhibitors reduced the DNA binding of Nrf2, suggesting that MAPKs pathways were not involved in the activation of this transcription factor in CuO NPs-treated HepG2 cells (Fig. 9A). In order to determine the mechanism by which Nrf2 could be activated following the exposure to CuO NPs, the effect of calphostin C (PKC inhibitor) or antioxidant NAC was studied on the nuclear translocation of Nrf2. While inhibition of PKC had no effect (ESI, Fig. S4†), preincubation with NAC reduced the Nrf2 translocation into the nucleus induced by both CuO NPs and positive control Sin-1 (Fig. 9D), confirming the implication of oxidative stress in the activation of Nrf2. DNA binding of NF-kB was not modified following the inhibition of p38MAPK. Inhibition of ERKs with U0126 and JNK/SAPK by SP600125 had only an effect on the DNA binding of NF-kB induced by spherical CuO NPs
Fig. 8 Effects of CuO NPs on MAPKs phosphorylation. Relative amounts of total and phosphorylated forms of p38MAPK, ERKs and JNK/SAPK after increasing the incubation time with 25 mg ml1 of rodshaped (R. CuO) or spherical (S. CuO) CuO NPs were assayed by western blotting. b-Actin was used as a reference protein.
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(Fig. 9B). Fig. 9C shows that inhibition of p38MAPK did not modify the DNA binding of AP-1. As expected, inhibition of JNK/SAPK reduced the DNA binding of AP-1 induced by both CuO NPs. Interestingly, U0126 decreased AP-1 DNA binding induced by both rod-shaped and spherical CuO NPs. Regulation of AP-1 activity by ERKs and JNK/SAPK was further confirmed by evaluating the impact of U0126 and SP600125 inhibitors on the phosphorylated status of c-Jun. Both inhibitors reduced the phosphorylation of c-Jun induced by CuO NPs (ESI, Fig. S5†).
3. Discussion In this study, we evaluated the toxicity of two types of CuO NPs characterized by similar chemical composition and size distribution, but different specific surface areas and shapes on the HepG2 hepatoma cell line. Both rod-shaped and spherical CuO NPs decreased cell viability dose-dependently. Dose-dependent cytotoxicity of CuO NPs was also observed in several other cell lines such as human alveolar A549 cells or human intestinal Caco-2 cells, even if the magnitude of CuO NPs cytotoxicity depended on the cell type.10 In order to investigate whether CuO NPs cytotoxicity could result from cellular penetration, intracellular CuO NPs were monitored by NanoSIMS50. The NanoSIMS technique is well suited for analysing the distribution of trace elements at high resolution because of its unique combination of high sensitivity (ppm), high lateral resolution (down to 50 nm) and detectability of all elements (from hydrogen to uranium) and isotopes.39 Previously, NanoSIMS50 has been used successfully in ecotoxicology for analysing the uptake of environmental xenobiotics into living organisms with high resolution.40,41 In HepG2 cells, NanoSIMS50 analysis revealed the cytoplasmic presence of Cu after 2 h of incubation in the presence of CuO NPs. Analysis of 31P distribution in HepG2 cells suggests no intranucleus localization of Cu in all the images analyzed. Interestingly, an intracellular overlapping between 63 Cu and 32S signals was observed, from 6 h to 24 h of incubation. This suggests a co-localization of copper with a sulfur-rich compound in the same area. One speculative hypothesis to explain this observation could be that CuO NPs, once inside the cells, are direct oxidants or they generate ROS which oxidize cytoplasmic sulphur-rich proteins. This could favour the formation of protein disulphur bonds and/or bonds42 between proteins and NPs, leading to the formation of intracellular protein–NPs complexes as suggested previously.43 Expression of metallothionein (MT) has recently been described to increase after exposure to CuO NPs.44,45 This cysteine-rich protein is able to bind heavy metals through the thiol group of its cysteine residues (representing nearly 30% of its amino acid composition).46 MT might co-localize with CuO NPs due to its capability to bind Cu and to act as a scavenger of oxygen species,47 participating in an antioxidant response to protect cells against oxidative stress induced by NPs.44,45,48 Indeed, CuO NPs were shown to generate oxidative stress in HepG2 cells as assayed by the 2,7-DCF probe oxidation measurement. An increase in ROS production was measured between 4 and 6 h of incubation, indicating that oxidative stress appeared after intracellular penetration of CuO NPs. Overexpression of oxidative stressrelated HMOX-1 protein was also observed after 4 h of exposure Nanoscale, 2012, 4, 7168–7184 | 7177
Fig. 9 Effects of MAPK inhibitors on DNA binding or nuclear translocation of redox-sensitive transcription factors. Analysis of Nrf2 (A), NF-kB (B) and AP-1 (C) DNA binding to their respective consensus sequence using a colorimetric assay. HepG2 cells were preincubated for 1 h with specific inhibitors of ERKs (U0126, 5 mM), JNK/SAPK (SP600125, 10 mM) or p38MAPK (SB203580, 20 mM) before exposure for 6 h in the presence or absence (control, CTL) of rod-shaped (R. CuO) or spherical (S. CuO) CuO NPs (25 mg ml1) with or without MAPK inhibitors. Sin-1 (2.5 mM, 6h), IL-1b (0.01 ng ml1, 30 min) or PMA (0.1 ng ml1) + ionomycin (1 mM, 6 h) were used as respective positive controls. Results in optical density (O.D.) are expressed as means s.d. for n ¼ 3 and normalized to control. *: P < 0.05, **: P < 0.01 or ***: P < 0.001 vs. control. #P < 0.05 or ##: P < 0.01 vs. the corresponding condition without inhibitor. (D) Nuclear translocation of Nrf2 (green) was studied by immunofluorescence after 6 h of incubation with CuO NPs in the presence or absence of NAC (10 mM).
to CuO NPs. These results confirm the oxidant potential of CuO NPs previously described in other cell lines such as Caco-2, A549, HEp-2, or human cardiac microvascular endothelial cells.5,7,9,10 Moreover, antioxidant NAC reduced cell death drastically and the amount of extracellularly released IL-8, indicating that oxidative stress plays an important role in CuO NPs cytotoxicity and pro-inflammatory effects. Oxidative stress can activate transcription factors like Nrf2, NF-kB or AP-1.32 The transcriptomic data indicated that CuO NPs increased the abundance of some transcripts like HMOX1 and IL-8 that were known to be regulated by redox-sensitive transcription factors.34–38 DNA binding activity of Nrf2 increased from 4 h of exposure to CuO NPs, parallel to ROS overproduction and accumulation of HMOX-1 and IL-8 proteins. Implication of Nrf2 in the inflammatory and anti-oxidant responses of HepG2 following the exposure to CuO NPs was confirmed using specific siRNA. Inhibition of Nrf2 expression reduced the abundance of transcripts coding for HMOX1 or IL-8 correlated with the reduction of protein abundance for HMOX1 and IL-8. Interestingly, Nrf2 is also activated upon exposure to titanium dioxide49 or carbon nanotubes,19 suggesting that Nrf2 could play an important role in response to the exposure to NPs. In parallel to the Nrf2 7178 | Nanoscale, 2012, 4, 7168–7184
activation, both NF-kB and AP-1 were shown to bind their consensus DNA sequence between 4 and 6 h of incubation with CuO NPs. Implication of these two transcription factors in the inflammatory and anti-oxidant responses was investigated using siRNA. Inhibition of AP1 and NF-kB reduced the quantity of extracellular released IL-8 indicating that these different transcription factors (Nrf2, AP-1 and NF-kB) could act synergically to regulate the expression of some CuO NPs-induced genes. A coordinate regulation of IL-8 transcription by AP-1 and NF-kB in response to different external stimuli was previously described.50 Interestingly, AP-1 was shown to be implicated in the response of A549 cells to the toxicity of CuO NPs,51 suggesting a major role of this transcription factor in cell response to exposure to CuO NPs. All together, these data indicate that several transcription factors are activated after CuO NPs incubation in HepG2 cells. Implication of MAPKs in the activation of downstream transcription factors was evaluated using specific inhibitors. Indeed, several studies have described the involvement of MAPKs in cell responses to exposure to NPs. For instance, CeO2 or SiO2 NPs-activation of Nrf2 has been described to depend on p38MAPK or ERKs signalling pathways respectively, even if no This journal is ª The Royal Society of Chemistry 2012
direct link was demonstrated between the MAPKs and the transcription factor.17,18 In addition, CuO NPs were shown to activate p38MAPK in endothelial cells due to oxidative stress.52 Here, we showed that CuO NPs transiently activated p38MAPK, ERKs and JNK/SAPK MAPKs. MAPKs can be implicated in the activation of Nrf2.53,54 The use of specific inhibitors revealed no implication of MAPKs in the regulation of Nrf2 DNA binding activity in CuO NPs-exposed HepG2 cells. Interestingly, phosphorylation of MAPKs decreased after 24 h of incubation with CuO NPs while activation of Nrf2 and oxidative stress was still observed. However, antioxidant NAC reduced nuclear translocation of Nrf2 suggesting that CuO NPs-induced oxidative stress could directly activate Nrf2. Indeed, oxidation of Keap1 thiol groups by ROS leads to the release of Nrf2 and its activation.15,16 In parallel, JNK/SAPK and ERKs were shown to activate AP-1 (and to a lesser extent NF-kB) after CuO NPs incubation. However, inhibition of p38MAPK did not modify the DNA binding of the CuO NPs-activated transcription factors. As shown by Hanagata et al., p38MAPK could be part of the cell survival process activated by CuO NPs.51 Interestingly, we observed a decrease of phosphorylated p38MAPK after 24 h of incubation. This decrease could be explained by the incapacity of the cell to face the stress triggered by CuO NPs after a long period of exposure, finally leading to cell death as observed after 24 h of incubation. Further investigation will be required to determine the exact mechanism and identify upstream factors leading to the activation of MAPKs after exposure to CuO NPs. Comparison of rod-shaped and spherical CuO NPs revealed a greater cytotoxicity of rod-shaped CuO NPs on HepG2 cells at least at the higher concentrations. A similar conclusion was obtained on Caco-2 cell monolayers incubated with the same CuO NPs.10 Moreover, secretion of IL-8, induction of HMOX1 and activation of MAPKs were generally more important after incubation with rod-shaped CuO NPs than spherical CuO NPs independent of the incubation time. Both CuO NPs have the same chemical composition and similar hydrodynamic diameter once dispersed in culture medium but are characterized by different specific surface areas, different shapes and release different amounts of Cu2+. These three variable parameters may be considered to explain fully the differential toxicity observed between the two CuO NPs. The specific surface area was predicted to directly affect toxicity via the reactive atoms and molecules on the NPs surface.55 When measured by BET, the specific surface area of rod-shaped CuO NPs (18.4 m2 g1) is 2.7 fold larger than spherical CuO NPs (6.8 m2 g1). Thus a greater surface contact is expected in the case of rod-shaped CuO NPs. However, the relationship between the specific surface area and cell viability (ESI, Fig. S6†) shows that the specific surface area cannot explain the difference of toxicity between rod-shaped and spherical CuO NPs. Indeed, more cytotoxicity was found at a given specific surface area value for spherical CuO NPs than for rod-shaped CuO NPs, as described previously.10 The relative involvement of the released cations and a specific NP effect in toxicity is still debated in the literature. For instance, several studies have shown that free Cd2+ cations were responsible for the cytotoxicity of CdTe quantum dots (QDs)56,57 while others showed that their cytotoxicity cannot depend solely on the toxic effect of free Cd2+ suggesting that specific properties of NPs This journal is ª The Royal Society of Chemistry 2012
(a specific ‘‘nano effect’’) could have an impact on their toxicity.58,59 In addition, an ecotoxicological study has concluded that the toxicity of CuO NPs was largely due to soluble Cu2+ cations60 while an in vitro study showed that CuO NPs toxicity was not due to the release of Cu2+ cations in the cell culture medium.5 Potential implication of Cu2+ cations in the CuO NPs cytotoxicity on HepG2 cells was investigated. It must be considered that ROS might be formed firstly by the Cu2+ cations released in the culture medium, generating thereby extracellular oxidative stress with potential deleterious effects. At all concentrations tested, more Cu2+ cations were released in cell culture medium by CuCl2 than CuO NPs but the cytotoxic effect of CuO NPs was always more important. Cu2+ cations from CuCl2 only decreased cell viability moderately at the highest concentration while the effect of CuO NPs was more pronounced suggesting that Cu2+ cations released from CuO NPs could only participate in part to the toxic effect of CuO NPs as previously described.5,8,10 Secondly, upon internalization, CuO NPs could deliver Cu2+ cations and exert cytotoxic effects through intracellular ROS production as suggested by Hanagata et al.51 Su et al. have observed a greater toxicity of CdTe QDs in comparison with CdCl2 while intracellular Cd2+ concentration was identical in HEK293 cells treated with either QDs or CdCl2. These authors observed the intracellular accumulation of QDs in perinucleus areas where QDs could release locally Cd2+ ions leading to a ‘‘concentration effect’’. Cd2+ ions, concentrated in these small areas, might then diffuse into the nuclei and damage cells.59 We also observed accumulation of CuO NPs in localized intracellular areas. Differential intracellular release of Cu ions by internalized CuO NPs could explain in part the different toxicity observed between the two CuO NPs, the rod-shaped CuO NPs releasing a higher quantity of Cu2+. More and more convincing data reveal the importance of the NPs shape in toxicity, cell internalization, blood flow and drug delivery.61–63 Here, we studied two CuO NPs characterized by different shapes – rod or spherical. Interestingly, CuO NPs with rod shape induced a greater cytotoxicity than spherical CuO NPs suggesting that shape could be implicated in the CuO NP toxicity as suggested previously.10 Shape could have an impact on the NPs cytotoxicity, for example, by modulating cellular dose, i.e. the ‘‘real’’ delivered dose of nanoparticles to cells.64 Indeed, the NPs shape could modify the NPs settling, diffusion and aggregation state, significantly affecting the final dose in contact with the cells or internalized. In addition, several authors described that NPs with larger aspect ratio (rod shaped) are taken up in greater amounts and at faster internalization rates.63,65 In this study, both CuO NPs were detected inside HepG2 cells independent of the incubation time even if we cannot exclude a difference in the internalization rates of the two NPs at shorter incubation times or in the amount of intracellular CuO NPs, the NanoSIMS technique being not straightforward in quantitative measurements. Indeed, the correlation of secondary ion intensities with the amount of matter in the sputter volume is not perfect and requires a complex approach (standards, curve of calibration, .).66 A faster internalization or a greater amount of internalized NPs would trigger a quicker production of ROS and activation of signaling pathways leading to a more rapid production of a greater amount of HMOX-1 and IL-8 induced by rod-shaped CuO NPs. Nanoscale, 2012, 4, 7168–7184 | 7179
4. Conclusion In conclusion, even if the mechanism(s) responsible for differential toxicity between CuO NPs remains unclear, both CuO NPs were shown to be able to enter into HepG2 cells and exert their cytotoxicity through oxidative stress. Activation of MAPKs and redox-sensitive transcription factors was demonstrated suggesting that MAPK pathways and redox-sensitive transcription factors could be major factors in CuO NPs toxicity. All together, these data allow a better comprehension of the molecular mechanism and signalling pathways induced by CuO NPs in HepG2 cells.
NPs (i.e., 5–100 mg ml1 CuO NPs) or CuCl2 (i.e., 11–215 mg ml1 CuCl2). To evaluate the potential implication of MAPKs in the cell response to CuO NP treatment, HepG2 cells were preincubated for 1 h in the presence of specific kinase inhibitors: SB203580 (an inhibitor of p38MAPK; 20 mM; Alexis), U0126 (an inhibitor of MEK1/2, upstream MAPK kinases activating ERKs; 5 mM; cell signalling) or SP600125 (an inhibitor of JNK/SAPK 1/2, 10 mM; Tocris bioscience) before co-treatment for 6 h in the presence or absence of CuO NPs. Under some conditions, HepG2 cells were preincubated for 1 h with 10 mM of N-acetyl-cystein (NAC, Sigma) or calphostin C (200 nM, Calbiochem) before a 6 or 24 h co-incubation with or without CuO NPs.
5. Experimental 5.1 Material characterization
5.4 2,7-DCF oxidation assay
Two categories of CuO NPs were tested. Cu(II) oxide NPs powder 12 nm and 50–80 nm were obtained from IoLiTec. CuO NPs size distribution and morphology were analyzed respectively by centrifugal liquid sedimentation (Disc Centrifuge DC24000, CPS) at 24 000 rpm and transmission electron microscopy (TEM) at 80 keV (FEI Tecnai 10, Philips) using TEM grids (Agar) covered with non-porous formvar. The specific surface area was determined by the Brunauer–Emmett–Teller (BET) method (ASAP 2010, Micrometrics). The chemical composition was analyzed by energy dispersive X-ray (EDX) spectrometry at 20 keV (a JED 2300 detector coupled to a field emission gun scanning electron microscope JSM 7500F, ZAF corrections, standardless analysis, Jeol). X-ray photoelectron spectroscopy (XPS) was used for extreme surface analysis. XPS spectra were recorded at a 35 take-off angle with an SSX-100 spectrometer using the monochromatized X-ray Al Ka radiation, 1486.6 eV and treated with the CasaXPS version 2.3.10 Dev 8 software.
After CuO NP incubation, cells were incubated for 15 min with 10 mM 2,7-dichlorofluorescein (2,7-DCF, Eastman Kodak) diluted in PBS (Phosphate Buffer Saline, pH 7.4). H2O2 (500 mM, Merck) was added in PBS for 15 min and used as positive control of oxidative stress. Cells were rinsed with PBS. Fluorescence intensities (485 nm excitation; 520 nm emission) were recorded with a fluorimeter (Fluoroskan Ascent, Thermo Scientific). Protein concentrations were assayed on cell lysates with the Pierce 660 nm protein assay reagent (Thermo Scientific) to normalize the fluorescence intensities to cellular protein content. Results are expressed as relative fluorescence unit (RFU) per mg of proteins and normalized to the control.
5.2 CuO NP suspensions Stock solutions of CuO NPs (1 mg ml1) were prepared in water by stirring CuO NPs for 90 min. To achieve the different concentrations used in this study, the stock CuO NPs suspensions were diluted in DMEM cell culture medium plus fetal bovine serum before cell incubations.
5.5 MTS reduction assay The conversion of MTS tetrazolium salt (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2Htetrazolium) into its reduced formazan form was assessed with the CellTiter 96 AQueous Non-Radioactive Cell Proliferation Assay kit (Promega) according to the manufacturer’s instructions. Briefly, HepG2 cells were incubated with MTS solution for 3 h at 37 C before measuring 490 nm absorbance with a spectrophotometer (xMark, Biorad). Results are expressed as percentages of controls.
5.3 Cell culture and incubation
5.6 Nrf2, p65 and c-Jun siRNAs transfection
Human HepG2 cells used at passages X + 1 to X + 20 were maintained in culture in 75 cm2 polystyrene flasks (Corning) with 15 ml of Dulbecco’s modified Eagle’s medium (DMEM) liquid (Gibco) containing 1% penicillin–streptomycin (BioWhittaker) and 10% fetal bovine serum (Gibco) and incubated at 37 C under an atmosphere of 5% CO2. 50 000 or 400 000 cells were seeded in 24 well cell culture plates (Corning) for cytotoxicity assays (MTS assay, see below) or for 2,7-DCF oxidation assays, respectively. The next day, cells were incubated as indicated (between 2 h and 24 h) with 500 ml per well of culture medium containing increasing concentrations of CuO NPs (5 to 100 mg ml1). For a comparison of cytotoxicity between CuO NPs and Cu ions, HepG2 cells were exposed to concentrations between 4 mg ml1 and 80 mg ml1 Cu of CuO
Knockdown of Nrf2, p65 and c-Jun expression was achieved using ON-TARGET plus SMART pool human NFE2L2 (L-003755-00, Dharmacon), ON-TARGET plus SMART pool human p65 (L-0035-33-00, Dharmacon) and ON-TARGET plus SMART pool human c-Jun (L-003268-00, Dharmacon) respectively. ON-TARGET plus Non-Targeting SMART pool (D-001810-10, Dharmacon) was used as a control for transfection of non-specific effects. 106 HepG2 cells seeded in 25 cm2 flasks (Corning) were transfected during 24 h under standard culture conditions with 10 nM (Nrf2 and p65) or 50 nM (c-Jun) siRNA using the DharmaFECT 1 transfection reagent (Dharmacon dilution 1 : 500) according to the manufacturer’s instructions. After 3 h in fresh medium, cells were then incubated with or without CuO NPs for 24 h.
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5.7 qPCR and TLDA analyses
5.9 Sample preparation and NanoSIMS analysis
Total RNA was extracted using the ‘‘TRI Reagent Soln’’ (Ambion). For real-time syber green quantitative PCR (qPCR) analysis in 96-well plates, mRNA contained in 1 mg of total RNA was reverse transcribed (RT) using the ‘‘Transcription First Strand cDNA Synthesis’’ kit (Roche) and the supplied materials. Forward and reverse primers for human heme oxygenase-1 (HMOX1), tumor growth factor-b1 (TGF-b1), interleukine-8 (IL-8) and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) were designed using the Primer Express 1.5 software (Applied Biosystems) as follows: HMOX1-forward, 50 -CCA GCA ACA AAG TGC AAG ATT-30 , HMOX1- reverse, 50 -CAC ATG GCA TAA AGC CCT ACA-30 ; TGF-b1forward, 50 -AGG GCT ACC ATG CCA ACT TCT-30 ; TGFb1-reverse, 50 -CCG GGT TAT GCT GGT TGT ACA-30 ; IL-8forward, 50 -CTG GCC GTG GCT CTC TTG-30 , IL-8-reverse, 50 -GGG TGG AAA GGT TTG GAG TAT G-30 ; GAPDHforward, 50 -ACC CAC TCC TCC ACC TTT GAC-30 , GAPDH-reverse, 50 -GTC CAC CAC CCT GTT GCT GTA-30 . Amplification reaction assays contained primer pairs at 300 nM or 900 nM and SYBR Green PCR Mastermix (both from Applied Biosystems). A hot start at 95 C for 5 min was followed by 40 cycles at 95 C for 15 s and 65 C for 1 min using an ABI PRISM 7000 SDS thermal cycler (Applied Biosystems). GAPDH was used as the reference gene for normalization and relative mRNA steady-state level quantification based on cycle thresholds (Ct). Melting curves were generated after amplification, and data were analyzed using the thermal cycler SDS 2.2.2 software by the 2DDCt method.67 For real time Taqman PCR low density arrays (TLDA) analysis, mRNA contained in 1 mg of total RNA was reverse transcribed using the ‘‘High-Capacity cDNA Reverse Transcription’’ kit (Applied Biosystems). Then multiplex qPCR amplifications were performed with an ABI PRISM 7000 SDS thermal cycler, using 384-well ‘‘microfluidic card’’ plates (Human Immune Array, Applied Biosystems) containing various primer pairs for parallel analyses of 96 transcripts on 4 samples. GAPDH mRNA was used as a reference for normalization and relative mRNA steady-state level quantification based on cycle thresholds (Ct). Melting curves were generated after amplification, and data were analyzed using the thermal cycler SDS 2.2.2 software by the 2DDCt method.67 Control (CTL) mean values were calculated from triplicate Ct values for each mRNA. Each NP-treated triplicate value was then calculated by the 2DDCt method in comparison with the CTL mean value and s.d. were computed. When calculable, statistical significance was considered for P < 0.05 using Student’s t-tests.
2 106 cells were seeded in 25 cm2 flasks (Corning) and incubated for increasing times with or without CuO NPs (25 mg ml1). Cells were harvested by centrifugation after trypsinisation in pyramidal beem capsule (Agar Scientific). Cell pellets were fixed for 150 min at 4 C in 2.5% (w/v) glutaraldehyde (Agar Scientific) in 0.1 M cacodylate buffer (pH 7.4). Cells were then washed, post-fixed in 1% (v/w) osmium tetroxide (Merck) in 0.1 M cacodylate buffer (pH 7.4) for 60 min at 4 C, dehydrated in a graded series of ethanol solutions (30, 50, 70, 85 and 100%), infiltrated and embedded in epon resin LX 112 (LADD Research Industries). The samples were cut into 200 nm semi-thin sections with an ultramicrotome (Leica) and placed on silicon wafers (Siltronix) for NanoSIMS analysis. SIMS experiments were performed using the Cameca NanoSIMS50 (Gennevilliers, France) in the raster imaging mode. The ionic images were obtained using a beam of Cs+ primary ions with an energy impact of 16 keV and a current of around pA on the sample. The images were recorded in 256 256 pixels. Under these analytical conditions, a lateral resolution of 100–80 nm is expected. Negative secondary ion signals for 12C14N, 31P, 34 63 S , Cu and 65Cu were collected simultaneously, thanks to the multi-collection system.69 The images were acquired with a counting time of 30 ms per pixel. Mass resolution (M/DM) was above 4000 to resolve mass interference with polyatomic ions such as between 63Cu (m ¼ 62.9296 atomic mass unit – amu) and 31 16 P O2 (m ¼ 629 636 amu). Both the isotopes of copper, 63Cu and 65Cu (68.98% and 31.05% of the natural abundance, respectively), were recorded and the mass calibration was achieved using standard references (copper foil). The ratio of the intensities of the 63Cu and 65Cu (ratio ¼ 2.22) was systematically checked.
5.8 Interleukin-8 and colony stimulating factor-1 assays Culture media collected after exposure to CuO NPs were centrifuged for 5 min at 13 000 rpm before assaying the concentration of IL-8 or CSF-1 (pg ml1) in the supernatants by specific sandwich ELISA (Quantikine human IL-8/CSF-1 Immunoassay, R&D Systems) according to the manufacturer’s protocol. Interleukin-1b (IL-1b, 0.01 ng ml1 R&D System), a well-known inducer of IL-8, was used as a positive control.68 This journal is ª The Royal Society of Chemistry 2012
5.10 Cu cation assay Cu cations released in cell culture medium were assayed in triplicate with Quantichrom copper assay (Gentaur). This method uses a chromogen that forms a colored complex specifically with bivalent copper ions. After 24 h of incubation at 37 C in the presence of CuO NPs or CuCl2, DMEM medium containing fetal bovine serum was centrifuged for 30 min at 13 000 rpm. The supernatant was collected, diluted 50 in distilled water and assayed as recommended by the manufacturer. Briefly, 35 ml of trichloroacetic acid was mixed with 100 ml of each sample and centrifuged for 2 min at 13 000 rpm. 150 ml of working solution (ascorbic acid, 4,40 -dicarboxy-2,20 -biquinoline, sodium hydroxide and hepes) was mixed with 100 ml of sample and incubated 5 min at room temperature. The amount of Cu++ was assayed by measuring of absorbance at 359 nm. Results were normalized in comparison to the control and expressed as fold increase. 5.11 Nuclear protein extraction and DNA-binding assay HepG2 cells seeded in 25 cm2 flasks (Corning) at a density of 106 cells per flask were incubated with NPs for 2, 4, 6 or 24 h. After incubation, flasks were placed on ice. Cells were rinsed with PBS containing 1 mM Na2MoO4 and 5 mM NaF, incubated for 3 min Nanoscale, 2012, 4, 7168–7184 | 7181
with cold Hypotonic Buffer (HB, 20 mM HEPES, 5 mM NaF, 1 mM Na2MoO4, 0.1 mM EDTA) and harvested in 200 ml HB containing 0.5% NP-40. Lysats were centrifuged for 30 s at 13 000 rpm. Sedimented nuclei were resuspended in 30 ml RE completed buffer: HB containing 20% glycerol, a protease inhibitor cocktail (Roche Molecular Biochemicals, 1 : 25 dilution) and phosphatase inhibitors (1 mM Na3VO4, 5 mM NaF, 10 mM p-nitrophenylphosphate, 10 mM b-glycerophosphate) added at a 1 : 25 dilution. Extraction was performed for 30 min at 4 C with gentle agitation by the addition of 30 ml SA completed buffer (HB containing 20% glycerol, 0.8 M NaCl and protease/phosphatase inhibitors). Nuclear extracts were obtained by centrifugation for 10 min at 13 000 rpm. The protein concentration of nuclear extracts was quantified by the Pierce method (Thermo Scientific). Nuclear extracts of HepG2 cells incubated for 30 min in the presence of IL-1b (0.01 ng ml1) or 6 h in the presence of 3-morpholinosydnonimine hydrochloride (Sin-1, 2.5 mM; Invitrogen), or a mix of phorbol-12-myristate13-acetate (PMA, 0.1 ng ml1, Sigma) and ionomycin (1 mM, Sigma) were used as positive controls for investigation of DNA binding of NF-kB, Nrf2 or AP-1 respectively. Nuclear extracts from Jurkat cells incubated for 6 h in the presence of phorbol-12myristate-13-acetate (PMA, 0.1 ng ml1) and ionomycin (1 mM) were also used as internal positive controls for DNA binding activity of AP-1. DNA-binding assays using a TransAM ELISA kit (Active Motif) for assaying NF-kB, Nrf2, HIF-1a, p53, and AP-1 DNAbinding activities were performed according to the manufacturer’s recommendations. Briefly, nuclear proteins (10 mg for NFkB and Nrf2 and 5 mg for HIF-1a, p53 and AP-1) were incubated for 60 min under agitation at room temperature in a 96-well plate coated with a double-stranded oligonucleotide containing a specific consensus sequence (4 pmol per well). The transcription factor bound to DNA was detected using a specific primary antibody: NF-kB p65 sc-372 (Santa Cruz), Nrf2 sc-13032 (Santa Cruz), HIF-1a BD 610958 (BD Biosciences), p53 sc-6243 (Santa Cruz) and AP-1 phospho-c-Jun sc-822 (Santa Cruz). Colorimetric reaction was performed with a horseradish peroxidase (HRP)-conjugated IgG antibody (anti-rabbit sc-2004 for NF-kB, Nrf2, p53 (Santa Cruz) and anti-mouse sc-2005 for HIF-1a and AP-1 (Santa Cruz)). After addition of developing solution for 10 min and stopping the colorimetric reaction, absorbance was determined using a spectrophotometer at 450 nm with a reference wavelength of 655 nm.
Proteins separation (15 mg to 25 mg proteins per well) was performed by SDS-PAGE on 12% polyacrylamide gel. Separated proteins were then transferred to an Immobilon-FL membrane (Millipore) for 2 h at 1 mA cm2. The membrane was then blocked using a blocking agent purchased from Odyssey (Licor, dilution 1 : 2 in PBS) for 2 h at room temperature or overnight at 4 C, followed by an incubation for 1 h at room temperature or overnight at 4 C with the primary antibody in blocking buffer containing 0.1% Tween (Sigma). After 4 washes of 5 min in PBS– Tween 0.1%, the incubation with the secondary antibody (IR Dye Licor) was performed for 1 h at room temperature in blocking buffer Tween 0.1%, followed by 4 washes of 5 min in PBS–Tween 0.1% and 2 washes of 5 min in PBS. Protein detection was performed using the Odyssey Infrared Imaging System (Licor). For the revelation of phospho-c-Jun and native or phosphorylated forms of JNK/SAPK, the super signal western blot enhancer (#46641 Thermoscientific) was used according to the manufacturer’s recommendations. Primary and secondary antibodies are listed in Table S2 (ESI†). a-Tubulin and b-actin western blotting was performed to assess for the total amount of proteins loaded on the gel. 5.13 Immunofluorescence Cells were seeded on glass cover slides 1 day before the incubation (50 000 cells per cover slide). After 6 or 24 h of incubation, cells were fixed 10 min with 4% paraformaldehyde in PBS, washed 3 times with PBS, permeabilized in PBS + 1% Triton X-100 (Merck) for 5 min and washed 3 10 min in PBS + 3% BSA (PAA). The primary antibody (anti-HMOX-1 ab52947 from Abcam, 1 : 500 dilution or anti-Nrf2 sc-13032 from Santa Cruz, 1 : 100 dilution) was added in PBS + 3% BSA overnight at 4 C in a wet room. The next day, cells were washed 3 times in PBS + 3% BSA before incubation for 1 h with the secondary antibody (Alexa Fluor 488 goat anti-rabbit IgG (H + L) conjugate (Molecular Probes); at 1 : 1000 dilution) in PBS + 3% BSA; room temperature. Cells were washed 3 times in PBS + 3% BSA. To visualize the nucleus, cells were incubated for 25 min at room temperature in the presence of TO-PRO-3 (1 : 80 dilution in PBS + RNase 2 mg ml1; Molecular Probes). The coverslips were mounted in Mowiol (Sigma) and observed using a confocal microscope TCS (Leica) with a constant photomultiplier. Sin-1 was used as a positive control to monitor the nuclear translocation of Nrf2.70,71 5.14 Statistical analysis
5.12 Western blotting 2
6
HepG2 cells seeded in 25 cm flasks (Corning) at a density of 10 cells per flask were incubated with CuO NPs for 2, 4, 6 or 24 h. After incubation, cells were placed on ice and scrapped in 200 ml of lysis buffer (Tris 40 mM pH 7.5, KCl 150 mM, EDTA 1 mM, 1% triton X-100) containing a protease inhibitor cocktail (Roche Molecular Biochemicals, added at a 1 : 25 dilution) and phosphatase inhibitors (1 mM Na3VO4, 5 mM NaF, 10 mM p-nitrophenylphosphate, 10 mM b-glycerophosphate; 1 : 25 dilution). Lysates were centrifuged 5 min at 13 000 rpm at 4 C and supernatants were harvested. The protein concentration was evaluated by the Pierce assay (Thermo Scientific). 7182 | Nanoscale, 2012, 4, 7168–7184
Results are expressed as mean standard deviation (s.d.). Data were analyzed by Student’s t-tests.
6. Declaration of interest The authors would like to report no conflict of interests.
Acknowledgements This work was supported by the DGO6 (Direction Generale Operationnelle de l’Economie, de l’Emploi et de la Recherche) of the Walloon Region of Belgium (‘‘Nanotoxico’’ Pole of Excellence, RW/FUNDP research convention no. 516252, and This journal is ª The Royal Society of Chemistry 2012
Silicalloy project, RW/FUNDP research convention no. 6144). The research leading to these results has received funding from the European Commission Seventh Framework Programme (FP7/2007-2013) under grant agreement no. 262163 (‘‘QNano’’ INFRASTRUCTURE) and no. 263147 (Large Scale Integrated Project ‘‘Nanovalid’’). O. Toussaint is a Senior Research Associate of the Belgian FNRS. E. Boilan was a fellow of the Belgian FRIA.
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