Effect of temperature on the release of intentionally and non ...

Food Chemistry 139 (2013) 672–680

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Effect of temperature on the release of intentionally and non-intentionally added substances from polyethylene terephthalate (PET) bottles into water: Chemical analysis and potential toxicity Cristina Bach a,c,⇑, Xavier Dauchy a, Isabelle Severin b, Jean-François Munoz a, Serge Etienne c, Marie-Christine Chagnon b a

ANSES, Nancy Laboratory for Hydrology, Water Chemistry Department, 40 rue Lionnois, 54000 Nancy, France Derttech ‘‘Packtox’’, Nutox team, Inserm U866, AgroSupDijon Nord, 1 Esplanade Erasme, 21000 Dijon, France c Institute Jean Lamour, UMR 7198, Department SI2M, Ecole des Mines de Nancy, University of Lorraine, Parc de Saurupt, CS 14234, 54042 Nancy, France b

a r t i c l e

i n f o

Article history: Received 21 August 2012 Received in revised form 30 November 2012 Accepted 15 January 2013 Available online 29 January 2013 Keywords: PET-bottled water By-products Chemical mixtures Cyto-genotoxicity Endocrine disruption

a b s t r a c t The purpose of this study was to investigate the impact of temperature on the release of PET-bottle constituents into water and to assess the potential health hazard using in vitro bioassays with bacteria and human cell lines. Aldehydes, trace metals and other compounds found in plastic packaging were analysed in PET-bottled water stored at different temperatures: 40, 50, and 60 °C. In this study, temperature and the presence of CO2 increased the release of formaldehyde, acetaldehyde and antimony (Sb). In parallel, genotoxicity assays (Ames and micronucleus assays) and transcriptional-reporter gene assays for estrogenic and anti-androgenic activity were performed on bottled water extracts at relevant consumer exposure levels. As expected, and in accordance with the chemical formulations specified for PET bottles, neither phthalates nor UV stabilisers were present in the water extracts. However, 2,4-di-tert-butylphenol, a degradation compound of phenolic antioxidants, was detected. In addition, an intermediary monomer, bis(2-hydroxyethyl)terephthalate, was found but only in PET-bottled waters. None of the compounds are on the positive list of EU Regulation No. 10/2011. However, the PET-bottled water extracts did not induce any cytotoxic, genotoxic or endocrine-disruption activity in the bioassays after exposure. Ó 2013 Elsevier Ltd. All rights reserved.

1. Introduction Today, the most common polymer used for the bottling of drinking water is polyethylene terephthalate (PET). Since migration can occur between packaging and foodstuffs, consumers may be exposed to the potentially harmful chemicals (additives, un-reacted monomers, and processing aids) used in manufacturing the packaging. These intentionally-added substances (IAS) are listed and controlled by European Regulation No. 10/2011 Abbreviations: BHT, butylated hydroxytoluene; BHET, bis(2-hydroxyethyl) phthalate; DBP, dibutyl phthalate; DEHA, di-2-ethylhexyl adipate; DEHP, di-2ethylhexyl phthalate; DEP, diethyl phthalate; DHT, dihydrotestoterone; DiBP, diisobutyl phthalate; DMP, dimethyl phthalate; DMSO, dimethyl sulfoxide; 2,4-dtBP, 2,4-di-tert-butylphenol; ECCVAM, European Center for the Validation of Alternative Methods; EFSA, European Food Safety Authority; IAS, intentionally added substances; LOQ, limit of quantification; NIAS, non-intentionally added substances; SML, specific migration limit. ⇑ Corresponding author at: ANSES, Nancy Laboratory for Hydrology, Water Chemistry Department, 40 rue Lionnois, 54000 Nancy, France. Tel.: +33 (0)3 83 38 87 29; fax: +33 (0)3 83 38 87 21. E-mail address: [email protected] (C. Bach). 0308-8146/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.foodchem.2013.01.046

and do not pose any risk to humans. However, over 50% of compounds migrating from food contact materials are nonintentionally added substances (NIAS) (Grob, Biedermann, Scherbaum, Roth, & Rieger, 2006). Indeed, Mittag and Simat (2007) reported that 98% of the toxicity evidenced by several epoxy coating migrates was due to NIAS and/or reaction products. European Regulation No. 10/2011 concerning plastics and multilayers recently became more strict, stating that ‘‘the risk assessment of a substance should cover the substance itself, relevant impurities and foreseeable reaction and degradation products in the intended use’’ (EU, 2011). PET is characterised by a limited range of additives and low diffusion of potential migrants in the polymer matrix (EFSA, 2011b). However, in PET-bottled waters non-polymer origins of NIAS also exist, namely the water itself, the bottling process, disinfection agents and environmental pollutions. Furthermore, PET can be degraded due to several exposure factors under normal conditions of use (heat and UV light). In addition, certain physicochemical properties of bottled water, such as inorganic composition, carbonation or bacterial presence, influence

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the leaching of constituents from PET bottles into water. It has also been established that the migration of several compounds (formaldehyde, acetaldehyde and Sb) from PET packaging to water is a thermally activated process (see review in Bach, Dauchy, Chagnon, and Etienne (2012)). However, little or nothing is known about the release of other NIAS (Grob et al., 2006) from the PET bottles into water and the final effect in terms of toxicity of all the migrated substances. Over the last few years, certain studies have reported finding chemical mixtures with cytogenotoxic effects and endocrine disruption activity in PET-bottled water (see review and comments in Bach et al. (2012)). Toxic effects, and especially endocrine disruption, could be attributed to a ‘‘cocktail effect’’ due to compound mixtures (Muncke, 2009). However, migration studies of PET-bottled water rarely combine chemical analysis with toxicological assessments. Therefore when bioassays demonstrate positive responses, analytical data to identify the responsible compounds are always lacking and conclusions are difficult to draw. The current European Regulatory framework states that an individual toxicological evaluation for substances used in the manufacturing of food contact materials is required. However, potential interactions (dose additivity, synergism, supra-additivity, etc.) between compounds may also occur at very low doses. These two points (low doses and interactions) are not often taken into account and represent new paradigms in toxicology. Furthermore, another current challenge is the development of analytical methods able to detect a wide range of analytes present in bottled water at very low levels (see review in Diduch, Polkowska, and Namiesńik (2011)). The aim of this study was to determine the chemical composition of various bottled waters and, in parallel, to perform in vitro bioassays to check the potential toxicity of these waters when exposed to high temperatures. The effect of temperature on the migration of aldehydes, trace metals and several other potential migrants present in plastic packaging was monitored in PET-bottled waters. The migration tests were performed under realistic conditions of human exposure according to the EU Regulation migration criteria (1 kg of water in contact with 6 dm2 of packaging material). The toxicological evaluation of bottled water extracts was carried out using toxicological endpoints of concern at low

concentrations. The bioassays retained in this study were the Ames test (using prokaryotes) and the micronucleus assay using P53 competent human cells (HepG2 cell lines) to assess genotoxicity. Gene reporter assays were also performed for endocrine disruption activities (estrogenic and anti-androgenic) using human HepG2 and MBA-MB453-kb2 cell lines. All the assays were chosen for their performance and feasibility and in accordance with EFSA and/or ICCVAM recommendations (EFSA, 2011a; ICCVAM, 2003). 2. Materials and methods 2.1. Bottled water samples and storage conditions Two French brands of non-carbonated water (brand A) and carbonated water (brand B) bottled in PET and in glass directly purchased from a local store were analysed. Water samples were bottled at the same time and were from identical batches. Three samples were derived from each brand by replacing the commercial water with ultrapure water: non-carbonated water in PET and glass (brand A), ultrapure water in PET (brand A), carbonated water in PET and in glass (brand B), ultrapure water in PET (brand B). Water samples were analysed (i) before the experiments (after 10 days at 20 °C), and (ii) after 10 days of storage at three different temperatures: 40, 50 and 60 °C. 2.2. Solid phase extraction (SPE) Fourteen compounds, presented in Table 1, were extracted using Oasis HLB glass cartridges (6 cc/200 mg, Waters, Milford, USA). These compounds were previously identified in PET-bottled waters using a preliminary GC–MS screening method (see Additional Data section). Prior to SPE extraction, three internal standards were added to water samples as surrogates (Table 1), namely 2,6-di-tert-butyl-d9-4-methylphenol-3,5-d2, benzophenone-d5 and,di-2-ethylhexyl-phthalate-3,4,5,6-d4 (CDN isotopes, Pointe-Claire, Quebec, Canada) at a concentration of 0.4 lg/L. Carbonated water was degassed by ultrasonication. Cartridges were conditioned with 5 mL of ethyl acetate, methanol and UPLC grade water (Biosolve, Valkenswaard, the Netherlands), and

Table 1 Analytical parameters for the 14 compounds related to plastic packaging. Ions monitored limits of quantification (LOQ) and average recoveries and standard deviations (SD) are indicated. Compound

Dimethyl phthalate (DMP) 2,6-Di-tert-butyl-p-benzoquinone 2,6-Di-tert-butyl-d9–4-methylphenol-3,5-d2a Butylated hydroxytoluene (BHT) 2,4-Di-tert-butylphenol (2,4-dtBP) Ethyl-4-ethoxybenzoate Diethyl phthalate (DEP) Benzophenone-d5a Benzophenone 4-Nonylphenol (NP) 3,5-Di-tert-butyl-4-hydroxybenzaldehyde (BHT-CHO) Di-iso-butyl phthalate (DiBP) Dibutyl phthalate (DBP) 2-Ethylhexyl-p-methoxycinnamate Di-2-ethylhexyl adipate (DEHA) Di-2-ethylhexyl-phthalate-3,4,5,6-d4a Di-2-ethylhexyl phthalate (DEHP) a b c d

Ionsb

LOQ

% Recovery (SD)

(m/z)

lg/L

0.1 lg/L (0.3 lg/Lc)

0.5 lg/L (1.6 lg/Ld)

163, 177, 222, 205, 191 194, 149 187, 182, 135, 219, 149 149 178, 129, 153, 149,

0.1 0.1 – 0.1 0.3 0.1 0.1 – 0.1 0.1 0.1 0.1 0.1 0.3 0.1 – 0.1

107 (12) 76 (17) 57 (6) 63 (8) 85 (10) 102 (15) 114 (13) 106 (13) 99 (10) 85 (16) 56 (10) 93 (11) 100 (13) 44 (11) 46 (8) 46 (5) 60 (5)

100 (7) 65 (11) 66 (4) 73 (12) 81 (4) 97 (10) 95 (15) 103 (9) 92 (10) 79 (11) 65 (13) 87 (11) 87 (11) 45 (6) 41 (5) 48 (7) 60 (4)

77 220, 135 240 220, 177 121, 166 110, 82 105, 77 121, 107 191

161 111 171 167

Internal standard. In bold: quantification ions. For 2,4-di-tert-butylphenol and 2-ethylhexylmethoxycinnamate, average recovery was calculated for a spiked level of 0.3 lg/L. For 2,4-di-tert-butylphenol and 2-ethylhexylmethoxycinnamate, average recovery was calculated for a spiked level of 1.6 lg/L.

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sample loading of 1L and elution with 2 mL of ethyl acetate was carried out. Extracts were analysed by GC–MS. In parallel, bottled water samples were extracted for the bioassays following the protocol described above but without internal standards. GC–MS analysis and toxicological tests were then performed on aliquots of ethyl acetate extracts after concentration (factor 500) in order to reproduce realistic consumer exposure in vitro. Preliminary cytotoxicity tests on ethyl acetate alone were performed on human cell lines (HepG2 and MDA-MB453-kb2 cells). Ethyl acetate was not cytotoxic at the final concentration of 1% in the culture medium (data not shown). 2.3. GC–MS analysis Instrumental analyses were performed using gas chromatography coupled with ion trap mass spectrometry (GC–MS) (Varian GC 4400-MS 240). Separation was achieved with an Rxi-5MS column (30 m 0.25 mm ID; 0.25 lm) connected with a 5 m 0.53 mm deactivated pre-column (Restek, Bellefonte, USA) with the following oven programme: 40 °C (hold 1 min) to 280 °C at 8 °C/min and 280 °C (hold 15 min). Helium flow rate was 1 mL/min. Large volume injections (4 lL) in the split mode (1:25) were carried out. The inlet temperature programme was: 40 °C (hold 1 min) to 300 °C at 100 °C/min and 300 °C (hold 15 min). Acquisitions were in Full Scan mode with ion extraction at specific m/z (Table 1). Calibration was performed in the 10–1000 lg/L concentration range, depending on the target compound. Table 1 shows the limits of quantification (LOQ) determined on the basis of a signal-to-noise ratio of 10. However, LOQ for DiBP, DBP and DEHP were calculated in such a way as to never reach 1/3 LOQ of the blanks values. Blanks were prepared with UPLC water (1L) spiked with the three deutered internal standards (Table 1) at 0.4 lg/L following the same protocol used for the samples. The background contributions of laboratory glassware, TeflonÒ connexions, solvents and UPLC water were monitored. For each sample batch, several samples were fortified with target compounds in order to detect a matrix effect due to the mineralisation and/or carbon dioxide in the water. In addition, UPLC water blanks were also prepared for each sample batch. The recovery experiments were determined in UPLC grade water. The recoveries obtained and spiked levels for each compound are presented in Table 1. Mean recoveries ranged from 44% to 114% for the lower spiked level (0.1 and 0.3 lg/L depending on the compound), from 40% to 103% for the higher spiked level (0.5 and 1.6 lg/L depending on the compound). Due to the lower extraction performance, around 40% for 2-ethyl-p-methoxycinnamate, the DEHA and DEHP concentration values obtained in bottled water were corrected using the recovery value obtained through fortification of the corresponding samples. 2.4. Aldehyde analysis in PET-bottled waters Aldehyde analysis (formaldehyde, acetaldehyde, propanal, butanal, crotonaldehyde, butanal, pentanal, hexanal, heptanal, octanal, nonanal, decanal) in water was based on derivatisation with 2,4dinitrophenylhydrazine (2,4-DNPH) prior to SPE and liquid chromatography/diode array detector (HPLC/DAD) analysis. A 40 mg 2,4-DNPH in 20 mL of acetonitrile (AcCN) reagent solution was purified to reduce background formaldehyde contamination in accordance with Zwiener, Glauner, and Frimmel (2002). Derivatisation reaction in water samples (550 mL) was carried out following the EPA 554 standard method (USEPA, 1992) with some modifications: 500lL of 2,4-DNPH reagent were added to samples, and reaction conditions were set up at 60 °C for 4 h. After derivatisation, carbonated water samples were degassed by ultrasonication for 1 h. Aldehydes-DNPH were extracted and concentrated with Oasis HLB cartridges (200 mg adsorbent, 6 cc; Waters,

Milford, MA, USA) using a Gilson’s GX-274 ASPEC™ instrument (Middleton, USA). The cartridges were conditioned successively with 10 mL AcCN (2 5 mL) and, 10 mL of citrate buffer 1 M (2 5 mL). Water samples were sucked through the adsorbent at a flow rate of about 5 mL/min. The elution consisted of 6 mL AcCN (2 3 mL). Extracts were adjusted to 7 mL with ultrapure water. Chromatographic separation was performed using an Agilent 1200 HPLC system equipped with a diode array detector (DAD). A SunFire™ C18 column (250 4.6 mm I.D.; particle size, 5 lm) was used for the separation of DNPH derivates with a flow rate of 1.2 mL/min and detection was performed at a wavelength of 360 nm. The mobile phase was a binary mixture of AcCN (A) and water (B). The gradient program was as follows: isocratic at 60% A for 20 min, 60% A to 90% A for 15 min, isocratic at 90% A for 15 min. Calibration was performed in the 1–10 lg/L concentration range. Quantification limits (LOQ) were defined as the tenfold value of the results obtained with ultrapure water blanks. The LOQ was 3.5 lg/L for formaldehyde, 2 lg/L for acetaldehyde and octanal, 3 lg/L for nonanal and decanal and 1.5 lg/L for the other aldehydes. 2.5. Analysis of trace metals Bottled water samples were analysed using a Series XII inductively coupled plasma mass spectrometer (ICP-MS) (Thermo, Germany) following the ISO 17294-2 standard method (ISO, 2003). The operation conditions were as follows: RF power was 1318 W, the carrier, the auxiliary and the nebuliser argon gas flow were 13.0, 0.88, and 0.69 dm3/min respectively. Rhodium in a concentration of 1 lg/L was used as the internal standard. The LOQ for trace metals was 1 lg/L, except for Sb (0.2 lg/L), Pb (0.1 lg/L) and V (0.5 lg/L). 2.6. Human cells Routine monitoring has shown the cells to be mycoplasma-free (Mycoalert kit from Cambrex, Verviers, France). Stocks of cells were routinely frozen and stored in liquid N2. All experiments were performed using the cell lines on 10 passages after thawing. 2.6.1. HepG2 cell line The HepG2 cell line was obtained from the ECACC (European Collection of Cell Cultures, UK). The cells were grown in monolayer culture in MEM supplemented with 2 mM L-glutamine, 1% nonessential amino acids and 10% FBS in a humidified atmosphere of 5% CO2 and at 37 °C. Continuous cultures were maintained by subculturing flasks every 7 days at 2.2 106 cells/75 cm2 flask by trypsination (trypsin (0.05%)–EDTA (0.02%)). 2.6.2. MDA-MB453-kb2 cell line This stable transfected human mammary cancer cell line was obtained from the ATCC (LGC Promochem, Molsheim, France). The cells were grown in monolayer culture in Leibovitz medium (L15) supplemented with 10% FBS in a humidified atmosphere at 37 °C. Continuous cultures were maintained by subculturing flasks every 7 days at 4.0 106 cells/75 cm2 flask by trypsination (trypsin (0.05%)–EDTA (0.02%)). 2.6.3. Cells exposure to extracts Bioassays were performed with concentrated bottled water extracts after 10 days at 60 °C. Extracts were tested in bioassays under realistic consumer exposure conditions (1 kg of foodstuff/ 6 dm2 of packaging material) in accordance with EU Regulation (EU, 2011). Cells sensitivity differs depending on the origins and protocols performed: transfected cells are more sensitive to vehicle. Due to


this, for the Ames test and micronucleus assay, the final concentration of bottled water extract was five times more concentrated (1% of ethyl acetate) than for endocrine disruption assays (0.2% of ethyl acetate). 2.7. Genotoxicity assays 2.7.1. Ames test The Ames test was carried out using the plate incorporation method with or without metabolic activation, with two histidinedependent auxotrophic mutants of Salmonella typhimurium strains, TA 98, TA 100, essentially as described by Maron and Ames (1983). The S. typhimurium strains were provided by B. Ames (University of California, Berkeley). The S9 mix was purchased from Trinova Biochem (Giessen, Germany).The test strains were cultured in the liquid broth medium for 10 h at 37 °C under agitation. After incubation, 0.5 mL of 0.1 M sodium phosphate buffer (pH 7.4) (absence of metabolic activation) or 0.5 mL of S9 mix (presence of metabolic activation), 0.1 ml of bacterial culture and 10, 25 or 50 lL of ethyl acetate extracts were placed in a test tube. Two mL of semi-liquid superficial agar was added to the mixture and poured onto a minimal glucose agar plate. The top agar was supplemented with 10 mL of 0.5 mM histidine/biotin solution per 100 mL agar, and mutations to histidine independence were scored on minimal glucose agar plates. The plates were incubated for 48 h at 37 °C and then revertant colonies were counted. All experiments were carried out in triplicate using the three extract concentrations. Mutagenic activity was expressed as an induction factor, i.e. as multiples of the background levels. 2.7.2. Micronucleus assay This assay was performed following the protocol of Séverin, Jondeau, Dahbi, and Chagnon (2005). HepG2 cells were seeded at 2.5 105 cells/well. After 24 h, cells were treated with 1% of the ethyl acetate extract and cytochalasin B (4.5 lg/mL) for 44 h. Cells were then washed with PBS and allowed to recover for 1.5 h in MEM with 10% FBS. The cells were then washed with PBS, trypsinised (trypsin (0.05%)-EDTA (0.02%) solution from Invitrogen laboratories (Cergy-Pontoise, France)), fixed in two steps with acid acetic/methanol (1/3) (v/v), spotted on a glass slide and stained with acridine orange (0.1%) diluted in Sorensen buffer (1/15, v/v) just before reading. Micronuclei were counted visually in 1000 binucleated cells (BNC)/slide using a fluorescence microscope (Olympus CK40) and two slides/concentration were counted. To identify micronuclei, the criteria of Kirsch-Volders et al. (2000) were applied: the diameter of micronuclei should be under onethird that of the main nucleus. They should be clearly distinguishable from the main nucleus and should have the same staining as the main nucleus. 2.8. In vitro endocrine disruptor potential 2.8.1. Estrogenic activity: Transcriptional activation assay with HepG2 cell line 2.8.1.1. Seeding. The estrogenic potential of extracts was determined using HepG2 cells transiently transfected with hERa and with an ER-responsive luminescent reporter gene. HepG2 cells were seeded at a density of 1.2 105 cells per well in 24-well tissue culture plates (Dutscher, France) and maintained in MEM medium without phenol red, supplemented with 10% dextran-coated charcoal foetal calf serum (DCC-FCS), 1% L-glutamin and 1% nonessential amino acids. The microplates were then incubated at 37 °C in a humidified atmosphere of 5% CO2 for 24 h. 2.8.1.2. Transient transfection. Plasmids ERE-TK-Luc and pRST7-hERa were kindly provided by Dr. D. McDonnell (Ligand Pharmaceutical,

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San Diego, USA). Plasmids pCMVb-Gal and pSG5 were kindly provided by Pr M. Cherkaoui-Malki (LBMC, Dijon, France). Each plasmid was first diluted in 0.15 M NaCl (sterile) to a final concentration of 100 ng/lL. HepG2 cells were transiently transfected using the Exgen500 procedure (Euromedex) with the following plasmid mix: 100 ng ERE-TK-Luc and 100 ng hERa, 100 ng of pCMVb-Gal and pSG5 to a final concentration of 0.5 lg DNA. Then, 2 lL of Exgen500 diluted in NaCl 0.15 M was added to the DNA. After vortex shaking, the microtubes were incubated at room temperature for 10 min. The Exgen500-DNA mixture was then added to OptiMEM without phenol red medium and distributed into the wells (300 lL/well). The microplate was then incubated at 37 °C in a humidified atmosphere of 5% CO2 for 1 h (Dumont, 2010). 2.8.1.3. Treatment and analysis. After incubating HepG2 cells for 1 h, the OptiMEM was removed and replaced by 1 mL of treatment medium (MEM without phenol red, without FCS, 1% L-glutamin and 1% non-essential amino acids), containing the water extract, or the vehicle ethyl acetate (1%, negative control), or 17b-estradiol (10 8 M, positive control). The plate was then incubated for 24 h. At the end of the treatment, the medium was aspirated and the wells were rinsed with 100 lL PBS and then lysed using 100 lL Reporter Lysis Buffer 1X (Promega). The microplate was then frozen at 80 °C for at least 15 min. After thawing, cells were scraped and placed into microtubes. The cells then underwent 3 freezing/ thawing cycles in liquid nitrogen and at 37 °C in a water bath. After centrifugation (5 min at 10,600g), luciferase and b-galactosidase activities were determined. For luciferase measurement, 10 lL from each well were transferred into an opaque white-walled plate (Perkin Elmer, Courtaboeuf, France) and mixed with 50 lL of luminol. The plate was quickly covered with an adhesive seal and the mixture was immediately analysed using a luminometer (TopCountNT, Packard). For b-galactosidase activity, 10 lL of each well were used to measure the chlorophenol-red b-D-galactopyranoside (CPRG) (Roche) product with a spectrophotometer at 570 nm (MRX Dynex). Protein absorbance was also measured using 2 lL of the lysate according to the Bradford method on a spectrophotometer at 595 nm (Bradford, 1976). Luciferase induction responses for each treatment group were normalised for b-galactosidase activity and protein level (Luc Prot/Gal) and the results of the different tested extracts were compared with the response observed for 17b-estradiol 10 8 M (set at 100%). 2.8.2. Anti-androgenic activity: Transcriptional activation assay using the human MDA-MB453-kb2 cell line The MDA-MB-453 (AR+) cell line was stably transfected with MMTV-neo-Luc with an (anti)-AR-responsive luminescent reporter gene (Wilson, Bobsein, Lambright, & Gray, 2002). Cells were seeded into a 24-well plate (Dutscher, France) in 1 mL of L15 medium without phenol red, supplemented with 5% of dextran-coated charcoal fetal calf serum (FCS), at a density of 5 104 cells/well. For anti-androgenic activity, 24 h after seeding, the medium was removed and cells were exposed to ethyl acetate extracts (0.05, 0.1 and 0.2%) in the presence of dihydrotestosterone (DHT), the androgenic reference (4 10 10 M, prepared in ethyl acetate). Nilutamide (Nil) (10 6 M, prepared in ethyl acetate) was used as a positive control for anti-androgenic activity. After 24 h treatment, luciferase activity was measured (Stroheker, Picard, Lhuguenot, Canivenc-Lavier, & Chagnon, 2004). Cells were washed once with 1 mL of phosphate buffered saline. Following 30 min incubation with 200 lL/well lysis buffer at room temperature with shaking, lysate were briefly vortexed and centrifuged at 3000g at 4 °C (Stroheker et al., 2004). Ten microlitre from each well were transferred into an opaque white-walled plate and mixed with 40 lL of luciferase assay reagent. The plate was quickly covered with an

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adhesive seal and the mixture was immediately analysed using a luminometer (TopCountNT, Packard). Results are expressed as a percentage of the androgenic positive control (DHT).

3. Results 3.1. Migration of 14 compounds linked to plastic packaging PET- and glass-bottled waters exposed to the worst-case conditions in this study (10 days at 60 °C) were analysed hypothesising that the highest temperature would promote maximal migration. 2,4-di-tert-butylphenol (2,4-dtBP) was detected in bottled waters, while the other compounds were below the quantification limits. As shown in Fig. 1, before experiments (storage time = 0) 2,4-dtBP was already present (0.4 lg/L) in glass-bottled non-carbonated water (brand A) and in PET-bottled water or glass-bottled carbonated water (brand B). After 10 days of exposure at 60 °C, at least a twofold increase of 2,4-dtBP was observed in these bottled waters and the same increase was also found in the ultrapure water placed in brand B bottles (0.7 lg/L). In the same conditions (10 days at 60 °C), an unexpected chromatographic peak appeared exclusively in PET-bottled water extracts. Bis(2-hydroxyethyl) terephthalate (BHET), an intermediate monomer in PET synthesis, was identified with the spectral library (NIST 98 Mass Spectral Library). BHET was not quantified because the analytical standard was not available in the laboratory. The BHET chromatogram and mass spectrum are shown in Fig. 1A in the Additional Data section. According to the molecular ion of BHET (m/z 254), the most intensive peak (m/z 193) corresponds to ion M-61 (M-C2H5O2) due to a double rearrangement of acetate. The m/z 211 peak corresponds to ion M-43 (M-CH3CO) and m/z 149 (typical of phthalate esters) corresponds to ion C8H5O3+.

3.2. Migration of aldehydes 3.2.1. Effect of temperature on formaldehyde and acetaldehyde migration The effect of temperature on aldehydes migration into PET-bottled water was assessed with results obtained in ultrapure water for both brands of PET bottles. With ultrapure water, formaldehyde migration was higher at 60 °C (Fig. 2A) while the release of acetaldehyde (Fig. 2B) had already begun at 50 °C. Acetaldehyde migration appears to be more sensitive to temperature than formaldehyde. At 60 °C, formaldehyde concentration was 4 times lower than acetaldehyde in ultrapure water packaged in brand A bottles and 13 times lower in water in brand B bottles.

3.2.2. Effect of water type (carbonated or non-carbonated) on formaldehyde and acetaldehyde migration In non-carbonated water, as observed before, acetaldehyde migration (Fig. 2B) was higher at 50 °C while formaldehyde migration (Fig. 2A) was higher only at the highest temperature (60 °C). However, data showed a higher dispersion of concentrations especially with formaldehyde in carbonated water, probably due to the presence of residual CO2 not eliminated during sample degasification. In carbonated water, formaldehyde and acetaldehyde were already present at 20 °C. Their concentrations were always higher in carbonated water than in non-carbonated water. At 60 °C, a weak increase (1.6 times) in aldehyde concentrations in carbonated water was observed. Formaldehyde concentrations were 4 times lower than acetaldehyde concentrations in non-carbonated water and 8.5 times lower in carbonated water. 3.3. Migration of trace metals Most of the inorganic elements found in both mineral waters (carbonated and non-carbonated) were those naturally present, except Sb. 3.3.1. Effect of temperature on Sb migration When the water was ultrapure, Sb was not detected in PET-bottled water at 20 °C (Fig. 3). In contrast, Sb concentration gradually increased in ultrapure water between 10 days of storage at 40 °C and at 60 °C from 0.5 to 3.5 lg/L. Therefore, higher temperatures induce higher levels of Sb migration. 3.3.2. Effect of the water type (carbonated or non-carbonated) on Sb migration Sb was already present (around 1 lg/L) in both PET-bottled waters (carbonated and non-carbonated) at 20 °C (Fig. 3). After 10 days at 60 °C, an increase in Sb concentration of 4.8 times and 7.3 times was observed in non-carbonated and carbonated water respectively. Sb concentration in carbonated water was twice as high as non-carbonated water, suggesting that Sb release was accelerated by carbon dioxide. 3.4. Genotoxicity assays 3.4.1. Ames test Induction factors obtained for negative and positive controls performed on the TA 98 and TA 100 strains with and without S9 mix were consistent with the laboratory’s historical data. Bottled waters were not mutagenic (induction factor twice as many as the negative control). As shown in Fig. 4, the cytotoxicity rates calculated for all water extracts were below the maximum recommended value of 55% (OECD, 2010). Bottled water extracts did not induce any chromosome aberration or genomic effect in the HepG2 cells after exposure.

Fig. 4. Micronucleus data in the HepG2 cell line after 10 days of exposure at 60 °C to PET- and glass-bottled water extracts. The solvent control (SC) was DMSO (0.25% final concentration). The negative control (NC) was ethyl acetate (1% final concentration) and the positive control (PC) was a solution of vinblastine sulphate in DMSO (0.005 lM final concentration). ApT and AvT represent non-carbonated mineral water bottled in brand A PET- and glass-bottles respectively. BpT and BvT represent carbonated mineral water in brand B PET- and glass-bottles respectively.

to the HepG2 cells in our experimental conditions (Table A.3 in the Additional Data section). 3.5.2. Anti-androgenic activity As expected, Nilutamide (NIL), the anti-androgenic reference, significantly decreased the androgenic activity of dihydrotestosterone (DHT). In contrast, extracts of the PET-bottled water did not modify the androgenic effect of DHT, suggesting they were not anti-androgenic even at 0.2%, the initial concentration for bottled water (Fig. 5A–D). However, when cells are co-exposed to glass-bottled carbonated water extracts and DHT, a weak but significant increase of the AR transcriptional activity in the MDA-MB456-kb2 cells was observed (Fig. 5D) when compared to the response of DHT alone. The effect was observed at two concentrations (0.05%, 0.2%) without dose-dependency.

3.5. Potential endocrine-disruption activity 4. Discussion 3.5.1. Estrogenic activity The maximum activity (100%) corresponds to the estrogenic activity of 10 8 M 17b-estradiol (E2). Activity of the negative control and extracts was calculated relative to E2. ERa transcriptional activity did not increase when bottled water extracts were exposed

Concerning chemical analysis, formaldehyde and acetaldehyde, well-known PET degradation products, were detected in all the PET-bottled waters. At 60 °C, the migration of these compounds was highly accelerated. This could be explained by the storage


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Fig. 5. Anti-androgenic activity in MDA-MB453-kb2 cell line exposed to bottled-water extracts (10 days at 60 °C). Figures A and B represent non-carbonated water in PET and glass respectively (brand A). Figures C and D represent carbonated water in PET and glass respectively (brand B). MDA-MB453-kb2 cell line was treated with extract concentrations of 0.05%, 0.1% and 0.2%. Ethyl acetate (EA) represents the negative control (0.25% final concentration). Maximum activity (100%) corresponds to the activity of dihydrotestoterone (DHT 4 10 10 M), the androgenic reference. Nilutamide (Nil) (10 6 M) is the positive control for anti-androgenic activity. The sign indicates results statistically different from the DHT positive control (androgenic reference) using the ANOVA statistical test and then a Dunnett’s multiple comparison method. All experiments were performed in triplicate.

temperature which was near the PET glass transition temperature (around 80 °C for semi-crystalline PET), which increases the mobility of polymeric chains directly linked to the migration phenomenon (Bach, Dauchy, David, & Etienne, 2011). We demonstrated that carbon dioxide in bottled water contributed to the increase of migration of both aldehydes. Indeed, formaldehyde and acetaldehyde were substantially present in the bottled carbonated waters before temperature experiments were conducted (10 days at 20 °C). Nijssen, Kamperman, and Jetten (1996) reported that the relatively acid pH of carbonated waters might be responsible for aldehyde migration. In this study, aldehydes were not detected in PET-ultrapure water with a pH very close to carbonated water. This confirms that the pressure exerted on the bottle wall by carbon dioxide could activate aldehyde migration, as previously suggested by Dabrowska, Borcz, and Nawrocki (2003). The same observations were drawn for Sb migration into carbonated waters; in this study Sb migration was also affected by temperature. The presence of Sb in PET-bottled waters (carbonated or non-carbonated) before the experiments were conducted (10 days at 20 °C) suggests that the water’s natural composition may promote Sb migration. However, poor transport conditions and storage cannot be excluded. With regard to EU Regulations, under the worst-case scenario of this study (10 days at 60 °C), concentrations of formaldehyde, acet-

aldehyde and Sb in PET-bottled water were largely below the specific migration limit (SML) (EU, 2011). However, it is worth noting that the formaldehyde level in carbonated water exceeded the French quality standard for bottled mineral waters (5.0 lg/L) (JORF, 2011). In this study, and consistent with chemical formulations of PET (Enneking, 2006; Guart, Bono-Blay, Borrell, & Lacorte, 2011), neither UV stabilisers nor phthalates were detected in bottled water before or after experiments. This is in accordance with Ceretti et al. (2010). In contrast, Montuori, Jover, Morgantini, Bayona, and Triassi (2008) found higher phthalate concentrations (20 times higher) in PET-bottled water than in glass-bottled water. Leivadara, Nikolaou, and Lekkas (2008) detected DEHP in bottled water, but this plasticiser was not identified in tap water stored in the same type of PET bottles. More recently, Amiridou and Voutsa (2011) quantified phthalates in PET-bottled water but they concluded that plasticiser migration was not significant. Phthalate pollution in the bottling line cannot be ruled out. 2,4-dtBP was detected in the ethyl acetate extracts of PET- and glass-bottled water after 10 days at 60 °C. Its presence was previously reported by the German Federal Institute for Risk Assessment (BfR, 2011). The twofold increase of 2,4-dtBP in both PET- and in glass-bottled water suggests that its occurrence was not directly linked to PET. Its presence could be due to the plastic material in


the bottle caps, as a large number of caps are made of polyethylene (PE) and polypropylene (PP) plastics (ILSI, 2003). Simoneau, Van den Eede, and Valzacchi (2012) recently quantified 2,4-dtBP leaching from PP, polyamide and silicone baby bottles. Indeed, 2,4-dtBP is a by-product of tris (2,4-di-tert-butylphenyl) phosphate (Irgafos 168), used to produce PE and PP. As recycled PET is authorised in water bottles, chemicals from non-food application containers found in collection systems of post-consumer PET are a possible source of PET contamination (EFSA, 2011b). This phenolic compound is not on the EU’s positive list (EU, 2011). 2,4-dtBP was cytotoxic when exposed to tumour cells or plants models (Kadoma, Ito, Atsumi, & Fujisawa, 2009; Malek, Shin, Wahab, & Yaacob, 2009). It is also suspected to be an endocrine disruptor. 2,4-dtBP has been shown to be an androgen antagonist in CHO-K1 cells and in rainbow trout (Satoh, 2007; Tollefsen, Eikvar, Finne, Fogelberg, & Gregersen, 2008) and is a compound of very high concern in substance assessment under the REACH Regulation (CoRAP, 2011; EU, 2006). In this study, BHET was also identified in the extracts of PETbottled water, but no specific migration limit (SML) has been established in EU Regulation No. 10/2011 for this substance. Dimethyl terephthalate, a compound of the same chemical family, has been shown to have estrogenic activities in animals and humans (Cheung, Lam, Shi, & Gu, 2007). No other compounds were detected in this study, although chemical analyses are never exhaustive since it is difficult to detect all the substances present (Bradley et al., 2009). Therefore, other non-identified NIAS may be present in PET-bottled water. As this study concerns low concentrations of compounds, two relevant toxicological endpoints, cyto/genotoxicity and endocrine disruption potential, were checked. Bottled water extracts were neither cytotoxic (data not shown), nor genotoxic for HepG2 cells. The extracts were not mutagenic for the S. typhimurium strains TA98 and TA100. This is not in agreement with Biscardi et al. (2003), who showed an eightfold increase in micronuclei when Tradescantia cells were exposed to PET-bottled non-carbonated water over 2 and 3 months as compared to distilled water. A substantial increase in micronuclei was also observed using Allium Cepa cells with carbonated water bottled in PET and glass after 10 days at 40 °C by Ceretti et al. (2010). But the genotoxic effect of bottled water was not reproduced by the latter, when they performed a Comet assay on Tradescantia cells. When using in vitro models, it is important to control the pH, the osmolarity of the culture medium and the cellular survival during compound exposure. Also, CO2 gas in water can create cytotoxic effects (Jondeau, 2006) that would give false positive responses in the genotoxicity assays. In addition, according to international guidelines, plants are not considered as primary ‘‘screening’’ tools for extrapolation to mammals. (EFSA, 2011a). With regard to potential endocrine disruption activities, no estrogenic activity using HepG2 cells was detected in PET- or glass-bottled water extracts prepared with Oasis HLB cartridges under realistic consumer exposure conditions. Our data are not in accordance with Pinto and Reali (2009), Wagner and Oehlmann (2009) or Wagner and Oehlmann (2011), who showed estrogenic activity PET-bottled water extracts. Divergent storage conditions, sample preparation methods (concentration factors, SPE, etc.), cell models (yeasts, snails, etc.) and bioassays (E-screen, YES assay, etc.) were used, making comparison of the results difficult. Wagner and Oehlmann (2011) reported that the use of C18 cartridges and the addition of dimethyl sulfoxide (DMSO) was the most efficient sample preparation method for extracting estrogen-like compounds. Unfortunately, the authors did not identify the compounds at the origin of the positive results they observed. More recently, Plotan, Frizzell, Robinson, Elliott, and Connolly (2012) reported that the level of hormonal activity in bottled water, estimated via daily intake, is not a matter of concern for consumers.

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The potential anti-androgenic activity of concentrated bottled water was also checked because compounds with weak estrogenic activity can also have anti-androgenic activity (Sohoni & Sumpter, 1998) as observed with BPA and phthalates (Stroheker et al., 2005; Xu et al., 2005). As mentioned before, 2,4-dtBP can also have antiandrogenic activity (Satoh, 2007). However, in our experimental conditions, no anti-androgenic activity was detected when MDAMB456-kb2 cells were exposed to PET- or glass-bottled water extracts. The data only showed a weak increase of the androgenic response of DHT in the presence of glass-bottled water extracts compared to DHT alone. This effect could be due to an androgenic activity of the extract itself. As the extract when tested alone was not androgenic (data not shown), this biological response could be due to an interaction such as a potentiation effect of the extract on DHT activity. 5. Conclusions The migration of IAS and NIAS in various bottled waters exposed to different temperatures was investigated. In parallel, in vitro bioassays were performed to check the potential toxicity of chemical mixtures in PET-bottled water. As expected, formaldehyde, acetaldehyde and Sb were detected in PET-bottled waters. We demonstrated that high temperatures and carbonation increased migration in the worst-case scenario used in this study (60 °C for 10 days). 2,4-dtBP and BHET were also identified as NIAS. These two compounds are not on the EU Regulation positive list. However, PET-bottled water after 10 days at 60 °C did not induced any toxic activity (cytotoxicity, genotoxicity or potential endocrine disruption) in vitro. Chemical analysis and global approaches using pertinent and sensitive bioassays are complementary tools to identify the potential hazard of all compounds able to migrate. Acknowledgements This research was supported by the French Agency for Food, Environmental and Occupational Health & Safety (ANSES) and the Institute Jean Lamour of the University of Lorraine. The authors wish to thank the Water Chemistry Department of ANSES’s Nancy Laboratory for Hydrology for their excellent technical assistance. The authors are grateful to Coralie Dumont, Khadija Raja, Anne Novelli, Valérie Fessard, Christian Tricard, and Eric Barthélémy for their collaboration. References Amiridou, D., & Voutsa, D. (2011). Alkylphenols and phthalates in bottled waters. Journal of Hazardous Materials, 185(1), 281–286. Bach, C., Dauchy, X., Chagnon, M. C., & Etienne, S. (2012). Chemical compounds and toxicological assessments of drinking water stored in polyethylene terephthalate (PET) bottles: A source of controversy reviewed. Water Research, 46(3), 571–583. Bach, C., Dauchy, X., David, L., & Etienne, S. (2011). Physico-chemical study of PET bottles and PET bottled water (in French). Matériaux & Techniques, 99, 391–408. BfR (2011). Opinion No. 007/2011. BfR assesses analyse of substances with hormone-like activity in natural mineral waters. Available at: Accessed 13.06.11. Biscardi, D., Monarca, S., De Fusco, R., Senatore, F., Poli, P., Buschini, A., Rossi, C., & Zani, C. (2003). Evaluation of the migration of mutagens/carcinogens from PET bottles into mineral water by Tradescantia/micronuclei test, Comet assay on leukocytes and GC/MS. Science of the Total Environment, 302(1–3), 101–108. Bradford, M. M. (1976). A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein–dye binding. Analytical Biochemistry, 72(1–2), 248–254. Bradley, E. L., Stammati, A., Salkinoja-Salonen, M., Andersson, M., Bertaud, F., Hoornstra, D., et al. (2009). Test procedures for obtaining representative extracts suitable for reliable in vitro toxicity assessment of paper and board intended for food contact. Food Additives & Contaminants, 27(2), 262–271. Ceretti, E., Zani, C., Zerbini, I., Guzzella, L., Scaglia, M., Berna, V., Donato, F., Monarca, S., & Feretti, D. (2010). Comparative assessment of genotoxicity of mineral

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