effect of sunlight exposure on the release of intentionally and/or … safety/pet...

Food Chemistry 162 (2014) 63–71

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Food Chemistry

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Effect of sunlight exposure on the release of intentionallyand/or non-intentionally added substances from polyethyleneterephthalate (PET) bottles into water: Chemical analysisand in vitro toxicity

http://dx.doi.org/10.1016/j.foodchem.2014.04.0200308-8146/� 2014 Elsevier Ltd. All rights reserved.

⇑ Corresponding author at: ANSES, Nancy Laboratory for Hydrology, WaterChemistry Department, 40 rue Lionnois, 54000 Nancy, France.

E-mail address: [email protected] (C. Bach).

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, Franceb Derttech ‘‘Packtox’’, Nutox team, AgroSupDijon Nord, 1 Esplanade Erasme, 21000 Dijon, Francec Institute Jean Lamour, UMR 7198, Department SI2M, Ecole des Mines de Nancy, University of Lorraine, Parc de Saurupt, CS 14234, 54042 Nancy, France

a r t i c l e i n f o

Article history:Received 18 October 2013Received in revised form 3 March 2014Accepted 3 April 2014Available online 13 April 2014

Keywords:PET-bottled watersMigrationSunlightNIASGenotoxicityEndocrine disruptionAldehydesAntimonyChemical analysis

a b s t r a c t

The effect of sunlight exposure on chemical migration into PET-bottled waters was investigated. Bottledwaters were exposed to natural sunlight for 2, 6 and 10 days. Migration was dependent on the type ofwater. Formaldehyde, acetaldehyde and Sb migration increased with sunlight exposure in ultrapurewater. In carbonated waters, carbon dioxide promoted migration and only formaldehyde increasedslightly due to sunlight. Since no aldehydes were detected in non-carbonated waters, we conclude thatsunlight exposure has no effect. Concerning Sb, its migration levels were higher in carbonated waters.No unpredictable NIAS were identified in PET-bottled water extracts. Cyto-genotoxicity (Ames andmicronucleus assays) and potential endocrine disruption effects (transcriptional-reporter gene assays)were checked in bottled water extracts using bacteria (Salmonella typhimurium) and human cell lines(HepG2 and MDA-MB453-kb2). PET-bottled water extracts did not induce any toxic effects (cyto-geno-toxicity, estrogenic or anti-androgenic activity) in vitro at relevant consumer-exposure levels.

� 2014 Elsevier Ltd. All rights reserved.

1. Introduction

PET is a polymer with very few additives used for its manufac-ture; plasticisers and antioxidants are not necessary to producePET bottles and colorants are added only in small quantities.Acetaldehyde scavengers are used to minimise the formation ofacetaldehyde during the melt-process. Also, titanium nitride nano-particles can be incorporated into PET bottle grade (EFSA, 2012).Even if starting substances and additives are strictly regulated byEU Regulation No. 10/2011, several substances known as NIAS(non-intentionally added substances) can be found in the finalplastic material, due to complex formulations of polymers,processes and storage (e.g. impurities, degradation products,breakdown products, etc.) (EU, 2011). These substances can also

migrate into foodstuffs. In addition, physical stress applied to aplastic material can modify the structure of its chemical ingredi-ents (with no toxicological concern) and generate NIAS whichmay have potential estrogenic and/or anti-androgenic activities(Yang, Yaniger, Jordan, Klein, & Bittner, 2011). According to EU Reg-ulation No. 1935/2004 (EU, 2004), ‘‘food contact materials must nottransfer their constituents to food in quantities which could endangerhuman health’’. Furthermore, EU Regulation No. 10/2011 (EU, 2011)states that ‘‘the risk assessment of a substance should cover thesubstance itself, relevant impurities and foreseeable reaction anddegradation products in the intended use’’.

A polymer exposed to sunlight may undergo photochemicalaging, which is the case with PET, which absorbs sunlight at awavelength (k) range located at the end of UV light spectra(300 nm 6 k 6 330 nm). Exposing PET bottles to sunlight, whichalso increases the water’s temperature, raises questions aboutthe formation of by-products and their migration into water, as apossible source of health hazards for the consumers. Few studies

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64 C. Bach et al. / Food Chemistry 162 (2014) 63–71

are available on photoproducts released into PET-bottled waterexposed to sunlight, and when they are available, case toxicitieshave not always been assessed in parallel. The presence ofaldehydes, phthalates, bisphenol A and 4-nonylphenol in PET-bottled waters following sunlight exposure were observed, butwith a wide range of concentrations and storage times whichmakes data comparison difficult. Furthermore, compounds werenot systematically presents or their levels were not statistically dif-ferent in the water samples before and after exposure to sunlight(see review in Bach, Dauchy, Chagnon, and Etienne (2012)). In vitrogenotoxicity using plant and eukaryote cell models has beenobserved in PET-bottled waters exposed to sunlight, but thechemicals responsible for these effects were not identified(Corneanu, Corneanu, Jurescu, & Toptan, 2010; Ubomba-Jaswa,Fernandez-Ibanez, & McGuigan, 2010).

PET containers are sometimes exposed to direct sunlight due topoor storage conditions in retail stores and consumers’ homes,which causes degradation of the polymer through thermo-mechanical and thermo-oxidative processes, generating NIASwhich can migrate into the bottled water (Bach et al., 2012). In fact,in a previous study we demonstrated that high temperaturesincrease migration of formaldehyde, acetaldehyde and Sb intoPET-bottled waters. In addition, we identified two NIAS (2,4-di-tert-butylphenol and bis(2-hydroxyethyl)terephthalate) in bottledwaters. However, bottled water extracts were not found to becyto/genotoxic, estrogenic or anti-androgenic when using in vitrobioassays (Bach et al., 2013).

The objective of the study was to investigate the effect of sun-light on chemical migration into PET-bottled waters and to checkthe potential toxicities of water extracts using in vitro bioassaysin order to avoid any hazard due to unpredictable NIAS (Muncke,2011). The release of formaldehyde, acetaldehyde and Sb was mon-itored in bottled waters exposed to sunlight for 2, 6 and 10 days.Other potential migrants linked to plastic packaging (phthalates,nonylphenols, etc.) were also checked. Experiments wereperformed under realistic conditions of human exposure accordingto the EU Regulation No. 10/2011 (EU, 2011). Sax (2010) and Yanget al. (2011) mentioned that all plastics may yield endocrine dis-ruptors under regular conditions of use. Next, relevant toxicologi-cal endpoints such as cyto/genotoxicity and also endocrinedisruption potential were tested in bottled water extracts as acomplementary approach to chemical analysis. Bioassays are use-ful tools to check potential toxicity due to unpredictable NIASand/or chemical mixtures. Indeed, exhaustive analytical identifica-tion and confirmation of all compounds present in the migrates isdifficult (Nerín, Alfaro, Aznar, & Domeño, 2013). Ames and micro-nucleus assays were performed to assess cyto/genotoxicity usingprokaryotes and a human cell line (HepG2), respectively. Endocrinedisruption potential (estrogenic and anti-androgenic) was assessedby gene reporter assays using human HepG2 and MBA-MB453-kb2cell lines. Bioassays were chosen in accordance with EFSA andICCVAM recommendations (EFSA, 2011; ICCVAM, 2003) for theirperformance. Results of bioassays were then correlated to chemicalanalysis.

Table 1Radiation values and mean temperatures reached in PET-bottled waters.

Exposure duration (days) Irradiation (MJ/m2) Water tempera

Brand A bottle

Mean

2 47.43 25.36 119.79 26.310 237.90 27.6

* Not available.

2. Material and methods

2.1. Samples and storage conditions

Two French brands of non-carbonated (brand A) and carbon-ated (brand B) water bottled in PET and in glass purchased froma local store were investigated. Brand A bottles had a light blue col-our and were made up of a single PET layer with a pattern in reliefon the surface. Brand B bottles had a green colour with a smoothsurface and were made up of an immiscible lamellar polyamide(PA) phase within the PET. This PA phase reduces the permeabilityof O2 and CO2. This type of PET bottle was usually used for carbon-ated water. Water samples for each brand were from identicalbatches. For the experiments, three samples were derived fromeach brand by replacing mineral water by ultrapure water.

Bottled waters were exposed to sunlight for 2, 6 and 10 daysduring July and August 2010 in the Bandol Weathering Station,Southern France. Samples were placed south-facing with aninclination of 45 degrees following the protocol described in thestandard method ISO 877 (ISO, 2009). During the experiments,the solar irradiation received by the packaging material for eachexposure duration was measured and the temperature of thebottled water was monitored using Thermo-tracers� (Oceasoft,Montpellier, France) (Table 1).

2.2. Solid phase extraction (SPE)

The presence of 14 compounds found in plastic packaging was eval-uated, namely: dimethyl phthalate (DMP), butylated hydroxytoluene(BHT), 2,6-di-tert-butyl-p-benzoquinone, 2,4-di-tert-butylphenol(2,4-dtBP), ethyl-4-ethoxybenzoate, diethyl phthalate (DEP), benzo-phenone, 4-nonylphenol (NP), 3,5-di-tert-butyl-4-hydroxybenzalde-hyde (BHT-CHO), di-iso-butyl phthalate (DiBP), dibutyl phthalate(DBP), 2-ethylhexyl-p-methoxycinnamate, di-2-ethylhexyl adipate(DEHA) and di-2-ethylhexyl phthalate (DEHP) (Table 1S,Supplementary data). One litre of water was spiked with surrogatestandards (2,6-di-tert-butyl-d9-4-methylphenol-3,5-d2, benzophe-none-d5 and,di-2-ethylhexyl-phthalate-3,4,5,6-d4) at concentra-tions ranging from 0.5 to 1.6 lg/l depending on the targetcompounds. The water samples were then loaded on Oasis HLB glasscartridges (6 cc/200 mg, Waters, Milford, USA) previously condi-tioned with 5 ml of ethyl acetate (EA), methanol (MeOH) andUPLC-grade water (Biosolve, Valkenswaard, the Netherlands).Analytes were eluted with 2 ml of EA directly analysed by GC–MS(Section 2.3). In parallel, bottled samples were extracted for toxico-logical tests following the same procedure, although deuteredstandards were not added to the water samples.

Chemical analysis and bioassays were then carried out on theEA extracts obtained (concentration factor 500). Preliminary toxic-ity tests of EA extracts were carried out for the cell lines used inthis study (HepG2 and MDA-MB453-kb2 cells) to check thecytotoxicity of the solvent. EA was not cytotoxic at the finalconcentration of 1% in the culture medium (data not shown).

tures (�C)

s Brand B bottles

Min. Max. Mean Min. Max.

16.5 42.5 * * *

17.0 43.5 27.4 16.5 45.516.5 45.5 * * *

C. Bach et al. / Food Chemistry 162 (2014) 63–71 65

2.3. GC–MS analysis

A Varian 450 gas chromatograph (GC) coupled to a Varian 240ion trap mass spectrometer (MS) (Walnut Creek, CA, USA) was usedto analyse EA extracts. Large injection volumes (4 ll) in the splitmode (1:25) were carried out. The inlet temperature was pro-grammed as follows: 40 �C (hold 1 min) to 300 �C at 100 �C/minand hold at 300 �C for 15 min. Analytes were separated on anRxi-5MS column (30 m � 0.25 mm; 0.25 lm film thickness) con-nected with a 5 m � 0.53 mm deactivated pre-column (Restek,Bellefonte, USA). The oven program was: 40 �C (hold 1 min) to280 �C at 8 �C/min and 280 �C (hold for 15 min). Helium (carriergas) was set at 1 ml/min. The transfer line, source and trap temper-ature were 310 �C, 220 �C and 200 �C, respectively. Data wasacquired in full scan mode at a range of 40–600 m/z. The list of ionsselected for the quantification is provided in Table 1S (Supplemen-tary data).

The LOQs were set on the basis of a signal-to-noise ratio of 10.However, phthalates were observed in blanks. Consequently, thephthalates’ LOQs were calculated to never exceed three times theLOQs of the blank values in order to ensure that the backgroundcontamination level remained lower that their limit of detection(LOD). Blanks were prepared with 1 l of UPLC-grade water (Bio-solve, Dieuze, France) spiked with the labelled standards at0.4 lg/l following the extraction procedure described (Section 2.2).The LODs for the analytes were defined as LOQ/3 (ISO/TS13530—Guidance on Analytical Quality Control for Chemical and Physico-chemical Water Analysis). For the method employed here, theLOQ ranged from 0.1 lg/l (for most of the target compounds) to0.3 lg/l (2,4-dtBP and 2-ethylhexyl-p-methoxycinnamate)(Table 1S).

The concentration ranges for performing external calibrationwere from 10 to 1000 lg/l depending on the target compounds.Recovery experiments were carried out with spiked ULPC waterand ranged from 44% to 114% (Table 1S). To ensure the validityof quantification during GC–MS analysis, calibration verificationswere run for each sample batch. Analytical runs were acceptableif analyte concentrations in the calibration verifications werewithin ± 20% of the average concentration determined for eachcompound. For each sample batch, several water samples were for-tified (concentrations from 0.5 to 1.6 lg/l depending on targetcompounds) with labelled standards and analytes to improve theefficiency of extraction and to detect matrix effects, respectively.UPLC blanks were also prepared for each sample batch in orderto ensure that the contamination of lab glassware, connections,solvents and the analytical instrument were lower than the LODs.

2.4. Aldehyde analysis in bottled waters

Aldehyde (formaldehyde, acetaldehyde, propanal, butanal, cro-tonaldehyde, pentanal, hexanal, heptanal, octanal, nonanal anddecanal) analysis in bottled waters was performed following theprotocol previously described by Bach et al. (2013). A derivatisa-tion reaction was carried out with 500 ll of 2,4-dinitrophenylhy-drazine (2,4-DNPH) reagent solution (2 mg/ml in acetonitrile(AcCN)) added to water samples (550 ml). The reaction conditionswere 4 h at 60 �C without agitation. Carbonated water sampleswere degassed after derivatisation. The DNPH derivatisedaldehydes were loaded through Oasis HLB cartridges (200 mgadsorbent, 6 cc; Waters, Milford, MA, USA) previously conditionedwith AcCN (2 � 5 ml) and citrate buffer solution at 1 M (2 � 5 ml).The elution was carried out with 6 ml AcCN (2 � 3 ml). Ultrapurewater was used to adjust the extracts to 7 ml prior to analysis.An Agilent 1200 HPLC system with an Agilent 1200 diode arraydetector (Palo Alto, CA, USA) was used for the aldehyde-DNPHanalysis. Chromatographic separation was achieved with a

SunFire™ C18 column (250 � 4.6 mm I.D.; particle size, 5 lm;Waters, Milford, MA, USA) with a binary mixture of AcCN (A) andultrapure water (B). The gradient program was as follows: isocraticelution at 60% A for 20 min, increase A to 90% over 15 min, and iso-cratic elution at 90% A. Detection was performed at a wavelengthof 360 nm. Matrix-matched calibration was prepared with concen-trations from 1 to 10 lg/l. The quantification limits (LOQ) weredefined as the tenfold value of results obtained with ultrapurewater blanks. The LOQ was 3.5 lg/l for formaldehyde, 2 lg/l foracetaldehyde and octanal, 3 lg/l for nonanal and decanal and1.5 lg/l for the other aldehydes.

2.5. Analysis of trace metals

Bottled water samples were analysed using Series XII inductivelycoupled plasma mass spectrometry (ICP-MS) (Thermo, Germany)following the ISO 17294-2 standard method (ISO, 2003). The oper-ating conditions were as follows: RF power was 1318 W, the carrier,the auxiliary and the nebulizer argon gas flow were 13.0, 0.88 and0.69 dm3/min, respectively. Rhodium at a concentration of 1 lg/lwas used as the internal standard. The LOQ was 1 lg/l for tracemetals, except for Sb (0.2 lg/l), Pb (0.1 lg/l) and V (0.5 lg/l).

2.6. Human cells

Routine monitoring showed the cells to be mycoplasma-free(Mycoalert kit from Cambrex, Verviers, France). Stocks of cellswere routinely frozen and stored in liquid N2. All experiments wereperformed using the cell lines on 10 passages after thawing.

2.6.1. HepG2 cell lineThe HepG2 cell line was obtained from the ECACC (European

Collection of Cell Cultures, UK). The cells were grown in monolayerculture in MEM supplemented with 2 mM L-glutamine, 1% non-essential amino acids and 10% FBS in a humidified atmosphere of5% CO2 at 37 �C. Continuous cultures were maintained by subcul-turing flasks every 7 days at 2.2 � 106 cells/75 cm2 flask by trypsi-nation (trypsin (0.05%)–EDTA (0.02%)).

2.6.2. MDA-MB453-kb2 cell lineThis 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 at37 �C. Continuous cultures were maintained by subculturing flasksevery 7 days at 4.0 � 106 cells/75 cm2 flask by trypsination (trypsin(0.05%)–EDTA (0.02%)) solution from Invitrogen laboratories(Cergy-Pontoise, France).

2.6.3. Cell exposure to extractsBioassays were performed with concentrated bottled water

extracts after 10 days of sunlight exposure (238 MJ/m2 irradiation).Extracts were tested in bioassays under realistic consumer expo-sure conditions (1 kg of foodstuff/6 dm2 of material surface) inaccordance with EU Regulation No. 10/2011 (EU, 2011).

Cell sensitivity differs depending on the origins and protocolsfollowed. Since transfected cells are more sensitive to vehicle, forthe Ames test and micronucleus assay the final concentration ofbottled water extract was 5 times more concentrated (1% of EA)than for the endocrine disruption assays (0.2% of EA).

2.7. Genotoxicity assays

2.7.1. Ames testThe Ames test was carried out using the plate incorporation

method with or without metabolic activation, with two histidine-


dependent auxotrophic mutants of Salmonella typhimurium strains,TA 98 and TA 100, essentially as described by Maron and Ames(1983). The S. typhimurium strains were provided by B. Ames (Uni-versity of California, Berkeley, USA). The S9 mix was purchasedfrom Trinova Biochem (Giessen, Germany). The protocol usedwas described by Bach et al. (2013). All the experiments were car-ried out in triplicate using three extract concentrations. Mutagenicactivity was expressed as an induction factor, i.e. as a multiple ofthe background level.

2.7.2. Micronucleus assayThis assay was performed following the protocol by Severin,

Jondeau, Dahbi, and Chagnon (2005). HepG2 cells were seeded at2.5 � 105 cells/well. After 24 h, cells were treated with 1% of theEA extract and cytochalasin B (4.5 lg/ml) for 44 h. Cells were thenwashed with PBS and allowed to recover for 1.5 h in MEM with 10%FBS. After washing with PBS, the cells were trypsinised (trypsin(0.05%)–EDTA (0.02%)) solution from Invitrogen laboratories(Cergy-Pontoise, France), fixed in two steps with acetic acid/MeOH(1/3) (v/v), spotted on a glass slide and stained with acridineorange (0.1%) diluted in Sorensen Buffer (1/15, v/v) just beforereading. Micronuclei were counted visually in 1000 binucleatedcells (BNC) per slide using a fluorescence microscope (OlympusCK40) and two slides per concentration were counted. To identifymicronuclei, the criteria established by Kirsch-Volders et al.(2000) was applied: the diameter of micronuclei should be underone-third of that of the main nucleus, they should be clearly distin-guishable from the main nucleus and they should have the samestaining as the main nucleus.

2.8. In vitro endocrine disruptor potential

2.8.1. Estrogenic activity: Transcriptional activation assay with HepG2cell line

The protocol used was recently described by Bach et al. (2013).Briefly, HepG2 cells were seeded at a density of 1.2 � 105 cells per

Fig. 1. Formaldehyde and acetaldehyde mean concentrations with standard deviations ialdehyde migration into ultrapure water stored in PET bottles of brands A and B, respe(brand A) and in carbonated water (brand B), respectively. All analyses were performed

well in 24-well tissue culture plates (Dutscher, France) and main-tained in MEM medium without phenol red, supplemented with10% dextran-coated charcoal fetal calf serum (DCC-FCS), 1% L-glu-tamin and 1% non-essential amino acids. The microplates werethen incubated at 37 �C in a humidified atmosphere of 5% CO2 for24 h. HepG2 cells were transiently transfected using the Exgen500procedure (Euromedex) with the following plasmid mix: 100 ngERE-TK-Luc and 100 ng hERa, 100 ng of pCMV-Gal and pSG5 to afinal concentration of 0.5 lg DNA. Then, 2 ll of Exgen500 dilutedin NaCl 0.15 M was added to the DNA. After vortex shaking, themicrotubes were incubated at room temperature for 10 min. TheExgen500-DNA mixture was then added to OptiMEM without phe-nol red medium and distributed into the wells (300 ll/well). Themicroplate was then incubated at 37 �C in a humidified atmo-sphere of 5% CO2 for 1 h. After incubation, the OptiMEM wasremoved and replaced by 1 ml of treatment medium (MEM with-out phenol red, without FCS, 1% glutamin and 1% non-essentialamino acids), containing the water extract, or the vehicle EA (1%,negative control), or 17-estradiol (10�8 M, positive control). Theplate was then incubated for 24 h. At the end of the treatment,luciferase and -galactosidase activity was determined.

2.8.2. Anti-androgenic activity: Transcriptional activation assay usingthe human MDA-MB453-kb2 cell line

The MDA-MB-453 (AR+) cell line was stably transfected withMMTV-neo-Luc with an (anti)-AR-responsive luminescent reportergene (Wilson, Bobsein, Lambright, & Gray, 2002). Cells were seededinto a 24-well plate (Dutscher, France) in 1 ml of L15 mediumwithout phenol red, supplemented with 5% of dextran-coated char-coal fetal calf serum (FCS), at a density of 5 � 104 cells/well. Foranti-androgenic activity, 24 h after seeding, the medium wasremoved and cells were exposed to EA extracts (0.05%, 0.15% and0.2%) in the presence of the androgenic reference dihydrotestoster-one (DHT), (4 � 10�10 M, prepared in EA). Nilutamide (NIL)(10�6 M, prepared in EA) was used as a positive control for anti-androgenic activity. After 24 h treatment, cells were washed once

n PET-bottled waters exposed to sunlight for 2, 6 and 10 days. (A and B) Representctively. (C and D) Correspond to the aldehyde migration in non-carbonated waterin quintuplicate (water from five different bottles).


with 1 ml of phosphate buffered saline. Following 30 min incuba-tion with 200 ll/well lysis buffer at room temperature with shak-ing, the lysates were briefly vortexed and centrifuged at 3000g at4 �C for luciferase activity measurement, as described byStroheker, Picard, Lhuguenot, Canivenc-Lavier, and Chagnon(2004). Ten ll from each well was transferred into an opaquewhite-walled plate and mixed with 40 ll of luciferase assayreagent. The plate was quickly covered with an adhesive seal andthe mixture was immediately analyzed using a luminometer (Top-CountNT, Packard). Results were expressed as a percentage of theandrogenic positive control (DHT).

3. Results

3.1. Migration of 14 compounds linked to plastic packaging

In PET- and glass-bottled waters exposed to the worst-caseconditions (10 days of direct sunlight), 2,4-di-tert-butylphenol(2,4-dtBP) was detected but could not be quantified because itscontent was between the limit of detection (LOD) and the LOQ ofthe analytical method.

Fig. 2. Sb mean concentrations with standard deviations in ultrapure water (A) andin mineral water (carbonated or non-carbonated) (B) packaged in PET bottles ofbrands A and B after 2, 6 and 10 days of sunlight exposure. All analyses wereperformed in quintuplicate (water from five different bottles).

3.2. Migration of aldehydes

Aldehydes were not detected in glass-bottled water before orafter sunlight exposure. Only formaldehyde and acetaldehyde werefound in PET-bottled water. The migration results are presented inthe following subsections.

3.2.1. Effect of sunlight exposure on formaldehyde and acetaldehydemigration

Impact of sunlight exposure on aldehyde migration into bottledwater was assessed with ultrapure waters for both brands of PETbottles. In brand A bottles, formaldehyde and acetaldehyde migra-tion increases to 11 lg/l and to 15 lg/l, respectively, after 10 daysof exposure (Fig. 1A). However, in brand B bottles, formaldehydemigration was observed only after 10 days while acetaldehyderelease was already observed at day 2 (Fig. 1B). At day 10, acetal-dehyde concentrations were still higher than formaldehyde (1.4times higher in brand A bottles and twice as high in brand Bbottles).

3.2.2. Effect of water type (non-carbonated or carbonated) onformaldehyde and acetaldehyde migration

In non-carbonated water (Fig. 1C), aldehyde migration was notobserved, while in carbonated water (Fig. 1D), both aldehydes werealready present before exposure (day 0) at 5 lg/l and 45 lg/l,respectively. A weak effect of sunlight on carbonated water withregard to formaldehyde migration was observed only at day 10with a two-fold increase. In contrast, for acetaldehyde no sunlighteffect was observed. This was due to the presence of carbon diox-ide, which had already promoted its migration before the exposureexperiments. Otherwise a steady concentration of acetaldehydewas observed.

3.3. Migration of trace metals

3.3.1. Effect of sunlight exposure on Sb migrationWith ultrapure water, sunlight exposure slightly increased Sb

migration between 0 and 2 days, and then reached a plateau forboth bottle brands (Fig. 2A), leading to a 0.5-fold increase.

3.3.2. Effect of the water type (non-carbonated or carbonated) on Sbmigration

At day 0, Sb was already present in non-carbonated and carbon-ated waters at concentration levels of 0.7 lg/l and 1.1 lg/l, respec-tively (Fig. 2B). In non-carbonated water, a weak effect of sunlighton the migration of Sb was observed (1.4 times concentrationincrease at day 10). Sb migration was more pronounced in carbon-ated water (1.8 times concentration increase), probably due to thepresence of carbon dioxide.

3.4. Genotoxicity assays

3.4.1. Ames testThe results of the Ames test on water extracts are presented in

Table 2S (Supporting information). Negative and positive controlswere consistent with the laboratory’s historical data. No mutageniceffect due to extracts was observed (induction factors

Fig. 3. Micronucleus data in HepG2 cells treated with bottled water extracts after10 days of sunlight exposure. ApUV and AvUV represent non-carbonated water inPET and in glass, respectively (brand A). BpUV and BvUV represent carbonatedwater in PET and in glass, respectively (brand B). The solvent control (SC) was DMSO(0.25% final concentration). The negative control (NC) was ethyl acetate (1% finalconcentration) and the positive control (PC) was a solution at 0.005 lM ofvinblastine sulphate in DMSO.


maximum recommended value of 55% (OECD, 2010). Bottled waterextracts did not induce any chromosome aberrations or genomiceffects in the HepG2 cells after exposure.

3.5. Potential endocrine-disrupting activity

3.5.1. Estrogenic activityEstrogenic activity measured in ERa transiently transfected

HepG2 cells exposed to water extracts are presented in Fig. 4A-D.The maximum activity (100%) was attributed to luciferase activityin the presence of 10�8 M 17b-estradiol (E2) (positive control).Activity of the negative control and extracts was expressed relativeto E2. Under our experimental conditions, no substantial increase inERa transcriptional activity was observed when HepG2 cells wereexposed to PET bottle extracts, suggesting that the waters are not

Fig. 4. Estrogenic activity in HepG2 cell line exposed to bottled water extracts (10 days orespectively (brand A). (C and D) represent carbonated water in PET and in glass, respectivand 0.2%. Ethyl acetate (EA) was the negative control (0.25% final concentration). Maximpositive control. The sign ⁄ indicates results statistically different from the control negativAll experiments were performed in triplicate.

estrogenic even at 0.2% (initial concentration of bottled waters).However, compared to the control, a weak but significant decreaseof the transcriptional activation was observed for the two highestconcentrations (0.1% and 0.2%) without dose dependency in non-carbonated water in PET (Fig. 4A) and for only one concentration(0.05%) in non-carbonated water in glass (Fig. 4B). No changes wereobserved with carbonated water extracts (Fig. 4C and D).

3.5.2. Anti-androgenic activityThe positive control, (Nilutamide at 10�6 M), decreased the

luciferase activity significantly when MDA-MB453-kb2 cells wereco-treated with the androgenic reference (DHT) (4 � 10�10 M).Extracts of glass-bottled water did not modify the AR transcrip-tional activity of DHT, suggesting that they were not anti-andro-genic (Fig. 5A, B and D). In contrast with extracts of PET-bottledwaters, a significant (1.5-fold) increase of the AR transcriptionalactivity (Fig. 5C) at 0.1% concentration was observed with carbon-ated waters compared to the DHT response alone. However, thissignificance could be due to the standard deviation which wasquite high.

4. Discussion

This is the first study in which potential endocrine disruption(estrogenic and anti-androgenic activity) was assessed in PET-bot-tled waters after sunlight exposure along with chemical analyses.Bioassays are useful tools for identifying the potential hazards ofall compounds present in the migrates (IAS and NIAS (known orunpredictable)). Comprehensive information on hazard and qualityassessment of chemical mixtures and their potential interactions(cocktail effects) can be obtained, as has been done for endocrinedisruptors, which have been shown to produce mixture effects(Kortenkamp, 2007). Biological assays are also particularly usefulfor non-threshold toxicity.

The effect of sunlight on the release of formaldehyde and acet-aldehyde in ultrapure waters was observed for both brands of PET

f sunlight exposure). (A and B) Represent non-carbonated water in PET and in glass,ely (brand B). HepG2 cell line was treated with extract concentrations of 0.05%, 0.1%um activity (100%) corresponds to the activity of 17b-estradiol (E2) at 10�8 M, thee EA using the ANOVA statistical test and a Dunnett’s multiple comparison method.

Fig. 5. Anti-androgenic activity in MDA-MB453-kb2 cell line exposed to extracts of bottled water after 10 days of sunlight exposure. (A and B) Represent non-carbonatedwater in PET and in glass, respectively (brand A). (C and D) Represent carbonated water in PET and in glass, respectively (brand B). MDA-MB453-kb2 cell line was treated withextract concentrations of 0.05%, 0.1% and 0.2%. Ethyl acetate (EA) was the negative control (0.25% final concentration). Maximum activity (100%) corresponds to the activity ofdihydrotestoterone (DHT) at 4 � 1010 M, the androgenic reference. Nilutamide (NIL) at 10�6 M was the positive control for anti-androgenic activity. The ⁄ sign indicatesresults statistically different from DHT control using the ANOVA statistical test and a Dunnett’s multiple comparison method. All experiments were carried out in triplicate.


bottles with the highest concentration at day 10. While in brand Abottles, aldehyde release started after 2 days of sunlight, and inbrand B bottles, formaldehyde release occurred only after 10 expo-sure days. Indeed, as described in Section 2.1, brand B bottles pres-ent a PA phase in PET which may slow down the aldehydemigration. As shown in others studies, the chemical quality ofthe raw material and the manufacturing technologies used in theproduction of PET bottles could be the reason that different alde-hyde levels were generated in the PET bottle wall (Mutsugaet al., 2006). In contrast, with non-carbonated mineral waters, nosunlight effect was observed. Neither formaldehyde nor acetalde-hyde was detected, suggesting that heterotrophic bacteria inmineral water and/or water-hardness may have led to their degra-dation and/or affected their migration, respectively (Mutsuga et al.,2006). This is not in accordance with Wegelin et al. (2001), whoidentified 2 lg/l of acetaldehyde in non-carbonated mineralwaters. However, the irradiation dose was 2.3-fold higher. In car-bonated mineral waters, we demonstrated that aldehyde migrationdepended more on water carbonation than on sunlight, especiallyfor acetaldehyde. Indeed, aldehydes were already observed at day0 due to the carbon dioxide, as mentioned by Dabrowska, Borcz,and Nawrocki (2003).

Moreover, acetaldehyde concentrations were always higherthan those of formaldehyde regardless of the water type or sun-light exposure, as already mentioned in a previous study (Bachet al., 2013) in which the impact of temperature was assessed inthe same samples of PET-bottled waters. However, after 10 daysof sunlight, higher formaldehyde levels were observed in ultrapurewaters than after 10 days at 60 �C (between 2 and 5 times higher)(Bach et al., 2013). Therefore, the migration of formaldehydeappears more dependent on sunlight (with a mean temperatureof 27.6 �C in bottled waters).

In contrast, Sb migration is less affected by sunlight. Indeedafter sunlight exposure, Sb concentrations were 4 times lower inmineral waters than for high storage temperatures (Bach et al.,2013). However, as shown for aldehyde migration, carbon dioxidecontributed to Sb migration more than sunlight. Sb concentrationsin this study are of the same order of magnitude (from 0.25 to0.34 lg/l) as in the study of Hungarian PET-bottled waters whichunderwent illumination for 5 days with a daylight lamp, asreported by Keresztes et al. (2009). In contrast, Cheng, Shi,Adams, and Ma (2010) observed higher Sb levels (up to 2.4 lg/l)in ultrapure waters after 7 days of sunlight exposure. The residualconcentration of Sb remaining on the PET bottle surface may varyaccording to the manufacturing process.

Concerning EU Regulation No. 10/2011 on food contact materi-als, formaldehyde, acetaldehyde and Sb concentrations neverreached the specific migration limits of 15 mg/kg, 6 mg/kg and0.04 mg/kg, respectively (EU, 2011). However, under the worst-case conditions (10 days of sunlight), formaldehyde concentrationsin carbonated waters exceeded the French quality limit (5 lg/l) formineral waters twice (JORF, 2011). Formaldehyde confers an off-flavour to mineral waters, deteriorating their organoleptic charac-teristics. Indeed, UV light exposure produced plastic-like off-odours in mineral water packaged in plastic materials (Strube,Buettner, & Groetzinger, 2009).

In this study, neither phthalates, 4-nonylphenol (NP) or UV sta-bilisers were detected in extracts of PET- and glass-bottled watersbefore or after sunlight exposure. This is in accordance with arecent publication which emphasises the fact that plasticisers arenot introduced during the PET manufacturing process (Dévieret al., 2013). Furthermore, phthalates may come from a wide vari-ety of sources (Bach et al., 2012). Contradictory results have beenpublished on the occurrence of phthalates and NP in PET-bottled


waters. Several phthalates (DMP, DEP, DBP and DEHP) weredetected in PET water samples after 10 weeks of sunlight exposureby Casajuana and Lacorte (2003), but they were also found in glasscontainers. Background pollution cannot be excluded. No substan-tial differences in DEHP concentrations in PET-bottled water (con-centrations ranging from 0.10 to 0.38 lg/l) after sunlight exposure(2 days at 34 �C) were observed by Schmid, Kohler, Meierhofer,Luzi, and Wegelin (2008). Other authors even observed a decreasein concentrations after long-term sunlight exposure. This is thecase of Amiridou and Voutsa (2011) who observed lower concen-trations of DEHP, DEP and DBP after storing PET-bottled watersfor 30 days in daylight. Similarly, Leivadara, Nikolaou, and Lekkas(2008) reported that DEHP was not present in PET-bottled waterafter 3 months of exposure to sunlight, although it was initiallypresent in the water samples. The same phenomenon was alsoobserved for NP. Amiridou and Voutsa (2011) also reported a1.25 concentration decrease of NP in bottled water after 30 daysof exposure to sunlight. Neamtu and Frimmel (2006) observed deg-radation of nonylphenol in water caused by solar UV-irradiation.Therefore, the fact that solar irradiation can cause degradation oforganic compounds via photoreactions cannot be excluded.

In our previous temperature study (Bach et al., 2013), a 2-foldconcentration increase of 2,4-dtBP in both PET- and glass-bottledwaters was observed after 10 days at 60 �C. In sunlight exposureexperiments, 2,4-dtBP was only detected as traces (LOD> traces3 months), suggesting that genotoxic compounds undergo degra-dation in non-genotoxic substances.

With plants models, contradictory results and conclusions usingthe Allium Cepa test were reported after sunlight exposure. While a2-fold increase in chromosomal aberrations was showed byEvandri, Tucci, and Bolle (2000) and Corneanu et al. (2010) attrib-uted the chromosomal mutations observed to the mineral salt con-tent of the water as well as the technology used for manufacturingthe PET bottles. Our results are not in accordance with these previ-ous studies due to differences in the bioassays, cell models (plants,human cell lines, etc.) and conditions used to perform them. Incontrast with ecotoxicology, plants systems are not considered asprimary screening tools for extrapolation to mammalian systems(EFSA, 2011). In addition, different sample preparations (cartridges,solvent polarities, etc.) could give different compound extractionefficiencies as demonstrated by Wagner and Oehlmann (2011).

Several authors suggest that PET bottles may yield endocrinedisruptor chemicals under regular conditions of use, such aslong-term storage, high temperatures and exposure to sunlight(Sax, 2010; Wagner & Oehlmann, 2011; Yang et al., 2011). In thisstudy, after 10 days of sunlight exposure, no estrogenic or anti-androgenic activity was detected in PET- and glass-bottled waterextracts using HepG2 and MDA-MB453-kb2 cells, respectively.

5. Conclusions

The effect of sunlight exposure on chemicals release into PET-bottled waters and the potential hazard of water extracts wereinvestigated using in vitro bioassays. The migration of aldehydesand Sb into ultrapure waters increased with sunlight especiallyafter 10 days of exposure without exceeding the current specificmigration limits set in Regulation No. 10/2011. However, an off-flavour can occur due to the level of formaldehyde in carbonatedwaters after 10 days in sunlight. In carbonated mineral water,carbon dioxide contributed to migration more than sunlight. Waterextracts did not induce any cyto-genotoxic or endocrine-disruptionactivity in the bioassays under our experimental conditions. Chem-ical analysis and global approaches using bioassays are comple-mentary tools to identify the potential toxic effects due tounpredictable NIAS and/or chemical mixtures.

Acknowledgements

This research was financed by the French Agency for Food,Environmental and Occupational Health & Safety (ANSES) andthe Institute Jean Lamour of the University of Lorraine. The authorswish to thank the Bandol Weathering Station (SEVN) and theWater Chemistry Department of ANSES’ Nancy Laboratory forHydrology for their excellent technical assistance. The authorsare grateful to C. Dumont, K. Raja, A. Novelli, V. Fessard, C. Tricard,and E. Barthélémy for their collaboration.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at http://dx.doi.org/10.1016/j.foodchem.2014.04.020.

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Effect of sunlight exposure on the release of intentionally and/or non-intentionally added substances from polyethylene terephthalate (PET) bottles into water: Chemical analysis and in vitro toxicity1 Introduction2 Material and methods2.1 Samples and storage conditions2.2 Solid phase extraction (SPE)2.3 GC–MS analysis2.4 Aldehyde analysis in bottled waters2.5 Analysis of trace metals2.6 Human cells2.6.1 HepG2 cell line2.6.2 MDA-MB453-kb2 cell line2.6.3 Cell exposure to extracts

2.7 Genotoxicity assays2.7.1 Ames test2.7.2 Micronucleus assay

2.8 In vitro endocrine disruptor potential2.8.1 Estrogenic activity: Transcriptional activation assay with HepG2 cell line2.8.2 Anti-androgenic activity: Transcriptional activation assay using the human MDA-MB453-kb2 cell line

3 Results3.1 Migration of 14 compounds linked to plastic packaging3.2 Migration of aldehydes3.2.1 Effect of sunlight exposure on formaldehyde and acetaldehyde migration3.2.2 Effect of water type (non-carbonated or carbonated) on formaldehyde and acetaldehyde migration

3.3 Migration of trace metals3.3.1 Effect of sunlight exposure on Sb migration3.3.2 Effect of the water type (non-carbonated or carbonated) on Sb migration

3.4 Genotoxicity assays3.4.1 Ames test3.4.2 Micronucleus assays

3.5 Potential endocrine-disrupting activity3.5.1 Estrogenic activity3.5.2 Anti-androgenic activity

4 Discussion5 ConclusionsAcknowledgementsAppendix A Supplementary dataReferences

effect of sunlight exposure on the release of intentionally and/or … safety/pet...

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