Food Chemistry 162 (2014) 63–71
Contents lists available at ScienceDirect
Food Chemistry
journal homepage: www.elsevier.com/locate/foodchem
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 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, 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 18 October 2013
Received in revised form 3 March 2014
Accepted 3 April 2014
Available online 13 April 2014
Keywords:
PET-bottled waters
Migration
Sunlight
NIAS
Genotoxicity
Endocrine disruption
Aldehydes
Antimony
Chemical analysis
a b s t r a c t
The effect of sunlight exposure on chemical migration into PET-bottled waters was investigated. Bottled
waters were exposed to natural sunlight for 2, 6 and 10 days. Migration was dependent on the type of
water. Formaldehyde, acetaldehyde and Sb migration increased with sunlight exposure in ultrapure
water. In carbonated waters, carbon dioxide promoted migration and only formaldehyde increased
slightly due to sunlight. Since no aldehydes were detected in non-carbonated waters, we conclude that
sunlight 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 and
micronucleus 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-genotoxicity, estrogenic or anti-androgenic activity) in vitro at relevant consumer-exposure levels.
Ó 2014 Elsevier Ltd. All rights reserved.
1. Introduction
PET is a polymer with very few additives used for its manufacture; plasticisers and antioxidants are not necessary to produce
PET bottles and colorants are added only in small quantities.
Acetaldehyde scavengers are used to minimise the formation of
acetaldehyde during the melt-process. Also, titanium nitride nanoparticles can be incorporated into PET bottle grade (EFSA, 2012).
Even if starting substances and additives are strictly regulated by
EU Regulation No. 10/2011, several substances known as NIAS
(non-intentionally added substances) can be found in the final
plastic material, due to complex formulations of polymers,
processes and storage (e.g. impurities, degradation products,
breakdown products, etc.) (EU, 2011). These substances can also
⇑ Corresponding author at: ANSES, Nancy Laboratory for Hydrology, Water
Chemistry Department, 40 rue Lionnois, 54000 Nancy, France.
E-mail address: [email protected] (C. Bach).
http://dx.doi.org/10.1016/j.foodchem.2014.04.020
0308-8146/Ó 2014 Elsevier Ltd. All rights reserved.
migrate into foodstuffs. In addition, physical stress applied to a
plastic material can modify the structure of its chemical ingredients (with no toxicological concern) and generate NIAS which
may have potential estrogenic and/or anti-androgenic activities
(Yang, Yaniger, Jordan, Klein, & Bittner, 2011). According to EU Regulation No. 1935/2004 (EU, 2004), ‘‘food contact materials must not
transfer their constituents to food in quantities which could endanger
human health’’. Furthermore, EU Regulation No. 10/2011 (EU, 2011)
states that ‘‘the risk assessment of a substance should cover the
substance itself, relevant impurities and foreseeable reaction and
degradation products in the intended use’’.
A polymer exposed to sunlight may undergo photochemical
aging, which is the case with PET, which absorbs sunlight at a
wavelength (k) range located at the end of UV light spectra
(300 nm 6 k 6 330 nm). Exposing PET bottles to sunlight, which
also increases the water’s temperature, raises questions about
the formation of by-products and their migration into water, as a
possible source of health hazards for the consumers. Few studies
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C. Bach et al. / Food Chemistry 162 (2014) 63–71
are available on photoproducts released into PET-bottled water
exposed to sunlight, and when they are available, case toxicities
have not always been assessed in parallel. The presence of
aldehydes, phthalates, bisphenol A and 4-nonylphenol in PETbottled waters following sunlight exposure were observed, but
with a wide range of concentrations and storage times which
makes data comparison difficult. Furthermore, compounds were
not systematically presents or their levels were not statistically different in the water samples before and after exposure to sunlight
(see review in Bach, Dauchy, Chagnon, and Etienne (2012)). In vitro
genotoxicity using plant and eukaryote cell models has been
observed in PET-bottled waters exposed to sunlight, but the
chemicals 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 to
poor storage conditions in retail stores and consumers’ homes,
which causes degradation of the polymer through thermomechanical and thermo-oxidative processes, generating NIAS
which can migrate into the bottled water (Bach et al., 2012). In fact,
in a previous study we demonstrated that high temperatures
increase migration of formaldehyde, acetaldehyde and Sb into
PET-bottled waters. In addition, we identified two NIAS (2,4-ditert-butylphenol and bis(2-hydroxyethyl)terephthalate) in bottled
waters. However, bottled water extracts were not found to be
cyto/genotoxic, estrogenic or anti-androgenic when using in vitro
bioassays (Bach et al., 2013).
The objective of the study was to investigate the effect of sunlight on chemical migration into PET-bottled waters and to check
the potential toxicities of water extracts using in vitro bioassays
in order to avoid any hazard due to unpredictable NIAS (Muncke,
2011). The release of formaldehyde, acetaldehyde and Sb was monitored 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 were
performed under realistic conditions of human exposure according
to the EU Regulation No. 10/2011 (EU, 2011). Sax (2010) and Yang
et al. (2011) mentioned that all plastics may yield endocrine disruptors under regular conditions of use. Next, relevant toxicological endpoints such as cyto/genotoxicity and also endocrine
disruption potential were tested in bottled water extracts as a
complementary approach to chemical analysis. Bioassays are useful tools to check potential toxicity due to unpredictable NIAS
and/or chemical mixtures. Indeed, exhaustive analytical identification and confirmation of all compounds present in the migrates is
difficult (Nerín, Alfaro, Aznar, & Domeño, 2013). Ames and micronucleus assays were performed to assess cyto/genotoxicity using
prokaryotes and a human cell line (HepG2), respectively. Endocrine
disruption potential (estrogenic and anti-androgenic) was assessed
by gene reporter assays using human HepG2 and MBA-MB453-kb2
cell lines. Bioassays were chosen in accordance with EFSA and
ICCVAM recommendations (EFSA, 2011; ICCVAM, 2003) for their
performance. Results of bioassays were then correlated to chemical
analysis.
2. Material and methods
2.1. Samples and storage conditions
Two French brands of non-carbonated (brand A) and carbonated (brand B) water bottled in PET and in glass purchased from
a local store were investigated. Brand A bottles had a light blue colour and were made up of a single PET layer with a pattern in relief
on the surface. Brand B bottles had a green colour with a smooth
surface and were made up of an immiscible lamellar polyamide
(PA) phase within the PET. This PA phase reduces the permeability
of O2 and CO2. This type of PET bottle was usually used for carbonated water. Water samples for each brand were from identical
batches. For the experiments, three samples were derived from
each brand by replacing mineral water by ultrapure water.
Bottled waters were exposed to sunlight for 2, 6 and 10 days
during July and August 2010 in the Bandol Weathering Station,
Southern France. Samples were placed south-facing with an
inclination of 45 degrees following the protocol described in the
standard method ISO 877 (ISO, 2009). During the experiments,
the solar irradiation received by the packaging material for each
exposure duration was measured and the temperature of the
bottled water was monitored using Thermo-tracersÒ (Oceasoft,
Montpellier, France) (Table 1).
2.2. Solid phase extraction (SPE)
The presence of 14 compounds found in plastic packaging was evaluated, 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), 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) and di-2-ethylhexyl phthalate (DEHP) (Table 1S,
Supplementary data). One litre of water was spiked with surrogate
standards (2,6-di-tert-butyl-d9-4-methylphenol-3,5-d2, benzophenone-d5 and,di-2-ethylhexyl-phthalate-3,4,5,6-d4) at concentrations ranging from 0.5 to 1.6 lg/l depending on the target
compounds. The water samples were then loaded on Oasis HLB glass
cartridges (6 cc/200 mg, Waters, Milford, USA) previously conditioned with 5 ml of ethyl acetate (EA), methanol (MeOH) and
UPLC-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 toxicological tests following the same procedure, although deutered
standards were not added to the water samples.
Chemical analysis and bioassays were then carried out on the
EA extracts obtained (concentration factor 500). Preliminary toxicity tests of EA extracts were carried out for the cell lines used in
this study (HepG2 and MDA-MB453-kb2 cells) to check the
cytotoxicity of the solvent. EA was not cytotoxic at the final
concentration of 1% in the culture medium (data not shown).
Table 1
Radiation values and mean temperatures reached in PET-bottled waters.
Exposure duration (days)
Irradiation (MJ/m2)
Water temperatures (°C)
Brand A bottles
2
6
10
*
Not available.
47.43
119.79
237.90
Brand B bottles
Mean
Min.
Max.
Mean
Min.
25.3
26.3
27.6
16.5
17.0
16.5
42.5
43.5
45.5
*
*
Max.
*
27.4
16.5
45.5
*
*
*
C. Bach et al. / Food Chemistry 162 (2014) 63–71
2.3. GC–MS analysis
A Varian 450 gas chromatograph (GC) coupled to a Varian 240
ion trap mass spectrometer (MS) (Walnut Creek, CA, USA) was used
to analyse EA extracts. Large injection volumes (4 ll) in the split
mode (1:25) were carried out. The inlet temperature was programmed as follows: 40 °C (hold 1 min) to 300 °C at 100 °C/min
and hold at 300 °C for 15 min. Analytes were separated on an
Rxi-5MS column (30 m 0.25 mm; 0.25 lm film thickness) connected with a 5 m 0.53 mm deactivated pre-column (Restek,
Bellefonte, USA). The oven program was: 40 °C (hold 1 min) to
280 °C at 8 °C/min and 280 °C (hold for 15 min). Helium (carrier
gas) was set at 1 ml/min. The transfer line, source and trap temperature were 310 °C, 220 °C and 200 °C, respectively. Data was
acquired in full scan mode at a range of 40–600 m/z. The list of ions
selected for the quantification is provided in Table 1S (Supplementary data).
The LOQs were set on the basis of a signal-to-noise ratio of 10.
However, phthalates were observed in blanks. Consequently, the
phthalates’ LOQs were calculated to never exceed three times the
LOQs of the blank values in order to ensure that the background
contamination level remained lower that their limit of detection
(LOD). Blanks were prepared with 1 l of UPLC-grade water (Biosolve, Dieuze, France) spiked with the labelled standards at
0.4 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 Physicochemical Water Analysis). For the method employed here, the
LOQ ranged from 0.1 lg/l (for most of the target compounds) to
0.3 lg/l
(2,4-dtBP
and
2-ethylhexyl-p-methoxycinnamate)
(Table 1S).
The concentration ranges for performing external calibration
were from 10 to 1000 lg/l depending on the target compounds.
Recovery experiments were carried out with spiked ULPC water
and ranged from 44% to 114% (Table 1S). To ensure the validity
of quantification during GC–MS analysis, calibration verifications
were run for each sample batch. Analytical runs were acceptable
if analyte concentrations in the calibration verifications were
within ± 20% of the average concentration determined for each
compound. For each sample batch, several water samples were fortified (concentrations from 0.5 to 1.6 lg/l depending on target
compounds) with labelled standards and analytes to improve the
efficiency of extraction and to detect matrix effects, respectively.
UPLC blanks were also prepared for each sample batch in order
to 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, crotonaldehyde, pentanal, hexanal, heptanal, octanal, nonanal and
decanal) analysis in bottled waters was performed following the
protocol previously described by Bach et al. (2013). A derivatisation reaction was carried out with 500 ll of 2,4-dinitrophenylhydrazine (2,4-DNPH) reagent solution (2 mg/ml in acetonitrile
(AcCN)) added to water samples (550 ml). The reaction conditions
were 4 h at 60 °C without agitation. Carbonated water samples
were degassed after derivatisation. The DNPH derivatised
aldehydes were loaded through Oasis HLB cartridges (200 mg
adsorbent, 6 cc; Waters, Milford, MA, USA) previously conditioned
with 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). Ultrapure
water was used to adjust the extracts to 7 ml prior to analysis.
An Agilent 1200 HPLC system with an Agilent 1200 diode array
detector (Palo Alto, CA, USA) was used for the aldehyde-DNPH
analysis. Chromatographic separation was achieved with a
65
SunFire™ C18 column (250 4.6 mm I.D.; particle size, 5 lm;
Waters, Milford, MA, USA) with a binary mixture of AcCN (A) and
ultrapure water (B). The gradient program was as follows: isocratic
elution at 60% A for 20 min, increase A to 90% over 15 min, and isocratic elution at 90% A. Detection was performed at a wavelength
of 360 nm. Matrix-matched calibration was prepared with concentrations from 1 to 10 lg/l. The quantification limits (LOQ) were
defined as the tenfold value of 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 Series XII inductively
coupled plasma mass spectrometry (ICP-MS) (Thermo, Germany)
following the ISO 17294-2 standard method (ISO, 2003). The operating conditions were as follows: RF power was 1318 W, the carrier,
the auxiliary and the nebulizer argon gas flow were 13.0, 0.88 and
0.69 dm3/min, respectively. Rhodium at a concentration of 1 lg/l
was used as the internal standard. The LOQ was 1 lg/l for trace
metals, 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 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 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%)) solution from Invitrogen laboratories
(Cergy-Pontoise, France).
2.6.3. Cell exposure to extracts
Bioassays 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 exposure conditions (1 kg of foodstuff/6 dm2 of material surface) in
accordance with EU Regulation No. 10/2011 (EU, 2011).
Cell sensitivity differs depending on the origins and protocols
followed. Since transfected cells are more sensitive to vehicle, for
the Ames test and micronucleus assay the final concentration of
bottled 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 test
The Ames test was carried out using the plate incorporation
method with or without metabolic activation, with two histidine-
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C. Bach et al. / Food Chemistry 162 (2014) 63–71
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 (University of California, Berkeley, USA). The S9 mix was purchased
from Trinova Biochem (Giessen, Germany). The protocol used
was described by Bach et al. (2013). All the experiments were carried out in triplicate using three extract concentrations. Mutagenic
activity was expressed as an induction factor, i.e. as a multiple of
the background level.
2.7.2. Micronucleus assay
This assay was performed following the protocol by Severin,
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
EA 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. 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 acridine
orange (0.1%) diluted in Sorensen Buffer (1/15, v/v) just before
reading. Micronuclei were counted visually in 1000 binucleated
cells (BNC) per slide using a fluorescence microscope (Olympus
CK40) and two slides per concentration were counted. To identify
micronuclei, the criteria established by Kirsch-Volders et al.
(2000) was applied: the diameter of micronuclei should be under
one-third of that of the main nucleus, they should be clearly distinguishable from the main nucleus and they 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
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
well in 24-well tissue culture plates (Dutscher, France) and maintained in MEM medium without phenol red, supplemented with
10% dextran-coated charcoal fetal calf serum (DCC-FCS), 1% L-glutamin and 1% non-essential amino acids. The microplates were
then incubated at 37 °C in a humidified atmosphere of 5% CO2 for
24 h. 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 pCMV-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. After incubation, the OptiMEM was
removed and replaced by 1 ml of treatment medium (MEM without phenol red, without FCS, 1% glutamin and 1% non-essential
amino acids), containing the water extract, or the vehicle EA (1%,
negative control), or 17-estradiol (10 8 M, positive control). The
plate 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 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 EA extracts (0.05%, 0.15% and
0.2%) in the presence of the androgenic reference dihydrotestosterone (DHT), (4 10 10 M, prepared in EA). Nilutamide (NIL)
(10 6 M, prepared in EA) was used as a positive control for antiandrogenic activity. After 24 h treatment, cells were washed once
Fig. 1. Formaldehyde and acetaldehyde mean concentrations with standard deviations in PET-bottled waters exposed to sunlight for 2, 6 and 10 days. (A and B) Represent
aldehyde migration into ultrapure water stored in PET bottles of brands A and B, respectively. (C and D) Correspond to the aldehyde migration in non-carbonated water
(brand A) and in carbonated water (brand B), respectively. All analyses were performed in quintuplicate (water from five different bottles).
C. Bach et al. / Food Chemistry 162 (2014) 63–71
with 1 ml of phosphate buffered saline. Following 30 min incubation with 200 ll/well lysis buffer at room temperature with shaking, the lysates were briefly vortexed and centrifuged at 3000g at
4 °C for luciferase activity measurement, as described by
Stroheker, Picard, Lhuguenot, Canivenc-Lavier, and Chagnon
(2004). Ten ll from each well was transferred into an opaque
white-walled plate and mixed with 40 ll of luciferase assay
reagent. The plate was quickly covered with an adhesive seal and
the mixture was immediately analyzed using a luminometer (TopCountNT, Packard). Results were expressed as a percentage of the
androgenic 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-case
conditions (10 days of direct sunlight), 2,4-di-tert-butylphenol
(2,4-dtBP) was detected but could not be quantified because its
content was between the limit of detection (LOD) and the LOQ of
the analytical method.
67
3.2. Migration of aldehydes
Aldehydes were not detected in glass-bottled water before or
after sunlight exposure. Only formaldehyde and acetaldehyde were
found in PET-bottled water. The migration results are presented in
the following subsections.
3.2.1. Effect of sunlight exposure on formaldehyde and acetaldehyde
migration
Impact of sunlight exposure on aldehyde migration into bottled
water was assessed with ultrapure waters for both brands of PET
bottles. In brand A bottles, formaldehyde and acetaldehyde migration increases to 11 lg/l and to 15 lg/l, respectively, after 10 days
of exposure (Fig. 1A). However, in brand B bottles, formaldehyde
migration was observed only after 10 days while acetaldehyde
release was already observed at day 2 (Fig. 1B). At day 10, acetaldehyde concentrations were still higher than formaldehyde (1.4
times higher in brand A bottles and twice as high in brand B
bottles).
3.2.2. Effect of water type (non-carbonated or carbonated) on
formaldehyde and acetaldehyde migration
In non-carbonated water (Fig. 1C), aldehyde migration was not
observed, while in carbonated water (Fig. 1D), both aldehydes were
already present before exposure (day 0) at 5 lg/l and 45 lg/l,
respectively. A weak effect of sunlight on carbonated water with
regard to formaldehyde migration was observed only at day 10
with a two-fold increase. In contrast, for acetaldehyde no sunlight
effect was observed. This was due to the presence of carbon dioxide, which had already promoted its migration before the exposure
experiments. Otherwise a steady concentration of acetaldehyde
was observed.
3.3. Migration of trace metals
3.3.1. Effect of sunlight exposure on Sb migration
With ultrapure water, sunlight exposure slightly increased Sb
migration between 0 and 2 days, and then reached a plateau for
both bottle brands (Fig. 2A), leading to a 0.5-fold increase.
3.3.2. Effect of the water type (non-carbonated or carbonated) on Sb
migration
At day 0, Sb was already present in non-carbonated and carbonated waters at concentration levels of 0.7 lg/l and 1.1 lg/l, respectively (Fig. 2B). In non-carbonated water, a weak effect of sunlight
on the migration of Sb was observed (1.4 times concentration
increase at day 10). Sb migration was more pronounced in carbonated water (1.8 times concentration increase), probably due to the
presence of carbon dioxide.
3.4. Genotoxicity assays
3.4.1. Ames test
The results of the Ames test on water extracts are presented in
Table 2S (Supporting information). Negative and positive controls
were consistent with the laboratory’s historical data. No mutagenic
effect due to extracts was observed (induction factors <2) with
either of the S. typhimurium strains (TA 98 and TA 100), with or
without the S9 mix under the experimental conditions used.
Fig. 2. Sb mean concentrations with standard deviations in ultrapure water (A) and
in mineral water (carbonated or non-carbonated) (B) packaged in PET bottles of
brands A and B after 2, 6 and 10 days of sunlight exposure. All analyses were
performed in quintuplicate (water from five different bottles).
3.4.2. Micronucleus assays
The validity of the test was checked against the positive
response obtained using the reference control, showing an increase
in the number of micronuclei in binucleated cells (greater than
twice that of the negative control). As shown in Fig. 3, the cytotoxicity rates calculated for all water extracts were below the
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C. Bach et al. / Food Chemistry 162 (2014) 63–71
estrogenic even at 0.2% (initial concentration of bottled waters).
However, compared to the control, a weak but significant decrease
of the transcriptional activation was observed for the two highest
concentrations (0.1% and 0.2%) without dose dependency in noncarbonated water in PET (Fig. 4A) and for only one concentration
(0.05%) in non-carbonated water in glass (Fig. 4B). No changes were
observed with carbonated water extracts (Fig. 4C and D).
Fig. 3. Micronucleus data in HepG2 cells treated with bottled water extracts after
10 days of sunlight exposure. ApUV and AvUV represent non-carbonated water in
PET and in glass, respectively (brand A). BpUV and BvUV represent carbonated
water 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% final
concentration) and the positive control (PC) was a solution at 0.005 lM of
vinblastine sulphate in DMSO.
maximum recommended value of 55% (OECD, 2010). Bottled water
extracts did not induce any chromosome aberrations or genomic
effects in the HepG2 cells after exposure.
3.5. Potential endocrine-disrupting activity
3.5.1. Estrogenic activity
Estrogenic 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 activity
in the presence of 10 8 M 17b-estradiol (E2) (positive control).
Activity of the negative control and extracts was expressed relative
to E2. Under our experimental conditions, no substantial increase in
ERa transcriptional activity was observed when HepG2 cells were
exposed to PET bottle extracts, suggesting that the waters are not
3.5.2. Anti-androgenic activity
The positive control, (Nilutamide at 10 6 M), decreased the
luciferase activity significantly when MDA-MB453-kb2 cells were
co-treated with the androgenic reference (DHT) (4 10 10 M).
Extracts of glass-bottled water did not modify the AR transcriptional activity of DHT, suggesting that they were not anti-androgenic (Fig. 5A, B and D). In contrast with extracts of PET-bottled
waters, a significant (1.5-fold) increase of the AR transcriptional
activity (Fig. 5C) at 0.1% concentration was observed with carbonated waters compared to the DHT response alone. However, this
significance could be due to the standard deviation which was
quite high.
4. Discussion
This is the first study in which potential endocrine disruption
(estrogenic and anti-androgenic activity) was assessed in PET-bottled waters after sunlight exposure along with chemical analyses.
Bioassays are useful tools for identifying the potential hazards of
all compounds present in the migrates (IAS and NIAS (known or
unpredictable)). Comprehensive information on hazard and quality
assessment of chemical mixtures and their potential interactions
(cocktail effects) can be obtained, as has been done for endocrine
disruptors, which have been shown to produce mixture effects
(Kortenkamp, 2007). Biological assays are also particularly useful
for non-threshold toxicity.
The effect of sunlight on the release of formaldehyde and acetaldehyde in ultrapure waters was observed for both brands of PET
Fig. 4. Estrogenic activity in HepG2 cell line exposed to bottled water extracts (10 days of sunlight exposure). (A and B) Represent non-carbonated water in PET and in glass,
respectively (brand A). (C and D) represent carbonated water in PET and in glass, respectively (brand B). HepG2 cell line was treated with extract 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 of 17b-estradiol (E2) at 10 8 M, the
positive control. The sign ⁄ indicates results statistically different from the control negative EA using the ANOVA statistical test and a Dunnett’s multiple comparison method.
All experiments were performed in triplicate.
C. Bach et al. / Food Chemistry 162 (2014) 63–71
69
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-carbonated
water 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 with
extract 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 of
dihydrotestoterone (DHT) at 4 1010 M, the androgenic reference. Nilutamide (NIL) at 10 6 M was the positive control for anti-androgenic activity. The ⁄ sign indicates
results 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 A
bottles, aldehyde release started after 2 days of sunlight, and in
brand B bottles, formaldehyde release occurred only after 10 exposure days. Indeed, as described in Section 2.1, brand B bottles present a PA phase in PET which may slow down the aldehyde
migration. As shown in others studies, the chemical quality of
the raw material and the manufacturing technologies used in the
production of PET bottles could be the reason that different aldehyde levels were generated in the PET bottle wall (Mutsuga
et al., 2006). In contrast, with non-carbonated mineral waters, no
sunlight effect was observed. Neither formaldehyde nor acetaldehyde was detected, suggesting that heterotrophic bacteria in
mineral water and/or water-hardness may have led to their degradation and/or affected their migration, respectively (Mutsuga et al.,
2006). This is not in accordance with Wegelin et al. (2001), who
identified 2 lg/l of acetaldehyde in non-carbonated mineral
waters. However, the irradiation dose was 2.3-fold higher. In carbonated mineral waters, we demonstrated that aldehyde migration
depended more on water carbonation than on sunlight, especially
for acetaldehyde. Indeed, aldehydes were already observed at day
0 due to the carbon dioxide, as mentioned by Dabrowska, Borcz,
and Nawrocki (2003).
Moreover, acetaldehyde concentrations were always higher
than those of formaldehyde regardless of the water type or sunlight exposure, as already mentioned in a previous study (Bach
et al., 2013) in which the impact of temperature was assessed in
the same samples of PET-bottled waters. However, after 10 days
of sunlight, higher formaldehyde levels were observed in ultrapure
waters than after 10 days at 60 °C (between 2 and 5 times higher)
(Bach et al., 2013). Therefore, the migration of formaldehyde
appears more dependent on sunlight (with a mean temperature
of 27.6 °C in bottled waters).
In contrast, Sb migration is less affected by sunlight. Indeed
after sunlight exposure, Sb concentrations were 4 times lower in
mineral waters than for high storage temperatures (Bach et al.,
2013). However, as shown for aldehyde migration, carbon dioxide
contributed to Sb migration more than sunlight. Sb concentrations
in this study are of the same order of magnitude (from 0.25 to
0.34 lg/l) as in the study of Hungarian PET-bottled waters which
underwent illumination for 5 days with a daylight lamp, as
reported 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 residual
concentration of Sb remaining on the PET bottle surface may vary
according to the manufacturing process.
Concerning EU Regulation No. 10/2011 on food contact materials, formaldehyde, acetaldehyde and Sb concentrations never
reached the specific migration limits of 15 mg/kg, 6 mg/kg and
0.04 mg/kg, respectively (EU, 2011). However, under the worstcase conditions (10 days of sunlight), formaldehyde concentrations
in carbonated waters exceeded the French quality limit (5 lg/l) for
mineral waters twice (JORF, 2011). Formaldehyde confers an offflavour to mineral waters, deteriorating their organoleptic characteristics. Indeed, UV light exposure produced plastic-like offodours in mineral water packaged in plastic materials (Strube,
Buettner, & Groetzinger, 2009).
In this study, neither phthalates, 4-nonylphenol (NP) or UV stabilisers were detected in extracts of PET- and glass-bottled waters
before or after sunlight exposure. This is in accordance with a
recent publication which emphasises the fact that plasticisers are
not introduced during the PET manufacturing process (Dévier
et al., 2013). Furthermore, phthalates may come from a wide variety of sources (Bach et al., 2012). Contradictory results have been
published on the occurrence of phthalates and NP in PET-bottled
70
C. Bach et al. / Food Chemistry 162 (2014) 63–71
waters. Several phthalates (DMP, DEP, DBP and DEHP) were
detected in PET water samples after 10 weeks of sunlight exposure
by Casajuana and Lacorte (2003), but they were also found in glass
containers. Background pollution cannot be excluded. No substantial differences in DEHP concentrations in PET-bottled water (concentrations 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 decrease
in concentrations after long-term sunlight exposure. This is the
case of Amiridou and Voutsa (2011) who observed lower concentrations of DEHP, DEP and DBP after storing PET-bottled waters
for 30 days in daylight. Similarly, Leivadara, Nikolaou, and Lekkas
(2008) reported that DEHP was not present in PET-bottled water
after 3 months of exposure to sunlight, although it was initially
present in the water samples. The same phenomenon was also
observed for NP. Amiridou and Voutsa (2011) also reported a
1.25 concentration decrease of NP in bottled water after 30 days
of exposure to sunlight. Neamtu and Frimmel (2006) observed degradation of nonylphenol in water caused by solar UV-irradiation.
Therefore, the fact that solar irradiation can cause degradation of
organic compounds via photoreactions cannot be excluded.
In our previous temperature study (Bach et al., 2013), a 2-fold
concentration increase of 2,4-dtBP in both PET- and glass-bottled
waters was observed after 10 days at 60 °C. In sunlight exposure
experiments, 2,4-dtBP was only detected as traces (LOD> traces
<LOQ) in bottled water extracts. 2,4-dtBP is a by-product of antioxidants (Irgafos 168 and Irgafos 1010) generated when food packaging is exposed to oxidation conditions such as high temperatures
(Alin & Hakkarainen, 2011). Therefore, the mean temperature
(27.6 °C/day) reached by bottled waters during 10 days of sunlight
exposure (see Table 1) may be not high enough to start the oxidation reaction. Furthermore, the photodegradation of this phenolic
compound in water cannot be ruled out (Neamtu & Frimmel,
2006).
No other compounds were detected in this study, but unpredictable NIAS may be present in PET-bottled water extracts. Indeed,
unidentified NIAS represent more than half of the compounds present in the migrates (Grob, Biedermann, Scherbaum, Roth, & Rieger,
2006). Moreover, chemical analysis could not always explain the
toxicity observed (Honkalampi-Hämäläinen et al., 2010), suggesting that testing the whole migrate therefore offers an opportunity
to reduce uncertainty (Muncke, 2011). Identification of NIAS is a
great challenge and a difficult task. Sample treatment procedures
and instrumental analysis are never exhaustive for all substances
present in migrates. In addition, low concentrations of migrants
and complex mixtures with co-eluting compounds makes the possibility of unequivocal analytical identification and confirmation
difficult (Nerín et al., 2013).
In this study, the potential toxicity due to unpredictable NIAS
and/or chemical mixtures present in bottled water extracts was
checked using two relevant toxicological endpoints (cyto/genotoxicity and endocrine disruption potential). Bottled water extracts
were neither mutagenic, nor cyto/genotoxic for S. typhimurium
strains (TA98 and TA100) and for the HepG2 cells, respectively.
This is not in accordance with De Fusco, Monarca, Biscardi,
Pasquini, and Fatigoni (1990), who reported weak mutagenic activity (only for TA98 strains with S9 mix) in PET-bottled water
extracts following exposure to daylight. These results were later
not confirmed by the same researchers (Monarca et al., 1994),
probably due to different types of PET bottles and mineral waters
used in the different studies. More recently, mutagenic activity
was observed in mineral water exposed to sunlight for 2 months
using the Ames fluctuation test (Ubomba-Jaswa et al., 2010). However, this response was not confirmed for longer periods of storage
(>3 months), suggesting that genotoxic compounds undergo degradation in non-genotoxic substances.
With plants models, contradictory results and conclusions using
the Allium Cepa test were reported after sunlight exposure. While a
2-fold increase in chromosomal aberrations was showed by
Evandri, Tucci, and Bolle (2000) and Corneanu et al. (2010) attributed the chromosomal mutations observed to the mineral salt content of the water as well as the technology used for manufacturing
the PET bottles. Our results are not in accordance with these previous studies due to differences in the bioassays, cell models (plants,
human cell lines, etc.) and conditions used to perform them. In
contrast with ecotoxicology, plants systems are not considered as
primary screening tools for extrapolation to mammalian systems
(EFSA, 2011). In addition, different sample preparations (cartridges,
solvent polarities, etc.) could give different compound extraction
efficiencies as demonstrated by Wagner and Oehlmann (2011).
Several authors suggest that PET bottles may yield endocrine
disruptor chemicals under regular conditions of use, such as
long-term storage, high temperatures and exposure to sunlight
(Sax, 2010; Wagner & Oehlmann, 2011; Yang et al., 2011). In this
study, after 10 days of sunlight exposure, no estrogenic or antiandrogenic activity was detected in PET- and glass-bottled water
extracts using HepG2 and MDA-MB453-kb2 cells, respectively.
5. Conclusions
The effect of sunlight exposure on chemicals release into PETbottled waters and the potential hazard of water extracts were
investigated using in vitro bioassays. The migration of aldehydes
and Sb into ultrapure waters increased with sunlight especially
after 10 days of exposure without exceeding the current specific
migration limits set in Regulation No. 10/2011. However, an offflavour can occur due to the level of formaldehyde in carbonated
waters after 10 days in sunlight. In carbonated mineral water,
carbon dioxide contributed to migration more than sunlight. Water
extracts did not induce any cyto-genotoxic or endocrine-disruption
activity in the bioassays under our experimental conditions. Chemical analysis and global approaches using bioassays are complementary tools to identify the potential toxic effects due to
unpredictable NIAS and/or chemical mixtures.
Acknowledgements
This research was financed 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 Bandol Weathering Station (SEVN) and the
Water Chemistry Department of ANSES’ Nancy Laboratory for
Hydrology for their excellent technical assistance. The authors
are 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, in
the online version, at http://dx.doi.org/10.1016/j.foodchem.2014.
04.020.
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