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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
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 * * *
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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-
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66 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
(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).
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C. Bach et al. / Food Chemistry 162 (2014) 63–71 67
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.
68 C. Bach et al. / Food Chemistry 162 (2014) 63–71
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.
C. Bach et al. / Food Chemistry 162 (2014) 63–71 69
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
-
70 C. Bach et al. / Food Chemistry 162 (2014) 63–71
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.
References
Alin, J., & Hakkarainen, M. (2011). microwave heating causes
rapid degradation ofantioxidants in polypropylene packaging,
leading to greatly increased specificmigration to food simulants as
shown by ESI–MS and GC–MS. Journal ofAgricultural and Food
Chemistry, 59(10), 5418–5427.
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 andtoxicological assessments of drinking water
stored in polyethyleneterephthalate (PET) bottles: A source of
controversy reviewed. Water Research,46(3), 571–583.
http://dx.doi.org/10.1016/j.foodchem.2014.04.020http://dx.doi.org/10.1016/j.foodchem.2014.04.020http://refhub.elsevier.com/S0308-8146(14)00565-2/h0005http://refhub.elsevier.com/S0308-8146(14)00565-2/h0005http://refhub.elsevier.com/S0308-8146(14)00565-2/h0005http://refhub.elsevier.com/S0308-8146(14)00565-2/h0005http://refhub.elsevier.com/S0308-8146(14)00565-2/h0010http://refhub.elsevier.com/S0308-8146(14)00565-2/h0010http://refhub.elsevier.com/S0308-8146(14)00565-2/h0020http://refhub.elsevier.com/S0308-8146(14)00565-2/h0020http://refhub.elsevier.com/S0308-8146(14)00565-2/h0020http://refhub.elsevier.com/S0308-8146(14)00565-2/h0020
-
C. Bach et al. / Food Chemistry 162 (2014) 63–71 71
Bach, C., Dauchy, D., Severin, I., Munoz, J.-F., Etienne, S.,
& Chagnon, M.-C. (2013).Effect of temperature on the release of
intentionally and non-intentionallyadded substances from
polyethylene terephthalate (PET) bottles into water:Chemical
analysis and potential toxicity. Food Chemistry, 139, 672–680.
Casajuana, N., & Lacorte, S. (2003). Presence and release of
phthalic esters and otherendocrine disrupting compounds in drinking
water. Chromatographia, 57(9–10),649–655.
Cheng, X., Shi, H., Adams, C. D., & Ma, Y. (2010).
Assessment of metalcontaminations leaching out from recycling
plastic bottles upon treatments.Environmental Science and Pollution
Research, 17(7), 1323–1330.
Corneanu, M., Corneanu, G., Jurescu, N., & Toptan, C.
(2010). Evaluation of thegenotoxicity of water bottled in PET
recipients. Environmental Engineering andManagement Journal, 9(11),
1531–1537.
Dabrowska, A., Borcz, A., & Nawrocki, J. (2003). Aldehyde
contamination of mineralwater stored in PET bottles. Food Additives
and Contaminants, 20(12),1170–1177.
De Fusco, R., Monarca, S., Biscardi, D., Pasquini, R., &
Fatigoni, C. (1990). Leaching ofmutagens into mineral water from
polyethyleneterephthalate bottles. Science ofthe Total Environment,
90, 241–248.
Dévier, M.-H., Le Menach, K., Viglino, L., Di Gioia, L.,
Lachassagne, P., & Budzinski, H.(2013). Ultra-trace analysis of
hormones, pharmaceutical substances,alkylphenols and phthalates in
two French natural mineral waters. Science ofthe Total Environment,
443, 621–623.
EFSA (2011). Scientific opinion on genotoxicity testing
strategies applicable to foodand feed safety assessment. EFSA
Journal, 9(9), 2379. 2368 pp.
EFSA (2012). Scientific Opinion on the safety evaluation of the
substance, titaniumnitride, nanoparticles, for us in food contact
materials. EFSA Journal, 10(3),2641–2649.
EU (2004). Regulation No. 1935/2004 of the European Parliament
and of the councilof 27 October 2004 on materials intended to come
into contact with food andrepealing directives 80/590/EEC and
89/109/EEC.
EU (2011). Commission regulation (EU) No. 10/2011 of 14 January
2011 on plasticmaterials and articles intended to come in contact
with food.
Evandri, M. G., Tucci, P., & Bolle, P. (2000). Toxicological
evaluation of commercialmineral water bottled in polyethylene
terephthalate: A cytogenetic approachwith Allium cepa. Food
Additives and Contaminants, 17(12), 1037–1045.
Grob, K., Biedermann, M., Scherbaum, G., Roth, M., & Rieger,
K. (2006). Foodcontamination with organic materials in perspective:
Packaging materials asthe largest and least controlled source? A
view focusing on the Europeansituation. Critical Reviews in Food
Science and Nutrition, 46(7), 529–536.
Honkalampi-Hämäläinen, U., Bradley, E. L., Castle, L., Severin,
I., Dahbi, L., Dahlman,O., et al. (2010). Safety evaluation of food
contact paper and board usingchemical tests and in vitro bioassays:
role of known and unknown substances.Food Additives &
Contaminants, 27(3), 406–415.
ICCVAM (2003). Evaluation of In vitro test methods for detection
potential endocrinedisruptors: Estrogen receptor and androgen
receptor binding and transcriptionalactivation assays. May, 2003.
NIH Pub 03-4503. Available:
http://iccvam.niehs.nih.gov/docs/endo_docs/edfinalrpt0503/edfinrpt.pdf
[accessed 7July 2012].
ISO (2003). ISO 17294-2. Water quality. Application of
inductively coupled plasmamass spectrometry (ICP-MS). Part 2:
Determination of 62 elements.
ISO (2009). International Standard ISO 877:2099. Plastics –
Methods of exposure tosolar radiation – Part 2: Direct weathering
and exposure behing window glass.
JORF (2011). Arrêté du 28 décembre 2010 modifiant l’arrêté du 14
mars 2007 relatifaux critères de qualité des eaux conditionnées,
aux traitements et mentionsd’étiquetage particuliers des eaux
minérales naturelle et des eaux de sourceconditionnées ainsi que de
l’eau minérale naturelle distribué en buvettepublique. Ministère du
travail, de l’emploi et de la santé. Journal Officiel de
laRépublique Française, Texte 42.
Keresztes, S., Tatár, E., Mihucz, V. G., Virág, I., Majdik, C.,
& Záray, G. (2009). Leachingof antimony from polyethylene
terephthalate (PET) bottles into mineral water.Science of the Total
Environment, 407(16), 4731–4735.
Kirsch-Volders, M., Sofuni, T., Aardema, M., Albertini, S.,
Eastmond, D., Fenech, M.,et al. (2000). Report from the In Vitro
Micronucleus Assay Working Group.Environmental and Molecular
Mutagenesis, 35(3), 167–172.
Kortenkamp, A. (2007). Ten years of mixing cocktails: a review
of combinationeffects of endocrine-disrupting chemical.
Environmental Health Perspectives,115(1), 98–105.
Leivadara, S. V., Nikolaou, A. D., & Lekkas, T. D. (2008).
Determination of organiccompounds in bottled waters. Food
Chemistry, 108(1), 277–286.
Maron, D. M., & Ames, B. N. (1983). Revised methods for the
Salmonellamutagenicity test. Mutation Research, 113(3–4),
173–215.
Monarca, S., De Fusco, R., Biscardi, D., De Feo, V., Pasquini,
R., Fatigoni, C., et al.(1994). Studies of migration of potentially
genotoxic compounds into waterstored in pet bottles. Food and
Chemical Toxicology, 32(9), 783–788.
Muncke, J. (2011). Endocrine disrupting chemicals and other
substances of conerne infood contact materials: and updated review
of exposure effect and risk assessment.Journal of Steroid
Biochemistry and Molecular Biology, 127(1–2), 118–127.
Mutsuga, M., Kawamura, Y., Sugita-Konishi, Y., Hara-Kudo, Y.,
Takatori, K., &Tanamoto, K. (2006). Migration of formaldehyde
and acetaldehyde into mineralwater in polyethylene terephthalate
(PET) bottles. Food Additives andContaminants, 23(2), 212–218.
Neamtu, M., & Frimmel, F. H. (2006). Photodegradation of
endocrine disruptingchemical nonylphenol by simulated solar
UV-irradiation. Science of the TotalEnvironment, 369(1–3),
295–306.
Nerín, C., Alfaro, P., Aznar, M., & Domeño, C. (2013). The
challenge of identifyingnon-intentionally added substances from
food packaging materials: A review.Analytica Chimica Acta, 775,
14–24.
OECD (2010). Guideline for the testing of chemicals. No 487: In
Vitro MammalianCell Micronucleus Test.
http://iccvam.niehs.nih.gov/SuppDocs/FedDocs/OECD/OECD-TG487.pdf.
Sax, L. (2010). Polyethylene terephthalate May yield endocrine
disruptors.Environmental Health Perspectives, 118(4), 445–448.
Schmid, P., Kohler, M., Meierhofer, R., Luzi, S., & Wegelin,
M. (2008). Does the reuseof PET bottles during solar water
disinfection pose a health risk due to themigration of plasticisers
and other chemicals into the water? Water Research,42(20),
5054–5060.
Severin, I., Jondeau, A., Dahbi, L., & Chagnon, M. C.
(2005). 2,4-Diaminotoluene (2,4-DAT)-induced DNA damage. DNA repair
and micronucleus formation in thehuman hepatoma cell line HepG2.
Toxicology, 213(1–2), 138–146.
Stroheker, T., Picard, K., Lhuguenot, J.-C., Canivenc-Lavier,
M.-C., & Chagnon, M.-C.(2004). Steroid activities comparison of
natural and food wrap compounds inhuman breast cancer cell lines.
Food and Chemical Toxicology, 46(6), 887–897.
Strube, A., Buettner, A., & Groetzinger, C. (2009).
Characterization and identificationof a plastic-like off-odor in
mineral water. Water Science and Technology: WaterSupply, 9,
299–309.
Ubomba-Jaswa, E., Fernandez-Ibanez, P., & McGuigan, K. G.
(2010). A preliminaryAmes fluctuation assay assessment of the
genotoxicity of drinking water thathas been solar disenfected in
polyethylene terephthalate (PET) bottles. Journalof Water and
Health, 8(4), 712–719.
Wagner, M., & Oehlmann, J. (2011). Endocrine disruptors in
bottled mineral water:Estrogenic activity in the E-Screen. Journal
of Steroid Biochemistry and MolecularBiology, 127(1–2),
128–135.
Wegelin, M., Canonica, S., Alder, C., Marazuela, D., Suter, M.
J. F., Bucheli, T. D., et al.(2001). Does sunlight change the
material and content of polyethyleneterephthalate (pet) bottles?
Journal of Water Supply: Research and Technology -AQUA, 50(3),
125–133.
Wilson, V. S., Bobsein, K., Lambright, C. R., & Gray, L. E.
(2002). A novel cell line,MDA-kb2, that stably expresses an
androgen- and glucocorticoid-responsivereporter for the detection
of hormone receptor agonist and antagonist.Toxicological Science,
66(1), 69–81.
Yang, C. Z., Yaniger, S. I., Jordan, V. C., Klein, D. J., &
Bittner, G. D. (2011). Most PlasticProducts Release Estrogenic
Chemicals: A Potential Health Problem That Can BeSolved.
Environmental Health Perspectives, 119(7), 982–996.
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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