Environmental Pollution xxx (2010) 1e6
Contents lists available at ScienceDirect
Environmental Pollution
journal homepage: www.elsevier.com/locate/envpol
A rapidly equilibrating, thin film, passive water sampler for organic
contaminants; characterization and field testing
Tiffany St. George b, e, Penny Vlahos a, b, *, Tom Harner c, Paul Helm d, Bryony Wilford c
a
Department of Chemistry, University of Connecticut, 55 Eagleville Road, Storrs, CT 06269, USA
Department of Marine Science, University of Connecticut, 1080 Shennecossett Road, Groton, CT 06340, USA
c
Science and Technology Branch, Environment Canada, 4905 Dufferin Street, Toronto, Ontario M3H 5T4, Canada
d
Environmental Monitoring & Reporting Branch, Ontario Ministry of the Environment, 125 Resources Rd, Toronto, Ontario M9P 3V6, Canada
e
Department of Science, United States Coast Guard Academy, 27 Mohegan Ave., New London, CT 06320, USA
b
An ethylene vinyl acetate (EVA), thin-film passive sampler for the detection of organic compounds in marine environments is calibrated
and field tested.
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 26 July 2010
Received in revised form
8 October 2010
Accepted 13 October 2010
Improving methods for assessing the spatial and temporal resolution of organic compound concentrations in
marine environments is important to the sustainable management of our coastal systems. Here we evaluate
the use of ethylene vinyl acetate (EVA) as a candidate polymer for thin-film passive sampling in waters of
marine environments. Log KEVA W partition coefficients correlate well (r2 ¼ 0.87) with Log KOW values for
selected pesticides and polychlorinated biphenyls (PCBs) where Log KEVA W ¼ 1.04 Log KOW þ 0.22. EVA is
a suitable polymer for passive sampling due to both its high affinity for organic compounds and its ease of
coating at sub-micron film thicknesses on various substrates. Twelve-day field deployments were effective in
detecting target compounds with good precision making EVA a potential multi-media fugacity meter.
Published by Elsevier Ltd.
Keywords:
Passive sampler
Ethylene vinyl acetate
Pesticides
PCBs
Chemical monitoring
Marine
1. Introduction
Contamination of marine and freshwater systems is of global
environmental concern; however, long-term monitoring of these
compounds remains a challenge (Namiesnik et al., 2005; Vrana
et al., 2005). Coastal observation networks are being introduced
along European and North American coastlines in order to provide
continuous monitoring of environmental parameters. Currently
these networks are restricted in the amount of chemical monitoring that can be conducted due to limitations in chemical sensor
technology (Stuer-Lauridsen, 2005). Expanding coastal observation
networks to include chemical sensors for organic contaminants in
the local environment would provide essential data to both scientists and policy makers. Currently, two general strategies exist for
long-term monitoring of organic pollutants in aqueous environments: discrete sampling and time-integrated or equilibrium
passive sampling (Zhao et al., 2005).
* Corresponding author.
E-mail address: penny.vlahos@uconn.edu (P. Vlahos).
Discrete sampling in the water phase provides a snapshot of the
concentration in the environment at the time of sampling (StuerLauridsen, 2005). This approach suffers from the limitations of
sampling artifacts, temporal deficiencies, and high costs. One
solution to this problem is to increase the sampling frequency or to
install sampling systems that automatically acquire a distinct
number of water samples in any given period (Mayer et al., 2003;
Vrana et al., 2005). However, this may be cumbersome and
prohibitively expensive when implemented in long-term water
quality monitoring programs.
A range of passive samplers have been successfully applied to
marine systems. They are based on the free flow of an analyte from
the sampled medium to a receiving phase in a sampling device and
collect target compounds in situ without disrupting the bulk solution. They may represent a time-averaged concentration such as
semi-permeable membrane devices (SPMDs) (Mayer et al., 2003;
Stuer-Lauridsen, 2005; Vrana et al., 2005) in which a compound
accumulates into the sampler, often non-linearly, and the final
concentration in the sampler is averaged over the deployment time
to estimate an average concentration of the source. These samplers
often mirror uptake in organisms and can be related to bioconcentration. Equilibrium samplers contain a sorbent such as
0269-7491/$ e see front matter Published by Elsevier Ltd.
doi:10.1016/j.envpol.2010.10.030
Please cite this article in press as: St. George, T., et al., A rapidly equilibrating, thin film, passive water sampler for organic contaminants;...,
Environmental Pollution (2010), doi:10.1016/j.envpol.2010.10.030
2
T. St. George et al. / Environmental Pollution xxx (2010) 1e6
a thin piece of polyethylene (Adams et al., 2007) or a thin film of
polymer coated onto a substrate that reaches equilibrium with the
surrounding medium (Wilcockson and Gobas, 2001) and the
ambient concentration can be calculated based on an equilibrium
partition constant. Both samplers provide insight to the environment. When used together the samplers should yield similar estimates of concentrations in the medium provided 1) they have been
well parameterized and 2) the ambient concentrations are steady
over the deployment period. If ambient concentrations differ over
the deployment periods or there are episodic fluctuations, the
concentration estimates may diverge. The divergence is useful in
assessing that the conditions above have or have not been met.
Equilibrium samplers are particularly useful in that, when calibrated, they can function as fugacity meters in the environment to
help determine both the concentration and flux of target
compounds in various media (Meloche et al., 2009). The calibration
of thin-film devices is simpler than their time-averaged counterparts and the film may be adjusted (surface area to volume ratio)
and targeted to different compounds over a given deployment
period. This approach has the potential to become a reliable and
cost-effective tool that may be utilized in ongoing efforts to
improve the detection of priority pollutants (Allan et al., 2006).
Lohmann and Muir (2010) have advocated the need for global scale
monitoring strategies for organic pollutants in aquatic environments to meet national and international monitoring obligations
(e.g. Stockholm Convention on Persistent Organic Pollutants).
This study reports on the calibration of an equilibrium-type
passive sampler for water, comprising of micron-thick layers of the
copolymer ethylene vinyl acetate (EVA) (Fig. 1). EVA was first
proposed as a fugacity sampler for organic tissue by Wilcockson and
Gobas (2001) and then adapted as an air-side fugacity meter by
Harner et al. (2003). This study extends the application of EVA as
a water-side fugacity meter to expand the application of this
medium (EVA) and test its potential as a multi-media fugacity
meter. EVA is elastomeric in softness and flexibility but may be
processed as other thermoplastics. It has good clarity, a utilization
temperature range of 60 to 55 C, stress-crack resistance (elongation 800%) such that it can be used in deep sea environments at
high pressures, is water proof (0.07% water saturation) and is
resistant to UV radiation thus maintaining its integrity in surface
waters. EVA is non-toxic and commonly used in drug delivery, hot
glue sticks and plastic wraps. Equilibrium samplers based on polyethylene have been recently tested for marine applications (Adams
et al., 2007) and are particularly effective for hydrophobic
compounds. The advantages of EVA include that it can be readily
spiked with internal reference compounds at exact concentrations
and can then be coated onto a variety of surfaces (Harner et al.,
2003; Farrar et al., 2005; Wu et al., 2008b). EVA may be readily
coated at various film thicknesses at the sub-micron range to optimize sampler uptake. It is expected that EVA has an expanded range
of target compounds that bridges the current gap in aquatic passive
sampling due to its polar acetate group, while providing a sufficient
non-polar surface area to match the chemical affinity of existing
samplers.
Fig. 1. Base unit for ethylene vinyl acetate (EVA) copolymer.
EVA-water partition coefficients are reported for a suite of
pesticides and polychlorinated biphenyls (PCBs) and a coastal
deployment using the derived partition constants for the detection
of currently used pesticides in natural waters is presented.
2. Theory
The difference in chemical potentials of an analyte across
separate phases results in its net flow from one medium to another.
For environmental marine monitoring, this occurs from seawater to
the polymer used in the passive sampler (Huckins et al., 1990). Net
movement continues until thermodynamic equilibrium is achieved
or the sampling period is halted (Vrana et al., 2005). The exchange
kinetics between sampler and water can be described by a firstorder one-compartment model:
CS ðtÞ ¼ CW
k1
1
k2
e
k2 t
(1)
CS(t) is the contaminant concentration in the sampler polymer
as a function of time, t, CW is the contaminant concentration in the
sampled medium (i.e., seawater), and k1 and k2 are the uptake and
offload rate constants respectively (Allan et al., 2006). Passive
samplers follow one of two phases in the accumulation of organic
pollutants during field deployments. These phases are kinetic or
equilibrium and can be shifted by altering the sampler design (i.e.
surface area and film thickness). The time to reach equilibrium for
the EVA sampler is a function of the partition coefficient Ksampler,medium and the uptake rate k1 of the compound. Uptake follows
a standard saturation curve in which the time to reach 90% of
equilibrium can be expressed as (Mayer et al., 2003):
t90% ¼
k1
k2
ln 10
ln 10
¼ Ksampler;medium
k2
k1
(2)
In the equilibrium phase, as ð1 e k2 t Þ approaches 1, and
¼ Ksampler;medium Equation (1) can be reduced to:
CW ¼
CS
KEVA
(3)
W
where KEVA W is the partition coefficient of the target analyte
between EVA and water.
3. Methods
The EVA sampler is based on the principles used to monitor semivolatile organic
compounds (SVOC) in the atmosphere as an alternative to high volume air sampling
(Harner et al., 2003). The design consists of a thin film of EVA (w1 mm) (Elvax 40W,
DuPont) on a substrate. The sampler versatility was recognized and demonstrated as
it could be used to sample in either the kinetic or equilibrium phases simply by
varying the thickness of the polymer coating (Harner et al., 2003; Farrar et al., 2005).
3.1. Determination of partition coefficients
The generator column method was used to measure KEVA W for a group of
pesticides including several legacy organochlorine pesticides (OCPs), currently used
pesticides (CUPs) and PCBs (see Fig. S1, Supplementary information). This is analogous to the methods used by Wu et al. (2008b) for the experimental determination
of KEVA A for PCBs in air. In this approach, water was passed through and equilibrated with a column of 3 mm glass beads coated with EVA that had been spiked
with target analytes.
The EVA solution was made by dissolving 16 g EVA in 200 mL DCM. This solution
was then spiked with the target compounds (Table 1). Glass beads were coated by
immersion in the EVA solution for approximately 1 min and decanting excess
solution. The ‘wet’ beads were then immediately transferred to a large stainless steel
bowl and swirled rapidly for 1e2 min to allow excess DCM to evaporate and to
prevent beads from clumping together (Wu et al., 2008a,b). The coated beads were
loaded to the 20 mL level of a 10 mm i.d. glass column. A sub-set of the beads was
retained for the determination of analyte concentration in the EVA (Cs)
(see Supplementary information).
Please cite this article in press as: St. George, T., et al., A rapidly equilibrating, thin film, passive water sampler for organic contaminants;...,
Environmental Pollution (2010), doi:10.1016/j.envpol.2010.10.030
T. St. George et al. / Environmental Pollution xxx (2010) 1e6
Table 1
Log KOW values of target compounds and experimentally derived values of KEVA
using the generator column method.
Compound
Log Kow
Log KEVA
Simazine
Carbofuran
Atrazine
Metolachlor
Alachlor
Phorate
a-HCH
Diazinon
Disulfoton
Terbufos
Trifluralin
Chlorothalonil
Metribuzin
Malathion
Chlorpyrifos
Dacthal
Pendimethalin
PCB 3
PCB 15
PCB 18
2.2b
2.3e
2.8f
3.1c
3.5e
3.6g
3.8g
3.8d
4.0d
4.5e
5.3c
2.9g
1.7a
2.4e
5.0d
4.4e
5.2g
4.5e
5.3e
5.6e
2.7
2.6
2.9
3.0
3.1
4.0
4.4
3.8
3.2
4.4
6.1
3.6
2.5
3.1
5.5
4.8
5.6
5.4
6.1
6.3
W
W
KOW values were selected based on recommended values of the Canadian National
Committee for CODATA(CNC/CODATA).
a
Draber et al. (1969).
b
Liu and Qian (1995).
c
Kenaga and Goring (1980).
d
Bowman and Sans (1983).
e
Hansch and Leo (1987).
f
Finizio et al. (1991).
g
Hansch et al. (1995).
3
during August 2006. This is an urbanized and industrialized tidal estuary along Long
Island Sound located in southeast Connecticut (CT), USA. The Estuary is approximately 20 km long with the head of the river located in Norwich, CT. The Yantic and
Shetucket Rivers comprise the major freshwater inflow to the Estuary. The watershed encompasses 3800 km2. There are numerous agricultural areas within the
watershed and area of the estuary, making it a good test location for investigating
the application of the thin-film passive samplers for legacy and current-use
pesticides.
The samplers (n ¼ 3 per station) were suspended on a stainless steel frame
approximately 50 cm above the bottom sediments and left in situ for 12 days (Fig. S2
Supplementary information). Equilibration times were determined in the lab and
ranged from 1 to 12 d for compounds with Log Kow values between 1 and 6
(St. George, 2008). Field blanks were exposed on ship deck until the sampler entered
the water. The blanks were then wrapped in pre-cleaned foil, placed in an airtight
container and stored at 4 C until ready for extraction and analysis. Sampler
recovery was conducted by divers at the end of the 12 d deployment period. Once
retrieved, the samplers were handled as described above. Conductivity, temperature, and depth (CTD) profiles were recorded at each station using a Seabird CTD.
Bottom water salinity and temperature ranges varied only slightly between stations.
Bottom water depths were 10e12 m and temperatures ranged from 18 to 20 C.
Samplers were rinsed with de-ionized water and placed in a 125 mL sealed
vessel and soaked in 120 mL analytical grade methanol (Fischer Scientific) for a 24 h
period before pouring the extract into a pre-rinsed 500 mL flask. This process was
repeated a second time and the extracts combined before being reduced to w2 mL
via rotary evaporator before transferring to a pre-rinsed centrifuge tube through
3 cm of sodium sulfate. Extracts were evaporated to approximately 0.5 mL under
a gentle stream of nitrogen and then solvent exchanged to ethyl acetate followed by
isooctane. Upon completion of solvent exchange, 10 mL of Mirex (10 ng mL 1) was
added as an internal standard. The internal standard is used to verify and quantify
instrument responses. Extracts were placed in a centrifuge at 5000 RPM for 3 min to
remove any EVA or sodium sulfate that may have remained in the sample. The
supernatant was transferred into a GC vial and the volume further reduced to 1 mL
for analysis.
4. Analysis
Ultra-high purity water was introduced to the top of the column at a rate of
5 mL min 1 until the beads were completely covered. The same delivery rate was
used during the collection of twelve 10 mL samples to assess Cw and to ensure that
equilibrium was achieved in the generator column. Further details are provided in
the Supplementary information. Experimental KEVA W values were calculated using
Cw, Cs and substituting into Equation (3). All 10 mL water samples were extracted by
liquideliquid extraction with 3 subsequent 5 mL DCM aliquots. After DCM addition,
samples were agitated using a vortex mixer for 1 min before centrifugation to
separate the phases and removal of the lower DCM phase into a clean boiling tube.
After the third DCM rinse, the composite DCM fraction was reduced under a gentle
stream of nitrogen and solvent exchanged into isooctane for analysis.
3.2. Preparation of samplers for field deployment
A coating solution of EVA was created by dissolving 2 g of EVA pellets into
100 mL of analytical grade DCM. The solution was stirred for 2 h ensuring that the
EVA was completely dissolved. Pre-combusted (420 C, 6 h) Whatman glass fiber
filters (12.5 cm i.d., 0.7 mm nominal retention size) were used as the coated
substrates. These were baked at 420 C for 6 h, allowed to cool to room temperature
and weighed. Substrates were then dipped into the coating solution for 5 s. Upon
removal, the DCM was evaporated, leaving a thin coating of EVA on the substrate
with an approximate surface area of over 245 cm2. The dry filters were weighed to
determine the mass of the EVA. Average mass of the EVA on the sampler was
0.40 g 0.01 g (density of EVA, rEVA ¼ 0.93 g mL 1) which corresponds to an average
film thickness of approximately 18 mm (based on the planar surface area of the filter).
It should be noted that the GFFs are a fiber mesh and therefore the true film
thickness along the mesh will be less than 18 mm due to the larger effective surface
area of the mesh. GFFs were used (rather than beads) as a direct comparison to
previous studies in air (Wu et al., 2008a,b) and to simplify the samplerewater
boundary to avoid diffusion limitations in a more quiescent medium. Filters were
then inserted into a solvent-rinsed stainless steel cage, wrapped in aluminum foil,
placed in an airtight container and frozen until deployment. During this procedure,
care was taken to ensure that the exposure of the samplers to ambient air was
minimized (typically less than 5 min) in order to limit contamination. Titanium
plates were also coated in a separate study (St. George, 2008) and showed no
difference in EVA partitioning due to substrate.
3.3. Field deployment of EVA samplers
EVA samplers were deployed in triplicate at three stations (Station 1: 41 26.60 Ne072 -05.90 W, Station 2: 41 -22.90 Ne072 -05.30 W, Station 3: 41 19.70 Ne072 -04.90 W) eastward of the channel along the Thames River Estuary (TRE)
Extracts from laboratory samples (generator column experiments) and field samples were analyzed for current-use pesticides
as per Yao et al. (2006). Briefly, phorate, a-HCH, dazoment, simazine, carbofuran, atrazine, terbufos, diazinon, disulfoton, alachlor,
and metolachlor were analyzed by gas chromatography mass
spectroscopy (GCeMS) in electron-ionization (EI) mode. Other
CUPs: trifluralin, dimethoate, chlorothalonil, metribuzin, malathion, chlorpyrifos, dacthal and pendimethalin were analyzed by
GCeMS in negative chemical ionization (NCI) mode. Legacy pesticides a-, b-, g-, and d-HCH, HEPT, HEPX, aldrin, dieldrin, TC, TN, CC,
Endosulfan I and II, and endosulfan sulfate, the isomers of DDT and
their breakdown products were also analyzed by NCI. Generator
column tests were also analyzed for PCB congeners 3, 8, 15,
and 18 in EI mode. For both types of analysis a J&W DB-5 30 m
column (J&W Scientific, Rancho Cordova, CA), i.d. 0.25 mm, 0.25 mm
film thickness was used, operated with helium as the carrier gas,
connected to a HewlettePackard 5973 MS (in NCI mode, methane
was used as the reagent gas). A 2 mL sample was injected in splitless
mode, with the split opened after 1 min. Samples were quantified
by comparison to a prepared set of standards of known concentration. Peaks were quantified when S/N 3, and when the
quantifier/qualifier ratio was within 15% of the standard (Yao et al.,
2006).
5. Results and discussion
5.1. Quality assurance/control
The limit of detection (LOD) was equal to the mean blank level
plus three standard deviations of the mean. The instrument
detection limits (DL) for the samples were calculated by extrapolation from the lowest level calibration standard to the point where
the signal to noise ratio was equal to three. Calibration experiment
blank levels were <10% of sample concentrations. Method
Please cite this article in press as: St. George, T., et al., A rapidly equilibrating, thin film, passive water sampler for organic contaminants;...,
Environmental Pollution (2010), doi:10.1016/j.envpol.2010.10.030
4
T. St. George et al. / Environmental Pollution xxx (2010) 1e6
recoveries ranged from 85 to 102%. Reported concentrations were
blank and recovery-corrected. Field sample blank levels were
below detection for all pesticides with the exception of Trifluralin;
that was as high as 7.7 pg L 1. All stations showed concentrations of
Trifluralin above the LOD. The standard deviation of the blanks
(n ¼ 3) was less than 1 pg L 1 (or w20%).
5.2. KEVA
W
determinations
Results of equilibration tests showed that the generator column
came to equilibrium for all compounds with Log KOW > 3.5.
Concentrations on beads for these compounds remained relatively
constant. In these instances the average value of CS and CW was
used to determine KEVA W. For compounds with lower KOW values
(based on results for alachlor and metolachlor), the EVA-coated
beads became depleted of analyte during the time-course of the
measurements. In these cases, partition coefficient determinations
were based on EVA concentrations adjusted for the corresponding
loss. The Log KEVA W values calculated from these experiments are
summarized in Table 1.
A plot was constructed and used to define Log KEVA W partitioning coefficients for the 2 classes of compounds examined
(Equation (4)). Strong correlations were observed (r2 ¼ 0.87;
p < 0.005) such that EVA could be an effective passive sampling
material with an affinity for organic compounds that is in consistent with that of octanol for the compounds tested. The Log KEVA W
versus Log KOW plot yielded the following relationship:
Log KEVA
W
¼ 1:04 Log KOW þ 0:22
(4)
Calibrations were in good agreement both within and among
sampled pesticides and PCBs. Results indicate the effectiveness of
EVA as a sampling medium for hydrophobic compounds. The
relationship shows that EVA has an affinity for organic compounds
slightly greater than that of octanol and other polymers used in
equilibrium samplers (Adams et al., 2007) and possibly with
a broader target range. These observations are in agreement with
those of Golding et al. (2007, 2008) which also show the somewhat
higher solubility of organic compounds in EVA compared with
octanol. EVA allows both non-polar:non-polar van der Waals
interactions and polar interactions at the acetate ends resulting in
both stronger attractive forces and a wider range of target organics.
Though a large range of hydrophobicities are represented here it
would be useful to investigate additional compound classes and
compounds with an extended range of KOW values to test these
correlations further.
5.4. Calculation of CW in field tests
Experimental KEVA W values (Table 1) were applied to assess CW
in equilibrated TRE field samples using Equation (3). Equilibrium for
all target compounds was predicted to be approached within 12
days based on an 18 mm EVA film and a diffusivity in water on the
order of 0.01 cm s 1. This was also confirmed experimentally in
previous batch uptake experiments (Wilford et al., 2006; St. George,
2008). The mass of the target analyte detected during GCeMS
analysis and the volume of EVA on the sampler was used to determine CS, which was blank corrected. Blank corrections in equilibrium sampling are conceptually difficult because exposure prior to
deployment is not expected to linger since the sampler re-equilibrates in the field. Here we assume that any residual blank signals
are due to post-deployment exposure. This is a conservative
approach and fortunately blank concentrations were not a problem.
5.5. Field testing of EVA samplers
The purpose of the field trials was to assess the experimental
parameters and sample reproducibility. Results showed that over
the deployment period, sample integrity was maintained and
minimal visible biofouling was observed. Of the nineteen pesticides
that were targeted, six were detected in the TRE. Compound
concentrations are shown in Fig. 2 and are divided into two groups
based on their ranges (ng L 1 and pg L 1).
Highest average EVA-derived water concentrations across the
three stations in the estuary were for metribuzin and atrazine, 110
and 72 ng L 1, respectively. Atrazine and metribuzin are triazine
herbicides and are commonly used to control broadleaf and grassy
weeds in vegetable and field crops as well as non-cropped industrial lands (Cox, 1997). Chlorothalonil was detected at 6.8 ng L 1.
This is one of the most common fungicides in agricultural and
household uses with lawn treatment accounting for approximately
one third of its use (Cox, 1997). Water concentrations for metribuzin and atrazine reported here are in the range of reported values
from water quality studies for the same study area (Garabedian
et al., 1998), reflecting the ongoing use and presence of these
pesticides in the estuary. In comparison with pesticide concentrations from other eastern US estuaries, the results from this study
are slightly higher than mean concentrations of atrazine (48 ng L 1)
and chlorothalonil (2.7 ng L 1) measured in the C-111 canal system
and Florida Bay (Scott et al., 2002), but substantially lower than the
1.29 mg L 1 atrazine concentration detected in the Patuxent River by
McConnell et al. (2004). Concentrations of these analytes varied
5.3. Detection limits
The detection limits of the EVA sampler can be tuned to the
environmental concentrations based on the amount of EVA applied
and an adjustment of the surface area to maintain a thin film.
Equation (4) above provides the critical parameter for field applications and KEVA W can be derived using a known or estimated
KOW. The required mass of EVA MEVA can be determined using the
expected field concentration ranges (CW) and the instrumental
detection limits of the proposed analysis in grams or moles (Ntarget).
MEVA ¼ Ntarget rEVA ðKEVA
W CW Þ
1
(5)
3
where rEVA is the density of EVA (0.93 g cm ). For example, in this
application the sampler contains 0.40 0.01 g EVA and the
detection limits of the MS analysis are in the ng range. For this
configuration, compounds with a KEVA W on the order of 103 have
ambient detection limits in ng L 1 and those with a KEVA W of >105
can be detected in the <pg L 1 range.
Fig. 2. Passive sampler-derived water concentrations of selected CUPs in the Thames
River Estuary. Range and blank-corrected average values depicted are for a 12 d
sampler deployment of the EVA sampler at three stations. Water concentrations were
calculated using Equations (3) and (4); Log KOW value for a-Endosulfan ¼ 3.83 (Hansch
and Leo, 1985).
Please cite this article in press as: St. George, T., et al., A rapidly equilibrating, thin film, passive water sampler for organic contaminants;...,
Environmental Pollution (2010), doi:10.1016/j.envpol.2010.10.030
T. St. George et al. / Environmental Pollution xxx (2010) 1e6
only slightly between stations indicating relatively even distribution in the bottom waters of the sampled area.
The second group of CUPs detected in the pg L 1 range, included
pendimethalin, a-endosulfan, and trifluralin. Average concentrations of pendimethalin and trifluralin were 35 and 0.91 pg L 1
respectively. These herbicides are used in a variety of agricultural
and residential settings to control and prevent broadleaf weeds.
a-Endosulfan, detected at 63e120 pg L 1, is a broad-spectrum,
contact insecticide and is registered for use on a wide variety of
commercial agricultural settings. The concentration of a-endosulfan
at Station 1 was considerably higher than at the other two stations
that were closer to the mouth of the estuary. Pendimethalin and
trifluralin were detected previously in the study area; however, the
concentrations reported at that time were higher with maximum
concentrations of 100 ng L 1 and 20 ng L 1 respectively (Garabedian
et al., 1998).
The EVA sampler has a high affinity for organic compounds both
polar and non-polar and may bridge the gap between current
sampler polymers which tend to work optimally for a particular
range of hydrophobicities. EVA shows much promise as a multimedia fugacity sampler that may be applied in diverse environmental conditions.
6. Implications and perspectives
The EVA sampler described and successfully tested here has
potential for future applications in environmental research and
monitoring. The sampler is flexible in that EVA can be coated onto
a variety of substrates and thicknesses (down to less than
a micrometer if required). EVA has been applied successfully as an
equilibrium lipid and air sampler. Application of the sampler in
coupled airewater lipidewater systems would allow fugacity
gradients to be determined to provide an assessment of airewater
exchange. If KOA and KOW values are known, concentrations in air
and water can be calculated. By analogy, this approach can also be
extended to measure chemical fugacities across sedimentewater
interfaces.
Future work includes determining KEVA W for a broader range of
target analytes, assessing KEVA W as a function of temperature,
salinity, and applying depuration compounds (labeled analogs
spiked into the EVA prior to deployment) for field deployments in
water to confirm the equilibrium status of target analytes. Analogous to the successful use of depuration compounds in passive air
sampling programs, these compounds will enable the broader use
of this marine passive sampler in the kinetic phase by providing an
estimate of effective water sample volume. The EVA sampler meets
the call for a cost-effective and simple tool for measuring persistent
organic pollutants in aquatic environments (Lohmann and Muir,
2010). When used as a multi-media fugacity meter, the EVA
sampler can provide additional information on inter-media equilibrium status and fluxes (e.g. airewater, sedimentewater).
Acknowledgements
This work was supported by the University of Connecticut’s
Center for Environmental Science and Engineering, CT Sea Grant,
Environment Canada and the Ontario Ministry of the Environment.
The authors would like to thank Captain Turner Cabaniss, Adam
Houk, and the diving team at UConn for deployment and retrieval
of the samplers. TMS acknowledges the U.S. Coast Guard for
funding. The research described herein does not necessarily reflect
the position of the U. S. Coast Guard and no official endorsement
should be inferred.
5
Appendix. Supplementary information
Supplementary information related to this article can be found
online at doi:10.1016/j.envpol.2010.10.030.
References
Adams, R.G., Lohmann, R., Fernandez, L.A., Macfarlane, J.K., Gschwend, P.M., 2007.
Polyethylene devices: passive samplers for measuring dissolved hydrophobic
organic compounds in aquatic environments. Environmental Science and
Technology 41 (4), 1317e1323.
Allan, I.J., Vrana, B., Greenwood, R., Mills, G.A., Roig, B., Gonzalez, C., 2006. A
“toolbox” for biological and chemical monitoring requirements for the European Union’s Water Framework Directive. Talanta 69, 302e322.
Bowman, B.T., Sans, W.W., 1983. Determination of octanol-water partitioning
coefficients (Kow) of 61 organophosphorus and carbamate insecticides and
their relationship to respective water solubility (S) values. Journal of Environmental Science Health, Part B 18 (6), 667e683.
Cox, C., 1997. Chlorothalonil e fungicide factsheet. Journal of Pesticide Reform 17
(4), 14e21.
Draber, W., Buchel, K.H., Dickore, K., Trebst, A., Pistorius, E., 1969. Structure-activity
correlation of 1,2,4-triazinones, a new group of photosynthesis inhibitors. Prog.
Photosyn. Res., Proc. Int. Congr. 3, pp. 1789e1795.
Farrar, N.J., Harner, T.J., Sweetman, A.J., Jones, K.C., 2005. Field calibration of rapidly
equilibrating thin film passive air samplers and their potential application for
low volume air sampling studies. Environmental Science and Technology 39 (1),
261e267.
Finizio, A., Di Guardo, A., Arnoldi, A., Vighi, M., Fanelli, R., 1991. Different approaches
for the evaluation of Kow for s-triazine herbicides. Chemosphere 23 (6),
801e812.
Garabedian, S.P., Coles, J.F., Grady, S.J., Trench, E.C.T., Zimmerman, M.J., 1998. Water
Quality in the Connecticut, Housatonic, and Thames River Basins, Connecticut,
Massachusetts, New Hampshire, New York, and Vermont, 1992e1995. U.S.
Geological Survey Circular 1155.
Golding, C.J., Gobas, F.A.P.C., Birch, G.A., 2008. A fugacity approach for assessing the
bioaccumulation of hydrophobic organic compounds from estuarine sediment.
Environmental Toxicology and Chemistry 27 (5), 1047e1054.
Golding, C.J., Gobas, F.A.P.C., Birch, G.A., 2007. Characterization of polycyclic
aromatic hydrocarbon bioavailability in estuarine sediments using thin-film
extraction. Environmental Toxicology and Chemistry 27, 1047e1054.
Hansch, C., Leo, A.J., 1985. Medchem Project Issue No. 26. Pomona College, Claremont, CA.
Hansch, C., Leo, A., 1987. The Log P Database. Claremont, CA. p. 286.
Hansch, C., Leo, A.J., Hoekman, D., 1995. Exploring QSAR: Hydrophobic, Electronic
and Steric Constants. American Chemical Society, Washington.
Harner, T.J., Farrar, N.J., Shoeib, M., Jones, K.C., Gobas, F.A., 2003. Characterization of
polymer-coated glass as a passive air sampler for persistent organic pollutants.
Environmental Science and Technology 37 (11), 2486e2493.
Huckins, J.N., Tubergen, M.W., Manuweera, G.K., 1990. Semipermeable membrane
devices containing model lipid: a new approach to monitoring the bioavailability of lipophilic contaminants and estimating their bioconcentration
potential. Chemosphere 20 (5), 533e552.
Kenaga, E., Goring, C.A., 1980. Relationship between water solubility, soil sorption,
octanol-water partitioning and concentration of chemicals in biota. In: Proc. 3rd
Annual Symp., ASTM Special Technical Publication 707. Aquatic Toxicology.
ASTM, Philadelphia, pp. 78e115.
Liu, J., Qian, C., 1995. Hydrophobic effects of s-triazine and phenyl urea herbicides.
Chemosphere 31 (8), 3951e3959.
Lohmann, R., Muir, D., 2010. Global Aquatic Passive Sampling (AQUA-GAPS): using
passive samplers to monitor POPs in the waters of the world. Environmental
Science and Technology 44 (3), 860e864.
Mayer, P., Tolls, J., Hermens, J.L., Mackay, D., 2003. Equilibrium sampling devices.
Environmental Science and Technology 37 (9), 184Ae191A.
Meloche, L.M., Debruyn, A.M.H., Otton, S.V., Ikonomou, M.G., Gobas, F.A.P.C., 2009.
Assessing exposure of sediment biota to organic contaminants by thin-film
solid phase extraction. Environmental Toxicology and Chemistry 28 (2),
247e253.
McConnell, L.L., Harman-Fetcho, J.A., Hagy, J.D., 2004. Measured concentrations of
herbicides and model predictions of atrazine fate in the Patuxent River estuary.
Journal of Environmental Quality 33, 594e604.
Namiesnik, J., Zablegala, B., Kot-Wasik, A., Partyka, M., Wasik, A., 2005. Passive
sampling and/or extraction techniques in environmental analysis: a review.
Analytical and Bioanalytical Chemistry 381 (2), 279e301.
Scott, G.I., Fulton, M.H., Wirth, E.F., Chandler, G.T., Key, P.B., Daugomah, J.W.,
Bearden, D., Chung, K.W., Strozier, E.D., DeLorenzo, M., Sivertsen, S., Dias, A.,
Sanders, M., Macauley, J.M., Goodman, L.R., LaCroix, M.W., Thayer, G.W.,
Kucklick, J., 2002. Toxicological studies in tropical ecosystems: an ecotoxicological risk assessment of pesticide runoff in South Florida estuarine ecosystems. Journal of Agricultural and Food Chemistry 50 (15), 4400e4408.
St. George, T., 2008. Organic Contaminants in the Thames River Estuary: Development of a Thin-Film Passive Sampler. University of Connecticut, Master of
Science.
Please cite this article in press as: St. George, T., et al., A rapidly equilibrating, thin film, passive water sampler for organic contaminants;...,
Environmental Pollution (2010), doi:10.1016/j.envpol.2010.10.030
6
T. St. George et al. / Environmental Pollution xxx (2010) 1e6
Stuer-Lauridsen, F., 2005. Review of passive accumulation devices for monitoring
organic micropollutants in the aquatic environment. Environmental Pollution
136 (3), 503e524.
Vrana, B., Allan, I.J., Greenwood, R., Mills, G.A., Dominiak, E., Svensson, K.,
Knuttson, J., Morrison, G., 2005. Passive sampling techniques for monitoring
pollutants in water. Trends in Analytical Chemistry 24 (10), 845e868.
Wilcockson, J.B., Gobas, F.A.P.C., 2001. Thin-film solid-phase extraction to measure
fugacities of organic chemicals with low volatility in biological samples. Environmental Science and Technology 35 (7), 1425e1431.
Wilford, B., Harner, T., Helm, P., 2006. POGS (POlymer Coated Glass Samplers) for water
and air sampling. Society of Environmental Toxicology and Chemistry (SETAC)
North America, 27th Annual Meeting. Montreal, Canada, 5e9 November, 2006.
Wu, R.W., Harner, T., Diamond, M.L., 2008a. Evolution rates and PCB content of
surface films that develop on impervious urban surfaces. Atmospheric Environment 42, 6131e6143.
Wu, R.W., Harner, T., Diamond, M.L., Wilford, B., 2008b. Partitioning characteristics
of PCBs in urban surface films. Atmospheric Environment 42 (22), 5696e5705.
Yao, Y., Tuduri, L., Harner, T., Blanchard, P., Waite, D., Poissant, L., Murphy, C.,
Belzer, W., Aulagnier, F., Li, Y., Sverko, E., 2006. Spatial and temporal distribution
of pesticide air concentrations in Canadian agricultural regions. Atmospheric
Environment 40 (23), 4339e4351.
Zhao, W., Ouyang, G., Alaee, M., Pawliszyn, J., 2005. On-rod standardization technique for time-weighted average water sampling with a polydimethylsiloxane
rod. Journal of Chromatography 1124, 112e120.
Please cite this article in press as: St. George, T., et al., A rapidly equilibrating, thin film, passive water sampler for organic contaminants;...,
Environmental Pollution (2010), doi:10.1016/j.envpol.2010.10.030