Marine Pollution Bulletin 64 (2012) 1005–1011
Contents lists available at SciVerse ScienceDirect
Marine Pollution Bulletin
journal homepage: www.elsevier.com/locate/marpolbul
Determination of deployment specific chemical uptake rates for SPMD
and PDMS using a passive flow monitor
Dominique O’Brien a,⇑, Tatiana Komarova b,1, Jochen F. Mueller a,2
a
b
The University of Queensland, National Research Centre for Environmental Toxicology (Entox), 39 Kessels Rd, Coopers Plains, QLD 4108, Australia
Queensland Health Forensic and Scientific Services (Inorganics), 39 Kessels Rd, Coopers Plains, QLD 4108, Australia
a r t i c l e
Keywords:
Herbicide
Passive sampling
SPMD
PDMS
Flow
Calibration
i n f o
a b s t r a c t
Passive sampling techniques facilitate the time-integrated measurement of pollutant concentrations
through the use of a selective receiving phase. Accurate quantification using passive sampling devices
rely on the implementation of methods that will negate the effects of environmental factors (flow, temperature, etc.) or that will allow the calculation of the chemical specific rates of uptake (Rs) into the passive sampler employed. We have applied an in situ calibration technique based on the dissolution of
gypsum to measure the average water velocity to which a sampler has been exposed. We demonstrate
that the loss of gypsum from the passive flow monitor (PFM) can be applied to predict changes in Rs
dependent on flow when using the absorbent SPMD (semipermeable membrane device) and PDMS (polydimethyl siloxan) passive samplers. The application of the PFM will enhance the accuracy of measurements made when calculating and reporting environmental pollutant concentrations using a passive
sampling device.
Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction
Passive sampling devices are environmental monitoring tools
that have been developed to facilitate the assessment of chemical
concentrations in environmental medium (such as water: Cw) from
the mass of targeted analytes sorbed within a sequestering phase
(Cs) (Greenwood et al., 2007; Seethapathy et al., 2008; StuerLauridsen, 2005; Vrana et al., 2005). Advantages associated with
the use of passive sampling techniques are derived from the potential to obtain a time-weighted average (TWA) measurement of a
chemical concentration in the environment that may be at or below the detection limit of conventional grab sampling techniques.
The theory describing the uptake of a chemical by passive sampling devices has been the subject of considerable investigation
and review. The models employed in this work are found in the
fundamental work of Booij et al. (2007). These models may be used
to describe the accumulation of a chemical in terms of the mass
transfer coefficients (k0 the overall mass transfer coefficient, and
Abbreviations: PFM, passive flow monitor; PRC, performance reference com
pound; rPFM, daily mass lost from the PFM; SPMD, semipermeable membrane
device; PDMS, polydimethyl siloxan.
⇑ Corresponding author. Tel.: +61 7 3274 9060; fax: +61 7 3274 9003.
E-mail addresses: [email protected] (D. O’Brien), [email protected] (T. Komarova), [email protected] (J.F. Mueller).
1
Tel.: +61 7 3274 9070; fax: +61 7 3274 9181.
2
Tel.: +61 7 3000 9197; fax: +61 7 3274 9003.
0025-326X/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved.
doi:10.1016/j.marpolbul.2012.02.004
KSW a sample/water partition coefficient) when taking into consideration the sampler volume (VS), the interfacial exchange area (A).
However at the beginning of a passive samplers deployment the
rate of uptake into the sampler is large, the rate of elimination is
very small and the sampler device is said to be operating in the linear phase of uptake. During the linear phase of uptake, the calculation of a time integrated average pollutant concentration in water
may be simplified by relating the accumulated analyte mass in the
sampler (Ms) to the sampling rate (Rs) and deployment time (t),
where
CW ¼
MS
RS t
ð1Þ
The Rs of a specific passive sampler may be governed by the rate at
which a solute diffuses through either the receiving phase and any
overlying synthetic diffusion barrier, such as a membrane or gel; or
the overlying water boundary layer (WBL). The rate of diffusion of
an analyte within the sampler is relatively constant and depends
primarily on both the thickness and type of material used in the
construction of the receiving phase employed. Alternatively, the
thickness of the WBL is a function of the ambient environmental
condition to which the passive sampler is exposed when deployed
and the flow/turbulence near the surface of the receiving phase is
a key parameter. Methods have been developed to ensure that uptake into a passive sampler is either controlled (i.e. through the
introduction of an artificial diffusion barrier (Zhang and Davison,
1006
D. O’Brien et al. / Marine Pollution Bulletin 64 (2012) 1005–1011
1999), or the effect of flow can be assessed for each deployment
undertaken (Huckins et al., 2002a; Vrana et al., 2006).
The introduction of so called performance reference (or depuration) compounds (PRC) during the 1990s provided a tool for the
in situ prediction of sampling rates (Booij et al., 2002) or alternatively the ‘‘correction’’ of laboratory based sampling rates (Huckins
et al., 2002a). The PRC method facilitates the calculation of a dissipation rate from measurable loss of an internally spiked reference
standard from within the absorbent phase of the sampler. When
used in association with calibration data, the dissipation rate can
be directly related to the uptake of chemicals with similar
properties through the assumption that the uptake and release of
chemicals from the passive sampler is governed by the same
rate-controlling mechanisms (Huckins et al., 2002a) (typically isotropic kinetics).
The use of PRCs in the correction of Rs has been successful when
applied during the use of the semipermeable membrane device
(SPMD) and polydimethyl siloxan (PDMS) passive samplings for
the monitoring of hydrophobic priority pollutants (Booij et al.,
2002, 2006; Huckins et al., 1993, 2006, 2002a; Ouyang and
Pawliszyn, 2007; Petty et al., 2004; Vrana et al., 2007). However
difficulties have been reported when applying this method in association with adsorbent sequestering phases and in the modelling of
the diffusion kinetics of compounds with a log Kow of >5.5; as the
target analytes will remain partitioned within the sampling phase
and not desorbed into the adjacent water phase following isotropic
(first order) kinetics (Huckins et al., 2002a). An alternative in situ
calibration approach has been presented that employs a passive
flow monitor (the PFM) constructed through the casting of dental
plaster (gypsum) into a plastic holder. O’Brien et al. (2009) found
that the mass of gypsum lost per day (rPFM) from the PFM could
be used in the calculation of both water velocity (cm s1) and the
sampling rate (Rs) of a co-deployed phosphate passive sampler.
Further, the PFM calibration technique has been successfully applied in association with the adsorbent styrenedivinylbenzene-reverse phase sulfonated receiving phase when exposed within a
chemcatcher type passive sampler housing (O’Brien et al., 2011a).
During the presented study we aimed to undertake a laboratory
based calibration where the uptake of hydrophobic pesticides by
the SPMD and PDMS when exposed to different water velocities
is correlated against the rPFM recorded from co-deployed PFM devices. The herbicides and insecticides selected as part of this study
are of significance for ecosystem monitoring as they are routinely
applied in the agricultural industry. The transportation of these
chemicals from the site of application to environmental sensitive
areas such as estuaries and marine systems poses a threat to
non-target organisms and as such routine monitoring of pesticides
in surface waters is often undertaken. The results generated from
this study are aimed to improve the current methods for the quantification of pesticide concentrations in waterways when employing a passive sampling device.
2. Methods
2.1. Methods
Any glassware, tools or equipment that were in contact with
either the sampling phase or the extracting reagents were cleaned
using acetone (Merck Australia) and washed with dilute acid (3%
HCl) before a final rinse with ultra pure MilliQ H2O prior to use.
insulated, 1400 L stainless steel calibration tank. The sampling devices were attached to arms that extend 32 cm from the rotor of a
custom built stainless steel turntable, driven by a 12 V DC motor
(Hitachi, Japan). The rotation of the turntable can range between
2.5 and 50 rpm to produce a mechanical movement of between
0.7 and 110 cm s1 or a corresponding water flow rate of 1.7 and
73.8 cm s1 relative to the movement of the deployed samplers.
A Flowwatch – Air or Liquid Flow Measurement Instrument (ALFMI) from JDC Electronic SA (Australia) was used in the assessment
of the water velocity within the experimental chamber and its use
in determining the water movement relative to the deployed samplers has been described previously (O’Brien et al., 2011b).
The tank was kept closed throughout the experiments to prevent atmospheric deposition and photodegradation of chemicals
within the system. This also reduces variations in water temperature, which was not controlled. The water temperature was monitored through the use of a Thermochron iButton (Maxim
Integrated Products, USA), part number DS1921L-F50) that was attached to the rotation turntable.
Prior to the undertaking of these experiments the tank was
cleaned using acetone and rinsed with a wash of dilute acid (3%
HCl) and flushed with mains water. The tank was filled with
1400 L of mains water for the undertaking of the experiments.
2.3. Experimental procedure
Five experiments, each consisting of a ten day deployment,
were carried out. During each deployment PFM (in triplicate) were
secured and allowed to hang under their own weight from rotating
arms within the tank which allowed limited movement of the PFM
in the water flow. The SPMD and PDMS passive samplers were exposed in duplicate within cages (as described below) and attached
in a fixed position to the arms of the tank. The sampling devices
were exposed to either negligible flow or simulated flow conditions, which equated to a water flow velocity of ‘‘0’’-negligble flow,
3.4, 6.0, 16, 24 cm s1 relative to the movement of the samplers
through the water.
Agricultural chemicals with a wide range of log Kow (2.6–6.1)
were selected for inclusion in the study undertaken. The mass of
each chemical added was determined based on their solubility in
water and the final volume to be achieved in the experimental
chamber. A stock of chemicals was prepared in the laboratory
within a glass beaker before being diluted into the 1400 L within
the experimental chamber. The chamber was then closed and
mixed for 24 h prior to the collection of water samples for the
assessment of the concentration of each chemical achieved.
The mass of the individual PFMs was recorded prior to deployment and up to twice daily over the deployment period after excess water was removed with a cellulose based tissue (Kimwipes,
Kimberly–Clark) through the use of a mobile balance (Quad Beam
311 g). A new set of three PFMs were introduced to the tank at the
beginning of each consecutive deployment period. Six SPMD and
PDMS were prepared per deployment, four of which were deployed
into the tank and retrieved in duplicate after 5 and 10 days of exposure. The remaining two replicates were analysed as blanks to ensure that the deployed samplers were free of contamination at the
time of preparation.
Grab water samples (2 L unfiltered) were obtained at each time
point at which samplers were deployed and retrieved from the
tank.
2.4. Plaster flow monitors (PFM)
2.2. Experimental chamber
The uptake of chemicals by the passive samplers and loss of
plaster mass from the PFM, respectively were undertaken in an
The PFM devices are prepared from Dental Plaster (BORAL) cast
into 120 mL specimen containers [internal dimensions: 42 mm Ø,
105 mm high] (SARSTEDT) that were altered with the addition of
D. O’Brien et al. / Marine Pollution Bulletin 64 (2012) 1005–1011
1007
a cable tie (Weller) secured in place with a construction adhesive
(Selleys clear liquid nails) to allow for the PFM to be attached to
a mooring when deployed in the field (Fig. 1). The altered containers were then filled with a 1:2 plaster mix prepared using 80 mL
deionised water and 160 g plaster powder. Once firm (5 min after
the plaster is first mixed with water), the containers were capped
to prevent further drying and stored at 4 °C.
chamber. After deployment the cages were placed in the same cans
and returned to the laboratory where each SPMD and PDMS strip
was taken out of the cage, individually wrapped in acetone rinsed
aluminium foil, and stored at 17 °C together with corresponding
blanks until analysis.
2.5. SPMD preparation
Prior to dialysis, the surface of each SPMD (including blanks) was
cleaned according to standard methods by scrubbing with water,
dipping in hexane for 30 s and 0.5 M HCl for 20 s followed by rinsing
with acetone and isopropanol (Huckins et al., 1996). The extraction
of pesticides from SPMDs was performed using an Accelerated Solvent Extractor Dionex ASE 300 (pressure: 500 psi; temperature:
40 °C, static time: 20 min; flush volume: 60%; cycles: 5) using a mixture of n-hexane (Lichrosolv, 99.8%)/acetone (Lichrosolv, 99.8%)
(90:10). This method was adopted from the one developed by Wenzel et al. (2004) or SPMD extraction on ASE. To ensure that each
SPMD was extracted efficiently they were first placed inside
stainless steel mesh frames designed to support the sampler during
extraction before being loaded into 33 mL cells.
Lay-flat low density polyethylene tubing of approximately
60 lm thickness, 2.5 cm wide and 98 cm long was cut and pre-extracted in redistilled hexane based on the method described by
Huckins et al. (1996). Each strip was injected with 1 mL of 99%
pure triolein (Sigma–Aldrich T7140–50 g) that was heat seeled
within a length of 92 cm of tubing (an overall surface area of 460
cm2). The overhanging 4 cm at each end of the SPMD strip was
used in the creation of loops that could be passed over the stainless
steel rods used to support the strips when deployed within the
deployment cages. Individual SPMDs were wrapped in solvent
rinsed aluminium foil immediately after fabrication and stored at
17 °C until deployment.
2.8. SPMD extraction
2.9. PDMS extraction
2.6. PDMS preparation
PDMS sheets of 410 lm thickness were supplied by Purple Pig
Australia. Strips of 92 cm long and 2.5 cm wide were made by cutting PDMS sheets with a whole punched at each end. Prior to use
PDMS strips (3 pcs) were pre-extracted on a shaker in 900 mL of
fresh redistilled hexane for three consecutive 24 h periods. Then
PDMS strips were dried, wrapped in solvent rinsed aluminium foil
after fabrication and stored in plastic bags at 25 °C until deployment. Prior to deployment of the PDMS strip an acetone cleaned
cable tie was passed through each of the punched holes; this addition facilitated the securing of the strips within the deployment
cages.
2.7. Passive sampler deployment and retrieval
SPMD and PDMS strips were deployed in the open stainless
steel cages within which the strips were supported by stainless
steel rods (see Fig. 1). All deployment devices were washed with
a solution of Mucasol detergent and rinsed with acetone. Sampling
strips were mounted inside the cages in the laboratory and away
from direct sunlight. The cages were enclosed in metal cans (precleaned with acetone) for transportation to the tank where they
were taken out of the cans and deployed within the experimental
Fig. 1. PDMS and SPMD configuration within the deployment cages.
Prior to dialysis, the loops at each end of the PDMS strips were
removed and the surface of each PDMS (including blanks) was
cleaned according to the standard methods by scrubbing with
water, dipping in hexane for 30 s and 0.5 M HCl for 20 s followed
by rinsing with acetone and isopropanol (Huckins et al., 1996).
Each PDMS strip was extracted in 180 mL of redistilled hexane
on a shaker at room temperature (21 °C) for two 24 h periods. The
combined extracts from each sampler were then reduced to about
1 mL using rotary evaporators. Then the extract was passed
through a column with about 2 g of sodium suphate to remove
moisture and small particles of powder that covered the surface
of original PDMS sheets.
2.10. Extraction of water samples
The combined grab samples collected during the first deployment were spiked with the recovery internal standard prior to
extraction using a preconditioned 3 M SDB Empore™ Extraction
Disks (Phenomenex, USA). The pooled water sample was divided
into 1 L parts to be extracted separately in order to minimize
obstruction of disk by particulate matter. After each sample was
drawn through a 3 M SDB Empore™ (ED), excess water was removed by maintaining a low vacuum for 10–15 min. The analytes
were eluted with 7 mL of acetone using an initial gravimetrical elution phase, followed by a vacuum assisted elution phase. All sampling bottles were rinsed twice with dichloromethane (DCM)
(Merck) and the rinsate was filtered through sodium suphate
(Merck) and added to the acetone eluate. All eluates were
combined to form one pooled sample. The sample was again dried
over sodium suphate, concentrated and transferred into hexane.
The extracts were then subjected to a purification step using
adsorption chromatography in a glass column (8 mm i.d.) filled
with 2 g of activated silica gel and toped with a layer anhydrous
Na2SO4 (Sigma). The prepared columns were rinsed with DCM
and then by cyclohexane (Merck) prior to the addition of each sample. The first fraction eluted with 10 mL of cyclohexane was discarded prior to the elution of the absorbed compounds using
cyclohexane:dichloromethane mixtures (95:5; 90:10; and 85:15,
10 mL each). The samples were then concentrated, transferred to
an insert, spiked with the analytical internal standard and further
reduced under a gentle stream of nitrogen gas to a final volume
of 100 lL.
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D. O’Brien et al. / Marine Pollution Bulletin 64 (2012) 1005–1011
2.11. Analysis of extracts
one phase exponential association was used to describe the relationship between Rs and velocity, where:
SPMD and PDMS sample extracts were concentrated to 1 mL
using a Büchi Syncore Q-101 evaporation unit, filtered through a
Millex 0.45 lm syringe driven filter unit and then made up to
7 mL with dichloromethane (DCM) and spiked with an internal
standard (naphthalene D8, acenaphthene D10, phenanthrene
D10, Chrysene D12 and Perylene D12) before being subjected to
size exclusion chromatography (19 mm by 150 mm guard column,
followed by a 19 mm by 300 mm main column, packed with Envirogel [100 Å pore size, 15 lm particle size, Waters] as the stationary phase and with dichloromethane DCM as the mobile phase).
The flow rate was 4.5 mL min1 and the sample fraction was collected between 15.00 to 24.20 min after sample injection onto
the GPC column. Samples were then concentrated under a gentle
stream of nitrogen, transferred to an insert and further reduced
to a final volume of 100 lL. The separation and quantification of
all herbicides following extraction from the passive samplers was
performed by GC–MS. Instrumental analysis was conducted by
Queensland Health Scientific Services on a Varian 3400 GC
equipped with a Finnigan A200S liquid autosampler (splitless;
injector temperature 295 °C; GC column: J&W DB-1, originally
20 m, 0.2 mm i.d., 0.33 lm film thickness; temperature programme: 65 °C (isothermal 2 min), 20 °C min1 to 295 °C (isothermal 10 min) and coupled to a Finnigan SSQ 710 single stage
quadrupole mass selective detector.
The separation and quantification of the pesticides extracted
from water was performed by GC–MS. All samples were spiked
with an internal standard (naphthalene D8, acenaphthene D10,
phenanthrene D10, Chrysene D12 and Perylene D12) before instrumental analysis was conducted by Queensland Health Forensic and
Scientific Services on a Shimadzu QP5050A GCMS splitless; injector
temperature 250 °C; GC columns: Phenomenex ZB5 and SGE HT5,
30 m, 0.25 mm i.d., 0.25 lm film thickness.
Rs ¼ Rsð0 cm s1 Þ þ ðRsðmaxÞRsð0 cm s1 Þ Þð1 expðK v v ÞÞ
2.12. QA/QC
The quantification criteria included confirmation of the retention times and selective ion monitoring of the labelled internal
standards and respective analyte. Routinely, the mass fragment
with the highest intensity (base peak) was used for quantification.
20% of the samples consisted of QC samples (lab blanks). Detection
limits for all samples were defined by the mean amount in the laboratory blanks plus three times the standard deviation. Where a
compound could not be identified in the blank, the detection limit
was set as three times the average noise peak area (2 ng sampler1). Reproducibility was measured by calculating the mean
normalised difference between replicates (n = 2). The% normalised
differences were calculated for replicates A and B according to
h
i
jvalueAvalueBj
100.
ððvalueAþvalueBÞ=2Þ
ð2Þ
where Rsð0 cm s1 Þ is the Rs when exposed to still waters, Rs(max) is the
maximum Rs for the chemical of interest, v is velocity expressed in
cm s1 and Kv is a rate constant expressed in reciprocal of the units
of velocity. The chemical specific constants for the use of Eq. (4) in
the calculation of Rs dependent on flow has been given in Table 1
(PDMS) and Table 2 (SPMD).
The observed approach to a maximum Rs with velocity corresponds with the uptake limiting resistance shifting from the WBL
control to the diffusion of the analytes within the membrane of
the passive samplers (Huckins et al., 2006). As the rate of diffusion
across the membrane is independent from environmental conditions, the maximum Rs achievable will be equal to the constant diffusion coefficient specific to the analytes targeted and the
properties and dimensions of the membrane employed.
3.2. Correlation of RS for changes in water velocity: PFM calibration
The mass lost from the exposed PFM over each deployment period remained linear over time and ranged from 4.95 to 40.1 g or
0.46 to 3.9 g day1 dependant on the flow rates to which the
PFM were exposed. A significant linear regression produced when
the daily mass lost (or rPFM) was plotted against velocity (r2 = 0.99)
allows an estimation of the water velocity where:
v ðcm s1 Þ ¼ ðrPFM 0:065Þ=0:164
ð3Þ
The use of the PFM is unable to distinguish between velocities of
less than 3.4 cm s1 [observed in the present study (see Table 3)
and discussed elsewhere (O’Brien et al., 2011a,b, 2009, 2011c)]. As
such an assessment of the performance of the PFM technique for
the correction of PDMS and SPMD sampling rates when exposed
to a change in velocity was undertaken excluding the PFM results
obtained at no-flow conditions (i.e. no mechanical rotation of the
table).
Fig. 3 shows a plot of PDMS and SPMD sampling rates against
rPFM. While there is a clear one phase exponential association between the SPMD Rs and rPFM [r2 ranged between 0.84 and 1], the
relationship between PDMS Rs and rPFM can be described as linear
[r2 ranged between 0.69 and 0.98] but a relationship of greater significance is achieved when described by a one phase exponential
association [r2 ranged between 0.92 and 0.99] for all chemicals
but prothiophos.
As such the change in Rs can be calculated where:
Rs ¼ Rsð0 cm s1 Þ þ ðRsðmaxÞ Rsð0 cm s1 Þ Þð1 expðK rPFM rPFM ÞÞ
ð4Þ
and K rPFM is a rate constant expressed in reciprocal of the units of
rPFM. The chemical specific constants for the use of Eq. (4) in the calculation of Rs dependant on flow has been given in Table 4 (PDMS)
and Table 5 (SPMD).
3. Results
4. Discussion and conclusion
3.1. Correlation of Rs with water velocity
The uptake of bifenthrin, dieldrin, oxadiazon, pendimethalin,
permethrin, prothiophos and trifluralin into both the PDMS and
SPMD remained linear over time when exposed to still and flowing
water [r2 ranged between 0.9 and 0.99] (Fig. 2). The Rs for each
chemical at all flow velocities investigated was calculated through
the use of Eq. (3). Change in the rate of chemical uptake (Rs) into
the PDMS and SPMD passive samplers as a function of water velocity is shown in Fig. 2 (Ai and Bi). As the change in Rs with velocity
for all chemicals, except trifluralin by the PDMS, was not linear, a
In the present study the uptake of chemicals by the PDMS and
SPMD has been shown to be influenced by the water velocity to
which the sampling device has be exposed to when the sampler
is operating within the integrative phase of chemical uptake. Few
studies have investigated the change in Rs of the PDMS or SPMD
when exposed to a range of flow velocities as the research has
largely focused on the influence of temperature on the sampler
kinetics. Studies into the effect of environmental factors on the
performance of the SPMD have shown that while temperature does
influence Rs, there will be little effect on the uptake of organic
1009
D. O’Brien et al. / Marine Pollution Bulletin 64 (2012) 1005–1011
SPMD
PDMS
40
20
30
Rs (L d-1)
Rs (L d-1)
15
20
10
10
5
0
0
0
10
20
Water velocity (cm
BIFENTHRIN
10
s-1)
20
Water velocity (cm
PERMETHRIN
PROTHIOPHOS
OXADIAZON
PENDIMETHALIN
DIELDRIN
0
30
30
s-1)
TRIFLURALIN
Fig. 2. PDMS and SPMD sampling rates (Rs) as a function of water velocity.
Table 1
Parameters for the use when calculating the PDMS Rs dependant on water velocity using Eq. (4).
Chemical analyte
Bifenthrin
Dieldrin
Oxadiazon
Pendimethalin
Permethrin
Prothiophos
Trifluralina
Rsð0 cm s1 Þ
0.30
0.15
0.57
0.77
0.34
4.19
0.57
Std. error
Rs(max)
0.71
2.72
1.52
1.91
3.86
4.30
3.72
Std. error
4.87
29.8
37.3
68.6
35.1
27.4
31584
0.70
15.7
71.6
333
17.4
5.23
5.3e + 007
Kv
0.17
0.06
0.02
0.01
0.06
0.24
<0.01
Std. error
0.09
0.06
0.05
0.06
0.06
0.16
0.06
Goodness of fit
R2
Absolute sum of squares
0.93
0.95
0.96
0.94
0.94
0.94
0.93
1.03
17.2
5.91
9.53
34.2
18.7
37.3
a
The regression obtained is considered ambiguous as the top plateau could not be constrained to a constant value (i.e. change in Rs remained linear with a change in
velocity).
Table 2
Parameters for the use when calculating the SPMD Rs dependant on water velocity using Eq. (4).
Chemical analyte
Bifenthrin
Dieldrin
Oxadiazon
Pendimethalin
Permethrin
Prothiophos
Trifluralin
Rsð0 cm s1 Þ
0.24
0.28
0.38
0.32
0.52
3.63
0.27
Std. error
Rs(max)
0.54
0.26
3.11
0.61
0.73
0.55
0.11
Std. error
3.42
4.03
19.5
5.74
7.27
20.8
0.74
0.50
0.21
3.51
1.23
2.22
0.69
0.12
Kv
0.18
0.24
0.15
0.10
0.08
0.23
0.16
Std. error
0.10
0.05
0.08
0.06
0.06
0.03
0.13
Goodness of Fit
R2
Absolute sum of squares
0.92
0.99
0.92
0.95
0.95
1.00
0.84
0.61
0.14
20.2
0.82
1.18
0.31
0.03
Table 3
Water velocity and temperature within the experimental tank and the corresponding rate of mass loss from PFM devices.
Velocity (cm s1)
0
3.4
8
16.1
24.1
Temperature (°C)
average (min/max)
rPFM
16.2
16.5
18.5
19.3
22.0
0.56
0.57
1.43
2.74
3.99
(16.0–16.5)
(16.0–19.5)
(17.0–19.0)
(18.0–20.5)
(21.0–22.5)
1
(g day
)
Velocity (cm s1) predicted from rPFM
ND (%) between measured
and predicted velocity
1.5
1.6
7.3
16.1
24.4
–
72
8.7
0.2
1.2
2
SD
r
0.01
0.02
0.02
0.03
0.03
0.98
0.94
0.99
0.99
0.99
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D. O’Brien et al. / Marine Pollution Bulletin 64 (2012) 1005–1011
SPMD
PDMS
40
20
30
Rs (L day-1)
Rs (L day-1)
15
20
10
10
5
0
0
10
20
0
30
0
10
rPFM (g day-1)
20
30
rPFM (g day-1)
BIFENTHRIN
OXADIAZON
PERMETHRIN
DIELDRIN
PENDIMETHALIN
PROTHIOPHOS
TRIFLURALIN
Fig. 3. PDMS and SPMD sampling rates (Rs) as a function of water velocity rPFM.
Table 4
Parameters for the use when calculating the PDMS Rs when correcting for changes in water velocity using rPFM and Eq. (4).
Chemical analyte
Rsð0 cm s1 Þ
Std. error
Rs(max)
Std. error
K rPFM
Std. error
Goodness of fit
R2
Bifenthrin
Dieldrin
Oxadiazon
Pendimethalin
Permethrin
Prothiophos
Trifluralina
1.52
3.14
0.08
0.08
2.34
Too few points
1.43
1.30
3.19
1.82
2.78
6.24
7.20
28.9
51.3
101
37.3
7.60
7.22
105
650
16.8
7.03
132836
9.6e + 008
Absolute sum of squares
0.26
0.40
0.09
0.04
0.37
0.60
0.21
0.22
0.31
0.34
0.92
0.99
0.99
0.98
0.98
0.40
1.89
1.01
2.51
7.61
<0.01
0.42
0.96
16.98
a
The regression obtained is considered ambiguous as the top plateau could not be constrained to a constant value (i.e. change in Rs remained linear with a change in
velocity).
Table 5
Parameters for the use when calculating the SPMD Rs when correcting for changes in water velocity using rPFM and Eq. (4).
Chemical analyte
Bifenthrin
Dieldrin
Oxadiazona
Pendimethalin
Permethrin
Prothiophosb
Trifluralin
a
b
Rsð0 cm s1 Þ
0.89
0.58
0.42
0.05
12.0
9.03
1.14
Std. error
0.54
1.05
0.17
0.92
8.25
2.73
Rs(max)
4.14
9.25
348
6.25
18.6
24.7
3.36
Std. error
K rPFM
0.18
5.00
1.2e + 006
1.35
1.29
0.38
0.96
0.28
<0.01
0.49
1.37
0.49
1.44
Std. error
0.28
0.30
0.89
0.28
0.47
1.07
Goodness of fit
R2
Absolute sum of squares
0.99
0.98
0.84
0.98
0.99
1.00
0.94
0.02
0.25
0.01
0.13
1.73
0.00
0.16
The regression obtained is considered ambiguous as the top plateau could not be constrained to a constant value.
There was a perfect fit of the regression to the data set obtained.
contaminants unless large geographical and temporal scales are involved (Booij et al., 2003). Of the chemicals investigated in this
study, Huckins et al. (2002b) and Sabalinas and Sodergren (1997)
have reported SPMD sampling rates for dieldrin and trifluralin,
respectively. Huckins et al. observed a sampling rate 1.3, 2.6 and
4.6 L d1 when SPMD were exposed to a flow of 0.004 cm s1 and
temperatures of 10, 19 and 24 °C, respectively. Sabalinas and
Sodergren (1997) reported SPMD sampling rated of 6.8 and
3.6 L d1 for dieldrin and trifluralin, respectively when exposed
to a flow of 0.006 cm s1 however the temperature at which the
experiments were undertaken was not reported. The sampling
rates of dieldrin and trifluralin observed during this study ranged
from between 0.5 and 6.5 L d1 and 0.3 and 3.6 L d1, respectively
when exposed to still water and flow rates of up to 24 cm s1. As
the temperatures were not controlled during these experiments
it is possible that temperature had an influenced on Rs and it would
be advisable that further experiments are undertaken to assess
change in Rs with temperature when SPMD are exposed to the
higher flow rates that have been used in these experiments and
are encountered when the devices are deployed into surface
D. O’Brien et al. / Marine Pollution Bulletin 64 (2012) 1005–1011
waters such as rivers and estuaries. However the application of the
PFM for the measurement of water flow velocities has been clearly
demonstrated as a cost effective, practical in situ calibration method that can be used in association with the PFM method or as an
alternative when PRCs can not be applied.
Acknowledgements
The work described was undertaken as a part of an ARC Discovery project (DP0556205) at the National Research Centre for Environmental Toxicology (Entox). Entox is a research center of the
University of Queensland which is co-funded by Queensland
Health Forensic and Scientific Services. The authors wish to
acknowledge the assistance of Andrew Dunn in undertaking the
described experiments and during the analysis of samples
collected.
Appendix A. Supplementary data
Supplementary data associated with this article can be found, in
the online version, at doi:10.1016/j.marpolbul.2012.02.004.
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