Accepted Preprint
Environmental Chemistry PREDICTING BIOACCUMULATION OF POLYCYCLIC AROMATIC HYDROCARBONS IN SOFT-SHELLED CLAMS (MYA ARENARIA) USING FIELD DEPLOYMENTS OF POLYETHYLENE PASSIVE SAMPLERS
Loretta A. Fernandez and Philip M. Gschwend
Environ Toxicol Chem., Accepted Article • DOI: 10.1002/etc.2892
Accepted Article
"Accepted Articles" are peer-reviewed, accepted manuscripts that have not been edited, formatted, or in any way altered by the authors since acceptance. They are citable by the Digital Object Identifier (DOI). After the manuscript is edited and formatted, it will be removed from the “Accepted Articles” Web site and published as an Early View article. Note that editing may introduce changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. SETAC cannot be held responsible for errors or consequences arising from the use of information contained in these manuscripts.
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Environmental Chemistry
Environmental Toxicology and Chemistry DOI 10.1002/etc.2892
PREDICTING BIOACCUMULATION OF POLYCYCLIC AROMATIC HYDROCARBONS IN SOFT-SHELLED CLAMS (MYA ARENARIA) USING FIELD DEPLOYMENTS OF POLYETHYLENE PASSIVE SAMPLERS
Running title: Predicting PAH bioaccumulation using PE passive samplers
Loretta A. Fernandez*†‡ and Philip M. Gschwend‡
† Departments of Civil and Environmental Engineering and Marine and Environmental Sciences, Northeastern University, Boston, Massachusetts, USA ‡ Department of Civil and Environmental Engineering, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA
* Address correspondence to [email protected]
Additional Supporting Information may be found in the online version of this article.
This article is protected by copyright. All rights reserved Submitted 11 September 2014; Returned for Revision 11 January 2015; Accepted 13 January 2015
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Abstract: Biota-sediment accumulation factors (BSAF), frequently used to predict tissue concentrations of organisms living within and above sediments contaminated with hydrophobic organic chemicals (HOCs), often produce inaccurate estimates. Hence, freely-dissolved porewater concentrations, CW, have also been investigated as predictors of organism tissue concentrations, but they are more difficult to measure than bulk sediment concentrations (used with BSAF). In situ passive sampling methods make it possible to deduce CW with less effort than is required to measure the value directly, and make it possible to relate CW with tissue concentrations of undisturbed, native organisms. In the present work, polyethylene (PE) passive samplers, containing performance reference compounds (PRCs) (d10-phenanthrene, d10-pyrene, and d12-chrysene), were deployed in diverse sediment beds near Boston, MA for a one-week period. Clams (Mya arenaria) and sediments were then collected from the same sediments. Concentrations of three PAHs (phenanthrene, pyrene, and chrysene) were measured in the pore waters, in clam tissues, and in the bulk sediment. BSAF and PE-deduced CW were used to predict organism tissue concentrations. Ratios of predicted-to-measured values showed that the BSAF method over-predicted tissue concentrations in M. arenaria by up to two orders of magnitude. The PE-deduced CW method resulted in average ratios closer to 1 (0.43 ± 0.26, 3.7 ±
2.5, and 1.1 ± 1.2 for phenanthrene, pyrene, and chrysene, respectively, N=26, uncertainty = ±1σ). This article is protected by copyright. All rights reserved
Keywords: PAH, Passive sampling, Polyethylene, BSAF, Bioavailability
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INTRODUCTION In the search for approaches to determine the ecological risks associated with sediments
contaminated with hydrophobic organic chemicals (HOCs), several indicators of the potential for compounds to be transferred from sediments to biological receptors have been studied. These include biota-sediment accumulation factors (BSAFs) [1-3], and freely-dissolved porewater water concentrations [2, 4-7]. The general idea behind both approaches is that, in an equilibrated
system, knowing the concentration in a single compartment (i.e., sediment organic carbon or porewater), one can estimate concentrations in organism tissues (often assumed to be primarily
in lipids). Organism tissue concentration can be estimated using BSAFs that relate lipidnormalized organism concentrations to organic carbon normalized sediment concentrations Corg = flip BSAF (CSED / fOC)
(1)
where Corg is the concentration in organism tissues (mol/g tissue), flip is the lipid fraction of the
organism tissue (g lipid/g tissue), CSED is the concentration in sediments (mol/g sediments), and fOC is the organic carbon (OC) fraction of the sediments (g OC/g sediment). Similarly, organism concentrations would be predicted from freely-dissolved porewater concentrations, CW (mol/mL water)
Corg = flip Klip-w CW
(2)
where Klip-w is the lipid-water equilibrium partitioning coefficient ((mol/g lipid) / (mol/mL water)).
There are challenges to using either approach. BSAF values depend on organic carbon-
normalized sediment concentrations. However, assuming OC is the only component regulating sediment-water transfer may not be applicable to all sediments. Equilibrium partitioning coefficients for organic carbon and water, KOC, have been observed to range over three orders of
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magnitude for PAHs in field sediments [8-11], most likely due to differing sorptive properties of the many types of materials measured as fOC. Subfractions, including black carbon (BC), oil, pitch, and chars, have been used to refine equilibrium partitioning (EqP) models [12-17], but it is
not always clear how best to assess these subfractions (e.g., Hammes et al. 2007 for BCs). Arp et al. [17] found that KOC tuned to sorption between coal tar and water predicted PAH sediment-
water distribution of PAHs better than other KOC values available in the scientific literature that
have been tuned to other organic carbon pools such as peat, coal, natural organic matter, octanol, or granular activated carbon. As an alternative to organic carbon-normalized sediment concentrations, biological
effects have been compared to concentrations in a more uniform phase of sediments, the pore waters [2, 7, 19]. Directly measuring CW by extracting HOCs from porewater samples, however,
is complicated by the need to physically separate water from sediment solids, then either remove or account for the colloids remaining in porewater samples [20-21]. For this reason, porewater concentrations are often estimated using equilibrium partitioning (EqP) techniques similar to those described above, where CW is assumed to be equilibrated with organic carbon-normalized sediment concentrations CW = CSED / (fOC KOC)
(3)
This method, of course, suffers from the same problems described above, where organic carbon
sub-fractions may have very different sorptive properties. Passive sampling methods offer an alternative by introducing an easily separable and
well-defined organic phase to sediment-water systems, from which CW may be deduced. [22-25] CW = Cpolymer / Kpolymer-w
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(4)
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where, Cpolymer is the concentation in the polymeric sampler, and Kpolymer-w is the polymer-water partition coefficient. This method is easier than the porewater extraction approach and results in accurate CW [26-27]. While passive sampling techniques have been used in laboratory studies to
investigate the relationship between CW, toxicity endpoints, and organism tissue concentrations [28-35], recent advances in passive sampling allow measurement of HOC concentrations in pore waters of intact sediment beds [26, 36-38]. This makes comparison of porewater HOC
concentrations with tissue concentrations of undisturbed, native organisms possible. In several studies, laboratory-raised worms have been exposed to in situ sediments and various methods, including both field and laboratory deployed passive samplers (solid-phase microextraction (SPME) and polyoxymethylene (POM) strips), have been used to predict PAH and PCB uptake
by the organisms [39-41]. Van der Heijden and Jonker [39] found that field deployed SPME performed better than the laboratory measurements in predicting uptake by the oligochaete (Lumbriculus variegatus). In the present study, polyethylene strips containing three performance reference
compounds (PRCs) (d10-phenanthrene, d10-pyrene, and d12-chrysene), which allow for shorter exposure times than are required for full equilibration, were used to measure CW profiles with depth, for three PAHs (phenanthrene, pyrene, and chrysene) in sediments beds at six locations. Field sites were selected for their diverse carbon fractions, variable PAH sources, and the
presence of native organisms, the marine clam Mya arenaria. Sediment PAH concentrations, organic carbon fractions, and published BSAF for M. arenaria, along with PE deduced porewater concentrations, were used to predict organism tissue concentrations. Concentrations
predicted using the two methods were compared to those measured in the native organisms.
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MATERIALS AND METHODS All solvents were Baker Ultraresi-analyzed. Laboratory water was treated with an ion-
exchange and activated carbon system (Aries Vaponics) until 18 MOhm-cm resistance was achieved, followed by UV exposure (TOC reduction unit, Aquafine Corporation). All PAH standards and PRCs were purchased from Ultra Scientific in methanol, acetone, or dichloromethane. Polyethylene (PE) strips were prepared from low-density polyethylene (LDPE) sheets (25
µm from Trimaco). All PE was soaked twice in dichloromethane for 24 h and twice in methanol for 24 h, before soaking twice in water for 24 h. Finite-difference model calculations for diffusion of d12-chrysene from stirred water, through a water-side boundary layer, and into PE indicated that 1 month is sufficiently long for 25 µm thick PE to equilibrate with the solution; but to be sure of even PRC distribution throughout the polymer films, the PE used in the present work was in contact with PRC solution for >7 months (~20 g PE in 1 L aqueous PRC solution
that was agitated by hand ~once per day). Field sampling PE strips were deployed in, and clams and sediments were collected from, six locations
near Boston, MA (Table 1, Supplemental Data, Figure S1) in November 2008. The sites were
selected based on the presence of M. arenaria, previous measurements of PAH concentrations in the sediments, and historical information concerning industrial use of the areas. From north to south, the locations included the following: (a) Collins Cove, Salem, MA – a large but shallow cove approximately 700 m east if the Salem Harbor Power Station, a coal- and oil-fired power plant, (b) Pioneer Village, Salem, MA – a sandy beach along Salem Harbor, approximately 1500
m south of Salem Harbor Power Station, (c) Forest River (Lead Mills), Marblehead, MA – at the
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mouth of a tidal river down-stream of a 19th century lead mill that burned down in 1968, (d)
Pines River, Saugus, MA – along a tidal river adjacent to a closed land fill and a waste-to-energy plant, (e) Island End, Chelsea, MA – a coal-tar contaminated site, and (f ) Dorchester Bay,
Quincy, MA – downwind of the Southeast Expressway (I-93) and near the site of a former garbage incinerator on Spectacle Island. It was expected that these sites would provide a wide range of PAH concentrations and varying organic and black carbon fractions in the sediments. Sampling of all sites was conducted over a period of two weeks, in November 2008,
during low tides, in the intertidal zone. First, PE samplers were deployed at each site directly
adjacent to what appeared to be clam siphon holes in the sediment. PE samplers in aluminum frames were pushed into sediments to a depth of between 4 and 12 cm depending on how far they would go in before meeting resistance (Supplemental Data, Figures S2 and S3). As many of these sites are located in urban areas, a one-week deployment time was selected in order to minimize chances that samplers would be vandalized. Mathematical modeling and the results of previous field testing indicated that this exposure time would be sufficient for the transfer of
measureable masses of target and PRC compounds between sediments and samplers. After being in place for one week later, PE samplers were collected. At that time, clams from the adjacent sediments were collected using a spade, and sediment samples were taken from the surface (approximately top 10 cm) and from a lower layer (approximately 10-20 cm deep). PE strips were rinsed with clean water in the field and placed between aluminum plates, which were then wrapped in aluminum foil. Clams and sediments were placed in glass jars. All samples were returned to the lab on ice. Clams were stored at -20ºC until dissection and extraction, sediments were stored at 4ºC until extraction, and PE were sectioned at 4 cm intervals and
immediately extracted. PE samplers deployed at Pioneer Village were not found on the retrieval
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trip. Instead, clams and adjacent sediment were collected and returned to the lab where three PE strips (approximately 2 cm x 2 cm) were inserted directly into jarred sediments using forceps, and exposed for 32 days. At Island End, clams were not found at the time of PE recovery, so clams collected from 2-3 m away, at the time sampler deployment, were used instead. PE extraction Upon return to the laboratory, PE strips were again rinsed in clean water and swabbed
with a wipe (a hexane-soaked wipe in the case of Island End samplers) to ensure that only absorbed molecules, but not those associated with adhering sediment particles or tarry films, would be quantified. Strips were cut into 4 cm sections, surrogate standards (d10-anthracene, d10-fluoranthene, and d12-benz(a)anthracene) were added, and strips were extracted three times by soaking in 15 mL of dichloromethane overnight. The combined extracts were exchanged into hexane and concentrated to approximately 0.5 mL under a gentle stream of ultra pure grade
nitrogen. Injection standards (d10-acenaphthene, m-terphenyl, and d12-perylene) were added to the extracts before gas chromatography/mass spectrometry (GC/MS) analysis. Calculation of CW from PRC and target analyte concentrations Freely dissolved porewater concentrations were calculated using previously described
methods [26]. Briefly, using PE strips impregnated with PRCs that match target compounds in terms of partition and diffusion coefficients, CW may be calculated from the concentrations of
target chemical (CPE) and PRC (CPRC) in the PE after time, t CW = (CPE (t) × CPRC, init) / (CPRC (t) × KPEW)
(5)
where CPRC, init is the initial concentration of PRC in the sampler, and KPEW is the polyethylene-
water partition coefficient for the PRC and target chemical (Table 2). Clam extractions
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Extraction of PAHs from clams was performed using a method modified from that
described by Yusa et al. [42]. Clams were shucked and dissected to remove stomach, intestines
and most internal organs. The remaining neck, foot, and adductor muscles were then sliced and chopped using two razor blades until a mushy consistency was achieved. Approximate 2 g of the chopped clam were then ground with 1-2 g of precombusted diatomaceous earth (Hyflo Supercel, Sigma Aldrich) until dry and crumbly. Three precombusted GF/B filters (Whatman International Ltd.) were used in the bottom of each 33 mL stainless steel extraction cell. The clam mixture was added to cells between two layers of precombusted Ottawa sand (EMD Chemicals). Surrogate standards were added to the top of the second sand layer before accelerated solvent extraction (ASE) was performed. The remaining homogenized clam tissue was weighed and dried to determine its water fraction. ASE was performed using a Dionex ASE 200 (Dionex Corporation). Each cell was
extracted three times, using dichloromethane:methanol (1:1), heated for 5 min (125ºC) at 1500 psi, and flushed with 60% of cell volume between each extraction. Extracts were collected in amber glass vials with aluminum-lined caps. Approximately 20 g anhydrous Na2SO4 were added to each vial, and extracts were stored overnight at 4ºC. Combined extracts were exchanged into approximately 2 mL hexane using a rotary evaporator (Buchi Rotavapor-R, Brinkman Instruments) before column chromatography and GC/MS analysis. Because ASE also extracted lipids and some proteins from the tissues, a lipid fraction
could not be measured by drying an aliquot of the extract as is normally done when using more gentle Soxhlet extraction methods that more closely mimic the classic Bligh and Dyer method [43]. In the present study, lipid and protein fractions for all clams are assumed to be 5% and 50%, respectively, based on measurements by other researchers. Lohmann et al. [24] measured
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lipid fractions between 5.3 and 9.0% dry weight in clams from Dorchester Bay and 5.8 and 7.6% dry weight in clams from the Saugus River (near Pines River) in 2001 on a dry weight basis, while protein fractions were calculated to be 48%. McDowell and Shea [44] measured lipid fractions between 3.7 and 6.1% for whole clams from the Massachusetts Bay, while Galassi et al. [45] measure an average of 5.1% in clams from the Gulf of Gdansk, in the Baltic Sea. Protein fractions for other aquatic invertebrates have been measured in the range of 50-70% [46]. Uncertainty in lipid-normalized clam tissue concentrations, Corg, was determined by
analyzing multiple clams taken from close proximity. The coefficients of variation (CV) of phenanthrene, pyrene, and chrysene in six clams taken from the Collins Cove, station 3 were 51%, 34%, and 50%, respectively. This uncertainty was assumed for all clam measurements.
Sediment extractions Sediments were homogenized in their collection jars by stirring with a spatula for 5 min,
while removing large stones and shells. Subsamples were taken for drying and weighing to determine water content. Additional subsamples (3-10 g) were taken for ASE. Two precombusted GF-B filters were used in the bottom of each 11-mL stainless steel extraction cell. Wet sediments were added to the cell between two layers of pre-combusted Ottawa sand. Surrogate standards were added to the top sand layer before ASE using the same method described above for clam extractions. Again, extracts were dried overnight at 4ºC using approximately 20 g anhydrous Na2SO4. Extracts were exchanged into hexane and reduced to approximately 2 mL using a rotary evaporator before column chromatography and GC/MS analysis. Column chromatography
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Chromatography columns were prepared in 20 cm long, 1 cm o.d., glass columns with
approximately 30 mL reservoirs. A small plug of glass wool, followed by approximately 2 cm of activated granular Cu0 (20-30 mesh, Baker Analyzed Reagent, J.T. Baker) for columns used with sediment extracts, followed by 5 g fully activated (475 ºC for 24 hr) silica gel (100-200 mesh, EMD Chemicals), followed by approximately 6 g anhydrous Na2SO4 were dry packed into each column [47]. Hexane, 30 mL, was used to flush each column before clam or sediment extracts were charged onto the column. PAHs were collected by running 100 mL of hexane:dichloromethane (9:1 v:v) through the column under slight pressure. The solvent
mixture was exchanged into hexane and reduced to approximately 0.5 mL for clam extracts and approximately 10 mL for sediment extracts. Injection standards (d10-acenaphthene, mterphenyl, and d12-perylene) were added to each extract immediately before GC/MS analysis to
determine sample volume. Gas chromatography/mass spectrometry analysis All extracts were analyzed using gas chromatography/mass spectrometry (GC/MS, JEOL
GCmate, JEOL Ltd.). Splitless 1-µL injections were made onto a 30 m J&W Scientific HP-5MS capillary column (0.25 mm internal diameter with a 0.25 µm film thickness). The injection port temperature was held at 305ºC. The initial column temperature of 70ºC was raised at 20ºC/min until a temperature of 180ºC was reached, and then the temperature was raised 6ºC/min until a
temperature of 300ºC was reached, and remained there for 7.5 min. The MS was operated in selected ion monitoring (SIM) and EI+ modes. Calibration standards containing 20 aromatic compounds including each of the target compounds, surrogates, and injection standards used in the present study, were run every 3 to 5 sample measurements to monitor instrument stability,
determine response factors, and confirm that measurements remained in the linear range for the
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instrument. Repeated observations using the calibration standard indicated the measurement uncertainty for the instrument was typically ±10% relative error. Quality control Target chemicals were above detection limits in all clam, PE, and sediment samples.
Percent recoveries for the surrogate standards ( 1 RSD) were 73 ± 39%, 71 ± 24%, and 60 ± 31% in PE extracts for d10-anthrancene, d10-fluoranthene, and d12-benz(a)anthracene,
respectively. Recoveries in clams were 54 ± 36%, 75 ± 26%, and 81± 29% for d10-anthrancene,
d10-fluoranthene, and d12-benz(a)anthracene, respectively. Recoveries in sediment extracts were 73 ± 36%, 88 ± 32%, and 91 ± 33% for d10-anthrancene, d10-fluoranthene, and d12benz(a)anthracene, respectively. Large ranges in recovery of surrogate standards were expected due to the many transfers and manipulations of the extracts. PRC and target compound concentrations were corrected for recoveries of the corresponding closest-eluting surrogate standard. Extraction and measurement methods were tested for accuracy by measuring standard
reference materials (SRM), NIST SRM 1974b – Organics in Mussel Tissue (Mytilus edulis) and
NIST SRM 1941a – Organics in Marine Sediment. While pyrene and chrysene matched within
uncertainty for the mussel tissue, phenanthrene measurements were 134% of the reported value. Repeated measurements yielded the same result. The high recovery of phenanthrene in the mussel tissues may have been due to contamination of the sample before extraction. For the sediment SRM, phenanthrene matched the reported value within uncertainty, while pyrene and chrysene were measured at 71% (± 4%) and 78% (±5%) of the reported values. OC and BC fraction analysis
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Dried (60ºC for 24 hr) and ground sediment sub-samples (~10 mg each) were analyzed
for their mass fraction of OC and the sub-fraction BC as well as C/N and C/H ratios, using a Vario EL III CHN elemental analyzer (Elementar). BC was determined by combusting samples at 375º C for 24 hr to remove the more thermally labile OC fractions (CTO375) [12, 24, 48]. Both OC and BC samples were acidified with 200 µL of 0.35 M sulfurous acid (H2SO3) (Baker Analyzed) and then dried at 60ºC for 24 h to remove carbonates before CHN analysis. Three analyses of each sediment sub-sample were performed for each of the two
measurements (BC and OC). Acetanilide (Elemental Microanalysis Limited) was used as a calibration standard for the analytical method. Blanks were run between every six samples. RESULTS AND DISCUSSION Concentrations of three PAHs (phenanthrene, pyrene, and chrysene) were measured in
sediments, clams, and pore waters (using passive samplers) at twelve stations distributed throughout the six sites: three stations at Collins Cove, one station at Pioneer Village, two stations at Lead Mills, three stations at Pines River, one station at Island End, and two stations at Dorchester Bay. All three target chemicals were found in both surface and deeper sediment samples from all twelve stations (n=23) (Supplemental Data, Table S1). Sediment concentrations across all stations ranged over 4 orders of magnitude from 5 ng/gdw for phenanthrene and chrysene at Pioneer Village to 23 µg/gdw phenanthrene at Collins Cove. Generally, sediment concentrations were greater in sediments with high fractions of organic carbon. Pyrene and chrysene concentrations correlated with the organic carbon fractions (fOC) of the sediments (R2 of 0.77 and 0.66), while phenanthrene concentrations were poorly correlated
(R2 = 0.10) Collins Cove and Island End had higher fractions of OC (and BC) and higher ratios of (methyl-phenanthrenes + methyl-anthracenes) to (phenanthrene + anthracene) (
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∑m/z192/∑m/z178) than other sites (1.0 to 1.7 in Collins Cove and Island End while
Environmental Chemistry PREDICTING BIOACCUMULATION OF POLYCYCLIC AROMATIC HYDROCARBONS IN SOFT-SHELLED CLAMS (MYA ARENARIA) USING FIELD DEPLOYMENTS OF POLYETHYLENE PASSIVE SAMPLERS
Loretta A. Fernandez and Philip M. Gschwend
Environ Toxicol Chem., Accepted Article • DOI: 10.1002/etc.2892
Accepted Article
"Accepted Articles" are peer-reviewed, accepted manuscripts that have not been edited, formatted, or in any way altered by the authors since acceptance. They are citable by the Digital Object Identifier (DOI). After the manuscript is edited and formatted, it will be removed from the “Accepted Articles” Web site and published as an Early View article. Note that editing may introduce changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. SETAC cannot be held responsible for errors or consequences arising from the use of information contained in these manuscripts.
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Environmental Chemistry
Environmental Toxicology and Chemistry DOI 10.1002/etc.2892
PREDICTING BIOACCUMULATION OF POLYCYCLIC AROMATIC HYDROCARBONS IN SOFT-SHELLED CLAMS (MYA ARENARIA) USING FIELD DEPLOYMENTS OF POLYETHYLENE PASSIVE SAMPLERS
Running title: Predicting PAH bioaccumulation using PE passive samplers
Loretta A. Fernandez*†‡ and Philip M. Gschwend‡
† Departments of Civil and Environmental Engineering and Marine and Environmental Sciences, Northeastern University, Boston, Massachusetts, USA ‡ Department of Civil and Environmental Engineering, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA
* Address correspondence to [email protected]
Additional Supporting Information may be found in the online version of this article.
This article is protected by copyright. All rights reserved Submitted 11 September 2014; Returned for Revision 11 January 2015; Accepted 13 January 2015
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Abstract: Biota-sediment accumulation factors (BSAF), frequently used to predict tissue concentrations of organisms living within and above sediments contaminated with hydrophobic organic chemicals (HOCs), often produce inaccurate estimates. Hence, freely-dissolved porewater concentrations, CW, have also been investigated as predictors of organism tissue concentrations, but they are more difficult to measure than bulk sediment concentrations (used with BSAF). In situ passive sampling methods make it possible to deduce CW with less effort than is required to measure the value directly, and make it possible to relate CW with tissue concentrations of undisturbed, native organisms. In the present work, polyethylene (PE) passive samplers, containing performance reference compounds (PRCs) (d10-phenanthrene, d10-pyrene, and d12-chrysene), were deployed in diverse sediment beds near Boston, MA for a one-week period. Clams (Mya arenaria) and sediments were then collected from the same sediments. Concentrations of three PAHs (phenanthrene, pyrene, and chrysene) were measured in the pore waters, in clam tissues, and in the bulk sediment. BSAF and PE-deduced CW were used to predict organism tissue concentrations. Ratios of predicted-to-measured values showed that the BSAF method over-predicted tissue concentrations in M. arenaria by up to two orders of magnitude. The PE-deduced CW method resulted in average ratios closer to 1 (0.43 ± 0.26, 3.7 ±
2.5, and 1.1 ± 1.2 for phenanthrene, pyrene, and chrysene, respectively, N=26, uncertainty = ±1σ). This article is protected by copyright. All rights reserved
Keywords: PAH, Passive sampling, Polyethylene, BSAF, Bioavailability
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INTRODUCTION In the search for approaches to determine the ecological risks associated with sediments
contaminated with hydrophobic organic chemicals (HOCs), several indicators of the potential for compounds to be transferred from sediments to biological receptors have been studied. These include biota-sediment accumulation factors (BSAFs) [1-3], and freely-dissolved porewater water concentrations [2, 4-7]. The general idea behind both approaches is that, in an equilibrated
system, knowing the concentration in a single compartment (i.e., sediment organic carbon or porewater), one can estimate concentrations in organism tissues (often assumed to be primarily
in lipids). Organism tissue concentration can be estimated using BSAFs that relate lipidnormalized organism concentrations to organic carbon normalized sediment concentrations Corg = flip BSAF (CSED / fOC)
(1)
where Corg is the concentration in organism tissues (mol/g tissue), flip is the lipid fraction of the
organism tissue (g lipid/g tissue), CSED is the concentration in sediments (mol/g sediments), and fOC is the organic carbon (OC) fraction of the sediments (g OC/g sediment). Similarly, organism concentrations would be predicted from freely-dissolved porewater concentrations, CW (mol/mL water)
Corg = flip Klip-w CW
(2)
where Klip-w is the lipid-water equilibrium partitioning coefficient ((mol/g lipid) / (mol/mL water)).
There are challenges to using either approach. BSAF values depend on organic carbon-
normalized sediment concentrations. However, assuming OC is the only component regulating sediment-water transfer may not be applicable to all sediments. Equilibrium partitioning coefficients for organic carbon and water, KOC, have been observed to range over three orders of
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magnitude for PAHs in field sediments [8-11], most likely due to differing sorptive properties of the many types of materials measured as fOC. Subfractions, including black carbon (BC), oil, pitch, and chars, have been used to refine equilibrium partitioning (EqP) models [12-17], but it is
not always clear how best to assess these subfractions (e.g., Hammes et al. 2007 for BCs). Arp et al. [17] found that KOC tuned to sorption between coal tar and water predicted PAH sediment-
water distribution of PAHs better than other KOC values available in the scientific literature that
have been tuned to other organic carbon pools such as peat, coal, natural organic matter, octanol, or granular activated carbon. As an alternative to organic carbon-normalized sediment concentrations, biological
effects have been compared to concentrations in a more uniform phase of sediments, the pore waters [2, 7, 19]. Directly measuring CW by extracting HOCs from porewater samples, however,
is complicated by the need to physically separate water from sediment solids, then either remove or account for the colloids remaining in porewater samples [20-21]. For this reason, porewater concentrations are often estimated using equilibrium partitioning (EqP) techniques similar to those described above, where CW is assumed to be equilibrated with organic carbon-normalized sediment concentrations CW = CSED / (fOC KOC)
(3)
This method, of course, suffers from the same problems described above, where organic carbon
sub-fractions may have very different sorptive properties. Passive sampling methods offer an alternative by introducing an easily separable and
well-defined organic phase to sediment-water systems, from which CW may be deduced. [22-25] CW = Cpolymer / Kpolymer-w
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(4)
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where, Cpolymer is the concentation in the polymeric sampler, and Kpolymer-w is the polymer-water partition coefficient. This method is easier than the porewater extraction approach and results in accurate CW [26-27]. While passive sampling techniques have been used in laboratory studies to
investigate the relationship between CW, toxicity endpoints, and organism tissue concentrations [28-35], recent advances in passive sampling allow measurement of HOC concentrations in pore waters of intact sediment beds [26, 36-38]. This makes comparison of porewater HOC
concentrations with tissue concentrations of undisturbed, native organisms possible. In several studies, laboratory-raised worms have been exposed to in situ sediments and various methods, including both field and laboratory deployed passive samplers (solid-phase microextraction (SPME) and polyoxymethylene (POM) strips), have been used to predict PAH and PCB uptake
by the organisms [39-41]. Van der Heijden and Jonker [39] found that field deployed SPME performed better than the laboratory measurements in predicting uptake by the oligochaete (Lumbriculus variegatus). In the present study, polyethylene strips containing three performance reference
compounds (PRCs) (d10-phenanthrene, d10-pyrene, and d12-chrysene), which allow for shorter exposure times than are required for full equilibration, were used to measure CW profiles with depth, for three PAHs (phenanthrene, pyrene, and chrysene) in sediments beds at six locations. Field sites were selected for their diverse carbon fractions, variable PAH sources, and the
presence of native organisms, the marine clam Mya arenaria. Sediment PAH concentrations, organic carbon fractions, and published BSAF for M. arenaria, along with PE deduced porewater concentrations, were used to predict organism tissue concentrations. Concentrations
predicted using the two methods were compared to those measured in the native organisms.
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MATERIALS AND METHODS All solvents were Baker Ultraresi-analyzed. Laboratory water was treated with an ion-
exchange and activated carbon system (Aries Vaponics) until 18 MOhm-cm resistance was achieved, followed by UV exposure (TOC reduction unit, Aquafine Corporation). All PAH standards and PRCs were purchased from Ultra Scientific in methanol, acetone, or dichloromethane. Polyethylene (PE) strips were prepared from low-density polyethylene (LDPE) sheets (25
µm from Trimaco). All PE was soaked twice in dichloromethane for 24 h and twice in methanol for 24 h, before soaking twice in water for 24 h. Finite-difference model calculations for diffusion of d12-chrysene from stirred water, through a water-side boundary layer, and into PE indicated that 1 month is sufficiently long for 25 µm thick PE to equilibrate with the solution; but to be sure of even PRC distribution throughout the polymer films, the PE used in the present work was in contact with PRC solution for >7 months (~20 g PE in 1 L aqueous PRC solution
that was agitated by hand ~once per day). Field sampling PE strips were deployed in, and clams and sediments were collected from, six locations
near Boston, MA (Table 1, Supplemental Data, Figure S1) in November 2008. The sites were
selected based on the presence of M. arenaria, previous measurements of PAH concentrations in the sediments, and historical information concerning industrial use of the areas. From north to south, the locations included the following: (a) Collins Cove, Salem, MA – a large but shallow cove approximately 700 m east if the Salem Harbor Power Station, a coal- and oil-fired power plant, (b) Pioneer Village, Salem, MA – a sandy beach along Salem Harbor, approximately 1500
m south of Salem Harbor Power Station, (c) Forest River (Lead Mills), Marblehead, MA – at the
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mouth of a tidal river down-stream of a 19th century lead mill that burned down in 1968, (d)
Pines River, Saugus, MA – along a tidal river adjacent to a closed land fill and a waste-to-energy plant, (e) Island End, Chelsea, MA – a coal-tar contaminated site, and (f ) Dorchester Bay,
Quincy, MA – downwind of the Southeast Expressway (I-93) and near the site of a former garbage incinerator on Spectacle Island. It was expected that these sites would provide a wide range of PAH concentrations and varying organic and black carbon fractions in the sediments. Sampling of all sites was conducted over a period of two weeks, in November 2008,
during low tides, in the intertidal zone. First, PE samplers were deployed at each site directly
adjacent to what appeared to be clam siphon holes in the sediment. PE samplers in aluminum frames were pushed into sediments to a depth of between 4 and 12 cm depending on how far they would go in before meeting resistance (Supplemental Data, Figures S2 and S3). As many of these sites are located in urban areas, a one-week deployment time was selected in order to minimize chances that samplers would be vandalized. Mathematical modeling and the results of previous field testing indicated that this exposure time would be sufficient for the transfer of
measureable masses of target and PRC compounds between sediments and samplers. After being in place for one week later, PE samplers were collected. At that time, clams from the adjacent sediments were collected using a spade, and sediment samples were taken from the surface (approximately top 10 cm) and from a lower layer (approximately 10-20 cm deep). PE strips were rinsed with clean water in the field and placed between aluminum plates, which were then wrapped in aluminum foil. Clams and sediments were placed in glass jars. All samples were returned to the lab on ice. Clams were stored at -20ºC until dissection and extraction, sediments were stored at 4ºC until extraction, and PE were sectioned at 4 cm intervals and
immediately extracted. PE samplers deployed at Pioneer Village were not found on the retrieval
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trip. Instead, clams and adjacent sediment were collected and returned to the lab where three PE strips (approximately 2 cm x 2 cm) were inserted directly into jarred sediments using forceps, and exposed for 32 days. At Island End, clams were not found at the time of PE recovery, so clams collected from 2-3 m away, at the time sampler deployment, were used instead. PE extraction Upon return to the laboratory, PE strips were again rinsed in clean water and swabbed
with a wipe (a hexane-soaked wipe in the case of Island End samplers) to ensure that only absorbed molecules, but not those associated with adhering sediment particles or tarry films, would be quantified. Strips were cut into 4 cm sections, surrogate standards (d10-anthracene, d10-fluoranthene, and d12-benz(a)anthracene) were added, and strips were extracted three times by soaking in 15 mL of dichloromethane overnight. The combined extracts were exchanged into hexane and concentrated to approximately 0.5 mL under a gentle stream of ultra pure grade
nitrogen. Injection standards (d10-acenaphthene, m-terphenyl, and d12-perylene) were added to the extracts before gas chromatography/mass spectrometry (GC/MS) analysis. Calculation of CW from PRC and target analyte concentrations Freely dissolved porewater concentrations were calculated using previously described
methods [26]. Briefly, using PE strips impregnated with PRCs that match target compounds in terms of partition and diffusion coefficients, CW may be calculated from the concentrations of
target chemical (CPE) and PRC (CPRC) in the PE after time, t CW = (CPE (t) × CPRC, init) / (CPRC (t) × KPEW)
(5)
where CPRC, init is the initial concentration of PRC in the sampler, and KPEW is the polyethylene-
water partition coefficient for the PRC and target chemical (Table 2). Clam extractions
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Extraction of PAHs from clams was performed using a method modified from that
described by Yusa et al. [42]. Clams were shucked and dissected to remove stomach, intestines
and most internal organs. The remaining neck, foot, and adductor muscles were then sliced and chopped using two razor blades until a mushy consistency was achieved. Approximate 2 g of the chopped clam were then ground with 1-2 g of precombusted diatomaceous earth (Hyflo Supercel, Sigma Aldrich) until dry and crumbly. Three precombusted GF/B filters (Whatman International Ltd.) were used in the bottom of each 33 mL stainless steel extraction cell. The clam mixture was added to cells between two layers of precombusted Ottawa sand (EMD Chemicals). Surrogate standards were added to the top of the second sand layer before accelerated solvent extraction (ASE) was performed. The remaining homogenized clam tissue was weighed and dried to determine its water fraction. ASE was performed using a Dionex ASE 200 (Dionex Corporation). Each cell was
extracted three times, using dichloromethane:methanol (1:1), heated for 5 min (125ºC) at 1500 psi, and flushed with 60% of cell volume between each extraction. Extracts were collected in amber glass vials with aluminum-lined caps. Approximately 20 g anhydrous Na2SO4 were added to each vial, and extracts were stored overnight at 4ºC. Combined extracts were exchanged into approximately 2 mL hexane using a rotary evaporator (Buchi Rotavapor-R, Brinkman Instruments) before column chromatography and GC/MS analysis. Because ASE also extracted lipids and some proteins from the tissues, a lipid fraction
could not be measured by drying an aliquot of the extract as is normally done when using more gentle Soxhlet extraction methods that more closely mimic the classic Bligh and Dyer method [43]. In the present study, lipid and protein fractions for all clams are assumed to be 5% and 50%, respectively, based on measurements by other researchers. Lohmann et al. [24] measured
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lipid fractions between 5.3 and 9.0% dry weight in clams from Dorchester Bay and 5.8 and 7.6% dry weight in clams from the Saugus River (near Pines River) in 2001 on a dry weight basis, while protein fractions were calculated to be 48%. McDowell and Shea [44] measured lipid fractions between 3.7 and 6.1% for whole clams from the Massachusetts Bay, while Galassi et al. [45] measure an average of 5.1% in clams from the Gulf of Gdansk, in the Baltic Sea. Protein fractions for other aquatic invertebrates have been measured in the range of 50-70% [46]. Uncertainty in lipid-normalized clam tissue concentrations, Corg, was determined by
analyzing multiple clams taken from close proximity. The coefficients of variation (CV) of phenanthrene, pyrene, and chrysene in six clams taken from the Collins Cove, station 3 were 51%, 34%, and 50%, respectively. This uncertainty was assumed for all clam measurements.
Sediment extractions Sediments were homogenized in their collection jars by stirring with a spatula for 5 min,
while removing large stones and shells. Subsamples were taken for drying and weighing to determine water content. Additional subsamples (3-10 g) were taken for ASE. Two precombusted GF-B filters were used in the bottom of each 11-mL stainless steel extraction cell. Wet sediments were added to the cell between two layers of pre-combusted Ottawa sand. Surrogate standards were added to the top sand layer before ASE using the same method described above for clam extractions. Again, extracts were dried overnight at 4ºC using approximately 20 g anhydrous Na2SO4. Extracts were exchanged into hexane and reduced to approximately 2 mL using a rotary evaporator before column chromatography and GC/MS analysis. Column chromatography
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Chromatography columns were prepared in 20 cm long, 1 cm o.d., glass columns with
approximately 30 mL reservoirs. A small plug of glass wool, followed by approximately 2 cm of activated granular Cu0 (20-30 mesh, Baker Analyzed Reagent, J.T. Baker) for columns used with sediment extracts, followed by 5 g fully activated (475 ºC for 24 hr) silica gel (100-200 mesh, EMD Chemicals), followed by approximately 6 g anhydrous Na2SO4 were dry packed into each column [47]. Hexane, 30 mL, was used to flush each column before clam or sediment extracts were charged onto the column. PAHs were collected by running 100 mL of hexane:dichloromethane (9:1 v:v) through the column under slight pressure. The solvent
mixture was exchanged into hexane and reduced to approximately 0.5 mL for clam extracts and approximately 10 mL for sediment extracts. Injection standards (d10-acenaphthene, mterphenyl, and d12-perylene) were added to each extract immediately before GC/MS analysis to
determine sample volume. Gas chromatography/mass spectrometry analysis All extracts were analyzed using gas chromatography/mass spectrometry (GC/MS, JEOL
GCmate, JEOL Ltd.). Splitless 1-µL injections were made onto a 30 m J&W Scientific HP-5MS capillary column (0.25 mm internal diameter with a 0.25 µm film thickness). The injection port temperature was held at 305ºC. The initial column temperature of 70ºC was raised at 20ºC/min until a temperature of 180ºC was reached, and then the temperature was raised 6ºC/min until a
temperature of 300ºC was reached, and remained there for 7.5 min. The MS was operated in selected ion monitoring (SIM) and EI+ modes. Calibration standards containing 20 aromatic compounds including each of the target compounds, surrogates, and injection standards used in the present study, were run every 3 to 5 sample measurements to monitor instrument stability,
determine response factors, and confirm that measurements remained in the linear range for the
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instrument. Repeated observations using the calibration standard indicated the measurement uncertainty for the instrument was typically ±10% relative error. Quality control Target chemicals were above detection limits in all clam, PE, and sediment samples.
Percent recoveries for the surrogate standards ( 1 RSD) were 73 ± 39%, 71 ± 24%, and 60 ± 31% in PE extracts for d10-anthrancene, d10-fluoranthene, and d12-benz(a)anthracene,
respectively. Recoveries in clams were 54 ± 36%, 75 ± 26%, and 81± 29% for d10-anthrancene,
d10-fluoranthene, and d12-benz(a)anthracene, respectively. Recoveries in sediment extracts were 73 ± 36%, 88 ± 32%, and 91 ± 33% for d10-anthrancene, d10-fluoranthene, and d12benz(a)anthracene, respectively. Large ranges in recovery of surrogate standards were expected due to the many transfers and manipulations of the extracts. PRC and target compound concentrations were corrected for recoveries of the corresponding closest-eluting surrogate standard. Extraction and measurement methods were tested for accuracy by measuring standard
reference materials (SRM), NIST SRM 1974b – Organics in Mussel Tissue (Mytilus edulis) and
NIST SRM 1941a – Organics in Marine Sediment. While pyrene and chrysene matched within
uncertainty for the mussel tissue, phenanthrene measurements were 134% of the reported value. Repeated measurements yielded the same result. The high recovery of phenanthrene in the mussel tissues may have been due to contamination of the sample before extraction. For the sediment SRM, phenanthrene matched the reported value within uncertainty, while pyrene and chrysene were measured at 71% (± 4%) and 78% (±5%) of the reported values. OC and BC fraction analysis
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Dried (60ºC for 24 hr) and ground sediment sub-samples (~10 mg each) were analyzed
for their mass fraction of OC and the sub-fraction BC as well as C/N and C/H ratios, using a Vario EL III CHN elemental analyzer (Elementar). BC was determined by combusting samples at 375º C for 24 hr to remove the more thermally labile OC fractions (CTO375) [12, 24, 48]. Both OC and BC samples were acidified with 200 µL of 0.35 M sulfurous acid (H2SO3) (Baker Analyzed) and then dried at 60ºC for 24 h to remove carbonates before CHN analysis. Three analyses of each sediment sub-sample were performed for each of the two
measurements (BC and OC). Acetanilide (Elemental Microanalysis Limited) was used as a calibration standard for the analytical method. Blanks were run between every six samples. RESULTS AND DISCUSSION Concentrations of three PAHs (phenanthrene, pyrene, and chrysene) were measured in
sediments, clams, and pore waters (using passive samplers) at twelve stations distributed throughout the six sites: three stations at Collins Cove, one station at Pioneer Village, two stations at Lead Mills, three stations at Pines River, one station at Island End, and two stations at Dorchester Bay. All three target chemicals were found in both surface and deeper sediment samples from all twelve stations (n=23) (Supplemental Data, Table S1). Sediment concentrations across all stations ranged over 4 orders of magnitude from 5 ng/gdw for phenanthrene and chrysene at Pioneer Village to 23 µg/gdw phenanthrene at Collins Cove. Generally, sediment concentrations were greater in sediments with high fractions of organic carbon. Pyrene and chrysene concentrations correlated with the organic carbon fractions (fOC) of the sediments (R2 of 0.77 and 0.66), while phenanthrene concentrations were poorly correlated
(R2 = 0.10) Collins Cove and Island End had higher fractions of OC (and BC) and higher ratios of (methyl-phenanthrenes + methyl-anthracenes) to (phenanthrene + anthracene) (
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∑m/z192/∑m/z178) than other sites (1.0 to 1.7 in Collins Cove and Island End while
