Skip to main content
NCBI home page
ACS AuthorChoice logoLink to ACS AuthorChoice
. 2024 Aug 5;58(33):14855–14863. doi: 10.1021/acs.est.4c05112

Exposure of Polycyclic Aromatic Hydrocarbons (PAHs) and Crude Oil to Atlantic Haddock (Melanogrammus aeglefinus): A Unique Snapshot of the Mercapturic Acid Pathway

Charlotte L Nakken †,‡,*, Marc H G Berntssen ‡, Sonnich Meier ‡, Lubertus Bijlsma §, Svein A Mjøs †, Elin Sørhus ‡, Carey E Donald ‡
PMCID: PMC11340023  PMID: 39101928

Abstract

graphic file with name es4c05112_0005.jpg

Fish exposed to xenobiotics like petroleum-derived polycyclic aromatic hydrocarbons (PAHs) will immediately initiate detoxification systems through effective biotransformation reactions. Yet, there is a discrepancy between recognized metabolic pathways and the actual metabolites detected in fish following PAH exposure like oil pollution. To deepen our understanding of PAH detoxification, we conducted experiments exposing Atlantic haddock (Melanogrammus aeglefinus) to individual PAHs or complex oil mixtures. Bile extracts, analyzed by using an ion mobility quadrupole time-of-flight mass spectrometer, revealed novel metabolites associated with the mercapturic acid pathway. A dominant spectral feature recognized as PAH thiols set the basis for a screening strategy targeting (i) glutathione-, (ii) cysteinylglycine-, (iii) cysteine-, and (iv) mercapturic acid S-conjugates. Based on controlled single-exposure experiments, we constructed an interactive library of 33 metabolites originating from 8 PAHs (anthracene, phenanthrene, 1-methylphenanthrene, 1,4-dimethylphenanthrene, chrysene, benz[a]anthracene, benzo[a]pyrene, and dibenz[a,h]anthracene). By incorporation of the library in the analysis of samples from crude oil exposed fish, PAHs conjugated with glutathione and cysteinylglycine were uncovered. This qualitative study offers an exclusive glimpse into the rarely acknowledged mercapturic acid detoxification pathway in fish. Furthermore, this furnishes evidence that this metabolic pathway also succeeds for PAHs in complex pollution sources, a notable discovery not previously reported.

Keywords: Bile metabolites, fish, high-resolution mass spectrometry, crude oil, screening, xenobiotic

Short abstract

Laboratory studies on Atlantic haddock present new insights into the detoxification of polycyclic aromatic hydrocarbons by the mercapturic acid pathway.

Introduction

Fish in the vicinity of offshore oil and gas production are constantly exposed to emissions, more specifically produced water (PW) which represents dispersed crude oil and the petroleum constituents polycyclic aromatic hydrocarbons (PAHs).1−3 To facilitate the elimination of PAHs, a series of Phase I and II reactions are initiated to enhance their polarity and aid excretion (Phase III). The primary Phase I reaction is mediated by the cytochrome P450 (CYP) enzymes to produce oxygenated metabolites.4 Further metabolic action is typically performed by Phase II biotransformations, which involve covalent bonding to large polar groups, predominantly sulfate, glucuronic acid, and glutathione (GSH) conjugates.5−8 These biotransformation processes primarily occur in the liver. Due to the subsequent storage of detoxification products in the gall bladder, biliary metabolite analysis have become a well-established biomarker of recent PAH exposure in fish.9 Thus, mapping the bile metabolites becomes crucial in assessing PAH exposure, as it provides a snapshot of PAHs prior to excretion. However, since these analyses have typically focused on hydroxylated metabolites along with deconjugated sulfate and glucuronide forms,10 little emphasis has been given to other metabolic products, including the GSH compounds.

Conjugation to the tripeptide GSH is initiated by a nucleophilic addition reaction with an electrophilic xenobiotic, like a Phase I PAH diol epoxide produced by CYP.11 Glutathione S-transferases (GST) typically catalyze this process (0, Figure 1), though the reaction can also occur spontaneously.12GSH can further enter the major biotransformation pathway known as the mercapturic acid pathway (MAP),13 which consists of four different conjugates: (1) GSH, (2) Cysteinylglycine (cysgly), (3) Cysteine (cys), and (4) N-acetyl-l-cysteine (Mercapturic acid, MA).13,14 Briefly, the cleavage of glutamic acid by gamma-glutamyl transferase (GGT) forms the second conjugate, cysgly. Next, cleavage of the glycine moiety by cysteinylglycine dipeptidase (CGDP) results in the amino acid cys. Acetylation catalyzed by N-acetyltransferase (NAT) leaves the ultimate conjugate, MA.(14,15)

Figure 1.

Figure 1

Open in a new tab

Proposed conjugates in the mercapturic acid pathway (MAP): (1) Glutathione (GSH), (2) Cysteinylglycine (cysgly), (3) Cysteine (cys), and (4) N-acetyl-l-cysteine (Mercapturic acid, MA). R = xenobiotic, GST = glutathione S-transferase, GGT = gamma-glutamyl transferase, CGDP = cysteinylglycine dipeptidase, NAT = N-acetyltransferase.14

Research in mammals indicates that the MAP is a central GSH-mediated metabolization pathway for PAHs as they can form various metabolites in the MAP across multiple biological contexts, encompassing human research, rodents, and cell studies.16−21 While glucuronide and sulfate metabolites are the main focus of Phase II pathways investigated in fish, GSH-derived metabolites have been comparatively less investigated.10 GST activity can serve as a biomarker to assess the effects of PAHs in fish,22 indicating that GSH plays a role in PAH detoxification. The tentative identifications of GSH metabolites from benzo[a]pyrene in English sole and starry flounder supports this conclusion.23−25 Still, our current knowledge base suggests limited insight into the potential transformation of GSH conjugates into cysgly, cys, or MA conjugates via the MAP in PAH exposed fish and their detection using mass spectrometry.

In this work, analysis by ion mobility combined with high-resolution mass spectrometry (IM-HRMS), aimed at mapping the conventional metabolites in fish bile, revealed numerous unexpected metabolites originating from the MAP. Based on our data set from individual PAH exposed fish, we developed a method to establish a MAP metabolite library designed for application in suspect screening analyses of fish exposed to complex oil mixtures. These metabolites are presented in an interactive library along with a modified nontarget screening strategy. The qualitative metabolite analysis herein contributes to improved insight into PAH metabolism and presents MAP metabolites not previously reported from oil-exposed fish. Hence, these newly identified products should be included in future investigations to better understand the impact of chronic emissions from oil production.

Methods

Chemicals

An instrument calibration solution (CCS Major Mix) and leucine enkephalin, used as the lock mass, were purchased from Waters (Manchester, U.K.). HPLC- and UHPLC-grade solvents (Chromasolv) were obtained from Honeywell (Seelze, Germany). Phenanthrene (CAS 85–01–8), dimethyl sulfoxide (DMSO), and tricaine methanesulfonate (MS-222) were purchased from Sigma-Aldrich (Oslo, Norway). 1-Methylphenanthrene (CAS 832–69–9), 1,4-dimethylphenanthrene (CAS 22349–59–3), benz[a]anthracene (CAS 56–55–3), chrysene (CAS 218–01–9), benzo[a]pyrene (CAS 50–32–8), and dibenz[a,h]anthracene (CAS 53–70–3) were purchased from Chiron (Trondheim, Norway). 1-Methylnaphtalene (CAS 90–12–0), 1,4-dimethylnaphtalene (CAS 571–58–4), anthracene (CAS 120–12–7), and 1-methylpyrene (CAS 2381–21–7) were purchased from LGC Standards AB (UK).

Two different complex petroleum related PAH mixtures were designed to represent the PAH composition in heavy weathered crude oil and production water (PW). Details about the composition and the making of these mixtures are given in Meier et al.26 The two petroleum mixtures contained significant quantities of various oil compounds, with PAHs contributing 1.5% (crude oil) and 1.1% (PW) of the weight. The PAH profile (% of total PAHs) in the weathered crude oil was dominated by 3-ring PAHs (93%), some 4-ring PAHs (5.3%) and 5-ring PAHs (1%). The PW-mixture was dominated by 2-ring PAHs (77%), with some 3-rings PAHs (22%) and trace amounts of heavy PAHs (>0.6% of 4–5 ring PAHs). The weathered crude oil was prepared using 45% of Gullfaks oil (Norwegian North Sea, Tampen area) distillation fractions (boiling points: 320–375 °C) and 55% distillation fractions (boiling points: 375–400 °C) with the addition of 0.06% pyrene. The PW-mixture was prepared using 90% of a cyclo-hexane extract of PW from Statfjord A (Norwegian North Sea, Tampen area) and 10% of distillation fractions (boiling points: 240–320 °C) of Gullfaks oil.

Fish Exposure and Bile Sample Preparation

The study was performed using Atlantic haddock (Melanogrammus aeglefinus) as it is an important ecological and commercial species with habitat in areas with chronic emissions of PAHs in the North Sea.2 Exposure experiments were performed similarly as described in Meier et al.,26 with adult haddock supplied from a brood stock at the Austevoll Research Station. All animal experiments were approved (FOTS ID 5924) by the Norwegian Animal Research Authority. All methods were performed in accordance with the approved guidelines. The Austevoll Research station has the following permission for the catch and maintenance of Atlantic haddock: H-AV 77, H-AV 78, and H-AV 79 (given by the Norwegian Directorate of Fisheries). The Austevoll Research station has a permit to run as a Research Animal facility using fish (all developmental stages) with code 93.

To establish the proof of concept for PAH detoxification, the study utilized injected exposure concentrations that would ensure the detection of all potential PAH metabolites. The study included 14 treatments representing a suit of petroleum relevant PAHs or complex oil mixtures: 5 alkylated PAHs, 6 unsubstituted PAHs, 2 complex oil mixtures, and a control. Fish were exposed to a high dose of an individual PAH or alkylated PAH compound dissolved in DMSO and fish oil corresponding to a dose of 5 mg/kg fish by intraperitoneal injection (number of bile extracts from individually exposed fish in brackets): 1-methylnaphthalene (2), 1,4-dimethylnaphtalene (1), anthracene (2), phenanthrene (3), 1-methylphenanthrene (2), 1,4-dimethylphenanthrene (2), chrysene (2), benz[a]anthracene (2), 1-methylpyrene (2), benzo[a]pyrene (2), dibenz[a,h]anthracene (2). The control treatment (1) had no PAHs administered. The two complex oil mixtures, crude oil (4) and PW (4), were injected directly without DMSO or fish oil. The exposure doses were 7.4 ± 0.3 mg PAH/kg fish for crude oil and 5.3 ± 0.3 mg PAH/kg fish for PW. Fish were anesthetized before injection (60 mg/L MS-222).

The fish were euthanized 3 days postinjection using a high dose of anesthetic. Bile was kept frozen in liquid nitrogen and stored at −80 °C until metabolite extraction following the protocol of da Silva et al.27 and was later stored in the dark at −20 °C until metabolite analysis. In brief, the sample preparation involved methanol extraction, including phospholipid removal, followed by hydrolysis with glucuronidase and sulfatase enzyme (pH 5, 1 h, 40 °C), and solid phase extraction. Each bile extract represents one fish, exposed to one PAH or mixture treatment. Two bile samples had less than the desired bile volume (<50 uL; one from each of the treatments 1-methylnaphthalene and chrysene) and were therefore discarded.

Instrumentation

Analyses were performed on a Waters Acquity UPLC system interfaced to an ion mobility quadrupole time-of-flight mass spectrometer (LC-IM-QTOF MS) (Waters, Milford, MA, USA). The reverse phase C18 ACQUITY UPLC BEH column (Waters, 100 × 2.1 mm, 1.7 μm) was kept at 45 °C. A binary mobile phase, composed by methanol (B) and water (A), was employed at a flow rate of 0.45 mL/min following the gradient program: 0.0 min, 30% B; 0.1 min, 30% B; 20.10 min, 80% B; 20.2 min, 98% B; 24.0 min, 98% B, 24.1 min, 30% B. The column equilibrated until 29.0 min. The sample manager was set at 10 °C, and the injection volume was 5 μL.

The LockSpray electrospray ionization was set at 2.6 kV in negative ionization mode; cone voltage of 40 V; desolvation temperature of 475 °C; source temperature of 110 °C; desolvation gas flow (N2) of 950 L/h; and cone gas flow (N2) of 50 L/h. Automatic lock mass correction (leucine enkephalin; m/z 554.2615) was used for accurate mass correction at half-minutes intervals. The analyzer mode was set to sensitivity, and the acquisition mode was in high definition MSE. A scan time of 0.3 s was set in the range of m/z 50–800. The low energy scan was set at 6 eV to monitor the deprotonated molecules, while the high energy ramp (8–45 eV) monitored their corresponding fragment ions. Nitrogen was employed as drift gas and argon as collision-induced dissociation gas.

Originally, samples from exposures to single compounds were analyzed. A year later, screening of the bile extracts collected from the crude oil and PW exposed fish were performed using the new established library.

Data Processing and Library Building

Data acquisition and processing were performed in UNIFI software (version 1.9.4.053, Waters). To enhance the data processing and reduce the number of false positives, the retention time and mass were constrained by the detection information on the heaviest and least polar PAH in the study. Accordingly, based on the detection information on dibenz[a,h]anthracene, data filtering and processing were focusing on compounds (<700 Da) eluting within 10 min. Intensity thresholds for low and high energy ions were set at 5 and maximum candidates (i.e., detected m/z) to keep for screening was set to 15 000 per sample. As presented in Figure 2, the experimental design involves a modified version of the refined nontarget workflow by Bijlsma et al.28 The single-exposure study and the sample preparation are described in the section Fish Exposure and Bile Sample Preparation.

Figure 2.

Figure 2

Open in a new tab

Schematic overview of the experimental workflow for constructing a custom library of novel metabolites, based on controlled single-exposure experiments and data processing with improved confidence from predicted collision cross section (CCS) values (adapted from Bijlsma et al.28).

Following the acquisition of the bile extracts, several data processing steps were carried out. First, a filter targeting the suspected base peaks in the spectra, i.e. [M+S–H]− and [M+S+H2O2–H]−, corresponding to a PAH-thiol and a dihydroxy-dihydro-PAH-thiol, was applied for peak picking. Concurrently, metabolites from MAP were explored using the transformation tool in UNIFI for predicting potential biotransformation products of the treatment compound. Phase I transformations included oxidation (+O), double oxidation (+O2), and dihydrodiol formation (+H2O2). Phase II transformations consisted of conjugation with GSH (+C10H15N3O6S, + C10H15N3O5S), cysgly (+C5H8N2O3S), MA (+C5H7NO3S), and cys (+C3H5NO2S, + C3H5NOS). The maximum number of transformations was set at two Phase I and one Phase II transformations.

Peaks matching both the mass of a biotransformation product and a target fragment ion were manually investigated for common characteristics, such as common neutral losses (CNLs) and common fragment ions. Several fragments and CNLs were noted but were excluded from the method due to a lack of specificity. This same procedure was applied to all single compound samples, resulting in the CNLs and fragments presented in the results (Table 1). These characteristics were incorporated into an in-house-built accurate mass screening workflow within the UNIFI software to facilitate tentative identifications. The chemical structures of two potential isomers fitting the accurate mass were drawn in ChemDraw (version 20.1.0.112) and uploaded to the UNIFI software to aid in fragment elucidation. Collision cross section (CCS) predictions for additional identification confidence were performed by the CCSH- model presented by Celma et al.29 Prediction accuracy for the CCSH- model was ±5.86% (relative error within the 95% confidence interval).

Table 1. Key Fragment Ions (m/z) and Common Neutral Losses (Monoisotopic Mass) Employed for Selective Screening of Metabolites in the Mercapturic Acid Pathway (MAP) in Negative Ionization Mode.

Conjugate Fragment ion (m/z) Common neutral loss (Da)
Glutathione (GSH) C10H14N3O6 (272.08881) C10H19N3O7S (325.09437)
C10H17N3O7 (291.10665)
C10H16N3O6 (274.10391)
C10H15N3O6 (273.09609)
C5H10N2O4 (162.06406)
C5H10N2O3 (146.06914)
Cysteinylglycine (cysgly) C5H9N2O3S (177.03394) C5H12N2O4S (196.05178)
C5H7N2O3 (143.04622) C5H10N2O4 (162.06406)
C5H8N2O3 (144.05349)
Cysteine (cys) C5H4NO2S (141.99682) C3H9NO5 (139.04807)
C3H5NO2 (87.03203)
N-acetyl-l-cysteine (Mercapturic acid; MA) C5H6NO3 (128.03532) C5H7NO3 (129.04259)
Open in a new tab

Due to the lack of available reference standards for any of the reported metabolites, we placed a high priority on reducing the risk of false positives and gaining the highest possible confidence in our identifications. Hence, we implemented strict identification criteria for inclusion in our custom scientific library in UNIFI: (i) mass accuracy ≤ 5 ppm for the suspect ion, (ii) a characteristic CNL, (iii) a characteristic conjugate fragment ion, (iv) CCS deviation within model limits, and (v) not detected in the control. Criteria (i)-iv) were automated in the data processing by using an accurate mass screening workflow in UNIFI. The custom library serves as a repository of detection information (m/z, fragment ions, retention time, CCS, spectra). Afterward, it can be imported and implemented in data processing in other batches, as demonstrated in our case with crude oil and PW samples.

All identified metabolites were collected in a library, presented here as interactive files in the Supporting Information. The library (.html and .pdf) presents the following information: (1) Structural information: formula, neutral mass, predicted CCS, suggested chemical structure, and simplified molecular-input line-entry system (SMILES), 2) Detection information: observed m/z, retention time, CCS, delta difference for acquired vs predicted CCS, adduct, and low- and high energy mass spectra. The exact position of the substitutions and conjugates are indeterminable. Compounds with the same conjugation were assigned numerical names with increasing number Phase I substitutions (e.g., cysteinylglycine I, II, and III). If isomers were detected, the library comprised the two largest peak areas, denoted as A and B, respectively. Detection information (m/z, retention time, and CCS) is reported in Table S1. Due to the lack of analytical standards and overall dose-dependent enzymatic deconjugation of most newly described metabolic pathways, the results are qualitative rather than quantitative.

Results and Discussion

Bile Extracts Captured Unique Insight into the Mercapturic Acid Pathway

Implementation of the screening strategy herein led to the detection of 33 bile metabolites involving all four conjugates in the MAP: (1) GSH, (2) cysgly, (3) cys, and (4) MA (Figure 1). Metabolites were identified in the following treatments: dibenz[a,h]anthracene, benzo[a]pyrene, chrysene, benz[a]anthracene, 1,4-dimethylphenanthrene, 1-methylphenanthrene, phenanthrene, and anthracene. Detection information and suggested structural details are presented in the library in the Supporting Information. No identifications were performed in bile extracts from the treatment groups 1-methylnaphthalene, 1,4-dimethylnaphtalene or 1-methylpyrene.

A visual overview of the 33 identified metabolites across the single-exposure treatments is provided in Figure 3. GSH and cysgly represent the first and second steps of the pathway and were notably the most commonly observed metabolite types overall.

Figure 3.

Figure 3

Open in a new tab

Graphical presentation of number metabolites from the mercapturic acid pathway (MAP) detected in the treatment groups dibenz[a,h]anthracene, benzo[a]pyrene, chrysene, benz[a]anthracene, 1,4-dimethylphenanthrene, 1-methylphenanthrene, phenanthrene, and anthracene.

The results from the benzo[a]pyrene treatment are of particular interest since it is a well-studied carcinogen known to produce highly reactive metabolites that can covalently bond to DNA, i.e. form DNA-adducts.30,31 The presence of DNA adducts provides an indication of previous exposure and the potential induction of carcinogenesis.32GSH can participate in conjugation reactions with benzo(a)pyrene metabolites and thus prevents potential DNA damage by reducing the available reactive species.33 In fact, there is a strong correlation between the level of GSH and DNA-adducts.34 In the benzo[a]pyrene treatment, two isomers of benzo[a]pyrene diol epoxide connected to cysgly were detected (library item Benzo[a]pyrene cysteinylglycine A-B). Our findings suggest that the MAP successfully converts potential mutagenic species into a detoxified metabolite. However, the exact configuration of the diol epoxide remains unknown.

Chrysene yielded metabolites of all four conjugation types, which allows for mapping of the entire pathway, as depicted in Figure 4. The metabolic route can be initiated by the cytochrome P450 enzymes which can form a highly reactive epoxide metabolite.12 By conjugation with GSH, the epoxides are detoxified (1) and can continue the pathway as a cysgly conjugate (2) in the MAP. In the case outlined in Figure 4, there is an additional dihydrodiol moiety present compared with the previous GSH step (1). Our hypothesis suggests that Phase I oxidations may occur both prior to and following the conjugation events. Moving forward to a cys metabolite, the same Phase I moieties were observed (3). Further metabolic processes can give rise to MA (4). However, no indication of other functional groups was observed in the mass spectra of this compound. A proposed explanation is the elimination of water, leaving the GSH residue singly on the ring.35,36 This observation, where only the MAP conjugate was observed on the PAH, occurred in 13 of the 33 compounds identified in our library.

Figure 4.

Figure 4

Open in a new tab

Chrysene (Chr) can be bioactivated by cytochrome P450 (CYP) enzymes to an epoxide and detoxified by the mercapturic acid pathway (MAP), producing metabolites with (1) Glutathione (GSH), (2) Cysteinylglycine (cysgly), (3) Cysteine (cys), and (4) Mercapturic acid (MA) conjugates.

The elimination of PAHs via bile is affected by environmental factors and dynamic physiological processes.37 It should be emphasized that the metabolism in MAP is a continuous process, and our sampling only captured a snapshot of the metabolic products. In principle, it is not anticipated that all these metabolites will persist simultaneously in the bile for an extended duration, as observed in the Chrysene treatment (Figure 4). The MAP is an interorgan process where the formation of the MA conjugate usually occurs in the kidney.15 These S-conjugates can also re-enter the liver-bile detoxification system and prolong their presence by enterohepatic circulation.13 Therefore, the detections presented in Figure 4 of Chrysene metabolites are unique, as all versions in the MAP are available in one bile extract. These conjugates exhibit high polarity, making them readily excretable in urine and less likely to exist in bile.

Analytical Workflow

The employed screening strategy integrated both a specific CNL and a diagnostic fragment ion for each conjugate, which allowed for selective data filtering. The characteristic fragmentation behaviors of the four conjugation types (Figure 1) are reported in Table 1. The CNL enables the detection of the characteristic PAH metabolite (Phase I; with or without sulfur retained), while a fragment ion confirms the identity of the conjugate.

The candidates matching the characteristics in Table 1 were further prioritized by incorporating predicted CCS values for extra identification confidence.38 The presented strategy accomplishes the detection of MAP metabolites in one injection of the sample using distinct combinations of compound characteristics. Based on the observed fragmentation patterns of the conjugates, we expect that the presented workflow also can be employed in xenobiotic scenarios beyond PAHs.

The conventional strategy for the detection of GSH and MA metabolites is a CNL of 129 Da.39,40 Another approach is negative precursor ion scanning of m/z 272 targeting GSHs.41−43 In contrast to the conventional methods that use CNL or precursor scans followed by product ion scans for structural elucidation work, additional injections were not necessary with MSE. Efforts to find well-documented analytical methods for cysgly and cys beyond the thorough elucidation presented by Levsen et al.36 were unsuccessful. We hypothesize that this diminished emphasis is due to the expected efficiency of the MAP, where the significance of intermediates is overshadowed by the initial GSH and the ultimate conjugate MA.

Two specific analytical challenges arise when analyzing the metabolites presented herein. First, it concerns the similarity in mass of conjugates: conjugation with cysgly (+C5H8N2O3S) and O-glucuronidation (+C6H8O6) adds 176.02556 and 176.03209 Da, respectively. Both have the same CNL strategy of monitoring 176 Da losses.36 Thorough investigation of spectra is therefore crucial to make the correct annotation based on fragment ions. Second, two key fragments of the metabolites have striking similarity in mass. Coincidentally, the mass of a biotransformed PAH into either a PAH-diol (containing two −OH groups) or a PAH-thiol (−SH group) differs by only 0.018 Da, where the difference appears in the second decimal. Hence, it is unlikely to make the same discovery with low resolution mass spectrometry since the technique lacks the ability to distinguish two peaks of slightly different mass-to-charge ratios. Benz[a]anthracene cysgly was the first metabolite to be elucidated after recognizing this small, but crucial mass difference. PAH-thiols were implemented in the workflow by reprocessing the data (see Data Processing and Library Building). This facilitated real-time investigations, eliminating the need for sample reinjection, and thereby conserving both time and chemicals.

The limited identifications of MA and cys (Figure 3) could be attributed to biological factors as discussed earlier or the specificity of the method. For instance, cys did not have diagnostic fragments beyond m/z 141, corresponding to the inclusion of carbon bonds from PAH. Ideally, we would have observed a fragment corresponding to the entire conjugate, but this was not present. However, not all conjugates exhibit the same characteristic loss during fragmentation44 or produce fragments at detectable levels. All of the involved conjugates are S-conjugates, also known as thioether conjugates. We observe variations in fragmentation pattern despite identical conjugation type, suggesting a strong influence of the aromatic system and the number of Phase I modifications on the PAH. Ma et al.45 reported 98% specificity for 121 CNL for cys. In our case, sulfur is retained, resulting in losses of 87 and 139 Da (Table 1).

Application of Library to Oil-Polluted Fish

Environmental samples typically contain PAHs from various sources. Hence, two relevant pollution sources were considered in this study: crude oil and PW. Increased expression of GST has been observed following oil exposure studies in early life stages of haddock.46 Thus, we anticipated the presence of MAP metabolites in marine fish exposed to crude oil mixtures. The detections corroborated the hypothesis as GSH and cysgly candidates matching alkylated 3-ring PAHs and a 4-ring PAH were identified (Table 2). This demonstrates that the formation of such metabolites is not limited to single compound exposures but also occurs in realistic and highly complex mixtures such as crude oil.

Table 2. Tentatively Identified Polycyclic Aromatic Hydrocarbon (PAH) Metabolites from the Mercapturic Acid Pathway (MAP) produced by Atlantic haddock after Exposure to Crude Oil.

PAH type Conjugate Formula Observed mass (m/z) Observed common neutral loss (Da) Observed diagnostic fragment(s) (m/z)
Methylated 3-ring PAH Cysgly C20H20N2O3S 367.1117 144.0532 143.0462
GSH C25H27N3O6S 496.1547 273.0960 254.0784, 272.0887
Dimethylated 3-ring PAH Cysgly C21H22N2O3S 381.1277 144.0533 143.0463
GSH C26H29N3O6S 510.1708 273.0961 254.0784, 272.0889
  C26H29N3O6S 510.1703 273.0960 272.0889
4-ring PAH Cysgly C23H20N2O3S 403.1124 144.0535 143.0465
Open in a new tab

Petrogenic PAHs dominate in crude oil polluted samples, characterized by a predominant presence of alkylated low molecular PAHs (2- and 3-ring) and modest contributions from heavier unsubstituted PAHs.47 Based on their large contributions in crude oil, the 3-ring PAHs are expected to be phenanthrene derivatives and the 4-ring is suspected to be pyrene or chrysene.26 Detection information (m/z, retention time, CCS) for all of the metabolites is reported in Table S2.

The formulas presented are the best fitted formula according to obtained m/z and isotope pattern using the elemental composition calculator in UNIFI. As observed in our metabolite library, the MAP metabolites exhibit a wide range of retention times, highlighting the significance of the substitution position in terms of polarity. Given the resemblance in detection information among isomers, identifying the compounds becomes impractical. We therefore assign generic names to the compound due to the numerous potential options available.

The second oil pollution treatment, PW, represents operational discharge containing the most water-soluble, low molecular weight PAHs.26 Given these considerations, we suspected a predominant presence of naphthalene metabolites in fish exposed to PW. Yet, no metabolites from C0-, C1-, or C2-naphthalene were elucidated in bile from either single compound, crude oil, or PW exposures. In fact, no MAP metabolites were detected by our workflow. Effective biotransformation processes can result in low internal concentration in fish despite high contribution in the polluted water,48 so we hypothesize that rapid detoxification following the exposure could be a contributing factor. We also acknowledge the possibility of loss of compounds during extraction (e.g. volatilization) or due to instrument limitations (i.e., inadequate level of detection). It is worth mentioning that it initially appeared to be a match with 1-methylphenanthrene, but upon closer examination of the base peak and isotopic pattern, the candidate was dismissed. Based on the detection of characteristic fragments m/z 272, 179, and 128, we are confident that the compound contains GSH. However, the exact source linkage from which parent compound it originated remains ambiguous.

Diversification of Metabolite Types Can Improve Environmental Monitoring

Enzymatic deconjugation to reconstruct the original Phase I metabolites is typically performed for quantitative investigations of PAH exposure.10 It is noteworthy that metabolites from the MAP could be present in samples using conventional extraction methods: the deconjugation enzymes (i.e., β-glucuronidase and aryl sulfatase) that recovers Phase I metabolites, do not hydrolyze these conjugates.10,25 As a result, they may potentially go unnoticed in the mixture. The inclusion of GSH metabolites can be accomplished by an extra enzymatic deconjugation step by addition of gamma-glutamyltranspeptidase49 or by performing acidic hydrolysis.50 This extra effort during sample preparation would give a more accurate overview of metabolite quantities and ultimately provide more thorough assessments of PAH exposure. An augmentation like this could enhance detection sensitivity at lower concentrations, thereby enabling the adoption of more refined analytical techniques.

The present study is a qualitative and nontargeted assessment of metabolites, identifying new MAP compounds previously unknown in fish but recognized in mammalian research.14 Ayala et al.16 quantitatively measured the urinary excretion of naphthalene metabolites and found that GSH and MA accounted for >60% of the total measured metabolites in mice. MAs have also been identified as urinary metabolites in humans.51 Contrary to the findings of Ayala et al.,16MAs have been characterized as minor metabolites in cigarette smokers.35 However, that study measured only one out of the four potential conjugates in the MAP. Similarly, Willet et al.49 observed a minimal presence of GSH metabolites in fish bile after exposure to benzo[a]pyrene. The present results imply that these limited detections might be due to the sequential metabolization of GSH into cysgly, cys, or MA derivates which are subsequently eliminated through Phase III pathways.14 Thus, the current identifications warrant further investigation into the enzymatic pathways initiated by GST.

This work demonstrates the discovery of novel metabolites by utilizing the power of IM-HRMS and the possibility to perform retrospective analysis. The controlled exposures of individual components in our study allowed us to develop a comprehensive metabolite library using our own sample set. This innovative approach proved instrumental in identifying previously undiscovered metabolites in fish exposed to crude oil. Specifically, our method enhances the scope of PAH metabolite analyses, offering a new approach for detecting and characterizing a wider range of metabolites compared to traditional methodologies.

The objective of the presented bile extracts was to qualitatively profile PAH metabolites to gain a deeper understanding of the metabolic pathways involved in their detoxification. Due to minimal reporting on metabolites of the MAP in the literature, these were not prioritized but discovered incidentally. Hence, it is likely that these xenobiotic metabolites frequently escape detection, leading to underestimated exposure assessments. In essence, this metabolite class should be incorporated into future investigations to enhance our understanding of the impact of chronic emissions of contaminants. By employing the methods outlined in this work, high confidence tentative identifications can be achieved.

Acknowledgments

This work was financed through the water column monitoring program (Norwegian State Pollution Control Agency) and by Statoil (now Equinor ASA). L.B. acknowledges grant RYC2020-028936-I funded by MCIN/AEI/10.13039/501100011033 and by “ESF Investing in your future”. The authors thank Denis da Silva for training in the extraction protocol, and Stig Ove Utskot at Austevoll Research Station for breeding and management of the fish. The authors used ChatGPT by OpenAI to rephrase and improve the language.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.est.4c05112.

  • Metabolite library (PDF, ZIP)

  • Detection results of identified metabolites and detection results from crude oil exposure (PDF)

The authors declare no competing financial interest.

Supplementary Material

es4c05112_si_001.pdf (1.9MB, pdf)
es4c05112_si_002.zip (1.9MB, zip)
es4c05112_si_003.pdf (168.9KB, pdf)

References

  1. Bakke T.; Klungsøyr J.; Sanni S. Environmental impacts of produced water and drilling waste discharges from the Norwegian offshore petroleum industry. Mar. Environ. Res. 2013, 92, 154–69. 10.1016/j.marenvres.2013.09.012. [DOI] [PubMed] [Google Scholar]
  2. Balk L.; Hylland K.; Hansson T.; Berntssen M. H.; Beyer J.; Jonsson G.; Melbye A.; Grung M.; Torstensen B. E.; Borseth J. F.; Skarphedinsdottir H.; Klungsøyr J. Biomarkers in natural fish populations indicate adverse biological effects of offshore oil production. PLoS One 2011, 6 (5), e19735 10.1371/journal.pone.0019735. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Beyer J.; Goksøyr A.; Hjermann D. Ø.; Klungsøyr J. Environmental effects of offshore produced water discharges: A review focused on the Norwegian continental shelf. Mar. Environ. Res. 2020, 162, 105155–105155. 10.1016/j.marenvres.2020.105155. [DOI] [PubMed] [Google Scholar]
  4. Goksøyr A.; Forlin L. The cytochrome P-450 system in fish, aquatic toxicology and environmental monitoring. Aquat. Toxicol. 1992, 22 (4), 287–311. 10.1016/0166-445X(92)90046-P. [DOI] [Google Scholar]
  5. de Maagd P. G.-J.; Vethaak A. D.. Biotransformation of PAHs and Their Carcinogenic Effects in Fish. In The Handbook of Environmental Chemistry Vol. 3 Part J. PAHs and Related Compounds; Neilson A. H., Ed.; Springer-Verlag: Berlin, Heidelberg; 1998; pp 265–309. [Google Scholar]
  6. Ikenaka Y.; Oguri M.; Saengtienchai A.; Nakayama S. M. M.; Ijiri S.; Ishizuka M. Characterization of phase-II conjugation reaction of polycyclic aromatic hydrocarbons in fish species: Unique pyrene metabolism and species specificity observed in fish species. Environ. Toxicol. Pharmacol. 2013, 36 (2), 567–578. 10.1016/j.etap.2013.05.018. [DOI] [PubMed] [Google Scholar]
  7. Solbakken J. E.; Palmork K. H.; Neppelberg T.; Scheline R. R. Urinary and Biliary Metabolites of Phenanthrene in the Coalfish (Pollachius virens). Acta Pharmacol. Toxicol. 1980, 46 (2), 127–132. 10.1111/j.1600-0773.1980.tb02431.x. [DOI] [PubMed] [Google Scholar]
  8. Pampanin D.; Sydnes M.. Polycyclic Aromatic Hydrocarbons a Constituent of Petroleum: Presence and Influence in the Aquatic Environment. In Hydrocarbon; Vladimir K.; Anton K., Eds.; IntechOpen: Rijeka, 2013; pp 83–118. [Google Scholar]
  9. Dearnley J. M.; Killeen C.; Davis R. L.; Palace V. P.; Tomy G. T. Monitoring polycyclic aromatic compounds exposure in fish using biliary metabolites. Crit. Rev. Environ. Sci. Technol. 2022, 52 (4), 475–519. 10.1080/10643389.2020.1825895. [DOI] [Google Scholar]
  10. Beyer J.; Jonsson G.; Porte C.; Krahn M. M.; Ariese F. Analytical methods for determining metabolites of polycyclic aromatic hydrocarbon (PAH) pollutants in fish bile: A review. Environ. Toxicol. Pharmacol. 2010, 30 (3), 224–244. 10.1016/j.etap.2010.08.004. [DOI] [PubMed] [Google Scholar]
  11. Sundberg K.; Widersten M.; Seidel A.; Mannervik B.; Jernström B. Glutathione Conjugation of Bay- and Fjord-Region Diol Epoxides of Polycyclic Aromatic Hydrocarbons by Glutathione Transferases M1–1 and P1–1. Chem. Res. Toxicol. 1997, 10 (11), 1221–1227. 10.1021/tx970099w. [DOI] [PubMed] [Google Scholar]
  12. Seidegard J.; Ekström G. The Role of Human Glutathione Transferases and Epoxide Hydrolases in the Metabolism of Xenobiotics. Environ. Health Perspect. 1997, 105, 791. 10.2307/3433285. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Cooper A. J. L.; Hanigan M. H.. Metabolism of Glutathione S-Conjugates: Multiple Pathways. In Comprehensive Toxicology, Third ed.; McQueen C. A., Ed.; Elsevier: Oxford, 2018; pp 363–406. [Google Scholar]
  14. Potęga A. Glutathione-Mediated Conjugation of Anticancer Drugs: An Overview of Reaction Mechanisms and Biological Significance for Drug Detoxification and Bioactivation. Molecules 2022, 27 (16), 5252. 10.3390/molecules27165252. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Hanna P. E.; Anders M. W. The mercapturic acid pathway. Crit. Rev. Toxicol. 2019, 49 (10), 819–929. 10.1080/10408444.2019.1692191. [DOI] [PubMed] [Google Scholar]
  16. Ayala D. C.; Morin D.; Buckpitt A. R. Simultaneous quantification of multiple urinary naphthalene metabolites by liquid chromatography tandem mass spectrometry. PLoS One 2015, 10 (4), e0121937 10.1371/journal.pone.0121937. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Hecht S. S.; Villalta P. W.; Hochalter J. B. Analysis of phenanthrene diol epoxide mercapturic acid detoxification products in human urine: relevance to molecular epidemiology studies of glutathione S-transferase polymorphisms. Carcinogenesis 2008, 29 (5), 937–943. 10.1093/carcin/bgn015. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Hecht S. S.; Berg J. Z.; Hochalter J. B. Preferential Glutathione Conjugation of a Reverse Diol Epoxide Compared to a Bay Region Diol Epoxide of Phenanthrene in Human Hepatocytes: Relevance to Molecular Epidemiology Studies of Glutathione-S-Transferase Polymorphisms and Cancer. Chem. Res. Toxicol. 2009, 22 (3), 426–432. 10.1021/tx800315m. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Horning M. G.; Sheng L. S.; Nowlin J. G.; Lertratanangkoon K.; Horning E. C. Analytical methods for the study of urinary thioether metabolites in the rat and guinea pig. J. Chromatogr. A 1987, 399, 303–319. 10.1016/S0021-9673(00)96133-8. [DOI] [PubMed] [Google Scholar]
  20. Upadhyaya P.; Rao P.; Hochalter J. B.; Li Z.-z.; Villalta P. W.; Hecht S. S. Quantitation of N-Acetyl-S-(9,10-dihydro-9-hydroxy-10-phenanthryl)-l-cysteine in Human Urine: Comparison with Glutathione-S-transferase Genotypes in Smokers. Chem. Res. Toxicol. 2006, 19 (9), 1234–1240. 10.1021/tx060096w. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Yang Y.; Griffiths W. J.; Sjövall J.; Gustafsson J. A.; Rafter J. Liquid chromatography-mass spectrometry with collision-induced dissociation of conjugated metabolites of benzol[ a]pyrene. J. Am. Soc. Mass Spectrom. 1997, 8 (1), 50–61. 10.1016/S1044-0305(96)00136-5. [DOI] [Google Scholar]
  22. Santana M. S.; Sandrini-Neto L.; Filipak Neto F.; Oliveira Ribeiro C. A.; Di Domenico M.; Prodocimo M. M. Biomarker responses in fish exposed to polycyclic aromatic hydrocarbons (PAHs): Systematic review and meta-analysis. Environ. Pollut. 2018, 242 (Pt A), 449–461. 10.1016/j.envpol.2018.07.004. [DOI] [PubMed] [Google Scholar]
  23. Varanasi U.; Nishimoto M.; Reichert W. L.; Le Eberhart B. T. Comparative metabolism of benzo(a)pyrene and covalent binding to hepatic DNA in English sole, starry flounder, and rat. Cancer Res. 1986, 46 (8), 3817–3824. [PubMed] [Google Scholar]
  24. Nishimoto M.; Yanagida G. K.; Stein J. E.; Baird W. M.; Varanasi U. The metabolism of benzo(a)pyrene by English sole (Parophrys vetulus): comparison between isolated hepatocytes in vitro and liver in vivo. Xenobiotica 1992, 22 (8), 949–961. 10.3109/00498259209049901. [DOI] [PubMed] [Google Scholar]
  25. Stein J. E.; Hom T.; Casillas E.; Friedman A.; Varanasi U. Simultaneous exposure of English sole (Parophrys vetulus) to sediment-associated xenobiotics: Part 2—chronic exposure to an urban estuarine sediment with added 3H-benzo[a]pyrene and 14C-polychlorinated biphenyls. Mar. Environ. Res. 1987, 22 (2), 123–149. 10.1016/0141-1136(87)90032-8. [DOI] [Google Scholar]
  26. Meier S.; Karlsen Ø.; Le Goff J.; Sørensen L.; Sørhus E.; Pampanin D. M.; Donald C. E.; Fjelldal P. G.; Dunaevskaya E.; Romano M.; Caliani I.; Casini S.; Bogevik A. S.; Olsvik P. A.; Myers M.; Grøsvik B. E. DNA damage and health effects in juvenile haddock (Melanogrammus aeglefinus) exposed to PAHs associated with oil-polluted sediment or produced water. PLoS One 2020, 15 (10), e0240307 10.1371/journal.pone.0240307. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. da Silva D. A. M.; Gates J. B.; O’Neill S. M.; West J. E.; Ylitalo G. M. Assessing hydroxylated polycyclic aromatic hydrocarbon (OHPAH) metabolites in bile of English sole (Parophrys vetulus) from Puget Sound, WA, USA by liquid chromatography/tandem mass spectrometry (LC-MS/MS). Sci. Total Environ. 2023, 865, 161229 10.1016/j.scitotenv.2022.161229. [DOI] [PubMed] [Google Scholar]
  28. Bijlsma L.; Berntssen M. H. G.; Merel S. A Refined Nontarget Workflow for the Investigation of Metabolites through the Prioritization by in Silico Prediction Tools. Anal. Chem. 2019, 91 (9), 6321–6328. 10.1021/acs.analchem.9b01218. [DOI] [PubMed] [Google Scholar]
  29. Celma A.; Bade R.; Sancho J. V.; Hernandez F. l.; Humphries M.; Bijlsma L. Prediction of Retention Time and Collision Cross Section (CCSH+, CCSH–, and CCSNa+) of Emerging Contaminants Using Multiple Adaptive Regression Splines. J. Chem. Inf. Model 2022, 62 (22), 5425–5434. 10.1021/acs.jcim.2c00847. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Villalta P. W.; Hochalter J. B.; Hecht S. S. Ultrasensitive High-Resolution Mass Spectrometric Analysis of a DNA Adduct of the Carcinogen Benzo[a]pyrene in Human Lung. Anal. Chem. 2017, 89 (23), 12735–12742. 10.1021/acs.analchem.7b02856. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Guo L.; Jiang X.; Tian H.-Y.; Yao S.-J.; Li B.-Y.; Zhang R.-J.; Zhang S.-S.; Sun X. Detection of BPDE-DNA adducts in human umbilical cord blood by LC-MS/MS analysis. J. Food Drug Anal. 2019, 27 (2), 518–525. 10.1016/j.jfda.2019.03.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Dunn B. Carcinogen Adducts As an Indicator for the Public Health Risks of Consuming Carcinogen-Exposed Fish and Shellfish. Environ. Health Perspect. 1991, 90, 111–6. 10.2307/3430852. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Kumar G. N.; Surapaneni S. Role of drug metabolism in drug discovery and development. Med. Res. Rev. 2001, 21 (5), 397–411. 10.1002/med.1016. [DOI] [PubMed] [Google Scholar]
  34. Jernström B.; Babson J. R.; Moldéus P.; Holmgren A.; Reed D. J. Glutathione conjugation and DNA-binding of (±)-trans-7,8-dihydroxy-7,8-dihydrobenzo[a]pyrene and (±)-7β,8α-dihydroxy-9α,10α-epoxy-7,8,9,10-tetrahydrobenzo[a]pyrene in isolated rat hepatocytes. Carcinogenesis 1982, 3 (8), 861–866. 10.1093/carcin/3.8.861. [DOI] [PubMed] [Google Scholar]
  35. Cheng G.; Zarth A. T.; Upadhyaya P.; Villalta P. W.; Balbo S.; Hecht S. S. Investigation of the presence in human urine of mercapturic acids derived from phenanthrene, a representative polycyclic aromatic hydrocarbon. Chem. Biol. Interact. 2017, 274, 80–88. 10.1016/j.cbi.2017.07.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Levsen K.; Schiebel H.-M.; Behnke B.; Dötzer R.; Dreher W.; Elend M.; Thiele H. Structure elucidation of phase II metabolites by tandem mass spectrometry: an overview. J. Chromatogr. A 2005, 1067 (1), 55–72. 10.1016/j.chroma.2004.08.165. [DOI] [PubMed] [Google Scholar]
  37. Meador J.; Stein J.; Reichert W.; Varanasi U. Bioaccumulation of Polycyclic Aromatic Hydrocarbons by Marine Organisms. Rev. Environ. Contam. T. 1995, 143, 79–165. 10.1007/978-1-4612-2542-3_4. [DOI] [PubMed] [Google Scholar]
  38. Celma A.; Sancho J. V.; Schymanski E. L.; Fabregat-Safont D.; Ibáñez M. a.; Goshawk J.; Barknowitz G.; Hernández F. l.; Bijlsma L. Improving Target and Suspect Screening High-Resolution Mass Spectrometry Workflows in Environmental Analysis by Ion Mobility Separation. Environ. Sci. Technol. 2020, 54 (23), 15120–15131. 10.1021/acs.est.0c05713. [DOI] [PubMed] [Google Scholar]
  39. Baillie T. A.; Davis M. R. Mass spectrometry in the analysis of glutathione conjugates. Biol. Mass Spectrom. 1993, 22 (6), 319–325. 10.1002/bms.1200220602. [DOI] [PubMed] [Google Scholar]
  40. Scholz K.; Dekant W.; Völkel W.; Pähler A. Rapid Detection and Identification of N-Acetyl- L-Cysteine Thioethers Using Constant Neutral Loss and Theoretical Multiple Reaction Monitoring Combined with Enhanced Product-Ion Scans on a Linear Ion Trap Mass Spectrometer. J. Am. Soc. Mass Spectrom. 2005, 16 (12), 1976–1984. 10.1016/j.jasms.2005.08.003. [DOI] [PubMed] [Google Scholar]
  41. Dieckhaus C. M.; Fernández-Metzler C. L.; King R.; Krolikowski P. H.; Baillie T. A. Negative Ion Tandem Mass Spectrometry for the Detection of Glutathione Conjugates. Chem. Res. Toxicol. 2005, 18 (4), 630–638. 10.1021/tx049741u. [DOI] [PubMed] [Google Scholar]
  42. Mahajan M. K.; Evans C. A. Dual negative precursor ion scan approach for rapid detection of glutathione conjugates using liquid chromatography/tandem mass spectrometry. Rapid Commun. Mass Spectrom. 2008, 22 (7), 1032–1040. 10.1002/rcm.3458. [DOI] [PubMed] [Google Scholar]
  43. Xie C.; Zhong D.; Chen X. A fragmentation-based method for the differentiation of glutathione conjugates by high-resolution mass spectrometry with electrospray ionization. Anal. Chim. Acta 2013, 788, 89–98. 10.1016/j.aca.2013.06.022. [DOI] [PubMed] [Google Scholar]
  44. Brink A.; Fontaine F.; Marschmann M.; Steinhuber B.; Cece E. N.; Zamora I.; Pähler A. Post-acquisition analysis of untargeted accurate mass quadrupole time-of-flight MSE data for multiple collision-induced neutral losses and fragment ions of glutathione conjugates. Rapid Commun. Mass Spectrom. 2014, 28 (24), 2695–2703. 10.1002/rcm.7062. [DOI] [PubMed] [Google Scholar]
  45. Ma Y.; Kind T.; Yang D.; Leon C.; Fiehn O. MS2Analyzer: A Software for Small Molecule Substructure Annotations from Accurate Tandem Mass Spectra. Anal. Chem. 2014, 86 (21), 10724–10731. 10.1021/ac502818e. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Sørhus E.; Incardona J. P.; Furmanek T.; Goetz G. W.; Scholz N. L.; Meier S.; Edvardsen R. B.; Jentoft S. Novel adverse outcome pathways revealed by chemical genetics in a developing marine fish. eLife 2017, 6, e20707. 10.7554/eLife.20707. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Boitsov S.; Petrova V.; Jensen H. K. B.; Kursheva A.; Litvinenko I.; Klungsøyr J. Sources of polycyclic aromatic hydrocarbons in marine sediments from southern and northern areas of the Norwegian continental shelf. Mar. Environ. Res. 2013, 87–88, 73–84. 10.1016/j.marenvres.2013.03.006. [DOI] [PubMed] [Google Scholar]
  48. Nepstad R.; Hansen B. H.; Skancke J. North sea produced water PAH exposure and uptake in early life stages of Atlantic Cod. Mar. Environ. Res. 2021, 163, 105203 10.1016/j.marenvres.2020.105203. [DOI] [PubMed] [Google Scholar]
  49. Willett K. L.; Gardinali P. R.; Lienesch L. A.; Di Giulio R. T. Comparative metabolism and excretion of benzo(a)pyrene in 2 species of ictalurid catfish. Toxicol. Sci. 2000, 58 (1), 68–76. 10.1093/toxsci/58.1.68. [DOI] [PubMed] [Google Scholar]
  50. Yu Y.; Wade T. L.; Fang J.; McDonald S.; Brooks J. M. Gas chromatographic-mass spectrometric analysis of polyclic aromatic hydrocarbon metabolites in Antarctic fish (Notothenia gibberifrons) injected with diesel fuel Arctic. Arch. Environ. Contam. Toxicol. 1995, 29 (2), 241–246. 10.1007/BF00212975. [DOI] [Google Scholar]
  51. Xie Z.; Chen J. Y.; Gao H.; Keith R. J.; Bhatnagar A.; Lorkiewicz P.; Srivastava S. Global Profiling of Urinary Mercapturic Acids Using Integrated Library-Guided Analysis. Environ. Sci. Technol. 2023, 57 (29), 10563–10573. 10.1021/acs.est.2c09554. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

es4c05112_si_001.pdf (1.9MB, pdf)
es4c05112_si_002.zip (1.9MB, zip)
es4c05112_si_003.pdf (168.9KB, pdf)

Articles from Environmental Science & Technology are provided here courtesy of American Chemical Society

RESOURCES