Water column monitoring near oil installations in the North Sea 2001–2004

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Marine Pollution Bulletin 56 (2008) 414–429

Water column monitoring near oil installationsin the North Sea 2001–2004

Ketil Hylland a,b,*, Knut-Erik Tollefsen a, Anders Ruus a, Grete Jonsson c,1, Rolf C. Sundt c,Steinar Sanni c, Toril Inga Røe Utvik d, Stale Johnsen e, Ingunn Nilssen e, Laurence Pinturier f,Lennart Balk g, Janina Barsiene h, Ionan Marigomez i, Stephen W. Feist j, Jan Fredrik Børseth c

a Norwegian Institute for Water Research (NIVA), Gaustadalleen 21, N-0349 Oslo, Norwayb Department of Biology, University of Oslo, Oslo, Norway

c IRIS and Akvamiljø, Mekjarvik, Norwayd Norsk Hydro, Bergen, Norway

e Statoil, Trondheim, Norwayf Total, EP, Stavanger, Norway

g ITM, Stockholm University, Stockholm, Swedenh Institute of Ecology, Vilnius University, Vilnius, Lithuania

i University of the Basque Country, Bilbo, Basque Country, Spainj Centre for Environment Fisheries and Aquaculture Science, Weymouth, UK

Abstract

Fisheries have been vital to coastal communities around the North Sea for centuries, but this semi-enclosed sea also receives largeamounts of waste. It is therefore important to monitor and control inputs of contaminants into the North Sea. Inputs of effluents fromoffshore oil and gas production platforms (produced water) in the Norwegian sector have been monitored through an integrated chemicaland biological effects programme since 2001. The programme has used caged Atlantic cod and blue mussels. PAH tissue residues in bluemussels and PAH bile metabolites in cod have confirmed exposure to effluents, but there was variation between years. Results for a rangeof biological effects methods reflected exposure gradients and indicated that exposure levels were low and caused minor environmentalimpact at the deployment locations. There is a need to develop methods that are sufficiently sensitive to components in produced water atlevels found in marine ecosystems.� 2007 Elsevier Ltd. All rights reserved.

Keywords: Produced water; Caging; Atlantic cod; Blue mussels; Biomarkers; Monitoring

1. Introduction

The North Sea is a semi-enclosed sea with ecosystemsthat are subject to intensive fishing pressure as well asreceiving inputs of environmental contaminants. The mainsources for contaminants have historically been land-based

doi:10.1016/j.marpolbul.2007.11.004

* Corresponding author. Address: Department of Biology, University ofOslo, Oslo, Norway. Tel.: +47 22857315; fax: +47 22854726.

E-mail address: ketilhy@bio.uio.no (K. Hylland).1 Present address: Stavanger University Hospital, Stavanger, Norway.

or riverine, but there has been an increasing contributionfrom offshore oil and gas production in the past decades.The main effluent from offshore oil and gas platforms dur-ing the production phase is generally termed producedwater. Produced water may contain varying concentrationsof production chemicals (e.g. complexing agents) or bio-cides in addition to natural components such as PAHs,alkylphenols and metals. Although discharged in large vol-umes, produced water is rapidly diluted in surrounding sea-water and marine organisms will generally be exposed tolow concentrations of produced water components. Toavoid conflict between fisheries and offshore activities it is

K. Hylland et al. / Marine Pollution Bulletin 56 (2008) 414–429 415

important that contaminant inputs and any biologicaleffects they may cause are monitored and controlled.

There are reasons to be concerned for effects of contam-inants in North Sea ecosystems. In the 1980s there wereclear indications that contaminants were related toincreased incidence of aberrations in fish embryos in thesouthern North Sea (von Westernhagen et al., 1987,1989). Secondly, extensive fish disease monitoring activitiesthat started in the late 1970s (Lang, 2002) has demon-strated decreases in some diseases and increases in others(Dethlefsen et al., 1987; Lang and Wosniok, 2003). It ishowever challenging to establish links from such observa-tions to contaminant exposure because of the multifacto-rial etiology of diseases and embryonal aberrations.There is a current concern as to possible impacts of off-shore oil and gas activities on North Sea fish populations.Recent data on sublethal health-related measurements(biomarkers) suggest that fish in areas with high producedwater inputs have increased incidence of DNA damagecompared to reference areas (Hylland et al., 2006b) andearlier studies have shown that North Sea fish could beinfluenced by estrogenic substances (Lye et al., 1997; Allenet al., 1999; Bateman et al., 2004; Stentiford and Feist,2005). Studies with model species in the laboratory in thelast few years have shown that that sex determinationand differentiation of some species can be extraordinarilysensitive to estrogen or androgen exposure (Orn et al.,2003). Based on those findings there is a need for moreinformation to evaluate the health of fish in the North Sea.

Following the Biological Effects of Contaminants inMarine Pelagic Ecosystems (BECPELAG) workshop in2001, the environmental impact of produced water effluentsfrom selected offshore production platforms (Troll B in2003 and Statfjord B in 2004) were monitored using a com-bination of chemical and biological effects methods. Themethods employed in the water column monitoring pro-grammes were chosen on the basis of recommendationsfrom the above workshop (Hylland et al., 2002) and focusedon the use of caged organisms, blue mussel (Mytilus edulis

L.) and Atlantic cod (Gadus morhua L.). The methods orig-inally selected for blue mussels included PAH concentra-tion, benzo[a]pyrene hydroxylase activity, lysosomalstability and selected histopathological endpointsand forAtlantic cod, bile PAH metabolite concentration, cyto-chrome P4501A activity (EROD), glutathione S-transferaseactivity, DNA adduct concentration, vitellogenin concen-tration and selected histopathological endpoints. An impor-tant component of the offshore water column monitoringprogramme was an option to try out new methods forfuture inclusion in the programme. One of the methods thathas been tested, micronucleus frequency in blue mussel hae-mocytes, is now included in the programme.

The methods listed above were partly chosen due toexpectations that the main contributors to toxicity in pro-duced water effluents would be natural components of oil,i.e. PAHs and alkylphenols, which is why PAH concentra-tion in blue mussels and PAH metabolites in cod bile were

used to provide information on exposure to the effluent.The effects methods were chosen on the basis of previousguidelines (e.g. JAMP, 1998) and experience from the2001 campaign (in which a large number of different meth-ods were used). It is well known that hepatic cytochromeP4501A (CYP1A) in fish is responsive to PAH exposure(see e.g. Hylland, 2006) and it was therefore included as acore method in the monitoring programme. Although notas widely applied as the phase-I enzyme CYP1A, thephase-II enzyme glutathione S-transferase (GST) wasincluded in the surveys as previous results have indicatedthat it may respond to PAHs and produced water compo-nents (Foureman, 1989; Kennedy et al., 1991; Danischew-ski, 2006). Exposure to some PAHs may lead to thegeneration of DNA adducts and subsequently cancer (Eric-son and Balk, 2000). The principle organ involved in detox-ification is the liver and changes in tissue integrity,including neoplasia, may then be elucidated using histopa-thological methods (Feist et al., 2004). All the above meth-ods have been linked specifically to PAH exposure and themethods included for cod were also the methods suggestedin the OSPAR JAMP guideline to monitor biologicaleffects of PAHs (JAMP, 1998).

As mentioned above, the presence of alkylphenols inproduced water has been a concern, mainly due to theirinteraction with estrogen-related processes. The egg yolkprecursor protein vitellogenin (vtg) is normally producedin female fish as a result of hepatic exposure to endogenousestrogens, but production in male fish can be artificiallyinduced by exposure to exogenous estrogens, includingenvironmental estrogens (Sumpter and Jobling, 1995).The use of vtg as a biomarker for environmental estrogensin ecologically relevant fish species has since then beenemployed in coastal and freshwater environmental moni-toring (Hylland et al., 1998, 1999) and for monitoring ofeffluents in areas receiving by discharges from oil produc-tion activities (Knudsen et al., 1997; Scott et al., 2006b).The results from studies with freshwater species suggestthat induction of vtg occurs at concentrations of environ-mental estrogens that also produce alteration in sexualdevelopment (Jobling et al., 1996; Orn et al., 2003).

In addition to the above, methods that reflect the gen-eral health of the caged organisms have been used, i.e. lyso-somal stability of blue mussel hepatopancreas cells (2001and 2003) or haemocytes (2003 and 2004) and micronu-cleus (MN) formation in cod kidney cells (2003 and2004) and blue mussel haemocytes (2004). Lysosomal sta-bility is a general health parameter and has earlier beenshown to respond to PAH and crude oil exposed mussels(Camus et al., 2000; Fernley et al., 2000). Extensive chro-mosomal rearrangements such as micronucleus formationare widely recognised consequences of genome instabilitythat may arise from contaminant stress (Fenech et al.,1999) and the MN assay may indicate DNA breakage orspindle dysfunction caused by clastogens and aneuploido-genic toxins (Heddle et al., 1983, 1991; Zoll-Moreux andFerrier, 1999).

416 K. Hylland et al. / Marine Pollution Bulletin 56 (2008) 414–429

The aim of this paper is to review results from threewater column monitoring campaigns in the Norwegian sec-tor of the North Sea with a special focus on the ability ofbiological effects methods to identify and quantify environ-mental effects of produced water effluents. This overviewdoes not include all methods used for the three years, butwill focus on methods viewed as particularly relevant formonitoring purposes.

2. Materials and methods

2.1. Areas

Water column monitoring in 2001 and 2004 took placein the Tampen region, close to the Statfjord oil field,whereas the water column monitoring campaign in 2003was done in the vicinity of the Troll B platform (Table 1,Fig. 1). Locations for caging deployment upstream ordownstream of platforms were chosen on the basis of dis-persion modelling using the DREAM model (see Durrellet al., 2006).

2.2. Origin of organisms

Blue mussels (M. edulis) were taken from clean locationsin southern Ireland in 2001 (Hylland et al., 2006c,d) andfrom Trøndelag (central Norway) in 2003 and 2004.Farmed Atlantic cod (G. morhua) to be used for deploy-ment in all 3 years were obtained from aquaculture facili-ties close to Bergen (Norway).

2.3. Cage deployment and retrieval

A description of the cages and a general discussion ofcaging technology can be found in Hylland et al. (2004).Cages were deployed at four locations in 2001 (StatfjordB/C), on five locations in 2003 (Troll B) and on five loca-tions in 2004 (Statfjord B/C). In 2001, cages were posi-

Table 1Positions for caging locations in 2001, 2002 and 2003 (only locationsdiscussed in the text are included)

Year Station code Organisms Position

2001 500 m Cod, blue mussel 61�1201800N 1�5003600E2000 m Cod, blue mussel 61�1104800N 1�5200600E10,000 m Cod, blue mussel 61�1000000N 1�5903000EReference Cod, blue mussel 60�0700000N 3�0300000E

2003 500 m Cod, blue mussel 60�4604400N 3�3003500E1000 m Cod, blue mussel 60�4605600N 3�3005900E5000 m Blue mussel 60�4701800N 3�3104600EReference-1 Cod, blue mussel 60�4604400N 3�3003500EReference-2 Cod, blue mussel 60�4605600N 3�3005900E

2004 500 m Cod, blue mussel 61�1201800N 1�5001800E1000 m Cod, blue mussel 61�1200600N 1�5004200E10,000 m Cod, blue mussel 61�1000000N 1�5903000EReference Cod, blue mussel 61�0700000N 1�5203000E

tioned in the direction assumed to be that of the mainresidual surface current from the platform at distances of500 m, 2000 m, 10,000 m and two cages at a reference loca-tion. In 2003, cages were positioned at 500 m, 1000 m (bothfish and mussels) in the direction assumed to be that of themain residual surface current, at 500 m and 1000 m in theopposite direction, as well as at 5000 m in the assumedmain direction for the plume (only mussel); two referencelocations were used in 2003, both 8000 m away from theplatform. Only data from locations with both fish and mus-sels will be included in this paper. In 2004, cages were posi-tioned in the same general direction as in 2001, but atdistances of 500 m, 1000 m, 2500 m and 10,000 m in addi-tion to two reference cages. None of the cages were lostin 2001, the cage at 2000 m (mussels only) in 2003, the2500 m cage and a reference cage in 2004 (both fish andmussels) were lost. Cages were deployed in May/June in2001 and in the autumn (August/September) in 2003 and2004. Cages were retrieved 5–6 weeks after deployment.

2.4. Sampling

Atlantic cod were sampled prior to deployment andimmediately following retrieval from cages. Blue musselswere sampled on board research vessels in 2001 and2003, but were brought back to shore for sampling andanalyses of lysosomal stability in 2004. Samples for bio-marker analyses were analysed immediately (lysosomal sta-bility in 2003), fixed (samples for lysosomal stability in2001, histopathology in each year and slides for micronu-cleus determination) or snap-frozen in liquid nitrogenand analysed later (all other methods). Hepatopancreasof blue mussel for histochemical and histological studieswere fixed in Davidson solution.

2.5. Chemical analyses

2.5.1. PAH concentrations in blue mussels

The biological matter was homogenised, internal stan-dards added and saponified. The compounds wereextracted with n-pentane and dried over sodium sulphate.The extraction volume was reduced and the extracts werecleaned by GPC and solvent exchanged to cyclohexane.The extracts were then analysed by GC/MS with the MSdetector operating in selected ion monitoring mode (SIM)and analyte concentrations in the standard solutions werein the range 5–1000 ng/ll. The GC was equipped with a30 m column with a stationary phase of 5% phenyl polysi-loxane (0.25 mm i.d. and 0.25 lm film-thickness), andinjections operated in splitless mode. The initial columntemperature was 40 �C, which after 2 min was raised to300 �C at a rate of 7 �C/min and thereafter raised to300 �C at a rate of 10 �C/min. The injector temperaturewas 300 �C, the transfer-line temperature was 280 �C, theMS source temperature was 230 �C and the column flowrate was 1.2 ml/min. Quantification of individual compo-nents was performed by using the internal standard

Fig. 1. Overview of locations referred to in the text. Production platforms are indicated by squares.

method. The alkylated homologues were quantified bybaseline integration of the established chromatographic

pattern and the response factors were set to equal withineach group of homologues.

2.5.2. PAH bile metabolites

2.5.2.1. FF-fixed wavelength fluorescence of bile. Bile sam-ples of cod were screened for PAH metabolites by directfluorescence analysis using fixed wavelength fluorescence(FF) as described by Aas et al. (2000). Bile samples werediluted 1:1600 in methanol:water (1:1) prior to analysis. Slitwidths were set at 2.5 nm for both excitation and emission,and samples were analysed by FF at the wavelength pairs290/335, 341/383 and 380/430 nm for the detection ofnaphthalene, pyrene and benzo[a]pyrene types of metabo-lites, respectively. The fluorescence signal was transformedinto pyrene fluorescence equivalents by use of a pyrene(Sigma, St. Louis, USA) standard calibration curve. Theconcentrations of PAH metabolites in bile samples couldthen be expressed as nanogram pyrene fluorescence equiv-alents per gram bile (Aas et al., 2000).

2.5.2.2. GC–MS gas chromatography–mass spectrometry of

bile. Fish bile was prepared for analysis as described byJonsson et al. (2003, 2004). Briefly, 25–30 ll of bile wasweighed accurately into a micro centrifuge vial. Internalstandards (2,6-dibromophenol, 3-fluorophenanthrene and1-fluoropyrene) and b-glucuronidase (3000 units) insodium acetate buffer (0.4 M, pH 5) were added and thesolution left at 40 �C for 2 h. The OH–PAHs wereextracted with ethylacetate (3 times 0.5 ml), the combinedextract dried with anhydrous sodium sulphate and concen-trated to 0.5 ml. Trimethylsilyl (TMS) ethers of OH–PAHswere prepared by addition of 0.2 ml BSTFA and heatingfor 2 h at 60 �C. Triphenylamine (TPA) was added as aGC–MS performance standard before transferring the pre-pared samples to capped vials.

Trimethylsilyl ethers of OH–PAHs (TMS–OH–PAHs)in fish bile samples were analysed by the GC–MS systemdescribed by Jonsson et al. (2003). Helium was used as car-rier gas and the column used was CP-Sil 8 CB-MS,50 m � 0.25 mm and 0.25 lm film-thickness (InstrumentTeknikk A.S., Oslo, Norway). Samples and calibrationstandards (1 ll) were injected on a split/splitless injectorwith splitless mode on for 1 min. The temperatures forthe injector, transfer-line and ion source were held at250 �C, 300 �C and 240 �C, respectively, and the GC oventemperature programme was as follows: 80–120 �C at15 �C min�1, 120–300 �C at 6 �C min�1 and held at300 �C for 30 min. Mass spectra were obtained at 70 eVin selected ion mode (SIM). Based on the fragmentationpattern of non-alkylated TMS–O–PAHs (Jonsson et al.,2003) and studies performed by Krahn et al. (1992) andYu et al. (1995) the molecular ions were selected for deter-mination of both alkylated and non-alkylated TMS–O–PAHs.

2.6. Biomarker analyses

2.6.1. Cytochrome P4501A activity (EROD)Cytochrome P4501A activity was measured as EROD

(ethoxyresorufin O-deethylase) activity during all three

years. In 2001 the analyses were done at the Universityof Gothenburg (Forlin and Hylland, 2006), in 2003 atNIVA and in 2004 at IRIS-Akvamiljø. The method usedin 2001 can be found in Forlin et al. (1994), methods usedin 2003 in Hylland et al. (2006c,d) and in 2004 as describedin Aas et al. (2000).

2.6.2. Glutathione S-transferase

Total glutathione S-transferase (GST) activity was mea-sured using 1-chloro-2,4-dinitrobenzene (CDNB,C6H3ClN2O4) as substrate (Habig et al., 1974; Habig andJakoby, 1981) during all three campaigns. When GSTactivities were quantified, the blank value was deductedand a molar extinction coefficient (2) for CDNB of9.6 mM�1 cm�1 was used. Microplate formats were usedduring all studies, but the analyses were done at Bundesamtfur Fischerei in 2001 and by IRIS at Akvamiljø in 2003 and2004. GST activities were expressed as moles of substrateconverted per minute per mg of protein in the cytosol.The method described by Bradford (1976) was used forprotein determinations.

2.6.3. DNA adducts

DNA adducts were determined at ITM, Stockholm Uni-versity, for all three campaigns. The method used was asdescribed in Ericson et al. (1998) and Ericson and Balk(2000). Briefly, liver samples were semi-thawed, gentlyhomogenised in 10 mM Tris–Cl, 100 mM etylenediamine-tetraacetic acid (EDTA) and then centrifuged to obtain apellet of crude nuclei. The pellet was resuspended and trea-ted with SDS, RNase T1, RNase A and a-amylase, fol-lowed by incubation with proteinase K. DNA adductswere enriched by the Nuclease P1 method, using 0.8 lgNuclease P1/lg DNA, and a 45 min incubation period(Reddy and Randerath, 1986; Beach and Gupta, 1992).As the final enzymatic treatment, DNA adducts wereradiolabelled using 50-[c-32P]triphosphate([c-32P]ATP) andT4 polynucleotide kinase. DNA for adduct analysis wasquantified on the basis of its absorption at 260 nm. TheDNA adducts were located and the levels quantified onthe TLC sheets with ImageQuant, 5.0 software, MolecularDynamics software, by the storage phosphor imaging tech-nique using a PhosphorImagerTM SI instrument (Sunnyvale,CA, USA), essentially according to the methodologydescribed by Reichert et al. (1992). DNA adducts were cal-culated relative to adduct labelling, which is the ratio ofadducts to the total number of analyzed nucleotides, ornmol adducts/mol normal nucleotides. The detection limitof DNA adducts was calculated for the individual samplesof cod liver tissue, from the background signal in a repre-sentative area of the autoradiogram. A spot/area/zone elec-tronic signal, corresponding to 1.5 times the backgroundlevel on the same autoradiogram, was considered to bethe limit of detection and the limit of quantification forDNA adducts.

2.6.4. Lysosomal stability

Lysosomal stability was quantified using histochemicalmethods on frozen digestive gland samples in 2001 and2003 and in fresh haemocytes in 2003 and 2004. Analysesin 2001 and 2003 were undertaken as described in Bilbaoet al. (2006). The determination of lysosomal membranestability was based on the time of acid labilisation treat-ment required to produce the maximum staining intensity.In the 2003 and 2004 field studies lysosomal response inhaemocytes was measured with the method described byLowe and Pipe (1994). Haemolymph samples were madeof 15 individuals at each station. 0.2 ml was sampled froma sinus in the posterior adductor muscle and mixed 1:1 witha physiological Ringer solution (pH 7.4). Forty microlitresdiluted haemolymph fluid was transferred to an objectglass and incubated 20 min in a light proof and highhumidity box prior to addition of 40 ll of the colour pig-ment neutral red (0.1 lg/ll). Time to 50% response wasdetermined.

In the 2004 field study mussels from the pre-exposuregroup and field groups were brought from to the labora-tory in Stavanger on ice prior to assessment of lysosomalstability. The mussels were further acclimatised in the labin aquaria with fresh supplies of seawater for 2 days priorto sampling (to prevent the influence of handling and trans-port stress). Haemolymph samples were obtained of 19individuals at each field station and 26 individuals fromthe pre-exposure group. 0.4 ml haemolymph was sampledfrom each mussel and mixed with filtered sea water at theration 2:1. Forty microlitres haemolymph/seawater-mix-ture was pipetted out on microscope-slides and incubatedin a light-proof box for 20 min before 35 ll neutral red(concentration 0.1 lg/ll) was added. All analyses were per-formed blind.

2.6.5. Frequency of micronuclei

Analyses for micronucleus formation were performed aspart of the monitoring programmes in 2003 and 2004.Mussel haemolymph and samples from cod liver werespread on slides, air dried and fixed for 15 min in methanol.Slides were then stained with 5% Giemsa solution for 10–20 min. Micronuclei were scored on coded slides withoutknowledge of the exposure status of the samples. The fre-quency of micronuclei was determined by scoring at a1000� magnification. Micronuclei were counted in 5000immature erythrocytes from each cod specimen and in2000 haemocytes from each mussel specimen and expressedas the number of micronuclei per 1000 cells scored (Bar-siene et al., 2004).

2.6.6. Vitellogenin

Vitellogenin in cod plasma were analysed by Cefas andBiosense Laboratories AS in parallel in 2001 (Scott et al.,2006b) and by NIVA in 2003 and 2004. Briefly, blood sam-ples were taken from the caudal artery of caged Atlanticcod and saithe by means of pre-cooled syringes containingheparin (10,000 IU/ml, Sigma) and the protease inhibitor

Aprotinin (5 TIU/ml, Sigma) and centrifuged at approxi-mately 2000g for 10 min at 4 �C. The supernatant was care-fully decanted, aliquots prepared and samples snap-frozenin liquid nitrogen and stored at �80 �C until analysis.Vitellogenin was determined by a competitive enzymelinked immuno sorbent assay (ELISA) as described inScott et al. (2006b), or as a sandwich ELISA accordingto the manufacturers specifications (Biosense LaboratoriesAS, Bergen, Norway).

2.6.7. Histochemistry and histology (blue mussel)

Fixed tissues were dehydrated in alcohols and embeddedin paraffin. Histological sections (7 lm) were cut with theaid of a rotary microtome, stained with haematoxylin–eosin (H&E) and mounted on glass slides. Histopatholo-gical examination was carried out under the light micro-scope. Prevalence of parasites, associated host response,haemocyte infiltration and pathological changes of thedigestive epithelium, the interstitial connective tissue andthe gonad tissue were recorded.

As an indication of whether cell-type replacementoccurred or not, the volume density of basophilic cells(VVBAS) in the digestive gland of mussels was determinedby means of stereology. A Weibel graticule (M-168; Wei-bel, 1979) was superimposed onto paraffin sections stainedwith H/E with the aid of a drawing tube attachment. Ran-domly selected five fields were counted in 10 individuals perexperimental group (20� objective). The volume density ofbasophilic cells was calculated as VVBAS = VBAS/VDT,where Vis the volume, BAS the basophilic cell and DT isthe digestive gland epithelium.

In order to quantify the structure of the digestivetubules, a planimetric procedure was applied on paraffinsections of digestive gland tissue (Vega et al., 1989). A totalof 25 tubule sections per individual were recorded in animage analysis system (Visilog 5.4 Noesis) attached to anOlympus BX50 light-microscope. The ratio between MET(mean epithelial thickness) and MLR (mean luminalradius) is then quantified.

2.7. Statistical analyses

Biological responses in individual mussel or fish weresubjected to analysis of variance (ANOVA) to clarifywhether there were differences between groups within eachyear of study (Sokal and Rohlf, 1981). Prior to analyses,homogeneity of variances was checked using Levene’s test.Variables were transformed as appropriate to attain homo-dascicity. Where this was not possible or there were fewobservations, the non-parametric Kruskal–Wallis analysiswas used (Sokal and Rohlf, 1981). Where the parametricANOVA indicated significant differences, groups werecompared using Tukey’s post-hoc test. The level of signifi-cance to reject H0: ‘‘no difference between groups” was setto 0.05.

3. Results

3.1. PAHs in blue mussels

Blue mussels deployed in the North Sea accumulatedPAHs in all three years: in 2003 and 2004 compared toboth reference and pre-deployment mussels, in 2001 com-pared to mussels at the reference location (Fig. 2). Theresults confirmed that cages were positioned in areas withan exposure to oil-derived contamination. Mussel PAH-concentrations were furthermore significantly higher atthe location closest to the oil platform (500 m) than at allother locations in 2001 and 2003, but the differences wereless obvious in 2004. Differences between locations weresimilar for 3-ring PAHs (Fig. 2A) and the sum of EPA-16 PAHs (Fig. 2B) for the three monitoring compaigns.Additional results for 2001 were reported in Ruus et al.(2006).

Fig. 2. Concentrations of PAH in caged blue mussels from the indicatedmonitoring campaigns. (A) Summed concentration of 3-ring components(ng/g wet weight). (B) EPA 16 PAHs summed (ng/g wet weight); median,quartiles and 10/90 percentiles are shown. *: significantly different frompre-exposure group.

3.2. PAH metabolites in cod bile

There were significant differences in bile PAH metabo-lite concentrations of cod caged near platforms comparedto cod at the reference location in 2001, but no signifi-cant differences the other 2 years (Table 2). Similar differ-ences were observed using both fixed fluorescence(expressed as pyrene equivalents) and for the concentra-tion of alkylated and non-alkylated naphthalene metabo-lites (RC0–C3 OH–NPHs) (Table 2). A similar gradientwas observed for phenanthrene metabolites (results notshown). No significant differences in PAH metaboliteconcentration was determined by analyses at wavelengthpair 380/430 nm (results not shown). In 2004 there wasa significantly higher concentration of bile PAH metabo-lites in the pre-exposure fish compared to cod held at thereference location, as measured using fixed fluorescence(Table 2).

3.3. Cytochrome P4501A in cod

There were no significant differences in hepatic ERODactivity for cod held at different locations during the threecampaigns (Table 3). The fish were juvenile and are pre-sented as a single group for each location although individ-uals were sexed. In 2004 there was a significant increase forall deployed cod compared to pre-exposure fish, due tovery low hepatic EROD activity in the farmed cod usedfor deployment (Table 2).

Table 2Biliverdin, pyrene-like metabolites measured by fixed fluorescence and thetotal concentration of alkylated and non-alkylated naphthalene metabo-lites (RC0–C3 OH–NPHs) measured by gas chromatography–mass spec-trometry (GC–MS) in cod bile from three different studies

Year Group PFE 341/383 (lg/ml) RC0–C3 OH–NPHs (ng/g)

Mean SD N Mean SD N

2001 Pre-exposure 3.13 0.77 15 489 117 8500 m 4.89a 1.77 15 3223a 1519 82000 m 4.51a 1.10 15 2157a 597 610,000 m 3.59a 0.56 15 1423a 675 6Reference 2.26 0.19 15 1036 191 5

2003 Pre-exposure 2.44 0.58 25 1515 317 25500 m 3.16 0.58 25 1847 265 251000 m 2.79 0.67 25 1677 267 25Reference-1 2.50 1.65 25 1761 1701 25Reference-2 2.07 0.37 25 1558 419 25

2004 Pre-exposure 5.14 1.02 25 1836 599 5500 m 3.12 0.50 25 994 303 51000 m 2.80 0.61 25 905 315 52500 m 2.17 0.29 25 1594a 481 510,000 m 2.73 0.53 24 839 153 5Reference 2.28 0.45 25 965 255 5

The number of analysed fish within each group was: biliverdin: n = 15–25,FF: n = 10–25 and GC–MS: n = 5–25.

a Significantly different from reference (p < 0.05).

Table 3Hepatic cytochrome P4501A activity (pmol/min/mg protein) in juvenileAtlantic cod sampled prior to exposure and following caging in 2001, 2003and 2004; median and quartiles are shown; groups significantly differentfrom pre-exposure group for that year in italics

Year Group N Median (quartiles)

2001 500 m 20 17.8 (12.3, 27.8)2000 m 22 25.5 (18.1, 30.3)10,000 m 12 24.6 (20.0, 31.2)Reference 21 18.5 (14.5, 22.9)

2003 Pre-exposure 18 3.9 (2.7, 7.5)500 m 25 14.3 (6.7, 20.7)

1000 m 25 15.0 (7.6, 19.4)

Reference-1 25 13.1 (7.5, 16.9)

2004 Pre-exposure 30 12.1 (8.5, 18.1)500 m 15 12.4 (7.2, 17.8)1000 m 16 15.6 (10.6, 22.0)10,000 m 12 13.4 (9.2, 18.1)Reference 15 11.7 (4.8, 19.4)

Data for 2001 from Forlin and Hylland (2006).

3.4. Glutathione S-transferase in cod

There was significantly higher hepatic glutathioneS-transferase (GST) activity in cod caged at 500 m and2000 m compared to cod held at the reference location in2001 (Danischewski, 2006). For cod deployed in 2003and 2004, there were no clear gradient-related differences,although GST was significantly different at some locationscompared to the pre-exposure groups in both years (Table4). There were no sex-related differences for any year (juve-nile cod; results not shown).

3.5. DNA adducts

Following cage exposure in 2001 the percentage of fishwith detectable DNA adducts increased with increasingdistance from the oil platform, from 0% to 71%, and thelevels of adducts were significantly higher at the referencelocation than at the locations in the exposure gradient inthe vicinity of the platform (Balk et al., 2006). No sex-

Table 4Hepatic glutathione S-transferase activity (nmol/min/mg protein) in maleand female cod sampled prior to exposure and following caging in 2003and 2004; groups significantly different from pre-exposure group for thatyear in italics

2003 Pre-exposure 25 687(545, 782)500 m 25 754 (664, 876)1000 m 25 799 (752, 973)

Reference-1 25 793 (750, 909)

Reference-2 25 744 (628, 830)

2004 Pre-exposure 30 942 (819, 1092)500 m 27 829 (645, 1035)1000 m 27 710 (525, 794)

10,000 m 27 885 (640, 1020)Reference 27 751 (626, 918)

dependent differences were observed. There were no signif-icant differences in hepatic DNA adduct concentration inpre-exposure fish or cod exposed in cages during the 2003campaign, although some individuals in all groups had ele-vated concentrations of adducts (results not shown). DNAadducts were not analysed in caged cod during the 2004campaign.

3.6. Lysosomal stability

In 2001 the lysosomal membrane labilisation period inmussel digestive gland cells was significantly lower for mus-sels caged 500 m and 2000 m from the platform comparedto mussels caged at the reference location (Bilbao et al.,2006). Some effects appeared to be present in 2003 as well,although the labilisation period was low for the pre-expo-sure group (Table 7).

Neutral red retention in blue mussel haemocytes was notreported in 2001, but included in 2003 and 2004. In 2003,mussels at one of the two reference locations (reference-1)had significantly decreased retention time compared tothe other two groups tested (Table 5). For all groups in2004 lysosomal responses were within the normal rangeof retention times usually observed for blue mussels in‘‘clean” locations (Table 5).

3.7. Frequency of micronuclei

Micronucleus formation in cod liver immature erythro-cytes was included in the programmes in 2003 and 2004 aswell as for blue mussel haemocytes in 2004. Results forboth cod and mussels indicated trends towards higher fre-quency of micronuclei in organisms caged closer to theplatforms (Figs. 3 and 4). There was a clear gradient-related increase in micronuclei in haemocytes of cagedmussels in 2004, although only at 500 m was there signifi-cantly higher frequency of micronuclei compared to therefence location. Similar results were observed for coderythrocytes in 2003 and 2004 (Fig. 4).

Table 5Lysosomal stability of haemocytes from blue mussels (retention time,minutes) following caging in 2003 and 2004; medians and quartiles areshown

2003 Pre-exposure 10 180 (120, 180)2000 ma 15 180 (180, 180)Reference-1 15 150 (90, 180)

2004 Pre-exposure 26 120 (90, 150)500 m 19 120 (90, 120)1000 m 19 120 (90, 150)10,000 m 19 120 (75, 150)Reference 19 120 (90, 150)

a In the opposite direction of the plume.

pre-exposure 500 m 1000 m 10000 m reference

Fig. 3. Micronucleus formation in haemocytes from blue mussels follow-ing caging in 2004; N: 10 for all groups except 20 for pre-exposure;Kruskal–Wallis p = 0.0012.

pre-exposure 500 m 1000 m reference

pre-exposure 500 m 1000 m 10000 m reference

Fig. 4. Micronucleus formation in immature erythrocytes from cod cagedin (A) 2003 (ANOVA p < 0.0001). (B) 2004 (no significant differences);medians (square symbols), quartiles and 10/90 percentiles are shown. *:significantly different from pre-exposure and reference groups.

Table 6APlasma vitellogenin levels (lg/ml) in caged male (juvenile) cod prior todeployment (start) and deployed at different distances from the dischargesource during the BECPELAG campaign (Statfjord B/C) in 2001, thewater column survey (WCS) in 2003 (Troll B) and 2004 (Statfjord B/C);medians and quartiles are shown; groups significantly different fromreference group in italicizes

2001 500 m 13 0.081 (0.037, 0.52)

2000 m 14 0.019 (0.011, 0.045)10,000 m 7 0.028 (0.020, 0.043)Reference 11 0.013 (0.010, 0.053)

2003 Pre-exposure 11 8.9 (3.3, 31)

500 m 10 3.5 (2.2, 6.6)1000 m 9 4.4 (4.1, 6.0)Reference-1 11 3.1 (2.3, 3.6)Reference-2 11 2.9 (2.4, 3.3)

2004 Pre-exposure 16 4.1 (1.0, 14)500 m 16 3.2 (1.0, 10)1000 m 15 5.1 (1.9, 9.2)10,000 m 14 2.7 (0.44, 12)Reference 14 1.3 (0.67, 4.0)

Data for 2001 from Scott et al. (2006b).

Table 6BPlasma vitellogenin levels (lg/ml) in caged female (juvenile) cod prior todeployment (pre-exposure) and deployed at different distances from thedischarge source during the BECPELAG campaign (Statfjord B/C) in2001 and the water column surveys (WCS) in 2003 (Troll B) and 2004(Statfjord B/C); groups significantly different from reference group initalicizes

2001 500 m 8 0.044 (0.011, 0.22)2000 m 10 0.033 (0.016, 0.16)10,000 m 7 0.071 (0.029, 0.16)Reference 11 0.051 (0.023, 0.076)

2003 Pre-exposure 12 27 (13, 38)500 m 7 30 (8.8, 67)1000 m 7 34 (12, 110)Reference-1 13 13 (10, 27)Reference-2 12 27 (11, 43)

2004 Pre-exposure 8 23 (2.3, 46)500 m 7 20 (15, 31)

1000 m 12 20 (10, 24)10,000 m 13 98 (41, 140)Reference 13 70 (46, 140)

Data for 2001 from Scott et al. (2006).

3.8. Vitellogenin

Vitellogenin (vtg) was determined in blood plasma fromcaged male and female cod in 2001, 2003 and 2004. Ele-vated levels of vtg compared to the reference site were onlyfound in male fish deployed at the 500 m station in theStatfjord B transect during the 2001 campaign (Table6A). No significant differences were found for the other sta-tion during the 2001 campaign and between deploymentstations during water column surveys performed in 2003and 2004. No significant differences were detected forgroups of female fish from the 2001 and 2003 deployments,although plasma from the female fish at the 1000 m station

during 2004 contained significantly lower levels of vtg thanfemales at the reference site (Table 6B). Interestingly, levelsof vtg in males sampled during 2003 were significantlyhigher in pre-exposure controls compared to the transectstations. Furthermore, plasma levels of vtg in males andfemales deployed during the 2001 deployment were up totwo orders of magnitude lower than those detected in thewater column surveys in 2003 and 2004.

3.9. Digestive gland histopathology

A range of histopathological changes were detected inmussels and reported elsewhere (Feist et al., 2006). Baso-

Table 7Overview of results for three selected histochemical and histologicalendpoints in blue mussels for the indicated years; stations in bold:significantly different from reference; stations in italics: significantlydifferent from pre-exposure mussels; ns: no significant differences betweengroups; N/I: not included in monitoring programme that year

Method 2001 2003 2004

Lysosomal stability 500 m 500 m N/I2000 m 1000 m

Basophilic cell volume ns 500 m 500 m

1000 m

Reference-1

Reference-2

Structure digestive tubuli(MLR/MET)

N/I ns 500 m

1000 m

10,000 m

philic cell density and the morphology of digestive aveoli inblue mussels will be addressed here. In 2001 the lysosomalmembrane labilisation period in mussel digestive glandcells was significantly lower for mussels caged 500 m and2000 m from the platform compared to mussels caged atthe reference location (Bilbao et al., 2006), but no sucheffect was seen in 2003 (Table 7). There was also a trendtowards higher relative proportion of basophilic cells inmussels caged in the plume compared to mussels caged atthe reference location (Bilbao et al., 2006). In 2003 thelabilisation period was shortest for the pre-exposure group.Similarly, in both 2003 and 2004 the pre-exposure groupsappeared to be more stressed than caged mussels asregarded basophilic cell volume (Table 7). This observationwas supported by differences in digestive tubuli morpho-metrics between the pre-exposure group and caged mussels(Table 7).

4. Discussion

4.1. PAH in blue mussels

There was no data for pre-exposure levels of PAHs in2001, but concentrations in mussels used for deploymentin 2003 and 2004 were within the background range estab-lished for the Norwegian coast (Green and Knutzen, 2003).The pattern for accumulation of PAHs in caged blue mus-sels along locations away from platforms were similar forthe three years, but there were nonetheless some intriguingdifferences, the most obvious being the difference in abso-lute levels between campaigns. The highest levels of PAHswere found in mussels deployed in 2001 (Statfjord B), fol-lowed by 2003 (Troll B) with mussels exposed in 2003 (Stat-fjord B) being almost a magnitude lower. The 2001 and2003-concentrations corresponded with earlier observa-tions (Utvik et al., 1999; Neff et al., 2006). Utvik et al.(1999) measured RPAH16 = 23 ng/g wet wt in musselscaged in the vicinity of Troll, as compared to our findingsof 14.1 ng/g wet wt, and 34.5 ng/g wet wt measured in mus-sels caged 0.5 km from Troll (2003) and Statfjord B (2001),

respectively (Fig. 2). Neff et al. (2006) did not measure allthe compounds constituting the EPA PAH16, however,the sum-concentration of naphthalene, fluorene, anthra-cene, phenanthrene, fluoranthene, pyrene, chrysene,benzo(b)fluoranthene, benzo(k)fluoranthene and benzo(-g,h,i)perylene in mussels caged 0.5 km downcurrent froma produced water discharge in the Ekofisk area was35.2 ng/g wet wt, as compared to our findings of 12.5 ng/g wet wt, and 31,9 ng/g wet wt measured in mussels caged0.5 km from Troll (2003) and Statfjord B (2001),respectively.

The difference between 2001 and 2003 presumablyreflects differences in total inputs and hydrographical con-ditions at Statfjord and Troll during the two campaigns.The same laboratory analysed the samples in all 3 yearsand inputs of PAHs from Statfjord B were of a similarmagnitude in 2001 and 2004. Since produced water at Stat-fjord has lower density than seawater at the release point itpredominantly distributes to the surface layers. A possibleexplanation for the low concentration found in blue mus-sels is related to the hydrographical conditions where thethickness of the upper water layer was much reduced inthe spring (2001) compared to the autumn (2004) (vanHaren and Howarth, 2004), thus providing a reduced dilu-tion volume for produced water in 2001. As mentionedabove cages were positioned in 2001 according to model-ling of the plume from Statfjord in May (using meteorolog-ical data from 2000). The same locations were used in 2004,but subsequent modelling using true wind data indicatedthat the plume had had an unexpected trajectory, initiallymoving south and then skirting the cage positions (Rye,personal communication). Exposure to effluent was there-fore lower in 2004 compared to 2001. A contributory factormay also be differences in the amount of particulate matterin the water column in 2001 and 2004. During spring (2001)the water column contained more particles (phytoplank-ton) than in the autumn (2004). Because PAHs have highparticle affinity and blue mussels are filter-feeding organ-isms, the amount of particles in the water column is knownto affect PAH-accumulation in mussels (Baumard et al.,1999). Since lipophilic compounds will be associated withorganic particles, the high turbidity during the springbloom would be expected to increase the bioavailabilityof PAHs for filter-feeding organisms such as mussels. Atthe same time the bioavailability of the same compoundswould be expected to decrease for non-filtering organismssuch as fish.

A second interesting observation was the difference inPAH concentrations accumulated in blue mussels cagedat the reference location in the 3 years. This differencemay to a large extent be explained by different depths ofthe pycnocline in May–June as compared to August–September. An upper water layer of 10–15 m as in May2001 would of course provide a much smaller volume fordilution of PAH released from diffuse sources than apycnocline of 40–50 m as is normally found in the autumn(van Haren and Howarth, 2004).

4.2. PAH metabolites in bile

Bile PAH metabolites in cod caged in 2001, measuredeither as pyrene equivalents or as naphthalene metabolites,were in good agreement with results for PAH tissue resi-dues in blue mussels. On the basis of those results onewould expect accumulation of PAH metabolites in codcaged downstream the platform in 2003, but this was notobserved. The most probable explanation is a non-contin-uous exposure of the cages to the effluent plume from TrollB, with lower levels the last few days or the week prior tosampling. The low concentrations of PAH metabolitesobserved in caged cod in 2004 corresponded with the lowlevels of PAH accumulated in mussels that year. The ele-vated concentration in pre-exposure cod was presumablyeither caused by oil contamination at the fish farm or bycomponents in the commercial feed, highlighting the needto handle organisms for caging with special care.

4.3. Cytochrome P4501A activity

Hepatic cytochrome P4501A activity in fish would beexpected to be induced by exposure to PAHs (Hylland,2006), but increased cytochrome P4501A activity in fishcaged closer to the platform was not observed for any ofthe campaigns reviewed here. The main components ofproduced water are phenols, naphthalenes and 3-ringPAHs, none of which are known to be strong inducers ofCYP1A. Earlier studies with other fish species have shownthat 2- or 3-ring PAHs may not induce (Gerhart and Carl-son, 1978) or may indeed inhibit hepatic CYP1A (McKeeet al., 1983; Willett et al., 2001). Results are not clear asa fourth study has shown a syngergistic interaction betweenfluoranthene and carcinogenic PAHs, albeit for a differentfish species (Basu et al., 2001). Produced water effluents docontain higher molecular-weight, carcinogenic PAHs thatmay be expected to induce CYP1A, but the levels were pre-sumably either too low to cause such induction or therewas a partial inhibition by low-molecular weight PAHs.

4.4. Glutathione S-transferase

The apparent gradient-related induction in hepatic glu-tathione S-transferase (GST) observed in 2001 was notfound in 2003 or 2004, although GST did vary significantlybetween some of the locations in the latter 2 years. Themechanisms of GST induction in fish are not well under-stood. The presence of multiple GST isoforms in fish hasrecently been confirmed through characterisation of genes(Donham et al., 2005a,b). In mammals, the induction ofGSTs involves promoter sequences including electrophileresponse elements (EpREs) and antioxidant response ele-ments (AREs) (Rushmore et al., 1991), as well as AhR-responsive elements. As will be evident, just about allorganic contaminants may therefore affect the expressionof GST isoforms (Perez-Lopez et al., 2002), but such differ-ences may not be evident in ‘‘total” cytosolic GST activity

(as measured using CDNB as substrate; George, 1994).Produced water is a complex mixture and the compositionwill differ between platforms and may change during thelifetime of a production field. There may be componentsthat affect GST in the Statfjord B effluent (2001 and2003) which were not present in the Troll B effluent(2003). A recent study has indicated that there is seasonalvariation in hepatic GST activity in cod (Hylland et al.,2006a), although this would not be expected to affect differ-ences (if present) between locations in any of thecampaigns.

4.5. DNA adducts

During the 2001 campaign higher concentrations ofhepatic DNA adducts were found in cod caged at the ref-erence location, away from any known point source ofPAHs, than at the locations in the effluent plume fromStatfjord B (Balk et al., 2006). In 2003, no differences werefound between any group, but some individuals in allgroups, including the pre-exposure group, had elevatedconcentrations of adducts. Experimental studies haveshown that adducts in fish, including cod, may be formedwithin days if the exposure concentration is sufficientlyhigh (Aas et al., 2000; Ericson et al., 1999; Ericson andBalk, 2000). An exposure period of 5–6 weeks as in thestudies described herein should therefore be sufficient forthe generation of adducts. As mentioned above there is lim-ited knowledge of how 2–3 ring PAHs are metabolised bydifferent fish species and studies indicate both inhibitionand potentiation of cytochrome P4501A (CYP1A), themain metabolising system for PAHs (Basu et al., 2001;Gerhart and Carlson, 1978; McKee et al., 1983; Willettet al., 2001). Since the formation of adducts depends onthe generation of PAH radicals by CYP1A, any inhibitionof this system by low-molecular weight PAHs would alsocause reduced adduct levels. The observation of adductsin pre-exposure fish in 2003 also has implications for theuse of farmed cod for caging studies. Further work isneeded to identify the sources of adducts in farmed cod.Oil spills, combustion engine exhaust (Tjarnlund et al.,1995) and components in feed are possible sources of sub-stances that may give rise to adducts.

4.6. Lysosomal stability in digestive cells

It has been well established that environmental stressorsreduce the stability (decreased labilisation period, LP) ofthe lysosomal membrane in mussel digestive cells (Marigo-mez et al., 2006). In 2001, LP values were lower in musselsaround the platform than in those from the reference site.Likewise, in 2003, particularly low LP values were obtainedin digestive gland of mussels from sites close Troll B, indi-cating that some source of stress was affecting musselhealth status. This response was not measured in 2004.Overall, it can be concluded that reduction in lysosomalmembrane stability in mussel digestive cells is a responsive

biomarker, even though source mussels appeared to be suf-fering some stress (LP values below 20 min; Viarengo et al.,2000). Indeed, histological data for 2003 actually indicatedthat pre-exposure mussels were spawning (see Section4.10).

4.7. Lysosomal stability in haemocytes

No contaminant-related differences were observedbetween groups of blue mussels caged at different locationsin lysosomal stability, measured as neutral red retention, ineither 2003 or 2004. The analyses were performed on boardthe vessel in 2003 (but for only three locations due to badweather) and for blue mussels transported back to the lab-oratory in 2004. Reduction in the lysosomal membrane sta-bility is a widely established biomarker of general stress(UNEP/RAMOGE, 1999; Cajaraville et al., 2000; Viar-engo et al., 2000), but.current techniques do not appearappropriate for use under the conditions generally encoun-tered in the North Sea.

4.8. Micronucleus frequency

Even at the very low exposure levels present in 2004there was an apparent exposure-dependent increase inhematocyte micronuclei in mussels caged closer to the plat-form. A similar trend was seen for cells from cod caged atthe same locations. Cytogenetic damage has earlier beendescribed in molluscs from harbours, oil terminal areasand follwing an accidental oil spill in the Baltic Sea (Bar-siene and Barsyte Lovejoy, 2000; Barsiene, 2002; Barsieneet al., 2006b). It has been suggested that increased genotox-icity in mussels from oil spill areas may primarily dependon the levels of water soluble components (Carls et al.,2001). The genotoxic potential of North Sea crude oil hasearlier been indicated in studies with turbot (Scophthalmus

maximus) blood (mature) and head kidney (immature)erythrocytes of Atlantic cod (Barsiene et al., 2006a).Short-term exposure of juvenile sea bass (Dicentrarchuslabrax) to 0.3, 0.9 and 2.7 lM of naphthalene causedincreased micronucleus formation in liver cells comparedto controls (Gravato and Santos, 2002) and exposure ofeel (Anguilla anguilla) to naphthalene similarly resulted inthe development of micronuclei and other nuclear abnor-malities (Teles et al., 2003). The sensitivity of micronucleifrequency as a biomarker of contaminant stress for musselshas earlier been indicated in experimental studies (Burgeotet al., 1995; Hylland et al., 2006c,d; Venier et al., 1997).

4.9. Vitellogenin

Plasma vitellogenin (vtg) was analysed following allsampling campaigns in the North Sea. Elevated levels ofvtg compared to the reference station were only observedat the station closest to the Statfjord B production plat-form (500 m station) during the 2001 campaign, suggestingthat discharges contain compounds capable of causing

estrogenic effects. The measured vtg levels were howeverseveral orders of magnitude lower than what was observedin female cod during the 2001 campaign and levels in malesexposed artificially to the potent estrogenic compound 17b-estradiol (Hylland, 2000). Extraction of water samplescombined with several in vitro bioassays in the same mon-itoring study has verified that low levels of estrogenic com-pounds were present in the waters of the Statfjord area andthat a isomeric mixture of low molecular weight alkylphe-nols and nonylphenol contribute to the observed estrogenicactivity (Thomas et al., 2004, 2006; Tollefsen et al., 2006).

The observed plasma levels of vtg in both males andfemales were found to be several order of magnitude higherduring the two water column surveys (2003 and 2004) thanthose reported for Atlantic cod in 2001. The discrepancybetween these studies may be explained by the fact thatthey were performed with fish of different origin, matura-tion status and age. In addition, the campaigns were per-formed at different times of the year. It is known thatvitellogenin production increase with age and maturity ofthe fish (Sole et al., 2002) as well as size (Scott et al.,2006a) and that there is a natural seasonal variation invtg levels in juvenile cod (Hylland, unpublished).

Several males sampled prior to the exposure period (pre-exposure group) had plasma levels of vtg that were compa-rable to background levels found in female fish. Similar ele-vation of vtg in males prior to exposure studies has beendocumented elsewhere (Knudsen et al., 1997) and has beensuggested to be caused by male fish being affected by 17b-estradiol (E2) excreted by females during close confine-ment, a typical situation occurring during high densityrearing of fish prior to deployment in the field.

Interestingly, plasma levels of vtg in females close to theStatfjord B production platform (2004) were found to belower than that of fish deployed at the reference station.The rationale for this observation is not immediately evi-dent, but it is well documented that certain PAHs such as3-methylcholanthrene and b-naphthoflavone are able toact as anti-estrogens and reduce the production of vtg infish in vitro studies (Navas and Segner, 2000). Althoughno similar effects were detected during the 2001 and 2003campaigns, interference of anti-estrogenic chemicals ofanthropogenic origin with normal estrogenic signalling infemale fish can not be ruled out.

4.10. Digestive gland histopathology

There appeared to be a clear, exposure-related effect forhistological endpoints in digestive gland of blue mussels in2001. No such effect was seen in either 2003 or 2004. Asmentioned above, mussels deployed in May 2001 werefrom Ireland, whereas mussels deployed in the autumn2003 and 2004 were from central Norway (Trøndelag).One reason to shift monitoring from spring to autumnwas to avoid spawning in the mussels, but the data for2003 and 2004 actually indicated that some of the pre-exposure mussels were spawning. This was possibly due

to handling and/or transport stress. Their health status wastherefore improved following field deployment in 2003 and2004 compared to the pre-exposure situation, even follow-ing exposure to effluent. Again, as has been stressed abovefor fish, it is crucial that any organism to be used in fieldcaging studies has low initial load of contaminants but itis just as important that they are in an appropriate physio-logical state and a good health status is established beforedeployment.

5. Summary and conclusions

Three years of water column monitoring near offshoreproduction platforms have shown the importance ofincluding both chemical analyses and biological effectsmethods in such surveys. The results suggest that the inputsdo not cause serious environmental impacts even thoughcomponents were detected in mussels and some biologicalresponses were observed,. Chemical analyses, i.e. PAH tis-sue residues in blue mussels and PAH bile metabolites incod, provided information about the effluent exposureand the bioavailability of effluent components and therebyincreased the scope for interpretation of biological effects.The effluent exposure for caged organisms was highest in2001 followed by 2003, with much lower levels in 2004compared to the other 2 years. Some effluent-relatedresponses were observed each of the three years and therange of biological effects methods that were found torespond corresponded to the observed environmental levelsof produced water components. The concentrations of con-taminants such as PAHs and alkylphenols reported in thesurveys were low compared to levels found in many coastalareas and appeared to approach the detection limit formany of the effects methods employed.

There were no easily interpretable responses in PAH-sensitive methods such as hepatic cytochrome P4501Aactivity or DNA adducts in cod from the three surveys.A possible explanation is the large proportion of 2- and3-ring PAHs compared to 4- and 5-ring PAHs in producedwater. In addition, the exposure of cod to contaminantswas low compared to levels found in polluted coastal areas,as indicated by low concentrations of bile PAH-metabo-lites. Produced water does appear contain sufficient levelsof xenoestrogens to influence fish, but this would only berelevant for some produced waters and close to platforms.Mussel histopathology, including lysosomal destabilisationin hepatopancreas, and haemocyte micronucleus frequencywere two methods that indicated that caged mussels wereaffected by components in produced water. In addition tobeing able to provide direct information about contami-nant impacts, histopathology also provided informationcritical for proper interpretation of results for other meth-ods, e.g. spawning status for mussels or parasite infectionfor fish.

The results from monitoring in 2001–2004 suggest thatsome of the methods used are either not appropriate or suf-ficiently sensitive to be used to monitor low levels of pro-

duced water inputs. This will become even more relevantin the future as North Sea production platforms implementcleaning processes that will decrease inputs, which willresult in lower environmental levels of the components inquestion.

Four issues have become apparent through the 3 yearsof monitoring and research reported here: (1) it is vital thatorganisms to be used for caging are in optimal conditionwith a known health status and gonadal maturation andpreviously unexposed, (2) monitoring activities shouldalways allow the inclusion of novel methods in additionto a core programme to facilitate the development of mon-itoring techniques, (3) the position and number of caginglocations is critical and should not make presumptionsabout hydrographical conditions, and finally (4) integratedmonitoring should include chemical analyses as well as arange of both contaminant-specific and general effectsmethods and not be limited to methods that would reflect‘‘expected” impacts.

Acknowledgements

The authors would like to thank the steering group,crews, cruise leaders and participants of the 2001 ICESworkshop on biological effects of contaminants in pelagicecosystems (BECPELAG). Thanks are also due to KarlHenrik Bryne, Statoil, who was in charge of field work in2003 and 2004, and Bjørn Serigstad, who built and main-tained cages for all three campaigns. The activity reviewedin this paper was partly funded by the Norwegian oil indus-try through OLF and the BECPELAG part (2001), whichwas also funded by the Research Council of Norway.

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