Chemosphere 84 (2011) 504–511

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Chemosphere

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Naphthenic acid biodegradation by the unicellular alga Dunaliella tertiolecta

Dean M. Quesnel a,b, Iyswarya M. Bhaskar a,b, Lisa M. Gieg b, Gordon Chua a,b,⇑a Institute for Biocomplexity and Informatics, University of Calgary, Alberta, Canada T2N 1N4b Department of Biological Sciences, University of Calgary, Alberta, Canada T2N 1N4

a r t i c l e i n f o

Article history:Received 10 December 2010Received in revised form 8 March 2011Accepted 9 March 2011Available online 2 April 2011

Keywords:Naphthenic acidsAlgaeOil sandsTailings pondsBiodegradationGas chromatography–mass spectrometry

0045-6535/$ - see front matter Crown Copyright � 2doi:10.1016/j.chemosphere.2011.03.012

⇑ Corresponding author at: Department of BioloCalgary, Alberta, Canada T2N 1N4. Tel.: +1 403 220 7

E-mail address: gchua@ucalgary.ca (G. Chua).

a b s t r a c t

Naphthenic acids (NAs) are a major contributor to toxicity in tailings waste generated from bitumen pro-duction in the Athabasca Oil Sands region. While investigations have shown that bacteria can biodegradeNAs and reduce tailings toxicity, the potential of algae to biodegrade NAs and the biochemical mecha-nisms involved remain poorly understood. Here, we discovered that the marine alga Dunaliella tertiolectais able to tolerate five model NAs (cyclohexanecarboxylic acid, cyclohexaneacetic acid, cyclohexaneprop-ionic acid, cyclohexanebutyric acid and 1,2,3,4-tetrahydro-2-naphthoic acid) at 300 mg L�1, a level whichexceeds that of any single or combination of NAs typically found in tailings ponds. Moreover, we showthat D. tertiolecta can metabolize four of the model NAs. Analysis of NA-amended cultures of D. tertiolectavia low resolution gas chromatography–mass spectrometry allowed us to quantify decreasing NA levels,identify metabolites, and formulate putative mechanisms of biodegradation. Degradation of cyclohexa-nebutyric acid and cyclohexanepropionic acid proceeded via b-oxidation and resulted in the transientaccumulation of cyclohexaneacetic acid and cyclohexanecarboxylic acid, respectively. Cyclohexanecar-boxylic acid was metabolized via 1-cyclohexenecarboxylic acid suggesting that further degradationmay occur by step-wise b-oxidation. When D. tertiolecta was inoculated in the presence of oil sands tail-ings water from the Athabasca region, biodegradation of single-ring NAs was observed relative to con-trols. This result corroborates the trend we observed with the single-ring model NAs.

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1. Introduction

The generation of tailings waste from the Athabasca Oil Sandsvia hot water extraction of bitumen poses challenges for the eco-logical and commercial sustainability of the region. Presently, tail-ings are stored in settling ponds on company leases so they canlater be reclaimed as there is a zero discharge policy regardingwastewater (Giesy et al., 2010). The lack of a suitable method totreat this wastewater has resulted in enormous ponds of liquidtailings containing naphthenic acids (NAs) that cover 130 km2

(Kean, 2009). As these ponds continue to expand, there is growingconcern over their environmental impact. This has led to intenseglobal scrutiny of the mining activities in the Athabasca Oil Sandsregion which has driven considerable reclamation efforts. One ofthe major hurdles involved in reclaiming these ponds is the pres-ence of recalcitrant NAs which accumulate in wastewater duringthe caustic hot water extraction of bitumen from surface-minedoil sands (Quagraine et al., 2005). NAs are naturally-occurring ele-ments of crude oil which exist as complex mixtures of alicyclic car-boxylic acids that have been classically defined by the molecular

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formula CnH2n+zO2. Typical tailings water contains NAs with anaverage Z number of �4 to �6 (Clemente and Fedorak, 2005).The Z designation of the compounds refers to the hydrogen defi-ciency introduced by the formation of ring structures. Recently ithas been suggested that this classical definition does not accu-rately describe the total fraction of toxic organic acids present intailings, many of which contain multiple hydroxyl/carboxyl groupsas well as heteroatoms like N or S (Grewer et al., 2010). These or-ganic acids are the leading contributor to tailings water toxicitywith acute and chronic detrimental effects observed in aquaticand terrestrial organisms (Frank et al., 2009). Toxicity associatedwith classical NA and organic acid contamination is difficult to mit-igate which has hindered reclamation efforts of the oil sands tail-ings ponds.

Several studies have shown that microbial biodegradation candecrease the toxicity of the tailings waters over time (Quagraineet al., 2005; Frank et al., 2008; Johnson et al., 2010) suggesting thatin situ bioremediation of NA and organic acid contaminated waste-water is a plausible means to address tailings management issuesfaced by Athabasca Oil Sands operators. A variety of microbialcommunities from the Athabasca region along with non-indigenous bacterial species have been investigated for theirpotential to specifically degrade classically-defined NAs (Hermanet al., 1994; Clemente et al., 2004; Scott et al., 2005; Del Rio

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et al., 2006; Hadwin et al., 2006; Biryukova et al., 2007). Predomi-nantly non-photosynthetic bacteria from the Athabasca regionwere able to readily degrade simple model NAs and NAs from com-mercial sources, but degradation of NAs from tailings water wasconsiderably less efficient (Clemente et al., 2004). Scott et al.(2005) demonstrated that the microbial degradation of a commer-cially-available NA mixture was almost complete after 14 dwhereas NAs in tailings water were more resistant to degradationwhere 75% still remained after 40 d. Further, it has been suggestedthat only a subset of tailings NAs are readily biodegradable, leadingto an enrichment of high molecular weight NAs once fresh effluentaddition has ceased (Holowenko et al., 2002). While studies withmodel NAs do not accurately estimate the potential for in situbioremediation, they have proven useful for the identification ofstructural factors influencing degradation and the elucidation ofdegradation pathways. Studies involving model NAs have revealedthat the extent of alkyl-chain branching and number of ring struc-tures are linked with NA persistence (Han et al., 2008; Smith et al.,2008; Johnson et al., 2010). Compounds such as cyclohexanecar-boxylic acid and cyclohexaneacetic acid have been investigatedfor their role in benzoate metabolism as well as for their potentialas NA surrogates. These investigations have led to the identifica-tion of a number of biodegradation mechanisms that allow for abasic understanding of possible fates of recalcitrant NAs presentin tailings water (Rho and Evans, 1975; Smith and Callely, 1975;Taylor and Trudgill, 1978; Ougham and Trudgill, 1982; Yoshizakoet al., 1991; Kuver et al., 1995; Elshahed et al., 2001; Iwaki et al.,2008). These elucidated pathways provide reference for compari-son of degradation products allowing putative mechanisms to beidentified.

Algal communities have been associated with the tailings pondsof the Athabasca Oil Sands (Leung et al., 2001, 2003). However,much less is known about the ability and mechanisms by whichthese algae can utilize NAs compared with indigenous bacteria. Arecent study indicated that algae may also be capable of degradingmodel and tailings associated NAs (Headley et al., 2008). Of the 12algal species characterized, only two distinct Naviculla strainsremoved the model NA trans-4-methylycyclohexaneacetic acid(4-MCHAA), while only one of these strains removed the cis-isomerof 4-MCHAA. The mechanisms by which these compounds weremetabolized have yet to be elucidated. In contrast, none of the algalspecies were able to effectively degrade a combination of NAsderived from oil sands tailings water. Furthermore, the green algaChlorella pyrenoidosa has been shown to metabolize cyclohexane-acetic acid and produce hydroxylated metabolites (Yoshizakoet al., 1991). While both studies begin to address the potential ofalgae to remediate NA-contaminated wastewater, more workneeds to be done including the characterization of algal prolifera-tion in tailings water and their contribution to the degradation ofrecalcitrant tailings NAs.

The aim of this study was to identify algal species capable ofdegrading classically-defined NAs and to characterize their mech-anisms of metabolism. It was initially hypothesized that algal tol-erance to NAs may indicate the presence of NA degradation activityor another method of detoxification. Using this rationale, wescreened algal and cyanobacterial species for growth in the pres-ence of five model NAs at 300 mg L�1, a concentration greatlyexceeding NA levels typically found in oil sands tailings water.Monitoring the biodegradation of these model NAs in algal culturesalso allowed for the identification of downstream metabolites andputative mechanisms involved in NA breakdown. Furthermore, al-gal growth in tailings water produced small shifts in the distribu-tion of tailings NAs suggesting that the alga investigated herewas also capable of mitigating a subset of classically-defined NAsfound in tailings. While the use of low resolution mass spectrom-etry is limited to the identification of classical NAs and is prone

to false positive identification of non-classical organic acids, thetechnique enabled us to observe general trends in tailings NA bio-degradation that were corroborated by model NA degradation data(Martin et al., 2008; Grewer et al., 2010). These results indicate thatcertain algal species may possess bioremediation capabilities thatcan be applied to NA-contaminated tailings water.

2. Materials and methods

2.1. Algal medium and growth conditions

The algal species Dunaliella tertiolecta (UTEX LB999) andChlorella vulgaris (CPCC 145) along with the cyanobacteriumSynechococcus leopoliensis (CPCC 102) were obtained from theUTEX Culture Collection of Algae (Austin, TX) and the CanadianPhycological Culture Center (Waterloo, ON). Algal and cyanobacte-rial species chosen have not been previously associated with oilsands tailings, although we are not certain whether C. vulgaris orD. tertiolecta are present in tailings as no eukaryotic communityanalysis has ever been done. Species were selected based on envi-ronmental tolerance to different conditions commonly encoun-tered in the Athabasca oil sands tailings such as high salt levels,low temperatures and alkaline pH. Geographic distribution of phy-la and availability of genome sequences/metabolic pathway datawas also considered when selecting species to screen. Stockcultures of D. tertiolecta were grown in 125 mL Erlenmeyer flaskscontaining 50 mL of liquid f/2 medium adjusted to pH 8(0.883 mM NaNO3, 0.0363 mM NaH2PO4� � �H2O, 0.107 mM Na2-

SiO3� � �9H2O, 10 lM FeCl3� � �6H2O, 10 lM Na2EDTA� � �2H2O, 40 nMCuSO4� � �5H2O, 30 nM Na2MoO4� � �2H2O, 80 nM ZnSO4� � �7H2O, 50nM CoCl2� � �6H2O, 0.9 lM MnCl2� � �4H2O, 0.1 nM cyanocobalamin,2 nM biotin and 0.3 lM thiamine� � �HCl) using an artificial seawa-ter base (10.379 g L�1 NaCl, 0.294 g L�1 KCl, 0.085 g L�1 NaHCO3,0.0373 g L�1 NaBr, 0.0113 g L�1 H3BO3, 0.0014 g L�1 NaF,2.75 g L�1 Na2SO4� � �10H2O, 4.698 g L�1 MgCl2� � �6H2O, 0.658 g L�1

CaCl2� � �2H2O and 0.0107 g L�1 SrCl2� � �6H2O). S. leopoliensis wasgrown in 50 mL of liquid Bold 3 N Medium and C. vulgaris wasgrown in 50 mL of liquid BG-11 medium as described by UTEX.org.Cultures were incubated at room temperature under 48 in. F40fluorescent bulbs (1900 lumens/bulb) (General Electric, Fairfield,CT) with an 8 h light: 16 h dark cycle and were capped with sterilefoam stoppers. Growth of all cultures was monitored via opticaldensity at 680 nm with a Spectramax Plus microplate reader(Molecular Devices, Sunnyvale, CA). Tailings water samples fromSuncor Pond 6 were generously provided by Suncor Energy Inc.

2.2. Naphthenic acid tolerance assays

D. tertiolecta cultures (25 mL) were spiked with stock solutions(7.5 g/L in Milli-Q water) of cyclohexanecarboxylic acid, cyclohex-aneacetic acid, cyclohexanepropionic acid, cyclohexanebutyricacid, or 1,2,3,4-tetrahydro-2-naphthoic acid (Sigma–Aldrich, St.Louis, MO) to a final concentration of 300 mg L�1. To dissolve themodel NA compounds in the stock solutions, two pellets of NaOHwere added per 50 mL. Under these conditions, the compounds ex-ist as sodium naphthenates which makes them more readily solu-ble in water. As a control, Milli-Q� water without NAs, adjusted toa pH of 12, was spiked into cultures at the same volume as with thestock solutions. The pH of spiked medium was approximately 9.5at the onset of growth measurements. The 25 mL incubations wereinoculated with 0.5 mL of a stock culture of D. tertiolecta(OD680 � 0.16) and grown under the conditions described abovein 50 mL Erlenmeyer flasks with sterile foam stoppers. Growth ofthe cultures was monitored daily by measuring the OD680 duringthe dark cycle.

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2.3. Biodegradation assays of model naphthenic acids

D. tertiolecta cultures were prepared in sterile 125 mL screw-topErlenmeyer flasks containing 50 mL of f/2 medium. Individualcultures were spiked separately with single stock solutions ofcyclohexanecarboxylic acid, cyclohexaneacetic acid, cyclohexane-propionic acid, cyclohexanebutyric acid or 1,2,3,4-tetrahydro-2-naphthoic acid to a final concentration of 200 mg L�1 and grownunder conditions described above. The pH after addition of stocksolutions was approximately 9.4. Sterile controls containing200 mg L�1 of the test compound and cultures lacking NAs wereperformed alongside experimental samples to rule out any non-biological mediated degradation (Fig. SM-1). After the cultureswere sealed, they were vented once a week to allow CO2 and O2

exchange. At 7 d intervals, 5 mL of culture fluids were removedfor analysis and acidified to pH 2 (using 5.2 M HCl). The OD680 ofthe cultures at each 7 d interval was also taken. The acidifiedsample was centrifuged at 2126 g for 10 min at room temperatureto pellet the cells. The supernatant was then transferred to clean4 oz Boston round bottles with Teflon cap liners. Two volumes ofdichloromethane (Sigma–Aldrich, St. Louis, MO) were added tothe bottles and the supernatant was extracted on an orbital rotator(Model: 260200, Boekel Scientific, Feasterville, PA) for 2 h. Follow-ing extraction, samples were poured through 125 mm phaseseparator Whatman 2200 filter papers previously rinsed withdichloromethane. The extraction bottle was then rinsed with10 mL of dichloromethane followed by 5 mL of dichloromethanewhich was then poured through the phase separator. The filter pa-per was again rinsed with dichloromethane and the volume of theflow through was reduced to 1.5 mL with a Rotovapor R II rotaryevaporator (BÜCHI Labortechnik AG, Flawil, St. Gallen, Switzerland).The sample was then transferred to a 1.5 mL auto-samplervial (Agilent Technologies, Santa Clara, CA) and derivatizedwith 150 lL of N,O-bis(trimethylsilyl)trifluoroacetamide (ThermoScientific, Waltham, MA) for 10 min at 60 �C. Naphthenic acid lev-els in extracted samples were measured using an Agilent 7890Agas chromatograph equipped with an Agilent HP-5MS 30 m �0.25 mm � 0.25 lm column and an Agilent 5975C mass selectivedetector (Agilent Technologies, Santa Clara, CA). One microlitre ofsample was injected via an auto-sampler with a 50:1 injector split.The oven was held at 50 �C for 5 min, increased to 250 �C at a rateof 8 �C/minute and held at 250 �C for another 5 min. Total ion chro-matogram integration and analysis of extracted compounds wereperformed using Enhanced MSD ChemStation E.02.00.493 RTEintegrator software (Agilent Technologies, Santa Clara, CA).

2.4. Biodegradation assays of tailings naphthenic acids

A mixture of 60% filter sterilized tailings pond water fromSuncor Energy Inc. (Pond 6, collected February 2010) and 40% f/2medium was used to set up 100 mL cultures. Filter sterilizationwas performed with Millipore Express� PLUS 0.22 lm Stericup�. Theexperimental cultures were inoculated with 1 mL of a D. tertiolectaculture with an OD680 � 0.16 and grown for 6 weeks in sealed250 mL Erlenmeyer flasks. A sterile control was performed concur-rently with the experimental cultures. These were vented weekly.After 6 weeks, the cultures were sacrificed and the NA fraction wasextracted. A sterile control consisting of 100 mL of tailings/mediummixture was also extracted immediately after preparation to estab-lish a time 0 baseline reading. All 100 mL cultures were acidified toa pH of 2 as described previously and centrifuged at 2126 g for10 min at room temperature to remove cellular debris. The super-natant was then transferred to a 16 oz Boston round bottle with aTeflon lined cap and prepared for GC–MS analysis as describedabove with the exception that NAs were derivatized with 200 lLof N-methyl-N-(tert-butyldimethylsilyl)-trifluoroacetamide

(Thermo Scientific, Waltham, MA) for 1 h at 60 �C. NAs were quan-tified according to the procedure described by Holowenko et al.(2002)

3. Results and discussion

3.1. Model naphthenic acid tolerance

The ability of algal strains to degrade NAs has not beenthoroughly investigated. To extend knowledge in this area, weexamined algal and cyanobacterial species from diverse habitatsnot previously tested for NA degradation activity. The organismsinvolved initially in our studies included a marine and a freshwateralga (D. tertiolecta and C. vulgaris, respectively) and the cyanobac-terium S. leopoliensis. The strains were first separately treated withfive model NAs and assayed for tolerance. We hypothesized thatstrains exhibiting higher tolerance to NAs would be more likelyto possess biochemical activities associated with NA degradation.The five model NAs tested in the tolerance assays were cyclohex-anecarboxylic acid (CHCA), cyclohexaneacetic acid (CHAA), cyclo-hexanepropionic acid (CHPA), cyclohexanebutyric acid (CHBA)and 1,2,3,4-tetrahydro-2-naphthoic acid. CHCA and CHAA werechosen as they have been investigated previously in other organ-isms, thereby providing a reference for comparison (Rho and Evans,1975; Smith and Callely, 1975; Taylor and Trudgill, 1978; Oughamand Trudgill, 1982; Yoshizako et al., 1991; Kuver et al., 1995;Elshahed et al., 2001; Iwaki et al., 2008). CHPA and CHBA wereselected as it was believed that b-oxidation may play a role in sidechain degradation, thus producing CHCA and CHAA, respectively.This would allow for easy identification of b-oxidation processesinvolved in the biodegradation of these two model NAs. Althoughnot a NA by the strict classical molecular definition (CnH2n+zO2),we chose the NA analogue 1,2,3,4,-tetrahydro-2-naphthoic acidto model double-ring NAs which may be more representative ofcomplex NAs found in tailings water.

Among the species screened for tolerance to these model NAs,only D. tertiolecta and S. leopoliensis strains exhibited robust growthat levels that exceed those found in NA-contaminated oil sands tail-ings water (Fig. 1; data not shown). The total NA content of tailingswater from this region typically ranges from 40 mg L�1 to120 mg L�1 (Clemente et al., 2004; Frank et al., 2009) while ourcultures were able to proliferate on single model NAs at aconcentration of 300 mg L�1. With respect to D. tertiolecta, thegrowth rate did not appear to be affected by single-ring modelNAs as these cultures grew similarly to controls after 14 d ofincubation (Fig. 1). The two-ringed 1,2,3,4-tetrahydro-2-naphthoicacid had the highest molecular weight among the model NAs testedand the most adverse effect on the overall growth of D. tertiolecta.The algal culture containing 300 mg L�1 of this model NA only grewto approximately 50% of the control culture after 14 d. While thiscompound demonstrated greater recalcitrance than the single-ringed NAs, the alga was still capable of proliferating andthereby exhibiting tolerance. Altogether, this data suggests that D.tertiolecta’s exhibited NA tolerance may be due to an ability to de-grade and/or neutralize the NAs to concentrations that allow signif-icant levels of growth (see below). In contrast, S. leopoliensis alsodisplayed tolerance to the model NAs, but displayed no evidenceof NA degradation after 2 weeks of incubation (data not shown).Therefore, further characterization of this species was not pursued.

3.2. Cyclohexanebutyric acid and cyclohexaneacetic acidbiodegradation

Cultures of D. tertiolecta containing 200 mg L�1 of CHBA wereanalyzed by GC–MS over a period of 35 d. During the time course

Fig. 1. Growth curves of D. tertiolecta amended with cyclohexanecarboxylic acid (CHCA), cyclohexaneacetic acid (CHAA), cyclohexanepropionic acid (CHPA), cyclohexa-nebutyric acid (CHBA), or 1,2,3,4-tetrahydro-2-naphthoic acid (TH-NA) at 300 mg L�1 (n = 3). With the exception of TH-NA, addition of model NAs did not inhibit growth incomparison to control cultures after 14 d.

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experiment, levels of CHBA dropped to below detectable levelswithin 21 d while a new compound with a retention time of15.640 min appeared transiently between day 7 and day 28(Fig. 2A, Fig. SM-2A-C). This compound was identified as CHAAby matching the GC retention time and mass spectral profile withan authentic standard of CHAA (data not shown). While only semi-quantitative, this was the first indication of NA metabolism by thisalga which allowed us to propose a simple mechanism by whichthe straight chain side group of CHBA could be metabolized, result-ing in CHAA (Fig. 2B). Further, CHAA in cultures containing CHBAtransiently appeared (Fig. 2A), indicating that CHAA was furtherdegraded. This subsequent degradation of CHAA in CHBA treatedalgal cultures cannot occur via b-oxidation (Han et al., 2008).Due to a tertiary carbon at the b position of CHAA, further degrada-tion must occur by a different mechanism (see below). These re-sults suggest that multiple mechanisms are being employed bythe alga to metabolize CHBA.

To further investigate the fate of CHAA by D. tertiolecta, culturescontaining 200 mg L�1 CHAA were grown for 49 d and periodically

A

B

Fig. 2. Decrease of CHBA and appearance of the metabolite CHAA in cultures of D. terti(n = 3) (A). Proposed b-oxidation of CHBA to CHAA with theoretical production obiotransformation observed (B).

sampled for GC–MS analysis. After 21 d, CHAA levels had decreasedby approximately 60% (Fig. 3A) confirming that degradation wasoccurring and that it was not initiated by the presence of a precur-sor NA such as CHBA. Between 42 and 49 d, CHAA was no longerdetected (data not shown). Several emergent peaks represent-ing metabolites were observed in cultures initially containing200 mg L�1 CHAA after 14 d of incubation (Fig. SM-3). Thesemetabolites were tentatively identified by their mass spectral pro-files as authentic standards were not available (Fig. 3B and C). Themass ion indicated by both spectra was at an m/z of 212, which wastwo mass units less than the mass ion observed for CHAA (Fig. 2B).This suggested a possible dehydrogenation step and the formationof a carbon–carbon double bond in the degradation process ofCHAA. Furthermore, two distinct fragmentation patterns were ob-served which indicated the formation of two unique products withthe same mass ion (Fig. 3B and C). One of these two products islikely cyclohexylidene acetic acid (Fig. 3B). Assuming that frag-mentation of the molecule occurs preferentially at the allylic posi-tion (Silverstein et al., 2005), electron bombardment of

olecta initially containing 200 mg L�1 CHBA as measured by low resolution GC–MSf acetate, which was not measured, likely describes the process yielding the

A

B C

Fig. 3. Decrease of CHAA levels in cultures of D. tertiolecta initially containing 200 mg L�1 CHAA as measured by low resolution GC–MS (n = 3) (A). Mass spectra of identifiedmetabolites both have mass ions of m/z = 212 (B,C), each with a distinct fragmentation pattern. Two possible dehydrogenation products of CHAA are cyclohexylidene aceticacid (B) and 1-cyclohexeneacetic acid (C) which are shown along with their respective spectra (spectra are of trimethylsilyl derivatives).

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cyclohexylidene acetic acid would result in a large proportion ofmolecules fragmenting between the keto and O-TMS groups ofthe trimethylsilyl derivative. This would produce m/z fragmentsof 123 and 89. We would also expect no fragments correspondingto ions that would have formed as a result of cleavage between thecyclohexane ring and the keto group of the side chain (expected m/zof 117 and 132). In Fig. 3B, the fragment with the highest abun-dance occurred at an m/z of 122, thus supporting cyclohexylideneacetic acid as the identified metabolite. The other product is likely1-cyclohexeneacetic acid (Fig. 3C). The allylic position distal to thering in 1-cyclohexeneacetic acid would result in fragmentationoccurring at a high frequency between the first carbon of the acetateside chain and the keto group. This would yield a fragment with anm/z of 117 as seen in Fig. 3C. There is also a low abundance peak atan m/z of 131.9 which suggests that a cleavage between the cyclo-hexane ring and the acetate side chain may also be occurring. Thispeak was not seen in Fig. 3B and may be lacking as a result of the car-bon–carbon double bond position in cyclohexylidene acetic acid.

The conversion of CHAA into cyclohexylidene acetic acid hasbeen previously suggested to occur in a bacterial strain, Arthrobactersp. strain CA1 (Ougham and Trudgill, 1982). Subsequent degrada-tion of cyclohexylidene acetic acid by Arthrobacter sp. CA1 involvesthe formation of a capro-lactone intermediate prior to ring opening.Our data indicate that D. tertiolecta may be able to metabolize CHAAin a manner similar to Arthrobacter, but also through another mech-anism that produces 1-cyclohexeneacetic acid. Although mecha-nisms allowing for the biotransformation of 1-cyclohexeneaceticacid have not been studied in algae, this compound is susceptibleto a-oxidation which would result in cyclohexenecarboxylic acid.Indeed, a-oxidation was believed to play a role in the degradation

of CHAA by the bacterium Alcaligenes sp. PHY12 where CHAA istransformed to CHCA followed by a dehydrogenation step to form1-cyclohexenecarboxylic acid (Rontani and Bonin, 1992). This dehy-drogenation product has also been associated with CHCA degrada-tion in mammalian systems which may be more relevant to D.tertiolecta than prokaryotic metabolism. In eukaryotic systems,cyclohexenecarboxylic acid isomers can be fully aromatized and ex-creted as benzoic acid and interestingly, only the 1-cyclohexencarb-oxylic acid isomer has been shown to be a downstream product ofCHCA via aromatization (Beer et al., 1951; Babior and Bloch, 1966).While CHAA degradation in rats is believed to occur via hydroxyl-ation and conjugation followed by excretion in the urine (Tulliezet al., 1981), neither hydroxylation nor conjugation products weredetectable in our extracts. In the case of D. tertiolecta, it appears thatbiotransformation of CHAA proceeds via dehydrogenation formingcyclohexylidene acetic acid and 1-cyclohexeneacetic acid whichmay be followed by capro-lactone formation or alpha-oxidation,respectively. However, no metabolites have yet been identified toconfirm that these latter compounds are being metabolized viaknown pathways.

3.3. Cyclohexanepropionic acid and cyclohexanecarboxylic acidbiodegradation

Biodegradation of CHPA proceeded in a manner very similar tothe degradation of CHBA described previously, although completetransformation of the compound and metabolites occurred within21 d rather than 35 d (Fig. 4A). After 7 d, we observed the emer-gence of CHCA, a metabolite with two fewer carbons, which againsuggested that b-oxidation was occurring with this model NA

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(Fig. 4A). The presence of CHCA was confirmed by comparison withan authentic standard (data not shown) and its presence was tran-sient. However, complete transformation of CHPA and CHCA oc-curred faster than CHAA which took more than 42 d as opposedto 21 and 35 d for CHPA and CHCA, respectively. These data qual-itatively suggest that both compounds are more susceptible to al-gal biodegradation than CHAA (Fig 3A, 4A-B). One explanation forthe accelerated degradation of CHCA is that the b carbon is not ter-tiary, therefore allowing step-wise b-oxidation to proceed. Thiswould allow ring opening and degradation of the resulting fattyacid-like compound by a simple respiratory process (Whitby,2010). This supports the idea that CHCA is less recalcitrant thanCHAA (Iwaki et al., 2008), although this algal species is capableof metabolizing both model NAs.

In addition, mass spectral analysis identified an emergentmetabolite from a 14 d algal culture initially containing 200 mg L�1

of CHCA (Fig. 4C). This metabolite was confirmed as 1-cyclohexen-ecarboxylic acid (Fig. 4C) by comparison with an authentic stan-dard (data not shown). Unlike the metabolism of CHAA by D.tertiolecta, only a single isomer was identified and this particularstructure likely represents the initial dehydrogenation step in-volved in the b-oxidation reaction (Poirier et al., 2006) or via thearomatization reaction described by Babior and Bloch (1966). Fur-ther, the specificity of the double bond formation at the b positionlikely rules out aromatization pathways seen in certain bacteria

A

C

Fig. 4. Decrease of CHPA levels and appearance of the secondary metabolite CHCA inresolution GC–MS (n = 3) (A). Cultures spiked with 200 mg L�1 CHCA also showed a15.352 min (C) from a 14 d old culture initially containing 200 mg L�1 CHCA yields a masas 1-cyclohexenecarboxylic acid (spectra is of the trimethylsilyl derivative).

(Iwaki et al., 2005). While we were unable to distinguish betweenthe two putative pathways, CHCA was readily biotransformed viathe 1-cyclohexenecarboxylic acid intermediate observed.

3.4. Biodegradation of 1,2,3,4-tetrahydro-2-naphthoic acid

While model NAs with a single-ring such as CHCA, CHAA, CHPAand CHBA reveal metabolic mechanisms employed by D. tertiolecta,they do not accurately represent NAs encountered in tailings gen-erated from bitumen extraction (Scott et al., 2005). NAs present inoil sands tailings ponds are comprised of a complex mixture ofcompounds that have a wide variety of ring structures. The mostabundant include compounds with a Z designation of �4 to �6(2–3 ring structures) (Clemente and Fedorak, 2005). Algal culturesincubated with the double-ringed, model NA 1,2,3,4-tetrahydro-2-naphthoic acid were monitored by GC–MS to investigate morecomplex NA degradation. In contrast to single-ring model NAs,algal incubation in the presence of 1,2,3,4-tetrahydro-2-naphthoicacid did not reveal degradation or bioconversion after a period of35 d suggesting that the Z family plays a significant role in therecalcitrant nature of NAs (Fig. SM-4). Both 1,2,3,4-tetrahydro-2-naphthoic acid and CHBA have similar molecular weights(176.22 g mol�1 and 170.25 g mol�1, respectively), but CHBA waseasily removed from algal cultures indicating NA persistence ismore influenced by ring composition than molecular weight.

B

cultures of D. tertiolecta initially containing 200 mg L�1 CHPA as measured by lowdecrease in compound levels after incubation (B). The mass spectra obtained ats ion of m/z = 198 suggesting dehydrogenation of CHCA with the structure identified

510 D.M. Quesnel et al. / Chemosphere 84 (2011) 504–511

Similar observations were made in studies investigating bacterialNA biodegradation (Han et al., 2008).

3.5. Compositional changes of tailings associated NAs

We next wanted to determine whether D. tertiolecta was able todegrade NAs from an active tailings pond still receiving effluentfrom the bitumen extraction process. One of the major hurdlesencountered when investigating NA biodegradation in oil sandstailings ponds is the multitude and structural complexity of thesecompounds. As a mixture of compounds with similar chemicalproperties, they are difficult to analyze singly but can viewed atas a group based on molecular weight using the method describedby Holowenko et al. (2002). While not completely accurate indetecting all individual NAs, the use of low resolution GC–MS al-lows for a fingerprint analysis of different families of NAs. By iden-tifying ions fitting the classically-defined formula CnH2n+zO2 fromthe total ion chromatogram (TIC), compositional changes of tail-ings NAs can be monitored. Cultures containing a mixture of 60%tailings and 40% f/2 medium in the presence and absence of D.tertiolecta were incubated for 6 weeks and the distribution of thetailings NAs in each sample was analyzed and compared after thistime (Fig. 5A and B). We found that this ratio of tailings to algalmedium was optimal in supporting growth of D. tertiolecta (datanot shown). Between the two samples, the most noticeable changewas the loss of 11–17 carbon compounds of the Z family = �2.These compounds are similar to the model NAs described earlieras they have a single-ring but have higher molecular masses. Asthere is no structural data about each compound fitting the for-mula CnH2n�2O2 where n = 11–17, the Z family is the most compel-ling property regarding biodegradation susceptibility. The other

A

B

Fig. 5. Distribution of NA compounds fitting the molecular formula CnH2n+zO2 from C5

inoculation with algae (A). The sum of all bars equals 100% as data was normalized to thchange includes compounds of the Z family �2 (single-ring) which were almost comple

key observation is the persistence of compounds with 10–15carbons and a Z designation of �4 to �6 (Fig 5A and B). We referto these compounds as the ‘‘recalcitrant fraction’’. This fractionbecomes enriched after algal incubation suggesting limited or nobiodegradation. Compounds with 15, 17 and 18 carbons with a Zdesignation of zero also appear to persist. However, this resultseems unlikely as these compounds fit the definition of a saturatedfatty acid which should readily break down via b-oxidation. It hasbeen shown previously that low resolution MS analysis artificiallyinflates Z = 0 NAs through the identification of false positives, thus,these compounds may be artefacts of the analysis process (Martinet al., 2008). It is also possible that these peaks indicatehighly-branched acyclic fatty acids which are more resistant tob-oxidation (Holowenko et al., 2002). There were also some unex-pected changes to compounds from the family Z = �8 (see SM-Ta-bles 1 and 2 for detailed information). With the exception of n = 24,all NAs from this family appeared to decrease. If increasing ringnumber correlates to decreased degradation susceptibility, wewould expect these compounds to persist after algal incubation.Compounds like benzoic acid also fit the molecular formula ofthe Z = �8 family of NAs, and thus these compounds may havebeen incorrectly identified in our assay (Grewer et al., 2010).Although the analysis method used suffers from some drawbacks,it was effective for identifying general trends in natural NA biodeg-radation by D. tertiolecta.

4. Conclusions

As reviewed in Quagraine et al. (2005), certain criteria must bemet for bioremediation of NAs to be considered as a feasible

to C33 in a culture mixture containing 60% tailings and 40% f/2 medium prior toe total abundance of selected ions. After 6 weeks of algal growth, the most notabletely removed (B). Tailings were obtained from Suncor Energy Inc. (Pond 6).

D.M. Quesnel et al. / Chemosphere 84 (2011) 504–511 511

solution to oil sands tailings reclamation issues, such as demon-strating biodegradation potential. Since the ability of photosyn-thetic algae to degrade NAs is rarely studied and poorlyunderstood, the first step undertaken in this investigation was toidentify species that exhibited potential NA degradation capabili-ties. Based on the premise that increased NA tolerance indicatesa means by which the algae are capable of detoxifying their sur-roundings, D. tertiolecta was identified and shown to be capableof degrading select model NAs. From our analyses of model NAsand their biodegradation, we identified transient metabolites andestablished putative mechanisms that initiate the process of NA re-moval. Although commercially-available NAs are more biodegrad-able than those found in oil sands tailings water (Scott et al., 2005),elucidating the mechanisms by which simple NAs are metabolizedgives some insight into how certain organisms can deal with thesetoxic compounds. Trials involving tailings NAs did reveal limitedbiodegradation after 6 weeks by D. tertiolecta, results of which clo-sely paralleled trends observed with model NAs. However, it wasincapable of degrading the recalcitrant fraction. Dealing with therecalcitrant fraction of NAs is the major limitation to any bioreme-diation effort as they have been shown to be the most difficult toattenuate both in situ and in vitro (Quagraine et al., 2005; Scottet al., 2005; Whitby, 2010). Overcoming this hurdle would validatethe potential of algae as a NA bioremediation agent. Further studiesusing algae indigenous to the Athabasca Oil Sands tailings pondsmay hold the potential to overcome the obstacles encounteredby D. tertiolecta as environmental pressures may select for speciesspecifically adapted to deal with complex tailings NAs and is thefocus of ongoing work.

Acknowledgements

This work was supported by the Canada School of Energy andEnvironment (Proof-of-Principle Grant to G.C.) and start-up fundsfrom the Faculty of Science, University of Calgary to G.C. andL.M.G. We thank Kate Chatfield–Reed for developing scripts forGC–MS data analysis of tailings NAs. Finally, we gratefullyacknowledge Suncor Energy Inc. for their in-kind contributions oftailings water samples.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at doi:10.1016/j.chemosphere.2011.03.012.

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