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Environ. Sci. Technol. 2010, 44, 8350–8356

Ozonation of Oil Sands Process-Affected Water Accelerates Microbial Bioremediation J O N A T H A N W . M A R T I N , * ,† THAER BARRI,† XIUMEI HAN,† PHILLIP M. FEDORAK,‡ MOHAMED GAMAL EL-DIN,§ LEONIDAS PEREZ,† ANGELA C. SCOTT,| AND JASON TIANGE JIANG† Division of Analytical & Environmental Toxicology, Department of Biological Sciences, Department of Civil & Environmental Engineering, and Department of Agricultural, Food & Nutritional Science, University of Alberta, Edmonton, Alberta, Canada

Received May 7, 2010. Revised manuscript received September 9, 2010. Accepted September 20, 2010.

Ozonation can degrade toxic naphthenic acids (NAs) in oil sands process-affected water (OSPW), but even after extensive treatment a residual NA fraction remains. Here we hypothesized that mild ozonation would selectively oxidize the most biopersistent NA fraction, thereby accelerating subsequent NA biodegradation and toxicity removal by indigenous microbes. OSPW was ozonated to achieve approximately 50% and 75% NA degradation, and the major ozonation byproducts included oxidized NAs (i.e., hydroxy- or keto-NAs). However, oxidized NAs are already present in untreated OSPW and were shown to be formed during the microbial biodegradation of NAs. Ozonation alone did not affect OSPW toxicity, based on Microtox; however, there was a significant acceleration of toxicity removal in ozonated OSPW following inoculation with native microbes. Furthermore, all residual NAs biodegraded significantly faster in ozonated OSPW. The opposite trend was found for ozonated commercial NAs, which are known to contain no significant biopersistent fraction. Thus, we suggest that ozonation preferentially degraded the most biopersistent OSPW NA fraction, and that ozonation is complementary to the biodegradation capacity of microbial populations in OSPW. The toxicity of ozonated OSPW to higher organisms needs to be assessed, but there is promise that this technique could be applied to accelerate the bioremediation of large volumes of OSPW in Northern Alberta, Canada.

Introduction One of the world’s largest deposits of petroleum is found in the oil sands in Northern Alberta, Canada. Many petroleum companies are investing in large operations to harvest this heavy oil, known as bitumen. The surface mining oil sands industry extracts bitumen from the oil sands using a caustic hot water extraction method (1, 2). The resulting large * Corresponding author phone: 780-492-1190; fax: 780-492-7800; e-mail: [email protected]. † Division of Analytical & Environmental Toxicology. ‡ Department of Biological Sciences. § Department of Civil & Environmental Engineering. | Department of Agricultural, Food & Nutritional Science. 8350

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volumes of oil sands process-affected water (OSPW) are recycled into the extraction process, but they are toxic and must be retained on site due to a zero-discharge policy (1, 3). Remediation of OSPW is difficult because it has both acute and chronic aquatic toxicity which is mainly attributable to the persistent class of organic acids known as naphthenic acids (NAs) (4-7). NAs are a broad group of alicyclic, or noncyclic, alkyl-substituted carboxylic acids having the general formula CnH2n+ZO2; where n is the carbon number and Z is zero or a negative even integer related to the hydrogen deficiency caused by the presence of rings or double bond equivalents (e.g., Z ) 0, no rings; Z ) -2, 1 ring etc....) (4). Substantial research has been conducted in an effort to remediate OSPW (8). However, to date there are no proven strategies that can fully eliminate OSPW NAs, or the toxicity of OSPW. Remediation of OSPW by in situ methods (i.e., relying only on natural microbial biodegradation) can slowly diminish NA concentrations, but a biopersistent NA fraction remains (8, 9) and acute toxicity to some aquatic organisms (e.g., larvae of Chironomus dilutus) at the outfall of constructed wetlands also remains (10). The in situ degradation half-lives for OSPW NAs were recently estimated to be 13 years in the field (11). Furthermore, very long hydraulic retention times are required for in situ bioremediation (8, 10), and given the large existing stores of OSPW, and ongoing expansion of the oil sands industry, we suggest that a more rapid approach for OSPW remediation is necessary. Recently, ozone treatment showed promise for rapidly degrading NAs and diminishing microbial toxicity, but even after extensive ozone treatment time a fraction of NAs remained, and total organic carbon was unaffected (12). Here we explore the hypothesis that ozonation of OSPW selectively degrades the most biopersistent NA fraction (i.e., the fraction with more rings and alkyl branching 9, 13) such that residual NAs can be more rapidly biodegraded. This structure-reactivity hypothesis was based on knowledge that ozone is unstable at the natural pH of OSPW (i.e., pH 8-9) and decomposes to produce reactive hydroxyl radicals (14) that can abstract hydrogen more readily from tertiary carbon (i.e., at a branching point) than either primary or secondary carbon atoms (15). We test this hypothesis by incubating untreated and ozone-degraded OSPW, and untreated and ozone-degraded commercial NAs, with the native microbial consortium from an active settling basin. Biodegradation kinetics, microbial toxicity, and oxidation byproducts were examined.

Experimental Section Chemicals and Reagents. Tetradecanoic acid-1-13C (C14H28O2; Z ) 0) was used as internal standard and was purchased from Sigma-Aldrich (Oakville, ON, Canada). Highpurity Merichem NAs were provided by Merichem Chemicals and Refinery Services LLC (Houston, TX). OSPW was collected on the site of Syncrude Canada Ltd. (Fort McMurray, AB, Canada) from the West-in-Pit active settling basin in December 2007 and stored at 4 °C. Ozone demand-free water was prepared by bubbling ozone through distilled water for half an hour and leaving it in a fume hood for 48 h before use. Ozonation of OSPW and Commercial NAs. An AGSO 30 Effizon ozone generator (WEDECO AG Water Technology, Herford, Germany) was used to produce ozone gas from extra dry, high purity oxygen. All glassware was cleaned and pretreated with ozone-demand-free water before experimentation. Ozonation reactions were performed at room temperature (approximately 20 °C) in open glass bottles on 10.1021/es101556z

 2010 American Chemical Society

Published on Web 10/07/2010

a stir plate. The intent of the ozone treatment was to reach various levels of NA degradation, thus ozone doses were not quantified. For preparation of commercial NAs, a 1000 mg/L stock solution of Merichem NAs was prepared in 0.1 M NaOH and adjusted to pH 10 with HCl. All solutions were prepared with ozone-demand-free water before addition of NAs. Three 1 L bottles containing 940 mL of 30 mM Na2HPO4 (pH 8.0) were prepared, and two of these were then bubbled with ozone for long enough to obtain 10 mg ozone/L (5 min with buffer at 20 °C) and 20 mg ozone/L (5 min with buffer at 5 °C), determined by the indigo colorimetric method (16). The third bottle was the control and did not receive any ozone. To all bottles, 60 mL of stock Merichem NA solution was then immediately added to obtain initial concentrations of 60 mg NAs/L. All treatments were left overnight and subsequent ultra pressure liquid chromatography/high resolution mass spectrometry (UPLC-HRMS) analysis indicated that the NAs in the ozonated bottles had degraded by 51% (10 mg ozone/L) and 72% (20 mg ozone/L), compared to the unozonated control. The unozonated control had a pH of 8.0, and the treated samples were not significantly different after ozonation. For preparation of OSPW, three 1 L batches of OSPW were filtered through a 0.45 µm cellulose membrane filter (Osmonics, Inc., Minnetonka, MN) to remove suspended solids, primarily clay and residual bitumen. One batch was left unozonated, for use as a control, and the two other batches were each placed in 2 L glass bottles for treatment by ozonation. Ozone was bubbled through the OSPW at a gas flow rate of 2 L/min (106 g/m3 ozone) and was stopped occasionally to take samples for UPLC-HRMS analysis. The ozonation treatment was continued until the total degradation of parent NAs in these two batches reached 54% (60 s treatment time) and 73% (90 s treatment time). The extent of degradation was purposely controlled to closely match the extent of degradation in the ozonated Merichem NAs. The pH of the untreated OSPW was 8.1, and the ozonation did not change this significantly. Microbial Biodegradation Incubations. Aerobic NA microbial biodegradation experiments were conducted using previously established protocols (17). Briefly, triplicate viable and sterile (heat-killed) controls for each treatment (unozonated and ozonated OSPW, and unozonated and ozonated Merichem NAs) were maintained at room temperature (approximately 20 °C) on a shaker at 200 rpm beginning in May 2008. Microorganisms indigenous to the OSPW, concentrated by centrifugation, were added to all treatments to result in microbial densities comparable to natural OSPW, as previously described (9). Final liquid incubation volumes were 250 mL and each culture was incubated in a screwcapped 500 mL Erlenmeyer flask to prevent evaporation of water during the extended incubation times. Calculations showed that oxygen in the headspace of these sealed flasks was adequate for complete mineralization of the organic material in each flask, but every 2 or 3 days each cap was loosened to allow air exchange to replenish oxygen. Triplicate 2 mL samples were collected from each treatment for microbial toxicity analysis on days 0, 6, 14, and 28 from Merichem NA incubations, and on days 0, 14, 28, 56, and 90 for OSPW incubations. These were frozen until time of the bioassay. An additional 3 mL of each incubation was sampled on days 0, 7, 14, 21, 28, 41, 56, 70, 84, and 98, for all OSPW treatments, and on days 0, 2, 6, 8, 10, 12, 14, 21, and 28 for all Merichem NA treatments. Due to the large number of samples requiring quantitative NA analysis (n ) 342 individual samples), the triplicate 3 mL samples for each treatment, for each sampling day, were pooled for analysis (n ) 114 pseudo average samples). The pooled samples (9 mL) were centrifuged, the supernatant was collected, and an

equivolume of methanol was added for preservation. Samples were then frozen until analysis by UPLC-HRMS. Microbial Toxicity Assay. Microtox toxicity bioassays were used to measure changes in toxicity using the basic protocol (18). Samples were thawed rapidly at 28 °C, vortexed for 10 s, centrifuged to remove suspended particles and microbes, and 2.5 mL of each supernatant was removed for the assay. The sample pH was not adjusted because ozonation did not alter the pH, and all samples were well buffered. A Microbics model M500 was used to measure the luminescence of reconstituted Vibrio fischeri before and after 15 min exposure to samples of OSPW or Merichem NAs. The percent (v/v) of sample that caused a 20% inhibition of luminescence was determined and reported as the IC20. As quality control, phenol toxicity was tested prior to analysis of real samples, and the measured toxicity was always within the recommended range (18). Analysis of NAs by UPLC-HRMS. A 1 mL portion of each sample was filtered through a 0.45 µm nylon syringe filter, and 10 µL of 10 µg mL-1 internal standard (tetradecanoic acid-1-13C) in methanol was added to 490 µL of each filtered sample. A Waters ACQUITY UPLC System (Waters, MA) was used for chromatographic separation of the NAs and their oxidized products. Chromatographic separations were run on a Waters UPLC Phenyl BEH column (150 × 1 mm, 1.7 µm) using a gradient mobile phase of (A) 10 mM ammonium acetate, and (B) 10 mM ammonium acetate in 40% acetone and 60% methanol. Gradient elution was as follows: 1% B for the first 2 min, ramped to 60% B by 3 min, ramped to 70% B by 7 min, ramped to 95% B by 13 min, held until 14 min, and finally a return to 1% B followed by a further 5.8 min equilibration time. The flow was constant at 0.11 mL min-1 and column temperature was 50 °C. Detection was performed with a high resolution (∼10,000 m/∆m) QSTAR Pulsar i mass spectrometer equipped with a TurboIon Spray source (Applied Biosystems/MDS Sciex, Concord, ON, Canada) operating in negative ion mode. Analyst QS 1.1 and Multiquant 1.1 software (Applied Biosystems, Foster City, CA) were used for data analysis, and the relative ratio of each analyte’s chromatographic peak (i.e., for each isomer class corresponding to each n and Z combination) area to the internal standard was calculated for subsequent kinetic analysis. Total NA degradation was estimated by the decrease in the sum response of all the UPLC-HRMS peak areas with exact masses corresponding to NAs (CnH2n+ZO2). NA Biodegradation Kinetics. Relative response (to internal standard) was plotted over time for the pseudo average NA concentration data, and a function was fit to determine the biodegradation kinetics for each NA, or NA+O, isomer class (i.e., for each n and Z combination). It is germane to note that OSPW is a complex mixture, and in the current work we only monitored for NAs and their oxidized products (i.e., NA+O) because they are known to be a major fraction. We previously demonstrated that biodegradation kinetics of untreated Merichem NAs were sigmoidal under these conditions, whereas untreated OSPW NAs degraded by first-order kinetics (9). Therefore, for Merichem NAs, SigmaPlot 2004 (Version 11, Systat Software, San Jose, CA) was used to fit a 5 parameter sigmoidal function and to calculate the time to 50% degradation (9), whereas for OSPW NAs linear regression of natural-log transformed data was performed and the halflife was calculated. Note that there are no authentic standards for chemicals in the complex NA mixtures, thus all kinetic concentration data was expressed relative to time zero, and the internal standard response.

Results and Discussion Microbial Toxicity. Compared to the unozonated control, ozonated Merichem NAs had significantly increased IC20 VOL. 44, NO. 21, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 1. Detoxification of ozonated and unozonated control (A) Merichem NAs and (B) OSPW following addition of viable microorganisms. Data are shown as mean increase in the IC20 ((1 standard deviation) as a percentage of full-strength, relative to time-matched sterile controls. (i.e., toxicity decreased) (Supporting Information (SI), Figure S1). Toxicity was lowest in the 72% ozone-degraded sample, but the IC20 of the 53 and 72% sample were not significantly different. In contrast, ozonation of OSPW did not significantly affect the IC20, even though the extent of parent NA degradation was similar to the Merichem solutions (SI Figure S1). Scott et al. (12) reported decreasing toxicity of ozonated OSPW, also using Microtox, but in that study the first sampling time was after 10 min of continuous ozone bubbling in OSPW, whereas here we used batch ozonation for a maximum of only 1.5 min. Thus the current results are not inconsistent with the literature (12), but a simple comparison of the two

studies is difficult because different apparatus were used and larger volumes were treated for much longer ozonation times in the previous work of Scott et al. After inoculation and incubation with the OSPW microbial community, toxicity decreased in all OSPW and Merichem treatments over time, but the rates differed by treatment (Figure 1). For all ozonated Merichem NAs, after 6 and 14 days of incubation, the toxicity was significantly reduced compared to the unozonated controls (Figure 1A). After 28 days, the 51% ozone-degraded sample continued to detoxify and was still much less toxic than the unozonated sample,

FIGURE 2. NA (CnH2n+ZO2) first order biodegradation half-lives (days (1 regression standard error) plotted by number of carbon (n) in unozonated control OSPW, 54% ozone-degraded NAs, and 73% ozone-degraded NAs grouped by Z number: (A) Z ) -2, (B) Z ) -4, (C) Z ) -6, (D) Z ) -8. In some instances, the 73% ozone-degraded OSPW samples contained too little starting material to accurately calculate a biodegradation half-life. 8352

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FIGURE 3. Oxidized NAs (NA+O; CnH2n+ZO3) in Merichem NAs before (A) and after 28 days of biodegradation (B), and in 51% ozone-degraded Merichem NAs before (C) and after 28 days of biodegradation (D). but curiously the 73% ozone-degraded Merichem sample was more toxic than the unozonated sample after 28 days. For OSPW, although mild ozonation itself did not affect microbial toxicity, increasing ozone treatment had clear benefits with regards to subsequent bioremediation by OSPW microorganisms (Figure 1B). Unozonated OSPW toxicity showed a long lag time of over 56 days before any significant decrease in toxicity was evident. In contrast, the 73% ozonedegraded sample had significantly detoxified after only 14 days, while the 54% ozone-degraded sample had significantly detoxified after only 28 days. Furthermore, the 73% ozonedegraded sample was always less toxic than the 54% ozonedegraded sample; albeit only significantly at days 14 and 28. As explained in the following section, the accelerated microbial toxicity removal shown here for ozonated OSPW was commensurate with an accelerated biodegradation of NAs in the same samples. Residual NA Biodegradation Kinetics. Han et al. (9) previously reported the biodegradation kinetics of OSPW and Merichem NAs incubated with a microbial consortia endogenous to OSPW tailings ponds. In general, OSPW NAs were much more persistent than Merichem NAs, possibly because of increased alkyl branching in OSPW NAs (9, 19). Here we used the same experimental approach but with an emphasis on comparing the relative biodegradability of unozonated NAs and residual ozonated NAs. As explained earlier, we hypothesized that ozonation should preferentially oxidize the NAs with more alkyl branching, thereby leaving a residual NA fraction with less branching which should be more susceptible to biodegradation. In ozonated OSPW, the average half-life of all residual NA classes were decreased (i.e., biodegradability increased) compared to untreated OSPW (Figure 2A-D). The extent of this effect depended on Z-series, and on the extent of

ozonation. For Z ) -2 NAs, average time to 50% biodegradation was 2.5-fold (1.7-3.3-fold) and 3.3-fold (2.6-3.9fold) faster for 54% ozone-degraded and 73% ozone-degraded treatments, respectively, compared to unozonated OSPW. For Z ) -4 NAs, average time to 50% biodegradation was 1.5-fold (1.4-1.7-fold) and 1.6-fold (1.4-1.8-fold) faster for 54% ozone-degraded and 73% ozone-degraded treatments, respectively, compared to unozonated OSPW. For Z ) -6 NAs, average time to 50% biodegradation was 1.4-fold (0.9-1.6-fold) and 1.5-fold (1.0-1.8-fold) faster for 54% ozone-degraded and 73% ozone-degraded treatments, respectively, compared to unozonated OSPW. For Z ) -8 NAs, average time to 50% biodegradation was 1.9-fold (1.2-2.4fold) and 2.5-fold (1.3-3.5-fold) faster for 54% ozonedegraded and 73% ozone-degraded treatments, respectively, compared to unozonated OSPW. Although the biodegradation kinetics of a few NAs in the Z ) -10 and Z ) -12 series could be determined in unozonated control samples, these NAs were not detectable in enough of the ozonated and biodegraded samples to allow for a comparison. Nonetheless, the Z ) -2, -4, -6, and -8 series of NAs are the most prominent NAs in OSPW. On the whole, total NA biodegradation half-life (i.e., taking the sum of all individual NAs) decreased from 83 days in unozonated OSPW, to 55 days in 54% ozone-degraded OSPW, and to 48 days in 73% ozonedegraded OSPW. The reason for improved biodegradability of NAs in OSPW following ozonation could be, as hypothesized, that the most persistent fraction of each NA homologue class was preferentially degraded by the ozone treatment (i.e., presumably the most alkyl branched fraction). The best available analytical methods for OSPW today do not allow the highly branched isomers to be separated from the more linear ones, thus this ozone structure reactivity hypothesis is currently being VOL. 44, NO. 21, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 4. Oxidized NAs (NA+O; CnH2n+ZO3) in OSPW before (A) and after 98 days of biodegradation (B), and in 54% ozone-degrade OSPW before (C) and after 98 days of biodegradation (D). investigated with model compounds in our lab. Nonetheless, an alternative explanation is that ozonation of OSPW enables the microbial population to grow more rapidly than in unozonated control OSPW due to increased biological oxygen demand, and the formation of more labile intermediate products (e.g., hydroxylated NAs or short chain acids). We did not monitor the biomass in the current study, but if the latter explanation was a significant factor, then it is difficult to explain why residual Merichem NAs did not also biodegrade more quickly following ozonation. In fact, the converse was generally true for Merichem NAs. For example, the time to 50% biodegradation was always longer in ozonated Merichem incubations, apart from a few exceptions in the Z ) -2 series (SI Table S1). Furthermore, time to 50% biodegradation for the 73% ozone-degraded Merichem treatments was generally longer than for the 51% ozonedegraded treatment, again opposite to what occurred in ozone-degraded OSPW. Unlike OSPW, which is composed of highly recalcitrant NAs (9, 13), Merichem NAs do not contain a substantial biopersistent fraction (i.e., much less highly branched fractions) (9), thus ozone or OH radical structure reactivity cannot be expected to improve the biodegradability of any residual NAs. The opposite phenomenon, whereby ozonated Merichem NAs biodegraded more slowly, may have occurred because NA degradation products were a preferred carbon source relative to the residual NAs. Formation and Fate of Oxidized NA Intermediates. The ozonation conditions used here were not anticipated to fully mineralize parent NAs, rather it was anticipated that various oxidized intermediates would be formed. We therefore targeted the appearance of mono-oxidized NAs (NA+O; CnH2n+ZO3) in ozonated samples of Merichem and OSPW, and we followed their subsequent fate under the microbial incubation conditions. To our knowledge, this is the first 8354

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time that profiles (i.e., by n and Z) or the biodegradability of these important compounds has been reported. Unozonated Merichem NAs contained only minor traces of NA+O (Figure 3A), with the profile showing greatest abundance in the C15-C20 range of Z ) -6 and Z ) -8 NAs. As we reported previously, oxidized NAs are intermediates in the biodegradation of NAs (9) and in the current Merichem NA study the response of these oxidized NAs increased 2.4fold over 28 days of biodegradation, largely in the C9-C16 and Z ) 0 to Z ) -8 range (Figure 3B). The shape and position of this profile for oxidized NAs (by n and Z), arising from microbial biodegradation, corresponded well with the profile for parent NAs in untreated Merichem (e.g., see profile published previously (20)), suggesting that the mode of microbial biodegradation includes simple insertion of oxygen. In the current work, ozonation of Merichem also produced abundant oxidized NAs. For example, in the 51% ozonedegraded sample, the abundance of oxidized NAs increased by 7.7-fold (Figure 3C). Over the subsequent 28 days of biodegradation, the oxidized NAs in the 51% ozone-degraded sample decreased by 46.7% (Figure 3D). Interestingly, the profile of oxidized NAs in the 51% ozone-degraded sample, before and after biodegradation (Figure 3C and 3D), was similar to the profile of oxidized NAs produced by simple microbial biodegradation of Merichem NAs (Figure 3B). The biodegradation kinetics of oxidized NAs produced by ozonation of Merichem NAs are difficult to calculate because, as just described, they are being both formed and degraded by the microbes during the course of the incubation. Nonetheless, pseudo degradation half-lives were calculated for each isomer class (i.e., for each combination of n and Z) of oxidized NAs, and half-lives ranged from 2 to 50 days overall. However, no clear pattern or structure-persistence relationship was evident. Nonetheless, overall in the ozonated Merichem samples the oxidized NAs had a net rate of

disappearance over the 28 day period, thus they too can be biodegraded by the indigenous microbial consortium of OSPW. As expected, OSPW contained a substantial fraction of oxidized NAs before ozonation (Figure 4A). We reported previously (11) that OSPW naturally contains a large proportion of oxidized NAs, and furthermore that the oldest OSPW on the industrial sites contained a higher relative fraction of these, relative to parent NAs. In the current work, the total mass spectrometer response (arbitrary units) for oxidized NAs in untreated OSPW was 59% of the response of parent NAs, thus the fate of these molecules is an important question under any OSPW reclamation strategy. For unozonated OSPW, over the subsequent 98 days of biodegradation the total response of oxidized NAs decreased by 30.0%, and the relative profile changed noticeably (Figure 4B). This was largely because Z ) -6 and Z ) -8 classes of oxidized NAs generally had no discernible decrease over the 98 day incubation with microbes, whereas the biodegradation half-lives of Z ) 0, -2, and -4 oxidized NAs were measurable, generally ranging from 20 to 100 days. There was otherwise no consistent structure-persistence relationship by Z series, but within the Z ) -2, -4, and -10 series, the shorter oxidized NAs were generally the most resistant to biodegradation (compare Figure 4A and B). Ozonation of OSPW had the overall effect of diminishing the total response of oxidized NAs in OSPW, presumably because their rates of degradation exceeded their rates of formation. For example, the total NA+O response diminished by 6.9% (Figure 4C) and 27% (compare SI Figure S2A and S2B) in the 54% and 73% ozone-degraded OSPW samples, respectively. In general, this was due to a combination of diminished response for Z ) 0, -2, -4, and -6 NA+O, and an increased response for Z ) -8, -10, and -12 NA+O, suggestive that parent NAs containing more rings formed more of the NA+O byproducts; assuming that oxygen is added to parent NAs without breaking or forming rings. Over the subsequent 98 days of biodegradation, the total response of oxidized NAs in 54% and 73% ozone-degraded OSPW decreased further by 34.7% (Figure 4D) and 30.4% (compare SI Figure S2B and S2C), respectively, and the relative profiles changed noticeably. As can be seen by comparison of Figures 4C and 4D, for 54% ozone-degraded NAs, the response of Z ) -6 oxidized NAs remained largely unaffected, while all oxidized NAs with fewer rings were significantly diminished. Furthermore, oxidized NAs in the C17-C21 range of Z ) -8, -10, and -12 were preferentially diminished, and many were not detectable by day 98. Interestingly, the NA+O profiles of unozonated and ozonated OSPW were very similar after biodegradation (i.e., compare Figure 4B an D). A similar trend was observed in the 74% ozone-degraded OSPW samples for oxidized NAs (compare Figure 4B and SI Figure S2C). Environmental Implications and Future Work. This study used OSPW from an active settling basin, and demonstrated that relatively light ozonation had the dual benefit of diminishing microbial toxicity while increasing the biodegradability of residual NAs by native microbial consortia. Furthermore, these benefits were more pronounced with higher extents of ozone degradation, and it is likely that the extent of NA degradation achieved here can be improved by optimizing ozonation conditions. Taken together with a previous report, which showed that much longer ozonation times can lead to NA degradation and Microtox toxicity reductions (12) without a subsequent bioremediation period, the potential for using this established oxidation system for OSPW remediation seems high. This study was a worst-case scenario, whereby the most toxic form of OSPW (i.e., OSPW fresh from an active settling basin) was treated by relatively light ozonation. Real world application may include treating the large stores of toxic OSPW with ozone at the inflow or

the outflow of constructed wetlands or end-pit-lakes containing partially aged OSPW. Nonetheless, it is important to keep in mind that all toxicity data reported here for ozonated OSPW was collected with a very simple, and perhaps environmentally irrelevant, microbial toxicity assay. Although the Microtox method is rapid, relatively inexpensive, and has been commonly applied in many studies to monitor the toxicity of OSPW and other NA mixtures (7), it cannot necessarily be used to predict the toxicity of ozonated OSPW to other relevant organisms. Such tests with relevant cell lines, aquatic invertebrates, fish, and mammals are ongoing in the laboratories of our collaborators (21), and are necessary before any real-world applications and release of ozonated OSPW to receiving environments can be justified. Furthermore, further advanced characterization of OSPW ozonation byproduct will complement toxicity tests in higher organisms. Only one class of intermediate oxidation byproduct was targeted here (i.e., the mono-oxidized NAs), but other important byproduct, including oxidized metals and inorganics, may be formed that should be identified before ozonated OSPW can be released to natural environments.

Acknowledgments This research was supported by an NSERC Strategic Grant (J.W.M., M.G. E.-D., and P.M.F.) and an Alberta Water Research Institute Grant (J.W.M., M.G.E.-D.). Syncrude Canada Ltd (Edmonton, AB) was a partner in both grants, and Michael MacKinnon and Warren Zubot are thanked for their keen cooperation, and provision of OSPW samples. J.W.M. further acknowledges Alberta Health and Wellness for daily support of laboratory activities. Debbi Coy (Department Biological Sciences, University of Alberta) is thanked for microcosm setup and sampling. J.W.M. acknowledges equal contribution to the experimental work by X.H. and T.B.

Supporting Information Available Biodegradation kinetics of control and ozonated Merichem NAs, and the microbial toxicity of OSPW and Merichem NAs before, and after, ozonation. This material is available free of charge via the Internet at http://pubs.acs.org.

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