15N-nitrate and 34S-sulfate isotopic fractionation reflects electron acceptor ‘recycling’ during hydrocarbon biodegradation

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N-nitrate and 34S-sulfate isotopic fractionation reflects electron acceptor ‘recycling’ during hydrocarbon biodegradation

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Martin Kern1, Andrea Watzinger2 and Kerstin E. Scherr1 1 University of Natural Resources and Life Sciences (BOKU), Department IFA-Tulln, Institute for Environmental Biotechnology, Konrad Lorenz Strasse 20, 3430 Tulln, Austria 2 AIT Austrian Institute of Technology GmbH, Energy Department, Environmental Resources and Technologies, Konrad Lorenz Straße 24, 3430 Tulln, Austria

The analysis of stable carbon isotopes for the assessment of contaminant fate in the aquifer is impeded in the case of petroleum hydrocarbons (TPH) by their chain length. Alternatively, the coupled nitrogen– sulfur–carbon cycles involved into TPH biodegradation under sulfate- and nitrate reducing conditions can be investigated using nitrogen (d15N) and sulfur (d34S) isotopic shifts in terminal electron acceptors (TEA) involved in anaerobic TPH oxidation. Biodegradation of a paraffin-rich crude oil was studied in anaerobic aquifer microcosms with nitrate (NIT), sulfate (SUL), nitrate plus sulfate (MIX) and nitrate under sulfate reduction suppression by molybdate (MOL) as TEA. After 8 months, TPH biodegradation was not different (around 33%) in experiments receiving only nitrate (NIT, MOL) versus under mixed TEA-conditions (MIX), despite higher biodiversity under mixed conditions (H0 NIT and H0 MOL  5.9, H0 MIX = 8.0). Molybdate addition effected higher nitrate depletion, possibly by increasing the production of nitrate reductase. Additional sulfate depletion under mixed conditions suggested bioconversion of polar intermediates. Microcosms only receiving sulfate (SUL) showed no significant TEA and TPH decrease. A Rayleigh kinetic isotope enrichment model for isotopic 15N/14N and 34S/32S shifts in residual TEA gave apparent enrichment factors eN,NIT and eN,MOL values of 16.7 to 18.0% for nitrate as sole TEA and eN,MIX of 6.0% and eS,MIX of 4.1% under mixed electron accepting conditions. The low isotopic fractionation under mixed terminal electron accepting conditions was attributed to lithotrophic, sulfide-dependent denitrification by Thiobacillus species, while it was hypothesized that Desulfovibrio replenished the reduced sulfur pool via oxidation of polar hydrocarbon metabolites. Concurrently, organotrophic denitrification was performed by Pseudomonas species, with isotopic fractionation expressed by eN,MIX representing the superposition of both denitrification processes. This is, to our knowledge, the first characterization of sulfur and nitrogen isotopic shifts associated to concurrent organotrophic and lithotrophic denitrification in a hydrocarbon-contaminated environment, and offers the prospect of improved understanding of biogeochemical cycles including in situ hydrocarbon biotransformation. Introduction The isotopic composition of educts and products of microbially mediated transformation reactions in the environment can be favorably used as an indicator of the occurrence and type of Corresponding author: Scherr, K.E. ([email protected])

transformation pathways in aquatic and terrestrial environments, and stable isotope data can provide unique information for understanding complex biogeochemical processes. Naturally occurring heavy isotopes are less likely to undergo biogeochemical transformation reactions [1] and, to a lesser extent, abiotic reactions [2] than their lighter counterparts. For example, during www.elsevier.com/locate/nbt

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S-sulfate isotopic fractionation reflects electron acceptor ‘recycling’ during hydrocarbon biodegradation, New

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biodegradation of organic contaminants, 12C-bearing molecules are expected to be consumed more rapidly than those bearing 13C, leaving the residual substrate pool with an elevated 13C/12C ratio. This carbon-based compound-specific isotope analysis is widely applied to determine the degradation of low molecular weight, simple hydrocarbons such as chlorinated and monoaromatic hydrocarbons [3,4], and attempts have been made to identify degradation pathways based on isotopic fractionation [5–7]. Carbon-based methods are, however, not applicable for higher molecular weight compounds, such as petroleum hydrocarbons (TPH), including crude oil and its products. Due to their relatively higher molecular weight and variability in chain length and structure, the prediction of their biogeochemical fate from 13 C/12C stable isotope ratios is not sensible. During biodegradation, the carbon isotope effect only occurs at carbon atoms involved in degradation, for example, subterminal or terminal carbon atoms during fumarate addition or carboxylation in the initial steps of anaerobic hydrocarbon activation [8,9]. This ‘dilution effect’ becomes more pronounced with increasing chain length and for molecules with a carbon number exceeding 12– 14, the determined isotopic shifts are not significantly different from analytical errors [10]. Moreover, TPH matrices with a different carbon isotopic composition may mix at a contaminated site, giving false positive or negative biodegradation fingerprints [11]. On the other hand, the analysis of stable oxygen, nitrogen and sulfur isotopes have contributed to the elucidation of nutrient and carbon cycles in aquatic environments [12,13]. Organotrophic denitrification has long been considered the dominant nitrogen eliminating process in most marine and lacustrine environments [14], but less attention has been paid to lithotrophic denitrification. The isotopic enrichment factor for organotrophic denitrification in soil and groundwater is around 25 to 30% [15,16], but considerably less fractionation is suggested in lithotrophic denitrification [17,18]. In the expression of ambient fractionation factors, the type of enzymes mediating denitrification, that is, respiratory or non-respiring nitrate reductases (periplasmatic, NAP; or membrane-bound, NAS) are deemed to dominate over N isotopic fractionation induced by transport processes. Nitrogen isotopic fractionation is expected to be dissimilar for organic and inorganic electron donors, but is not systematically different between different organotrophic denitrifiers [12]. While nitrogen fractionation during denitrification can be associated to the activity of specific enzymes, with NAP reductases effecting a lower nitrogen imprint than NAS [13], considerable differences exist between sulfate-reducing species in terms of sulfur fractionation. In a related study, species-dependent enrichment factors between 2 and 42% were determined, where less pronounced isotopic enrichment was associated with incomplete electron donor oxidation [19]. For these reasons, it may be more useful to predict TPH biodegradation using isotopic imprints during utilization of naturally occurring or anthropogenic terminal electron acceptors (TEA) for biodegradation, such as stable nitrogen (15N/14N), sulfur (34S/32S) and oxygen (18O/16O) isotopes in dissolved nitrate (NO3) and sulfate (SO42). This may enhance our understanding of the intertwined nutrient and carbon cycles and facilitate process monitoring in situ, for example, during monitored or enhanced natural attenuation (MNA, ENA, respectively). Few studies have 2

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investigated the fractionation of anaerobic electron acceptors during hydrocarbon biodegradation [19–22] and the mechanisms governing fractionation are not comprehensively understood. There is evidence that the nature of the electron donor for denitrification – organic or inorganic – is reflected on the 15N isotopic imprint [12,17], but this has not been studied in complex communities of hydrocarbon degraders. We thus hypothesize that the simultaneous occurrence of organotrophic and lithotrophic denitrification, co-occurring with sulfate-dependent hydrocarbon degradation, has a significant impact on both sulfur and nitrogen isotopic fractionation. This hypothesis was developed based on an earlier study, where we collected evidence for the ‘recycling’ of reduced sulfate coupled to denitrification in a hydrocarbon-contaminated aquifer [23]. Using the analysis of nitrogen and sulfur isotopic fractionation processes during cycling associated to hydrocarbon biodegradation, the objective of the present study is to investigate stable isotope fractionation effects associated with single and mixed terminal electron acceptors, reflecting in situ conditions and multiple co-occurring litho- and organotrophic redox-processes. The investigation of fractionation effects reflecting biodegradation of bulk complex hydrocarbon matrices may help in improving our understanding of the coupled nitrogen–sulfur–carbon cycles in contaminated aquifers and practically, facilitate the monitoring of MNA or ENA-treatment approaches based on the addition of TEA.

Materials and methods Chemicals All used chemicals were purchased from Sigma Aldrich (Sigma Aldrich, Vienna, Austria), Merck (Merck GmbH, Vienna, Austria) ¨ ren, or Macherey-Nagel (Macherey-Nagel GmbH & Co. KG, Du Germany) and were of analytical quality.

Aquifer material and crude oil The aquifer material used in the experiments was originally collected from the nitrate- and sulfate-reducing zone of a historically TPH-contaminated site in Lower Austria. It is of sandy texture and was used in previous anaerobic degradation experiments under nitrate- and sulfate-reducing conditions [23]. Thus, microbial consortia of anaerobic TPH-degraders were already present. For this experiment the aquifer material was sieved to <2 mm and carefully homogenized in an anaerobic glove-box (MECAPLEX, Grenchen, Switzerland) flushed with argon 5.0 (Messer GmbH, Gumpoldskirchen, Austria). Due to the depleted nature of the residual hydrocarbon matrix, the aquifer material was spiked with a paraffin-rich crude oil to a target concentration of approximately 18,000 mg/kg dry weight by adding the oil with a Pasteur pipette and careful mixing with a clean metal spatula in the anaerobic glove box. A more detailed description of the crude oil and its degradation characteristics are found in related publications [23–26].

Mineral medium For mineral medium macronutrients, trace elements and vitamins were dissolved in MilliQ1-Water (Merck Millipore, Darmstadt, Germany) with an electrical resistivity of 18.5 MV cm. To yield an approximate stoichiometric C (contaminant + aquifer TOC):N:P:Krelationship of 150:5:1:1 nutrients were added as NH4Cl, KH2PO4

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and Na2HPO4 as described elsewhere [27]. Additionally, 10 mL/L of a trace element solution consisting of (mg/L): H3BO3 (3), CaCl22H2O (703.2), CoCl2 (3.9), CuSO45H2O (17.3), FeSO47H2O (206), KI (101), MnSO41H2O (14), Na2MoO42H2O (5.3), Na2SeO4 (0.6), NiCl26H2O (4.6), ZnSO47H2O (20.5), H2SO4 (concentration 1 mL/L) and 1 mL/L of a vitamin-solution containing (mg/L): 4aminobenzoic acid (5), biotin (2), folic acid (2), lipoic acid (5), nicotinic acid (5), panthothenic acid (5), pyridoxine hydrochloride (10), riboflavin (5), thiamin hydrochloride (5) and vitamin B12 (0.1) were added (modified from [27,28]). Furthermore, 2.2 mg/L Resazurin as redox- and pH-indicator was supplemented. The medium was sterilized and degassed in a VARIOKLAV1 Series autoclave (H+P Labortechnik AG, Oberschleissheim, Germany) at an operating temperature of 1258C and an operating pressure of 1.4 bar, followed by gassing with argon 5.0 in the anaerobic glove-box.

Anaerobic microcosms Anaerobic biodegradation experiments with TPH-contaminated aquifer material under different electron accepting conditions (nitrate; sulfate; nitrate + sulfate; nitrate under sulfate reduction suppression) were carried out in 250 mL glass microcosms. Therefore each reactor was filled with 15 g of contaminated aquifer material and 220 mL mineral medium. Each of the 42 microcosms received nitrate (32.2 mM, designated as NIT), nitrate (32.2 mM) plus sulfate (20.8 mM, designated as MIX) and sulfate (20.8 mM, designated as SUL). Concentrations were chosen to allow for the biodegradation of a significant fraction of TPH, and were to be replenished if in limitation. Nitrate was supplemented as KNO3 and sulfate as Na2SO4. To detect the influence of sulfate reduction on nitrate depletion under nitrate-reducing conditions, another 42 nitrate-amended reactors were supplemented with Na2MoO4 (3 mM, designated as MOL) as sulfate reduction suppressor [29]. All bioreactors were filled in the glove box flushed with argon 5.0 and then sealed airtight with Mininert Valves1 (Sigma Aldrich, Vienna, Austria). Afterwards microcosms were incubated horizontally without shaking in the dark at 20  28C.

Identification of present eubacterial and archaeal microbiome For the determination of present eubacterial and archaeal microbiome in anaerobic microcosms, genomic DNA was extracted from the samples using PowerSoil1 DNA Isolation Kit (MO BIO Laboratories Inc., Carlsbad, USA). Since extracted genomic DNA-amounts were too small for further analysis, DNA was amplified via polymerase chain reaction (PCR) using universal primer pairs (515f/806r [30]) and native Taq-Polymerase (ThermoFisher Scientific, Waltham, USA). Further amplification of eubacterial and archaeal 16S rDNA, Illumina-Miseq sequencing, data processing and taxonomic assignment was conducted as described elsewhere [31]. Sequences are deposited in GenBank under SU1312167.

Analysis of TPH-concentrations Periodically, a triplicate of reactors was sacrificed and the solids were analyzed qualitatively and quantitatively for their total petroleum hydrocarbon (TPH)-content according to DIN EN ISO 16703:2011 as described elsewhere [26]. Briefly, TPH were extracted with n-heptane supplemented with n-C10 and n-C40 as

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retention time window (RTW) markers in an ultrasonic bath for 1 h. Clean-up was conducted using MgO3Si (Florisil1) following drying over Na2SO4. Extracts were analyzed by a HP 5890 Series II gas chromatograph with a flame ionization detector (Hewlett Packard, Palo Alto, USA) equipped with a DB-5HT capillary column (Agilent Technologies, Santa Clara, USA) at a constant helium carrier gas flow rate of 1.6 mL/min. The oven program was set to 608C for 1 min, then increased by 208C/min to 3908C and finally held 5 min at 3908C. To calculate TPH-concentrations chromatograms were integrated between n-C10 and n-C40 using the Agilent GC ChemStation Software. For conversion to mM molecular weight of n-C25 was used, since this alkane is located in the center of the investigated RTW.

Quantification of TEA-concentrations In all microcosms nitrate- and sulfate-concentrations were monitored on a weekly basis to calculate the percentage depletion of TEA. Therefore 30 mL of the aqueous supernatant were removed with a gas-tight syringe (Hamilton, Reno, USA) and analyzed ¨ NORM EN ISO 10304-1:2009 for nitrate- and sulaccording to O fate-concentrations on a Dionex ICS 900 ion chromatography system (ThermoFisher Scientific, Waltham, USA) with guard column AG14A, separation column AS14A, micro membrane suppressor AMMS, conductivity detector DS5 and an eluent of 8 mM Na2CO3 and 1 mM NaHCO3 at a flow rate of 1 mL/min. Also concentrations of chloride, nitrite and phosphate were analyzed by default, where chloride concentrations were used as a quality parameter for the measurement due to its high stability. After sampling, the microcosms were shaken lightly and then stored horizontally in the dark. Additionally, from a portion of microcosms 6 mL of the aqueous supernatant were removed, stabilized with HNO3 and analyzed via ULTIMA ICP-AES (Horiba Jobin Yvon, Kyoto, Japan) for its content of dissolved iron (Fe(II)), manganese (Mn(II)) and Mo concentrations as described elsewhere [27].

Sample preparation for stable isotope analysis Nitrate precipitation To avoid re-oxidation of nitrite to nitrate, precipitation was performed in an anaerobic glove-box (MECAPLEX, Grenchen, Switzerland) flushed with argon 5.0. First, the aqueous supernatants were carefully removed from the reactors with a Pasteur pipette without taking oil phase and aquifer material. Then the supernatants were filtered through Al2O3 for eliminating humic matter, followed by pH adjustment to 12 by adding 10 N KOH. Further, aqueous supernatants were dried first by heating and then for at least four hours at 708C in an incubator. After drying precipitation residues were crushed, filled in a tube and flushed with argon 5.0.

Sulfate precipitation After removal and filtration of the aqueous supernatants as described above, pH was set to 2–2.5 using 6 N HCl. Then the aqueous supernatants were heated to 758C and supplemented with 10 mL of a 0.2-M BaCl2-solution to precipitate sulfate as BaSO4, followed by incubation at 708C for 3 h. Furthermore, aqueous supernatants were sucked off over blue ribbon filters with 0.45 mm pore size (Schleicher & Schuell BioScience GmbH, Dassel, Germany) to collect precipitated BaSO4 and the loaded filters were

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dried at 958C in an incubator. After drying, BaSO4-precipitates were filled in a small tube and flushed with argon 5.0.

Analysis of stable isotope ratios

Research Paper

Stable isotope ratios (d15N for nitrate; d34S for sulfate) were measured with a Vario EL III Series elemental analyser (Elementar GmbH, Hanau, Germany) and a Delta Plus XP Series isotope ratio mass spectrometer (ThermoFisher Scientific, Bremen, Germany). Therefore, precipitation residues were oxidized in an oxidation furnace filled with W(IV)O3 at a temperature of 11508C. The NO3 formed in this process was reduced to N2 in a furnace filled with elementary copper at a temperature of 8508C, while the formed SO2 was hold in a trap. Subsequently, N2 and SO2 were passed to the isotope ratio mass spectrometer for isotope ratio measurement. Nitrate samples were calibrated against the international standard IAEA-NO-3 (KNO3) and d15N-values were based on N2 in air. Barium sulfate samples were calibrated against the standard NBS 127 (BaSO4) and d34S-values were based on Vienna Canyon Diablo Troilite (VCDT). For nitrogen isotope analysis the sample amount was aliquot to 1.3–1.5 mg KNO3 and for sulfur isotope analysis to 0.6–0.7 mg BaSO4. Device-specific measuring precisions amounted to 0.3% and 0.4% for nitrogen and sulfur isotope analysis and were accounted for via Gaussian error propagation in the statistical evaluation of the results.

Calculation of enrichment factors and statistical analysis For the determination of the enrichment factor of 15N in nitrate and 34S in sulfate the following form of the Rayleigh equation was used:     Rt Ct Rt ðdt þ 1000Þ ln ¼ ða1Þ  ln with : ¼ R0 C0 R0 ðR0 þ 1000Þ C0 and Ct are TEA concentrations initially and at time t, respectively. Similarly, R0 and Rt are initial and time-t isotopic ratios of 15 N/14N and 34S/32S. The term (a  1) is defined as the slope of a linear regression of ln(Rt/R0) over ln(Ct/C0). The isotope enrichment factor e (expressed in %) is calculated by multiplying (a  1) by 1000. For statistical analysis of correlations between TEA-decrease and measured shifts in TEA stable isotope ratios, linear regression analysis using the programming language R1 was performed. Statistical analysis was performed with TEA-depletion as independent variables (x) and the isotope ratio of 15N/14N and 34S/32S as dependent variables (y). To check whether the existing data sets fulfill all necessary conditions for the application of a linear regression, Durbin–Watson-Tests and analysis via diagnostic plots were performed. One-way ANOVA using Tukey post-tests (a = 0.05) was used for the analysis of final TPH and TEA concentrations.

Results and discussion Microbial phylogeny under different electron accepting conditions Archaeal and eubacterial community composition and biodiversity indices based on the analysis of aquifer 16S rDNA during incubation are shown in Table 1. Communities in microcosms receiving nitrate as sole electron acceptor (NIT and MOL) were very similar and had lower biodiversity than under mixed TEA-conditions. The addition of molybdate appeared not to profoundly influence community composition. Acidovorax and Pseudomonas, inter alia including organotrophic denitrifiers [17,32,33] were 4

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abundant in all experiments, where Pseudomonas represented one quarter of the microbial community with nitrate as sole electron acceptor, versus 6% under mixed conditions. There, Thiobacillus, whose members are capable of chemolithotrophic denitrification using reduced sulfur species as electron donors [17,34], Desulfovibrio, sulfate reducers known for the oxidation of hydrocarbon metabolites such as organic acids or alcohols [35,36], and Lactobacillus, biosurfactant-producing organisms aiding hydrocarbon degradation [37] were distinctly abundant. Although Thiobacillus includes some species capable of dissimilatory iron reduction, typical Fe(III) and Mn(IV) reducers (Geobacter, Shewanella) were absent. Methane-producing archaea Methanothermobacter and other Methanobacteriaceae formed about 1.3% of the community under mixed TEA-conditions, despite the unfavorably high RedOx-Potential, and were less abundant with nitrate as sole electron acceptor.

Petroleum hydrocarbon (TPH) degradation under nitrate and mixed TEA-conditions Figure 1 shows petroleum hydrocarbon (TPH) and terminal electron acceptor (TEA) concentrations in microcosms with nitrate as sole added electron acceptor (NIT, MOL) and under mixed terminal electron accepting conditions (combined nitrate and sulfate addition, MIX) over time. Degradation parameters are shown in Table 2. Within 252 days of incubation, nitrate reduction was significantly higher under addition of molybdate as sulfate reduction suppressor (8.04  0.37 mM/L), possibly due to a better expression of the molybdenum containing nitrate reductase [38], than without molybdate (NIT, 6.04  0.62 mM/L) and under mixed TEA-conditions (6.6  0.74 mM/L). In contrast, nitrate reduction was quantitatively independent from sulfate reduction when comparing NIT and MIX reactors. Although MOL reactors showed higher initial TPH degradation rates (first month), all microcosms exhibited a similar quantitative TPH decrease of around 16–19 mM/kg (related to n-C25) after 8 months of incubation. Thus, neither the addition of molybdate nor mixed terminal electron accepting conditions did increase TPH degradation. However, additional electron equivalents consumed by sulfate reduction under mixed conditions point towards the occurrence of more complete degradation, that is, the further conversion of partially oxidized metabolites – originating from organotrophic denitrification – using sulfate under mixed TEA conditions. Since the sulfate reducer Desulfovibrio, known for the oxidation of such polar compounds [36,39], was abundant under mixed conditions only, its responsibility for these processes appears likely. In contrast to MIX reactors, sulfate-only microcosms (SUL) did not show significant depletion of sulfate and TPH (one-way ANOVA tested against a = 0.05; not shown) after three months and were omitted from further analysis. This can be explained by the absence of sulfate reducers capable of activating hydrocarbons under sulfateonly conditions, and underline that sulfate reducing organisms were dependent on metabolites from denitrification under mixed TEA-conditions.

TPH fingerprint of sulfate reduction Earlier biodegradation studies of this crude oil revealed distinct qualitative, TEA-dependent degradation fingerprints. Higher molecular weight hydrocarbons were found to be preferentially

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TABLE 1

Relative microbial community composition (averages of n = 2 (NIT) and n = 3) on genus level based on 16S rDNA analysis after 113 (NIT, MOL) and 139 (MIX) days of incubation, and biodiversity indices. Relative contributions in all parallels <0.5% are omitted. Taxonomic assignment

NIT

MOL

MIX

Order

Class

Family

Genus

Euryarchaeota

Methanobacteria

Methanobacteriales

Methanobacteriaceae

Methanothermobacter Other

0.28 0.37

0.42 0.50

0.65 0.69

Acidobacteria Actinobacteria Actinobacteria

Holophagae Acidimicrobiia Micrococcales

iii1-8 Acidimicrobiales Cellulomonadaceae

Uncultured OCS155marinegroup Other Cellulomonas

Other Other Other Uncultured

6.61 1.12 0.42 0.36

5.07 3.93 0.57 0.97

5.03 0.24 0.18 0.13

Bacteroidetes

Bacteroidia SB-1 Sphingobacteriia

Bacteroidales Uncultured Sphingobacteriales

Porphyromonadaceae Other Other

Proteiniphilum Other Other

3.58 0.01 0.79

4.76 0.01 0.13

4.12 1.88 2.57

Chloroflexi

Anaerolineae

Anaerolineales

Anaerolineaceae

Levilinea Uncultured

0.38 8.58

0.31 6.91

0.65 2.16

Cyanobacteria

SHA-109

Uncultured

Other

Other

Firmicutes

Bacilli

Lactobacillales

Lactobacillaceae

Lactobacillus

Lentisphaerae

Lentisphaeria

RFP12gutgroup

unculturedbacterium

Other

Proteobacteria

a-Proteobacteria

Caulobacterales Rhizobiales Rhodobacterales Rhodospirillales Sphingomonadales Burkholderiales

Caulobacteraceae Rhodobiaceae Rhodobacteraceae KCM-B-15 Sphingomonadaceae Alcaligenaceae Alcaligenaceae Comamonadaceae Comamonadaceae Hydrogenophilaceae Hydrogenophilaceae Rhodocyclaceae Desulfovibrionaceae Other Enterobacteriaceae Other Pseudomonadaceae

b-Proteobacteria

Xanthomonadales

Sinobacteraceae Xanthomonadaceae

Uncultured Parvibaculum Other Uncultured Sandaracinobacter Bordetella Pusillimonas Acidovorax Other Thiobacillus Uncultured Azoarcus Desulfovibrio Other Citrobacter Other Pseudomonas Other Uncultured Thermomonas

Hydrogenophilales

d-Proteobacteria g-Proteobacteria

% of total community

Rhodocyclales Desulfovibrionales Acidithiobacillales Enterobacteriales PYR10d3 Pseudomonadales

1.66

2.38

0.25

13.18

12.46

22.73

0.02

0.01

1.47

0.00 0.40 0.02 0.84 0.04 0.04 4.20 9.25 0.51 0.61 0.44 0.79 0.06 12.46 0.60 1.78 24.73 0.54 1.11 0.64

0.01 0.49 0.01 0.83 0.07 0.52 3.26 9.58 0.62 0.49 0.88 0.50 0.02 7.87 0.49 1.87 25.49 0.54 2.84 0.97

0.80 1.57 1.09 1.52 1.97 0.17 0.89 13.76 0.71 11.00 0.04 0.09 4.12 5.74 0.01 1.18 5.55 0.11 0.09 0.93

Synergistetes

Synergistia

Synergistales

Synergistaceae

Uncultured

0.35

0.43

0.53

Verrucomicrobia

Spartobacteria

Chthoniobacterales

FukuN18FWG

Other

0.00

0.01

1.53

3.24

3.78

3.87

Contributors < 0.5% of total community Biodiversity indices

NIT

Chao1

3858

Shannon-Wiener (H’)

6.00

degraded with sulfate as TEA [23], contrary to the commonly observed inverse relation of chain length to actual degradation under aerobic [26], denitrifying and methanogenic conditions [23,25]. Figure 2 shows the relative abundance of four retention time based fractions based on 24,27 in the TPH-GC-FID chromatogram, each covering approximately one fourth of the total retention time between n-C10 and n-C40. Higher molecular weight compounds (equivalent carbon number between 32 and 40, Fig. 2) were similarly depleted under nitrate- and mixed electron accepting conditions. This suggests that sulfate reduction did not entail a significant qualitative fingerprint on hydrocarbon distribution, further corroborating the hypothesis that the initial steps

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MOL

MIX

3611

5942

5.80

8.00

of hydrocarbon degradation were nitrate-dependent also under mixed TEA-conditions.

Competing and predominant electron accepting conditions Besides added nitrate and sulfate, geogenic TEA appeared to be absent, with Fe(II) and Mn(II) being close to or below detection limit (0.005 mM), as was sulfate in NIT and MOL reactors, indicating the absence of additional geogenic sulfate by dissolution of gypsum (CaSO4). However, this may be masked by the precipitation of pyrite (FeS2), and the occurrence of Fe- and Mn-reduction appears not to be connected to abundant known Fe- and Mnreducers, as we recently observed in a groundwater study [31].

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40

6

TPH

40

4

NO3

20

-

2

NO2-

Research Paper

0

NO 2- [mM/L]

NO 3- [mM/L] TPH as n-C25 [mM/kg]

60

0 0

100

200

% relative abundance in TPH

NITRATE (NIT)

start NIT MIX

30

20

10

0 >10 to 17

>17 to 24

NITRATE + MOLYBDATE (MOL)

FIGURE 2

TPH 40

4

NO3-

20

NO2-

0

2

100

200

NITRATE + SULFATE (MIX) TPH

60

6

4

20

2 SO42NO2-

0 0

NO 2- and SO 32- [mM/L]

NO3-

40

Relative abundance of different hydrocarbon fractions at the start (blank bars) and after 233 days of incubation with nitrate as sole electron acceptor (NIT, light grey bars) and 242 days under mixed electron accepting conditions (MIX, dark grey bars). Fractions are based on chromatographic retention time of alkanes corresponding to the equivalent carbon number. Bars are 1 SD, n = 3.

denitrification [41] in all microcosms. Contrarily, the presence of the sulfur-oxidizing genus Thiobacillus (Table 1) under mixed conditions suggests the occurrence of nitrate-dependent sulfurreoxidation, that is, lithotrophic denitrification under mixed terminal electron accepting conditions.

0 0

NO 3- , SO 42- [mM/L] TPH as n-C25 [mM/kg]

>32 to <40

6

NO2- [mM/L]

NO 3- [mM/L] TPH as n-C25 [mM/kg]

60

0

100 200 Incubation Time [d]

SO32-

FIGURE 1

Petroleum hydrocarbon (TPH) concentrations converted to n-C25, and TEA during anaerobic incubation with nitrate (NIT), nitrate and molybdate (MOL) and nitrate and sulfate (MIX) in aquifer microcosms. Bars indicate one standard deviation (n = 3 for TPH and sulfite, n 30 for TEA and nitrite), some bars are smaller than the symbols.

In contrary to the redox-cycling of sulfur, the formation of N2 represents a nitrogen sink under groundwater conditions. For denitrification, different stoichiometries are plausible depending on the reaction products (NO2, NO, N2O or N2). Conversion to N2 has been observed for Pseudomonas [40], the most prevalent denitrifying strain in our experiments (Table 1). The biologically largely unavailable N2 can only be replenished to the nitrogen cycle via biological nitrogen fixation to NH4+. In addition, the small accumulation of nitrite (Fig. 1) points towards complete 6

>24 to 32

Equivalent carbon number

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Fate of hydrocarbon metabolites Theoretically, for the mineralization of 1 mM n-C25 a total of 30.4 mM NO3 are necessary [42]. The calculation of actual and theoretical electron donor/acceptor balances, and gaps therein, may indicate prevalent bioconversion processes, however disregarding microbial assimilation and possible abiotic losses of hydrocarbons via, for example, adsorption, adding to the imprecision arising from the reduction of a complex TPH matrix to one alkane (i.e. n-C25). Nevertheless, actual nitrate consumption amounted to only between 17 and 20% of the calculated stoichiometry (data from Table 2), suggesting the occurrence of partial oxidation, as we previously observed for denitrifying conditions [23]. Metabolic intermediates may have accumulated although the presence of a methanogenic community, most prominent under mixed conditions, supports further methanogenic conversion, despite unfavorable redox-conditions. On the other hand, methanogenesis may be inhibited by nitrate and sulfate concentrations even lower than presently used [43]. Low relative community contributions of methanogenic archaea are able to produce significant quantities of methane [31], and are not thus preclusive to methanogenesis.

Rayleigh model of

34

S-sulfate and

15

N-nitrate

The relation between the changes of isotope ratios during incubation in the residual TEA fraction (Rt/R0) and the corresponding decline in TEA concentration (Ct/C0) could be described well via linear regression in a Rayleigh-plot (Fig. 3). The Rayleigh kinetic enrichment model yielded isotopic enrichment factors

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TABLE 2

a

RedOx-potential [mV] pH [–]a

NIT

MOL

MIX

170 6.5–7

+38 6.5–7

170 6.5–7

TPH decreaseb

mM n-C25/kg  1SDc %  1SDc mM n-C25/MC  1SDc

16.4  1.8 31.6  3.5 0.25  0.03

18.7  0.4 43.2  0.9 0.28  0.01

18.9  1.7 35.7  3.2 0.28  0.03

TEA decreaseb

mM/L  1SDd

6.04  0.62

8.04  0.37

%  1SDd

19.4  2.0

25.3  1.2

mM/MC  1SDd

1.33  0.14

1.77  0.08

6.6  0.74e 3.14  0.29f 19.5  2.2e 13.7  1.3f 1.45  0.16e 0.69  0.06f

d15N [%]g

Start  1SDh End  1SDh

3.5  0.1 7.2  0.1

3.3  0.1 8.3  0.1

3.7  0.1 5.5  0.1

d34S [%]g

Start  1SDh End  1SDh

n.a. n.a.

n.a. n.a.

0.4  0.1i 1.0  0.1i

Isotopic enrichment factor [%]

eN  1 SD eS  1 SD

16.7  1.1 n.a.

18.1  0.1 n.a.

6.0  0.9 4.1  0.8

a

Day 165. Day 0–day 242 for MIX, day 233 for NIT and MOL. c n = 3. d n 30. e For nitrate. f For sulfate. g Device-specific measuring precision was 0.3 % for d15N and 0.4 % for d34S (based on measurement of international reference standards). h n = 9; device-specific measuring precision was taken into account via Gaussian error propagation. i Significant difference between d34S-values at start and end was tested against a = 0.05 via ANOVA and paired t-test. MC: microcosm, n.a.: not applicable. b

(Fig. 3, Table 2) of 16.7 and 18.1% for eN with nitrate as sole TEA and 6.0% and 4.1% for eN and eS, respectively, under mixed electron accepting conditions. Analytically detected value for the pure chemicals were 3.5% for KNO3 and 0.5% for Na2SO4, in good

agreement with day zero samples (Table 2). Tested against a significance level of a = 0.05, a significant linear dependence of d15N-values in the residual nitrate fraction and d34S for the residual sulfate to the corresponding TEA-decrease was found (p-values < 0.05; t-values > j1.96j). Furthermore, good model qualities for all performed linear regressions were reflected by an R2 > 0.82 and no outliers outside of Cook’s distance causing leverage effects were detected.

Fractionation of

FIGURE 3

Double logarithmic Rayleigh plot of the change in N and S isotopic composition (Rt/R0; d+1000) over TEA-concentrations (Ct/C0; NO3 and SO42) in different microcosms during incubation with petroleum hydrocarbons, calculated linear regression lines and R2-values; NIT (filled squares, green), MOL (filled diamonds, blue), and sulfate (empty circles, red) and nitrate (filled triangles, orange) under mixed terminal electron accepting conditions.

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15

N in nitrate

Few studies describe combined N and S fractionation associated to oxidative hydrocarbon transformation, and more data is available on nitrate as sole TEA. Nitrogen fractionation during denitrification was observed to be around, expressed as eN, between 25% and 30% in uncontaminated ground- and ocean water [12,13,16,44], but appears to be lower with hydrocarbons as electron donors. Two culture studies on aromatic compounds and metabolite fatty acids report eN well in the range of our observation, that is, around 17% during denitrification [22,45]. Qualitative analysis of chromatograms (not shown) revealed that straight-chain alkanes were degraded before more complex compounds, which was observed earlier under aerobic, denitrifying and methanogenic conditions for this crude oil [23,25,26]. An electron-donor structure-dependent extent of 15N fractionation, as suggested by Wunderlich and co-authors [22], would correspond to lower fractionation in the later stages of degradation of an increasingly complex petroleum hydrocarbon matrix. This was not evident in our data (Fig. 3).

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Research Paper

Depletion of TEA and TPH, nitrate-15N and sulfate-34S isotopic shifts and resulting kinetic enrichment factors eN and eS during incubation with nitrate (NIT) as sole added electron acceptor, under mixed terminal electron accepting conditions (MIX) and nitrate with sulfate reduction suppression (MOL).

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Under the addition of sulfate, fractionation of nitrate-15N was much lower (eN,MIX = 6%) than under nitrate-only conditions. While the relative decrease of nitrate (approximately 19.4%), was similar in NIT and MIX, different nitrogen fractionating processes appeared to be in effect. Under supplementation of molybdate, nitrate degradation proceeded faster, but effected similar fractionation (Table 2). Due to the similar nitrate concentrations applied in all microcosms, the reflection of the concentration dependency of 15N fractionation, as observed previously for marine environments [13,46] can be excluded, as is the generally reduced lower fractionation in sediments compared to open waters [47,48], which would similarly apply to all microcosms.

Evidence for lithotrophic, sulfur-dependent denitrification Based on our data, we explored the possibility of chemolithotrophic denitrification co-occurring with organotrophic denitrification under mixed terminal electron accepting conditions, and their reflections in isotopic fractionation. The (re)cycling of electron acceptors, that is, the interconnection of sulfur, nitrogen and carbon cycles is commonly known in sediments [49], but to our knowledge, isotopic investigation on the occurrence of lithotrophic versus organotrophic hydrocarbon biodegradation under different electron accepting conditions has not been published yet. Recently, chemolithotrophic denitrification and anaerobic ammonia oxidation (annamox) were identified to be associated with lower 15N fractionation than organotrophic denitrification, albeit in a limnic system [12,14]. Similar nitrite concentrations in the experiments (Fig. 1) are preclusive of the occurrence of substantial annamox. In contrast to organotrophic denitrification, nitrate reduction using inorganic electron donors, that is, lithotrophic denitrification, is reflected in significantly lower 15N fractionation. An eN of 7.5 to 9% was observed recently [12] in limnic, sulfide-dependent denitrification, and was attributed to either limited substrate availability or low fractionation inherent in the process. For our study, the former can be excluded since both electron donor and nitrate were present sufficiently. Lower inherent fractionation during lithotrophic dentrification is attributable at least in part to the mediation of lithotrophic denitrification by the periplasmic nonrespiring NAP, which was associated with lower N fractionation in Rhodococcus sphaeroides earlier [13]. In contrast, the respiratory membrane-bound NAR or NAS are responsible for the fractionation during organotrophic denitrification [13,46]. On the other hand, both sulfate reducing (Desulfovibrio) and sulfur-oxidizing (Thiobacillus) genera were more abundant under mixed terminal electron accepting conditions. Although biodegradation of n-alkanes by Desulfovibrio was reported in a few studies between 1940 and 1970 [50–52], since then all attempts to reproduce hydrocarbon activation performed by this bacterial strain failed [53]. Nevertheless, the capability of Desulfovibrio for the oxidation of TPH intermediates, like alcohols or fatty acids, is well known [36,39]. Members of Thiobacillus are capable of using sulfite, sulfide, elemental sulfur and pyrite as electron donors for denitrification [17,18,54], but our data do not allow for the identification of the main sulfur species driving lithotrophic denitrification. This has, however, interesting consequences for the identification of dominant electron accepting pathways in hydrocarbon biodegradation. With reduced sulfate recycling to nitrate, the actual amount of sulfate used as TEA for 8

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34

hydrocarbon biodegradation under mixed TEA conditions may be underestimated if sulfate reducers capable of TPH activation are absent.

Fractionation of

34

S in sulfate

Similarly to nitrogen, sulfur fractionating processes were expressed poorly or masked under mixed electron accepting conditions, where low but significant sulfur fractionation was observed (see Table 2). There, eS was estimated to be 4.1  0.8%, and d34S did not exceed 1% (Table 2) at the end of the experiments. Fractionation could not be determined with sulfate as sole electron acceptor (not shown) due to low sulfate depletion, however indicating that sulfate reducing organisms are profiting from nitrate addition. The absence of sulfate reducers capable of activating hydrocarbons in our sediments explains the lack of TEA and TPH degradation under sulfate-only conditions. Several studies have investigated 34 SO4 isotopic shifts during petroleum hydrocarbon biodegradation under sulfate-reducing conditions in the field and laboratory scale. Detmers et al. [19] could not identify a relation between sulfate reduction rate and magnitude of sulfur fractionation while in contrast, carbon source mineralization entailed higher 34S fractionation than partial oxidation. The impact of community composition on sulfur fractionation was underlined by the identification of sulfate-reducer-specific enrichment factors, which ranged up to eS = 42% in the mineralizing Desulfonema magnum but where as low as 2% for partial carbon oxidation by Desulfovibrio halophila, also representing the most abundant identified sulfate reducer in the present study. With sulfate reduction as dominant TEA, however, more pronounced fractionation was observed, with eS between 20 and 46% [20,21]. Sulfate isotopic fractionation is, to our knowledge, not fully investigated under mixed terminal electron accepting conditions, and the present results suggest significant interactions with nitrogen cycling.

Microbial team play in sulfur, nitrogen and carbon cycling The main drivers for the observed processes appear to be members of Desulfovibrio, typically reducing sulfate to hydrogen sulfide concurrent to using polar TPH intermediates as electron donors [36,39], with Thiobacillus performing lithotrophic, nitrate dependent sulfide reoxidation to sulfate. The sulfur isotopic imprint of these recycling processes can be expected to be generically low [55] in addition to the confined nature of our microcosms, where the previously 34S-depleted, reoxidized sulfate is simply added to the sulfate pool it originated from. An alternative explanation is offered by the low sulfur fractionation of abiotic processes such as pyrite precipitation [55] and subsequent reoxidation [17,56]. These processes are not prevalent in groundwater systems [21] and would have also occurred in the sulfate-only experiments. Thus, under mixed TEA-conditions, Thiobacillus were responsible for lithotrophic denitrification, with Pseudomonas and more pronouncedly Acidovorax and Lactobacillus performing organotrophic denitrification, that is, hydrocarbon degradation, as indicated by the relatively higher abundance in MIX versus NIT and SUP experiments (Table 1). Thus, the observed 15N isotopic enrichment represents a superposition of lithotrophic and organotrophic denitrification processes. The role of methanogens under relatively high redoxpotential is yet to be determined.

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Electron acceptor recycling under mixed terminal electron accepting conditions Under mixed terminal electron acceptors, the assessment of the relative contributions of TEA for TPH degradation is rendered difficult by lithotrophic denitrification. The stoichiometry for sulfatedependent TPH degradation is 19.0:1 of SO42:n-C25, assuming TPH mineralization and conversion of sulfate to H2S. If nitrate is present in abundance, as in our experiments, sulfides can be expected to be quickly reoxidized to sulfate [57], while incomplete oxidation to S0 can be expected under nitrate limiting conditions [58]. Stoichiometrically, for the production of 1 mM SO42 from reduced sulfur species, 1.6, 1.2 and 0.8 mM NO3 are required for the oxidation of S2, S0 and S2O32, respectively, assuming complete denitrification to N2. Thus, the observed depletion of nitrate under mixed terminal electron accepting conditions (1.45 mM) may mask the reoxidation of 0.9–1.8 mM reduced sulfur species to sulfate, thus linking sulfur and nitrogen cycles in hydrocarbon degradation. However, considering the presence of organotrophic denitrifiers such as Pseudomonas, it can be assumed that a significant portion of nitrate was in fact consumed during TPH oxidation, while the role of methanogenic archaea under mixed terminal electron accepting conditions remains to be elucidated.

Conclusions Few studies on the anaerobic electron acceptor isotopic fractionation during petroleum hydrocarbon biodegradation are available.

In our study, we collected evidence for concurrent lithotrophic and organotrophic denitrification in addition to sulfate-dependent TPH oxidation with an unquantified contribution of methanogenesis. These are a new aspect opposing the classical, thermodynamically driven redox-zonation concept postulating subsequent TEA depletion, and may aid in a better understanding of coupled nitrogen, sulfur and carbon cycles in the groundwater and active hydrocarbon plume fringes. In the light of these findings, the use of electron acceptor isotopic fractionation factors as a tool to quantify the biodegradation of complex petroleum hydrocarbon matrices that abstract themselves from carbon isotopic analysis due to hydrocarbon molecular weight and/or complexity is not straightforward if the number of natural or anthropogenic TEA exceeds one.

Acknowledgements Financial support for the project IsoMon by the European Regional Development Fund (EFRE) together with the Government of Lower Austria, contract No. WST3-T381/029/ 2011 and WST3-T81/034/2014 (Project IsoMon), is greatly ¨ llern and J. appreciated. M. Sumetzberger-Hasinger, K. Mu Heindler are thanked for laboratory assistance, D. Backes, T. Lindner and L. Lehner for molecular biological analyses and A. Loibner and T.G. Reichenauer for project administration. Two anonymous reviewers are thanked for their helpful suggestions to improve the manuscript.

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S-sulfate isotopic fractionation reflects electron acceptor ‘recycling’ during hydrocarbon biodegradation, New