Increased rates of dissimilatory nitrate reduction to ammonium (DNRA) under oxic conditions in a periodically hypoxic estuary

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ScienceDirect Geochimica et Cosmochimica Acta 133 (2014) 313–324 www.elsevier.com/locate/gca

Increased rates of dissimilatory nitrate reduction to ammonium (DNRA) under oxic conditions in a periodically hypoxic estuary Keryn L. Roberts, Adam J. Kessler, Michael R. Grace, Perran L.M. Cook ⇑ Water Studies Centre, School of Chemistry, Monash University, Clayton, Australia Received 17 September 2013; accepted in revised form 21 February 2014; available online 12 March 2014

Abstract The Yarra River Estuary is a salt wedge estuary prone to hypoxia in the bottom waters during low flow periods. Rates of denitrification and dissimilatory nitrate reduction to ammonium (DNRA) were quantified using 15N in relation to oxygen, nitrate and available reductants. Denitrification was the dominant nitrate reduction pathway under all oxygen conditions, however, DNRA increased from <1% under hypoxic conditions (<50 lmol L1 O2) to 18% of total nitrate reduction under oxygen saturation in the water column in intact core incubations. Microprofiles of nitrate reduction pathways in intact cores using diffusive equilibrium in thin layer (DET) gels showed significant rates of DNRA only occurred under oxic conditions in the presence of Fe2+. Cores incubated anoxically, developed free sulfide within the porewater, had very low concentrations of Fe2+ and low rates of DNRA. Slurry incubations with varying concentrations of NO 3 showed that denitrifying bacteria had a higher affinity than nitrate ammonifying bacteria with Km values of 49 and 86 lmol L1 for denitrification and DNRA, respectively, however, this could not explain the change in the rates of DNRA relative to denitrification observed. Further slurry incubations to investigate the relationship between DNRA and Fe2+ oxidation were inconclusive and complicated by very high backgrounds of sorbed (HCl extractable) Fe2+. Addition of Fe2+ to the slurry did not stimulate denitrification compared to a control (no Fe2+ addition), however, there was a significant decrease in the Fe2+ concentration over the period where DNRA occurred in the Fe2+ addition treatment, and no significant decrease in the control treatment. The ratio of DNRA to Fe2+ consumption was 15 ± 6 and 7 ± 3 for the Fe2+ and control treatments, respectively. We suggest reduced rates of DNRA under anoxic conditions can be explained by the binding of Fe2+ with free sulfides and the formation of FeS removing available Fe2+ for DNRA. Ó 2014 Elsevier Ltd. All rights reserved.

1. INTRODUCTION Nitrogen is a key limiting nutrient in the marine environment and increasing inputs from anthropogenic sources Abbreviations: DNRA, dissimilatory nitrate reduction ammonium; ADM, ammonium diffusion method; DO, dissolved oxygen; DET, diffusive equilibrium in thin films ⇑ Corresponding author. Address: G25a B19 Water Studies Centre, Chemistry Department, Monash University, Clayton, Victoria 3800, Australia. Tel.: +61 3 9905 4091. E-mail address: [email protected] (P.L.M. Cook). http://dx.doi.org/10.1016/j.gca.2014.02.042 0016-7037/Ó 2014 Elsevier Ltd. All rights reserved.

have led to an increasing incidence of eutrophication and hypoxia (<100 lmol-O2 L1) in coastal waters (Jorgensen and Sorensen, 1988; Conley et al., 2007; Diaz and Rosenburg, 2008). In addition to detrimental ecological impacts, hypoxia also significantly affects biogeochemical cycling pathways, especially for nitrogen. Hypoxic conditions are known to increase the release of bioavailable nitrogen, primarily in the form of ammonium (NH+ 4 ), at the expense of nitrogen removal processes such as denitrification (Kemp et al., 2005). The key mechanism for this is thought to be a decrease in nitrification resulting in less nitrate (NO 3)

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available for denitrification (Rysgaard et al., 1996; Roberts et al., 2012). In addition, more reducing conditions that accompany hypoxia are thought to favour dissimilatory + NO 3 reduction to NH4 (DNRA) over denitrification (and anammox), although the controls on this process remain poorly understood. Four key factors are thought to control the balance between nitrate reduction via denitrification and DNRA: (1) NO 3 availability, (2) temperature, (3) organic carbon loading, and (4) availability of reductants such as sulfide or iron(II) (Fe2+). The ability of denitrifiers or nitrate ammonifiers to rapidly respond to the presence of NO 3 can determine whether it is converted to N2 or NH+ under varying oxygen and 4 NO 3 concentrations. Unlike denitrifiers, nitrate ammonifiers possess constitutive enzymes and therefore are able to survive under variable NO 3 conditions, potentially outcompeting denitrifiers when nitrate is sporadically available (Jorgensen, 1989). In salt marsh sediments, King and Nedwell (1985) observed the proportion of DNRA increased up  to 50% of total NO 3 reduction when NO3 concentrations were low, a finding supported by several other researchers (Ogilvie et al., 1997; Childs et al., 2002; Strohm et al., 2007; Nizzoli et al., 2010). Under these conditions, nitrate ammonifiers are energetically favoured, producing more energy per mole of NO 3 than denitrifying bacteria (Childs et al., 2002; Nizzoli et al., 2010). Seasonal variation in temperature can determine the species composition of a microbial community (Tison et al., 1980; King and Nedwell, 1984; Ogilvie et al., 1997). In estuarine sediments, Ogilvie et al. (1997) observed a shift in the heterotrophic microbial population from NO 3 ammonifiers at high water temperatures (20 °C) to denitrifiers at low temperatures (5 °C) in laboratory enrichment experiments. Studies have reported that the NO 3 affinity of nitrate ammonifiers is higher than that of denitrifying bacteria at temperatures above 10 °C, whilst below these temperatures denitrifiers have a greater affinity for NO 3 (King and Nedwell, 1984; Ogilvie et al., 1997). Likewise, in the tropical Cisadane estuary, Indonesia, Dong et al. (2011) observed an increase in the rate of DNRA to 10fold greater than denitrification when NO 3 was in excess. In that study, the high rates of DNRA were explained by the higher affinity of nitrate ammonifiers for NO 3 at tropical temperatures (Dong et al., 2011). Heterotrophic DNRA is carried out by fermentative bacteria and is observed in environments with high labile organic carbon availability (Tiedje, 1988; Yin et al., 2002). Cole and Brown (1980) observed the accumulation of ace+ tate and formate, with NO 2 reduction to NH4 in strains of Escherichia coli sp., a pathway later confirmed by Bonin (1996). In soils, the addition of labile organic carbon increased DNRA from 2% to 25% of total NO 3 reduction when the C/N ratio was high (Fazzolari et al., 1998). This finding was supported by Yin et al. (2002), who noted that soils with high organic carbon inputs were able to support a large heterotrophic population of bacteria, 20% of which were nitrate ammonifiers compared to only 6% in carbon depleted soils. Under these condi+ tions >50% of the NO 3 added was converted to NH4

and organic N at the expense of denitrification (Yin et al., 2002). Chemoautotrophic DNRA has also been reported and is largely controlled by available reductants such as sulfide and Fe2+. DNRA has long been linked to sulfide oxidation with slurry experiments showing NH+ 4 production linked to sulfide oxidation accounting for up to 30% of total nitrate reduction (Brunet and Garcia-Gil, 1996). Intact sediment core incubations have also shown DNRA is the dominant nitrate reduction pathway in highly sulfidic sediments (An and Gardner, 2002). A number of sulfur-oxidising bacteria, for example Thioploca sp. and Beggiatoa sp., can store and + reduce NO 3 to NH4 as a secondary metabolic pathway (Otte et al., 1999; Schulz and Jorgensen, 2001). In the presence of Beggiatoa sp. in Aarhus Bay, Denmark, DNRA was the predominant pathway of nitrate reduction; however, in the absence of sulfur oxidising bacteria, DNRA ceased (Sayama et al., 2005). Numerous studies have linked iron oxidation to NO 3 reduction with N2 as the main product (Nealson and Saffarini, 1994; Straub et al., 1996; Benz et al., 1998); however, studies linking Fe2+ oxidation to NH+ 4 production via DNRA have been more limited. It has previously been shown that DNRA can be coupled to Fe2+ oxidation in wetland and river floodplain sediments and that Geobacter sp. and Betaproteobacteria mediate this process (Weber et al., 2006; Coby et al., 2011). Most recently, Hou et al. (2012) inferred that Fe2+ could be a controlling factor on DNRA rates based on a strong correlation between 15 NH+ 4 production and reactive iron oxide concentrations in Texas coastal sediments. The Yarra River Estuary is prone to periodic hypoxia in the bottom waters during low summer flows due to its typical salt wedge structure. A previous study showed the importance of DNRA relative to denitrification increased in the presence of oxygen in the water column, despite DNRA being expected to occur under more reducing and low NO 3 conditions (Roberts et al., 2012). In that study it was hypothesised that NO 3 concentration was a possible control on DNRA. The purpose of this study was to investigate these unexpected findings further and explore the behaviour of DNRA under different oxygen conditions using both slurry and intact core experiments. Specifically three research questions were addressed: (1) Can the behaviour of DNRA and denitrification in the previous study be replicated under controlled conditions? (2) How do denitrification and DNRA behave with respect to NO 3 concentration? and (3) Does the availability of Fe2+ change the relative contributions of denitrification and DNRA to total nitrate reduction? 2. METHODS 2.1. Site description Nitrate Reduction pathways were quantified in the Yarra River Estuary, Melbourne Australia in November 2009, May 2012 and January, February and May 2013 (Fig. 1). In November 2009 sediment samples were collected near Morell Bridge (55°320 27.2500 E 58°110 38.500 N), on all other

K.L. Roberts et al. / Geochimica et Cosmochimica Acta 133 (2014) 313–324 Maribyrnong River

Yarra River

N Dights Falls

A B Gardiners Creek 0

4 km

Port Phillip Bay

Fig. 1. The Yarra River Estuary, Australia. Sediment samples were collected from (A) Morell Bridge and (B) Scotch College in the salt wedge region of the estuary.

sampling occasions sediment samples were collected from 4.5 km upstream at Scotch College (55°320 63.4800 E 58°100 85.400 N) to determine the behaviour of denitrification and DNRA within the Yarra River Estuary (Fig. 1). The estuary exhibits a typical salt wedge structure and is prone to extended periods of hypoxia during low rainfall. The salt wedge region of the estuary is aphotic in the bottom waters owing to the high turbidity of the surface waters (Roberts et al., 2012). 2.2. Sampling Sediment cores were collected in 27.5  6.6 cm polyethylene cylinders and stoppered. Saline bottom water was collected using a bilge pump or a 5 L Niskin bottle. Sediment cores were transferred to a temperature-controlled water bath. Each core was mixed using a magnetic stirrer (40 rpm) suspended 2 cm above the sediment surface to prevent disturbance of the sediment. 2.3. Intact core incubations Sediment and water samples were collected on the 24th November 2009 at Morell Bridge. Rates of 15N reduction were calculated from the water-column accumulation of 29 15 N2, 30N2 and 15NH+ NO 4 after the addition of 3 in two treatments (100% air saturated and hypoxic water column) over a period of 21 days, measured in the water column on days 1, 7, 11, 13, 18 and 21. During the incubation phase, water was circulated through the cores using a peristaltic pump; for each treatment the reservoir was bubbled with air for oxic, and argon for hypoxic to maintain the desired oxygen concentration over the 21 day period. During measurements, the circulation was turned off and the cores were sealed. Four cores were kept oxic (200–300 lmol-O2 L1) for the 21 day incubation period as the control treatment, and eight cores were hypoxic (<50 lmol-O2 L1) for the first 11 days. On day 11 of the incubation, four of the eight hypoxic cores were terminated to determine the concentra-

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tion of 15NH+ 4 accumulated within the sediment. The remaining four hypoxic cores were made oxic by aerating the reservoir. Denitrification and DNRA were measured using the methods in Dalsgaard et al. (2000), however, to continue the experiment over the 21 day incubation period, the core was kept intact and a sample collected from the 29 water column to quantify 15NH+ N2 and 30N2. Water 4, removed was replaced with saline water and accounted for in calculations. At the start of the incubation, the dissolved oxygen (DO) and pH were measured in the water column of each core using a HQ-40d Hach multimeter (Germany). A  NOx (NO 3 + NO2 ) sample was filtered (0.45 lm; 30-mm polypropylene housing, Bonnet) and stored frozen until analysis. An aliquot of 0.4 mL of 0.05 mol L1 15NO 3 (Cambridge 98% 15N, 40 lmol L1 final concentration) was added to the overlying water in each core and mixed thoroughly without disturbing the surface of the sediment. To determine the final concentration of 15NO 3 in the overlying water, a 12 mL NOx sample was collected after stirring; the displaced water was replaced and the core sealed. The cores were incubated for 7 h, stirring and at in situ temperature. At the end of the incubation period, the DO and pH were measured in the overlying water and a 12 mL sample was filtered for the analysis of NOx. A 12.5 mL unfiltered sample was collected for N2 in a gas tight vial (Exetainer, Labco) and preserved with 250 lL ZnCl2 (50% w:v). A 200 mL unfiltered sample was also collected for analysis of 15NH+ 4 via the ammonium diffusion method (ADM) (see Section 2.7). After the incubation period, the core was slowly refilled with saline water from the reservoir to avoid disturbance to the sediment surface. The cores were circulated with reservoir water to allow for re-equilibration before nitrate reduction pathways were measured again. To determine the background concentration and account for the carry-over of 15NO 3 for subsequent measurements on day 7, 11, 13, 18 and 21, a sample for 15N–N2 and 15NH+ 4 was collected from the reservoir at the start of the incubation. After sampling on day 21, the water was removed from the sediment surface of the remaining 8 cores and the cores were extruded. The surface 3 cm of sediment was homogenised and a 20 mL subsample extracted with 1:1 (v:v) 2 mol L1 KCl, shaken at 120 rpm for an hour and then centrifuged. The supernatant was measured for 15NH+ 4 via the ADM (see Section 2.7). We found the vast majority of the 15NH+ 4 produced accumulated in the water column (>95%). We note that N2 accumulation was only measured in the water column and while we would expect some to accumulate in the sediment, this would be less than 15 NH+ 4 and therefore, neglecting this would not alter our results appreciably. Denitrification (D15) and DNRA (DNRA15) were calculated from the accumulation of 15 N labelled N2 and 15NH+ 4 , respectively, over time (see Section 2.7). 2.4. Nitrate slurry experiments Sediment and water samples were collected from Scotch College on the 15th May 2012 (see Section 2.2). The

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sediment cores were covered with site water and aerated overnight before slurries were prepared the following day. Sediment slurries were prepared by removing the surface 1 cm of sediment from the core after extrusion. The 1 cm slice was mixed with bottom water at a ratio of 1:1 (v:v). Slurries were prepared in duplicate; denitrification in a 12.5 mL vial (Exetainer, Labco) containing 8 mL of slurry, and DNRA in a 30 mL serum bottle containing 18 mL of slurry. The bottles were sealed with a butyl rubber septum and purged with He for 5 min to remove oxygen. It was observed that any residual oxygen after this time was consumed within a few minutes. Additions of 15NO 3 were made to the slurries to a final concentration of 10, 30, 60, 100 and 200 lmol L1. Five slurries were prepared for each treatment and mixed on a shaker table at 125 rpm for the duration of the incubation period. Slurries were terminated at 20 min intervals via the addition of 250 and 500 lL ZnCl2 (50% w:v) for the denitrification and DNRA slurries, respectively. The total incubation period was <2 h. After termination of bacterial activity, the denitrification samples were pre-treated with 50 lL of air to increase the background concentration of 14N in the sample; this was done so that excess 15N could be calculated from the ratios of 14N/15N. Samples for 15NH+ 4 analysis were pre-treated with 90 lg 14NH+ 4 to increase the background concentration of nitrogen before analysis via the ADM (see Section 2.7). 2.5. Micro-profiles in intact cores – oxygen treatments Sediment cores were collected in January and February 2013 (see Section 2.2). Two treatments were undertaken: oxic, and anoxic. Six cores were collected for each treatment, 3 for H2S and O2 measurements and 3 for nitrate reduction pathways, nutrients and total Fe measurements. Oxic conditions were maintained during pre-incubation by circulating aerated site water above the sediment; during incubations the core was gently bubbled with air. Anoxic cores were circulated with Ar-purged site water and then sealed during incubations. The isotopically labelled products of denitrification (29N2 and 30N2) and DNRA  (15NH+ 4 ) were profiled as well as O2, total Fe, NO3 , total + NH4 and H2S in triplicate cores for each treatment. Profiles were undertaken in open cores whilst being bubbled with air for oxic treatments and argon for the anoxic treatment. Profiles of O2 were measured in triplicate for each core using a fibre-optic sensor and a computer controlled micromanipulator (Pyroscience OXR230 connected to a Firesting O2 metre and MUX2 profiler). The O2 sensor was calibrated with 0 and 100% O2 saturated water of site salinity. H2S was measured in triplicate using an amperometric microsensor (Unisense H2S-100 connected to a Unisense microsensor amplifier). The H2S microsensor was calibrated in accordance with the Unisense procedure; calibration standards were prepared under anoxic conditions in pH 4 buffer over the linear sensor range of 0 to 300 lmol L1. Profiles of pH (Unisense pH-100) were measured under oxic and anoxic conditions in September 2012 however the microsensor broke during use. As such, we

have based the calculation of total sulfide (S2 tot ) on the assumption the averaged pH profiles will be similar in these current experiments. Nitrate reduction pathways were measured using diffusive equilibrium in thin layer (DET) gels combined with 15 N enrichment experiments. DNRA was measured as described in Stief et al. (2010) and denitrification measured via a 15N–N2 DET gel method (Kessler et al., 2013). The acrylic gel probes were constructed as described previously in Krom et al. (1994) with the dimensions 8.5 (h)  6 (w)  0.8 (d) cm and a window where the gel is in contact with the sediment of 4.6 (w)  5.0 (h) cm. Gels were cast as described in Stief et al. (2010); 80 mL acrylamide (15% w:v) was mixed with 40 mL N,N-methylenebisacrylamide (2% w:v) and slowly stirred. 1.5 mL dipotassium peroxodisulfate (0.11 mol L1) was added to initiate the polymerisation reaction and 120 lL N,N,N0 ,N0 -tetramethylethylenediamine was used as the catalyst. The gel was cast between two glass plates (pre-heated 40 °C) with 1 mm spacers and then clamped into position. The mould was heated to 40 °C for 20 min until polymerisation was complete. After the gel was cast, it was hydrated in NaCl solution at the same salinity as the site water for 24 h. The gel was 2 mm thick after hydration. The gel was then cut to size, placed inside the gel probe and covered with 0.2 lm pore size polyethersulfone filter membrane (Pall, Australia) to prevent bacterial and physical degradation of the gel. The constructed gel probe was then stored at 4 °C in NaCl solution (site salinity) until experiments. The overlying water column of the sediment core was 1 dosed with 1.5 mL 0.05 mol L1 15NO fi3 (150 lmol L nal concentration) 24 h prior to the deployment of the gel. The probe storage solution was purged for 6 h with N2 before deployment to remove oxygen from the gel surface and then the gels were inserted vertically into the sediment. Oxic conditions were maintained within the core by gently bubbling with air and anoxic cores were kept sealed. Two gel probes were inserted back to back in each core, one for the measurement of nitrate reduction pathways and the other for total NH+ 4 , NOx and total Fe. The gels were allowed to equilibrate for 30 h before the gels were removed rinsed and sliced into 2 mm slices with depth. The gel was cut on an acid washed glass plate using a homemade stainless steel cutting device acid washed in between samples. Gel slices for nutrients and total Fe were placed into 10 mL of ultrapure water and frozen until analysis (see Section 2.7). Denitrification and DNRA were determined from the accumulation of 15N–N2 and 15NH+ 4 within the gel, respectively. Gels slices were immediately placed into pre-filled 3 mL vials (Exetainer, Labco) containing ultrapure water and 100 lL ZnCl2 (50% w:v). The process was filmed to calculate the exposure time of the gel to air and these times were used in the correction of N2 concentration after calibration of the method (see Section 2.7). At least 24 h after the gel slice was placed in the vial, a 1.5 mL helium headspace was introduced for the analysis of 29N2 and 30N2 gas. The 1.5 mL sample removed was placed in a separate vial (Exetainer, Labco) and purged with helium and 15NH+ 4 was collected from the sample using the hypobromite method (see Section 2.7).

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2.6. Iron slurry experiments Sediment samples were collected on May 9th, 2013 from Scotch College (see Section 2.2). Cores were aerated in site water overnight prior to the preparation of slurries. Slurries comprising of 5% wet sediment were prepared with site bottom water and the surface 1 cm of sediment. A volume of 15 mL slurry was sealed in a 20 mL gas tight glass serum bottle and purged with He for 5 min to remove oxygen. Two treatments were prepared: a control treatment with 1 2+ added 15NO treatment 3 only (1 mmol L ), and an Fe 15  1 2+ with both NO3 (1 mmol L ) and Fe (5 mmol L1) addition. The incubation was carried out over 8 days and samples were collected at time = 0, 3, 12 and 24 h and time = 2, 3, 4, 6 and 8 days thereafter. Throughout the incubation period, slurries were shaken in the dark at 180 rpm at 22 °C. Since 2+ it has been shown that NO at 2 spontaneously oxidises Fe acidic pH, the anoxic slurry was centrifuged at 228 G for 2 min and the supernatant analysed for Fe2+ before extraction of the sediment with HCl to avoid this potential artefact (Weber et al., 2001; Klueglein and Kappler, 2013). Before the slurry was uncapped, a 1.5 mL gas sample was collected from the headspace and introduced into a vial (Exetainer, Labco) containing ultrapure water. The slurry was then opened and an unfiltered water sample was collected rapidly in a 3 mL vial (Exetainer, Labco) for 15 NH+ 4 and preserved with 100 lL ZnCl2 (50% w:v). The remaining water was filtered (0.2 lm pore size, Advantec) + immediately for analysis of NOx, NO 2 and NH4 , and of this, a 0.5 and a 0.1 mL filtered subsample were preserved in ferrozine for the analysis of Fe2+ for the control and Fe treatment respectively. Nutrient samples were frozen until analysis (see Section 2.7). The remaining sediment was homogenised and 0.2 g weighed into a centrifuge tube containing 20 mL 0.5 mol L1 HCl, to determine the amount of easily reducible iron in the sediment (HCl extractable Fe2+). The sample was shaken at 200 rpm for 2 h, then filtered and a 0.5 and 0.1 mL subsample preserved with ferrozine for the control and Fe treatments, respectively. For the determination of 29N2 and 30N2, a 100 lL subsample of the gas collected was injected into a purged vial (Exetainer, Labco) containing 1.5 mL helium headspace and 50 lL of air to increase the background of 14N as described previously. Samples for 15NH+ 4 were determined via the hypobromite method (see Section 2.7). 2.7. Analytical methods + Nutrient samples (NOx, NO 2 and NH4 ) were analysed via flow injection analysis (FIA) (Lachat Quickchem 8000 Flow injection Analyser, spectrophotometric detector). Standards (0–1 mg-N L1), spikes and reference materials were within acceptable limits and analysed following the procedures in Standard Methods for Water and Wastewater (2005). For both the slurry and gel experiments, iron was determined via the ferrozine method (Stookey, 1970; Voillier et al., 2000). Standards were prepared in the range

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0–1 mg L1 Fe and triplicate absorbance readings taken at 632 nm using a UV–Visible Spectrophotometer (GBC UV Visible Spectrophotometer – 918), all reagents were prepared as described in Voillier et al. (2000). Water column samples for iron were fixed with 0.5 mL ferrozine immediately to preserve Fe2+ in the sample; if the expected concentration of Fe2+ was high (e.g. slurry samples), the amount of ferrozine was doubled. Only Fe2+ was determined for the slurry samples because Fe2+ was 90% of total Fe. As a result, the final two reagents for total Fe (hydroxylamine hydrochloride and the ammonium acetate buffer) were omitted. For the gels, total Fe was determined by the addition of 0.5 mL ferrozine and 1 mL hydroxylamine hydrochloride and allowed to react 20 min before the addition of 0.25 mL of ammonium acetate buffer. After the reagents were added, the samples were left for at least 1 h for colour development before analysis. To account for diffusive loss of N2 in the gel samples during the handing process after incubation, a set of N2 calibration standards were prepared. The gels were prepared as described previously, but were then hydrated and stored in ultrapure water. Five standards were prepared in gas tight glass jars to final concentrations of 5, 10, 25, 50  and 100 lmol L1 15NO 2 . To convert NO2 to N2, sulfamic acid prepared in 5% HCl was added to the standards to a final concentration of 16.5 mmol L1 and a final pH of 1.5–2. Standards were then stoppered and shaken overnight. The pH of the standard was neutralised using 2 mol L1 NaOH to stop the conversion of NO 2 to N2 after which the gel probe was inserted. The standards were stoppered again for >24 h and a N2 gas sample was collected in a 3 mL vial (Exetainer, Labco) and preserved with 100 lL ZnCl2 (50% w:v) to determine the initial concentration of 15 N–N2 (t = 0). The gels were then removed, sliced and analysed as described previously. The process was filmed to calculate the exposure time of the gels to air. To account for the loss of N2 over time, a multi-variable regression was used to fit the concentration observed in the gels as a function of initial concentration and exposure time. The amount of 15N–N2 detected after exposure (ct in lmol L1) was related to the initial concentration of 15N–N2 (ci in lmol L1) and the time the gel was exposed to air (t in s) (Eq. (1)). ct ¼ ðAci þ BÞ lnðt þ 1Þ þ ðCci þ DÞ

ð1Þ

The fitted constants A = 0.0713, B = 0.5939, C = 0.5459 and D = 3.247 were used to determine the initial concentration from Eq. (1). The R2 of the calibration was 0.97 and the uncertainty was greatest at high 15N–N2 concentrations, however experimental data were not in this upper range. The correction was applied to the samples to determine actual concentrations in the gel strips. 15 NH+ 4 was collected using two different techniques. In November 2009, the ADM was used and 15NH+ 4 collected in the acid trap was measured using a gas chromatograph – isotope ratio mass spectrometer coupled to an elemental analyser at 1050 °C (Sigman et al., 1997). In 2013 15NH+ 4 was measured using the hypobromite method (RisgaardPetersen et al., 1995); samples were purged with He to remove background N2 then 60 lL of 12 mol L1 NaOH was added to the 3 mL vial (Exetainer, Labco) containing

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the sample, shaken, then 60 lL of hypobromite added. The samples were shaken for 24 h before analysis to ensure all the NH+ 4 was converted to N2. Both the ADM and hypobromite method achieved P90% recovery of 15NH+ 4 . Isotope analysis of nitrogen was undertaken using either a Shimadzu GCMS-QP5050 (N2 in intact core experiments), or Sercon 20–22 continuous flow isotope ratio mass spectrometer coupled either to a gas chromatograph (N2 in gels) or elemental analyser (ADM samples). 3. RESULTS 3.1. Intact core experiments

N-Denitrification (µmol 15N m-2 h -1)

During the control oxic treatment, mean DNRA was 28 ± 2 lmol m2 h1 and denitrification initially (<7 days) 280 ± 100 lmol m2 h1, decreasing to a mean of 76 ± 14 lmol m2 h1 after day 7 with a mean denitrification:DNRA ratio of 3.2 ± 0.6 after day 7 (Fig. 2). During the hypoxic treatment, however, mean denitrification:DNRA ratio was 130 ± 20 and dropped significantly

1000

A

Oxic

3.2. Nitrate slurry experiments

800

600 400 200 0

15

N-DNRA (µmol m -2 h -1) 15

Denitrification : DNRA

upon re-aeration (5.3 ± 0.5). The ratio increased significantly throughout the hypoxic incubation (Denitrification:DNRA = 8.3(day) + 73.1, R2 = 1.00, n = 3, p < 0.01) owing to a decrease in DNRA activity and subsequent increase in denitrification in comparison to the control treatment (Fig. 2C). Mean denitrification under hypoxic conditions (740 ± 40 lmol m2 h1) decreased significantly when conditions were made oxic (155 ± 12 lmol m2 h1, df = 2, p < 0.05) (Fig. 2A). Mean DNRA under hypoxic conditions (6.7 ± 1.0 lmol m2 h1) increased significantly (5-fold), when the water column was oxygenated (33.1 ± 1.2 lmol m2 h1, df = 2, p < 0.05, t-test) (Fig. 2C). Sedimentary 15NH+ 4 detected after the final sampling date showed relatively small additional pools of ammonium, which we could not confidently ascribe to being either from the final tracer addition or accumulation from previous tracer additions. Assuming that the sediment 15NH+ 4 originates entirely from the final tracer addition, the values depicted in Fig. 2B may underestimate DNRA by 12 lmol m2 h1 under oxic conditions, and 3 lmol m2 h1 under anoxic conditions, however if any of the sedimentary 15NH+ 4 has accumulated, the underestimation may be less than this.

B

50 40 30 20 10

DNRA and denitrification rates both increased to a maximum with increasing NO 3 concentration (Fig. 3). Denitrification increased to a maximum (Vmax) of 43.5 ± 3.3 lmol L-slurry1 h1 whereas DNRA reached a Vmax of 26.4 ± 7.1 lmol L-slurry1 h1 (Fig. 3). The half saturation constant (Km) for denitrification and DNRA 1 were 49.0 ± 10.5 lmol-NO and 85.9 ± 52.2 lmol3 L  1 NO3 L , respectively. The kinetics for both denitrification (R2 = 0.98, p < 0.01) and DNRA (R2 = 0.91, p < 0.05) followed the Michaelis–Menten kinetic model (Fig. 3). DNRA averaged 28 ± 4% of total nitrate reduction over the nitrate concentration range. 3.3. Micro-profiles in intact cores – oxygen treatments

0 200

C

150 Control Oxic Hypoxic Oxic

100 50 0 0

5

10 Day

15

20

Fig. 2. Rates of denitrification and dissimilatory nitrate reduction to ammonium (DNRA) in laboratory incubated cores. The control treatment (d) was oxic for the 21 day period. The treated cores are represented by open shapes showing hypoxic conditions for the first 11 days (h) and subsequently oxic (s). The dotted line represents the change from hypoxic to oxic conditions on day 11 of the treated cores. (A) Denitrification (lmol m2 h1), (B) DNRA (lmol m2 h1) and (C) Denitrification:DNRA ratio over the incubation period.

To investigate the spatial distribution of denitrification and DNRA within the sediment, profiles of the process rates and ancillary variables were undertaken in intact oxic and anoxic cores. For the anoxic treatment, the 15N–N2 concentration profile, indicative of denitrification, displayed a maximum concentration (190 ± 20 lmol L1) at the sediment–water interface and decreased with sediment depth (Fig. 4A). Concentrations of 15N–N2 in the oxic treatments, however, increased with depth, reaching a peak (highest 15N–N2) at 1.0 cm depth below the oxic surface layer of the sediment (Fig. 4E). For the anoxic treatment, the 15NH+ 4 profile, was uniform with sediment depth, with 15 1 NH+ (Fig. 4A). In contrast, 4 averaging 3.4 ± 0.1 lmol L + 15 under oxic conditions, NH4 reached a maximum concentration at 1.2 cm depth, with a concentration of 7.0 ± 2.0 lmol L1 (Fig. 4E). The 15NH+ 4 profile for the oxic treatments followed a similar trend to the 15N–N2 profile, however it was an order of magnitude lower (Fig. 4E). The 15N–N2: 15NH+ 4 ratio was much higher in the anoxic

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1 1 1 Fig. 3. 15N–NO h ) with respect to NO 3 reduction rate (lmol L slurry 3 concentration (lmol L ) measured in anaerobic slurries.  Denitrification (s) and DNRA (d) follow the Michaelis–Menten model increasing with respect to NO3 concentration (p < 0.01 and p < 0.05, respectively). Denitrification:DNRA (N) with respect to 15N -NO 3 concentration.

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1 15 Fig. 4. Micro-profiles of 15NH+ N–N2 (±SE, n = 3) (lmol L1, indicating denitrification); 4 (±SE, n = 3) (lmol L ; indicating DNRA); 15 1  N–N2:15NH+ (indicating denitrification:DNRA) (±SE, n = 3), NO (±SE, n = 3) (lmol L1), NH+ 4 3 4 (±SE, n = 3) (lmol L ), Fetot (±SE, tot 1 2 1 n = 3) (lmol L ), Stot (±SE, n = 3) (lmol L ) and oxygen (±SE, n = 3) (% O2 saturation). (A–D) represents anoxic conditions, and (E–H) oxic conditions. Profiles taken in February 2013.

treatment indicating high denitrification and low DNRA; the maximum denitrification:DNRA was observed at the sediment surface (53 ± 7, Fig. 4B). The 15N–N2: 15NH+ 4 ratio in the oxic treatment averaged 14.4 ± 0.6 in the sedi-

ment and was significantly different from the mean under anoxic conditions (33 ± 2, p  0.001) (Fig. 4F). The NO 3 concentration decreased with sediment depth in the oxic treatment, however it was completely depleted

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for the Fe treatment was 1.24 lmol L-slurry1 h1 (15NH+ 4 = 1.24(h) + 3.46, R2 = 0.95, p < 0.01) (Fig. 5B). The 15NH+ 4 concentration plateaued in both treatments suggesting DNRA ceased after 48 h. Denitrification (illustrated by the 15N–N2 concentration) was comparable to DNRA in the control (15N–N2 = 1.08(h)  0.22, R2 = 1.00, p < 0.001) (Fig. 5A) in the first 48 h and half the rate of 15NH+ 4 production in the Fe treatment (15N–N2 = 0.66(h)  0.31, R2 = 0.99, p < 0.001) (Fig. 5B). After 2 days, the denitrification rate increased 6 and 13-fold in the control and Fe treatment, respectively, becoming the dominant nitrate reduction pathway (Fig 5A and B). NOx consumption was 2.27 lmol L-slurry1 h1 (15NO 3 = 2.27(h) + 877, R2 = 0.89, p < 0.05) in the first 48 h for the control and 2.16 lmol L-slurry1 h1 (15NO 3 = 2.26(h) + 756, R2 = 0.70, p = 0.08) for the Fe treatment (Fig. 5E and D). NOx consumption increased significantly after 48 h, following the denitrification trend in both the control and Fe treatment. The NO 2 concentration spiked before denitrification increased, reaching 75% of total NOx at day 3 and 4 for the control and Fe treatment, respectively. Total Fe2+ (HCl extractable Fe2+ + dissolved Fe2+) decreased in the first 48 h at a rate of 18.8 lmol L-slurry1h1 and 7.5 lmol L-slurry1 h1 for the Fe (Total Fe2+ = 19(h) + 3974, R2 = 0.82, p < 0.05) and control (Total

at the surface in the anoxic treatment. The total NH+ 4 concentration increased with sediment depth in all treatments. Free sulfide was observed under anoxic conditions and increased with depth reaching a maximum of 695 lmol L1 at 2.5 cm; sulfide was not observed at high concentrations in the water column (Fig. 4D). In the oxic treatment, total sulfide was <40 lmol L1 (Fig. 4H). Total Fe averaged 8.2 ± 0.9 lmol L1 in the anoxic treatment (Fig. 4D). In contrast, the total Fe concentration in the oxic treatment reached a maximum of 500 ± 200 lmol L1 at 1.2 cm (Fig. 4H). The oxygen concentration was <1% O2 saturation in the anoxic treatment in both the water column and the sediment (Fig. 4D). The oxygen in the oxic treatment reached 92% O2 saturation in the water column and penetrated 0.3 cm into the sediment (Fig. 4H). 3.4. Iron slurry experiments The 15NH+ 4 production in both the control and Fe treatment were comparable, reaching maximum concentrations of 55 and 60 lmol L-slurry1 after 48 h, respectively (Fig. 5). The rate of 15NH+ 4 production over the first 48 h for the control was 1.15 lmol L-slurry1 h1 (15NH+ 4 = 1.15(h) + 2.45, R2 = 0.97, p < 0.01) (Fig. 5A) and the rate

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Fig. 5. (A and B) (C and D) and total Fe (dissolved + 0.5 M HCl extractable, E and F) (lmol L-Slurry1) over time in anoxic slurries of sediment (5% wet sediment/water) with (right panels) and without (left panels) the addition of Fe2+. The dotted line 15 N–N2 production and NOx and Fe2+ consumption were determined. represents the point at which rates of 15NH+ 4, 15 NH+ N–N2 4,

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Fe2+ = 7.5(h) + 2641, R2 = 0.55, p > 0.1) treatments respectively (Fig. 5E and F). For the Fe treatment, the total Fe2+ increased after 48 h at a rate of 20 lmol L-slurry1 h1 (Total Fe2+ = 7.9 (h) + 2998, R2 = 0.63, p = 0.05), which corresponded to the cessation of DNRA activity, however, for the control treatment, HCl extractable Fe2+ plateaued. 4. DISCUSSION 4.1. Methodological considerations In this study we applied a variation of a diffusional equi15 librium gel technique to sample 15NH+ N–N2 based 4 and on the previous work of Stief et al. (2010) and Kessler et al. (2013) respectively. This approach gave us new insight into the relative rates of denitrification and DNRA with depth in the sediment. However, it is important to recognise that this approach leads to smearing of the profiles within the gel (Harper et al., 1997). Here, this is exemplified by the fact that the profiles are relatively broad with no sharp inflections at the oxic/anoxic boundary as one would expect for denitrification (Stief et al., 2010). Moreover, 15NH+ 4, 29 N2 and 30N2 are detected several millimetres below the depth of NO 3 penetration. Therefore, this approach can only be used to gain information on the relative concentrations of isotopically labelled NH+ 4 and N2 with depth, and hence, the relative rates of DNRA and denitrification, which was the aim of this study. Theoretically it would be possible to use an inverse modelling approach to derive rate profiles in a similar manner to the well established PROFILE routine of Berg et al. (1998). To do this would require the addition of a non-steady state component to the model to account for incubation time, NH+ 4 sorption to the sediment as well as accounting for the different tortuosities of the gel and sediment, which was beyond the scope of this study. In slurries, the accumulation and decomposition of NO 2 after NO 3 addition can lead to artefacts when interpreting trends in Fe2+ consumption. Under acidic conditions, NO 2 decomposes to HNO2 and further to NO2 and NO which are powerful oxidants and can readily re-oxidise Fe2+ to insoluble Fe3+ (Weber et al., 2001; Klueglein and Kappler, 2013). This problem was avoided by centrifuging the sample when conditions were anoxic and the supernatant immediately fixed with ferrozine. The remaining sediment sample was HCl extracted and the concentrations of NO 2 in the extractant over the first 48 h were <0.7 and 0.5 lmol L1 in the control and iron treatments, respectively, and over 1 the total incubation NO 2 did not exceed 5 lmol L . In 2+ addition, no relationship was observed between Fe and NO 2 , therefore, we do not believe the decomposition of 2+ NO is an artefact in this 2 and subsequent oxidation of Fe study. 4.2. DNRA under changing oxygen conditions Using a range of approaches we observed that DNRA decreases under anoxic conditions relative to denitrification. Under prolonged water column hypoxia, the denitrifi-

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cation:DNRA ratio increased, driven by high denitrification rates combined with low DNRA rates resulting in DNRA comprising <1% of total nitrate reduction. Re-oxygenation of the water column decreased the denitrification:DNRA ratio and DNRA represented 18% of total nitrate reduction (Fig. 2). Intact sediment profiles of denitrification and DNRA were also in agreement with this and showed the denitrification:DNRA ratio averaged 14.4 ± 0.6 under oxic conditions in the water column, and was significantly lower than the average denitrification:DNRA ratio under anoxia (33.1 ± 2.1). These results support previous observations in the Yarra River Estuary which showed that denitrification was the main nitrate reduction pathway under hypoxia within the water column (Roberts et al., 2012). In that study DNRA became more prevalent, increasing from <1% of total nitrate reduction under anoxia to 50% under oxic conditions in the water column. High rates of DNRA after re-oxygenation of the water column were unexpected because DNRA has typically been reported to occur to a greater extent under sulfidic conditions (Brunet and Garcia-Gil, 1996; Christensen et al., 2000; An and Gardner, 2002; Childs et al., 2002) which would be greater under anoxia. We have two possible explanations for this behaviour based on the profile data, 2+ which showed the highest concentrations of NO 3 and Fe + 15 coincided with the highest concentration of NH4 (implying high DNRA, Fig. 4). 1. Differing reaction kinetics (Vmax and Km) mean that the relative rates of denitrification and DNRA change in relation to NO 3 concentrations, which change under oxic and anoxic conditions. 2. DNRA is linked to Fe2+ oxidation. We now discuss the plausibility of these explanations based on the results of the slurry incubations. 4.3. The influence of NO 3 concentration on DNRA Both denitrification and DNRA followed similar Michaelis–Menten kinetics with respect to NO 3 concentration. Only a weak negative correlation (p = 0.22) was observed between the denitrification:DNRA ratio and increasing NO 3 concentration (Fig. 3, grey squares), however this is not strong enough to be considered the primary factor regulating DNRA in the Yarra River Estuary and does not explain the very low denitrification:DNRA ratios observed under oxic conditions in intact cores (Fig. 2). Several studies have observed the highest rates of DNRA when NO 3 is limiting (King and Nedwell, 1985; Brunet and Garcia-Gil, 1996; Childs et al., 2002; Nizzoli et al., 2010). For example, Childs et al. (2002) found diminished rates of denitrification in the Gulf of Mexico and hypothesised the decrease was due to a competitive advantage of DNRA under the hypoxic, NO 3 -limited conditions. Observations of DNRA in the Yarra River Estuary by contrast showed the highest rates of DNRA occurred when conditions were oxic in the water column and NO 3 was present at moderate concentrations. Similarly, in the tropical Cisadane estuary, Indonesia, DNRA increased 10-fold greater than denitrification when there was a surplus of NO 3 , however this was explained by the higher affinity of

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nitrate ammonifiers for NO 3 at high temperatures (Ogilvie et al., 1997; Dong et al., 2011). Unlike the Cisadane estuary, the Yarra River Estuary is a temperate estuary and DNRA was observed over a range of temperatures (14–22 °C, temperature range from rates measured in Roberts et al. (2012)). The concentration of NO is typically 3 <30 lmol L1 in the bottom waters of the Yarra River Estuary, however, to account for influxes of NO 3 during high flow events (when DNRA was observed) and nitrification within the sediment, the NO 3 concentration range tested in incubation experiments was 10 to 240 lmol L1.

although several studies have identified the role of sulfide in DNRA, we did not observe high DNRA rates in the presence of sulfide (Fig. 4) (Brunet and Garcia-Gil, 1996; An and Gardner, 2002). The addition of NO 3 to estuarine slurries resulted in the immediate onset of DNRA which was the dominant process for the first 48 h of the incubation. In the period of DNRA activity, total Fe2+ (dissolved + HCl extractable) decreased significantly in the Fe2+ treatment (p < 0.05) however, this trend was not significant in the control (no Fe2+ added) treatment, however, there was a modest R2 of 0.55 and the lack of significance is primarily driven by small sample size as opposed to high variability. Upon ces2+ sation of NH+ concentrations changed 4 production, Fe little, or increased slightly in the control and Fe treatment, respectively (Fig. 5E and F). The ratios of Fe2+ consumption to NH+ 4 production in the slurries were 1:15 ± 6 and 1:7 ± 3 for the control and iron treatments, respectively which is close within uncertainty to the expected stoichiometry of 1:8 (Eq. (2)), supporting the plausibility of the hypothesis that Fe2+ plays a role in NH+ 4 production.

4.4. Is DNRA linked to Fe2+ oxidation? Enzymatic nitrate dependent Fe2+ oxidation is well documented, with the main product being N2 gas, produced via denitrification (Nealson and Saffarini, 1994; Straub et al., 1996; Benz et al., 1998) or the more recently described iron coupled anammox process (Oshiki et al., 2013). Several studies have linked both abiotic (Hansen et al., 1996, 2001), and biotic (Weber et al., 2006; Coby et al., 2011) nitrate dependent, Fe2+ oxidation to the production of NH+ 4. We note, however, these studies were performed in slurries or culture experiments and the environmental significance of Fe driven DNRA remains unknown. Under oxic conditions in the water column, the sediment profiles of 15NH+ 4 coincided with the presence of high concentrations of Fe2+ in the sediment (Fig. 4). Under long-term anoxic conditions in the water column, the concentration of free sulfides increased to 695 lmol L1 and Fe2+ decreased to <20 lmol L1 (Fig. 4). High sulfide concentrations were characteristic of anoxic conditions and

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Fig. 6. Conceptual model of sediment nitrogen cycling under (A) oxic and (B) anoxic conditions, highlighting the links between the nitrogen, iron and sulfur cycles. (a) nitrification, (b) denitrification, (c) DNRA, (d) carbon oxidation (e) Fe2+ oxidation, (f) Fe2+ reduction and (g) FeS precipitation. Dashed lines represent diffusion and width of line represents the relative contribution of each process under oxic and anoxic conditions.

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(Fig. 5). The cause of this shift from DNRA to denitrification remains uncertain; however we speculate that the significant disruption of the sediment caused by slurrying led to a rapid and unpredictable shift in the microbial community. In this case we speculate that turnover of the microbial community initially present led to a release of labile organic carbon, which stimulated denitrification. The presence of DNRA under oxic rather than anoxic conditions can be explained by the availability of Fe2+ in the sediment (Fig. 6). Under oxic conditions the reduction of Fe3+ at the oxic-anoxic boundary supplies Fe2+ to the anoxic zone, as shown in the conceptual model (Fig. 6). Under these conditions denitrification and DNRA occur at similar rates and the removal of NO 3 as N2 is diminished due to the increase in DNRA. However, under anoxic conditions the formation of free sulfides within the sediment will rapidly bind with available Fe2+ to form iron sulfide precipitates (FeS) decreasing the bioavailability of Fe2+, a mechanism which is clear in the profiles of sulfide and Fe2+ (Figs. 4 and 6). While this study suggests Fe driven DNRA may be significant in the Yarra Estuary, the global importance of this process remains uncertain. We postulate that it may be significant in non-sulfidic sediments such as freshwater lakes and estuarine systems which accumulate large amounts of iron through coagulation and sedimentation of colloidal material (Hart and Davies, 1981). In large rivers such as the Amazon and Mississippi, significant amounts of Fe may be deposited on the continental shelf, leading to high rates of Fe reduction and Fe2+ accumulation within the porewaters (Aller et al., 1986; Sholkovitz, 1993; Powell and Wilson-Finelli, 2003). There appears to be circumstantial evidence in the literature to support this hypothesis. In Copano Bay, Texas, iron oxide concentrations were significantly correlated with 15NH+ 4 production (Hou et al., 2012). In contrast to this, however, another recent study found no correlation with DNRA and Fe dissolved in the porewater (Behrendt et al., 2013). The plausibility and widespread occurrence of Fe driven DNRA therefore remains uncertain and requires further investigation. ACKNOWLEDGMENTS We are grateful to Scotch College for providing private boat ramp access. We thank E. Richmond, K. Browne, V. Eate, T. Scicluna, and M. Bourke for assistance in the field and laboratory. K. Browne and V. Evrard undertook the isotope ratio mass spectrometry analyses. We would like to thank R. Roberts and H. Sarkissian for their help with the design and assembly of the gel cutting device. We appreciate the helpful advice of B. Thamdrup, R.N. Glud and E. Robertson. We would like to acknowledge the support of the Australian Research Council (LP0991254), Melbourne Water Corporation and the Environment Protection Authority Victoria. A. Kessler was supported by a Sir James McNeill Postgraduate Research Scholarship. We thank 3 anonymous reviewers whose constructive comments greatly helped improve the manuscript. REFERENCES Aller R. C., Mackin J. E. and Cox, Jr., R. T. (1986) Diagenesis of Fe and S in Amazon inner shelf muds: apparent dominance of Fe reduction and implications for the genesis of ironstones. Sediment. Proc. Amazon Cont. Shelf 6, 263–289.

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An S. and Gardner W. S. (2002) Dissimilatory nitrate reduction to ammonium (DNRA) as a nitrogen link, versus denitrification as a sink in a shallow estuary (Laguna Madre/Baffin Bay, Texas). Mar. Ecol. Prog. Ser. 237, 41–50. APHA (2005) APHA-AWWA-WPCF, Standard Methods for the Examination of Water and Wastewater, 20th ed. American Public Health Association, American Water Works Association and Water Environment Federation, Washington. Behrendt A., de Beer D. and Stief P. (2013) Vertical activity distribution of dissimilatory nitrate reduction in coastal marine sediments. Biogeosciences 10, 7509–7523. Benz M., Brune A. and Schink B. (1998) Anaerobic and aerobic oxidation of ferrous itron at neutral pH by chemoheterotrophic nitrate reducing bacteria. Arch. Microbiol. 169, 159–165. Berg P., Risgaard-Petersen N. and Rysgaard S. (1998) Interpretation of measured concentration profiles in sediment pore water. Limnol. Oceanogr. 43, 1500–1510. Bonin P. (1996) Anaerobic nitrate reduction to ammonium in two strains isolated from coastal marine sediment: a dissimilatory pathway. FEMS Microbiol. Ecol. 19, 27–38. Brunet R. C. and Garcia-Gil L. J. (1996) Sulfide-induced dissimilatory nitrate reduction to ammonia in anaerobic freshwater sediments. FEMS Microbiol. Ecol. 138, 131–138. Childs C. R., Rabalais N. N., Turner R. E. and Proctor L. M. (2002) Sediment denitrification in the Gulf of Mexico zone of hypoxia. Mar. Ecol. Prog. Ser. 240, 285–290. Christensen P. B., Rysgaard S., Sloth N. P., Dalsgaard T. and Schwaerter S. (2000) Sediment mineralization, nutrient fluxes, denitrification and dissimilatory nitrate reduction to ammonium in an estaurine fjord and sea cage trout farms. Aquat. Microb. Ecol. 21, 73–84. Coby A. J., Picardal F., Shelobolina E., Xu H. F. and Roden E. E. (2011) Repeated anaerobic microbial redox cycling of iron. Appl. Environ. Microbiol. 77, 6036–6042. Cole J. A. and Brown C. M. (1980) Nitrite reduction to ammonia by fermentative bacteria: a short circuit in the biological nitrogen cycle. FEMS Microbiol. Lett. 7, 65–72. Conley D. J., Carstensen J., Aertebjerg G., Christensen P. B., Dalsgaard T., Hansen J. L. S. and Josefson A. B. (2007) Longterm changes and impacts of hypoxia in Danish coastal waters. Ecol. Appl. 17, S165–S184. Dalsgaard T., Nielsen L. P., Brotas V., Viaroli P., Underwood G. J. C., Nedwell D. B., Sundback K., Rysgaard S., Miles A., Bartoli M., Dong L. F., Thornton D. C. O., Ottosen L. D. M., Castaldelli G. and Risgaard-Petersen N. (2000) Protocol Handbook for NICE – Nitrogen Cycling in Estuaries: A Project under the EU Research Programme: Marine Science and Technology (MAST III). Ministry of Environment and Energy, National Environmental Research Institute, Department of Lake and Estuarine Ecology, Denmark. Diaz R. J. and Rosenburg R. (2008) Spreading dead zones and consequences for marine ecosystems. Science 321, 926– 929. Dong L. F., Sobery M. N., Smith C. J., Rusmana I., Phillips W., Stott A., Osborn A. M. and Nedwell D. B. (2011) Dissimilatory reduction of nitrate to ammonium, not denitrification dominates benthic nitrate reduction in tropical estuaries. Limnol. Oceanogr. 56, 279–291. Fazzolari E., Nicolardot B. and Germon J. C. (1998) Simultaneous effects of increasing levels of glucose and oxygen partial pressures on denitrification and dissimilatory nitrate reduction to ammonium in repacked soil cores. Eur. J. Soil Biol. 34, 47–52. Hansen H. C. B., Koch C. B., Nancke-Krogh H., Borggaard O. K. and Sorensen J. (1996) Abiotic nitrate reduction to ammonium: key role of green rust. Environ. Sci. Technol. 30, 2053–2056.

324

K.L. Roberts et al. / Geochimica et Cosmochimica Acta 133 (2014) 313–324

Hansen H. C. B., Guldberg S., Erbs M. and Koch C. B. (2001) Kinetics of nitrate reduction by green rusts – effects of interlayer anion and Fe(II):Fe(III) ratio. Appl. Clay Sci. 18, 81–91. Harper M. P., Davison W. and Tych W. (1997) Temporal, spatial, and resolution constraints for in situ sampling devices using diffusional equilibration: dialysis and DET. Environ. Sci. Technol. 31, 3110–3119. Hart B. T. and Davies H. R. (1981) Trace metal speciation in the freshwater and estuarine regions of the Yarra River, Victoria. Estuar. Coast. Shelf Sci. 12, 353–374. Hou L., Liu M., Carini S. A. and Gardner W. S. (2012) Transformation and fate of nitrate near the sediment–water interface of Copano Bay. Cont. Shelf Res. 35, 86–94. Jorgensen K. S. (1989) Annual pattern of denitrification and nitrate ammonification in estuarine sediment. Appl. Environ. Microbiol. 55, 1841–1847. Jorgensen K. S. and Sorensen J. (1988) Two annual maxima of nitrate reduction and denitrification in estuarine sediment (Norsminde Fjord, Denmark). Mar. Ecol. Prog. Ser. 48, 147–154. Kemp W. M., Boynton W. R., Adolf J. E., Boesch D. F., Boicourt W. C., Brush G., Cornwell J. C., Fisher T. R., Glibert P. M., Hagy J. D., Harding L. W., Houde E. D., Kimmel D. G., Miller W. D., Newell R. I. E., Roman M. R., Smith E. M. and Stevenson J. C. (2005) Eutrophication of Chesapeake Bay: historical trends and ecological interactions. Mar. Ecol. Prog. Ser. 303, 1–29. Kessler A. J., Glud R. N., Cardenas M. B. and Cook P. L. M. (2013) Transport zonation limits coupled nitrification–denitrification in permeable sediments. Environ. Sci. Technol. 47, 13404–13411. King D. and Nedwell D. B. (1984) Changes in the nitrate-reducing community of an anaerobic saltmarsh sediment in response to seasonal selection by temperature. J. Gen. Microbiol. 130, 2935– 2941. King D. and Nedwell D. B. (1985) The influence of nitrate concentration upon the end-products of nitrate dissimilation by bacteria in anaerobic salt marsh sediment. FEMS Microbiol. Ecol. 31, 23–28. Klueglein N. and Kappler A. (2013) Abiotic oxidation of Fe(II) by reactive nitrogen species in cultures of the nitrate-reducing Fe(II) oxidizer Acidovorax sp. BoFeN1 – questioning the existence of enzymatic Fe(II) oxidation. Geobiology 11, 180– 190. Krom M. D., Davison P., Zhang H. and Davison W. (1994) Highresolution pore-water sampling with a gel sampler. Limnol. Oceanogr. 39, 1967–1972. Nealson K. H. and Saffarini D. (1994) Iron and manganese in anaerobic respiration: environmental significance, physiology, and regulation. Annu. Rev. Microbiol. 48, 311–343. Nizzoli D., Carraro E., Nigro V. and Viaroli P. (2010) Effect of organic enrichment and thermal regime on denitrification and dissimilatory nitrate reduction to ammonium (DNRA) in hypolimnetic sediments of two lowland lakes. Water Res. 44, 2715–2724. Ogilvie B. G., Rutter M. and Nedwell D. B. (1997) Selection by temperature of nitrate-reducing bacteria from estuarine sediments: species composition and competition for nitrate. FEMS Microbiol. Ecol. 23, 11–22. Oshiki M., Ishii S., Yoshida K., Fujii N., Ishiguro M., Satoh H. and Okabe S. (2013) Nitrate-dependent ferrous iron oxidation by anaerobic ammonium oxidation (anammox) bacteria. Appl. Environ. Microbiol. 79, 4087–4093. Otte S., Kuenen J. G., Nielsen L. P., Paerl H. W., Zopfi J., Schulz H., Teske A., Strotmann B., Gallardo V. A. and Jorgensen B. B. (1999) Nitrogen, carbon, and sulfur metabolism in natural Thioploca samples. Appl. Environ. Microbiol. 65, 3148–3157.

Powell R. T. and Wilson-Finelli A. (2003) Importance of organic Fe complexing ligands in the Mississippi River plume. Estuar. Coast. Shelf Sci. 58, 757–763. Risgaard-Petersen N., Rysgaard S. and Revsbech N. P. (1995) Combined microdiffusion-hypobromite oxidation method for determining N-15 isotope in ammonium. Soil Sci. Soc. Am. J. 59, 1077–1080. Roberts K. L., Eate V. M., Eyre B. D., Holland D. P. and Cook P. L. M. (2012) Hypoxic events stimulate nitrogen recycling in a shallow salt-wedge estuary: the Yarra River estuary, Australia. Limnol. Oceanogr. 57, 1427–1442. Rysgaard S., Risgaard-Peterson N. and Sloth N. P. (1996) Nitrification, denitrification and nitrate ammonification in sediments of two coastal lagoons in Southern France. Hydrobiologia 329, 133–141. Sayama M., Risgaard-Petersen N., Nielsen L. P., Fossing H. and Christensen P. B. (2005) Impact of bacterial NO 3 transport on sediment biogeochemistry. Appl. Environ. Microbiol. 71, 7575– 7577. Schulz H. N. and Jorgensen B. B. (2001) Big bacteria. Annu. Rev. Microbiol. 55, 105–137. Sholkovitz E. R. (1993) The geochemistry of rare earth elements in the Amazon River estuary. Geochim. Cosmochim. Acta 57, 2181–2190. Sigman D. M., Altabct M. A., Michener R., McCorkle D. C., Fry B. and Holmes R. M. (1997) Natural abundance-level measurement of the nitrogen isotopic composition of oceanic nitrate: an adaptation of the ammonia diffusion method. Mar. Chem. 57, 227–242. Stief P., Behrendt A., Lavik G. and De Beer D. (2010) Combined gel probe and isotope labeling technique for measuring dissimilatory nitrate reduction to ammonium in sediments at millimeter-level resolution. Appl. Environ. Microbiol. 76, 6239–6247. Stookey L. L. (1970) Ferrozine – a new spectrophotometric reagent for iron. Anal. Chem. 42, 779–781. Straub K. L., Benz M., Shinck B. and Widdel F. (1996) Anaerobic, nitrate-dependent microbial oxidation by ferrous iron. Appl. Environ. Microbiol. 62, 1458–1460. Strohm T. O., Griffin B., Zumft W. G. and Schink B. (2007) Growth yields in bacterial denitrification and nitrate ammonification. Appl. Environ. Microbiol. 73, 1420–1424. Tiedje J. M. (1988) Ecology of denitrification and dissimilatory nitrate reduction to ammonium. In Biology of Anaerobic Microorganisms (ed. A. J. B. Zehnder). Wiley, New York, pp. 179–244. Tison D. L., Pope D. H. and Boylen C. W. (1980) Influence of seasonal temperature on the temperature optima of bacteria in sediments of Lake George, New York. Appl. Environ. Microbiol. 39, 675–677. Voillier E., Inglett P. W., Hunter K., Roychoudhury A. N. and Van Cappellen P. (2000) The ferrozine method revisited: Fe(II)/ Fe(III) determination in natural waters. Appl. Geochem. 15, 785–790. Weber K. A., Picardal F. W. and Roden E. E. (2001) Microbially catalyzed nitrate-dependent oxidation of biogenic solid-phase Fe(II) compounds. Environ. Sci. Technol. 35, 1644–1650. Weber K. A., Urrutia M. M., Chruchill P. F., Kukkadapu R. V. and Roden E. E. (2006) Anaerobic redox cycling of iron by freshwater sediment microorganisms. Environ. Microbiol. 8, 100–113. Yin S. X., Chen D., Chen L. M. and Edis R. (2002) Dissimilatory nitrate reduction to ammonium and responsible microorganisms in two Chinese and Australian paddy soils. Soil Biol. Biogeochem. 34, 1131–1137. Associate editor: Jack J. Middelburg