Effects of Ferrous Iron and Hydrogen Sulfide on Nitrate Reduction in the Sediments of an Estuary Experiencing Hypoxia

Hypoxia is common feature of eutrophic estuaries and semi-enclosed seas globally. One of the key factors driving hypoxia is nitrogen pollution. To gain more insight into the effects of hypoxia on estuarine nitrogen cycling, we measured potential nitrate reduction rates at different salinities and levels of hypoxia in a eutrophic temperate microtidal estuary, the Neuse River Estuary, North Carolina, USA. We also tested the effect of hydrogen sulfide and ferrous iron additions on the nitrate reduction pathways. Overall, DNRA dominated over denitrification in this periodically hypoxic estuary and there was no correlation between the potential nitrate reduction rates, salinity, or dissolved oxygen. However, when hypoxia lasted several months, denitrification capacity was almost completely lost, and nearly all nitrate added to the sediment was reduced via DNRA. Additions of hydrogen sulfide stimulated DNRA over denitrification. Additions of ferrous iron stimulated nitrate consumption; however, the end product of nitrate consumption was not clear. Interestingly, substantial nitrous oxide formation occurred in sediments that had experienced prolonged hypoxia and were amended with nitrate. Given expanding hypoxia predicted with climate change scenarios and the increasing nitrate loads to coastal systems, coastal sediments may lose their capability to mitigate nitrogen pollution due to DNRA dominating over denitrification during extended hypoxic periods.


Introduction
The amount of reactive nitrogen (N) in the environment has increased dramatically during the past 150 years, which has promoted eutrophication of coastal waters (Paerl and Piehler 2008). One of the most severe symptoms of eutrophication is hypoxia (oxygen (O 2 ) 90 μmol l −1 at STP) Rosenberg 1995, 2008;Breitburg et al. 2018), which alters biogeochemical cycling of numerous key elements (e.g., carbon, nitrogen, sulfur, phosphorus).
Microbes in coastal sediments provide an important ecosystem service by converting reactive N through microbial pathways to dinitrogen (N 2 ) and nitrous oxide (N 2 O) gases (Seitzinger 1988;Dalsgaard et al. 2005). It has been estimated that globally, coastal and shelf areas remove approximately of 25% of the total fluvial reactive N input (Sharples et al. 2017). However, under hypoxic conditions, N removal in coastal sediments decreases, since nitrate (NO 3 − ) is also reduced to ammonium (NH 4 + ), instead of to N 2 and N 2 O, via the dissimilatory nitrate reduction to ammonia (DNRA) pathway (An and Gardner 2002;Gardner et al. 2006;Dong et al. 2011). DNRA can be performed by fermentative heterotrophic organisms, which use organic carbon as the electron donor and by chemolithoautotrophic organisms, which use reduced inorganic compounds as an electron donor (Giblin et al. 2010). In general, fermentative processes are slow and

Communicated by Marco Bartoli
Electronic supplementary material The online version of this article (https://doi.org/10.1007/s12237-020-00783-4) contains supplementary material, which is available to authorized users. fermentative DNRA cannot compete with dentification for NO 3 − unless the carbon the NO 3 − ratio is very high (Kraft et al. 2014). Chemolithoautotrophic DNRA in the other hand is not dependent on the availability of organic carbon (An and Gardner 2002;Gardner et al. 2006;Dong et al. 2011). Under benthic anoxia, sedimentary nitrification, which typically provides most substrate for the NO 3 − reducing processes (i.e., Hietanen and Kuparinen 2008), ceases and thus DNRA rates are also inhibited due to the low availability of electron acceptors. Hence, hypoxia can lead to a vicious cycle of eutrophication where the excess bioavailable N builds up, preventing recovery from this process (Jäntti and Hietanen 2012). Sulfate (SO 4 2− ) is highly available in marine ecosystems (Canfield 1989 and references therein) and a common consequence of hypoxia is the accumulation of toxic hydrogen sulfide (H 2 S) in the bottom water. H 2 S is produced when, in the absence of O 2 , organic matter is oxidized with SO 4 2− (Middelburg and Levin 2009). H 2 S affects sedimentary N cycling by inhibiting nitrification and anammox (Joye and Hollibaugh 1995;Hannig et al. 2007), while stimulating chemolithotrophic denitrification and DNRA (Brettar and Rheinheimer 1991;An and Gardner 2002;Gardner et al. 2006;Dong et al. 2011). Hence, hypoxia creates favorable conditions for DNRA; however, the presence of this process is still poorly quantified in estuarine systems.
H 2 S is often bound to iron (Fe) compounds, which can also be abundant in estuaries and coastal areas. The interactions between H 2 S, Fe, and various forms of N are currently poorly understood. The reactions between NO 3 − , nitrite (NO 2 − ), and reduced Fe, which regulate the bioavailability of N, were initially considered to be mainly abiotic (Moraghan and Buresh 1977;Buresh and Moraghan 1976). However, Straub et al. (1996) showed that Fe oxidation with NO 3 − can also be microbially mediated and since then the number of microbial taxa known to be capable for anaerobic Fe 2+ oxidation with NO 3 − under dark circumneutral conditions has increased substantially (Weber et al. 2006). The magnitude of microbial Fe oxidation with NO 3 − has, however, been questioned because of inappropriate analysis of N and Fe intermediates which can result in overestimated rates (Picardal 2012;Klueglein and Kappler 2013). Overall, the significance of Fe determining the relative importance of denitrification and DNRA in natural environments is currently poorly constrained. The purpose of this study was to investigate the fate of NO 3 − in a eutrophic, seasonally hypoxic, N limited estuary.
More specifically, we wanted to investigate how salinity and O 2 concentration affect the NO 3 − reduction pathways and how elevated availabilities of reduced S and Fe species affect the interplay between the different NO 3 − reducing pathways under anoxic conditions.

Sampling
Samples for the sediment incubations were collected from the Neuse River Estuary (NRE). The NRE is the largest subestuary of the lagoonal Albemarle-Pamlico Sound, which is the second largest estuarine complex in the lower USA. The average depth of the NRE is~3.5 m and the astronomical tidal range is < 0.1 m (Luettich Jr. et al. 2000). Sediment composition in the NRE changes from sandy at the shoreline to muddy in mid-channel. Its watershed drains rapidly expanding agricultural (animal and row crop operations), urban (Raleigh-Durham Research Triangle), and industrial centers within the piedmont and coastal plain regions of North Carolina. The NRE has over the past several decades undergone anthropogenic eutrophication leading to increased phytoplankton production and bloom frequency (Paerl et al. 2010). The saltwater wedge extending from the downstream Pamlico Sound up into the estuary naturally decreases ventilation of the benthic waters and resultant hypoxia is a seasonal (spring through fall) feature of this estuary (Paerl et al. 1998;Fear et al. 2005 ) and NH 4 + concentrations in the bottom water were determined by withdrawing a water sample from approximately 5 cm above the sediment surface in a core. The NO x − and NH 4 + samples were filtered (prewashed 0.8/0.2-μm double filter syringe, Sarsted, Nümbrecht, Germany) and frozen at − 20°C for subsequent flow-injection nutrient analysis (Lachat instruments, Loveland, CO, USA). Dissolved O 2 concentration was measured using the Winkler method (Grasshoff et al. 1999).

Intact Core Incubations
For the potential NO 3 − reduction measurements, each large core was sub-sampled using three or four smaller acryl plastic cores (diameter 2.5 cm, height 18 cm), so that there was approximately 5 cm of sediment and 13 cm of water in each subsample leaving no headspace in the cores. The sediment samples from different casts were randomized for the treatments in which they were enriched with potassium nitrate (K 15  NH 4 + concentrations. The capped cores were incubated in darkness for 3-4 h at in situ temperature with magnetic stirring bars placed in the caps. After incubation, the sediment was mixed with the overlying water and allowed to stand until most sediment particles had settled (typically 5-15 min). A 12-ml water sample for isotopic analysis of N 2 and N 2 O was withdrawn from the top of the core into a gas tight glass vial (Exetainer, Labco Scientific, High Wycombe, UK) containing 0.5 ml ZnCl 2 (100% w/v). The remaining core content was centrifuged (1500 rpm, 10 min) and the supernatant was filtered (prewashed 0.8/0.2-μm double filter syringe, Sarsted, Nümbrecht, Germany) and frozen at − 20°C for subsequent and NH 4 + and 15 NH 4 + analyses.

Slurry Incubations
To determine the effect of the availability of different electron donors on NO 3 − reduction rates, sediment-water slurries were prepared in May 2016 from sediment and water collected at St70. The slurry for the incubation experiments was comprised of the top 1 cm in the sediment cores and water collected directly above the sediment. The water and sediment were mixed at a 1:1 (vol:vol) ratio and purged with N 2 for 15 min to remove O 2 and background CO 2 and H 2 S gases. Purging of the slurry was likely to increase the sample pH by 1-2 units due to CO 2 leaving the sample. However, St70 experiences pH changes in that range when the saltwater wedge is intruding in the estuary; thus, the microbes present in the sample are likely to experience similar changes naturally. After purging, the slurry was transferred into a glove bag and handled under a N 2 atmosphere until the incubation vials were capped. The slurry was divided into for four 1-l bottles for the following additions in the final concentration: (1) 1000 μM Na 2 S (H 2 S treatment); (2) 1000 μM Fe(II)SO 4 (Fe 2+ treatment); (3) 1000 μM Na 2 S + 1000 μM Fe(II)SO 4 (H 2 S + Fe 2+ treatment); (4) no additional electron donors (control treatment). All treatments received approximately 100 μM K 15 NO 3 − . The treatment solutions were prepared in N 2 purged water and the Fe(II)SO 4 addition was verified to reduce the pH of the slurries by less than 0.5. The slurry from each treatment bottle was divided into 32 12-ml gas tight glass vials (Exetainer, Labco Scientific, High Wycombe, UK), resulting in a total of 128 samples. After preparation, the samples were incubated for approximately 6 h. Then four samples from each treatment were terminated for 15 N 2 and 15 N 2 O concentration measurements by creating a 4-ml He headspace and adding 100 μl of ZnCl 2 (100% w/v) to each sample. Simultaneously, incubations of four samples from each treatment were terminated by filtering the sample through prewashed 0.8/0.2-μM double filter syringes (Acrodisc, Pall Scientific, New York, NY, USA). Thereafter, samples were terminated every 6 h, until 24 h of incubation. The filtrate was analyzed within few hours for NO 3 − and NO 2 − , (Fawcett and Scott 1960;Miranda et al. 2001) and the rest of the samples were frozen at − 20°C and analyzed later for NH 4 + and 15 NH 4 + concentrations. For the 15 N 2 and 15 N 2 O gas samples, a 4 ml helium (He) headspace was created immediately before analysis and samples withdrawn from the headspace were analyzed with a ThermoScientific GasBench + Precon gas concentration system interfaced to a ThermoScientific Delta V Plus isotoperatio mass spectrometer (ThermoScientific, Bremen, Germany) at the University of California Davis Stable Isotope Laboratory (Davis, CA, USA) (May 2016) and with Isoprime100 IRMS coupled to an Isoprime TraceGas preconcentration unit (Elemental Analysensysteme GmbH, Langenselbold, Germany) at the University of Jyväskylä (Jyväskylä, Finland) (October 2016 and June 2017). The 15 NH 4 + analysis was modified from Sigman et al. (1997) and Holmes et al. (1998). First, NH 4 + concentrations of the samples were determined according Fawcett and Scott (1960) to ensure that a minimum of 10 μM NH 4 + was available for the extraction. Then, 10 ml of sample was placed in a 20-ml HDPE scintillation vial (Wheaton, Millville, NJ, USA) and the salinity of the samples was adjusted to 30 with sodium chloride (NaCl). Thereafter, 100 mg of magnesium oxide (MgO) was added to each sample. pH was measured after the addition of MgO to ensure that it was optimal (~10) pH for the conversion of NH 4 + to NH 3 . The liberated NH 3 was collected in diffusion packets which were constructed by pipetting 30 μl of 2.5 M potassium bisulfate (KHSO 4 ) on to a fiberglass filter (Whatman, GF/D, diameter 5 mm, Whatman, Maidstone, Kent, UK) that was placed between two pieces of Teflon tape. The diffusion packets were added prior to MgO addition to minimize NH 3 escaping before closing the vials. The vials were incubated for 3 days at + 37°C on a shaker table (150 rpm). Then, the diffusion packets were removed from the vials and placed in a desiccator under a sulfuric acid (H 2 SO 4 ) atmosphere to dry. After 2 days, the packets were disassembled, and the fiberglass filters were packed into silver foil cups (Elemental Microanalyses Ltd., Toft, Cambridge, UK). The isotopic ratio of the extracted N was analyzed using Thermo Finnigan Delta V plus (Thermo Scientific, Waltham, MA, USA) at the University of Eastern Finland (Kuopio, Finland).

Calculations and Statistics
Our initial aim was to measure in situ NO 3 − reduction rates with the isotope pairing method (IPT; Nielsen 1992, Christensen et al. 2000, Risgaard-Petersen et al. 2003, 2004, Master et al. 2005. However, two problems arose during the measurements: (1) The concentration of 15 N-labelled N species did not always linearly increase with the 15 NO 3 − concentration (supplementary figure) and (2) the O 2 concentration in the cores during all sampling times was so low that although the incubation time was kept as short as possible, the cores were nearly anoxic by the end of the incubation. These artifacts violate the fundamental assumptions behind the IPT (Nielsen 1992), and calculation of reliable in situ NO 3 − reduction rates was impossible. Therefore, we calculated only the potential NO 3 − reduction rates be averaging the 15 N production rates from all 15 NO 3 − concentrations.
Potential N 2 production is the average of excess 15 N 2 (p 29 N 2 + 2 × 30 N 2 ), potential N 2 O production is the average of excess 15 N 2 O (p 45 N 2 O + 2 × p 46 N 2 O), and potential DNRA is the average excess 15 NH 4 + production. Because in June 2017 there was an increasing trend in the 15 N 2 production with the 15 NO 3 − concentration, the potential during that sampling time provides only a minimum estimate and the actual potentials might be higher (supplementary figure). The differences in rates measured in the core samples were tested by using Whitney-Mann U test. The correlations between the potential rates and environmental parameters were determined by using non-parametric correlation analysis. The NO 3 − consumption and NO 2 − , 15 N 2 , 15 N 2 O, and 15 NH 4 + production rates were calculated from liner regression analysis between the N concentration of different N-species and time. Rates were considered significant at p = 0.05. The comparison between the rates was done by using two-way ANOVA.

Environmental Conditions
Both stations were hypoxic during all sampling times. However, there was always a small amount O 2 detected, except in Oct 2016 at st70 when the odor of H 2 S was detected in the bottom water and in May 2017 at St30 when O 2 was at the detection limit (~3 μmol l −1 ) of the Winkler method (Table 1).
In May 2016, the bottom water salinity was at its lowest at St70, most likely due to excessive rainfall and elevated freshwater runoff which pushed the salt wedge towards the outlet of the estuary. In October 2016, the bottom water salinity at St70 was higher compared with May; however, the saltwater wedge did not reach deep into the estuary and the salinity at St30 was 0. In June 2017, when the river flow had remained low and the saltwater wedge intruded deep into the estuary, the bottom water salinity was 12 at St70 and 10 at St30 (Table 1). The bottom water NO x − concentration was always higher at St30 than at St70, but the opposite was true for NH 4 + ( Table 1).

Potential Denitrification and DNRA Rates Measured on the Intact Cores
There was no evidence of anammox based on the IPT calculations (Risgaard-Petersen et al. 2003, 2004; hence, most N 2 must have originated from denitrification.  (Fig. 2). In October 2016, potential N 2 production was detected in only in seven samples at St70 and at twelve samples at St70, while the rest were below the detection limit. 15 N 2 O production was detected in 12 samples St70 and at all samples at St30. Where denitrification was detected, over 50% of it was N 2 O (Table 1). However, potential DNRA was still proceeding at measurable rates at both stations. There was no significant difference in denitrification and DNRA rates between May 2016 and June 2017 (n = 15 in May 2016 St70 and n = 16 in June 2017 St70, p = 0.545 for denitrification and p = 0.866 for DNRA) (Fig. 2). There were no statistically significant correlations (p < 0.05) between the production of 15 N-labelled products and environmental parameters (O 2 , temperature and salinity) in the core samples.

Potential Denitrification and DNRA Rates Measured from Slurry Samples
The addition of 1 mM H 2 S increased the 15 NO 3 − consumption rates, although not significantly, when compared with the control samples and with samples that had H 2 S + Fe 2+ (Table 2). Although the total 15 N 2 concentration remained lower than in the control samples, it was still increasing steadily over the entire incubation period (Fig. 3). There was also a steady increase in the 15 NH 4 + concentration and the total 15 NH 4 + production rates were significantly higher in the samples that were amended with H 2 S compared with all other treatments   Fig. 3). The 15 NH 4 + product increased, although not significantly, simultaneously to the decrease in NO 2 − concentration at the last sampling point in the control samples and H 2 S + Fe 2+ -treated samples (Fig. 3).

NO 3 − Reduction Rates in the Intact Cores
Overall, potential denitrification rates measured in the NRE were on the lower end of the range of average in situ denitrification rates in temperate coastal sediments (1.2-5 mmol m 2 day −1 ; Seitzinger 1988) but within the same range (0-6600 μM N m −2 day −1 ) previously measured in the NRE by Fear et al. (2005). The low rates in this study can be explained by sampling during warm months when hypoxia is present, which is when Fear et al. (2005) also measured the lowest denitrification rates. Table 2 The N consumption and production rates calculated from regression analysis between concentrations of N species and time. Linearly significant regressions (p ≤ 0.05) are presented in italics Although denitrification is an anoxic process, there typically is a positive correlation between denitrification rates and O 2 concentration because nitrification, which provides NO 3 − for denitrification, is stimulated by the availability of O 2 (Jenkins and Kemp 1984;Kemp et al. 1990). Since we only measured potentials, where denitrification is not limited by the availability of NO 3 − , we expected to find a negative correlation between denitrification rates and O 2 concentration. However, no such correlation was found, and this may be because under hypoxia the presence of H 2 S, rather than the negligible availability of O 2 , is likely to be the key regulating factor for denitrification.
N 2 O production in core samples collected in October 2016 was on the higher end of measured sedimentary N 2 O fluxes in estuarine environments (Murray et al. 2015 and the references there in). This can be partially explained by it being a potential rather than in situ rate since there was more NO 3 − available than under naturally occurring conditions. Presence of H 2 S in the bottom water clearly favored N 2 O production over N 2 production, since 99% of end product of denitrification was N 2 O (Table 1). This is consistent with the early findings of Sørensen et al. (1980) reporting inhibition of N 2 O reduction causing accumulation of N 2 O in the presence of H 2 S. The potential DNRA rates measured at St70 are comparable with DNRA rates measured in the other estuaries in the southern USA (0-2.4 mmol N m −2 day −1 ; Gardner et al. 2006); (0-8.2 mmol N m −2 day −1 ; Giblin et al. 2010) as well as rates measured in sulfide-rich sediments in Denmark (0-6.5 mmol N day −1 , Christensen et al. 2000). At St30, potential DNRA rates were substantially lower but still comparable with DNRA rates measured in periodically anoxic Baltic Sea sediments (0.03-1.1 mmol N m −2 day −1 ; Jäntti and Hietanen 2012). Surprisingly, there was also no correlation between the DNRA and O 2 concentration, and the highest DNRA rates were measured under conditions when little O 2 was available (May 2016). This may be because NO 3 − reduction to NO 2 − , the first step of DNRA, is suggested to be also be driven by denitrifying bacteria ) which can be inhibited by H 2 S. Hence, the DNRA rates were probably limited by the NO 2 − availability. It appears that if H 2 S is present near denitrification layer, the overall NO 3 − reduction rates are lower than in conditions when H 2 S is not present. However, the lowest denitrification to DNRA ratio was found when H 2 S was present in the bottom water (Oct 2016) suggesting that the presence of H 2 S inhibited denitrification relatively more than DNRA. Salinity did not correlate with any of the 15 N production rates, although increases in salinity have been linked to stimulation of DNRA (Gardner et al. 2006;Giblin et al. 2010).
Consequently, it appears that in the NRE, the duration of hypoxia, rather than salinity or O 2 availability, determines the dominance of DNRA over denitrification.

NO 3 − Reduction Rates in the Sediment Slurry Samples
In the slurry samples, the slopes of N 2 production and NO 3 − consumption proceeded steadily but in opposite directions in the control and H 2 S + Fe 2+ treatments, indicating presence of denitrification. Similar 15 N 2 production and NO 3 − consumption pattern between the control and H 2 S + Fe 2+ treatments can be explained by H 2 S chemically reacting with Fe 2+ and forming iron sulfide (FeS), which is a relatively stable compound and does not act as an electron donor for either DNRA or denitrification (Brunet and Garcia-Gil 1996). However, the 15 NH 4 + concentration was higher in the H 2 S + Fe 2+ -treated samples compared with the control samples and it appears that presence of additional electron donors caused a higher percentage of NO 3 − to be converted to NH 4 + over the entire incubation period. Because there was no NO 2 − accumulation and substantial 15 NH 4 + accumulation in the H 2 S + Fe 2+ -treated samples, it is likely that the additional electron acceptors stimulated NO 2 − reduction, the end product of which is NH 4 + . The simultaneously elevated concentrations of NO 2 − and N 2 O in nearly all treatments are in line with the results from wastewater treatment facilities where high N 2 O production during the denitrification phase has been linked with elevated NO 2 − concentrations, although the explanation for this relationship is not clear (Kampschreur et al. 2009).While NO 2 − and N 2 O concentrations began to decrease during the last measurement time, the 15 NH 4 + concentration began to increase, although the increase was not linearly significant, suggesting that DNRA has a higher affinity for NO 2 − than denitrification, similar to results of Kraft et al. (2014). However, the opposite has been observed in chemostat experiments (van den Berg et al. 2017), suggesting that the dominant NO 2 − reduction pathway is also dependent on NO 2 − reducing microbial community composition.
In the H 2 S-treated slurries, there was a steady increase in 15 N 2 , suggesting that denitrifying microbes were still reducing NO 3 − , although at rates lower than the controls and almost equal proportions of NO 3 − were reduced to N 2 and NH 4 + . This was expected, since stimulation of DNRA under elevated H 2 S concentrations has been previously demonstrated in several studies (Høgslund et al. 2009;Schutte et al. 2018;An and Gardner 2002;McCarthy et al. 2008;Jäntti and Hietanen 2012;Bernard et al. 2015;Murphy et al. 2020), as well as in our core incubations, where the highest DNRA to denitrification ratio was observed when the bottom water contained H 2 S in October 2016. However, the NO 3 − concentrations remained higher in the H 2 S-treated samples than in the control and H 2 S + Fe 2+ -treated samples, suggesting that H 2 S overall inhibits NO 3 − reduction, particularly immediately after it becomes present. This was indicated by much lower decrease in the NO 3 − concentration at the first 6 h of incubation in the H 2 Streated samples when compared with the control samples. Interestingly, the N 2 O accumulation in the H 2 S-treated samples was not substantially higher than in the control samples, although N 2 O accumulation was observed in the intact cores in presence of H 2 S. This could be explained by slurries being in a closed incubation system where intermediates, such as N 2 O, cannot escape from the active site (sediment surface) during the incubation, unlike in the intact cores and consequently the intermediates in denitrification/DNRA were consumed at the rate which they were formed. The effect of Fe 2+ on N cycling is not yet well understood because it is challenging to separate the roles of abiotic and biotic processes since they can combine, even within a single organism (Picardal 2012;Melton et al. 2014;Ionescu et al. 2015). Also, most studies have been done by using microbial cultures, and the role of micro-organisms oxidizing Fe 2+ with NO 3 − in natural environments is currently poorly known. In the slurry samples, there was a rapid accumulation of 15 N 2 at beginning of the incubation in Fe 2+ -treated samples. However, the concentration of 15 N 2 did not change after the initial increase; hence, denitrification was quickly brought to a halt. The rapid decrease in N 2 formation in the Fe 2+ -treated samples could be caused by Fe 2+ reacting with the organic carbon compounds present in the sediment producing poorly degradable compounds (Lalonde et al. 2012;Shields et al. 2016), and thus the low availability of labile organic carbon limited heterotrophic denitrification shortly after Fe 2+ was introduced. Fe 2+ can also disturb intracellular electron transport (Carlson et al. 2012) that inhibits denitrification rates and could explain the non-linear increase of 15 N 2 in the Fe 2+ -treated slurries. The low 15 NH 4 + formation in the Fe 2+ -treated samples was unexpected as several studies have demonstrated that Fe 2+ stimulates microbe-mediated DNRA (Robertson et al. 2016;Robertson and Thamdrup 2017;Kessler et al. 2018;Kessler et al. 2019) and abiotic NO 3 − reduction to NH 4 + (Hansen et al. 1994;Hansen et al. 1996;Guerbois et al. 2014

Conclusions
Based on the results of this and other experiments, we conclude that the length of hypoxia has a substantial effect on the N cycling processes in carbon and SO 4 2− rich sediments and that the role of Fe 2+ has to investigated more thoroughly in hypoxic estuarine sediments where H 2 S accumulation occurs. When O 2 is present in the sediment surface (Fig. 4A), nitrification proceeds at the rate which NH 4 + is produced by mineralization. There is no NH 4 + release in the bottom water because nitrification efficiently oxidizes the produced NH 4 + , and denitrification is tightly coupled to nitrification. The DNRA rates are low because in the absence of H 2 S, aerobic heterotrophy dominates, and DNRA microbes cannot compete with denitrification because the quality of organic carbon available for anaerobic processes, after intensive aerobic processes, is low (Kraft et al. 2014). When hypoxia first settles in (Fig. 4B), nitrification proceeds at high rates because nitrification can tolerate microaerophilic conditions (Laanbroek and Gerards 1993;Jäntti et al. 2018) although the end product of nitrification can switch to N 2 O (Kalvelage et al. 2011). The denitrification rates increase because denitrification is stimulated by decreasing O 2 concentrations (Hietanen and Lukkari 2007). At this stage, availability of H 2 S near the sediment surface is still low, and consequently chemolithotrophic DNRA rates remain moderate. When hypoxia is established (Fig. 4C), H 2 S reaches sediment surface and nitrification ceases because it does not tolerate H 2 S (Joye and Hollibaugh 1995). Denitrification rates decrease and the end product of denitrification changes from N 2 to N 2 O. At the same time, DNRA begins to dominate NO 3 − reduction. During persistent hypoxia (Fig. 4D), H 2 S reaches bottom water, and there is no nitrification and only very little denitrification. DNRA rates also decrease because of low NO 2 − availability.
Funding Information Open access funding provided by University of Eastern Finland (UEF) including Kuopio University Hospital. This research was funded by the Academy of Finland (grant nos. 275127 and 307331 (Jäntti), 310302 (Aalto)) and joint funding by Olvi Foundation, Jenny and Antti Wihuri foundation, and Saastamoinen funding (Jäntti). Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.