Acidification Enhances Hybrid N2O Production Associated with Aquatic Ammonia-Oxidizing Microorganisms

Ammonia-oxidizing microorganisms are an important source of the greenhouse gas nitrous oxide (N2O) in aquatic environments. Identifying the impact of pH on N2O production by ammonia oxidizers is key to understanding how aquatic greenhouse gas fluxes will respond to naturally occurring pH changes, as well as acidification driven by anthropogenic CO2. We assessed N2O production rates and formation mechanisms by communities of ammonia-oxidizing bacteria (AOB) and archaea (AOA) in a lake and a marine environment, using incubation-based nitrogen (N) stable isotope tracer methods with 15N-labeled ammonium (15NH4+) and nitrite (15NO2−), and also measurements of the natural abundance N and O isotopic composition of dissolved N2O. N2O production during incubations of water from the shallow hypolimnion of Lake Lugano (Switzerland) was significantly higher when the pH was reduced from 7.54 (untreated pH) to 7.20 (reduced pH), while ammonia oxidation rates were similar between treatments. In all incubations, added NH4+ was the source of most of the N incorporated into N2O, suggesting that the main N2O production pathway involved hydroxylamine (NH2OH) and/or NO2− produced by ammonia oxidation during the incubation period. A small but significant amount of N derived from exogenous/added 15NO2− was also incorporated into N2O, but only during the reduced-pH incubations. Mass spectra of this N2O revealed that NH4+ and 15NO2− each contributed N equally to N2O by a “hybrid-N2O” mechanism consistent with a reaction between NH2OH and NO2−, or compounds derived from these two molecules. Nitrifier denitrification was not an important source of N2O. Isotopomeric N2O analyses in Lake Lugano were consistent with incubation results, as 15N enrichment of the internal N vs. external N atoms produced site preferences (25.0–34.4‰) consistent with NH2OH-dependent hybrid-N2O production. Hybrid-N2O formation was also observed during incubations of seawater from coastal Namibia with 15NH4+ and NO2−. However, the site preference of dissolved N2O here was low (4.9‰), indicating that another mechanism, not captured during the incubations, was important. Multiplex sequencing of 16S rRNA revealed distinct ammonia oxidizer communities: AOB dominated numerically in Lake Lugano, and AOA dominated in the seawater. Potential for hybrid N2O formation exists among both communities, and at least in AOB-dominated environments, acidification may accelerate this mechanism.

Ammonia-oxidizing microorganisms are an important source of the greenhouse gas nitrous oxide (N 2 O) in aquatic environments. Identifying the impact of pH on N 2 O production by ammonia oxidizers is key to understanding how aquatic greenhouse gas fluxes will respond to naturally occurring pH changes, as well as acidification driven by anthropogenic CO 2 . We assessed N 2 O production rates and formation mechanisms by communities of ammonia-oxidizing bacteria (AOB) and archaea (AOA) in a lake and a marine environment, using incubation-based nitrogen (N) stable isotope tracer methods with 15 N-labeled ammonium ( 15 NH + ) and nitrite ( 15 NO − ), and also measurements of the 4 2 natural abundance N and O isotopic composition of dissolved N 2 O. N 2 O production during incubations of water from the shallow hypolimnion of Lake Lugano (Switzerland) was significantly higher when the pH was reduced from 7.54 (untreated pH) to 7.20 (reduced pH), while ammonia oxidation rates were similar between treatments. In all incubations, added NH + was the source of most of the N incorporated into N 4 2 O, suggesting that the main N 2 O production pathway involved hydroxylamine (NH 2 OH) and/or NO − produced by ammonia oxidation during the incubation period. A small but 2 significant amount of N derived from exogenous/added 15 NO − was also incorporated 2 into N 2 O, but only during the reduced-pH incubations. Mass spectra of this N 2 O revealed that NH + and 15 NO − each contributed N equally to N 2 O by a "hybrid-N 4 2 2 O" mechanism consistent with a reaction between NH 2 OH and NO − , or compounds derived 2 from these two molecules. Nitrifier denitrification was not an important source of N 2 O. Isotopomeric N 2 O analyses in Lake Lugano were consistent with incubation results, as 15 N enrichment of the internal N vs. external N atoms produced site preferences (25.0-34.4 ) consistent with NH 2 OH-dependent hybrid-N 2 O production. Hybrid-N 2 O formation was also observed during incubations of seawater from coastal Namibia with 15 NH + and NO − . However, the site preference of dissolved N 2 O here was low (4.9 ),

INTRODUCTION
Ammonia oxidizing bacteria (AOB) and archaea (AOA) are a source of the greenhouse gas nitrous oxide (N 2 O) (Goreau et al., 1980;Santoro et al., 2011;Löscher et al., 2012) in soils and aquatic environments. The rate at which these microorganisms produce N 2 O depends on the rate at which they carry out chemosynthetic reactions that oxidize ammonia (NH 3 ) to nitrite (NO − 2 ). However, other environmental factors can enhance their N 2 O production rate, such as reduced oxygen (O 2 ) concentrations (Goreau et al., 1980;Löscher et al., 2012), higher NO − 2 concentrations, and higher densities of ammonia-oxidizing cells (Frame and Casciotti, 2010). In soils, pH is another factor that influences N 2 O production, with acidic soils generally producing more N 2 O than alkaline soils (Martikainen, 1985). Certain lakes and marine environments also experience pH decreases, which may occur naturally as a result of rapid respiration of organic carbon to carbon dioxide (CO 2 ), or by the dissolution of acid-forming gases (e.g., CO 2 , sulfur dioxide, and nitrogen oxides) produced by human activities.
There are several ways in which reducing the pH of aquatic environments (i.e., acidification) may affect the rate of N 2 O production by ammonia oxidizers. Some evidence suggests that acidification will cause ammonia oxidation rates to decline. Specifically, the ammonia monooxygenase enzyme (AMO), which catalyzes conversion of NH 3 to the intermediate hydroxylamine (NH 2 OH), is thought to act on the free base form of the substrate (NH 3 ), rather than the protonated form, ammonium (NH + 4 ) (Suzuki et al., 1974;Stein et al., 1997). In the pH range of many natural aquatic systems (pH 6-8) NH + 4 /NH 3 is mostly present as NH + 4 (pKa = 9.25 at 25 • C). Any acidification will further reduce the fraction of NH + 4 /NH 3 that is present as NH 3 , and thus reduce the substrate concentration for ammonia oxidizers.
The net effect of ammonia oxidation is also acidifying, releasing protons (H + ) to the surrounding environment: so that, for example, AOB batch cultures that are actively consuming NH 3 are normally exposed to pH decreases as they grow. In these cultures, once the pH drops below ∼6.5, further ammonia oxidation is inhibited (Allison and Prosser, 1993;Jiang and Bakken, 1999b). However, the reason for this may not be decreased substrate availability, since decreases in the activity of AOB are not necessarily correlated with reductions in the NH 3 concentration (Jiang and Bakken, 1999a). It is more likely that inhibition is caused by other factors, such as toxic buildup of nitrous acid (HNO 2 ), nitric oxide (NO), and nitrogen dioxide (NO 2 ) under acidic conditions (Schmidt and Bock, 1997;Stein and Arp, 1998;Schmidt et al., 2002;Udert et al., 2003;Park and Bae, 2009). Recent environmental studies suggest that ammonia oxidation rates may not have a single relationship to pH. For example, ammonia oxidation rates in the open ocean are inhibited by acidification (from pH 8.1-8.2 down to pH 7.6-7.8; Beman et al., 2011;Rees et al., 2016), whereas sedimentary ammonia oxidation rates do not seem to be sensitive to acidification (from pH 8 down to 6; Kitidis et al., 2011). AOA may not be subject to the same growth inhibition as AOB at lower pH ranges. For example, an obligately acidophilic AOA with an optimum pH range of 4-5 was discovered in acidic soil (Lehtovirta-Morley et al., 2011), and in soil pH manipulation experiments, archaeal amoA transcript abundances outnumbered those of AOB in acidic soils (Nicol et al., 2008), suggesting that AOA may outcompete AOB in acidic environments. Marine AOA, which are generally regarded as more important than AOB to ammonia oxidation in the ocean (Wuchter et al., 2006), may also be more tolerant of acidic conditions. For example, certain marine AOA strains are capable of maintaining near-maximal growth rates down to a pH of 5.9 (Qin et al., 2014), perhaps because they express NH + 4 -transport proteins that actively transport NH + 4 into AOA cells, thus supplying AMO with NH 3 under acidic conditions (Lehtovirta-Morley et al., 2011. Unlike NO − 2 , N 2 O is not the major nitrogenous product of ammonia oxidation, and it is not known to what degree the reactions that produce N 2 O are convolved with the main energyharnessing reactions of ammonia oxidizers (i.e., NH 3 oxidation to NH 2 OH and then to NO − 2 ). This means that the impact of pH on the N 2 O production rate may be decoupled from its impact on the ammonia oxidation rate. That is, even if acidification decreases the ammonia oxidation rate, the N 2 O production rate may not necessarily also decrease proportionally. In fact, many of the reactive nitrogen oxides produced during ammonia oxidation undergo N 2 O-forming reactions over relevant timescales, with or without enzyme catalysis, and with their own pH-dependencies.
One of these nitrogen oxides is NH 2 OH, which is the enzymatic product of NH 3 oxidation by AMO in both AOB and AOA (Figure 1, blue box; Hofman and Lees, 1953;Vajrala et al., 2013). Although most NH 2 OH is converted to NO − 2 during active ammonia oxidation, NH 2 OH is also subject to abiotic autoxidation (Figure 1, pathway 1a) and disproportionation reactions (Figure 1, pathway 1b) that produce N 2 O as well as nitrogen (N 2 ), nitric oxide (NO), and NH 3 /NH + 4 . N 2 O yields during these reactions vary with alkalinity (Bonner et al., 1978), redox conditions (Moews and Audrieth, 1959;Pacheco et al., 2011), and the presence of certain transition metals (Anderson, 1964;Alluisetti et al., 2004). NH 2 OH may also react with NO − 2 /HNO 2 to produce N 2 O (Figure 1, pathway 2). This reaction occurs abiotically at a rate that accelerates as pH decreases (Döring and Gehlen, 1961;Bonner et al., 1983). It can also be catalyzed by the copper-and iron-containing NO −

Reference
Oxygen pH Reaction Solutions FIGURE 1 | Reactions between products of ammonia oxidation that produce N 2 0. The steps of ammonia oxidation are in the blue box and the steps of nitrifier denitrification are in the yellow box. Known abiotic pathways to N 2 0 formation are located outside these boxes.
N 2 in proportions that are pH-dependent (Figure 1, pathway 3; Bonner et al., 1978). In terms of tracing the source compounds contributing N to N 2 O, NO can be derived abiotically from HNO 2 through a disproportionation reaction (Figure 1, pathway 4; Park and Lee, 1988), and the reaction of HNO 2 -derived NO with NH 2 OH could also produce a hybrid type N 2 O. However, abiotic disproportionation HNO 2 tends to be most important only in very acidic environments (pKa HNO 2 = 2.8; Riordan et al., 2005). Reduction of NO − 3 and NO − 2 by trace metal ions (Buresh and Moraghan, 1976) and metal-containing minerals (e.g., Rakshit et al., 2008) is known as chemodenitrification (Figure 1, pathway 5). In this process, reduced metal species, particularly Fe 2+ (and possibly also Mn 2+ ) are oxidized, and NO, N 2 O, and N 2 are produced (Picardal, 2012). This pathway has a recognized importance in soils (Zhu-Barker et al., 2015), but is less studied in seawater and eutrophic lake water, which typically have much lower metal concentrations (Morel et al., 2003) than soil. Reducing sediments along productive continental margins may support significant rates of chemodenitrification (Scholz et al., 2016).
Enzymatic reduction of NO − 2 to NO and N 2 O in AOB is known as nitrifier denitrification (Figure 1, (Poth and Focht, 1985). However, in similar experiments with AOA cultures, Stieglmeier et al. (2014b) observed no 46 N 2 O production, even at low O 2 concentrations that are thought to stimulate 1 For clarity in this paper, we will reserve the term nitrifier denitrification for this specific chain of enzymatic reactions and will not use it for other forms of reductive N incorporation from NO − 2 into N 2 O. nitrifier denitrification in AOB (Goreau et al., 1980). Similarly, microrespirometry measurements of Nitrososphaera viennensis cultures indicate that this AOA does not reduce NO − 2 to N 2 O (Kozlowski et al., 2016).
In AOB, NO produced by nitrite reduction is converted to N 2 O by a membrane-bound NO reductase (NOR) that reduces 2NO to N 2 O (Figure 1, yellow box; Beaumont et al., 2004b;Kozlowski et al., 2014). In some denitrifiers, the NOR homolog that carries out the same reduction of NO to N 2 O, has a neutral to acidic pH optimum (5-7.6; Hoglen and Hollocher, 1989) raising the possibility that this step in nitrifier denitrification also has a slightly acidic pH optimum. Among AOA, however, no homologs for the catalytic subunit of bacterial NOR (norB) have been found in any sequenced genomes to date (Santoro et al., 2015), confirming tests of AOA cultures that indicate that nitrifier denitrification does not occur in these organisms (Stieglmeier et al., 2014b;Kozlowski et al., 2016).
NO is a precursor of N 2 O during bacterial nitrifier denitrification, but its production and consumption may be involved in other processes in ammonia oxidizers. For example, NO is an intermediate in the catalytic cycle of hydroxylamine oxidoreductase (HAO) (Cabail and Pacheco, 2003), an enzyme that oxidizes NH 2 OH to NO − 2 in AOB (Figure 1, blue box). NO production is also required for AOA to carry out their ammonia oxidation cycle (Shen et al., 2013;Martens-Habbena et al., 2015;Kozlowski et al., 2016), though no HAO homologs have been identified among AOA (Hallam et al., 2006;Walker et al., 2010).
Field studies assessing the importance of N 2 O production pathways in aquatic environments have relied on two approaches to date: (1) 15 N tracer incubation studies that track the incorporation of N derived from 15 N-labeled precursor molecules, and (2) dissolved N 2 O measurements of the bulk O and N stable isotopic composition as well as the intramolecular distribution of 15 N and 14 N between the internal and external N atoms of the linear, asymmetrical N 2 O molecule (known as site preference; SP = δ 15 N internal -δ 15 N external ; Toyoda and Yoshida, 1999). The SP signature can be useful for distinguishing N 2 O production pathways because it is often (but not always) independent of the isotopic composition of the starting compounds (Yang et al., 2014). Using the first approach, Nicholls et al. (2007) and Trimmer et al. (2016) may have observed hybrid N 2 O formation by an ammonia oxidizer community immediately above the oxygen minimum zone (OMZ) of the Arabian Sea and in the Eastern Tropical North Pacific, respectively, where they observed 45 N 2 O but not 46 N 2 O production during tracer incubations with 15 NH + 4 and NO − 2 with a natural abundance (NA) isotopic composition. In studies using the second approach, profiles of the SP of N 2 O have been used to distinguish N 2 O produced by NH 2 OH-dependent pathway(s), which have a distinctly higher SP (∼34 ; e.g., Sutka et al., 2006;Heil et al., 2014;Frame and Casciotti, 2010) than N 2 O that is formed during denitrification and nitrifier denitrification, which has a much lower SP (0 to −5 ; Toyoda et al., 2005;Sutka et al., 2006;Yamazaki et al., 2014).
Here we have used profiles of dissolved inorganic N concentrations (NH + 4 , NO − 3 , NO − 2 , and N 2 O) and the natural abundance isotopic composition of NO − 3 , NO − 2 , and N 2 O to locate depths where ammonia oxidation and/or N 2 O production are important in the water columns of Lake Lugano, a humanimpacted lake in southern Switzerland, and the marine upwelling zone off the Namibian coast of southwestern Africa. N 2 O isotope and site preference profiles were used to identify the likely pathways of N 2 O production and the involved substrates/intermediates in the two environments. Short (24-30 h) incubations with 15 N-tracers ( 15 NH + 4 and 15 NO − 2 ) at targeted depths revealed that hybrid N 2 O formation occurred in both the shallow hypolimnion of Lake Lugano, as well as in water from the Namibian upwelling zone. Furthermore, N 2 O yields produced during incubations of Lake Lugano water were significantly higher when the pH was reduced experimentally. The isotopic composition of the N 2 O that was produced indicated that the increase was due, at least in part, to enhanced incorporation of N derived from exogenous NO − 2 . Multiplex sequencing of microbial 16S rRNA from the incubation locations indicated that AOB numerically dominated the ammoniaoxidizing community in Lake Lugano whereas AOA dominated in the Namibian Upwelling zone. The lines of evidence presented here suggest that there is potential, at least over the short term, for acidification to enhance hybrid N 2 O formation in aquatic environments.

Sampling
Lake Lugano is separated into a permanently stratified northern basin and a monomictic southern basin. This study focuses on the 95 m-deep southern basin. Water samples and incubation water were collected with a 5L Niskin bottle at the Figino Station (45.95 • N, 8.90 • E) during a sampling campaign on November 5, 2013. Profiles of dissolved O 2 , temperature, salinity, and pH were collected by a conductivity, temperature, and depth sensor (CTD). O 2 profiles were calibrated by Winkler titration. Water from the Namibian Upwelling zone was collected by hydrocast with a 10L-Niskin bottle rosette at station 89 (20.65 • S, 10.95 • E) on January 28, 2014 during the NamUFil cruise of the R/V Meteor.

Geochemical Profiles
Water samples for NH + 4 and NO − 3 concentrations, as well as NO − 3 isotope measurements were immediately filtered through 0.22 µm-pore sterivex filters (Millipore) and then frozen within 2 h of sampling. NH + 4 concentrations were measured fluorometrically (Holmes et al., 1999). NO − 2 concentrations were determined by converting NO − 2 present in 10 ml of sample water to N 2 O by azide reduction (McIlvin and Altabet, 2005) and then quantifying the amount of N 2 O in each sample by gas chromatography-isotope ratio mass spectrometry (GC-IRMS, see below). NO − 2 concentration standards were prepared in 10 ml of distilled water and in lake-water or seawater, and were analyzed by GC-IRMS along with the samples. For NO − 3 concentration and isotopic measurements, sulfamic acid was used to remove NO − 2 prior to analysis (Granger and Sigman, 2009). NO − 3 concentrations were measured by reduction to NO with Vanadium (III) and chemiluminescence detection (Braman and Hendrix, 1989). Nitrate N and O isotope measurements of duplicate samples were performed by conversion of NO − 3 to N 2 O using the denitrifier method (Sigman et al., 2001;Casciotti et al., 2002) and subsequent purification and analysis of this N 2 O with a modified purge-and-trap gas bench GC-IRMS (Thermo Finnigan DeltaV Plus) system. Isotopic calibration was performed by concurrent analysis of NO − 3 isotope standards USGS 32, USGS 34, and USGS 35 (Casciotti et al., 2008) Samples for N 2 O concentration and isotope analyses were taken by overfilling 160 ml glass sample bottles twice from the bottom through a plastic hose connected to the Niskin outlet. The Lake Lugano N 2 O samples were preserved by adding 100 µl of saturated mercuric chloride solution (HgCl 2 ) after a headspace was added by pipetting 1 ml of water off the top of each bottle. Each bottle was then sealed with a butyl rubber septum (VWR, 5483369) and aluminum crimps (CS Chromatographie, 300219). The marine samples were preserved by adding 5 ml of 10 M sodium hydroxide (NaOH) to the bottom of each bottle with a syringe (Mengis et al., 1997), pipetting 1 ml of water off the top for headspace, sealing with butyl septa and aluminum crimps, and then shaking vigorously to distribute the NaOH. Lake Lugano N 2 O samples were analyzed within 1 week of collection. Marine samples were analyzed within 3 months of collection. The total N 2 O in each sample was purged with carrier helium directly into a customized purgeand-trap system (McIlvin and Casciotti, 2010) and analyzed by continuous-flow GC-IRMS. Duplicate N 2 O samples at each depth were collected for the Lake Lugano profile and one sample from each depth was analyzed for the Namibian Upwelling profile. N 2 O isotope ratios were referenced to N 2 O injected from a reference N 2 O tank (≥99.9986%, Messer) calibrated on the Tokyo Institute of Technology scale (Mohn et al., 2012) for bulk and site-specific isotopic composition by J. Mohn (EMPA, Switzerland). Ratios of m/z 45/44, 46/44, and 31/30 signals were converted to δ 15 N-N 2 O (referenced to N 2_AIR ), δ 18 O-N 2 O (referenced to Vienna Standard Mean Ocean Water), and site-specific δ 15 N α and δ 15 N β -N 2 O according to Frame and Casciotti (2010), with an additional two-point correction  using measurements of two isotopic mixtures of N 2 O in synthetic air (CA-06261 and 53504; kindly provided by J. Mohn). N 2 O concentrations were calculated by converting the N 2 O sample peak areas measured by GC-IRMS to N 2 O standards prepared by converting NO − 3 to a known quantity of N 2 O by the denitrifier method (McIlvin and Casciotti, 2010). At each depth, temperature and salinity data were used to calculate the N 2 O concentrations at equilibrium with the atmosphere according to Weiss and Price (1980), based on atmospheric partial pressures reported by NOAA ESRL Global Monitoring Division (http://esrl.noaa.gov/gmd/). The saturation disequilibrium ( N 2 O) was calculated as the difference between the measured N 2 O concentration and the atmospheric equilibrium concentration, with positive values corresponding to oversaturation.

Incubations
A list of all incubation treatments is provided in Table 1. Water for the Lake Lugano incubations that was collected at 17 m depth was poured into opaque 10 L HDPE canisters (Huber, 15.0250.03), stored in the dark for ∼5 h during transport back to the laboratory, and then amended with 15 N-labeled incubation reagents. Water for the seawater incubations was drawn from 200 m depth and immediately mixed with the 15 Nlabeled substrates inside 3.4 L LDPE drinking water containers (Campmor, 81027). A dilution (1:2) of 30% hydrochloric acid (Fluka TraceSelect, 96208) with milliQ water was added to water for reduced-pH incubations of Lake Lugano water and mixed immediately before NH + 4 and NO − 2 substrates were added. Tracer 15 NH 4 Cl (98.5%) and Na 15 NO 2 (99.2%) purchased from Cambridge Isotope Laboratories (NLM-658 and NLM-467) were paired, respectively, with NaNO 2 and NH 4 Cl with natural abundance (NA) isotopic compositions, so that each incubation received either 1 µM 15 NH + 4 + 1 µM NA NO − 2 or 1 µM NA NH + 4 + 1 µM 15 NO − 2 . For each set of experimental conditions (Table 1), 10 acid-washed 160 ml glass incubation bottles (Wheaton, 223748) were rinsed with milliQ and lake or sea water, and then filled with 120 ml of incubation water and closed with gray fluorobutyl PTFE-lined septa (National Scientific, C4020-36AP) and aluminum crimps. Headspace O 2 concentrations of the reduced-O 2 incubations were adjusted by displacement with either high purity (99.999%) helium for the Lake Lugano incubations, or N 2 for the Namibian Upwelling incubations. O 2 in the headspace was quantified using a gas chromatograph with an electron-capture detector (SRI 8610C), and O 2 concentrations were calculated according to Weiss and Price (1980). During the Lake Lugano incubations, one bottle was sacrificed at the beginning of the incubation, and three bottles each were sacrificed after ∼5, 17, or 30 h. During the Namibian Upwelling incubations, one bottle for each set of experimental conditions was sacrificed at the beginning of each incubation, and three bottles were sacrificed after ∼12 and ∼24 h. Bottles were incubated in the dark at 20-21 • C for both experiments.
Immediately before an incubation bottle was sacrificed, 40 ml of liquid was withdrawn by syringe for colorimetric pH measurements with phenol red (Robert-Baldo et al., 1985) or frozen at −80 • C and then stored at −20 • C prior to measurements of concentration and dissolved N isotope composition. Seawater incubation samples were also filtered through polycarbonate membrane filters (Whatman nuclepore) with 0.22 µm pores before freezing. NH + 4 and NO − 3 concentration measurements were made as described above. was calculated at the beginning of the incubation using the added tracer concentration and the ambient concentration of NH + 4 before tracer addition, and assuming that the δ 15 N of ambient NH + 4 was 0 . The 15 F of NH + 4 was not measured during the incubations. The 15 F of NO − 2 during the incubations was measured using the azide reduction/GC-IRMS method (McIlvin and Altabet, 2005), and the 15 F of NO − 3 was measured using the denitrifier method (Sigman et al., 2001) after removal of NO − 2 with sulfamic acid. NO − x -derived N 2 O with a 15 N composition <15% was analyzed using the IRMS manufacturer's resistor (3

DNA Extraction
Water from the incubation depth was filtered through a single 0.22 µm-pore size 47 mm polycarbonate Nuclepore filter (Whatman). The volume of water filtered from the Namibian Upwelling was 4 L and from the Lake Lugano was 0.3 L. Filters were frozen immediately and stored at ≤ −80 • C until extraction. DNA from either environment was extracted using the FastDNA spin kit for soil (MP Biomedicals).

Illumina 16S rRNA Library Generation
The polymerase chain reaction (PCR) was used to amplify the V4 region of the 16S ribosomal RNA (rRNA) gene of prokaryotes using universal 16S rRNA V4 primers F515 (5 ′ -GTGCCAGCMGCCGCGGTAA -3 ′ ) and R806 (5 ′ -GGACTACHVGGGTWTCTAAT-3 ′ ; Caporaso et al., 2011). Forward and reverse primers were barcoded and appended with Illumina-specific adapters (Kozich et al., 2013). PCR amplifications were carried out using BioReagents TM exACTGene TM Complete PCR Kit and Core Reagent Sets (Thermo Fisher Scientific) for 30 cycles (Caporaso et al., 2012). Agarose gel electrophoresis was used to separate PCR products of the correct size that were then bandexcised and recovered using a QIAquick gel extraction kit (Qiagen). For each library, triplicate PCR products were combined and quantified with a Qubit fluorometric assay (ThermoFisher Scientific) and pooled at equimolar ratios. The final pool was analyzed on an Agilent 2100 Bioanalyzer System (Agilent Technologies) using a High Sensitivity DNA chip to determine the average size of the amplicon pool. Quantitative PCR was performed on the pool using a Biorad IQ5 real time thermocycler (Bio-Rad Laboratories) and Illumina quantification standards (KAPA Biosystems). The samples were sequenced on an Illumina MiSeq (http:// www.illumina.com/systems/miseq.ilmn) using 201 nucleotide paired-end multiplex sequencing with 5% PhiX spiked into the run. The library was sequenced on a single flow cell using V2 sequencing reagents, generating paired reads of ∼400 bp, with ∼150 bp overlap between forward and reverse reads.

Bioinformatic Analyses
Raw 16S rRNA sequence data were initially processed in Basespace using Illumina's metagenomic pipeline (https:// basespace.illumina.com/home/index). Merging the paired reads and further analyses were carried out using Axiome with installed PANDAseq and the Quantitative Insights Into Microbial Ecology (QIIME v.1.8.0) software pipeline, including taxaplot for overall bacterial diversity, and calculations of Chao1 to estimate taxon richness and rarefaction curves to calculate species richness for a given number of individual sequences sampled (Caporaso et al., 2010;Masella et al., 2012;Lynch et al., 2013). On PANDAseq, the minimum overlap length was set at the threshold of 0.9. The read length maximum for all sequences was 253 bp. Since the V4 region of the 16S rRNA gene is conserved, reads were removed from further analysis if at least one of the following criteria was met: reads were >4 bp shorter than the maximum mentioned above, the number of ambiguous bases was >1, homopolymers with >4 bp were present, or sequences did not match any sequences in the database by more than 97% based on the percent coverage in BLAST. The clustering of the sequences into operational taxonomic units (OTUs) was initially performed using UCLUST (Edgar, 2010;Edgar et al., 2011) with a cutoff value of 97% sequence identity. The taxonomic identity of a representative sequence from each cluster was classified using the RDP classifier (Wang et al., 2007) and Greengenes datafiles compiled in October 2012 (downloaded from: http://greengenes.lbl.gov/), which includes chimera screening based on 16S rRNA gene records from GenBank (DeSantis et al., 2006). The confidence threshold was set at the default cutoff value of 80% in order to retrieve potential sequences from the nitrifying taxa Thaumarchaeota and Nitrosomonadaceae.

Phylogenetic Analyses
Phylogenetic trees were constructed using MEGA 7 (Kumar et al., 2016). The 16S rRNA gene sequences of representative AOA and AOB were downloaded from GenBank (Benson et al., 2015) for all phylogenetic analyses. Gene sequences were aligned using the MUSCLE algorithm in MEGA 7 and manually inspected. The final alignments consisted of 253 characters, comprising 481 and 108 taxa for sequences related to AOA and AOB, respectively. Phylogenetic reconstruction was implemented using Maximum Parsimony (MP) and Maximum Likelihood (ML). MP was implemented with complete deletion if gaps were present, and tree-bisection-reconnection (TBR) utilized for tree generation. The resulting trees were obtained via random stepwise addition of sequences, at MP search level 1, with 25 initial trees generated from a heuristic search. ML was implemented using Tamura-Nei model of nucleotide substitution rates, with tree inference based on Nearest-Neighbor-Interchange (NNI). Statistical support for MP and ML trees were obtained from 1000 bootstrap replicates under the same initial settings (only bootstrap values >50% are reported). Pairwise base comparisons of OTUs and their closest relatives were determined using BLAST (Altschul et al., 1990) and reported as % identity values.

Statistical and Ecological Analyses
The abundances (frequencies) of ammonia oxidizer OTUs in the Namibian Upwelling and Lake Lugano samples were counted using a program written in Python (v.2.7) (https://www.python. org/download/releases/2.7) to search for each OTU that was verified by the above phylogenetic analyses (Lau et al., 2015). The Shannon-Weiner Index and Pielou Evenness were calculated using the BiodiversityR package (Kindt and Coe, 2005)

Lake Lugano
The 17-m incubation depth in Lake Lugano corresponded to the shallow local minima in both pH (7.47) and O 2 concentration (58 µM; Figures 2A,B). Below 15 m, the respective NH + 4 and NO − 2 concentrations dropped from their surface values of ∼3.8 µM and ∼1.2 µM to ∼0.9 µM and ≤0.3 µM, respectively (Figures 2C,D). In contrast, N 2 O concentrations increased steadily from a surface concentration of 11 nM to 37 nM at 50 m (∆N 2 O = 22 nM; Figure 2E). The δ 15 N-N 2 O dropped from a surface value of 4.9 to a minimum of 2.4 at 15 m and then increased again to 4.9 at 50 m ( Figure 2F). With the onset of seasonal anoxia in the south basin of Lake Lugano (Lehmann et al., 2004;Blees et al., 2014), the sediments (95 m) and deep redox transition zone (70-90 m) become important in the production and consumption of deep N 2 O in this system (Freymond et al., 2013;Wenk et al., 2016). Here we have restricted our discussion to the top 50 m of the water column. See Wenk et al. (2014) and Wenk et al. (2016) for details about N cycle dynamics in the redox transition zone.

Namibian Upwelling Zone
The salinity (35.06) and potential temperature (11.74 • C) of the water at the 200-m incubation depth ( Figure 3A) was characteristic of a deeper tropical branch of South Atlantic Central Water that flows northwestward along the African continental shelf at this latitude (Brea et al., 2004) Figure 3G), and SP (4.9 ± 0.4 Figure 3H). NO − 3 concentrations were near zero in the surface water and increased with depth to a near-maximal concentration of 29 µM at 200 m ( Figure 3I). At this depth, NO − 3 had a δ 15 N (5.3 ± 0.2 , Figure 3J) similar to that of N 2 O and a δ 18 O (3.0 ± 0.3 , Figure 3K) that was much lower than that of N 2 O.

Lake Lugano
Ammonia oxidation rates were estimated in two ways, using data from either the 15 NH + 4 incubations or the 15 NO − 2 incubations.  lower ammonia oxidation rates than those at the untreated pH (0.325 ± 0.004 µM/day and 0.380 ± 0.005 µM/day, respectively). Because we did not measure the 15 F NH4+ over the course of the 15 NH + 4 incubations, we were not able to account for the dilution of the tracer 15 NH + 4 over time by regeneration of NH + 4 , or removal of NH + 4 from the system by processes other than ammonia oxidation (Ward and Kilpatrick, 1990). Therefore, in a second approach, we used the results of the incubations with 15 NO − 2 and NA NH + 4 to calculate zero-order rates of ammonia oxidation (R amm_ox ) and nitrite oxidation (R nit_ox ). Specifically, rates were calculated for each timepoint (t) using measurements of the 15 N atom fractions of NO − 2 ( 15 F NO − 2 , Figure 4B) and Figure 4C), and the total concentrations of NO − 2 ([NO − 2 ], Figure 4D) and NO − 3 ([NO − 3 ], Figure 4E) to solve the following equations: Ammonia oxidation rates calculated this way were similar but somewhat higher than those derived from the 15 NH + 4 incubation data. Average rates calculated for incubations terminated at the middle timepoint (∼17 h) and final timepoint (∼30 h) were 0.50 ± 0.02 µmol/day for the untreated-O 2 -untreated-pH incubations, 0.48 ± 0.02 µM/day for the reduced-O 2 -untreated-pH incubations, 0.54 ± 0.01 µM/day for the untreated-O 2 -reduced-pH incubations, and 0.54 ± 0.02 µM/day for the reduced-O 2 -reduced-pH incubations. Rates are given with the standard deviation among the calculated rates for each of the 17 and 30-h time points. Values of R nit_ox calculated for the 15 NO − 2 incubations were higher than R amm_ox under all conditions. The average R nit_ox was 0.64 ± 0.06 µmol/day among the untreated-O 2 -untreated-pH incubations, 0.59 ± 0.05 µmol/day among the reduced-O 2 -untreated-pH incubations, 0.72 ± 0.03 µM/day among the untreated-O 2reduced-pH incubations, and 0.72 ± 0.05 µM/day among the reduced-O 2 -reduced-pH incubations. Thus, there was net consumption of NO − 2 of 0.1-0.2 µM/day.

Namibian Upwelling Zone
In the Namibian Upwelling incubations, ammonia-oxidation rates calculated from linear regressions of 15 NO − x measured at 12 and 24 h during 15 NH + 4 incubations were two orders of magnitude lower than during the Lake Lugano incubations. Average rates were 3.0 ± 0.3 nM/day (r 2 = 0.96) at 220 µM O 2 , 2.4 ± 0.3 nM/day (r 2 = 0.91) at 50 µM O 2 , and 2.3 ± 0.2 nM/day (r 2 = 0.98) at 20 µM O 2 ( Table 2).   (Figures 5A,B) and then multiplying the molecular fractions by the total N 2 O: For each set of experimental conditions, the total 45 N 2 O and 46 N 2 O measured in incubations killed immediately after tracer addition (t 0 ) was subtracted from the measurements of incubations killed at all subsequent time points. The 45 R and 46 R measured in incubations killed at t 0 were similar to those of the background N 2 O, indicating that our preservation methods prevented further N 2 O production from either 15 NH + 4 or 15 NO − 2 . Error bars in Figure 5 represent the propagated error from measurements of the total N 2 O in each incubation, its isotope ratios, and the total volume of water and background N 2 O present in each incubation bottle. Total daily incorporation of 15 N into N 2 O during 15 NH + 4 incubations was calculated from these daily rates as (Rate 45N2O + 2 × Rate 46N2O ). Rates of 15 N incorporation were 0.0086 ± 0.003 nM-N/day for the untreated-O 2 -untreated-pH 15 NH + 4 incubation, 0.025 ± 0.005 nM-N/day for the reduced-O 2 -untreated-pH incubations, 0.028 ± 0.003 nM-N/day for the untreated-O 2 -reduced-pH incubations, and 0.043 ± 0.001 nM-N/day for the reduced-O 2reduced-pH incubation, where we have indicated ± one standard deviation from the daily average calculated using the final three incubations. Assuming that NH + 4 is the ultimate source of all the N 2 O produced during these incubations, multiplying the rates of 15 N incorporation by their respective isotope dilution factors (1/ 15 F NH + 4 0 ) would increase total N 2 O production rates by factors of 1.50 for the untreated-O 2 -untreated-pH and reduced-O 2 -untreated-pH incubations, and 1.61 for the untreated-O 2reduced-pH and reduced-O 2 -reduced-pH incubations.
Multiplying rates by 1/ 15 F NH + 4 0 does not account for incorporation of N from exogenous NO − 2 into N 2 O, which probably also contributed to N 2 O production, particularly in the reduced-pH incubations. Indeed, during incubations with 15 NO − 2 , significant amounts of 45 N 2 O formed during both the reduced-pH and reduced O 2 -reduced-pH incubations (t-test, p = 0.012 and 0.022, respectively; Figure 5D), indicating that 15 N derived from tracer 15 NO − 2 also contributed to N 2 O production. However, no significant 46 N 2 O production was observed among any of the 15 NO − 2 incubations (Figure 5C), including those that produced significant amounts of 45 N 2 O.
The yield of N 2 O during the 15 NH + 4 incubations was calculated as the rate of incorporation of 15 N into N 2 O relative to the rate of 15 NO − x production. The average yields (mol 15 N-N 2 O/mol 15 N-NO − x ) were 3.64 × 10 −5 for the untreated-O 2 -untreated-pH incubations, 10.0 × 10 −5 for the reduced-O 2 -untreated-pH incubations, 14.0 × 10 −5 for the untreated-O 2 -reduced-pH incubations and 21.0 × 10 −5 for the reduced-O 2 -reduced-pH incubations.

Namibian Upwelling Zone
Among the Namibian Upwelling incubations, there was detectable production of 45 N 2 O when 1 µM 15 NH + 4 + 1 µM NA NO − 2 was added, but not when 1 µM NA NH + 4 + 1 µM 15 NO − 2 was added. The increases in 45 N 2 O were too small to be converted to significant daily rates of 45 N 2 O production, and therefore results are reported in Table 2 using the more sensitive delta notation where the δ 15 N-  (Table 2). However, correction for the differences in background N 2 O among O 2 treatments explains less than half of the increase in δ 15 N-N 2 O during the 20 µM-O 2 incubations as compared to the 220 µM-O 2 incubations.

Overall Microbial Diversity and Abundances
The OTU abundances and their taxonomic affiliation, closest known cultured relatives, and the number of reads assigned to each OTU are summarized in Tables S1, S2. Three separate reactions with DNA extracted from the Lake Lugano water (17 m depth) yielded 382,374, 77,700, and 287,306 partial 16S rRNA gene sequence reads. Three separate reactions with DNA from the Namibian Upwelling incubation water yielded 246,743, 23,288, and 154,829 reads. These reads underwent quality filtration and de-noising to produce a total of 271,425 unique OTUs that were ∼253 bp long. Figure S5 summarizes the bacterial and archaeal phyla into which these OTUs fall. Comparison of the observed taxon richness to Chao1-estimated richness revealed that multiplex sequencing coverage was 47.9 ± 1.4% in the Lake Lugano sample and 52.3 ± 3.7% in the Namibian Upwelling sample. Rarefaction analyses that assess taxon richness in the Namibian Upwelling and Lake Lugano samples were generated with the QIIME pipeline (see Figure S4; Caporaso et al., 2012). The Shannon-Weiner diversity index (H), which is directly proportional to the number of taxa and inversely proportional to the number of sequences falling into each taxon, was an order of magnitude higher for AOB in Lake Lugano than for AOB in the Namibian Upwelling, whereas this index was higher for AOA in the Namibian Upwelling than in Lake Lugano ( Figure S6).

AOA and AOB Diversity and Abundances
A total of 442 unique OTUs related to AOA and 65 unique OTUs related to the AOB family Nitrosomonadaceae were identified. The AOA OTUs constitute 0.3 and 31.2% of total microbial OTUs in Lake Lugano and the Namibian Upwelling site, respectively. AOB OTUs constituted 0.6% of total microbial OTUs in the Lake Lugano sample, but were extremely rare (< 0.01%) in the Namibian Upwelling sample. Both ML and MP trees were nearly identical in their placement of AOA and AOB OTUs with respect to their closest relatives. The 65 AOB OTUs fell into a monophyletic cluster that included cultured members of the family Nitrosomonadaceae ( Figure 6A). The closest cultured representative to 27 of the 65 OTUs was the predominantly terrestrial species Nitrosospira briensis (with 91-100% 16S rRNA sequence identity), although their closest relatives were all uncultured freshwater organisms (see Figure 6A; Table S1). Most of the AOB OTUs (62) were detected only in the Lake Lugano sample and not the Namibian Upwelling sample, and the remaining 3 OTUs were present in both the Lake Lugano and Namibian Upwelling samples (Table S1).
The 442 unique AOA sequences fell into a clade with members of the Thaumarchaea (marine group I archaea) such as Nitrosopumilus maritimus SCM1 and Nitrososphaera sp. JG1, with moderate bootstrap support ( Figure 6B). The closest cultured relatives of these OTUs were mainly found in seawater, with the majority most closely related to Candidatus Nitrosopelagicus brevis strain CN25 (with 88-100% sequence identity), and a large number most closely related to Candidatus Nitrosopumilus sp. NF5 and Candidatus Nitrosopumilus sp. D3C (with 86-98% sequence identity; see Figure 6B; Table  S2). With a few exceptions, the uncultured closest relatives of these OTUs were also detected in seawater (Figure 6B). Of the 442 AOA OTUs identified, 339 were detected only in the Namibian Upwelling sample, 18 were detected only in Lake Lugano, and 85 were detected in both locations (Table  S2). Among the AOA OTUs unique to Lake Lugano, their closest cultured relatives include Candidatus Nitrosopelagicus brevis strain CN25 (Santoro et al., 2015) and Candidatus Nitrosopumilus sp. HCA1 (KF957663.1) (Bayer et al., 2016), which are both marine, and also Nitrososphaera viennensis EN76 (Stieglmeier et al., 2014a) and Candidatus Nitrososphaera evergladensis SR1 (Zhalnina et al., 2014), which were both found in soil.

Geochemical Profiles
The profiles of the N 2 O concentration and isotopic composition were useful as qualitative indicators of the depths of rapid nitrification in Lake Lugano. Although the depth of the water used for the Lake Lugano experiments (17 m) was shallower than the N 2 O concentration maximum, its coincidence with a clear minimum in the δ 15 N-N 2 O profile ( Figure 2F) suggests that there was rapid in situ N 2 O production there. However, the absence of a corresponding extremum in the SP profile ( Figure 2H) at this depth suggests that the δ 15 N-N 2 O minimum probably reflects a minimum in the δ 15 N of the precursor N molecule, rather than a change in the mechanism of N 2 O formation at this depth. While there is also a minimum in the δ 15 N-NO − 3 profile at the incubation depth, the δ 18 O-NO − 3 profile indicates that NO − 3 was probably not the precursor of N 2 O at this depth. More precisely, at this depth, the δ 18 O-N 2 O was 44 higher than the δ 18 O-NO − 3 , whereas the combination of a 42 branching isotope effect (Casciotti et al., 2007;Frame et al., 2014) and a −22 kinetic isotope effect (Granger et al., 2006) associated with NO − 3 reduction to N 2 O, should have produced N 2 O with a δ 18 O that was at most only 20 higher than the δ 18 O-NO − 3 . The similarity of the δ 15 N-N 2 O and δ 15 N-NO − 3 profiles suggests that both compounds are derived from a shared pool of relatively low-δ 15 N precursor N, which could be either NH + 4 or NO − 2 . The δ 15 N-NO − 2 in the top 15 m ranged between −29 and −27 ( Figure 2L). The NH 3 oxidized to NO − 2 at these depths could have been relatively depleted in 15 N because of rapid remineralization of isotopically lighter organic N. The 20 equilibrium isotope effect between NH + 4 and NH 3 (Hermes et al., 1985) and/or expression of the isotope effect(s) associated with ammonia oxidation (Casciotti et al., 2003) may have also contributed to production of 15 N-depleted N 2 O by ammonia oxidizers at this depth. N 2 O production in the deeper water of this basin has been linked to a NH 2 OH-decomposition pathway, largely by the SP value of the N 2 O, which approaches a value of ∼34 in the oxic water between 30 and 70 m (Wenk et al., 2016 and Figure 2H). This particular N 2 O formation mechanism happens during ammonia oxidation in aerobic conditions (Sutka et al., 2006;Frame and Casciotti, 2010;Santoro et al., 2011). Interestingly, such a high SP value is also observed in N 2 O that is formed abiotically by either the hybrid reaction of NH 2 OH with HNO 2 /NO − 2 or by the oxidation of NH 2 OH (Figure 1, pathways 1 and 2; Heil et al., 2014). Thus the production of high SP (∼34 ) N 2 O often observed among AOB and AOA cultures may not distinguish a pathway involving only NH 2 OH from a hybrid pathway where N 2 O is formed via the reduction of HNO 2 /NO − 2 by NH 2 OH. To our knowledge, no data has been reported on the SP of N 2 O produced by the enzyme-catalyzed reaction of NH 2 OH and HNO 2 described by Hooper (1968), but it seems reasonable to assume that it may also be ∼34 . Thus, while the SP of the N 2 O present in the shallower depths of Lake Lugano increases steadily between 5 and 50 m (Figure 2H), suggesting a NH 2 OH-dependent N 2 O formation mechanism, we cannot use SP alone to distinguish N 2 O produced by the reaction between NH 2 OH and NO − 2 from N 2 O produced by NH 2 OH autoxidation or disproportionation.
The high concentration of N 2 O that had accumulated at the depth of the Namibian Upwelling incubation had a relatively low SP (Figure 3H), suggesting that the source of this N 2 O was either NO − x reduction by denitrification or NO − 2 reduction by nitrifier denitrification (Toyoda et al., 2005;Frame and Casciotti, 2010). However, in our incubations we observed hybrid N 2 O formation rather than denitrification or nitrifier denitrification. The absence of denitrification and nitrifier denitrification during the incubations is unsurprising, given the relatively high O 2 concentrations (20, 50, or 220 µ M), all of which were well above thresholds that limit transcription of norB in denitrifiers (Dalsgaard et al., 2014) and initiation of nitrifier denitrification by AOB (Frame and Casciotti, 2010). The lower O 2 concentrations tested during these incubations were similar to the in situ O 2 concentration at this depth (56.8 µM),

FIGURE 6 | Continued
Frontiers in Microbiology | www.frontiersin.org suggesting that if denitrification or nitrifier denitrification had occurred in this water mass, it was not happening at the time and location where we sampled it. Frame et al. (2014) have argued that in this region of the South Atlantic, transport and mixing of continental shelf water that is O 2 -depleted and contains N 2 O produced by anaerobic or suboxic processes with relatively O 2rich offshore water, can produce water that contains relatively high O 2 concentrations and also N 2 O with isotopic signatures that are characteristic of low-O 2 processes like denitrification or nitrifier denitrification.

Nitrification Rates
The difference in ammonia oxidation rates (R amm_ox ) calculated during incubations with 15 NH + 4 vs. those with 15 NO − 2 is probably the result of dilution and loss of the added 15 NH + 4 due to rapid NH + 4 regeneration and uptake, which would both tend to reduce our estimate of R amm_ox during the 15 NH + 4 incubations. It also suggests that the rate reduction that we observed in the reduced-pH 15 NH + 4 incubations does not necessarily reflect an actual reduction in the rate of ammonia oxidation, but instead a more rapid reduction in the 15 F NH4+ over time as compared to the control-pH incubations.
In both Lake Lugano and the Namibian Upwelling, zeroorder reaction kinetics were assumed for ammonia oxidation, rather than first-order or Michaelis-Menten kinetics. This was probably a reasonable assumption in the Namibian Upwelling experiments, because AOA have an extremely high affinity for NH 3 /NH + 4 , with half-saturation constants (K m ) on the order of 100 nM (Martens-Habbena et al., 2009;Horak et al., 2013). In contrast, AOB have a lower affinity for NH + 4 than marine assemblages of AOA (Horak et al., 2013;Newell et al., 2013). The lowest reported K m among cultivated AOB representatives is 6 µM, and the typical range for cultivated AOB is 0.05-14 mM (Knowles et al., 1965;Keener and Arp, 1993;Martens-Habbena et al., 2009;Jiang and Bakken, 1999b). NH + 4 concentrations during the incubations remained well above the K m of AOA but below the K m reported for AOB. Since both AOB and AOA were present in the Lake Lugano incubations, we used the results of the 15 NO − 2 incubations to model first-order rate constants for ammonia oxidation that were 0.46 ± 0.04 M −1 day −1 for the untreated-O 2 -untreated-pH incubations, 0.45 ± 0.04 M −1 day −1 for the reduced-O 2 -untreated pH incubations, 0.47 ± 0.04 M −1 day −1 for the untreated-O 2 -reduced-pH incubations, and 0.42 ± 0.04 M −1 day −1 for the reduced-O 2reduced-pH incubations. Using the observed concentration of NH + 4 (0.92 µM) at 17 m in Lake Lugano and assuming that this concentration is in steady-state, the actual ammonia oxidation rate may be slightly lower than what we calculated using the zero-order reaction model.

N 2 O Yields and Mechanisms of N 2 O Formation in Lake Lugano
Total N 2 O yields measured in Lake Lugano (3.64 × 10 −5 to 21.0 × 10 −5 mol 15 N-N 2 O / mol 15 N-NO − x ) were comparable to those observed by Yoshida et al. (1989) in the western North Pacific, also using 15 N tracer techniques (8 to 54 × 10 −5 ). They were at the low end of the range observed during growth of batch cultures of the AOB Nitrosomonas marina (10 to 60 × 10 −5 ) at similar O 2 concentrations (Frame and Casciotti, 2010) and were lower than those observed for batch cultures of the AOA N. maritimus (60 to 100 × 10 −5 ) and N. viennensis (140 to 180 × 10 −5 ) in media buffered to pH 7.5 (Stieglmeier et al., 2014b). The N 2 O formed during the Lake Lugano incubations was largely derived from intermediates or products of the ammonia oxidation reactions, and relatively little N was incorporated from exogenous NO − 2 , as indicated by the higher rates of 15 N-N 2 O formation during the 15 NH + 4 incubations than during the 15 NO − 2 incubations (Figures 5A-D).
During all of the 15 NH + 4 incubations, the measured ratio of 46 N 2 O: 45 N 2 O production (0.38-0.67) was lower than the expected ratio produced by random pairing of N derived from NH + 4 with the isotope ratio 15 F NH4+0 (expected 46 N 2 O/ 45 N 2 O = ( 15 F NH4+0 ) 2 / (2 × 15 F NH4+0 × (1− 15 F NH4+0 )) = 1.0 for the untreated-O 2 -untreated-pH and reduced-O 2 -untreated-pH incubations and 0.82 for the untreated-O 2 -reduced-pH and reduced-O 2 -reduced-pH incubations). Interestingly, Jung et al. (2014) also report relatively high 45 N 2 O production compared to 46 N 2 O production during tracer incubations of AOB and soil AOA cultures in the presence of 99% 15 NH + 4 and excess N.A. NO − 2 . Because they were working with laboratory cultures, their experiments started with almost no background 14 NH + 4 and contained no ammonium regenerating processes that would increase 14 NH + 4 over the course of the incubations, allowing them to attribute the 45 N 2 O production to a reaction between 15 NH + 4 -derived N and NO − 2derived N.
In the present study, incubations with 15 NH + 4 at the reduced pH produced lower ratios of 46 N 2 O: 45 N 2 O than incubations at the untreated pH. Three factors may account for the difference: (1) the 15 F NH4+_0 was lower among the reduced-pH incubations (0.62) than it was among incubations at the untreated pH (0.67), (2) there may have been differences in the regeneration rates of NH + 4 among experimental treatments, which would have progressively reduced the 15 F NH4+ at different rates in the two different pH treatments, and (3) an increased contribution of N from an unlabeled N pool (i.e., exogenous NO − 2 ) enhanced 45 N 2 O production. Although we lack knowledge of the evolution of 15 F NH4+ during our experiments that would allow us to rule out factors (1) and (2), we can be certain that the more rapid production of 45 N 2 O during the reduced-pH 15 NO − 2 incubations was at least partly the result of additional N-incorporation from exogenous NO − 2 , given the production of 45 N 2 O in the 15 NO − 2 -amended experiment (Figure 5D), and that this argues in favor of factor (3) discussed above.
Nitrifier denitrification is unlikely to have contributed to N 2 O production during the Lake Lugano experiments. During the 15 NO − 2 incubations with untreated-O 2 -reduced-pH and reduced-O 2 -reduced-pH, the formation of 45 N 2 O and not 46 N 2 O (Figures 5C,D) suggests that nitrifier denitrification was not important. Although cultured representatives of Nitrosospira, the most abundant AOB genus in the Lake Lugano, are known to contain norB homologs (Garbeva et al., 2007), whose enzyme products reduce NO to N 2 O during nitrifier denitrification reactions (Figure 1, yellow box; Schmidt et al., 2004;Kozlowski et al., 2014), the incubation conditions, such as the relatively high O 2 concentrations (70 and 290 µM), were unlikely to have stimulated nitrifier denitrification. Rather, a hybrid N 2 O formation mechanism (Figure 1, pathway 2) that combines one N derived from exogenous NO − 2 with one N derived from a different, unlabeled N pool, explains the formation of 45 N 2 O in the absence of 46 N 2 O formation during the 15 NO − 2 incubations. Stieglmeier et al. (2014b) also observed hybrid N 2 O formation by AOA cultures, and have suggested that this other pool of N is NH 2 OH. NH 2 OH is known to form N 2 O in the presence of NO − 2 , both enzymatically (Hooper, 1968) and abiotically (Döring and Gehlen, 1961). Since both NO − 2 and NH 2 OH form in the periplasm (Hollocher et al., 1982), a reaction between these two compounds during the 15 NH + 4 incubations is also consistent with the relatively high rates of both 45 N 2 O and 46 N 2 O production (Figures 5A,B). However, without knowledge of 15 F NH4+ over the course of the incubations, we cannot rule out formation of some 15 N-N 2 O through NH 2 OH autoxidation or disproportionation (Figure 1, pathway 1). A third possibility is that two intracellular NO − 2 molecules react with each other to form N 2 O via nitrifier denitrification in a system where mixing between endogenous (i.e., periplasmic) NO − 2 and exogenous NO − 2 occurs so slowly that the two NO − 2 pools have distinct isotopic compositions. In the 15 NH + 4 incubations, nitrifier denitrification of the relatively 15 N-enriched periplasmic NO − 2 would produce more 45 N 2 O and 46 N 2 O than we would predict based on the measured isotopic composition of the total NO − 2 . In the 15 NO − 2 incubations, nitrifier denitrification of periplasmic NO − 2 would mainly produce 44 N 2 O, and only a small influx of exogenous 15 NO − 2 would be reduced to 45 N 2 O after mixing with unlabeled periplasmic NO − 2 (as we observed during the reduced-pH incubations; Figure 5D). In this way, it would be possible for nitrifier denitrification to produce 45 N 2 O and not 46 N 2 O during the 15 NO − 2 incubations. We believe that this pathway is unlikely given the relatively high O 2 concentrations in our incubations, but without more detailed knowledge of the size of the periplasmic NO − 2 pool maintained by the ammoniaoxidizing cells, and the rate at which this pool exchanges with the external NO − 2 pool, we cannot completely exclude the possibility of nitrifier denitrification.

Possible Explanations for the Influence of pH on N 2 O Production
Reducing the pH during the Lake Lugano incubations increased 15 N 2 O production in both the 15 NH + 4 experiments and the 15 NO − 2 experiments. There may be several reasons for this, including structural (e.g., the outer cell membrane may exchange HNO 2 /NO − 2 more rapidly between the periplasm and the outer environment at a lower pH), enzymatic (e.g., a shift toward the optimal pH of the N 2 O-producing enzymes in the periplasm), transcriptional (regulation of the genes encoding the enzymes involved in N 2 O production may be pH-sensitive), and chemical (due to acceleration in the rates of abiotic reactions that produce N 2 O from precursor molecules made by AOB). As discussed below, the most likely explanations are a shift toward the optimal pH of enzymes that catalyze N 2 O production and/or the involvement of a non-biological catalyst that accelerates the abiotic reactions that form N 2 O.
The majority of ammonia oxidizers in Lake Lugano are Gramnegative bacteria, which means that they have a periplasmic space that is bounded by inner and outer cell membranes. In other Gram-negative species such as Escherichia coli, the pH of the periplasm rapidly changes to reflect that of the external environment (Wilks and Slonczewski, 2007). To our knowledge, it is unknown whether AOB regulate the pH of their periplasm or not, but decreases in the periplasmic pH are likely to enhance the rates of the N 2 O-forming reactions of NH 2 OH and/or NO − 2 . Furthermore, if there are differences in the rate at which HNO 2 vs. NO − 2 cross the outer cell membrane, as suggested by Hollocher et al. (1982), then a pH shift could also alter the rate at which 15 N from the tracer 15 NO − 2 enters the periplasm and the rate at which the NO − 2 formed during ammonia oxidation is expelled from the periplasm into the outer environment. This effect would not necessarily change the actual rate of N 2 O production, just our ability to observe it with 15 N tracers. However, if this occurs, it is unlikely to be the dominant effect, because we observed increased 15 N-N 2 O production during the reduced-pH incubations with 15 NH + 4 , and in this case, more rapid exchange of NO − 2 /HNO 2 across the outer cell membrane in the reduced-pH incubations would dilute the periplasmic concentration of 15 NO − 2 , and therefore decrease the rate of 15 N-N 2 O production relative to total N 2 O production.
The results of the 15 NH + 4 -incubations are consistent with an acceleration of the periplasmic reactions that form N 2 O. Formation of 46 N 2 O, which is composed only of 15 NH + 4 -derived N (and should therefore be relatively independent of any influx of external NO − 2 into the periplasm), was also faster at the reduced pH (Figure 5A). At the enzyme level, Hooper (1968) observed acceleration of N 2 O formation with decreasing pH: AOB enzyme extracts converted NH 2 OH + HNO 2 to N 2 O with an optimum pH of 5.75, via a reaction whose rate increased steadily as the pH dropped from 7.5 to 6. Assuming that a similar reaction also occurs in intact AOB cells, the reaction rate increase observed by Hooper (1968) was large enough to explain the pH effect observed during the Lake Lugano incubations. Decreases in pH also have effects at the level of transcription and protein expression. For example expression of enzymes involved in handling nitrogen oxides in AOB, increases as the pH of the growth medium decreases from 8.2 to 7.2 (Beaumont et al., 2004a). The mechanism for this appears to depend on a transcriptional regulator whose repression is reversed by the presence of NO − 2 at lower pH values (Beaumont et al., 2004a).
Without some form of catalysis, the rate constants reported for the abiotic hybrid N 2 O formation reaction between NH 2 OH and NO − 2 are too small for these reactions to have contributed significantly to N 2 O production during our incubations. In particular, the set of reactions thought to produce N 2 O that contains one NH 2 OH-derived N and one HNO 2 -derived N, has a second-order dependence on [HNO 2 ] (Döring and Gehlen, 1961;Bonner et al., 1983;Schreiber et al., 2012): (Park and Lee, 1988) (10) NO + NO 2 ↔ N 2 O 3 k = 9.5 × 10 7 M −1 day −1 (Grätzel et al., 1970) (11) N 2 O 3 + NH 2 OH → N 2 O + HNO 2 + H 2 O k = 1.7 × 10 13 M −1 day −1 (Döring and Gehlen, 1961;Casado et al., 1983) (12) This rate-limiting step (i.e., Equation 10, the formation of NO by disproportionation of HNO 2 ) becomes important at pH values <4.5 (Hooper, 1968), but it is not fast enough to explain 45 N 2 O production during the incubations, given the low rate constant for HNO 2 disproportionation and the low [HNO 2 ] during the incubations (1.8 × 10 −11 M at the untreated pH and 4.0 × 10 −11 M at the reduced pH). Thus a role for a catalyst, whether enzymatic or non-biological, is indicated. It is important to note that the NO reacting in Equation (11) may be formed through mechanisms other than the rate-limiting abiotic HNO 2 disproportionation step. As mentioned earlier, a number of processes in ammonia oxidizers release NO. For example, nitrite reductases can convert NO − 2 to NO. In AOB, NO is an intermediate in the catalytic cycle of hydroxylamine oxidoreductase (HAO) (Cabail and Pacheco, 2003) and may be released from NH 2 OH in HAO enzyme preparations (Hooper and Nason, 1965;Ritchie and Nicholas, 1972;Hooper and Terry, 1979). In AOA, NO is needed to oxidize NH 2 OH to NO − 2 (Kozlowski et al., 2016). Abiotic reactions between NH 2 OH and NO observed by Bonner et al. (1978) (pH 7.8, anaerobic conditions), produced N 2 O that was ∼75% composed of equal proportions of NH 2 OHderived N and NO-derived N, and ∼25% composed of only NOderived N. If this reaction occurs during ammonia oxidation, then depending on whether NO is derived from NH 2 OH, NO − 2 , or both, these reactions could produce N 2 O that is entirely derived from NH 2 OH, or some mixture of hybrid N 2 O and N 2 O that is entirely derived from NO − 2 . Chemodenitrification, the reduction of NO − 3 or NO − 2 coupled to oxidation of ferrous iron (Fe 2+ ) (Buresh and Moraghan, 1976;Rakshit et al., 2008;Picardal, 2012) was an unlikely source of N 2 O during the Lake Lugano incubations. In the top 20 m of the lake, concentrations of metals involved in chemodenitrification (Fe and possibly manganese, Mn) were less than the 1.7 µM detection limit when measured by induction coupled plasma optical emission spectrometry (J. Tischer, U. Basel, unpublished data). Furthermore, the 15 F NO2− values during incubations with 15 NO − 2 ( Figure 4B) were high enough that if chemodenitrification of NO − 2 to N 2 O had been significant, the observed production of 45 N 2 O ( Figure 5C) would have been accompanied by detectable 46 N 2 O production (Figure 5D), and it was not. The 15 F NO3− values were lower than 15 F NO2− values (Figure 4C), so that mixed reduction of both NO − 2 and NO − 3 by Fe 2+ could explain detectable 45 N 2 O production in the absence of detectable 46 N 2 O production. However, NO − 3 reduction by Fe 2+ is much slower than oxidation by NO − 2 , except when Cu 2+ > 1.6 µM (Buresh and Moraghan, 1976;Picardal, 2012). We did not measure Cu concentrations in Lake Lugano, but the total dissolved Cu measured in Lake Greifen, a similarly eutrophic lake in northeastern Switzerland, were much lower than this (0.5-2.8 × 10 −8 M; Xue and Sigg, 1993).
Although trace metal concentrations were low in Lake Lugano, metal ions could have played a role in accelerating the hybrid N 2 O reaction. Harper et al. (2015) have reported that Cu 2+ can drive abiotic hybrid N 2 O formation rates in activated sludge that are faster than the biologicallycatalyzed reactions. Furthermore, NH 2 OH disproportionation and oxidation reactions may be driven by the presence of copper and iron ions (Anderson, 1964;Alluisetti et al., 2004).

Evidence for Hybrid N 2 O Formation in the Namibian Upwelling Zone
The relationship between the changes in δ 15 N-N 2 O and δ 18 O-N 2 O during the Namibian Upwelling incubations were too small to convert to N 2 O production rates, but they still contain information about the mechanism of N 2 O formation. In the Eastern Tropical South Pacific, above the OMZ (O 2 ≥ 10 µM), the rates of NH + 4 -derived N incorporation into N 2 O (0.01-0.02 nM/day) were similar to what was observed in Lake Lugano, although the yield (as defined in this paper) was substantially higher (80 × 10 −5 ; Ji et al., 2015). In the North Pacific Gyre, Wilson et al. (2014) observed no changes in δ 15 N-N 2 O during incubations with either 1 µM NA NH + 4 + 1 µM 15 NO − 2 or 1 µM 15 NH + 4 + 1 µM NA NO − 2 . Critically, however, when they reduced the NA NH + 4 addition to 100 nM and added 1 µM 15 NO − 2 , the δ 15 N-N 2 O increased significantly over the course of the incubation. This suggests that the rate of ammonia oxidation influences the degree to which N derived from exogenous NO − 2 can be incorporated into N 2 O, with higher rates of ammonia oxidation perhaps flooding the intracellular NO − 2 pool and preventing N derived from exogenous NO − 2 from being incorporated into N 2 O.
It is difficult at this point to determine whether the same hybrid N 2 O reaction mechanism(s) can explain the results of both the Lake Lugano incubations, which were numerically dominated by AOB, and the Namibian Upwelling incubations, which were dominated by AOA. To date, it is not known whether AOA enzymes also catalyze the reaction between NH 2 OH and NO − 2 (Figure 1, pathway 2) as observed for AOB by Hooper (1968). The original AOB periplasmic enzyme complex purified in that study included HAO as well as other enzyme components. No homologs of hao have been identified in AOA genomes, though alternatives have been proposed (Stahl and de la Torre, 2012). Furthermore, if a periplasmic reservoir of NO − 2 plays a role in the incorporation of NO − 2derived N into N 2 O, as we hypothesize here, then differences in the NO − 2 permeability of the outer membranes/cell walls of AOB vs. AOA could contribute to differences in the degree to which N derived from NH + 4 vs. exogenous NO − 2 contribute to N 2 O formation. Unlike AOB, almost all archaea tested have only a single cell membrane bounding the cytoplasm (Albers and Meyer, 2011). Rather than having an outer membrane, AOA have an S-layer protein cell wall separating a pseudoperiplasm from the surrounding environment (Stieglmeier et al., 2014a). Model predictions of AMO protein structure in the soil AOA Candidatus Nitrosotalea devanterra suggest that the membrane-bound enzyme faces outward into the pseudoperiplasm (Lehtovirta-Morley et al., 2016). For future reference during 15 N tracer studies of N 2 O production, it would be helpful to confirm that NH 2 OH and NO − 2 both form in the pseudoperiplasm of AOA, and investigate what controls the rates at which exogenous NO − 2 enters, and periplasmic NO − 2 exits, this compartment.

A Putative Link to O 2
During both the Lake Lugano and Namibian Upwelling incubations, more 15 N-N 2 O was produced at the reduced-O 2 concentrations (O 2 = 70 and 20 µM, respectively) than at the untreated-O 2 concentrations (O 2 = 290 and 220 µM, respectively). It is well known that N 2 O yields by AOB increase during growth at suboxic O 2 concentrations (Goreau et al., 1980), and previous work on the mechanisms causing this increase implicated induction of the nitrifier denitrification pathway at very low O 2 concentrations (e.g., Frame and Casciotti, 2010). Reducing O 2 from 290 to 70 µM in the Lake Lugano incubations nearly tripled the yield of 45 N 2 O and 46 N 2 O during the 15 NH + 4 incubations. However, the reason for this increase was probably not increased nitrifier denitrification, since there was no 46 N 2 O production during any of the incubations with 15 NO − 2 ( Figure 5C).
The mechanism conferring this O 2 sensitivity may not necessarily involve direct regulation of enzyme activity. In particular, NO removal by O 2 is a possible abiotic NO-sink that would become more important at higher O 2 and NO concentrations: 2NO (aq) + O 2 (aq) → 2NO 2 (aq) k = 1.8 × 10 11 M −2 day −1 (Awad and Stanbury, 1993) (13) Once NO 2 is formed, in aqueous solutions it tends to react with water to form NO − 3 and NO − 2 (Park and Lee, 1988), or react with NO to form N 2 O 3 (Grätzel et al., 1970). Martens-Habbena et al. (2015) measured NO concentrations of ∼50-80 nM during oxic incubations of N. maritimus with 10 µM NH + 4 . If similar NO concentrations were produced during incubations in the present study, liquid-phase reactions between NO and O 2 may deplete NO concentrations significantly, with the rate of depletion increasing in proportion to [O 2 ] as well as [NO] 2 . Thus, abiotic reaction with O 2 may compete for NO with N 2 O-forming reactions that also consume NO, particularly when incubation O 2 concentrations are high.
Like O 2 , NO tends to partition into the gas phase over the aqueous phase (Schwartz and White, 1981). If it is NO (aq), rather than NO − 2 , that participates in biological hybrid N 2 O formation, an implication is that inclusion of a headspace during incubations of ammonia oxidizers suspended in water will reduce aqueous NO concentrations, and therefore slow down liquid-phase NO-dependent reactions (such as the reaction of NO with NH 2 OH to form N 2 O). Differences in aqueous NO concentrations might contribute to the discrepancy in the literature over whether reduced-O 2 growth conditions increase the yields of N 2 O produced by AOA (e.g., Löscher et al., 2012;Stieglmeier et al., 2014b), particularly if there is variation in aeration procedures and ratios of headspace to liquid volumes.

CONCLUSIONS
Previous studies have shown that decreases in pH can increase N 2 O production by AOB cultures (e.g., Jiang and Bakken, 1999a) but did not separate the effect of pH-dependent NH 3 limitation from the influence of pH on the N 2 O production mechanisms.
Here we have shown that acidification enhances the N 2 O yields of ammonia oxidizers even when it does not substantially change the ammonia oxidation rates. We have demonstrated that hybrid N 2 O formation (i.e., the combination of NH + 4and NO − 2 -derived N) occurs among the Nitrosospira-dominated ammonia oxidizer community in the shallow hypolimnion of Lake Lugano and that this mechanism contributes to the increased yield of N 2 O under acidified conditions. The NH + 4derived reactant in this hybrid N 2 O production pathway is probably NH 2 OH, while the NO − 2 -derived reactant could be one of several inter-convertible nitrogen oxides (NO − 2 /HNO 2 , NO, N 2 O 3 ). Our results suggest that nitrifier denitrification was not an important source of N 2 O in this environment. While N derived from exogenous NO − 2 contributed significantly to N 2 O formation under acidified conditions, N derived from NH + 4 was always a more important contributor to N 2 O. Finally, we report preliminary isotopic evidence that hybrid N 2 O formation also occurs among the subsurface AOA-dominated nitrifier community present in the Namibian Upwelling zone.
Our results are not necessarily predictive of the long-term influence of acidification on N 2 O production by ammonia oxidizers, since acidification may also change ammonia oxidizer community composition (Bowen et al., 2013) and pH decreases may have cascading chemical and biological effects in lake and ocean ecosystems. However, our results are applicable to environments that experience rapid changes in pH such as stratified lakes that undergo episodic mixing or rapid influx of acidified precipitation, and ocean upwelling zones where CO 2 -rich, low-pH deeper water may enhance N 2 O production when it comes in contact with shallower ammonia-oxidizing communities.

AUTHOR CONTRIBUTIONS
CF conceived of and performed experiments. EL and EN analyzed and interpreted genetic sequence data. TG provided instrumentation support and sample analysis. CF and ML performed chemical data analysis and interpretation. All authors contributed to writing this paper.