Hydroxylamine as an intermediate in ammonia oxidation by globally abundant marine archaea

Abstract

The ammonia-oxidizing archaea have recently been recognized as a significant component of many microbial communities in the biosphere. Although the overall stoichiometry of archaeal chemoautotrophic growth via ammonia (NH₃) oxidation to nitrite (NO₂⁻) is superficially similar to the ammonia-oxidizing bacteria, genome sequence analyses point to a completely unique biochemistry. The only genomic signature linking the bacterial and archaeal biochemistries of NH₃ oxidation is a highly divergent homolog of the ammonia monooxygenase (AMO). Although the presumptive product of the putative AMO is hydroxylamine (NH₂OH), the absence of genes encoding a recognizable ammonia-oxidizing bacteria-like hydroxylamine oxidoreductase complex necessitates either a novel enzyme for the oxidation of NH₂OH or an initial oxidation product other than NH₂OH. We now show through combined physiological and stable isotope tracer analyses that NH₂OH is both produced and consumed during the oxidation of NH₃ to NO₂⁻ by Nitrosopumilus maritimus, that consumption is coupled to energy conversion, and that NH₂OH is the most probable product of the archaeal AMO homolog. Thus, despite their deep phylogenetic divergence, initial oxidation of NH₃ by bacteria and archaea appears mechanistically similar. They however diverge biochemically at the point of oxidation of NH₂OH, the archaea possibly catalyzing NH₂OH oxidation using a novel enzyme complex.

Microbial oxidation of ammonia (NH₃) to nitrite (NO₂⁻), the first step in nitrification, plays a central role in the global cycling of nitrogen. Recent studies have established that marine and terrestrial representatives of an abundant group of archaea, now classified as Thaumarchaeota, are autotrophic NH₃ oxidizers (1–5). Despite increasing evidence that ammonia-oxidizing archaea (AOA) generally outnumber ammonia-oxidizing bacteria (AOB), and likely nitrify in most natural environments, very little is known about their physiology or supporting biochemistry (6, 7). Genome sequence analyses have pointed to a unique pathway for NH₃ oxidation, likely using copper as a major redox active metal, and coupled to a variant of the hydroxypropionate/hydroxybutyrate cycle (8). However, the only genome sequence feature that associates the archaeal pathway for NH₃ oxidation with that of the better characterized AOB is a divergent variant of the ammonia monooxygenase (AMO), which may or may not be a functional equivalent of the bacterial AMO. Thus, the supporting biochemistry of a biogeochemically significant group of microorganisms remains unresolved (8, 9).

Among the AOB, as represented by the model organism Nitrosomonas europaea, NH₃ is first oxidized to hydroxylamine (NH₂OH) by AMO, an enzyme composed of three subunits encoded by amoC, amoA, and amoB genes (7). NH₂OH is subsequently oxidized to NO₂⁻ by the hydroxylamine oxidoreductase (HAO) (7), a heme-rich enzyme encoded by the hao gene (7). Of the four electrons released from the oxidation of NH₂OH to NO₂⁻, two are transferred to the terminal oxidase for respiratory purposes and two are transferred to AMO for further oxidation of NH₃ (7). Although all available genome sequences for the AOA contain homologs of the bacterial AMO (amoB, amoC, and amoA), there are no obvious homologs of AOB-like HAO, or cytochromes c554 and c_M552 critical for energy conversion in AOB (8–15). Thus, either the product of NH₃ oxidation is not NH₂OH or, alternatively, these phylogenetically deeply branching thaumarchaea use a novel biochemistry for NH₂OH oxidation and electron transfer (8).

In an attempt to gain further insights into the biochemistry and physiology of these unique archaeal nitrifiers, we here investigated the role of NH₂OH in Nitrosopumilus maritimus metabolism. These studies were complicated by the extremely oligotrophic character of this organism contributing to very low cell densities in culture (16). To overcome the challenge of working with low cell density cultures of N. maritimus, we established a method to concentrate cells on nylon membrane filters such that the cells remained competent for NH₃-dependent NO₂⁻ formation and oxygen (O₂) uptake. This method enabled us to carry out relatively short duration physiological studies and stable isotope tracer experiments directed at determining if N. maritimus can oxidize exogenous NH₂OH to NO₂⁻ while consuming O₂ and producing ATP, and if NH₂OH is an intermediate in NH₃ oxidation pathway of N. maritimus.

Results

N. maritimus Oxidizes NH₂OH to NO₂⁻.

The apparent lack of genes encoding a recognizable AOB-like HAO complex in the genome of N. maritimus has generated considerable interest in the pathway of NH₃ oxidation in AOA (8). Because purification and direct biochemical characterization of AMO enzymes has thus far been unsuccessful, we aimed to determine if N. maritimus could convert NH₂OH to NO₂⁻. To determine if N. maritimus can oxidize NH₂OH to NO₂⁻, it was necessary to expose the N. maritimus cells to NH₂OH in the absence of the NH₄⁺ used to grow the culture. Hence, a technique was developed to concentrate the cells onto 0.2-µm pore size nylon membrane filters using a vacuum manifold system. The manifold system allowed simultaneous filtration of several liters of culture. In addition to concentrating the cells and separating them from residual NH₄⁺ in the medium, the nylon membranes served as a solid support for convenient handling of concentrated cells in subsequent incubation assays. The membrane-associated N. maritimus cells actively oxidized NH₃ for several hours.

To investigate whether NH₂OH can be oxidized by N. maritimus, a series of short-term NO₂⁻ accumulation assays was conducted, in which membrane-associated N. maritimus cells were exposed to various concentrations of NH₂OH. Irrespective of the initial concentration of NH₂OH (50–200 µM), N. maritimus oxidized ∼50 µM NH₂OH to NO₂⁻ over 20 h (Fig. 1). During the first 30 min of exposure to 200 µM NH₂OH, the rate of NH₂OH-dependent NO₂⁻ production was equal to the rate of NH₃-dependent NO₂⁻ production [0.19 ± 0.01 µmol/(min × mg protein)]. With longer incubation times, the rate of NH₂OH oxidation decreased gradually and finally stopped after accumulation of ∼50 µM of NO₂⁻ (Fig. 1). Addition of 1 mM NH₂OH led to a complete inhibition of the NO₂⁻ formation (Fig. 1), reflecting a toxicity at higher concentrations also observed for AOB (17). Attempts to grow N. maritimus on NH₂OH as sole energy source failed as previously observed in N. europaea (18).

Fig. 1.

Accumulation of NO₂⁻ by N. maritimus. Determinations in the presence of 200 µM NH₄⁺ (dashed lines, squares), 200 µM NH₂OH (solid lines, diamonds), 50 µM NH₂OH (solid lines, triangles), 1 mM NH₂OH (solid lines, circles), and no substrate (dashed lines, crosses). Inset shows accumulation of NO₂⁻ after 20 h of incubation. Data are means of triplicates, with variation less than 10%. The experiment was repeated at least three times and produced similar results. Error bars represent SEM.

Acetylene (C₂H₂) (19) and allylthiourea (ATU) (20) are specific inhibitors of NH₃ oxidation by AMO in AOB and AOA (19–21). In the present work, the ability of N. maritimus to oxidize NH₃ or NH₂OH in presence of either C₂H₂ or ATU was analyzed by monitoring NO₂⁻ accumulation. C₂H₂ (0.1%) was added to the headspace of sealed bottles containing membrane-attached N. maritimus cells in synthetic crenarchaeota medium (SCM) containing either NH₄⁺ or NH₂OH. C₂H₂ completely inhibited NH₃-dependent NO₂⁻ accumulation but had no effect on NH₂OH-dependent NO₂⁻ production (Fig. S1 A and B) as previously observed in N. europaea (19). Similarly, the addition of ATU also inhibited NH₃-dependent NO₂⁻ production but did not inhibit NH₂OH-dependent NO₂⁻ production (Fig. S1 A and B) (20). These results clearly indicate that N. maritimus can oxidize NH₂OH and its oxidation is independent of the AMO activity (Fig. S1 A and B).

NH₂OH Oxidation Is Coupled to O₂ Uptake in N. maritimus.

Further experiments were conducted to determine if oxidation of NH₂OH to NO₂⁻ by N. maritimus is accompanied by consumption of a stoichiometric amount of O₂. To test this, NH₂OH consumption, O₂ uptake, and NO₂⁻ accumulation were independently determined (Fig. 2). The rate of NH₂OH-dependent O₂ consumption by N. maritimus remained constant for 8 min at a rate of 0.20 ± 0.01 µmol O₂/(min × mg protein) and the ratio of O₂ consumed to NO₂⁻ produced was 1 ± 0.04 (Fig. 2). This stoichiometry is similar to that observed in AOB (22). By comparison, the rate of O₂ consumption in the presence of NH₄⁺ (200 µM) was 0.28 ± 0.03 µmol O₂/(min × mg protein) and reflects the stoichiometry of 1 NH₃:1.5 O₂ previously reported for both the bacterial and archaeal oxidation of NH₃ to NO₂⁻ (16, 23). Substrate depleted N. maritimus cells consumed only 0.003 ± 0.001 µmol O₂ per minute per milligram protein. This endogenous rate of O₂ uptake by N. maritimus was 50–100 times lower compared with the O₂ uptake rate observed with NH₃ or NH₂OH and consequently provides a good measure of the O₂ uptake rates observed during NH₃ or NH₂OH oxidation.

Fig. 2.

Stoichiometry of NH₂OH oxidation in N. maritimus. O₂ uptake (solid squares), NH₂OH consumed (open circles), NO₂⁻ produced (gray triangles); Data are means of triplicates, with variation of less than 10%. Incubations were carried out for 8 min while the conditions were at optimum (i.e., the unavoidable degradation of NH₂OH did not interfere with the stoichiometry). The experiment was repeated at least three times and produced similar results. Error bars represent SEM.

To evaluate the possibility that NO₂⁻ was produced from NH₂OH before inhibition by C₂H₂ or ATU was maximal, NH₃-dependent and NH₂OH-dependent O₂ consumption were studied in the presence of either C₂H₂ or ATU. Membrane-immobilized N. maritimus cells were treated with C₂H₂ for 2 h before placing them in the O₂ electrode chamber. C₂H₂-exposed cells commenced respiring immediately after the addition of NH₂OH (Fig. S2A). Similarly, ATU-inhibited N. maritimus cells immediately resumed O₂ consumption upon addition of NH₂OH (Fig. S2B). In agreement with previous observations (2), higher concentrations of ATU (2.5 mM) were required in our studies as well to completely stop NH₃-dependent NO₂⁻ accumulation and O₂ uptake in N. maritimus, whereas as little as 10 µM ATU concentrations inhibited NH₃-dependent NO₂⁻ accumulation and O₂ uptake in N. europaea (20). The fact that C₂H₂ or ATU did not inhibit NH₂OH-dependent O₂ uptake clearly indicates that NH₂OH oxidation is not dependent on AMO (Fig. S2 A and B).

NH₂OH Oxidation Is Coupled to ATP Production in N. maritimus.

If NH₂OH is an intermediate of NH₃ oxidation in N. maritimus analogous to the AOB, then the oxidation of NH₂OH should be coupled to the respiratory electron transfer chain and ultimately to the reduction of O₂, and to ATP synthesis. Indeed, we demonstrated that NH₂OH oxidation is coupled stoichiometrically (1:1) to reduction of O₂ (see above), suggesting electron transfer from NH₂OH to O₂. Given the limited N. maritimus biomass available, direct measurement of electron transport or proton translocation in response to NH₂OH was impossible. Thus, to confirm the metabolic coupling of NH₂OH oxidation, we measured ATP levels and NO₂⁻ production rates in the presence of different substrates. Cellular ATP levels were measured following addition of SCM media containing either no substrate, NH₄⁺, or NH₂OH to membrane-immobilized N. maritimus cells. Cells incubated in the presence of NH₂OH showed statistically significantly higher levels of ATP compared with cells incubated without substrate (P value 0.006). Furthermore, cells incubated with NH₂OH showed statistically significantly higher ATP levels compared with cells incubated with NH₄⁺ (P value 0.007) (Fig. 3A) although the NO₂⁻ produced in presence of either NH₄⁺ or NH₂OH is almost similar (Fig. 3B). Addition of either carbonyl cyanide m-chlorophenyl hydrazone (CCCP), a well-characterized uncoupler of ATP synthesis, or use of heat-killed cells, prevented NH₃-dependent or NH₂OH-dependent increase in ATP levels in all treatments (Fig. 3A). Furthermore, CCCP addition diminished NO₂⁻ production from NH₃, but not NO₂⁻ production from incubation with NH₂OH (Fig. 3B). These results suggest that the respiratory reduction of O₂ associated with NH₂OH oxidation is coupled to ATP production in N. maritimus, as has been reported in N. europaea (24). Inhibition of NO₂⁻ production from NH₃ by CCCP further suggests that analogous to the AOB, the oxidation of NH₃ in N. maritimus by AMO also depends on input of electrons from electron carriers (7). Consistent with this dual sink of electrons in the NH₃ oxidation metabolism, the cellular level of ATP is lower during incubation with NH₄⁺ compared with incubations with NH₂OH only (Fig. 3A).

Fig. 3.

ATP formation by N. maritimus. (A) ATP and (B) NO₂⁻ concentration in N. maritimus cells incubated in presence of either no substrate (white bars), 200 µM NH₄⁺ (gray bars), or 200 µM NH₂OH (black bars). Concentration of ATP and NO₂⁻ in heat-killed cells and cells treated with 100 µM CCCP for 20 min is also shown. The NO₂⁻ trace amounts detected in the controls are likely due to leaching of residual traces of NO₂⁻ in the nylon filters used to harvest N. maritimus cells. Error bars represent the SEM.

Generation of NH₂OH from NH₃.

We used stable isotope tracer experiments to further establish that NH₂OH is an intermediate in the NH₃ oxidation pathway of N. maritimus. Membrane-immobilized cells were incubated with natural abundance NH₄Cl and NH₂OH (200 µM each). The ¹⁵N isotopic composition of NH₂OH and accumulated NO₂⁻ were measured over time by isotope ratio mass spectrometry (IR-MS) following chemical conversion of NH₂OH or NO₂⁻ to nitrous oxide (N₂O) by iron acetate (FeAc) or sodium azide, respectively (25). NH₂OH-derived N₂O and N₂O present in the control vials before chemical conversion of NH₂OH to N₂O were quantified by gas chromatography (GC). During 60 min of incubation, NO₂⁻ concentrations steadily increased and NH₂OH concentrations decreased (Fig. 4 A and B). After addition of 20 µM ¹⁵NH₄Cl, the relative abundance of ¹⁵N increased over time in the NH₂OH and NO₂⁻ pools (Fig. 4 A and B). After 120 min, NH₃ oxidizing activity was blocked by addition of C₂H₂. Accumulation of ¹⁵N stopped completely in the NH₂OH pool within 20 min of C₂H₂ addition, indicating complete inhibition of metabolism of highly ¹⁵N-labeled NH₃. In contrast, uptake and oxidation of then significantly ¹⁵N-enriched NH₂OH pool continued, leading to further accumulation of ¹⁵NO₂⁻ at a lower rate (Fig. 4 A and B). Due to rapid decomposition of NH₂OH in SCM media, we could not establish the mass and isotope balance among NH₄⁺, NH₂OH, and NO₂⁻ in our ¹⁵N experiments. Moreover, presence of high concentrations of NH₂OH was toxic to N. maritimus and slowed down the rates of NH₄⁺ oxidation and NO₂⁻production.

Fig. 4.

Kinetics of N. maritimus NH₂OH consumption and accumulation of ¹⁵N in a stable isotope tracer experiment. N. maritimus cells were incubated in the presence of 200 µM NH₄Cl and 200 µM NH₂OH. After 60 min (arrow) 20 µM ¹⁵NH₄Cl (99.5% purity) was added to the cell suspension. After 120 min NH₃-oxidation activity was blocked by addition of C₂H₂ (0.1%). (A) Consumption of NH₂OH (dashed lines, filled circles) and accumulation of ¹⁵N in NH₂OH pool (solid lines, filled diamonds). (B) Accumulation of NO₂⁻ (dashed lines, filled circles), ¹⁵N in NO₂⁻ pools (solid lines, filled diamonds).

Because the determination of δ¹⁵N signatures of NH₂OH and NO₂⁻ entailed a chemical conversion step affording the more stable N₂O for subsequent stable isotope analysis, a series of controls were conducted to exclude other sources of ¹⁵N accumulation in the N₂O pool. Neither NH₄⁺ itself, nor N₂H₄ were found to yield N₂O by the chemical conversion method used here, thus, ruling out a possible interference by NH₄⁺ itself or a role of N₂H₄ in the N. maritimus metabolism. We further eliminated the possibility of generation of N₂O in N. maritimus cell suspensions from chemical decomposition of unstable N species, such as nitric oxide (NO), nitroxyl (HNO), or nitrosyl (NO⁺), or their rapid reaction with added NH₂OH. Although such chemical reactions cannot be completely excluded, their significance during our experiments must be negligible because N₂O concentrations in the control vials did not increase over time, indicating no significant new production of N₂O through either of these reactions. Thus, although NH₂OH does not accumulate at measurable concentrations during normal growth of N. maritimus in culture, production and consumption of NH₂OH during oxidation of NH₃ remains the only plausible explanation for our observed results, indicating that NH₂OH is an intermediate in the NH₃ oxidation pathway of N. maritimus.

Discussion

The lack of an identifiable gene encoding HAO in the genome of N. maritimus (8) necessitates either a novel enzyme for the oxidation of NH₂OH or, alternatively, an initial product of NH₃ oxidation other than NH₂OH (8). We earlier speculated that the immediate oxidation product of the archaeal variant of AMO might be a reactive N species, such as nitroxyl, instead of NH₂OH (8). However, the data presented in this study now clearly implicate NH₂OH as the immediate product of AMO, with properties similar to those previously observed in N. europaea (20). NO₂⁻ accumulation and O₂ uptake during NH₂OH oxidation by N. maritimus were stoichiometrically coupled (Fig. 2) and closely paralleled NH₂OH disappearance and therefore fulfilled the expected 1:1 stoichiometry of NH₂OH and O₂ consumption. Metabolism of NH₂OH was not inhibited by C₂H₂ or ATU, two well-characterized inhibitors of the bacterial AMO. Both archaeal and bacterial NH₃ oxidation are inhibited by C₂H₂, a mechanism-based inactivator of NH₃ oxidation in the AOB (Fig. S1A) (21, 26). As observed in AOB, C₂H₂ did not inactivate NH₂OH oxidation in N. maritimus (Fig. S1A). Previously, C₂H₂ was shown to be a suicidal substrate for AMO in N. europaea and that protein synthesis was essential for recovery from C₂H₂ inactivation (19, 26). N. maritimus cells inactivated by C₂H₂ also require a recovery period, not showing any respiratory capacity until 60–90 min following NH₄⁺ addition. In addition, the oxidation of NH₂OH by N. maritimus remained virtually unaffected by concentrations of ATU, an inhibitor of the bacterial AMO, that completely inhibits AOB and AOA NH₃ oxidation at the concentrations used in our experiments (Fig. S1B) (20). Thus, although the enzymes and metabolic pathway of AOA have not yet been characterized (8–12), we have now shown that N. maritimus is capable of oxidizing NH₂OH in presence of C₂H₂ similar to N. europaea. Thus, we can also discard the possibility that C₂H₂ interferes with components other than the AMO protein in the AOA.

Having demonstrated that NH₂OH can be oxidized to NO₂⁻ by N. maritimus, we then considered whether this oxidation provided a benefit to the cells. This is a necessary consequence of NH₂OH serving as an intermediate in NH₃ oxidation, because all electrons for respiration are derived from the oxidation of NH₂OH. Indeed, a benefit was observed by marked ATP production in response to NH₂OH addition relative to resting cells without substrate (Fig. 3A). Similar, but lower, ATP production coupled to oxidation of NH₂OH was measured in N. europaea (24, 27). Although we have no explanation for this differential response, N. maritimus may maintain a slightly higher energy charge than that characterized for AOB.

Direct evidence that the oxidation of NH₃ to NO₂⁻ by N. maritimus proceeds via NH₂OH came from stable isotope tracer experiments in which ¹⁵NH₂OH production was detected while N. maritimus was oxidizing ¹⁵NH₃ (Fig. 4A). One possible caveat to this interpretation is that accumulation of ¹⁵NH₂OH was measured by IR-MS after its conversion to ¹⁵N₂O. Recent reports demonstrated that archaeal NH₃ oxidation produces N₂O (28) albeit no biochemical pathway for its production in AOA has been identified (29). Production of N₂O either by N. maritimus itself or chemical reactions affording N₂O in our growth media, e.g., through decomposition of NH₂OH, could potentially yield N₂O that would interfere with NH₂OH isotopic analysis. We attempted to quantify N₂O concentrations and isotopic composition in parallel in iron acetate-converted and -unconverted samples under identical conditions. However, N₂O concentrations in the unconverted samples were too low and outside the calibration range of our IR-MS setup, even after increasing the injection volume 10-fold. We estimate the highest N₂O concentration in any unconverted sample was 6–10 nM (median 7.75, SD 1.6, n = 9) and it neither increased nor decreased significantly over time. Albeit, we cannot state its isotopic composition with confidence, the measured δ¹⁵N of unconverted and converted samples were in a similar range throughout the experiments (−28.8 to + 66.3, and −32.7 to + 140.0 for NH₂OH-derived and direct N₂O, respectively). Thus, ¹⁵N accumulating in the N₂O pool contributed a very minor fraction of ¹⁵N accumulating in the NH₂OH pool. Previous studies demonstrate that N₂O production during NH₃ oxidation in enrichment culture from oceanic water samples, enrichments, and N. maritimus only contribute up to two parts per thousand of NO₂⁻ produced (29). A similar rate of N₂O production by N. maritimus cells during our experiments would yield an estimated 70 nM N₂O over the course of our entire experiments. Because the N₂O concentrations in the unconverted samples were below detectable levels, we conclude that any direct N₂O produced in our experiments would not affect the NH₂OH isotopic analysis.

The isotopic signature of N₂O produced by AOA strongly indicated that it is a product of NH₃ oxidation and not from reduction of NO₂⁻ (28). In addition, C₂H₂ (∼10%) is known to inhibit N₂O reductase but not nitrite reductase (NirK) or nitrous oxide reductase (NOR) activities of nitrifier denitrification pathways in AOB (30, 31). In our ¹⁵N tracer experiments, the decline of NH₂OH-derived ¹⁵N₂O accumulation after addition of 0.1% C₂H₂, indicated that the ¹⁵N₂O is derived from oxidation of ¹⁵NH₃ and not from reduction of ¹⁵NO₂⁻ (Fig. 4A). We also note the formal possibility that exogenously added NH₂OH could be converted chemically or biologically forming short-lived nitrogen radicals, such as nitroxyl (HNO^•), nitrogen monoxide (NO^•), or nitrosyl (NO⁺), all of which under O₂ limitation could undergo further reactions yielding N₂O to varying degrees (32). N₂O generation can also result from biological or chemical conversion of NO₂⁻ and NH₂OH into N₂O (33). However, a role of such cross-reactions involving NH₂OH itself as well as nitrogen-free radicals can also be ruled out because addition of C₂H₂ stops further accumulation of ¹⁵N₂O in total N₂O pool. Together, these results clearly demonstrate that NH₂OH is a product of NH₃ oxidation in N. maritimus.

We are thus left with a unique enzyme system for the oxidation of NH₂OH in the AOA. All data now implicate a variety of unusual proteins, rather than the c-type cytochrome system of NH₂OH oxidation and respiration in the AOB, comprising the remaining oxidative and energy-harvesting steps in the AOA. Apart from providing additional support for the role of a unique biochemistry in global nitrification, continued characterization of this pathway may point to environmental factors, such as specific trace metal availability, limiting or promoting nitrification in different marine and terrestrial provinces.

Methods

Culture Conditions and Cell Harvesting.

All physiological experiments were carried out with N. maritimus strain SCM1 in Hepes-buffered synthetic crenarchaeota medium (SCM) prepared as described (16). C₂H₂ was produced from barium carbide (BaC₂) as described by Hyman and Arp (34). Other reagents, such as NH₄Cl, NH₂OH, and ATU were research-grade products and were obtained from Sigma. N. maritimus cells were grown to midlog phase (∼550–650 µM NO₂⁻ accumulated) and were concentrated by filtration onto nylon membrane filters (Whatman; 47 mm diameter, 0.2 µm pore size; 7402-004) using a vacuum manifold system at 600 mm Hg pressure. The N. maritimus cells harvested in this fashion actively metabolized NH₃ for at least 8 h, enabling short-term assays to be conducted. Multiple liters of N. maritimus cultures could be filtered simultaneously and the nylon membranes with concentrated cells served as solid support for the experimental incubations. The N. maritimus cells on the nylon membranes were incubated in SCM with either NH₄⁺ or NH₂OH added as a substrate.

NH₃- and NH₂OH-Dependent O₂ Uptake Measurements.

Rates of NH₃ and NH₂OH-dependent O₂ uptake of the N. maritimus cells adsorbed onto the nylon membrane filters were measured with a Clark-type O₂ electrode (Yellow Springs Instrument) mounted in an all-glass, water-jacketed reaction vessel (18 mL volume) and operated at 30 °C. All measurements were carried out in NH₄⁺-free SCM media with added NH₄⁺ or NH₂OH (200 µM each).

Determination of NH₂OH, NO₂⁻_, N₂O and Protein Concentration.

NH₂OH in SCM was oxidized to nitrous oxide (N₂O) by the addition of Fe (III) as described (25) and with a minor modification. In our study, the FeAc reagent consisted of 8% (wt/vol) colloidal iron oxide and 6% (vol/vol) glacial acetic acid. This modification adjusted the pH of the sample below 1.4 and assured complete conversion of NH₂OH to N₂O. The FeAc reagent (1 mL) was injected into 20 mL amber glass serum bottles sealed with Teflon caps containing 2 mL of sample and the bottles were mixed thoroughly. After 1 h incubation at 30 °C, saturated NaCl solution (1 mL) was injected into the sealed bottles to displace the N₂O in the liquid phase into the headspace. The bottles were then flash frozen on dry ice and stored upside down until further analysis. The N₂O gas in the headspace was measured by gas chromatography using a thermal conductivity detector. Samples were analyzed before and after incubation with FeAc reagent to determine if any N₂O accumulated during NH₂OH oxidation before incubation with FeAc reagent. The conversion of NH₂OH to N₂O proceeded in a 2:1 stoichiometry and was above 95% in a concentration range of 1 µM to 1 mM NH₂OH. Neither NH₄⁺ nor NO₂⁻ interfered with the conversion and quantification of NH₂OH.

NO₂⁻ concentration was determined with the Griess reagent (sulfanilamide and N-naphthylethylenediamine) as described (35). The rate of NO₂⁻ accumulation has been shown previously to be proportional to the density of N. maritimus cell culture (16) and was used as an indicator of growth stages.

Protein concentrations of N. maritimus cells adsorbed onto the filter were determined with the micro BCA protein assay kit from Pierce after protein solubilization at 60 °C for 60 min.

ATP Measurements.

For ATP measurements, the N. maritimus cultures were grown to midlog phase and filtered using the manifold setup. The filters carrying the N. maritimus cells were immersed in NH₄⁺-free SCM in a serum vial capped with a gray-butyl rubber stopper. The bottles were purged with argon gas and cells were allowed to incubate for 2 h to deplete their endogenous ATP levels, allowing detection of newly synthesized ATP upon addition of the substrates. After 2 h of anaerobic treatment, identical filter pieces were cut and distributed into wells of a white 96-well assay plates (White w/lid, tissue culture-treated; BD Biosciences). The plate wells contained O₂-saturated SCM with (i) no substrate, (ii) 200 µM NH₄⁺, or (iii) 200 µM NH₂OH. ATP accumulation was measured over a time course using a commercial kit based on the luciferase enzyme activity (BactTiter Glo; Promega). To detect ATP generation, 100 µL of the reagent was added per well, incubated 5 min, and the bioluminescence was measured with a multifunction plate reader (Infinite M200; Tecan) with 1-s integration and 10-ms settle time. Statistical differences in ATP values between NH₂OH and NH₄⁺ or no substrate were evaluated using Student’s t test. An appropriate ATP standard curve was used to estimate the concentration of ATP in the samples. Samples for NO₂⁻ determination were collected before addition of ATP reagent and analyzed as described above.

Detection of NH₂OH Formation in N. maritimus Cells.

Natural abundance NH₄Cl and NH₂OH (200 µM each) were added to SCM containing N. maritimus cells adsorbed onto the nylon membrane filters. The bottles were incubated in the dark at 30 °C. Samples were taken at 20-min intervals for determination of NO₂⁻, N₂O concentrations, and isotopic compositions of NH₂OH and NO₂⁻. A total of 20 µM ¹⁵NH₄Cl (Cambridge Isotopes; 99 atom % ¹⁵N) was added at 60-min time point and 0.1% C₂H₂ was added at 120-min time point. NO₂⁻ concentration was determined by Griess reagent as described above.

Total NH₂OH was determined by GC-thermal conductivity detector (TCD) after conversion to N₂O as described above. Isotopic composition of N₂O was determined using a continuous flow ThermoFinnigan DeltaPlus stable isotope ratio mass spectrometer (Thermo) coupled to a Precon (Thermo) and Gasbench II (Thermo) relative to N₂O and NO₃⁻ standards as described previously (36, 37). Even though the δ¹⁵N/¹⁴N in N₂O pools and the AT% ¹⁵N/¹⁴N increased during incubations, the rapid disappearance of NH₂OH caused the µM ¹⁵N-NH₂OH concentrations to decrease over time. Hence µM ¹⁵N-NH₂OH concentrations were calculated from AT % excess ¹⁵N/¹⁴N values. The AT % excess ¹⁵N and µM concentrations of ¹⁵NH₂OH are calculated as defined below:

• i) AT % ¹⁵N excess in a sample = AT % ¹⁵N in labeled NH₂OH pool of sample – AT % natural abundance ¹⁵N in unlabeled NH₂OH pool at the beginning of the experiment; and

• ii) µM ¹⁵N-NH₂OH = (µM total NH₂OH)/(AT % ¹⁵N excess in a sample × 100).