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Ammonia oxidation pathways and nitrifier denitrification are significant sources of N2O and NO under low oxygen availability Xia Zhua,b,c,1, Martin Burgerb,1, Timothy A. Doaneb, and William R. Horwathb,1
aCenter for Ecological Studies, Chengdu Institute of Biology, Chinese Academy of Sciences, Chengdu 610041, China; bBiogeochemistry and Nutrient Cycling Laboratory, Department of Land, Air and Water Resources, University of California, Davis, CA 95616; and cUniversity of Chinese Academy of Sciences, Beijing 100049, China
Edited by Mark H. Thiemens, University of California at San Diego, La Jolla, CA, and approved March 8, 2013 (received for review November 16, 2012)
The continuous increase of nitrous oxide (N2O) abundance in the atmosphere is a global concern. Multiple pathways of N2O pro- duction occur in soil, but their significance and dependence on oxygen (O2) availability and nitrogen (N) fertilizer source are poorly understood. We examined N2O and nitric oxide (NO) pro- duction under 21%, 3%, 1%, 0.5%, and 0% (vol/vol) O2 concen- trations following urea or ammonium sulfate [(NH4)2SO4] additions in loam, clay loam, and sandy loam soils that also con- tained ample nitrate. The contribution of the ammonia (NH3) oxi- dation pathways (nitrifier nitrification, nitrifier denitrification, and nitrification-coupled denitrification) and heterotrophic de- nitrification (HD) to N2O production was determined in 36-h incu- bations in microcosms by 15N-18O isotope and NH3 oxidation inhibition (by 0.01% acetylene) methods. Nitrous oxide and NO production via NH3 oxidation pathways increased as O2 concentra- tions decreased from 21% to 0.5%. At low (0.5% and 3%) O2 con- centrations, nitrifier denitrification contributed between 34% and 66%, and HD between 34% and 50% of total N2O production. Heterotrophic denitrification was responsible for all N2O produc- tion at 0% O2. Nitrifier denitrification was the main source of N2O production from ammonical fertilizer under low O2 concentrations with urea producing more N2O than (NH4)2SO4 additions. These findings challenge established thought attributing N2O emissions from soils with high water content to HD due to presumably low O2 availability. Our results imply that management practices that increase soil aeration, e.g., reducing compaction and enhancing soil structure, together with careful selection of fertilizer sources and/or nitrification inhibitors, could decrease N2O production in agricultural soils.
Nitrous oxide (N2O) and nitric oxide (NO) are key trace gasesthat play major roles in atmospheric chemistry. Nitrous oxide is a significant greenhouse gas contributing to positive radiative forcing and ozone destruction in the stratosphere (1), and NO is a relevant factor in atmospheric photochemistry and air quality issues, particularly tropospheric ozone production (2). Globally, agricultural soils are a major source of anthropogenic N2O and NO emissions (3), and there is little doubt that the use of nitrogen (N) fertilizer and manure is driving the increase in atmospheric N2O (4). Nitrous oxide and NO are produced through the microbial
processes of denitrification and nitrification and through abiotic chemodenitrification reactions (5–7). Heterotrophic denitrification (HD) is performed by heterotrophic bacteria using nitrate (NO3
−) or nitrite (NO2
−) as alternate electron acceptors to oxygen (O2). The first step of nitrification is the oxidation of ammonia (NH3) with ammonia monooxygenase to nitrite (NO2
−) via hydroxylamine (NH2OH) (6). This biotic process, carried out under aerobic con- ditions by chemoautotrophs using NH3 as an energy source (8), can produce N2O and NO by several pathways, namely nitrifier nitri- fication (NN), nitrifier denitrification (ND), and nitrification-cou- pled denitrification (NCD) (Fig. 1).
Previous research on N2O and NO emissions from soil has identified N application rates, fertilizer types, soil moisture, and soil texture as main factors affecting N2O and NO emissions (9, 10). Although the importance of O2 as a controlling factor in regulating the magnitude and pathway of N2O and NO pro- duction has been recognized (6, 11, 12), O2 concentrations are rarely measured, and soil moisture content has generally been accepted as a measurable proxy of O2 availability (13). For ex- ample, optimum conditions for N2O emissions via denitrification have been assumed to exist at water-filled pore space (WFPS) of 70–90% (9, 14), whereas N2O emissions at lower WFPS have often been attributed to nitrification (14–16). However, soil moisture controls not only the diffusion of O2, but also substrate availability (17) and microbial activity. In previous studies, it has therefore been difficult to distinguish between the effects of O2 and substrate availability on N2O production. For these reasons, the role of O2 in regulating N2O and NO production in soil has been challenging to explain (18–20). In pure culture studies, the reduction of NO2
− to NO and N2O by autotrophic NH3 oxidizers (21) occurred at increasing rates as O2 concentration declined (22, 23). Thus, under limiting O2 concentrations ND could contribute significantly to N2O pro- duction in soil (20, 24). However, differentiating between ND and other N2O-producing processes has been methodologically challenging. Acetylene (C2H2) at concentrations of 0.1–10 Pa (0.01% by volume) inhibits NH3 oxidation by autotrophs, such as Nitrosomonas europaea (25, 26), and therefore ND, NN, and NCD; at these concentrations, C2H2 does not inhibit NH3 oxi- dation by heterotrophs (26), which might be the source of N2O under certain conditions such as low pH, high O2 availability, and ample carbon substrates (27), nor does it inhibit nitrous oxide reductase activity (28) (Fig. 1). Wrage et al. (24) conceived a dual isotope (15N and 18O) approach to distinguish the pathway of ND from other N2O-forming processes. Kool et al. (29–31) further refined this method by including 18O-labeled NO3
− to quantify the exchange of oxygen atoms (O) between H2O and N oxides during denitrification and nitrification in soil. Although these methods have elucidated pathways and sources of N2O production, there is still a dearth of knowledge of the environ- mental factors (particularly O2 availability) and substrates (fer- tilizer N source) controlling soil N2O production through ND. Among N fertilizers, which serve as a substrate for NH3 oxi-
dation, urea is the most widely used in agricultural production
Author contributions: X.Z., M.B., and W.R.H. designed research; X.Z. and T.A.D. per- formed research; X.Z. analyzed data; and X.Z., M.B., and W.R.H. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission. 1To whom correspondence may be addressed. E-mail: [email protected], wyjzhu@ ucdavis.edu, or [email protected].
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10. 1073/pnas.1219993110/-/DCSupplemental.
6328–6333 | PNAS | April 16, 2013 | vol. 110 | no. 16 www.pnas.org/cgi/doi/10.1073/pnas.1219993110
(32). Its application, like those of aqueous and anhydrous NH3, leads to a short-term increase in soil pH and soluble organic carbon (33), and may result in the accumulation of NO2
−, which is a substrate contributing to N2O and NO production (20, 34). Additions of the less commonly used acidic-forming fertilizers, such as (NH4)2SO4, have often resulted in lower N2O emissions (10). However, both types of N fertilizers have been shown to produce N2O from nitrification (35), and, in general, to promote denitrification (33). Here, we investigate the influence of fertilizer type and O2
availability on N2O and NO production pathways in three agri- cultural soils varying in texture. To quantify the contribution of NN, ND, NCD, and HD to N2O production under a range of O2 availability, a dual-label 15N-18O isotope method was used in short-term (36-h) incubations of urea- and (NH4)2SO4 - amended soil. We also inhibited NH3 oxidation by 0.01% C2H2 to de- termine the relative importance of the NH3 oxidation pathways and HD to total N2O and NO production. We hypothesized that the majority of N2O released from ammonical fertilizer additions is via the NH3 oxidation pathways and that fertilizer source significantly affects the magnitude of N2O release. We further hypothesized, based on the results of pure culture studies, that O2 regulates both pathways and magnitude of N2O production. The overall objective was to improve understanding of how N2O production following N fertilizer additions in low-O2 soil envi- ronments is regulated. Such information could be used in de- signing agricultural and fertility management with the goal of mitigating N2O emissions from agriculture practices.
Results The contribution of the NH3 oxidation pathways and HD to N2O production under ambient and low (≤3% by volume) O2 was determined by two independent methods. In these methods, NO3
− and NH4 + as either urea or (NH4)2SO4 were present in the
soil in equal quantities in the fertilized treatments. By the first method, a 15N-18O isotope pool dilution and tracer
experiment in clay loam soil, N2O production via the different pathways, gross nitrification rates, and N2O generated as per- centage of NO3
−-N produced were determined. The quantities of N2O produced increased 19-fold as O2 concentration de- creased from 21% to 3% (Fig. 2). At low (3% and 0.5%) O2 concentrations, the contribution of ND to total N2O production ranged from 48% to 66% in urea- and from 34% to 57% in (NH4)2SO4-amended soil, whereas HD contributed 34–44% and 43–50% to total N2O production in urea- and (NH4)2SO4-
amended soil. At these low O2 levels, NN contributed 0–18% and NCD 0–6% to total N2O production. Between 0.5% and 0% O2, N2O production increased threefold to sixfold. In the absence of O2, HD was the only source of N2O. None of the 15N-labeled N2O was further reduced to N2 (Table S1). Gross nitrification rates decreased significantly as O2 con-
centration decreased, and no nitrification was measured under anaerobic conditions (0% O2) (Table 1). At ambient O2 con- centrations, gross nitrification rates were higher in the urea than the (NH4)2SO4 treatment, but these rates were not different between urea and (NH4)2SO4 at subambient O2 concentrations. As the O2 concentration decreased from 21% to 0.5%, the percentage of N2O-N emitted per unit NO3
−-N produced in- creased from 0.1% to 8.3% in urea- and from 0.08% to 6.9% in (NH4)2SO4-fertilized soils (Table 1). Overall, more N2O was produced from urea than from (NH4)2SO4 (P < 0.01). By the second method, 0.01% (vol/vol) C2H2 inhibition of
autotrophic NH3 oxidation in loam, clay loam, and sandy loam soils, N2O and NO production via the NH3 oxidation pathways was calculated from the difference in N2O and NO yields be- tween treatments with (+) and without (−) 0.01% (vol/vol) C2H2. According to the terminology of this article, the NH3 oxidation pathways of N2O and NO production include N2O and NO generated through NN, ND, and NCD. Nitrous oxide and NO produced in +C2H2 was assumed to be due to HD. Production of N2O increased in all fertilizer-amended soils as headspace O2 concentration decreased from 21% to 0% and was 20, 7, and 12 times greater in the 0% than the 0.5% O2 treatments in loam, clay loam, and sandy loam soils, respectively (Fig. 3). At 0% O2 headspace concentration, HD was the sole pathway of N2O production. However, when headspace O2 concentration was between 0.5% and 3%, in all but one treatment [sandy loam soil fertilized with (NH4)2SO4 under 0.5% O2], at least 50% of total N2O production was via the NH3 oxidation pathways. At O2 concentrations >0%, the amount of N2O produced in loam and clay loam soils fertilized with urea was significantly higher than with (NH4)2SO4 (Fig. 3). In the sandy loam soil, N2O production was similar between the two fertilizer treatments.
NH3 0.01% v/v C2H2 blocks this stepO2
NH2OH N2O/NON2O/NO H2O
--------
†NO2 -
NO - NO - NO N O N
NO N2O N2N2O/NO
H2O
†------
NO3 NO2 NO N2O N2
Ammonia oxida�on, by autotrophic NH3 -oxidizers
d f d
Nitrite oxida�on, by NO2 --oxidizers
Denitrifica�on by autotrophic NH -oxidizers
O2>0%
O ≥0%
Nitrifier Nitrifica�on, NN
Nitrifier Denitrifica�on ND
------ Byproduct of ammonia oxida�on
Denitrifica�on, by autotrophic NH3-oxidizers
Denitrifica�on, by denitrifiers
O2≥0%
O2<0.2%
Nitrifier Denitrifica�on, ND
Nitrifica�on-Coupled Denitrifica�on, NCD
Chemodenitrifica�on, CDSoil pH< 5.47 or Fe2+ or Cu2+ or Mn2+ Heterotrophic Denitrifica�on, HD
†: To date, this process a ributed only to the autotrophic nitri er-Nitrosomonas sp
Fig. 1. The main pathways of nitrous oxide (N2O) and nitric oxide (NO) production in soil.
0% 20% 40% 60% 80% 100%
3 %
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3 %
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co nc
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at io
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(NH4)2SO4
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48
0 11.5 48.4
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0 2.8 3.5
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29.4 ng N g-1
568 ng N g-1
708 ng N g-1
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311 ng N g-1
440 ng N g-1
27.3 34.4
0
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43.9
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100 0 %
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27.3 62.7
10 0
0 43.1
50.30 3.5
100
0% 20% 40% 60% 80% 100%
2146 ng N g-1
2683 ng N g-1
HD % of total N2O
Fig. 2. Relative contributions of nitrifier denitrification (ND), nitrifier nitrification (NN), nitrification-coupled denitrification (NCD), heterotrophic denitrification (HD) to N2O production in clay loam soil.
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In all three soils, the N fertilizers promoted NO production (P < 0.01) (Fig. 4). At 0% O2 concentration, HD was the sole source of NO produced from fertilized soils. However, at O2 concentrations >0%, the primary source of NO was by NH3 oxidation pathways, which accounted for 72–97% of the total NO production from N-fertilized soils.
Discussion Our study shows that ND is the main pathway of N2O production from ammonical fertilizer applications in soil under limited O2 availability (Fig. 2). According to the results of our 15N-18O ex- periment, ammonical fertilizer was the main source of the ND- generated N2O, whereas NO3
−, present in equal abundance as the initial ammonical fertilizer N, provided the substrate for the N2O produced by HD, as shown by the N2O-
15N isotopic data (Table S1). The NN and NCD pathways were relatively minor sources of N2O under subambient O2 concentrations. The results obtained by the NH3 oxidation inhibition method
showed similar trends with regard to total N2O production at the different O2 levels among the three soils and in comparison with the 15N-18O isotope experiment in clay loam soil. Values of total N2O production (±SE) in clay loam soil [630±230 and 220±56 ng N2O-N·g
−1 after urea and 710±87 and 72±25 ng N2O-N·g −1
after (NH4)2SO4 fertilization at 0.5 and 3% O2, respectively] in the –C2H2 treatment agreed reasonably well with those of the isotope experiment (Fig. 2). At 0% O2, both methods showed that HD alone was responsible for N2O production. However, the amount of N2O attributed to HD in clay loam soil at low (0.5% and 3%) O2 concentrations in the presence of 0.01% C2H2 was, in general, lower (17±1.2 − 230±16 ng N2O-N·g
−1) than the HD contribution to total N2O determined by the 15N-18O isotope method (130±50 − 310±22 ng N2O-N·g
−1), and this increased the estimate of N2O production via the NH3 oxi- dation pathways in the NH3 oxidation inhibition approach. Thus, the extent of the contribution of NH3 oxidation pathways to total N2O production as calculated in the
15N-18O experiment can be regarded as the more conservative measure than the one determined by the NH3 oxidation inhibition approach. Emissions of N2O produced via NH3 oxidation pathways in
response to ammonical fertilizer additions can be significant. Many field experiments have shown that the bulk of seasonal N2O emissions occurs shortly after fertilizer N applications (36, 37) when N2O production through the NH3 oxidation pathways is most likely because almost all synthetic N fertilizers supply N in the ammonical form. Our results support those of recent analyses of global trends of the N2O molecule’s
15N, 18O, and site pref- erence (δ15Nα-δ15Nβ) signature that point to the contribution of ammonical N fertilizers and possibly nitrification as the principal sources responsible for the rise in atmospheric N2O (38).
Effect of O2 Availability on N2O and NO Production. The significance of the NH3 oxidation pathways leading to N2O production has been poorly understood partly because the influence of different N fertilizer sources and oxygen availability has not been well quantified. The role of O2 in evoking the various N2O and NO production pathways has been predicated based on the easily measured metric of WFPS. However, WFPS does not take into account the distribution of macropores and micropores, which is controlled by soil texture, and therefore is not adequate to estimate true O2 availability. In field and laboratory experiments, nitrification has often been
presumed to be the major source of N2O at low soil moisture contents, and HD, the major source at high soil moisture contents (14–16, 39). However, our results show that as O2 availability decreased, N2O production from NH3 oxidation pathways in- creased, which is contrary to the established thought on the reg- ulation of N2O production by WFPS. Our results confirm the conclusions of pure culture studies (22) and results from soil microcosm studies showing a ninefold increase in N2O produced per unit NH3 oxidized with a decrease in O2 concentration from 20% to 0.8% (19). In the present experiment, gross nitrification rates decreased significantly as O2 concentration decreased, but concurrently, the amount of N2O generated per unit NO3
−-N
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Fig. 3. Total nitrous oxide (N2O) produced at different levels of headspace oxygen (percentage O2) over 36 h after application of fertilizers to soils. For each soil (A–C), different uppercase letters indicate a significant difference in total N2O emission across all treatments. Different lowercase letters in- dicate a significant difference in the proportion of N2O derived from am- monia oxidation.
Table 1. Soil gross nitrification rates and the percentage of nitrous oxide (N2O) emitted per unit nitrate (NO3
−) produced in clay loam soil at the end of the experiment
Oxygen content
Nitrification rate, ng N·g−1·h−1
Percentage of N2O emitted per unit NO3
− produced
Urea (NH4)2SO4 Urea (NH4)2SO4
21% 560Aa (17) 360Ba (26) 0.11Ac (0.00) 0.080Bc (0.010) 3% 260Ab (220) 200Ab (63) 2.9Ab (2.2) 0.52Bb (0.22) 0.5% 120Ac (13) 99Ac (7.1) 8.3Aa (1.1) 6.9Aa (1.1) 0% 0.00 (0.00) 0.00 (0.00) 0.00 (0.00) 0.00 (0.00)
SDs are shown in parentheses. For each N transformation variable, differ- ent uppercase letters indicate a significant difference between fertilizer treatments within each oxygen (O2) level, whereas different lowercase let- ters indicate significant differences among O2 levels (P < 0.05). n = 4. Two significant figures are shown.
6330 | www.pnas.org/cgi/doi/10.1073/pnas.1219993110 Zhu et al.
produced increased, and the absolute amount of N2O produced by ND increased 81-fold as O2 concentration decreased from 21% to 0.5%. Thus, NH3 oxidation via ND can be a significant source of N2O production—especially in low-O2 soil environments. Our results suggest that the source of most of the NO present
at subambient O2 concentrations was ammonical fertilizer. In the +C2H2 treatments of the clay loam and sandy loam soil, a small amount of NO, which was probably produced during the short time span (∼20 min) between fertilizer application and head- space treatment imposition, was measured at 4 h (Fig. S1). At later sampling times in the +C2H2 treatments, the NO had likely been consumed. Analogous to N2O production, at 0% O2, none of the NO was derived from NH3 oxidation. However, the in- crease in NO production at 0% O2 was not as pronounced nor as consistent as that of N2O because N2O was probably a sink for NO (34). Unlike NO, N2O was not reduced further even under 0% O2 as evidenced by the N2 isotopic data (Table S1), most likely because in the present experiment denitrifiers were pro- vided with ample NO3
−, a preferred electron acceptor over N2O (6, 40). Although HD-related N2O production at 0% O2 was three to nine times higher than that occurring through NH3 oxidation and HD pathways in low O2 environments, our results
suggest that only totally anoxic conditions in the presence of NO3
− will lead to such elevated N2O production from HD. It should also be noted that the relatively low water content [50% water holding capacity (WHC)] of the experimental conditions may have resulted in less opportunity for N2O consumptive processes via denitrification at 0% O2 than a soil with a higher water content would.
Effect of Fertilizer Type and Soil Texture on N2O and NO. Ammonium has previously been identified as the main source of N2O after ammonical fertilizer additions (41), but the conditions regulating the N2O production pathways remain a subject of debate and investigation (9, 18), and the interactive effects of the soil envi- ronment and fertilizer N sources on N2O production have not been systematically studied. In the present study, urea fertilizer increased N2O and NO production at >0% O2 concentration compared with (NH4)2SO4 (Figs. 3 and 4). The production of N2O per unit NO3
− produced was 1.2–5.5 times higher in the urea than the (NH4)2SO4 treatments. This greater loss as N2O in the urea than (NH4)2SO4 treatment is in agreement with the findings of metaanalyses of field experiments (10, 42). According to our results, ND was the pathway responsible for the higher production of N2O in the urea-amended soil. Nitrous oxide de- rived from HD was similar between N fertilizer treatments, which is consistent with our assertion based on the 15N results that NO3
−, rather than the ammonical fertilizers, was the source of N2O at 0% O2. The greater N2O and NO production in loam and clay loam
soils treated with urea also produced higher soil NO2 − concen-
trations compared with (NH4)2SO4-treated soil (Table S2). Hy- drolysis of urea increases soil pH (Table S2), which can lead to NO2
− accumulation due to impaired Nitrobacter spp.-mediated nitrite oxidation (43). In pure culture studies with nitrifying bac- teria, N2O production at low O2 levels has been shown to be correlated with an increase in NO2
− (44) suggesting that the ac- cumulation of NO2
− in our soils, as in the pure cultures, promoted ND. Other workers have shown nitrite to promote N2O and NO production via abiotic reactions, such as with metals and with organic matter, especially in acidic soils (5, 20, 34). The N2O derived from these processes, termed chemodenitrification, would have the same isotopic signature as that generated by ND because NO2
− is the source of N2O in both processes. However, our experiment with sterilized soil (SI Materials and Methods) showed that the N2O derived from chemodenitrification com- prised only 0.1–1.3% of the total N2O production (Table S3). In the sandy loam soil, the originally high soil pH (Table S4) may explain why there were no differences in production of NO, N2O, and NO2
− between urea and (NH4)2SO4-amended soil at O2 concentrations >0%. Nitrous oxide production was greater in the clay loam than in the loam and sandy loam soils, which likely relates to greater soil nitrifier populations and nitrification rate (45). Nitrifier populations inhabiting clay surfaces have been shown to be protected from the effects of H+ produced from NH3 oxidation (46). This study showed that N fertilizers, the vast majority of which
supply N in the form of NH4 +, promote N2O and NO production
and that ND is the dominant pathway of N2O production from ammonical fertilizer application at O2 concentrations ≥0.5%. The results demonstrate how O2 concentration and fertilizer N source regulate the magnitude and pathways of N2O production following N fertilizer applications. The findings also explain why the size of NO3
− pools alone is, generally, a poor predictor of N2O emissions because the NH3 oxidation pathways contribute a significant portion to total N2O production under low O2 avail- ability. Moreover, our results readily explain why N2O emissions for a given cropping season are often highest following ammon- ical fertilizer additions. The results of this research can be used to design management practices that reduce N2O production from
(NH4)2SO4
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Fig. 4. Total nitric oxide (NO) at different levels of headspace oxygen (percentage O2) 4 h after application of fertilizers to soils. For each soil (A–C), different uppercase letters indicate a significant difference in total NO production across all treatments, and different lowercase letters indicate a significant difference in the proportion of NO derived from ammonia oxidation.
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SC IE N C ES
NH3 oxidation pathways during this critical period and to revise process-based models by including N2O production via NH3 oxidation pathways in low-O2 environments. Refocusing research on N2O production through NH3 oxidation pathways will bring forth guidelines for careful selection of fertilizer source and/or nitrification inhibitors (47) suitable for particular soil properties and environmental conditions. Specifically, applications of urea should be avoided in soil conditions prone to low O2 availability and in low-pH soils. In general, agricultural practices that main- tain aerobic soil conditions, e.g., by reducing soil compaction and enhancing soil structure through organic matter management, are likely to mitigate N2O emissions following N fertilizer additions.
Materials and Methods Soil Samples. Soil samples were collected from the 0- to 15-cm layer of three agricultural fields in the Sacramento Valley and Central Coast area in Cal- ifornia (Table S4). Composite soil samples from numerous auger borings were sieved to 2 mm, and refrigerated (4 °C) until the experiment began. The soils include a loam, a sandy loam, and a clay loam. The soils were preincubated at 40% WHC for 7 d. The purpose of the preincubation was to avoid the pulse of respiration associated with wetting dry soils (48).
Experiment I: 15N-18O Experiment. Experimental setup. The preincubated clay loam soil was used to measure gross nitrification rates and N2O production derived from NH3 oxidation pathways (NN, ND, and NCD) and HD. At the beginning of the incubation, 5-g (dry weight) preincubated soil samples were placed into 60-mL glass serum vials (Supelco) and four different treat- ments for each ammonical fertilizer [(NH4)2SO4 or urea] were established. The following treatments enriched in 18O and 15N were applied: (i) 18O−H2O + NO −3 + NH
+ 4 , (ii) H2O +
18O−NO −3 + NH + 4 , (iii) H2O +
15NO3 + NH + 4 , and (iv)
H2O + NO − 3 +
15NH +4 . All treatments received (NH4)2SO4 or urea and KNO3 to bring the concentrations of mineral N in soils to 50 mg of NH4
+-N and 50 mg of NO3
−-N kg−1 soil and were incubated at 50% of WHC at 22 °C. The labeled 15N compound was added to bring the NH4
+ and NO3 − pools to an
enrichment of 6.0 atom% 15N excess; the labeled 18O compound was added to bring the H2O and NO3
− pools to an enrichment of 1.0 atom% 18O excess. Sample vials were crimp top sealed following the 15N- and 18O-labeled additions and immediately adjusted to four different levels of headspace O2 contents (21%, 3%, 0.5%, and 0% by volume). The O2 treatments were applied after evacuating the headspace in each vial three times to <0.1 bar and flushing with 100% ultra–high-purity helium in between evacuations. Aliquots of pure O2 were injected to achieve the desired O2 levels after re- moving the same volume of gas from the headspace before injection. The experiment was set up as a fully randomized design, with four replicates per treatment. Gas sampling and analysis for 15N and 18O. Gas samples were removed from the headspace of the vials at the end of the incubation at 36 h and transferred to exetainers (Labco) that were flushed with helium and evacuated before use. Gas samples were analyzed at the University of California, Davis, Stable Isotope Facility. The concentration of N2O and its
15N and 18O signature were measured on a ThermoFinnigan GasBench + PreCon trace gas concentra- tion system interfaced to a Thermo Scientific Delta V Plus isotope-ratio mass spectrometer. Soil extraction and mineral 15N analysis. After removal of headspace gas samples at 36 h, all treatment soils were extracted with 1 M KCl (10:1 extractant volume-to-soil mass ratio). The extracted soils were centrifuged at 13,000 × g for 20 seconds and the supernatant was frozen at −20 °C. Isotope analysis of soil mineral N was performed on aliquots of the extracts using a diffusion technique (49). The 15N isotopic analyses were performed at the University of California, Davis, Stable Isotope Facility on a PDZ Europa ANCA-GSL ele- mental analyzer interfaced to a PDZ Europa 20–20 isotope ratio mass spec- trometer (Sercon). Calculations. We calculated the N2O produced by each pathway based on the 18O and 15N isotopic enrichment of the accumulated N2O, the
18O signature in H2O and NO3
−, and the 15N signatures of NH4 + and NO3
−. The contribution of NH3 oxidation pathways and HD to N2O production was calculated based on 15N data following a modified method reported by Stevens et al. (50) and Senbayram et al. (36). Then, the contribution of the different NH3 oxidation pathways (NN, ND, and NCD) to N2O production was calculated based on the combined 18O and 15N data according to the dual-isotope method of Kool et al. (29, 31). The method is based on the principle that N2O produced via NN, ND, and NCD obtains 0, one-half, and two-thirds of the O from H2O, respectively (24, 51) (Fig. 1) and also takes O exchange between N oxides and
H2O (52, 53) into account. Briefly, the theoretical O incorporation from H2O into N2O is compared with the measured actual O incorporation from H2O into N2O based on certain assumptions regarding the relative contributions of NN, ND, NCD to N2O production and the occurrence of O exchange be- tween N oxides and H2O, which in our experiment was between 70% and 89% (Table S1). Depending on the outcome of this comparison, the assump- tions regarding the NN, ND, and NCD contributions to N2O production are evaluated (i.e., maintained or rejected), and the minimum and maximum contributions of each pathway to N2O production are determined.
The gross nitrification rates (n) were calculated by the 15N isotope pool dilution method (54). The N2O produced by the NH3 oxidation pathways expressed as a percentage of the produced NO3
− (N %) was calculated as follows:
N% = 100 * N2OðNH3Þ * N2Ototal=Nn;
where N2O(NH3) is the N2O-N generated by NH3 oxidation pathways and Nn is the gross nitrification rate multiplied by the incubation time.
Experiment II: Inhibition of NH3 Oxidation by 0.01% C2H2. Experimental setup. Preincubated 5-g (dry weight) samples of the loam, clay loam, and sandy loam soils were placed into 60-mL glass serum vials. Each soil received three fer- tilizer treatments: none (control), urea, or (NH4)2SO4, applied at the rate of 50 mg fertilizer-N·kg−1 soil, which is equivalent to about 75 kg N·ha−1. The fertilizer solution or deionized water (control) was sprayed onto the surface of the soil by using a syringe with a fine-tipped needle to achieve a soil moisture content of 50% WHC with uniform distribution. To eliminate the substrate limitation for HD, nitrate as KNO3 was applied to fertilizer treat- ments to bring NO3
− concentrations to 50 mg N·kg−1 in each soil. All of the additions were carried out at 21%, 3%, 1%, 0.5%, and 0% O2. A separate set of treatments received C2H2 at 0.01% (vol/vol) at the same time as O2. The incubation conditions were the same as in the 15N-18O experiment. The ex- periment was set up as a completely randomized design with 90 treatments, each with three replicates. Gas sampling and analysis. For N2O analysis, headspace gas was removed with syringes at 36 h after the onset of the experiment because the N2O pro- duction in −C2H2 treatments remained linear over the measurement period, whereas no significant change was observed in +C2H2 soils (except for the 0% O2 treatment) (SI Discussion; Fig. S2). The gas samples were transferred to evacuated exetainers previously flushed with helium and were then an- alyzed by a gas chromatograph (HP6890; Hewlett Packard) with a 63Ni electron capture detector and a 2-m HayeSep Q column (Supelco). The sys- tem was calibrated with 10 N2O standards prepared by diluting a commer- cial standard of 2,009 ppmv N2O (Airgas) in pure He gas (coefficient of determination r2 = 0.99). For NO analysis, 0.25-mL headspace samples were removed from each vial using a gas-tight syringe at 4 h after the onset of the experiment. Aliquots of 0.1- or 0.25-mL samples and certified NO stan- dard gases (Scott-Marrin) were injected into a NO-free gas stream flowing through a chromium trioxide column converting NO to NO2 and then through a Scintrex LMA-3 luminol chemiluminescence detector (Unisearch Associates). The chemiluminescence detector showed no interference with 0.01% (vol/vol) C2H2 in preliminary tests.
To verify headspace O2 levels, the O2 concentrations in the C2H2-treated and control sets were measured at 0, 24, and 36 h in 0.5-mL aliquots by an O2 analyzer (S-3A/I; Applied Electrochemistry). The analyzer has an accuracy of 0.01% (vol/vol) O2. Results showed that the change in headspace O2 during the 36-h incubation was <10% of the target O2 concentration. Soil extraction and mineral N analysis. Soil was extracted after gas sampling as described above except that the supernatants were stored at 4 °C. Nitrite analysis was conducted immediately following the extraction, and NH4
+ and NO3
− analyses were performed within 24 h after extraction using colori- metric methods (55, 56). Calculations. Nitrous oxide and NO produced in +C2H2 were attributed to HD, whereas N2O and NO generated via the NH3 oxidation pathways was calculated by subtracting the N2O produced in +C2H2 from that produced in –C2H2. Statistical analysis. Analysis of variance was performed using PROC MIXED procedures in SAS (SAS 9.2; SAS Institute). Fertilizer and O2 concentration were treated as fixed and replicate as random factors. Nitrous oxide and NO production data were natural log transformed for the statistical analysis due to the nonnormal distribution of residuals of the untransformed data, as evaluated by Shapiro–Wilk test. Means separation was performed using pdiff when ANOVA showed significant effects (α < 0.05).
6332 | www.pnas.org/cgi/doi/10.1073/pnas.1219993110 Zhu et al.
ACKNOWLEDGMENTS. We gratefully acknowledge the assistance of those who collected the soil samples. We thank Dennis E. Rolston and Benjamin Z.
Houlton for their comments on earlier versions of this manuscript. We thank the J. G. Boswell Endowed Chair in Soil Science for supporting this research.
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Zhu et al. PNAS | April 16, 2013 | vol. 110 | no. 16 | 6333
A G R IC U LT U R A L
SC IE N C ES
ShawPaper.pdf
Environmental Microbiology (2006)
8
(2), 214–222 doi:10.1111/j.1462-2920.2005.00882.x
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd
Blackwell Science, LtdOxford, UKEMIEnvironmental Microbiology 1462-2912Society for Applied Microbiology and Blackwell Publishing Ltd, 20058
2214222
Original Article
Nitrifier denitrification by Nitrosospira spp.L. J. Shaw
et al.
Received 17 February, 2005; accepted 3 June, 2005. *For correspon- dence. E-mail [email protected]; Tel. (
+
44) 1224 272691; Fax (
+
44) 1224 272703.
Nitrosospira
spp. can produce nitrous oxide via a nitrifier denitrification pathway
Liz J. Shaw,
1
Graeme W. Nicol,
2
Zena Smith,
2
Jon Fear,
1
James I. Prosser
2
and Elizabeth M. Baggs
2
*
1
Imperial College London, Wye Campus, Department of Agricultural Sciences, Wye, Kent TN25 5AH, UK.
2
School of Biological Sciences (Plant and Soil Science), University of Aberdeen, Cruickshank Building, St Machar Drive, Aberdeen AB24 3UU, UK.
Summary
Nitrous oxide (N
2
O) emission from soils is a major contributor to the atmospheric loading of this potent greenhouse gas. It is thought that autotrophic ammo- nia oxidizing bacteria (AOB) are a significant source of soil-derived N
2
O and a denitrification pathway (i.e. reduction of NO
2 –
to NO and N
2
O), so-called nitrifier denitrification, has been demonstrated as a N
2
O pro- duction mechanism in
Nitrosomonas europaea.
It is thought that
Nitrosospira
spp. are the dominant AOB in soil, but little information is available on their ability to produce N
2
O or on the existence of a nitrifier den- itrification pathway in this lineage. This study aims to characterize N
2
O production and nitrifier denitrifica- tion in seven strains of AOB representative of clusters 0, 2 and 3 in the cultured
Nitrosospira
lineage.
Nitrosomonas europaea
ATCC 19718 and ATCC 25978 were analysed for comparison. The aerobically incu- bated test strains produced significant (
P
<<<<
0.001) amounts of N
2
O and total N
2
O production rates ranged from 2.0 amol cell
----
1
h
----
1
, in
Nitrosospira tenuis
strain NV12, to 58.0 amol cell
----
1
h
----
1
, in
N. europaea
ATCC 19718.
Nitrosomonas europaea
ATCC 19718 was atyp- ical in that it produced four times more N
2
O than the next highest producing strain. All AOB tested were able to carry out nitrifier denitrification under aerobic conditions, as determined by production of
15
N-N
2
O from applied
15
N-NO
2 –
. Up to 13.5% of the N
2
O pro- duced was derived from the exogenously applied
15
N- NO
2 –
. The results suggest that nitrifier denitrification could be a universal trait in the betaproteobacterial AOB and its potential ecological significance is discussed.
Introduction
Nitrous oxide (N
2
O) is a greenhouse gas with 296 times the global warming potential of carbon dioxide (Houghton
et al
., 2001). Nitrous oxide also contributes to the destruc- tion of the stratospheric ozone layer (Conrad, 1996). Atmospheric concentrations of N
2
O have been increasing from pre-industrial values of 0.27 ppmv to current concen- trations that approach 0.32 ppmv (Conrad, 1996). Both natural and agricultural soils are major sources of N
2
O (Conrad, 1996; Houghton
et al
., 2001) and have recently been estimated to contribute 57% to the total global N
2
O budget (Mosier
et al
., 1998; Kroeze
et al
., 1999). It is thought that soil sources have contributed to the post- industrial revolution upturn in atmospheric N
2
O loading through increased soil nitrogen availability as a result of nitrogen deposition and increased nitrogen fertilization and biological nitrogen fixation, through agricultural expansion and intensification (Nevison and Holland, 1997).
Microbial transformations of ammonium and nitrate are considered the main processes responsible for gaseous nitrogen emissions from soil. However, chemodenitrifica- tion may be important, particularly in acid soils (VanCle- emput and Samater, 1996). The best characterized biotic pathway of N
2
O production is heterotrophic denitrification in bacteria [reviewed in detail by Zumft (1997)], whereby, under conditions of low oxygen tension, nitrogen oxides serve in place of dioxygen as terminal electron acceptors in electron transport phosphorylation. Heterotrophs pos- sessing the complete denitrification pathway produce N
2
O as an intermediate in the stepwise dissimilatory reduction of nitrate to dinitrogen gas (NO
3 –
Æ
NO
2 –
Æ
NO
Æ
N
2
O
Æ
N
2
) with oxidation of an organic carbon source used as reducing power (Hochstein and Tomlinson, 1988). Thus denitrification completes the soil nitrogen cycle by return- ing dinitrogen gas to the atmosphere.
The ammonia oxidizing bacteria (AOB) also play a central role in the nitrogen cycle. They catalyse the first step in nitrification, the conversion of ammonia via hydroxylamine to nitrite. As chemolithoautotrophs, they use ammonia as a sole source of energy and reducing power and obtain carbon for biosynthesis by fixation of CO
2
. Pure culture studies, mainly conducted with
Nitrosomonas europaea
strains (Ritchie and Nicholas, 1972; Hynes and Knowles, 1984; Poth and Focht, 1985; Anderson
et al
., 1993; Kester
et al
., 1997; Beaumont
Nitrifier denitrification by
Nitrosospira
spp.
215
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd,
Environmental Microbiology
,
8
, 214–222
et al
., 2002), but also with
Nitrosospira
spp. (Goreau
et al
., 1980; Remde and Conrad, 1990; Jiang and Bakken, 1999a; Dundee and Hopkins, 2001; Wrage
et al
., 2004a), have shown that AOB also produce N
2
O. In fact, it has been estimated that ammonia oxidizers can contribute up to 80% to total soil N
2
O emissions, depending on soil type, temperature and water content (Webster and Hopkins, 1996; Godde and Conrad, 1999).
The mechanism of N
2
O production by nitrifiers is not completely characterized although traditionally two differ- ent routes have been proposed. The first relates to the activity of hydroxylamine oxidoreductase (HAO), which mediates the conversion of hydroxylamine to nitrite, the second step in ammonia oxidation. It has been shown,
in vitro
, that HAO can also catalyse the oxidation of hydroxylamine to N
2
O (Hooper and Terry, 1979), possibly indirectly through the decomposition of unstable interme- diates (Ritchie and Nicholas, 1972). The second N
2
O- yielding route relates to the existence of a denitrification pathway, so-called nitrifier denitrification, where nitrite is reduced to nitric oxide and N
2
O (Wrage
et al
., 2001; Arp and Stein, 2003), as in the classical heterotrophic denitri- fication pathway. Recently, Schmidt and colleagues (2004) have identified denitrification as the major source of N
2
O produced by
N. europaea
ATCC 19718, although mutants lacking the denitrification pathway released N
2
O during oxidation of ammonia to nitrite (most likely as a result of auto-oxidation and chemodenitrification of hydroxylamine).
Several authors have considered the likely contribution of nitrifier denitrification to the total N
2
O emission from soil and the factors that influence its relative importance, for example, local ammonium and oxygen concentrations
(Robertson and Tiedje, 1987; Webster and Hopkins, 1996; Wrage
et al
., 2001, 2004b). However, the funda- mental determinant of intrinsic nitrifier denitrification potential in soil will be the abundance of ammonia oxidiz- ers with denitrifying ability (Wrage
et al
., 2001). The nitrifier denitrification pathway has only been unequivo- cally demonstrated in
Nitrosomonas
spp. (Colliver and Stephenson, 2000), whereas there is mounting evidence that nitrosomonads are not representative of AOB groups found in soil (Arp and Stein, 2003). Specifically, analysis of clone libraries suggests that betaproteobacterial AOB in the
Nitrosospira
lineage may be most abundant (Stephen
et al
., 1996; Kowalchuk and Stephen, 2001; Smith
et al
., 2001). To our knowledge, it is not known whether nitrifier denitrification is universal among AOB or confined to the nitrosomonads. The aim of this study therefore was to determine N
2
O production and nitrifier denitrification potential in AOB cultures more representa- tive of phylotypes dominating soil AOB communities.
Results
Nine different cultures of AOB (Table 1) were incubated under ambient air concentrations of O
2
in medium con- taining ammonium and nitrite at initial concentrations of 3.5 mM and 1 mM, respectively, and the production of N
2
O and nitrite was quantified during incubation for 8 h. Total (
14
+
15
N) N
2
O production was quantified and the ability of the AOB strains to produce
15
N-N
2
O by reduction of exogenously applied
15
N-NO
2 –
during nitrifier denitrifica- tion was determined. Results are exemplified with assay data obtained with
Nitrosospira
sp. strain En13, because this strain has not previously been the subject of physio-
Table 1.
Details and 16S-rRNA gene-based phylogenetic affiliation of AOB strains used in this study.
AOB strain NCBI accession number for 16S rRNA gene sequence; isolation details (if known) and available literature references
a
16S-rRNA gene – based phylogenetic cluster/lineage
b
Nitrosomonas europaea
ATCC 19718
BX321856; Isolated from soil; Source: H. J. E. Beaumont, Vrijie University, Amsterdam, the Netherlands (Beaumont
et al
., 2002, 2004a,b; Chain
et al
., 2003)
Nitrosomonas europaea
/
Nitrosococcus mobilis’
lineage
Nitrosomonas europaea
ATCC 25978
NB070982; Isolated by soil enrichment; (Lewis and Pramer, 1958).
Nitrosomonas europaea
/ ‘
Nitrosococcus mobilis’
lineage
Nitrosospira briensis
strain 128 L35505; (Teske
et al
., 1994)
Nitrosospira
lineage, cluster 3
Nitrosospira multiformis
ATCC 25196
L35509; Isolated from soil, Surinam, South America; (Watson
et al
., 1989).
Nitrosospira
lineage, cluster 3
Nitrosospira tenuis
strain NV12 M96405
Nitrosospira
lineage, cluster 3
Nitrosospira
sp. strain 40KI X84656; Isolated from cultivated clay loam soil (pH 6.5), Norway; (Jiang and Bakken, 1999a,b).
Nitrosospira
lineage, cluster 0
Nitrosospira
sp. strain B6 X84657; Isolated from pebbles (pH 6.3), nitrifying reactor, sewage treatment plant, Norway (Jiang and Bakken, 1999a,b).
Nitrosospira
lineage, cluster 2
Nitrosospira
sp. strain En13 Isolated from Craibstone soil (pH 4.5), Aberdeen, UK; This study.
Nitrosospira
lineage, cluster 3
Nitrosospira
sp. strain NpAV Y10127; Isolated from soil, Apple Valley, Minnesota; (McCarty
et al
., 1991)
Nitrosospira
lineage, cluster 3
a.
Unless otherwise stated, the immediate source of the strains was the AOB culture collection at the University of Aberdeen, UK.
b. According to the classification proposed by Purkhold and colleagues (2003). Not all 16S rRNA gene sequences of the strains used in the present study were included in the analysis of Purkhold and colleagues (2003). Accordingly, the 16S-rRNA gene-based phylogenetic tree of Purkhold and colleagues (2003) was reconstructed to include sequences for those strains not previously analysed. The resulting tree is available as Supplementary material (Fig. S1).
216 L. J. Shaw et al.
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 214–222
logical characterization. The results of a typical assay are shown in Fig. 1. Over the time course, significant (P < 0.001) N2O (Fig. 1A) and nitrite (Fig. 1B) were pro- duced in comparison to the non-inoculated control. All other strains tested also produced concentrations of N2O significantly above (P < 0.05) control levels. Nitrosospira sp. B6 was found to be contaminated with nitrite oxidizers, as determined by detection of nitrate in the growth media and successful polymerase chain reaction (PCR)-amplifi- cation of 16S rRNA gene fragments using nitrite oxidizer- specific primers (T. Freitag and J.I. Prosser, unpublished). Therefore, this strain was not used for quantitative analy- sis of N2O and
15N-N2O production rates, but the ability of this co-culture to produce 15N-N2O from
15N-NO2 – was
assessed. For the remaining eight strains, the kinetics of both cumulative N2O and nitrite production could be esti- mated using a linear function (see Fig. 1A and B for strain
En13; R2 ≥ 0.92, P < 0.01 in all cases), the exception being nitrite production by Nitrosospira sp. 40KI, for which the rate parameter was not significantly different from zero.
For each strain the N2O production assay illustrated in Fig. 1A was repeated three times with different batches of cells and data were used to estimate the linear rate of N2O production, correcting to take account of the initial cell number in each repetition. These data are presented in Fig. 2 as a function of maximum specific growth rate, which ranged from 0.018 h-1 (Nitrosospira sp. 40KI) to 0.070 h-1 (N. europaea ATCC 19718). The rate of N2O production ranged from 0.45 ± 0.02 amol cell-1 h-1 in Nitro- sospira briensis strain 128–85 ± 0.5 amol cell-1 h-1 in N. europaea ATCC 19718: a 186-fold difference. Analysis of variance using all data (specific N2O production rate for eight AOB strains, repeated three times with four within- repetition replicates, i.e. 95 d.f.) revealed that both AOB strain (P < 0.001) and repetition (P < 0.001) had a highly significant effect on the specific N2O production rate. One- way ANOVA using the repetition means (23 d.f.) also sup- ported the highly significant effect (P < 0.001) of AOB strain on N2O production rate, but pairwise comparison of repetition means (Table 2) revealed that this was due to the significantly (P < 0.05) elevated N2O production rate of N. europaea ATCC 19718; there was no difference in N2O produced between the remaining strains. When expressed as a percentage of the nitrite produced during
Fig. 1. Typical kinetics of (A) 14+15N-N2O and (B) nitrite production by Nitrosospira sp. En13 cell suspensions (�) in comparison to non- inoculated controls (�). The initial cell concentration of Nitrosospira sp. En13 in the assay was 1.63 ± 0.09 ¥ 107 cells ml-1. Data points are mean ± SE (n = 4). Solid lines represent linear regression function (P < 0.001, R 2 ≥ 0.991) Y = b1X + b0; where Y = cumulative concen- tration of N2O or NO2
– produced (nmol ml-1), X = time, b1 = linear rate (nmol ml-1 h-1) and b0 = intercept (nmol). Inset in A shows production of 15N-N2O in medium containing 1 mM
15N-NO2 – at 25 atom% excess
15N.
C u m
u la
tiv e n
m o l N
2 O
m l–
1
m ic
ro b ia
l c u ltu
re
0.0
0.2
0.4
0.6
0.8 A
B
Time (h)
0 2 4 6 8 10
C u m
u la
tiv e n
m o l N
O 2
– m
l– 1
m ic
ro b ia
l c u ltu
re
960
980
1000
1020
1040
1060
1080
1100
1120
1140
0 2 4 6 8 10 0.00
0.02
0.04
0.06
0.08
0.10
Fig. 2. Production of N2O by ammonia oxidizing bacterial strains: Nitrosospira sp. 40KI (�), Nitrosospira briensis 128 (�), Nitrosospira sp. En13 (�), Nitrosospira multiformis ATCC 25196 (�), Nitrosomo- nas europaea ATCC 19718 (�), Nitrosomonas europaea ATCC 25978 (�), Nitrosospira sp. NpAV (�) and Nitrosospira tenuis NV12 (�). The N2O production rate is expressed as a function of the maximum specific growth rate for each strain. The estimated linear rate of N2O produced during the 8- h assay was corrected for the initial number of cells in each assay. Each data point represents the mean (n = 4) N2O production rate for cells harvested from an inde- pendent flask. Bars represent ±1 SEM.
Specific growth rate (h–1)
0.01 0.02 0.03 0.04 0.05 0.06 0.07 0.08
N itr
o u s
o xi
d e p
ro d u ce
d (
a m
o l N
2 O
h – 1 c
e ll–
1 )
1
10
100
Nitrifier denitrification by Nitrosospira spp. 217
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 214–222
each assay, N2O production ranged from 0.03 ± 0.002% (N. briensis) to 0.70 ± 0.08% (Nitrosospira sp. En13).
All of the strains tested produced 15N-N2O at concen- trations significantly (P < 0.05) greater than the non- inoculated control. As in the case of total N2O production, kinetics of 15N-N2O production could be fitted to a linear function, an example of which is shown in Fig. 1A (inset). Estimated specific rates of 15N-N2O production are pre- sented in Table 2. Nitrosospira sp. B6 also produced significant amounts of 15N-N2O, but this culture was con- taminated with nitrite oxidizers. Freitag and colleagues (1987) have shown that Nitrobacter strains can grow by dissimilatory nitrate reduction with production of N2O. Thus the role of the contaminating nitrite oxidizers in 15N- N2O production in incubations with Nitrosospira sp. B6 can not be ruled out and quantitative data are not presented. With regard to total (14+15N) N2O production, N. europaea ATCC 19718 had the highest specific 15N-N2O production rate, which was more than double that of the next highest producing strain (Nitrosospira sp. 40KI). When the amount of 15N-N2O was expressed as a percentage of the
14+15N- N2O produced (Table 2), mean values for the AOB strains varied fourfold from 3.1 ± 0.04% for Nitrosospira multifor- mis to 13.5 ± 0.3% for Nitrosospira sp. 40KI.
Discussion
Most previous attempts to characterize the physiology of N2O production by AOB have concentrated on N. euro- paea strains (Ritchie and Nicholas, 1972; Hynes and Knowles, 1984; Poth and Focht, 1985; Anderson et al., 1993; Kester et al., 1997; Beaumont et al., 2002). Here, we quantified N2O production in aerobic incubations by six AOBs representing different phylogenetic clusters in the Nitrosospira lineage [clusters 0, 2 and 3 in the classifica-
tion according to Purkhold and colleagues (2003) based on cultured strains, Table 1] in addition to two different strains of N. europaea. All eight AOB strains tested produced N2O at detectable levels in the experimental system employed. Results show that, in addition to nitrosomonads, nitrosospiras also produce N2O, in agreement with observations reported by Goreau and col- leagues (1980), Remde and Conrad (1990), Jiang and Bakken (1999a), Dundee and Hopkins (2001) and Wrage and colleagues (2004a).
Nitrous oxide production by our strains was not always comparable to production reported elsewhere in the liter- ature. Published data on a marine Nitrosomonas sp., stud- ied by Goreau and colleagues (1980), lead to a N2O production rate of 46 amol h-1 cell-1, a value similar to those reported here. However, data obtained by Remde and Conrad (1990) for N. europaea (strain 28) suggest a production rate of 229 amol N2O h
-1 cell-1, i.e. fourfold and 15-fold higher than rates reported here for N. europaea strains ATCC 19718 and ATCC 25978 respectively. Simi- larly, data in Hynes and Knowles (1984) suggest a value for a N. europaea strain of 833 amol N2O cell
-1 h-1. The discrepancies in the rate of N2O production between stud- ies probably reflect differences in incubation conditions used because the amount of N2O formation depends on ammonium concentration (Hynes and Knowles, 1984), dissolved oxygen concentration (Goreau et al., 1980) and cell density (Remde and Conrad, 1990). Low oxygen con- ditions are known to induce clear increases in N2O pro- duction rates by Nitrosomonas sp. (Goreau et al., 1980; Kester et al., 1997; Dundee and Hopkins, 2001). The dis- cussed data sets were all obtained using aerobic incuba- tions. However, N. europaea (strain 28) was incubated in medium containing 10 mM ammonium (compared with 3.5 mM in this study), and Hynes and Knowles (1984)
Table 2. Ammonia oxidizer 14+15N-N2O and 15N-N2O production rates and molar yields expressed as a percentage of the NO2
– and 14+15N-N2O pro- duction rate respectively.†
AOB strain Total N2O production rate (amol 14+15N-N2O h
-1 cell-1)‡ Yield of 14+15N-N2O on a nitrite basis§ (%)
15N-N2O production rate (amol 15N-N2O h
-1 cell-1)§ Yield of 15N2O on a total (14+15N-) N2O basis
§ (%)
N. europaea ATCC 19718 58.0a* 0.45a 3.29a 7.8a
N. europaea ATCC 25978 15.5b 0.19b 0.34b 5.6b
N. briensis strain 128 4.2b 0.03c 0.02c 3.5c
N. multiformis ATCC 25196 7.6b 0.08d 0.20d 3.1c
N. tenuis strain NV12 2.0b 0.06d 0.06e 4.5d
Nitrosospira sp. strain 40KI 4.6b –** 0.88f 13.5e
Nitrosospira sp. strain En13 5.7b 0.70e 0.66g 12.4f
Nitrosospira sp. strain NpAV 3.9b 0.29f 0.33b 8.9g
*Mean values in columns superscripted by different letters are significantly different (P > 0.05). **Rate of nitrite production by Nitrosospira sp. 40KI was not significantly different from 0 (P > 0.05); therefore, percentage yield of 14+15N-N2O on a nitrite basis was not calculated. †Calculations are based on N2O concentration measured in the headspace; no correction is made for the concentration of N2O dissolved in the medium. ‡Values are the means of three independent experiments. §Values are the means (n = 4) for cells harvested from a single independent flask.
218 L. J. Shaw et al.
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 214–222
conducted assays at cell densities of 1.2 ¥ 109 ml-1 (com- pared with ~1 ¥ 107 in this study). In contrast to N2O pro- duction rate, the values for molar yield of N2O expressed as a percentage of moles of NO2
– produced are of the same order of magnitude (0.03–0.7%) as those reported by Remde and Conrad (1990) for N. europaea strain 28 (0.05–1.95%) and by Jiang and Bakken (1999a) for Nitrosomonas ATCC 25978 (0.2%) and Nitrosospira sp. strains (~0.05–1%).
One characteristic of the N2O assays employed in this study was the significant variation in the rate of N2O pro- duction between different repeats of the assay for some of the AOB strains. Variation between assay repeats (and for within-repeat replicates) has been recorded previously during experiments to quantify consumption of NO by N. europaea ATCC 19718 (Beaumont et al., 2004a) and the effect of inhibitors on N2O production by N. europaea ATCC 19718 and N. briensis strains (Wrage et al., 2004a). It is known that N2O production depends on cell density (Remde and Conrad, 1990) and differing initial cell con- centrations in assays has been suggested as a reason for variable N2O production (Wrage et al., 2004a). In our assays the initial cell concentration was between 0.63 and 2.6 ¥ 107 ml-1, but did not consistently influence the N2O production rate (data not shown). An alternative explana- tion is variation in the activity of different batches of cells, although care was taken to ensure that cells used were in a comparable physiological state, by using those harvested in the same growth phase (75–90% of the available ammonium used, as judged by nitrite analysis). However, in spite of the variation between cell batches, N. europaea ATCC 19718 consistently produced sig- nificantly (P < 0.001) greater amounts of N2O (four times more than the next highest producing strain). The genome of N. europaea ATCC 19718 has recently been sequenced (Chain et al., 2003) and this strain is becoming a model for reverse genetics approaches to study the role of denitrification-like genes in inorganic nitrogen metabo- lism (Beaumont et al., 2002, 2004a; Schmidt et al., 2004). The atypical behaviour of N. europaea ATCC 19718 in terms of N2O production should be borne in mind if extrap- olating findings gained with this strain to other soil AOB.
Most strikingly, all of the strains tested, both nitrosomonads and nitrosospiras, were able to reduce nitrite to N2O, i.e. to carry out nitrifier denitrification. Appli- cation of 15N-NO2
– (25 atom percentage excess 15N) enabled distinction of N2O produced by nitrite reduction from that potentially produced by other mechanisms, such as the formation of N2O by abiotic reactions involv- ing intermediates of ammonia oxidation. The ability of N. europaea to mediate nitrifier denitrification has been demonstrated previously, either through use of 15N analy- sis (Poth and Focht, 1985; Remde and Conrad, 1990) or artificial electron donors (Remde and Conrad, 1990;
Anderson et al., 1993). Genes putatively involved in nitri- fier denitrification [nirK (aniA) and norB, encoding nitrite reductase and nitric oxide reductase, respectively] have been annotated on the N. europaea ATCC 19718 genome (Chain et al., 2003) and detected by PCR ampli- fication in pure cultures of marine Nitrosomonas and Nitrosococcus spp. (Casciotti and Ward, 2001, 2005). It should be noted that Arp and Stein (2003) suggest, based on evidence gained using a nirK– deficient N. euro- paea ATCC 19718 mutant, that the nirK gene product is not wholly responsible for nitrite reduction. However, recent experiments show that N. europaea ATCC 19718 nirK– and norB– mutants are incapable of 15N-gas produc- tion from 15N-NO2
–, indicating a key role for NirK and NorB in nitrifier denitrification (Schmidt et al., 2004). Interest- ingly, despite the efforts of Casciotti and Ward (2001, 2005), nirK and norB homologues have yet to be identi- fied in Nitrosospira spp. by PCR amplification. By con- trast, genomic DNA from Nitrosospira sp. NpAV and Nitrosospira (formerly Nitrosolobus) sp. 24-C has been shown to hybridize at low stringency to probes specific for nirK in Pseudomonas sp. G-179 (Bruns et al., 1998). Regardless of the molecular basis of nitrifier denitrifica- tion in Nitrosomonas and Nitrosospira spp., this is the first demonstration of the ability of nitrosospiras to undertake this function.
Up to 13.5% (Nitrosospira sp. strain 40KI, Table 2) of the N2O produced was enriched in
15N derived from exog- enously applied 15N-NO2
–. If enzymes responsible for nitrite reduction reside in the periplasmic space, as seems most likely from knowledge of other denitrification systems (Zumft, 1997), the exogenously -applied 15N-NO2
– must have been able to cross the outer membrane for the first step in the denitrification pathway to occur. The pKa of nitrous acid (HNO2) is 3.3; therefore, at the pH of medium used in this study (7.5), less than 0.01% of exogenous nitrite would have been present in the protonated form, enabling free passage through the outer membrane (Moir and Wood, 2001). It is more likely that the majority of the nitrite entered the periplasmic space via outer membrane porins (Nikaido, 2003). A gene putatively encoding gen- eral diffusion outer membrane porins has been identified on the N. europaea ATCC 19718 genome (http://www.ncbi. nlm.nih.gov/entrez/viewer.fcgi?db=nucleotideandval= 30248031). Given the fact that 15N-NO2
– was applied at only 25 atom%, the actual percentage of total (14+15N) N2O produced through denitrification of total exogenous (14+15N) NO2
– may have been much higher (i.e. four times that of the values obtained on a 15N-N2O basis, assuming negligible isotopic discrimination against 15N). Thus, up to 54% of the N2O produced may have been derived from denitrification of exogenous nitrite. Whether the remaining N2O was produced by reduction of nitrite produced endog- enously (by ammonia oxidation) or through other sources
Nitrifier denitrification by Nitrosospira spp. 219
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 214–222
(e.g. abiotic reaction of intermediates of ammonia oxida- tion) is not known.
Detection of nitrifier denitrification activity in six nitro- sospiras (possibly a seventh, including strain B6), which are representatives of three major phylogenetic clusters (Purkhold et al., 2003) in the cultured Nitrosospira lin- eage, in addition to the previously known activity of nitrosomonads, suggests that the ability to denitrify is a widespread, if not ubiquitous, trait in ammonia oxidizers, at least in those phylogenetic clusters with cultured rep- resentatives. The widespread nature of the denitrification trait prompts conjecture regarding the ecological signifi- cance of nitrifier denitrification. The benefits to ammonia oxidizers are beginning to be understood. One suggestion is that reduction of nitrite may be a strategy to reduce competition for oxygen from nitrite oxidizers by removing their substrate (Poth and Focht, 1985). Another sugges- tion is that nitrite is reduced to conserve oxygen and produce energy in low oxygen environments (Poth and Focht, 1985; Schmidt and Bock, 1997) as in classical heterotrophic denitrification. If the role of denitrification in ammonia oxidizers is the same as in heterotrophs, the pathway should be regulated in the same way, i.e. by concentrations of available oxygen and nitrite (Zumft, 1997). However, NorB in N. europaea ATCC 19718 is expressed under aerobic conditions (Beaumont et al., 2004a) and NirK is expressed aerobically and in response to increasing concentrations of nitrite (Beaumont et al., 2004b). The latter finding suggests that a major role for the NirK enzyme is to help to protect N. europaea cells from toxic nitrite produced during nitrification (Beaumont et al., 2002, 2004b; Arp and Stein, 2003).
Several studies have reported oxygen sensitivity of N2O production in ammonia oxidizers (Goreau et al., 1980; Hynes and Knowles, 1984; Poth and Focht, 1985; Remde and Conrad, 1990; Kester et al., 1997; Dundee and Hop- kins, 2001). For example, N2O production was shown to be greatly stimulated under anaerobic conditions (approx- imately 700 times greater that in aerobic conditions) in both N. europaea strain 28 and Nitrosospira (formerly Nitrosovibrio) strain K71 (Remde and Conrad, 1990). Thus, the potential of the N. europaea and Nitrosospira spp. strains tested here for N2O production may also be significantly greater at oxygen tensions lower than those studied. Increased N2O production by ammonia oxidizers at low oxygen tensions has previously been attributed to increased rates of nitrifier denitrification (Dundee and Hopkins, 2001); presumably even if oxygen concentration does not significantly repress the expression of denitrify- ing enzymes in AOBs (Beaumont et al., 2004b), denitrifi- cation activity could be greater under low O2 due to reduced competition with nitrite from O2 as a preferred electron acceptor. However, studies have not specifically quantified the contribution of nitrifier denitrification above
other possible processes that could be occurring. Here, the application of 15N-NO2
– as a tracer, has demonstrated that significant nitrite reduction occurs in well-aerated cul- tures (in one experiment, the dissolved oxygen content was estimated as 5.6 ± 0.1 mg l-1); a finding in agreement with the studies with N. europaea (Beaumont et al., 2004a,b) which demonstrate the aerobic expression of denitrification pathway enzymes. Thus, soil-based studies which attempt to identify the contribution of various biological pathways to total N2O production from soils through use of oxygen suppression of denitrification (Webster and Hopkins, 1996; Wrage et al., 2004b) should consider the possibility that this approach may not com- pletely suppress the nitrifier denitrification pathway.
Experimental procedures
Ammonia oxidizer bacteria (AOB) strains and maintenance
Nine betaproteobacterial AOB strains were used in this study. Details of their source and affiliation with ammonia oxidizer 16S rRNA gene-based phylogenetic clusters proposed by Purkhold and colleagues (2003) are given in Table 1. All strains were maintained as static batch liquid cultures at 28∞C in modified Skinner and Walker (1961) (S and W) medium containing (per litre): (NH4)2SO4 (0.235 g); KH2PO4 (0.200 g); CaCl2.2H2O (0.040 g); MgSO4.7H2O (0.04 g); FeSO4 (0.5 mg); Na2EDTA (0.5 mg) and phenol red (0.5 mg) as a pH indicator. After autoclaving, a sterile solution of Na2CO3 (5% w/v) was added dropwise to the medium until the colour changed from yellow to pink, corresponding to a pH of 7.5– 8. Growth of AOB cultures results in acidification of the medium (through production of nitrous acid) and a change in colour of the indicator from pink to yellow. Cultures were checked weekly for pH indicator colour change and, if required, the pH was adjusted by addition of Na2CO3. Ammo- nia oxidizing bacteria cultures were subcultured into fresh S and W medium after the pH had been adjusted three times. Cultures were maintained in at least triplicate and, at each subculture, were screened for heterotroph contamination by drop plating on nutrient agar. Nutrient agar plates were incu- bated at 28∞C for 28 days. Cultures showing contamination were discarded.
Determination of AOB specific growth rates
Stationary phase cultures, maintained in S and W medium, were used to inoculate (1%, v/v) quadruplicate 250 ml conical flasks containing 150 ml SWH medium, consisting of S and W medium with added HEPES (free acid, Sigma-Aldrich, Dorset, UK) buffer (20 mM, pH 7.5) and without phenol red. After autoclaving, the medium was amended with 1.78 ml l-1
of 5% (w/v) Na2CO3 solution. Flasks were incubated at 28∞C in the dark on an orbital shaker (120 r.p.m.). At regular inter- vals, aliquots (500 ml) were removed for quantification of nitrite production (see below) as a surrogate measure of growth.
220 L. J. Shaw et al.
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 214–222
Ln nitrite concentration was plotted against time and the specific growth rate (m) estimated over the linear range by regression analysis.
Colorimetric quantification of nitrite
Nitrite concentrations in culture supernatant samples were determined using a miniaturized version of the assay of Keeney and Nelson (1982). Briefly, 20 ml diazotizing reagent (50 mg sulfanilamide ml-1 in 2.4 M HCl) was added to 40 ml sample diluted in 920 ml d.d. H2O. After vortexing and incubation for 5 min, 20 ml of coupling reagent [30 mg N-(1-naphthyl)-ethylenediamine HCl ml-1 in 0.12 M HCl] was added and the solution vortexed and incubated for a further 10 min prior to determination of absorbance at 540 nm. A standard curve was constructed using NaNO2 in the range 0–5 mg NO2–N ml-1.
Assay of 14+15N-N2O and 15N-N2O production by AOB
strains
Ammonia oxidizing bacteria cultures for N2O assays were grown up in 250 ml SWH medium in 1-l conical flasks at 28∞C and shaken at 120 r.p.m. (Innova 4330 refrigerated incubator shaker, New Brunswick Scientific). Cells were har- vested by filtration (0.2 mm cellulose acetate filter) using a vacuum manifold when 75–90% of the available ammonium had been utilized (as judged by analysis of nitrite and, for strain B6, nitrate and nitrite). Cells were washed off the filter and resuspended in an equal volume of fresh SWH medium. Aliquots (50 ml) of the resulting cell suspensions were decanted to quadruplicate 125 ml narrow-necked bot- tles. A solution of Na15NO2 (200 ml, 250 mM, 25 atom per- centage 15N) was pipetted into each bottle, resulting in a final NO2
– concentration of 1 mM. Na15NO2 was purchased at 98–99 atom percentage 15N (Sercon, Cheshire, UK) and mixed with NaNO2 to give the desired
15N enrichment. The bottles were sealed with gas-tight silicone rubber septa and placed on a rotary shaker (120 r.p.m., 28∞C). The head- space contained ambient air concentrations of O2. After incubation for 2, 4, 6 and 8 h, headspace samples (12 ml) were removed using a gas-tight syringe and stored in 12 ml evacuated gas-vials. From the 12 ml of sample, 1 ml was taken for total (14+15N) N2O determination by gas chromatog- raphy (GC) and the remainder was used for 15N-N2O analy- sis by isotope ratio mass spectrometry (IRMS). After gas sampling, the bottles were opened, a subsample (300 ml) of the culture removed for nitrite analysis and the headspace allowed to exchange with the atmosphere before resealing. The initial number of cells used in each assay was deter- mined by direct counts using epifluorescence microscopy and staining with 4¢,6-diamidino-2-phenylindole (DAPI) (Hobbie et al., 1977).
Gas subsamples were analysed for total (14+15N) N2O using an Agilent 6890 GC equipped with a 63Ni electron capture detector and a Haysep Q 60–80 mesh packed 8 ft (2.44 m) column. The carrier gas was N2 and the isothermal column and detector temperatures were 40∞C and 250∞C respec- tively. The GC was calibrated using laboratory air, 1.1 and 5.2 ppmv standards.
Isotope ratio mass spectrometry analysis was conducted by cryofocusing using an ANCA TGII trace gas preparation module connected to a 20–20 isotope ratio mass spectrom- eter (PDZ/Europa). Headspace sample in the 12-ml vial was transferred to pre-evacuated and He-flushed gas-tight 125- ml glass bottles and cut with laboratory air prior to analysis. 15N-N2O atom percentage values, volume of headspace and sample analysed and total concentration of N2O in the culture headspace, as determined by GC analysis, were used to calculate the mmoles of 15N-N2O produced in the culture headspace. The concentration of 15N-N2O was then con- verted to a per cell basis using the DAPI count data.
Statistical treatment of data
Analysis of Variance (General Linear Model) and regression analysis were performed using Minitab v. 13.1.
Acknowledgements
This work was funded by a Biotechnology and Biological Sciences Research Council, UK, Grant (32/D19035) to E.M.B. and J.I.P.
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Supplementary material
The following supplementary material is available for this article online: Fig. S1. Phylogenetic trees showing the placement of nine strains (in bold) used in this study within the Nitrosospira (A) and Nitrosomonas (B) lineages of the Betaproteobacteria subgroup ammonia oxidizing bacteria.
DenitrificationAsASourceOfNitricOxideEmissionsFromIncubatedSoilCoresFromA-UKgrasslandSoil.pdf
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Soil Biology & Biochemistry 95 (2016) 1e7
Contents lists avai
Soil Biology & Biochemistry
journal homepage: www.elsevier.com/locate/soilbio
Denitrification as a source of nitric oxide emissions from incubated soil cores from a UK grassland soil
Nadine Loick a, Elizabeth R. Dixon a, Diego Abalos b, Antonio Vallejo b, G. Peter Matthews c, Karen L. McGeough d, Reinhard Well e, Catherine J. Watson d, Ronnie J. Laughlin d, Laura M. Cardenas a, *
a Rothamsted Research, North Wyke, Okehampton, Devon EX20 2SB, UK b Technical University of Madrid, Chemistry and Agricultural Analysis, Madrid, Spain c School of Geography, Earth and Environmental Sciences, University of Plymouth, Davy Building, Drake Circus, Plymouth, Devon PL4 8AA, UK d Agri-Food and Biosciences Institute, Newforge Lane, Belfast BT9 5PX, UK e Thünen-Institut für Agrarklimaschutz, Bundesallee 50, 38116 Braunschweig, Germany
a r t i c l e i n f o
Article history: Received 22 September 2015 Received in revised form 11 December 2015 Accepted 17 December 2015 Available online 3 January 2016
Keywords: Nitrous oxide Flow-through system Isotopes Grassland Nitrogen cycle Greenhouse gas (GHG) emissions
* Corresponding author. Tel.: þ44 (0) 1837 883 500 E-mail address: [email protected]
http://dx.doi.org/10.1016/j.soilbio.2015.12.009 0038-0717/© 2016 Published by Elsevier Ltd.
a b s t r a c t
Agricultural soils are a major source of nitric oxide (NO) and nitrous oxide (N2O), which are produced and consumed by biotic and abiotic soil processes. The dominant sources of NO and N2O are microbial nitrification and denitrification. While N2O emissions have been attributed to both processes, depending on the environmental conditions such as substrate availability, pH and water filled pore space (WFPS), NO emissions are thought to predominantly derive from nitrification. Although attributing gaseous emissions to specific processes is still difficult, recent findings challenge the latter of those assumptions. Using the gas-flow-soil-core method, i.e soil cores incubated under a He/O2 atmosphere at constant surface gas flow, combined with 15N labelled isotopic techniques, the present study investigated the role of denitrification on NO, N2O and N2 emissions in a UK grassland soil under high soil moisture and an aerobic headspace atmosphere. With the application of KNO3 and glucose to support denitrification, denitrification was the source of N loss of between 0.61 and 0.67% of the added N via NO emissions, 1.60 e1.68% via N2O and 0.03e0.05% via N2 emissions. Overall, our study showed that denitrification has been overlooked as a source of NO emissions.
© 2016 Published by Elsevier Ltd.
1. Introduction
Agricultural soils are the dominant source of nitrous oxide (N2O), a potent greenhouse gas and a major cause of ozone layer depletion (IPCC, 2007; Ravishankara et al., 2009). Other gaseous forms of nitrogen (N) are lost from agricultural soils, such as N2 which together with N2O represents less N available for crop growth. Soils also act as a significant source of nitric oxide (NO), which catalyses the formation of ground level ozone, affecting human health and vegetation (Crutzen, 1981), and contributes to the formation of acid rain and the eutrophication of semi-natural ecosystems. Microbial denitrification is often the dominant pro- cess generating N2O, and as such, intense investigations (i.e. >1000 published studies) have led to a good understanding of the abiotic
. (L.M. Cardenas).
factors regulating N2O emissions via denitrification (Beaulieu et al., 2011). However, the role of this process on NO emissions remains largely unexplored, apart from a few studies (Wang et al., 2011, 2013), even though NO is an obligatory intermediate of N2O for- mation in denitrification (Wolf and Russow, 2000; Russow et al., 2009).
Most experiments suggest that NO emitted from soils is mainly produced through nitrification (Skiba et al., 1997), whereas that produced from denitrification is further reduced to N2O before it escapes to the soil surface (Skiba et al., 1997). This is attributed to high soil water content (it has been shown that at a WFPS above 70%, N2O was produced solely by denitrification (Bateman and Baggs, 2005)), soil compaction and fine soil texture (sieved to <2 mm) creating low diffusivity for gases, which increases the residence time and the potential for further reduction when deni- trification conditions dominate. Recent findings, however, chal- lenge these assumptions. Using the gas-flow-soil-core technique, which has been proven to be a reliable tool for quantifying
N. Loick et al. / Soil Biology & Biochemistry 95 (2016) 1e72
emissions from denitrification, Wang et al. (2013) observed sig- nificant NO fluxes from nitrate (NO3
�) amended soils. Attributing these emissions specifically to denitrification has remained elusive due to methodological constraints to elucidate the underlying mi- crobial production and consumption processes. Previous efforts to identify these processes have mostly relied on acetylene inhibition and isotope labelling techniques (Baggs, 2008).
Isotope analysis has emerged as a way to identify the source and thereby the processes from which N2O is being produced (Arah, 1997). It is also known that microorganisms discriminate against the heavier molecule (e.g 15N vs. 14N), preferring to use the lighter molecule which requires less energy to break the bonds (Kendall and Caldwell, 1998). This should be considered when applying labelled substrate to investigate microbial processes.
The aim of this study was to explore the potential role of deni- trification as a significant source of NO emissions. We hypothesise that denitrification can be a major source of NO emissions in a UK grassland soil under high moisture content. This study uses the gas- flow-soil-core technique (C�ardenas et al., 2003), further developed to include NO measurements, combined with isotopic analyses. A 15N labelled substrate as well as an unlabelled substrate at the same application ratio was used to determine whether there was an ef- fect of the labelled N on the investigated processes at a 5 atom% enrichment. Additionally to adding potassium nitrate (KNO3) as N source, glucose was added to supply a readily available C source and thereby promote denitrification. During denitrification C is used as electron donor and C availability is one factor controlling denitri- fication rates and compared to other C-compounds, denitrification tends to be most stimulated after addition of ethanol or glucose (Morley and Baggs, 2010).
2. Materials and methods
2.1. Soil preparation
A clayey pelostagnogley soil of the Hallsworth series (Clayden and Hollis, 1984) (44% clay, 40% silt, 15% sand (w/w), Table 1) was collected on the 4th of November 2013 from a typical grassland in SW England, located at Rothamsted Research, North Wyke, Devon, UK (50�4601000N, 3� 5400500W). Spade-squares (20 � 20 cm to a depth of 15 cm) of soil were taken from 12 locations along a ‘W’ line across a field of 600 m2 size. After sampling, the soil was air dried to ~30% H2O (dry basis), roots and plant residue were removed and the soil sieved to <2 mm and stored at 4 �C for 5 days before packing into cores and starting the incubation.
2.2. Experimental setup
The incubation was carried out using the DENItrification System (DENIS), a specialized gas-flow-soil-core incubation system (C�ardenas et al., 2003). Twelve cores were packed with soil to a bulk
Table 1 Soil characteristics (before amendment application). Mean ± standard error (n ¼ 3).
Parameter Amount
pH water [1:2.5] 5.6 ± 0.27 Available Magnesium (mg kg�1 dry soil) 100.4 ± 4.81 Available Phosphorus (mg kg�1 dry soil) 10.4 ± 1.10 Available Potassium (mg kg�1 dry soil) 97.5 ± 12.83 Available Sulphate (mg kg�1 dry soil) 51.7 ± 0.62 Total N (% w/w) 0.5 ± 0.01 Total Oxidised N (mg kg�1 dry soil) 15.1 ± 0.07 Ammonium N (mg kg�1 dry soil) 9.2 ± 0.09 Organic Matter (% w/w) 11.7 ± 0.29
density of 0.8 g cm�3 and a height of 75 mm into stainless steel vessels of 140 mm diameter. To ensure denitrification conditions, the soil moisture was adjusted to 85% WFPS, taking the later amendment into account. This WFPS was similar to those used in previous studies to promote denitrification processes (Meijide et al., 2010; Bergstermann et al., 2011). In order to measure N2 fluxes the native atmosphere was removed by flushing the soil cores from the bottom with a mixture of He:O2 (80:20) at 30 ml min�1 for 14 h Flow rates were then decreased to 12 ml min�1 and the flow re-directed over the surface of the soil core for three days before amendment application to measure baseline emissions. O2 was kept in the gas mixture at atmospheric levels as the objective was to investigate denitrification achieved by high WFPS instead of forcing anaerobic conditions by preventing any O2 diffusion.
The following treatments were applied to four replicate vessels: (a) labelled (15N-labelled KNO3 at 5 atom% and glucose); (b) unla- belled (KNO3 and glucose); (c) control (water only). The labelled and unlabelled treatments contained nitrogen at a rate equivalent to 75 kg N ha�1 (i.e. 121.5 mg N kg�1 dry soil) and C as glucose at 400 kg C ha�1 (i.e. 648 mg C kg�1 dry soil), which is similar to previous studies (Meijide et al., 2010; Bergstermann et al., 2011). The amendment for each core was dissolved in 50 ml distilled water, and the controls also received 50 ml distilled water each. The vessels were kept at 20 �C during the whole incubation period, which lasted for 10 days after amendment application.
2.3. Gas analyses and data manipulation
Gas samples were taken every two hours for each vessel. Fluxes of N2O and CO2 were quantified using a Perkin Elmer Clarus 500 gas chromatograph (Perkin Elmer Instruments, Beaconsfield, UK) equipped with an electron capture detector (ECD) for N2O, and with a flame ionization detector (FID) and a methanizer for CO2. N2 emissions were measured by gas chromatography with a helium ionisation detector (VICI AG International, Schenkon, Switzerland) (C�ardenas et al., 2003), while NO concentrations were determined by chemiluminescence (Sievers NOA280i, GE Instruments, Colo- rado, USA). All gas concentrations were corrected for the surface area and flow rate going through the vessel (measured daily). Fluxes were calculated on a kg N or C ha�1 day�1 basis.
2.4. Isotopic analyses of N2O
Gas sampling times for 15N analysis were pre-determined based on data from previous experiments (data not shown). Samples were taken just before (0 h) and 4 h after amendment application, then every 24 h for the first week, followed by a final sample at day 10. This sampling strategy was decided on from previous experi- mental results to cover changes in isotopic signature before amendment application, as well as during the NO and N2O peaks (4e5 h and 3e4 d, respectively), and after emissions returned to background levels. Samples were taken from the outlet line of each vessel using 12 ml exetainers (Labco) which had previously been flushed with He and evacuated. 15N enrichment of N2O was measured using a TG2 trace gas analyser (Europa Scientific, now Sercon, Crewe, UK) and Gilson autosampler, interfaced to a Sercon 20e22 isotope ratio mass spectrometer (IRMS). Solutions of 6.6 and 2.9 atom% ammonium sulphate ((NH4)2SO4) were prepared and used to generate 6.6 and 2.9 atom% N2O (Laughlin et al., 1997) which were used as reference and quality control standards.
The process leading to the formation of the measured N2O, i.e. whether it is produced by nitrification or denitrification, was determined by calculating how much of the N2O was derived from NO3
� as the parent molecule. When 15N labelled NO3 � is added, it is
N. Loick et al. / Soil Biology & Biochemistry 95 (2016) 1e7 3
assumed that it completely mixes with the native soil NO3 � pool to
form a single uniformly labelled NO3 � pool. The 15N content of the
N2O was calculated from either 45R or 46R, with 45R being the ratio
of the ion currents (I) for mass 45/44 (45R ¼ 45I/44I) and 46R for mass 46/44 (46R ¼ 46I/44I). If the 15N contents of the measured N2O calculated from either 45R or 46R are equal, then the distribution of the 15N atoms in the N2O molecules is random, and therefore the N2O originated from a single uniformly labelled NO3
� pool (Stevens et al., 1997; Stevens and Laughlin, 1998). When the NO3
� pool is labelled and the N2O concentration is greater than the IRMS method detection limit (2 ppm), calculations of the fraction of N2O derived from the denitrifying pool (d0D) were performed. The sources of N2O were then apportioned into d0D and the fraction derived from the nitrifying pool (d0N ¼ (1 � d0D)) and calculated as described in Arah (1997). In Arah's equation N2O d0D is the fraction of the emitted N2O which is derived from the
15N labelled, deni- trifying NO3
� pool. A N2O d0D value of unity (1.00) indicates that 100% of the N2O emitted derived from the NO3
� pool. To determine the source of the measured N2O, i.e. how much of
it was derived from the amendment (N2O_Namend) rather than the native soil N, the following equation was used for the labelled treatments (Senbayram et al., 2009):
N2O Namend ¼ N2O Ntotal
15Nat%exsample 15Nat%exfert
! (1)
where N2O_Ntotal ¼ total emissions of N2O from the soil; 15Nat% exsample ¼ 15N atom% excess of the emitted N2O (15N atom% of the measured sample minus the mean natural 15N abundance of background N2O obtained in our experiment (0.366 atom %));
15Nat %exfert ¼ 15N atom% excess of the applied amendment solution.
2.5. Soil analyses
Soil samples were taken at the beginning and end of the incu- bation to determine the initial and final moisture contents and the NH4
þ and total oxidised N (TON: NO3 � þ NO2�) concentrations. Nitrite
(NO2 �) is generally thought to accumulate very rarely in nature, and
it has been shown that NO2 � is rapidly mineralised in soil (Paul and
Clark, 1989; Burns et al., 1995, 1996). It is therefore assumed that NO2
� concentrations in the soil samples are negligible, and TON is nearly exclusively made up of NO3
�. For the final soil analyses, each core was divided in half to separate the top section from the bottom section. WFPS was calculated from soil moisture contents by drying a subsample (50 g) at 105 �C overnight. Soil NH4
þ-N and TON were analysed by automated colorimetry from 2 M KCl soil extracts using a Skalar SANPLUS Analyser (Skalar Analytical B.V., Breda, Netherlands) (Searle, 1984). 15N abundance of NO3
� and NH4 þ was
measured by quadrupole mass spectrometer (GAM 200, InProcess, Bremen, Germany) (as described by Stange et al. (2007) at the Thünen Institute of Climate Smart Agriculture (Brauschweig, Ger- many)). Briefly, NO3
� was reduced to NO by Vanadium chloride (V(III)Cl3) and NH4
þ was oxidized to N2 by Hypobromite (NaOBr). NO and N2 were the gases measured.
Table 2 Cumulative emissions of NO, N2O, N2 as kg N ha
�1 and CO2 as kg C ha �1 over the time
of the respective peaks. N2 and CO2 emissions are baseline subtracted. Different letters indicate a significant difference between treatments for each measured gas (n ¼ 4, p < 0.05).
Gas Labelled (15NeKNO3 þ C) Unlabelled (KNO3 þ C) Control NO 0.46 ± 0.02A 0.50 ± 0.02A 0.03 ± 0.03B
N2O 1.20 ± 0.28 A 1.26 ± 0.08A 0.01 ± 0.01B
N2 0.30 ± 0.03 A 0.33 ± 0.07A 0.14 ± 0.06A
CO2 87.89 ± 3.73 A 92.68 ± 2.68A 5.50 ± 3.39B
2.6. Statistical analysis
Statistical analysis was performed using GenStat 16th edition (VSN International Ltd). Prior to the statistical tests all data were analyzed to proof their normal distribution (KolmogoroveSmirnov test) and equality of variance (Levene test). Cumulative emissions of NO, N2O, N2 and CO2 were calculated from the area under the curve after linear interpolation between sampling points. Differ- ences in total emissions for each gas measured between treatments
as well as differences in soil characteristics between treatments and between top and bottom of soil cores were assessed by ANOVA at P < 0.05. Where treatment effects proved to be significant, Fisher's Least Significant Test (LSD) was used as post hoc test to ascertain differences among treatment levels.
3. Results
3.1. Gas emissions
CO2 fluxes showed constant emissions of 10 kg C ha �1 d�1 before
and after the CO2 peak (day 0e6) in all vessels. N2 emissions increased at the moment the amendment was applied, but decreased immediately after until day 3.5 when they reached background levels, before increasing again. In order to show CO2 and N2 emissions attributed to amendment application only, the fluxes were adjusted by subtracting background emissions. There were no significant differences in fluxes, or cumulative emissions for any of the measured gases between the labelled and unlabelled treatments (Table 2). Both treatments, however, were significantly higher than the control for all gaseous emissions measured, except for N2.
Nitric oxide emissions peaked 14 h after amendment application (Fig. 1), with maximum average fluxes of 0.58 and 0.70 kg N ha�1 d�1, for the labelled and unlabelled treatment, respectively. Fluxes decreased afterwards resulting in values below 0.1 kg N ha�1 d�1 30 h after amendment application. Fluxes then decreased further to below 0.05 kg N ha�1 d�1, before showing a linear increase over 5 days to values of around 0.1 kg N ha�1 d�1
until the end of the experiment. Losses of N via NO emissions represented 0.61 and 0.67% of the N added. The control treatment showed negligible fluxes of NO over the whole experimental period.
Similar to NO, emissions of N2O increased immediately after amendment application. After 14 h, N2O showed a first maximum of 0.24 and 0.17 kg N ha�1 d�1 for the labelled and unlabelled treatment, respectively (Fig.1). In both treatments fluxes decreased over the following 12 h by 0.02 kg N ha�1 d�1 before increasing again to a maximum of 0.45 and 0.44 kg N ha�1 d�1, 3.3 and 3.8 days after amendment application, respectively. Total losses of N2O represented 1.60 and 1.68% of the N applied for the labelled and unlabelled treatment, respectively. Again the control treatment maintained significantly lower fluxes than the fertilized treatments over the whole experimental period.
Gaseous nitrogen (N2) fluxes (Fig. 1) were very similar in all treatments, and showed a decrease during the first 3.5 days of the experiment. After this initial phase, fluxes increased again to maxima of 0.09, 0.08 and 0.05 kg N ha�1 d�1 for the unlabelled, labelled and control treatment, respectively. Though not statisti- cally different (p ¼ 0.078), both of the amended treatments showed higher fluxes (maximum of 0.08 kg N ha�1 d�1) than the control (maximum of 0.05 kg N ha�1 d�1), before decreasing again to the level they had reached 3.5 days after amendment application. Total
N. Loick et al. / Soil Biology & Biochemistry 95 (2016) 1e74
N2eN losses attributed to the amendment were 0.05% and 0.03% of the N applied, for the labelled and unlabelled treatment, respectively.
Cumulative emissions over the course of the experiment (Table 2) show that about 2.5 times more N was lost via N2O emissions than NO emissions, and total N losses via NO and N2O were over 40 times higher in the amended treatments than in the control.
Carbon dioxide fluxes (Fig. 1) increased immediately after amendment application, reaching values of 27.3 kg C ha�1 d�1 for both labelled and unlabelled treatments 1.5 days after amendment application, and 1.5 kg C ha�1 d�1 for the control 2 days after amendment application. By day 4, CO2 fluxes had decreased to values of 6 kg C ha�1 d�1 for both fertiliser amended treatments, with further decreases to background levels. The control only showed slightly elevated fluxes that decreased back to background levels by day 3. Above background losses of CO2 represented 22.0 and 23.2% of C added with the amendment for the labelled and unlabelled treatments.
Fig. 2 shows the average of the fluxes of all measured gases emitted from the fertiliser amended treatments (mean of labelled and unlabelled). Emissions of NO, N2O and CO2 increased within the first 2 h after amendment application. As expected from the mechanistic pathway for denitrification, NO is the first gas to peak followed by N2O, and finally N2. The sequence of emissions and processes can be described in 3 phases. Phase I (day 0e1): NO peak and a first small N2O peak; Phase II (day 1e4): main N2O peak, maximum CO2; Phase III (day 4e10): N2 peak, NO small gradual increase.
3.2. Isotopic results
The 15N enrichment of the measured N2O was equal whether it was calculated from 45R or 46R, proving that N2O originated from a single uniformly labelled NO3
� pool (homogeneously mixed labelled amendment with native soil NO3
�). The N2O d0D values obtained from Arah's equation, were not significantly different from unity (data not shown); therefore the source of the N2O was the uni- formly mixed 15N labelled NO3
� pool. The emitted N2O of the labelled treatment was analysed for
15N enrichment, and results showed that up to day 5, around 85% of the emitted N2O was derived from the amendment and 15% originated from the native soil NO3
�.
3.3. Soil chemistry
Total oxidised nitrogen (TON) (which is assumed to be nearly exclusively made up of NO3
�) was significantly higher in the top half than in the bottom half of the cores, and while there was no sig- nificant difference between the labelled and unlabelled treatments, both had significantly higher concentrations of TON and NH4
þ-N than the control (Table 3). The initial soil TON content was about an eight of the added N (15.1 vs 121.5 mg N kg dry soil�1). At the end of the incubation the amended treatments showed a 16e19 fold in- crease in TON while the TON in the control increased 6e7 fold. The 15N enrichment of TON was significantly higher in the top (3.5803 ± 0.0496 atom%) than in the bottom (3.0708 ± 0.0536 atom %) half of the cores in the labelled treatment.
The soil NH4 þ-N concentrations were lower than TON concen-
trations at the end of the incubation in all treatments, with slightly
Fig. 1. Gaseous emissions over the course of the incubation (1 kg ha�1 d�1 ¼ 4.17 � 10�4 mg cm�2 h�1).
0
5
10
15
20
25
30
0.0
0.2
0.4
0.6
0.8
1.0
-1 0 1 2 3 4 5 6 7 8 9 10
C kg
h a-
1 d-
1
N k
g ha
-1 d-
1
day a er amendment
NO N2O N2 * 10 CO2
Phase I Phase II Phase III
Fig. 2. Evolution of gaseous emissions of NO, N2O, N2 and CO2 from the amended treatments; N2 flux from the amended treatment is multiplied by ten, to improve visibility on the graph. CO2 and N2 emissions are baseline corrected to show amend- ment effects only (1 kg ha�1 d�1 ¼ 4.17 � 10�4 mg cm�2 h�1). Phase I: NO peak and N2O shows first peak. Phase II: NO emissions decrease. Main N2O peak, high CO2 concen- trations decrease. Phase III: NO emissions steadily increase again; CO2 and N2O emis- sions decrease to background levels.
N. Loick et al. / Soil Biology & Biochemistry 95 (2016) 1e7 5
higher values in the bottom sections of the cores. By the end of the incubation, NH4
þ concentrations had increased from 9.2 mg N kg�1
dry soil to around 13.2 and 15.0 mg N kg�1 at the top and bottom of the core respectively. The enrichment of NH4
þ-N in the top (0.4624 ± 0.0164 atom%) was significantly different to the bottom (0.3941 ± 0.0130 atom%) and to natural abundance, but the enrichment of the NH4
þ-N at the bottom (though elevated) was not significantly higher than natural abundance.
Soil moisture was 85% WFPS at the start of the incubation and was maintained for the whole core at a similar level for all treat- ments throughout the experiment (top of cores 81.27 ± 1.319%, bottom of cores 88.90 ± 1.145). By the end of the experiment the WFPS was significantly higher at the bottom of the core than the top with ~5% of the water having been redistributed from the top to the bottom of the core.
4. Discussion
4.1. N2O emissions
Stable isotope ratios are determined by the isotope ratios of the precursor materials and the preferential use of lighter isotopes by microorganisms (Holland and Turekian, 2010; Hu et al., 2015). Re- sults showed that using 5 atom% enriched KNO3 had no influence on the use of the native vs. enriched N-pool, providing confidence that the isotope analysis used in this study was a good tool to further investigate the source process of the gaseous emissions.
Data from the 15N-labelled treatment indicate that 85% of N2O was derived from the exogenously applied NO3
�, whereas only 15% was produced from the native soil NO3
� pool and/or NO3 � formed by
mineralisation. This source apportioning was maintained until day 5, after which N2O emissions were negligible, and were similar to
Table 3 Total Oxidized N (TON) and ammonium (NH4
þ) at the end of the experiment. Different let Bottom (X/Y)]; * indicates significant differences between the Top and Bottom layer with
Parameter Layer Labelled (15NeKN
TON (mg N kg�1 dry soil) Top 271.8 ± 17.32*A
Bottom 246.0 ± 21.37*X
NH4 þ (mg N kg�1 dry soil) Top 13.4 ± 1.66*A
Bottom 15.2 ± 2.42*X
the initial apportioning of the soil NO3 �, with the native soil NO3
�
making up 11.1% of the total NO3 �, while the amendment repre-
sented 88.9%. This similarity suggests that the amendment NO3 �
was homogeneously mixed with the native soil NO3 �. The amount of
N2O derived from the native soil NO3 from the fertilizer amended treatments (0.18 kg N ha�1) was higher than that emitted from the control (<0.01 kg N ha�1, Fig. 2) also suggesting that the amend- ment (KNO3 and C) and the native soil NO3
� had mixed, becoming available to the microbial community.
The equation of Arah (1997) was used to determine the process leading to the formation of the measured N2O for data collected during the first 5 days after amendment application; after this period, N2O concentrations were too low to calculate d0D values. The determined d0D values for those first 5 days indicate that close to 100% of the emitted N2O derived from denitrification of the NO3
�
pool. Arah's equation assumes that nitrification and denitrification
are the only source processes occurring. Our results, however, suggest that it is possible that some of the N2O might have derived from dissimilatory nitrate reduction to ammonium (DNRA). In DNRA, NO3
� is reduced to NH4 þ under similar conditions as deni-
trification (Fazzolari et al., 1998) and is promoted at C:N ratios (glucose-C:NO3
�) higher than 4 (Smith, 1982; Fazzolari et al., 1998). The increase in soil NH4
þ in the N treatments and the increase in 15N enrichment by 0.092atom% indicates that some of the added NO3
�
was transformed to NH4 þ. Although it has been argued that N2O is
produced by DNRA via NO2 � reduction (Schmidt et al., 2011), the
contribution of DNRA to N2O production is still uncertain (Baggs, 2011). The C:N ratio following amendment in the current study was 5.3, and the formation of NH4
þ from NO3 � indicates the possi-
bility that some of the N2O was produced through DNRA.
4.2. NO emissions
Nitric oxide is an obligate intermediate of N2O production through denitrification (e.g. Ye et al. (1994)). However, if soil moisture content is high (WFPS > 80%), emission of NO is generally considered to be non-detectable due to slow diffusion of NO from denitrifier-cells to the soil atmosphere, and later to air (Russow et al., 2009), during which it is further reduced to N2O. Based on this assumption, most studies indicate that emitted NO is mainly produced from hydroxylamine (NH2OH) during nitrification by ammonium oxidisers, which occurs at low soil moisture levels (Skiba et al., 1997). The control treatment did not show any NO emissions. As both, control and N amended treatments, had similar initial soil NH4
þ contents (9e13 mg N kg�1), treatments should have had similar NO fluxes if nitrification of NH4
þ had been the only source of NO under our experimental conditions. As this is not the case it can be assumed that nitrification did not contribute to initial NO emissions.
The increase observed with KNO3 application in phase I (Fig. 2) indicates that NO came from denitrification in our experiment. Several studies have measured NO fluxes under anoxic/denitrifying conditions in the field or laboratory and have found increased NO emissions after fertilisation or irrigation (e.g. Liu et al., 2010a; Liu
ters indicate significant differences between treatments for each layer [Top (A/B) or in a single treatment (TON and NH4
þ: n ¼ 4, p < 0.05, p < 0.05).
O3 þ C) Unlabelled (KNO3 þ C) Control 292.6 ± 17.09*A 90.5 ± 3.61*B
239.5 ± 14.85*X 108.3 ± 5.22*Y
13.0 ± 1.25*A 8.5 ± 0.55*B
14.9 ± 2.11*X 9.5 ± 0.77*Y
N. Loick et al. / Soil Biology & Biochemistry 95 (2016) 1e76
et al., 2010b; Bakken et al., 2012). However, to date only our study and those of Russow et al. (2009) and Wang et al. (2011, 2013) have shown that significant NO emissions can be directly promoted by denitrification in soils. Those previous studies confirmed NO as a free intermediate product of denitrification, however, those find- ings were derived from experiments performed under O2 depleted atmospheres. The soil in our study had a high WFPS to create anaerobic conditions, and therefore promote denitrification within the soil, the atmosphere above the soil surface, however, was kept aerobic. To the best of our knowledge our study is the first one showing high NO emissions derived from denitrification processes under an aerobic atmosphere.
During phase III (Fig. 2) of the experiment, NO emissions started to gradually increase again. A possible explanation for this is that around day 5, at the point of the N2 maximum, the soil O2 would have been depleted to its lowest levels, with rapid reduc- tion of N2O to N2 as a result of anaerobic respiration. The CO2 fluxes were back to background levels showing aerobic respiration was back to pre-amendment application levels. The recovery of NO after this point, and the lack of N2O emissions suggest that the soil might be recovering some aerobicity due to diffusion of the atmospheric oxygen from the headspace, and that nitrification could have been the source of those later NO fluxes (day 5.5e10). The soil NO3
� increased during the incubation by about 125e130 mg N kg�1 dry soil (equivalent to ~10 mg N kg�1 dry soil d�1). This rate is similar to rates measured previously for the same soil (unpublished data). This increase shows that mineralisation and nitrification occurred at some point in the incubation and that the later increase in NO could have been the result of these processes.
4.3. N2 emissions
One indication of NO3 � reduction by denitrification is the emis-
sion of N2. The high N2 concentrations in our experiment directly after amendment application were most likely due to dissolved N2 contained in the amendment solution being released into the vessel and flushed out over the first few days, reducing the N2 concen- trations back to background levels before the actual N2 peak appeared after day 3.5. When N2O was depleted in the fertilizer treatments, N2 increased slightly (Fig. 2), but concentrations were very low and not significantly different from the control, indicating that the addition of water stimulated production of N2 in all treatments. Although there is scarce information regarding fluxes of N2 in agricultural soils in response to the application of C and N sources, the appearance of the N2 peak has also been observed 3e4 days after application of amendments in previous experiments (Cardenas et al., 2007; Meijide et al., 2010; Bergstermann et al., 2011).
The relatively low N2 emissions in comparison to high NO and N2O emissions can be explained by the physiology and metabolism of the denitrifying bacteria and the high soil NO3
� levels remaining at the end of the incubation. Energy yields from denitrification reactions lessen in order of their appearance, with the reduction of NO3
� via NO2 � to NO being more energetically favourable than the
reduction of NO to N2O and of N2O to N2 (Koike and Hattori, 1975).
4.4. Denitrification as the source process of emissions summarised
The aim of this study was to investigate gaseous emissions from denitrification under an atmosphere that still contained natural amounts of oxygen. To induce low oxygen conditions in the soil, while the above atmosphere was kept at normal O2 levels, the soil cores had been set to a high WFPS and NO3
� and a labile C source had been applied in excess.
The apex of the peaks of the measured gases appear in the order that would be expected from the denitrification pathway, i.e. NO3
� is transformed to NO, which is then transformed into N2O and finally N2. In our study NO was produced in the hours following NO3
�
application (Fig. 2, Phase I). These emissions start at the same time as those of N2O, but decline more rapidly (i.e. 2 vs 5 days after amendment application). The next gas to peak in its emissions is N2O (Fig. 2, Phase II) followed by a small increase in N2 (Fig. 2, Phase III).
Overall, the results of this study indicate that denitrification played the most significant role in gaseous emissions. Total deni- trification (sum of NO, N2O and N2) is normally affected by soil abiotic properties such as WFPS, NO3
� and available C. A high soil WFPS reduces O2 diffusion to the pore space (Parton et al., 2001) which, in combination with KNO3 and C addition, promotes deni- trifying conditions. The availability of C not only supports the ac- tivity of denitrifiers per se, but also has the indirect effect of causing soil microsite anaerobiosis, due to an increased respiratory demand for O2. The high amount of NO3
�, which acts as an electron acceptor for denitrifiers, favoures the production of gaseous N-oxides over other reduced forms such as N2. Additionally, even though the synergistic activities of microbial communities in soil can lead to complete denitrification of NO3
� to N2, the earlier steps in the denitrification process are energetically more favourable often resulting in N2O consequently becoming the final denitrification product, especially if NO3
� is not limiting (Saggar et al., 2013).
5. Conclusions
This study shows that denitrification can be a major source of NO from soils at high water content and under the presence of an easily available C source. Until now, most studies indicated that NO produced in soils during denitrification was consumed by de- nitrifiers forming N2O or N2. To the best of our knowledge, this study, on a UK grassland soil, is the first showing high NO emissions derived from denitrification processes in a soil under high WFPS (creating anaerobic soil conditions and promoting denitrification), but with aerobic conditions above the soil surface. Our findings have several implications for an array of research fields. For example, in simulation studies using process-based models, the contribution of denitrification to NO emissions has been over- looked and needs to be taken into account. Our results also show that NO was mainly produced when an external source of NO3
� was added to soils. N2O fluxes, which appeared when NO fluxes had diminished, were also affected by amendments. Complete denitri- fication from exogenous NO3
� to N2 did not occur, and consequently the N2O:N2 ratio increased with amendment addition. Further research combining molecular tools with isotopic analyses is needed to expand the findings of our study.
Acknowledgements
Rothamsted Research receives strategic funding by the Biotechnology and Biological Sciences Research Council (BBSRC, Grant number BB/J004286/1). This study was funded by BBSRC project BB/K001051/1. D. Abalos thanks the Spanish Ministry of Science and Innovation for economic support through the Project AGL2009-08412-AGR. The authors thank Wolfram Eschenbach for 15N analysis from soil extracts, and Enrique Cancer-Berroya and Denise Headon for analyses of soil extracts.
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- Denitrification as a source of nitric oxide emissions from incubated soil cores from a UK grassland soil
- 1. Introduction
- 2. Materials and methods
- 2.1. Soil preparation
- 2.2. Experimental setup
- 2.3. Gas analyses and data manipulation
- 2.4. Isotopic analyses of N2O
- 2.5. Soil analyses
- 2.6. Statistical analysis
- 3. Results
- 3.1. Gas emissions
- 3.2. Isotopic results
- 3.3. Soil chemistry
- 4. Discussion
- 4.1. N2O emissions
- 4.2. NO emissions
- 4.3. N2 emissions
- 4.4. Denitrification as the source process of emissions summarised
- 5. Conclusions
- Acknowledgements
- References
ApplEnvironMicrobiol.-2014-Kozlowski-4930-5.pdf
Revision of N2O-Producing Pathways in the Ammonia-Oxidizing Bacterium Nitrosomonas europaea ATCC 19718
Jessica A. Kozlowski, Jennifer Price,* Lisa Y. Stein
University of Alberta, Department of Biological Sciences, Edmonton, Alberta, Canada
Nitrite reductase (NirK) and nitric oxide reductase (NorB) have long been thought to play an essential role in nitrous oxide (N2O) production by ammonia-oxidizing bacteria. However, essential gaps remain in our understanding of how and when NirK and NorB are active and functional, putting into question their precise roles in N2O production by ammonia oxidizers. The growth phenotypes of the Nitrosomonas europaea ATCC 19718 wild-type and mutant strains deficient in expression of NirK, NorB, and both gene products were compared under atmospheric and reduced O2 tensions. Anoxic resting-cell assays and in- stantaneous nitrite (NO2
�) reduction experiments were done to assess the ability of the wild-type and mutant N. europaea strains to produce N2O through the nitrifier denitrification pathway. Results confirmed the role of NirK for efficient substrate oxidation of N. europaea and showed that NorB is involved in N2O production during growth at both atmospheric and reduced O2 tensions. Anoxic resting-cell assays and measurements of instantaneous NO2
� reduction using hydrazine as an electron do- nor revealed that an alternate nitrite reductase to NirK is present and active. These experiments also clearly demonstrated that NorB was the sole nitric oxide reductase for nitrifier denitrification. The results of this study expand the enzymology for nitro- gen metabolism and N2O production by N. europaea and will be useful to interpret pathways in other ammonia oxidizers that lack NirK and/or NorB genes.
Ammonia-oxidizing bacteria (AOB) are obligate chemolitho-trophs that oxidize ammonia (NH3) through the intermedi- ate hydroxylamine (NH2OH) to nitrite (NO2
�) as their primary energy metabolism. During ammonia oxidation AOB produce gaseous nitrogen oxides, including nitrous oxide (N2O), a green- house gas (GHG) with more than 300 times the global-warming potential of CO2 (1), across a wide range of substrate and oxygen concentrations (2–4). Genes that encode nitrogen oxide reducta- ses, including a periplasmic copper-containing nitrite reductase (nirK) and a membrane-bound nitric oxide reductase (norB), are present in many closed AOB genome sequences (5), including that of Nitrosomonas europaea strain ATCC 19718 (6), the model or- ganism for this study. Previous work has identified two N2O-pro- ducing pathways in N. europaea, the pathway of hydroxylamine oxidation and the pathway of nitrifier denitrification. Generally, hydroxylamine oxidation is favored at atmospheric O2 tension (7, 8) and nitrifier denitrification is favored at low O2 tension (4, 9, 10). Although previous work has been done to describe the roles NirK and NorB may play in electron flow during substrate oxida- tion and NO2
- reduction to N2O (11, 12), many questions remain about the functionality of these gene products, particularly under reduced O2 tension, at which nitrifier denitrification becomes en- vironmentally relevant (13, 14). Furthermore, screening by low- stringency Southern blotting and PCR to identify DNA sequences with similarity to nirK revealed no hybridization signals from genomic DNA of Nitrosococcus mobilis Nc2, Nitrosomonas cryotol- erans Nm55, or Nitrosomonas communis Nm2 (15). In addition, the genome of the recently sequenced Nitrosomonas sp. strain Is79 showed no homologues to the norCBQD gene cluster (16). These observations suggest either that NirK and NorB are nonessential to the ammmonia oxidizer lifestyle or that alternate mechanisms of reducing nitrogen oxides are present in AOB that lack these particular nitrogen oxide reductases.
Previous work on a NirK-deficient strain of N. europaea grown at atmospheric O2 tension showed NirK activity to be important
in tolerance of the bacteria to NO2 � (17) as well as for their effi-
cient oxidation of NH3 and NH2OH (9, 10). Work on a NorB- deficient strain of N. europaea suggested that NorB is important for reduction of nitric oxide (NO) but not for net N2O production under atmospheric O2 tension (18). However, previous studies present conflicting evidence regarding whether NorB is essential for efficient oxidation of NH3 and NH2OH (10, 18). Conflicting results are also present in work on NirK-deficient N. europaea, particularly the role of NirK in pathways of N2O production. When grown in a chemostat, NirK-deficient N. europaea cells were unable to reduce NO2
� as an alternate terminal electron acceptor (10), in contrast to batch growth, in which, at reduced O2 tension, there was no difference in the ability of NirK-deficient cells to reduce NO2
� to N2O compared to that of wild-type N. europaea (9).
The functional roles of NirK and NorB in growth, substrate oxidation, and N2O production of N. europaea across a range of O2 tensions have not been fully elucidated. In this study, we com- pared the phenotypes of N. europaea wild-type, NirK-deficient, NorB-deficient, and NirK- plus NorB-deficient strains to solidify our understanding of the enzymology for N2O production as a function of variable O2 levels and to determine the necessity of NirK and NorB for growth, substrate oxidation, and NO2
� reduc- tion to N2O.
Received 28 March 2014 Accepted 28 May 2014
Published ahead of print 6 June 2014
Editor: R. E. Parales
Address correspondence to Lisa Y. Stein, [email protected].
* Present address: Jennifer Price, 6850 Jamieson Ave., Reseda, California, USA.
Copyright © 2014, American Society for Microbiology. All Rights Reserved.
doi:10.1128/AEM.01061-14
4930 aem.asm.org Applied and Environmental Microbiology p. 4930 – 4935 August 2014 Volume 80 Number 16
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MATERIALS AND METHODS Bacterial strains. Wild-type Nitrosomonas europaea ATCC 19718 was used as the native strain for this study. The nirK::Kan (nirK gene locus NE 0924) strain of N. europaea was created in a previous study (17) and was received as a gift from H. J. E. Beaumont. Confirmation of the nirK::Kan strain was done by PCR using primers nir10f (5=-GGG CGA CAT ACC CAA GAG TG-3=), nir10r (5=-CAA GCC TAT GGG GGT TTA TAG-3=), and nir26r (5=-GTC ATA GCT GTT TCC TGT GTG AAA TT-3=) as de- scribed previously (17).
norB::Gen and nirK::Kan norB::Gen N. europaea strains were created by following a methodology described elsewhere (19). Briefly, the norB:: Gen strain was generated by amplifying the norB gene (NE 2004) from N. europaea ATCC 19718 genomic DNA using primers Ne_2004F (5=-ACC CAG AAG CTT GCT TAC CC-3=) and Ne_2004R (5=-TGT TCG GTG ACG ATG ACA CT-3=). The amplified fragment was purified and ligated into the pGEM-T vector (Promega, Madison, WI). The ligation mixture was transformed into competent E. coli cells negative for both dam and dcm (New England BioLabs Inc., Ipswich, MA), and transformants were selected via blue-white screening on LB agar plates containing 0.5 mM isopropyl-�-D-thiogalactopyranoside (IPTG), 80 �g/ml 5-bromo-4- chloro-3-indolyl-�-D-galactopyranoside (X-Gal), and 100 �g/ml of am- picillin. Plasmids from positive recombinants were purified using a Wiz- ard Plus SV Minipreps DNA purification system kit (Promega) and digested with the KpnI restriction enzyme (New England BioLabs Inc.). The digest was run on a 0.8% agarose gel, and linearized vector was gel purified using the Wizard SV gel and PCR clean-up system kit (Promega). The gentamicin resistance cassette from the pUGM vector was digested with KpnI and gel purified (QIAquick gel extraction kit; Qiagen, Venlo, the Netherlands). The purified gentamicin cassette was then ligated into the previously KpnI-digested pGEM-T vector to disrupt the norB gene at nucleotide position 699 to 1347. The ligation mixture was transformed into E. coli JM109 cells, and positive transformants were selected on LB plates containing 100 �g/ml of ampicillin and 10 �g/ml of gentamicin. Positive recombinants were verified by PCR and Sanger sequencing using the BigDye Terminator cycle sequencing kit (Applied Biosystems, Foster City, CA). The plasmid with the correct construct was electroporated into prepared N. europaea cells (19) using an E. coli Pulser transformation apparatus (Bio-Rad Laboratories, Hercules, CA). Electroporated cells were inoculated into mineral salts medium (MSM) (20) without antibi- otics and incubated at 28°C without shaking. After 24 h, 5 �g/ml of gen- tamicin was added and cultures were monitored until turbidity was evi- dent and approximately 10 mM NO2
� was produced. Cell culture (1 ml) was then inoculated onto nitrocellulose membranes overlaying agar-so- lidified mineral medium to select single recombinant colonies as de- scribed previously (19). Cultures were PCR screened to confirm the loca- tion and orientation of the gentamicin resistance cassette within the norB gene that had recombined in the chromosome using primers Ne_2004F (as reported above), GenF (5= TGC CTC GGG CAT CCA AGC AG-3=), and GenR (GAG AGC GCC AAC AAC CGC TTC T-3=). The methods for creation of the nirK::Kan norB::Gen strain were identical to generation of the norB::Gen strain except that the nirK::Kan strain of N. europaea (17) was used as the recipient instead of wild-type N. europaea and 5 �g/ml of gentamicin and 30 �g/ml of kanamycin were added to the MSM of elec- troporated cells after 24 h of incubation as described above.
All N. europaea strains were grown in 500-ml Erlenmeyer flasks with 250 ml of MSM containing 25 mM (NH4)2SO4 (20). Cultures were incu- bated at 30°C in the dark with shaking. Inoculation of fresh medium used a 1% volume of culture in stationary phase, which was determined by NO2
� concentration (21). Concentrations of NO2 � were determined us-
ing a standard curve from 1 mM to 20 mM NaNO2, and stationary phase was achieved at 10 mM NO2
�. Growth experiments. Wild-type and mutant N. europaea cultures (1
ml) were inoculated into MSM (100 ml) in Wheaton bottles (250 ml) sealed with caps inlaid with butyl rubber stoppers. Cultures were initiated at atmospheric (ca. 22%) or hypoxic (ca. 5%) levels of O2. Hypoxia was
achieved by aseptically sparging the bottles with nitrogen gas and injecting pure O2 into the headspace. Final headspace O2 levels were confirmed by gas chromatography (GC-thermal conductivity detector [TCD] from Shi- madzu and molecular sieve column from Alltech, Deerfield, IL). O2 was measured again at the experimental endpoint (72 h) to determine the amount consumed. N2O was measured in the gas headspace at 24, 48, and 72 h by GC-TCD (Hayesep Q column). Headspace concentrations of O2 and N2O in the cultures were determined by comparison to standard curves using pure gases (Sigma-Aldrich). Total cell counts were done at 0, 24, 48, and 72 h using a Petroff-Hausser counting chamber and contrast light microscopy to follow the cells from exponential into stationary phase of growth. NO2
� concentrations were determined at 0, 24, 48, and 72 h by colorimetric assay as described above. NH2OH concentration was mea- sured during growth between 0 h and 72 h in increments of 6 h using a colorimetric assay (22). Statistical differences between measured values among the N. europaea strains and experimental conditions were evalu- ated using Student’s t test at a P value of �0.05.
Resting-cell assays. The wild-type and mutant strains of N. europaea were grown to stationary phase as described above. For each experiment, culture (1 ml) was transferred to a 12-ml vial sealed with a rubber stopper and aluminum crimp seal. The vial was sparged with nitrogen gas to anoxia. An electron donor (ascorbic acid; 1 mM) and electron shuttle (phenazine methosulfate; 0.1 mM) (23) were added to the culture via Hamilton syringe. The vial was left to sit at 30°C in the dark for 72 h to allow adequate time for reduction of NO2
�and accumulation of N2O. Headspace N2O concentration was measured at 0 and 72 h as described above. To confirm consistent anoxia, O2 was measured at 0 and 72 h using gas chromatography (GC-TCD from Shimadzu and molecular sieve col- umn from Alltech).
MR measurements. In preparation for instantaneous O2 consump- tion and NO2
� reduction experiments, the wild-type and mutant strains of N. europaea were grown in 250-ml Wheaton bottles in 100 ml of MSM to stationary phase. Cells were harvested by filtration on Supor200 0.2-�m filters (Pall, Ann Arbor, WI) and rinsed three times with sodium phosphate buffer (50 mM NaH2PO4, 2 mM MgCl2; pH 8) to wash away remaining NO2
� produced during growth. Approximately 5 � 1010 total cells were resuspended into 10 ml of sodium phosphate buffer in a 10-ml two-port microrespiratory (MR) chamber with fitted injection lids (Unisense, Aarhus, Denmark). O2 concentration was measuring using an OX-MR 500-�m-tip-diameter MR oxygen electrode (Unisense, Aarhus, Denmark), and N2O concentration was measuring using an N2O-500 N2O minisensor electrode with a 500-�m tip diameter (Unisense). Hy- drazine (N2H4) was added to the chamber as an electron donor for NO2
�
reduction at the beginning of each experiment at a concentration of 250 �M and again at a concentration of 125 �M after the cells had consumed more than half of the available O2. Once the cells had consumed all avail- able O2 they were left to sit for 5 to 10 min under anoxia. An absence of N2O production confirmed that no endogenous NO2
� was present, after which 2 mM NaNO2 was added to the chamber through the injection port. Instantaneous NO2
� reduction to N2O was measured for approxi- mately 10 min.
RESULTS Growth phenotype of N. europaea strains. N. europaea wild-type and mutant strains were grown under atmospheric (ca. 22%) and reduced (ca. 5%) O2 tensions to evaluate and compare the pheno- types of strains deficient in NirK, NorB, or both gene products. Growth experiments beginning at atmospheric O2 tension re- vealed that only the double mutant (nirK::Kan norB::Gen) strain had a significantly slower doubling time with respect to the other three strains; however, the amounts of NO2
� produced by both the nirK::Kan and double mutant strains were significantly less than that of the wild type (Fig. 1; Table 1). In contrast, the norB:: Gen strain showed no significant difference in doubling time or
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NO2 � production relative to those of the wild type under atmo-
spheric O2 tension (Fig. 1; Table 1). None of the cultures initiated at atmospheric O2 tension reduced the O2 headspace level to be- low 6%; however, the double mutant consumed significantly less O2 than the other strains, in congruence with its slower doubling time (Table 1). Previous studies have shown that batch growth of N. europaea with ample O2 is limited by acidification of the me- dium, which reduces availability of NH3 to the cells (24). Hence, the cells entered stationary phase prior to consuming all of the available O2 in the present experiments due to medium acidifica- tion and not O2 limitation.
Growth experiments beginning at reduced O2 tension again revealed a significant reduction in doubling time for the double mutant compared with those of the other strains (Fig. 1; Table 1). The double mutant also showed a significant accumulation of NO2
� in comparison to those of the other strains (Table 1). Al- though NH2OH was assayed from all of the cultures under all O2 tensions, the assay was unable to detect significant differences over time or between strains (data not shown).
Together, the results suggest that regardless of initial O2 levels, NorB alone played no significant role in the growth phenotype or substrate oxidation efficiency of N. europaea; however, NirK was
essential for efficient substrate oxidation efficiency, especially dur- ing growth initiated at atmospheric O2 tension. The lack of both gene products significantly slowed growth of N. europaea and also allowed for significant accumulation of NO2
� during hypoxic growth relative to the other strains.
Effects of NirK and NorB absence on N2O production by N. europaea under variable O2 tensions. To evaluate the roles of NirK and NorB in the pathways of N2O production for N. euro- paea, N2O concentration in the gas headspace was measured dur- ing growth experiments initiated at both atmospheric and re- duced O2 tension and in resting-cell assays in the absence of O2. In confirmation of prior studies, the concentration of N2O in the headspace of the nirK::Kan strain at the endpoint (72 h) of growth in cultures initiated with atmospheric O2 was approximately 15 times that of the wild-type strain (Fig. 2) (9, 17). However, the resulting N2O measured after hypoxic growth and in the anoxic resting-cell assay revealed no difference between wild-type and nirK::Kan strains of N. europaea. In contrast, the norB::Gen strain produced ca. 20% less N2O than did the wild type following growth under atmospheric O2 and approximately 70% less N2O following growth under reduced O2 (Fig. 2). No N2O was detected in the anoxic resting-cell assay of norB::Gen cells. The double mu- tant produced an amount of N2O similar to that produced by the norB::Gen strain when grown under atmospheric O2 but was un-
T ot
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Time (h)
0 24 48 72
Wild type nirK::Kan norB::Gen nirK::Kan norB::Gen
Wild type nirK::Kan norB::Gen nirK::Kan norB::Gen
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Wild type nirK::Kan norB::Gen nirK::Kan norB::Gen
FIG 1 Growth curves for N. europaea strains initiated at 22% O2 (A) and 4.6% O2 (B). Data represent mean values � SEs (ca. 22% O2, n � 8, ca. 4.6% O2, n � 6).
TABLE 1 Doubling time, total nitrite production, and percent remaining headspace O2 for wild-type and mutant strains of N. europaea ATCC 19718 cultivated under atmospheric and reduced O2 tensions
a
Organism description
Value at indicated oxygen tension
Doubling time (h) Total NO2
�-N produced (mM � 1010 cells�1) Remaining O2 in headspace (%)
22% 4.6% 22% 4.6% 22% 4.6%
Wild type 6.5 d (0.6) 8.7 d (0.3) 7.3 bd (0.02) 4.2 d (0.4) 6.7 d (9.2e�5) 1.0 (6.7e�4) nirK::Kan 6.7 d (0.2) 9.3 d (0.8) 5.8 a (0.01) 4.1 d (0.8) 6.8 d (1.8e�3) 1.2 d (2.4e�3) norB::Gen 7.1 d (0.3) 9.2 d (0.2) 6.6 d (0.02) 4.3 d (0.4) 6.7 d (1.7e�3) 1.2 (2.4e�3) nirK::Kan norB::Gen 9.6 abc (0.4) 14.3 abc (1.9) 5.6 ac (0.04) 8.0 abc (1.1) 8.0 abc (1.8e�3) 0.8 b (1.1e�4) a Doubling times were calculated over the 0- to 48-h period of exponential growth. Total NO2
�-N produced and remaining O2 in headspace were determined at 72 h for all cultures. Averages and SEs (in parentheses) were calculated from 8 and 6 replicated experiments for cultures grown under 22 and 4.6% O2, respectively. Significant differences (P � 0.05) are denoted by different letters as follows: “a,” strain versus wild type; “b,” strain versus nirk::Kan strain; “c,” strain vs. norB::Gen strain; and “d,” strain versus nirK::Kan norB:: Gen strain.
22% 4.6% <1%
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FIG 2 Total N2O produced by all strains after 72 h of growth at high (n � 8) and low (n � 6) oxygen. N2O profiles under anoxia (n � 8) were collected during resting-cell assays. Data are presented as means � SEs.
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able to produce measurable N2O after 72 h when grown under hypoxia or in the anoxic resting-cell assay.
Instantaneous O2 consumption and N2O production by wild-type and mutant N. europaea strains. The anoxic resting- cell assays revealed that N. europaea lacking NorB expression did not produce a measureable quantity of N2O in the gas headspace from the reduction of available NO2
� (10 mM) in the medium. Therefore, instantaneous NO2
� reduction experiments were con- ducted to confirm whether NorB is essential to NO2
� reduction to N2O by N. europaea and whether an alternative nitrite reductase to NirK was operating in the nirK::Kan strain to produce the same amount of N2O as observed in the wild type (Fig. 2).
Instantaneous NO2 � reduction experiments were conducted
in microelectrode chambers with the use of a non-nitrite-forming intercellular electron donor, N2H4, and microelectrodes for O2 and N2O. The cells were allowed to consume all of the O2 in the microelectrode chamber via oxidation of N2H4, after which NaNO2
� was added. Both the wild type and the nirK::Kan strain produced approximately 8 �mol N2O per liter per min, confirm- ing the presence of an alternate nitrite reductase activity in nirK:: Kan cells (Fig. 3A and B). Both the norB::Gen and double mutant strains showed only background levels of N2O production from electrode drift upon the addition of NaNO2, confirming that ac- tivity of NorB is essential to this process (Fig. 3C and D).
DISCUSSION Function of NirK and NorB in efficient substrate oxidation and growth of N. europaea under variable O2 tensions. The reduced production of NO2
� by both the nirK::kan and double mutant strains of N. europaea when grown at atmospheric O2 tension confirm previous reports of the requirement of NirK for efficient oxidation of NH3 to NO2
� during batch (9, 17) and chemostat (10) cultivation. Slowed substrate oxidation in the NirK-deficient strain of N. europaea was previously suggested to be caused by interruption of electron flow from NH2OH to NO2
� due to the inability of HAO to pass electrons on to NirK through cytochrome c electron carriers (9). A diminished ability of NirK-deficient N. europaea to oxidize exogenous NH2OH during growth strength- ens the hypothesis that NirK functions aerobically to facilitate efficient substrate oxidation (9). Furthermore, extensive accumu- lation of N2O in the gas headspace of nirK::Kan cultures during growth under atmospheric O2 tension validates previous mea- surements from growth of this strain in batch (9, 17) and chemo- stat (10) cultures. A possible explanation for this phenotype is that in the presence of high NH2OH concentrations the HAO enzyme produces NO due to incomplete oxidation of NH2OH, which is then enzymatically reduced to N2O (14). Our data suggest that enzymatic reduction of NO via NorB could lead to production of N2O from NH2OH. During growth under atmospheric O2 ten- sion, the 20% reduction in N2O produced by the norB::Gen strain could be accounted for by the lack of NorB activity, with the re- maining N2O being produced from an alternate nitric oxide re- ductase (Fig. 2).
In the double mutant strain, growth and net NO2 � production
were likely slowed during growth at atmospheric O2 by both the lack of NirK in speeding substrate oxidation and also from the lack of NorB in preventing toxic accumulation of NO. When culti- vated under reduced O2, the alternate nitrite reductase activity could cause the double mutant cultures to accumulate an excess of NO that could not be removed as NorB is more active at low O2. This excess NO could both slow cellular growth and result in net NO2
� production due to chemical reactions of nitrogen oxides (NOx) in the culture medium. Future work comparing NO accu- mulation between the wild type and mutant strains of N. europaea under variable O2 tension would assist in validating these hypoth- eses.
The growth rate of N. europaea lacking NorB expression alone was not significantly impaired, which is in agreement with data from a previous study (18); however, in contrast to that study, N2O production by norB::Gen cells was significantly lower than that of the wild type (Fig. 2). Schmidt (10) showed that NorB- deficient N. europaea had a significantly lower growth rate and yield than did the wild type, an N2O production profile similar to that of the NirK-deficient strain, and significantly larger amounts of NH2OH released to the growth medium than did the wild type, sug- gesting similar inefficiency of substrate oxidation by both NirK- and NorB-deficient strains. Under the growth conditions of the present study, however, the results obtained by Schmidt (10) were not vali- dated. Rather, our results suggest that the absence of NorB expression alone in N. europaea had no effect on growth or substrate oxidation rates or on NH2OH accumulation but did result in diminished N2O production in comparison to that of the wild type.
NorB, but not NirK, is required for anoxic reduction of NO2
� to N2O in N. europaea. The inability of N. europaea strains
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FIG 3 Instantaneous oxygen consumption and nitrite reduction by wild-type (A), nirK::Kan (B), norB::Gen (C), and nirK::Kan norB::Gen (D) strains of N. europaea. Data are single representatives of reproducible results.
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lacking NorB expression to make measurable N2O in anoxic rest- ing-cell assays (Fig. 2) and instantaneous NO2
� reduction assays (Fig. 3) pointed to NorB as the essential nitric oxide reductase involved in NO reduction in the absence of O2. The significant accumulation of NO2
� only during hypoxic growth of the double mutant strain (Table 1) could be explained by chemical decay of highly reactive NO that may accumulate from activity of the alter- native nitrite reductase working in the absence of both NirK and NorB enzymes. However, the lack of similar results with the norB:: Gen strain suggests that the activity of NirK in the absence of NorB has an effect on nitrogen oxide metabolism that is substantially different from that of the alternative NO2
� reductase. Thus, ex- ploration of alternative nitrogen oxide reductases active in N. eu- ropaea with and without expression of NirK and/or NorB will be helpful to elucidate the enzymology behind these phenotypes.
Amended pathways of N2O production in N. europaea and other AOB. The most important finding of this study is the dem- onstration that NirK is not essential to the nitrifier denitrification pathway of N. europaea, as has been assumed for many years in the literature. The similar headspace N2O levels produced by both wild-type and nirK::Kan strains of N. europaea during growth un- der reduced O2 tension, in anoxic resting-cell assays, and in anoxic instantaneous nitrite reduction experiments all revealed that an alternate nitrite reductase to NirK is active in the production of N2O by N. europaea ATCC 19178 (Fig. 4). It was previously sug- gested that an alternate nitrite reductase may be active in N. euro- paea (9); however, no other known homologues to nitrite reduc- tase genes have been identified in its closed genome (6). One possible candidate for an alternate nitrite reductase in AOB is C-terminally truncated HAO (HaoA=; NE0962, 2044, 2339) (25). Evolutionary reconstructions showed that HAO evolved from an octaheme cytochrome c nitrite reductase (26), and gene expres- sion of HaoA= in the methanotrophic strain Methylococcus capsu- latus strain Bath was induced in the presence of ammonia (25). M. capsulatus strain Bath can reduce NO2
� to N2O in the presence of NH3 and NO2
� (27) even though homologues to both nirK and nirS NO-forming nitrite reductases are absent from the closed genome sequence (28). Although nirK genes have been found in the genomes of most AOB (15) and ammonia-oxidizing archaea (29, 30) and nirK has long been used as a marker for denitrifica- tion activity in the field of microbial ecology, the present study shows that at least in Nitrosomonas europaea ATCC 19718, nirK is not a marker for denitrification but rather should be considered a marker for ammonia oxidation. It should be noted that due to differences in gene phylogenies and neighborhoods in the Ni- trosospira spp. and Nitrosococcus spp. (9), the specific roles of NirK and NorB should be physiologically examined within strains of
these genera to determine if the nitrifier denitrification pathways share similar inventories and are similarly regulated among the ammonia-oxidizing bacteria.
In addition to an alternate nitrite reductase, our results also demonstrate that NorB activity plays a role in the hydroxylamine oxidation pathway of N2O production by N. europaea ATCC 19718 (Fig. 4). It is also possible that alternate nitric oxide reduc- tases are active in Nitrosomonas spp. For instance, a complete tran- scriptome of the nirK::Kan strain showed increased expression of genes for norSY (originally annotated as coxAB2), an alternative nitric oxide reductase, in comparison to the wild-type strain when grown under normal oxic conditions (7). Furthermore, Nitro- somonas eutropha C91 grown under continuous cultivation for 3 months in the presence of nitrogen dioxide (NO2) gas showed increased expression of NorY protein (31). These observations suggest that NorY nitric oxide reductase could potentially con- tribute to N2O production along with NorB during growth of N. europaea particularly under atmospheric O2 levels (Fig. 2); how- ever, this hypothesis remains to be validated.
Our results showing the inability of the norB::Gen and double mutant strains of N. europaea to reduce NO2
� to N2O in instan- taneous NO2
� reduction experiments (Fig. 3C and D), even with a readily available source of electrons, demonstrate that NorB is the sole enzyme involved in N2O production through the nitrifier denitrification pathway (Fig. 4). While these results are in agree- ment with those of Cantera and Stein and of Schmidt (9, 10), a discrepancy remains regarding the role of NirK in this pathway.
Conclusions. This study is unique in its comparison of pheno- types of N. europaea lacking expression of NirK, NorB, and both enzymes together. Furthermore, our assays allowed comparison of phenotypes under O2 initially present at atmospheric, hypoxic, and anoxic levels, each having a different effect on N2O produc- tion by the two characterized pathways in N. europaea ATCC 19718 (1–4, 14). The main conclusions from this study are that (i) NirK, but not NorB, plays an essential role in efficient substrate oxidation under atmospheric O2 tension; (ii) an alternate nitrite reductase to NirK is active in N. europaea under both hypoxic and anoxic conditions; (iii) NorB and/or other NOR enzymes are ac- tive in N. europaea during growth under atmospheric O2 tension; and (iv) NorB is the only nitric oxide reductase active in the nitri- fier denitrification pathway. These results suggest that AOB have diverse enzymology beyond NirK and NorB leading to N2O pro- duction that remains to be characterized.
ACKNOWLEDGMENTS
This work was supported by a graduate fellowship awarded to J.A.K. by Alberta Innovates Technology Futures and NSERC Discovery Award 371544-09 to L.Y.S.
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NH3
NO2 -
NH2OH
AMO
HAO 1
N2O aNorB Nitrifier Denitrification
N2O Hydroxylamine Oxidation NO
HAO2
aNIR
aNorB
NirK
NO
FIG 4 Amended pathways of N2O production by Nitrosomonas europaea ATCC 19718. AMO, ammonia monooxygenase; HAO, hydroxylamine oxi- doreductase; NirK, nitrite reductase; NorB, nitric oxide reductase; NIR, un- identified alternate nitrite reductase. The role of enzymes in gray were charac- terized in previous studies as follows: AMO, reference 32; HAO1, reference 33; HAO2, reference 34; and NirK, reference 9). The roles of enzymes denoted with superscript “a” are from the present study.
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16. Bollmann A, Sedlacek C, Norton J, Laanbroeck HJ, Suwa Y, Stein LY, Klotz MG, Arp D, Sayavedra-Soto L, Lu M, Bruce D, Detter C, Tapia R, Han J, Woyke T, Lucas S, Pitluck S, Pennacchio L, Nolan M, Land M, Huntemann M, Deshpande S, Han C, Chen A, Kyrpides N, Mavromatis K, Markowitz V, Szeto E, Ivanova N, Mikhailova N, Pagani I, Pati A, Peters L, Ovchinnikova G, Goodwin L. 2013. Complete genome se- quence of Nitrosomonas sp. Is79 —an ammonia oxidizing bacterium adapted to low ammonium concentrations. Stand. Genomic Sci. 7:469 – 482.
17. Beaumont HJE, Hommes NG, Sayavedra-Soto LA, Arp DJ, Arciero DM, Hooper AB, Westerhoff HV, van Spanning RJM. 2002. Nitrite reductase of Nitrosomonas europaea is not essential for production of gas- eous nitrogen oxides and confers tolerance to nitrite. J. Bacteriol. 184: 2557–2560. http://dx.doi.org/10.1128/JB.184.9.2557-2560.2002.
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28. Ward N, Larsen Ø, Sakwa J, Bruseth L, Khouri H, Durkin AS, Dimitrov G, Jiang L, Scanlan D, Kang KH, Lewis M, Nelson KE, Methé B, Wu M, Heidelberg JF, Paulsen IT, Fouts D, Ravel J, Tettelin H, Ren Q, Read T, DeBoy RT, Seshadri R, Salzberg SL, Jensen HB, Birkeland NK, Nelson WC, Dodson RJ, Grindhaug SH, Holt I, Eidhammer I, Jonasen I, Vanaken S, Utterback T, Feldblyum TV, Fraser CM, Lillehaug JR, Eisen JA. 2004. Genomic insights into methanotrophy: The complete genome sequence of Methylococcus capsulatus (Bath). PLoS Biol. 2(10):e303. http: //dx.doi.org/10.1371/journal.pbio.0020303.
29. Bartossek R, Nicol GW, Lanzen A, Klenk H-P, Schleper C. 2010. Homologues of nitrite reductases in ammonia-oxidizing archaea: Diver- sity and genomic context. Environ. Microbiol. 12:1075–1088. http://dx .doi.org/10.1111/j.1462-2920.2010.02153.x.
30. Hatzenpichler R. 2012. Diversity, physiology, and niche differentiation of ammonia-oxidizing archaea. Appl. Environ. Microbiol. 78:7501–7510. http://dx.doi.org/10.1128/AEM.01960-12.
31. Kartal B, Wessels HJCT, van der Biezen E, Francoijs K-J, Jetten MSM, Klotz MG, Stein LY. 2012. Effects of nitrogen dioxide and anoxia on global gene and protein expression in long-term continuous cultures of Nitrosomonas eutropha C91. Appl. Environ. Microbiol. 78:4788 – 4794. http://dx.doi.org/10.1128/AEM.00668-12.
32. Hyman MR, Wood PM. 1983. Methane oxidation by Nitrosomonas eu- ropaea. Biochem. J. 212:31–37.
33. Hooper AB, Maxwell PC, Terry KR. 1978. Hydroxylamine oxidoreductase from Nitrosomonas: absorption spectra and content of heme and metal. Biochemistry 17:2984 –2989. http://dx.doi.org/10.1021/bi00608a007.
34. Pacheco AA, McGarry J, Kostera J, Corona A. 2011. Techniques for investigating hydroxylamine disproportionation by hydroxylamine oxi- doreductases. Methods Enzymol. 486:447– 463. http://dx.doi.org/10.1016 /B978-0-12-381294-0.00020-1.
Revision of N2O-Producing Pathways in N. europaea
August 2014 Volume 80 Number 16 aem.asm.org 4935
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- Revision of N2O-Producing Pathways in the Ammonia-Oxidizing Bacterium Nitrosomonas europaea ATCC 19718
- MATERIALS AND METHODS
- Bacterial strains.
- Growth experiments.
- Resting-cell assays.
- MR measurements.
- RESULTS
- Growth phenotype of N. europaea strains.
- Effects of NirK and NorB absence on N2O production by N. europaea under variable O2 tensions.
- Instantaneous O2 consumption and N2O production by wild-type and mutant N. europaea strains.
- DISCUSSION
- Function of NirK and NorB in efficient substrate oxidation and growth of N. europaea under variable O2 tensions.
- NorB, but not NirK, is required for anoxic reduction of NO2− to N2O in N. europaea.
- Amended pathways of N2O production in N. europaea and other AOB.
- Conclusions.
- ACKNOWLEDGMENTS
- REFERENCES
Pathways and key intermediates required for obligate aerobic ammonia-dependent chemolithotrophy.pdf
ORIGINAL ARTICLE
Pathways and key intermediates required for obligate aerobic ammonia-dependent chemolithotrophy in bacteria and Thaumarchaeota
Jessica A Kozlowski1, Michaela Stieglmeier2,3, Christa Schleper2, Martin G Klotz4,5 and Lisa Y Stein1 1Department of Biological Sciences, University of Alberta, Edmonton, Alberta, Canada; 2Department of Ecogenomics and Systems Biology, Division Archaea Biology and Ecogenomics, University of Vienna, Wien, Austria; 3Department of Biology I, Ludwig-Maximilians-University of Munich, Biocenter, Planegg-Martinsried, Germany; 4Department of Biology, Queens College, The City University of New York, Flushing, NY, USA and 5Institute of Marine Microbes & Ecospheres and State Key Laboratory of Marine Environmental Science, Xiamen University, Xiamen, China
Chemolithotrophic ammonia-oxidizing bacteria and Thaumarchaeota are central players in the global nitrogen cycle. Obligate ammonia chemolithotrophy has been characterized for bacteria; however, large gaps remain in the Thaumarchaeotal pathway. Using batch growth experiments and instantaneous microrespirometry measurements of resting biomass, we show that the terrestrial Thaumarchaeon Nitrososphaera viennensis EN76T exhibits tight control over production and consumption of nitric oxide (NO) during ammonia catabolism, unlike the ammonia-oxidizing bacterium Nitrosospira multiformis ATCC 25196T. In particular, pulses of hydroxylamine into a microelectrode chamber as the sole substrate for N. viennensis resulted in iterative production and consumption of NO followed by conversion of hydroxylamine to nitrite. In support of these observations, oxidation of ammonia in growing cultures of N. viennensis, but not of N. multiformis, was inhibited by the NO-scavenger PTIO. When based on the marginal nitrous oxide (N2O) levels detected in cell-free media controls, the higher levels produced by N. multiformis were explained by enzyme activity, whereas N2O in N. viennensis cultures was attributed to abiotic reactions of released N-oxide intermediates with media components. Our results are conceptualized in a pathway for ammonia-dependent chemolithotrophy in Thaumarchaea, which identifies NO as an essential intermediate in the pathway and implements known biochemistry to be executed by a proposed but still elusive copper enzyme. Taken together, this work identifies differences in ammonia-dependent chemolithotrophy between bacteria and the Thaumarchaeota, advances a central catabolic role of NO only in the Thaumarchaeotal pathway and reveals stark differences in how the two microbial cohorts contribute to N2O emissions. The ISME Journal advance online publication, 16 February 2016; doi:10.1038/ismej.2016.2
Introduction
Ammonia-oxidizing archaea, in the phylum Thau- marchaeota, and ammonia-oxidizing bacteria are abundant and diverse microorganisms that control the oxidation of ammonia (NH3) to nitrite (NO2−) in the global biogeochemical nitrogen cycle. Through many decades of research, the biochemical pathway for chemolithotrophic growth of ammonia-oxidizing bacteria has been principally elucidated (Sayavedra- Soto and Arp, 2011); however, this pathway has yet
to be characterized in the more recently discovered thaumarchaeotal ammonia-oxidizers. This lesser understanding is largely due to the difficulty of growing reliable and sufficient biomass from pure cultures for performing physiological experiments, thus making identification of the genetic inventory that supports chemolithotrophic growth of the thaumarchaeotal ammonia-oxidizers a challenge. In contrast, the pathways for the autotrophic assimila- tion of carbon have been identified in both cohorts (Arp et al., 2007; Könneke et al., 2014).
Previous experiments with the marine isolate Nitrosopumilus maritimus SCM1 indicated that ammonia oxidation is dependent on the activity of the ammonia monooxygenase enzyme and (an) unknown enzyme(s) that convert(s) hydroxylamine (NH2OH) to NO2− and provide electrons for energy
Correspondence: LY Stein, Department of Biological Sciences, University of Alberta, CW 405 Biological Sciences Building, Edmonton, Alberta T6G2E9, Canada. E-mail: [email protected] Received 17 April 2015; revised 14 December 2015; accepted 24 December 2015
The ISME Journal (2016), 1–10 © 2016 International Society for Microbial Ecology All rights reserved 1751-7362/16 www.nature.com/ismej
conservation (Vajrala et al., 2012). In ammonia- oxidizing bacteria, this second step is performed by hydroxylamine dehydrogenase (EC 1.7.2.6); however, no homologues of hydroxylamine dehydrogenase- encoding genes have been identified in genome sequences obtained from any pure or enrichment culture of Thaumarchaea (Walker et al., 2010; Kim et al., 2011; Tourna et al., 2011; Spang et al., 2012). In addition to NH2OH, there is also evidence that nitric oxide (NO) plays an important role in the Thaumarch- aeotal but not in the bacterial ammonia oxidation pathway (Shen et al., 2013; Martens-Habbena et al., 2015). Martens-Habbena et al. (2015) demonstrated that NO accumulated in N. maritimus SCM1 cultures during active oxidation of NH4Cl in a closed microrespirometry chamber, and was released at higher levels under saturating versus non-saturating availability of NH4Cl. Exposure to increasing concen- trations of an NO-scavenging compound over a 24 h period resulted in decreased levels of nitrite produc- tion in batch cultures of ammonia-oxidizing Thau- marchaea, but not bacteria (Martens-Habbena et al., 2015). The authors concluded that NO was either released as a free intermediate during ammonia oxidation by N. maritimus, or it could serve a functional role as an electron delivery mechanism to ammonia monooxygenase, an idea that has been proposed previously (Schleper and Nicol, 2010).
Although the detection of nitrous oxide (N2O) has been reported for both enrichments and pure cultures of Thaumarchaea engaged in ammonia oxidation (Santoro et al., 2011; Loscher et al., 2012; Jung et al., 2014; Stieglmeier et al., 2014b), the isotope data reported by Stieglmeier et al. (2014b) revealed that ammonia-oxidizing Thaumarchaea cannot enzymatically reduce NO2− to N2O via NO in the pathway known as ‘nitrifier denitrification’. Several publications have suggested that ammonia- oxidizing Thaumarchaea are a major source of N2O to the environment based on their relative abun- dance in oxic environments, the isotopic signature of the detected N2O, and that the authors failed to detect known bacterial denitrification genes and pertinent activities (Santoro et al., 2011; Loscher et al., 2012; Jung et al., 2014). Yet, control experiments to verify or falsify chemical formation of N2O facilitated by interaction of Thaumarchaeo- tal metabolites with components of the cultivation or incubation media or assay solutions remain absent from the literature. It should be noted that interactions of ammonia oxidation intermediates with iron, manganese, and organic compounds could generate substantial amounts of N2O under environmentally relevant conditions (Zhu-Barker et al., 2015).
The present study addresses critical ecophysiolo- gical questions about how two different cohorts of microorganisms, simultaneously involved in the biogeochemical nitrogen cycle through ammonia- oxidation, vary in their contributions, particularly to production of nitrous oxide. This study also furthers
the observation of NO as an intermediate for ammonia chemolithotrophy in the terrestrial Thau- marchaeon Nitrososphaera viennensis strain EN76T (Stieglmeier et al., 2014a) by examining its complete profile of NO production and consump- tion during substrate oxidation at oxic conditions and the transition into an extended period of anoxia. In contrast to the above referenced studies of ‘N2O production’ by ammonia-oxidizing Thau- marchaea, our results do not support any scenario in which N. viennensis enzymatically reduces NO to N2O through a denitrification pathway. Instead, the results support that N2O was formed abiotically from NO by interaction with media components or with debris in killed cell controls. We further demonstrated that NO is an active and necessary intermediate during the oxidation of NH2OH to NO2− in ammonia-oxidizing Thaumarch- aea rather than participating directly in the oxida- tion of NH3 to NH2OH as suggested previously (Schleper and Nicol, 2010). Based on these results, a new pathway for obligate ammonia-dependent chemolithotrophy for ammonia-oxidizing Thau- marchaea is proposed that implicates a novel copper enzyme to perform a biochemistry known to occur in ammonia-oxidizing bacteria facilitated by heme-containing cytochrome c.
Materials and methods Strains and cultivation N. viennensis strain EN76T was maintained at 37 °C in 50 ml freshwater medium (FWM) supplemented with 2 mM NH4Cl, 0.5 mM sodium pyruvate and 50 μg/ml carbenicillin and buffered with HEPES (Tourna et al., 2011; Stieglmeier et al., 2014a) and inoculated at 4% v/v. Cultures were grown in Wheaton bottles (150 ml) sealed with caps inlaid with grey butyl rubber stoppers. Nitrosospira multiformis ATCC 25196T was maintained at 28 °C in 100 ml HEPES-buffered HK medium (HKM) (Krümmel and Harms, 1982) containing 3 mM ammonium and phenol red as pH indicator (pH of 7.5–8) and inoculated at 5% v/v into 250 ml Wheaton bottles. The pH of N. multiformis cultures was maintained with regular additions of 10% NaHCO3.
Growth experiments with NO-scavenger PTIO For monitoring activity in the presence of an NO- scavenging compound, N. viennensis was culti- vated in 20 ml FWM. N. multiformis was cultivated in 20 ml phosphate-buffered mineral medium (Skinner and Walker, 1961) amended with 1 mM NH4Cl and pH was adjusted regularly with 5% Na2CO3. In early to mid-exponential phase of growth, 150 μM of 2-phenyl-4,4,5,5,-tetramethylimidazoline-1- oxyl 3-oxide (PTIO; Sigma-Aldrich, Vienna, Austria), a chemical that scavenges NO (Goldstein et al., 2003)
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was injected into the cultures. Ammonium con- sumption and nitrite production were measured over a period of 6–8 days using standard colorimetric assays (Clesceri et al., 1998) and N2O was measured via GC (AGILENT 6890N, Vienna, Austria; injector: 120 °C, detector: 350 °C, oven: 35 °C, carrier gas: N2) in connection with an automatic sample-injection system (DANI HSS 86.50, Head-space-Sampler, Sprockhövel, Germany). Detailed sampling and sample preparation has been described previously (Stieglmeier et al., 2014a).
Instantaneous measurement of NO and N2O during oxidation of ammonia In preparation for experiments measuring instanta- neous O2 consumption and either NO or N2O production, N. viennensis was inoculated at 4% v/v into 2 l of HEPES-buffered FWM and N. multiformis was inoculated at 5% v/v into 250 ml HKM. Cells were harvested at late exponential phase (N. viennen- sis, 1–1.5 mM NO2−; N. multiformis, 2–2.5 mM NO2−) by filtration on Supor200 0.2 μm filters (Pall, Ann Arbor, WI, USA) and rinsed three times with substrate-free media (N. viennensis, FWM; N. multiformis, HKM). Washed cells (N. viennensis, ca. 1 × 1011 total cells; N. multiformis, ca. 1 × 1010 total cells) were re- suspended into 10 ml of substrate-free growth medium for each strain in a 10 ml two-port microrespiratory (MR) chamber with fitted injection lids (Unisense, Aarhus, Denmark). Cell concentrations for microre- spirometry experiments were chosen on the basis of comparable oxygen consumption rates between the two strains. O2 concentration was measured using an OX-MR 500 μm tip diameter MR oxygen electrode (Unisense), N2O concentration was measured using an N2O-500 N2O minisensor electrode with 500 μm tip diameter (Unisense), and NO was measured using an ami-600 NO sensor with 600 μm tip diameter (Innovative Instruments Inc., Tampa, FL, USA). The availability of O2 in the MR chamber, a closed system, corresponded to either ca. 207 μM O2 (FWM) or ca. 243 μM O2 (HKM) respectively, based on equili- brium O2 concentration at operating temperatures and medium salinities. For microrespirometry experiments involving ammonia oxidation, cells were provided 2 mM NH4Cl. The microrespirome- try chamber was maintained at 37 °C and 28 °C for measurements with N. vienennsis and N. multi- formis cells, respectively, reflecting their optimal growth temperatures.
Instantaneous measurement of NO from N. viennensis during oxidation of NH2OH For experiments measuring the oxidation of NH2OH (99.999% purity, Sigma-Aldrich, St Louis, MO, USA), N. viennensis was provided with multiple additions of 200 μM NH2OH (based on chamber volume) to maintain a steady rate of O2 consumption. NO production was measured until O2 was
undetectable in the chamber. Samples were taken post-experiment for NO2− measurements.
Instantaneous ammonia and hydroxylamine oxidation by N. viennensis in the presence of PTIO Microrespirometry experiments with the NO- scavenger PTIO were performed with N. viennensis cells harvested as described above; cells were incubated with 200 μM PTIO in the dark with shaking at 37 °C for 1 h prior to adding the cells to a 2 ml 1-port MR chamber at 37 °C for the measure- ment of NH4+- and NH2OH-dependent O2 consump- tion (Supplementary Figure S1). Confirmation of the NO-scavenging activity of PTIO was confirmed chemically by addition of 1 μl PAPA NONOate ((Z)- 1-[N-(3-aminopropyl)-N-(n-propyl)amino]diazen-1- ium-1,2-diolate; Cayman Chemical, Ann Arbor, MI, USA; half-life of 15 min at 37 °C liberating 2 moles of NO per mole of parent compound) to FWM in the 2 ml MR chamber with the NO sensor at 37 °C. Once the rate of NO release from PAPA NONOate slowed, 200 μM PTIO was added to the chamber and NO disappearance was immediately measured. After ~ 7 min of NO-chelation by PTIO, another 1 μl of PAPA NONOate was added but NO levels remained below detection levels (Supplementary Figure S2).
Instantaneous measurement of N2O from media and killed-cell controls To measure the abiotic production of N2O from either FWM or HKM, 10 ml of cell-free media was added to the 10 ml MR chamber. The NO-donor MAHMA NONOate (6-(2-Hydroxy-1-methyl-2-nitro- sohydrazino)-N-methyl-1-hexanamine, NOC-9; Cay- man Chemical) was added to the MR chamber in increasing additions of 20–100 μl, which is equiva- lent to the release of ca. 1.1–5.5 μM NO, or in a single addition of 100 μl (ca. 5.5 μM NO). The half-life of MAHMA NONOate at pH 7.4 is 1 min and 3 min at 37 °C and 22–25 °C, respectively. N2O production was measured using the N2O microelectrode during the decay of 1 mol MAHMA NONOate into 2 moles NO in either FWM or HKM. Experiments were performed at 37 °C and 28 °C for FWM and HKM, respectively, after sparging to ca. 0–3% O2 saturation with N2 (Praxair) as determined by O2 electrode. Chemical controls to confirm that FWM alone did not react with NH2OH to form measureable NO involved addition of 200, 400 and 600 μM NH2OH to sparged (ca. 0–3% O2) FWM or FWM+200 μM NO2−
to reflect maximum concentration available once cells depleted MR chamber O2 (Supplementary Figures S3a and b). Control experiments with heat killed N. viennensis (ca. 1 × 1011 cells) were per- formed in sparged FWM+NaNO2 (200 μM) with measurement of NO and N2O upon addition of 200 μM NH2OH (Supplementary Figure S4) to deter- mine whether NH2OH interacts with cellular debris.
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Results
Effects of the NO-scavenger PTIO on N2O levels measured in cultures of N. viennensis and N. multiformis To investigate the role of NO in chemolithotrophic oxidation of ammonia to nitrite and generation of N2O in these two organisms, batch cultures of N. viennensis or N. multiformis were grown to mid-log phase, at which point PTIO (150 μM) was added (Figure 1). Addition of PTIO resulted in the immediate saturation of N2O levels in either culture as would be expected in the absence or decreasing levels of NO intermediates. However, in cultures of N. viennensis, PTIO also caused an inhibition of both ammonium consumption and nitrite production (Figure 1a), whereas in cultures of N. multiformis, ammonia oxidation and nitrite production continued at the same rate as before PTIO addition (Figure 1b). These results indicated that NO is an essential, dynamic, intermediate in the process of ammonia oxidation to nitrite and thus ammonia-dependent chemolithotrophy for N. viennensis, but not for N. multiformis.
Dynamics of NO and N2O production during and following ammonia oxidation The effect of PTIO on growing cultures of N. viennensis and N. multiformis indicated differ- ent requirements for NO during ammonia oxida- tion. Using a MR chamber, the dynamics of NO production and consumption were measured dur- ing and after ammonia oxidation (2 mM NH4Cl) by N. viennensis or N. multiformis as determined by O2 consumption profiles (Figures 2a and c). To achieve an equivalent rate of O2 consumption for rate comparison, 10 times more N. viennensis than N. multiformis cells were required in the MR chamber. Whereas N. multiformis showed a linear rate of O2 consumption during ammonia oxidation (Figure 2c), the initial rate of O2 consumption by N. viennensis was quite rapid, followed by a slower, linear rate (Figures 2a and b). N. viennensis produced a maximum of ca. 1.41 nM of NO per 1 × 1010 cells (n = 4) at the beginning of substrate oxidation, concomitant with the initial rapid rate of O2 consumption. The NO was immediately re- consumed as the cells achieved the slower, linear rate of O2 consumption (Figure 2a). After ca. 3 min from the point at which O2 became undetectable, N. viennensis cells began to release NO reaching a maximum of ca. 1.39 nM per 1x1010 cells (n = 4). None of the NO released by N. viennensis follow- ing O2 depletion was re-consumed. In contrast, N. multiformis produced a maximum of ca. 92.15 nM NO per 1 × 1010 cells (n = 4) and re- consumption of NO began once ca. 50% of the available O2 was consumed (Figure 2c).
Levels of N2O were measured during and follow- ing ammonia oxidation; however, no N2O was detectable during ammonia oxidation by cells of either microbe (Figures 2b and d). Assays including N. viennensis cells contained measurable N2O levels increasing at a non-linear rate after ca. 5 min following depletion of O2, yielding an average maximum at ca. 40 min of 0.19 μM per 1 × 1010 cells (Figure 2b; n = 4). In contrast, assays including N. multiformis cells contained measurable N2O levels immediately upon O2 depletion increasing at a linear rate to an average maximum of 5.6 μM per 1 × 1010 cells at ca. 40 min (Figure 2d; n = 4).
Dynamics of NO production and consumption during NH2OH oxidation by N. viennensis EN76T
Although the experiments described above indicated that N. viennensis cultures produce and consume NO during ammonia oxidation to nitrite, it was not clear whether NO acted as an intermediate in the ammonia- or hydroxylamine-oxidizing step of the pathway. Therefore, we examined production and consumption of NO by N. viennensis cells when fed with NH2OH instead of ammonium. NH2OH was added in 200 μM pulses to the MR chamber containing N. viennensis cells to support linear O2 consumption until all of the available O2 was
Figure 1 Inhibition of N. viennensis (a) and N. multiformis (b) with the NO-scavenger PTIO. Ammonium consumption (black squares, dotted line) and nitrite production (white triangles, solid line) as well as N2O production (gray circles, solid line) are plotted. The black arrow indicates the time point of PTIO addition (150 μgml− 1) to the cultures. Mean values of five-fold replicated experiments with standard deviations are shown.
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consumed (Figure 3). Each subsequent addition of equal aliquots of NH2OH led to the production of ca. 5 nM NO per 1 × 1010 cells (n = 3), followed by an immediate re-consumption of NO and O2 until the next addition of NH2OH (Figure 3). NO2− accumulated to ca. 206 μM (n = 3) in the culture medium, which matched the ca. 207 μM O2 consumed during the time course of the experiment. Importantly, free conversion of NH2OH to NO in the absence of cells was stochastic and insignificant (Supplementary Figure S3a). Addi- tion of NH2OH to FWM containing 200 μM NaNO2 resulted in the production of ca. 4 nM NO but only once a concentration of 1.2 mM NH2OH was reached in the MR chamber (Supplementary Figure S3b).
In an effort to demonstrate the requirement of NO by N. viennensis for the oxidation of either NH3 or NH2OH, washed N. viennensis cells were incubated with 200 μM PTIO for 1 h prior to measurement of NH4+- or NH2OH-dependent O2 consumption. The presence of PTIO did not prevent substrate- dependent O2 consumption of either substrate by N. viennensis cells (Supplementary Figure S1), although PTIO was able to effectively scavenge NO in cell-free FWM containing the NO-donating com- pound, PAPA-NONOate (Supplementary Figure S2).
Detection of abiotic N2O in growth media without viable cells Inspired by the observed differences in N2O produc- tion profiles between cultures of N. viennensis and N. multiformis, we performed abiotic experiments in
the MR chamber using cell-free FWM or HKM and the NO-donating compound, MAHMA NONOate. Addition of MAHMA NONOate to FWM released ca. 5.5 μM NO, 70% of which was converted to N2O (Figure 4a). In contrast, addition of an equal aliquot of MAHMA NONOate to HKM resulted in only a 20% conversion of the released NO to N2O (Figure 4c). Continuous addition of MAHMA NONOate to produce 1.1–5.5 μM NO in FWM or HKM resulted in a sustained high-efficiency conver- sion of released NO to N2O only in FWM, but not
Figure 2 Instantaneous measurements of O2 (black line), NO (a and c) and N2O (b and d) (gray dots) after addition of 2 mM NH4Cl in liquid phase suspensions of N. viennensis (a and b) and N. multiformis (c and d) cells. Panels are single representative measurements of reproducible results (n = 4). Note that y-axes for NO are on different scales for N. viennensis versus N. multiformis. N. viennensis cell concentration was 1011 cells per ml, whereas N. multiformis cell concentration was 1010 cells per ml to achieve equivalent rates of O2 consumption by the two strains.
Figure 3 Instantaneous measurements of O2 consumption (black line) and NO production (grey dots) from 200 μM pulses of NH2OH in liquid phase incubations of N. viennensis in the absence of NH4+. Plot is a single representative of replicable experiments (n = 4).
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HKM (Figures 4b and d). Reactivity of NH2OH in FWM+heat-killed N. viennensis cells was also explored (Supplementary Figure S4). When NH2OH was introduced into the MR chamber containing heat-killed cells, accumulation of NO reached ca. 110 nM NO over 5 min (Supplementary Figure S4a). After 30 min, N2O accumulated to levels of ca. 90 μM, demonstrating that NH2OH was eventually converted to N2O in the absence of physiologically active cells (Supplementary Figure S4b).
Discussion
The NO-scavenger, PTIO, stops ammonia-dependent chemolithotrophy of N. viennensis The measured cessation of ammonium consump- tion and nitrite production upon PTIO addition to growing cultures of N. viennensis demonstrates the requirement of free NO for ammonia chemolitho- trophy that was not observed for N. multiformis. These results confirm prior growth experiments with enrichment cultures (Jung et al., 2014) and reported effects on nitrite production and activity by ammonia-oxidizing Thaumarchaea and bacteria incubated with PTIO (Shen et al., 2013; Martens- Habbena et al., 2015). For both N. viennensis and N. multiformis, PTIO addition abolished N2O production, suggesting that the presence of the enzyme-generated free NO intermediate is required for formation of N2O production by both strains, regardless of whether NO is reduced biotically by enzyme activity or abiotically.
NO is produced and immediately consumed during active ammonia oxidation by N. viennensis EN76T
The initial, rapid production of NO followed by its equally rapid consumption during ammonia-dependent O2 consumption by N. viennensis differed from results in similar experiments with N. maritimus SCM1 (Martens-Habbena et al., 2015). In this prior study, N. maritimus SCM1 produced NO at a steady-state level prior to its consumption once NH4+ was depleted or its partial consumption at saturating concentrations of NH4+. A major difference in the two profiles observed for both cultures was that O2 levels remained quite high in assays with N. maritimus SCM1 such that complete consumption of NO was not observed as a function of time and O2 consumption as observed for N. viennensis EN76T. Even so, experiments with both N. viennensis and N. maritimus confirm that NO is being produced and consumed during ammonia oxidation. In addition, the present experiments demonstrate that NO is being released at the onset of anoxia. A likely fate of released NO at anoxia was its conversion to N2O, because 1000 times more N2O than NO was measured once the microrespirometry chamber reached anoxia, suggesting rapid conversion of released NO to N2O (Figures 2a and c). Another contributor to N2O levels measured in anoxic assays with N. viennensis cells could be the reactivity of cell components with released NH2OH as heat-killed cells showed a rapid conversion of exogenous NH2OH to measureable NO and N2O (Supplementary figure S4).
NO dynamics during ammonium-dependent O2 consumption by N. multiformis showed a vastly different profile compared to that of either N. viennensis
Figure 4 Abiotic production of N2O from the NO-donor MAHMA NONOate in either FWM (a and b) or HKM (c and d). Panels are single representative measurements of reproducible results (n = 3). The addition of varying concentrations of MAHMA NONOate is indicated by arrows.
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or N. maritimus, revealing ca. 10 times more NO released from the cells, some of which was slowly re-consumed during ammonia oxidation and through anoxia. The comparison of NOx profiles in microre- spirometry measurements with cells of N. multiformis and the ammonia-oxidizing Thaumarchaea reveals an intriguing difference in how NOx is metabol- ized during ammonia oxidation by bacteria and Thaumarchaea, which requires further investiga- tion. Unlike N. viennensis, N2O production by N. multiformis during anoxia was linear and 10 times more N2O was produced per number of cells. This is a confirmatory evidence that ammonia- oxidizing bacteria, but not N. viennensis, are capable of producing N2O enzymatically via nitri- fier denitrification (Stieglmeier et al., 2014b).
NO is produced and consumed during NH2OH oxidation to NO2− in N. viennensis The rapid production and consumption of NO during NH2OH oxidation by N. viennensis along with the stoichiometric production of NO2− with O2 consump- tion suggest that NO is directly participating in the dehydrogenation of NH2OH. Recent models have postulated that NO is involved in providing reductant to ammonia monooxygenase (Schleper and Nicol, 2010; Stahl and de la Torre, 2012); however, if this were the case then the rapid production/consumption cycle of NO during NH2OH oxidation would not be observed. Experiments to demonstrate the role of NO in NH2OH oxidation by pre-incubating washed cells with PTIO with the goal to observe quenching of NH3 or NH2OH-dependent O2 consumption were inconclusive. It is possible that PTIO is only effective during active growth of N. viennensis (which was observed; Figure 1), or perhaps PTIO was ineffective at chelating rapidly cycling NO at the high cell densities used in the MR chamber.
N2O in N. viennensis cultures originates from the abiotic reaction of biotic N-oxide intermediates with medium or cellular components Previous studies measuring N2O in pure and enrich- ments cultures of ammonia-oxidizing Thaumarchaea suggested an enzymatic origin of measured N2O (Santoro et al., 2011; Loscher et al., 2012; Jung et al., 2014); however, control experiments to test for abiotic reduction of NO to N2O, including by medium components, were not performed. We observed a high rate of NO reduction to N2O in FWM both in the presence of an NO-donating molecule and in the presence of NH2OH plus heat-killed cells. Based on the difference in metal content of both media, we propose that reduction of NO to N2O in FWM is facilitated by iron, which is present in FWM at a relatively high final concentration of 7.5 uMl− 1 in the form of FeNaEDTA but absent from HKM. Under anoxic conditions, in which the metal components of the medium are reduced, the Fe(II) and reduced trace
metals act as chemical catalysts for NO reduction to N2O. This ‘chemodenitrification’ process has been implicated by hypothesis in contributing to abiotic N2O production in reduced environments where Fe(II) is abundant (Samarkin et al., 2010; Kampschreur et al., 2011; Jones et al., 2015).
Proposed pathway for ammonia chemolithotrophy in ammonia-oxidizing Thaumarchaea in which NO facilitates NH2OH oxidation Our revised model of ammonia-dependent chemo- lithotrophy of the Thaumarchaeota places NO as a necessary co-reactant for the oxidation of NH2OH to NO2− (Figure 5a). This NO-dependent dehydro- genation of NH2OH to NO2− is not based on novel chemistry because ammonia-oxidizing bacteria, and others such as aerobic methane-oxidizing bacteria, utilize the heme-containing cytochrome P460 enzyme to facilitate this reaction (Figure 5b; Simon and Klotz, 2013). Instead, the central reaction in the Thaumarchaeotal nitrification pathway is based on a proposed novel copper enzyme capable of per- forming known P460 activity. This model achieves the proper substrate stoichiometry and reductant flow. In addition, the modelled rapid cycling of NO (and, concomitantly electrons) to support NH2OH oxidation would logically preclude any enzymology for NO reduction to N2O. In agreement with this requirement, none of the sequenced genomes of ammonia-oxidizing Thaumarchaeota revealed the presence of canonical and alternate inventory for NO reduction to N2O. Nitrite reductase (nirK) is encoded in the genomes of all published sequences of ammonia-oxidizing Thaumarchaea (Bartossek et al., 2010, 2012) and nirK transcripts have been detected at very high steady-state levels in environmental metatranscriptomes (Hollibaugh et al., 2011; Radax et al., 2012), which makes this enzyme the most parsimonious source of the NO needed to support ammonia-dependent chemolitho- trophy. The proposal that NO2− reduction and not NH2OH oxidation is the more likely source of the NO required for the oxidation of NH2OH to NO2−
is supported by the following logic and reasoning: (1) A two-step oxidation of NH2OH to NO2− via a NO
intermediate would require the operation of two enzyme complexes that feed extracted electrons (3+1) via two redox shuttles to two quinone-reactive enzymes. In addition to requiring additional unknown inventory, such a pathway would also not generate enough electrons needed to provide for effective linear electron flow (4−2− 1=1). In contrast, the proposed one-step model provides for effective linear electron flow (5− 2−1=2; Figure 5). The bioenergetics contrast stands in the context that an observed active NirK activity would draw one electron per reduced NO2− in both scenarios in addition to that the two-step model would include two linearly connected sources of NO production in a genomic background not encoding identifiable NO detoxification inventory and a scenario
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that should not lead to stoichiometric conversion of N-NH3 to N-NO2−.
(2) Isotopic measurements of 15N2O produced by N. viennensis suggest a ‘hybrid signature’ in that one N atom would originate from NH3 (contributed as NH2OH) and one atom would originate from NO2−
(contributed as NO) (Stieglmeier et al., 2014b). This finding also contradicts a two-step model and supports the model shown in Figure 5.
The proposed one-step model is parsimonious in that it requires the innovation of only one enzyme in ammonia-oxidizing Thaumarchaea. Based on exist- ing knowledge, this novel enzyme is copper-based and facilitates known redox chemistry in context with known enzyme complexes such as the NH2OH- producing ammonia monooxygenase, NO-producing NirK, plastocyanin redox carriers and a quinone- reactive membrane protein, all of which are copper proteins and have been identified in all sequenced genomes of ammonia-oxidizing Thaumarchaea (Walker et al., 2010; Bartossek et al., 2012; Stahl and de la Torre, 2012).
We propose that the model of catabolic electron flow presented here (Figure 5a) applies to all obligate chemolithotrophic ammonia-oxidizing Thaumarchaea because it is based on and supported by results from above referenced experiments with marine ammonia- oxidizing Thaumarchaea including N. maritimus SCM1 and experiments with terrestrial ammonia- oxidizing Thaumarchaeota including the data pre- sented here for N. viennensis EN76T.
Conclusion
The present study establishes that both ammonia- oxidizing Thaumarchaea and bacteria contribute to the production of N2O, although the mechanisms by which they do so are distinct. Whereas the ammonia-oxidizing bacteria produce N2O enzymati- cally through nitrifier denitrification, the ammonia- oxidizing Thaumarchaea release intermediates (NO and/or NH2OH), which are then reduced non- enzymatically to N2O in anoxic microenvironments
Figure 5 Proposed pathway for ammonia-dependent chemolithotrophy in the ammonia-oxidizing Thaumarchaea (a) compared with known pathways of N-oxide transformation in ammonia-oxidizing bacteria (b). The model presents a central role of NO in the oxidation of NH2OH, and its contribution to hybrid formation of N2O as proposed by Stieglmeier et al. (2014b). Due to the lack of heme proteins including HAO and quinone-reactive proteins such as cM552 (CycB), redox processes in ammonia-oxidizing archaea are likely mediated by Cu protein complexes (Walker et al., 2010; Stahl and de la Torre, 2012). The present literature suggests that NH3 is monooxygenated to NH2OH by ammonia monooxygenase (AMO) and that NH2OH is dehydrogenated to NO2− by activities of a number of unknown enzymes (Walker et al., 2010; Stahl and de la Torre, 2012; Vajrala et al., 2012). Based on existing chemistry facilitated by heme proteins in ammonia-oxidizing bacteria (b), the model in (a) proposes that the oxidation of NH2OH to NO2− and subsequent extraction of five electrons results from a reaction of NH2OH with NO and H2O facilitated by a novel Cu-containing enzyme. This could be one of the multi-copper oxidases encoded in all genomes of ammonia-oxidizing Thaumarchaeota (Bartossek et al., 2010, 2012; Walker et al., 2010). NO is provided by the Cu-containing NirK, which enzymatically reduces one NO2− per NH3 oxidized to NO. A fraction of the enzyme-produced NO and NH2OH could react to form N2O by hybrid formation. The figure was adapted from Simon and Klotz (2013). AMO/Cu-MMO, ammonia monooxygenase; c552, cytochrome c redox carrier; CytS: cytochrome c’-beta (see Simon and Klotz, 2013, and references therein); HAO, hydroxylamine dehydrogenase; HCO, heme-copper oxidase; HURM, hydroxylamine:ubiquinone redox module (see Simon and Klotz, 2013, and references therein); NirK, Cu-containing NO-forming nitrite reductase; NOR, nitric oxide reductase; P460, tetraheme cytochrome c protein P460 (CytL; see Simon and Klotz, 2013, and references therein); pcy, plastocyanin; pmf, proton-motive force; Q/QH2, quinone/ quinol pool.
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(Zhu-Barker et al., 2015). Due to the relatively high abundance and activity of Thaumarchaea across terrestrial, freshwater, and marine environments (Zhang et al., 2010; Pratscher et al., 2011; French et al., 2012; Berg et al., 2015) and their established tolerance of low ammonium and oxygen environ- ments (Martens-Habbena et al., 2009), their contribu- tions to NOx emissions is likely of high global significance (Babbin et al., 2015). For instance, marine Thaumarchaea may be essential in providing a substantial concentration of NO to denitrifying microorganisms within oxygen minimum zones, and in return, the denitrifiers could provide organic carbon to the Thaumarchaeota to establish a nitrifying- denitrifying consortium (Karner et al., 2001; Beman et al., 2012; Ganesh et al., 2015). The present study also supports that both ammonia-oxidizing Thau- marchaea and bacterial ammonia-oxidizers likely contribute to chemodenitrification in terrestrial envir- onments through the release and subsequent transfor- mation of metabolites (NH2OH, NO and NO2−) either abiotically or via denitrifying consortia (Jones et al., 2015), which dominate in less oligotrophic environ- ments. The elucidation of NO as an essential pathway intermediate and released metabolite of the ammonia- oxidizing Thaumarchaea in the absence of a nitrifier denitrificaiton pathway will allow refinement of the relative contributions of ammonia-oxidizing microor- ganisms to global N2O production.
Conflict of Interest
The authors declare no conflict of interest.
Acknowledgements JK was supported by graduate fellowship funds from Alberta Innovates Technology Futures. LYS was supported by a discovery grant from NSERC (RGPIN-2014-03745). Work in CS laboratory was supported by the Austrian Science Fund grant P25369. MGK was supported by NSF grant MCD1202648.
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Supplementary Information accompanies this paper on The ISME Journal website (http://www.nature.com/ismej)
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- title_link
- Introduction
- Materials and methods
- Strains and cultivation
- Growth experiments with NO-scavenger PTIO
- Instantaneous measurement of NO and N2O during oxidation of ammonia
- Instantaneous measurement of NO from N. viennensis during oxidation of NH2OH
- Instantaneous ammonia and hydroxylamine oxidation by N. viennensis in the presence of PTIO
- Instantaneous measurement of N2O from media and killed-cell controls
- Results
- Effects of the NO-scavenger PTIO on N2O levels measured in cultures of N. viennensis and N. multiformis
- Dynamics of NO and N2O production during and following ammonia oxidation
- Dynamics of NO production and consumption during NH2OH oxidation by N. viennensis EN76T
- Inhibition of N. viennensis (a) and N. multiformis (b) with the NO-scavenger PTIO. Ammonium consumption (black squares, dotted line) and nitrite production (white triangles, solid line) as well as N2O production (gray circles, solid line) are plotted. The
- Detection of abiotic N2O in growth media without viable cells
- Instantaneous measurements of O2 (black line), NO (a and c) and N2O (b and d) (gray dots) after addition of 2&znbsp;mM NH4Cl in liquid phase suspensions of N. viennensis (a and b) and N. multiformis (c and d) cells. Panels are single representative measur
- Instantaneous measurements of O2 consumption (black line) and NO production (grey dots) from 200&znbsp;μM pulses of NH2OH in liquid phase incubations of N. viennensis in the absence of NH4+. Plot is a single representative of replicable experiment
- Discussion
- The NO-scavenger, PTIO, stops ammonia-dependent chemolithotrophy of N. viennensis
- NO is produced and immediately consumed during active ammonia oxidation by N. viennensis EN76T
- Abiotic production of N2O from the NO-donor MAHMA NONOate in either FWM (a and b) or HKM (c and d). Panels are single representative measurements of reproducible results (n�=�3). The addition of varying concentrations of MAHMA NONOate is indicated by arro
- NO is produced and consumed during NH2OH oxidation to NO2− in N. viennensis
- N2O in N. viennensis cultures originates from the abiotic reaction of biotic N-�oxide intermediates with medium or cellular components
- Proposed pathway for ammonia chemolithotrophy in ammonia-oxidizing Thaumarchaea in which NO facilitates NH2OH oxidation
- Conclusion
- Proposed pathway for ammonia-dependent chemolithotrophy in the ammonia-oxidizing Thaumarchaea (a) compared with known pathways of N-�oxide transformation in ammonia-oxidizing bacteria (b). The model presents a central role of NO in the oxidation of NH2OH,
- JK was supported by graduate fellowship funds from Alberta Innovates Technology Futures. LYS was supported by a discovery grant from NSERC (RGPIN�-�2014-03745). Work in CS laboratory was supported by the Austrian Science Fund grant P25369. MGK was support
- ACKNOWLEDGEMENTS