Carbon dioxide (CO 2 ) receives primary attention as a driver for climate change, but methane (CH 4 ) and nitrous oxide (N 2 O) account for ∼20% and 7%, respectively, of the net radiative forcing in the atmosphere [1,2]. The central objective of the Paris Agreement, which is to hold the global average temperature increase to "well below 2 • C above preindustrial levels" [3,4], cannot be met without controlling CH 4 and N 2 O emissions. Global emissions of both CH 4 and N 2 O are ultimately controlled by microbial processes [2,5]; however, human activities have massively disturbed the natural balance of microbial production and consumption of these greenhouse gases [6]. The current estimated annual net atmospheric emission increases of ∼51 Tg of CH 4 [7,8] and 2.2 Tg of N 2 O [9] are both predicted to accelerate [1,10]. The key microbial guilds and biogeochemical processes responsible for CH 4 and N 2 O production and consumption are known, and their responses to climate change have been a matter of intense research [11,12]. One area of considerable uncertainty pertains to positive and negative feedback loops affecting greenhouse gas emissions and climate [13]. An infamous scenario for a positive feedback loop in response to a warming climate is permafrost thawing, which stimulates the microbial activity and the release of CO 2 , CH 4 , and N 2 O from newly bioavailable carbon and nitrogen pools. The massive release of greenhouse gases will further accelerate radiative forcing and in turn cause more thawing. This example illustrates why a comprehensive understanding of both positive and negative feedback loops is essential. Quantitative information is needed to meaningfully incorporate feedback effects into refined climate models.
Methanogenic archaea (methanogens) drive CH 4 production by utilizing acetate, H 2 /CO 2 , and methylated compounds as substrates, generating around 1 billion metric tons of CH 4 annually [14,15]. Approximately, two-thirds of biogenic CH 4 is produced from acetoclastic methanogenesis (i.e. the conversion of acetate to CH 4 and CO 2 ), with the remaining one-third attributed to hydrogenotrophic CO 2 reduction (hydrogenotrophic pathway) and methylated compound utilization (methylotrophic pathways) [16][17][18][19][20]. While the three major methanogenesis pathways share a core set of enzymes, several mechanistically distinct enzyme systems with corrinoid prosthetic groups (i.e. vitamin B 12 derivatives) are involved in methyl group transfer reactions, energy conservation, and CH 4 production ( Fig. 1) [18,19,21].
In hydrogenotrophic methanogenesis, CO 2 is sequentially reduced to a methyl group carried by tetrahydromethanopterin (H 4 MPT) with H 2 as electron donor. The corrinoid-dependent N 5 methyltetrahydromethanopterin:CoM methyltransferase (MTR) complex then transfers the methyl group onto another C 1 -carrier, coenzyme M, forming CH 3 -S-CoM, a step associated with energy conservation, followed by CH 4 production catalyzed by methyl coenzyme M reductase (MCR) [22]. Acetoclastic methanogens produce CH 4 by activating acetate to acetyl-CoA, which is then cleaved by the corrinoid-dependent enzyme system carbon monoxide dehydrogenase/acetyl-CoA synthase (CODH/ACS) to yield an enzyme-bound methyl group and a carbonyl group. The carbonyl moiety is oxidized to CO 2 and the methyl group is transferred to H 4 MPT [23]. The MTR complex catalyzes a methyl group transfer to form CH 3 -S-CoM, a step associated with energy conservation, before MCR mediates the reduction of the methyl group to CH 4 [16,24]. Methylotrophic methanogens either directly generate CH 3 -S-CoM from methylated compounds (e.g. MeOH) utilizing substrate-specific, corrinoid-dependent MTRs, or cleave the methyl group from methoxylated compounds (e.g. 2methoxybenzoate) and form CH 3 -CoM via H 4 MPT and the MTR complex [25], followed by CH 4 production via MCR. All three methanogenesis pathways utilize corrinoid-dependent enzyme systems for methyl group transfers and have strict requirements for super-reduced Co(I) to generate CH 4 and conserve energy [26,27]. These features render methanogenesis sensitive to oxidative stress (e.g. f luctuating redox conditions and oxygen intrusion) [19,28].
N 2 O is an even stronger oxidant than oxygen (E o '(N 2 O(g)/N 2 (g) = +1.355 V > E o '(O 2 (g)/H 2 O(l) = +0.818 V), reacts with Co(I), and has been shown to interfere with Co(I) cobamide-dependent enzyme systems [29,30]. Demonstrated metabolic consequences include ceased corrinoid-dependent methionine biosynthesis and impaired organohalide respiration [31][32][33]. Based on this information, elevated N 2 O in anoxic ecosystems would be expected to inhibit other corrinoid-dependent processes such as methanogenesis; however, available studies investigating the impacts of N 2 O on methanogenesis have led to inconsistent N 2 O inhibition patterns for axenic methanogen cultures and CH 4 producing microbial communities [34][35][36]. Laboratory incubations showed that Methanobacterium bryantii strain Bab1 grown with H 2 /CO 2 ceased CH 4 production in the presence of 95 µM N 2 O, whereas Methanosarcina barkeri strain MS maintained some methanogenic activity at 10-fold higher N 2 O concentrations under the same growth conditions [35]. Different sensitivities to N 2 O inhibition were also reported for mixed methanogenic cultures maintained with different substrates. For example, inhibition of CH 4 production in a mixed community bioreactor occurred only at N 2 O concentrations exceeding 700 µM [36], whereas 20-28 µM N 2 O completely inhibited methanogenic activity in salt marsh sediment and Amazon peatland enrichment cultures [34,37]. These variable sensitivities to N 2 O suggest that inhibition of methanogenesis by N 2 O is organism-and possibly substrate-specific; however, the available data are scarce and do not allow a robust, quantitative assessment of N 2 O inhibition on methanogenesis from relevant methanogenic substrates [34][35][36].
N 2 O fluxes in soil-water systems have risen sharply due to the intensified use of synthetic N fertilizer in agriculture [6,9,38]. As a result, elevated N 2 O concentrations occur more frequently in ecosystems with CH 4 production such as rice paddy soils [39], wastewater treatment plants [40], sediments [41], and groundwater aquifers [42]. Also, permafrost thawing accelerates N turnover, releasing large amounts of N 2 O [38,43]. More detailed knowledge about the interactions between N 2 O concentrations and methanogenesis is needed to advance the predictive capabilities of climate change impacts on future greenhouse gas emission scenarios. To address the existing knowledge gaps, we assessed the inhibitory effect of N 2 O on CH 4 production from major methanogenic substrates in growth experiments with axenic and mixed methanogenic cultures and in wholecell suspension assays. Using cultures performing acetoclastic, methylotrophic, and/or hydrogenotrophic methanogenesis, we determined kinetic parameters that quantitatively describe N 2 O inhibition on methanogenesis.
The methanogenic archaeon M. barkeri strain Fusaro metabolizes acetate, H 2 /CO 2 , and MeOH while employing different, substratespecific methanogenic pathways. To determine if M. barkeri exhibits varied sensitivities to N 2 O, cultures were pregrown with MeOH, H 2 /CO 2 , or acetate for at least three consecutive transfers before the impact of N 2 O on CH 4 production was examined. Also, we analyzed three methanogenic mixed cultures derived from anaerobic digester sludge, which is known to harbor a broad diversity of methanogenic archaea. Three different enrichment cultures from the same source material were obtained with MeOH, H 2 /CO 2 , or acetate as growth substrate. The mixed cultures were transferred at least six times on the respective substrate and were used to examine the impact of N 2 O on CH 4 production from different methanogenic substrates (see Supplemental Information for additional information on the mixed methanogenic cultures). Experiments were performed in triplicate 60-ml glass serum bottles with 30 ml N 2 /CO 2 (80/20, v/v) headspace and 30 ml bicarbonate-buffered (50 mM) mineral salt medium (pH 7.2) reduced with 0.2 mM sulfide and 0.2 mM Lcysteine [44]. Acetoclastic, methylotrophic, and hydrogenotrophic cultures received 20 mM acetate, 30 mM MeOH, and 1.24 mmol H 2 , respectively. To avoid overpressure in bottles with H 2 as electron donor, the headspace of culture bottles was replaced with 30 ml of filter-sterilized H 2 . For M. barkeri cultures, 0.1-1.0 ml of N 2 O gas (undiluted or 10-fold diluted in N 2 ) was directly added to the incubation vessels to achieve final aqueous N 2 O concentrations of 100 and 200 µM in cultures with MeOH, 50 and 100 µM in cultures with H 2 , and 20 and 50 µM in cultures with acetate. For the methanogenic mixed cultures grown with 30 mM MeOH, 30 ml H 2 (1.24 mmol), and 20 mM acetate, 0.5-1.6 ml of 10-fold diluted N 2 O gas (in N 2 ) was introduced to achieve final aqueous phase N 2 O concentrations of 10 and 30 µM. More detailed information about N 2 O additions and concentration calculations is provided in the Supplemental Methods. All cultures were incubated without agitation at 37 • C in the dark with the stoppers facing up, and replicates without N 2 O and without inoculum served as positive and negative controls, respectively. CH 4 and N 2 O were analyzed throughout the growth experiments by injecting 100 µl headspace samples into an Agilent 3000A Micro gas chromatograph equipped with thermal conductivity detectors and a molecular sieve column and a PLOT Q column for CH 4 and N 2 O measurements, respectively.
The M. barkeri and the methanogenic mixed cultures were first grown in 1.6 l medium and harvested via centrifugation when about two-thirds of the initial substrate (i.e. MeOH, H 2 /CO 2 , or acetate) had been converted to CH 4 , as calculated based on CH 4 production according to Equations ( 1)-(3) (see below). The cell pellets collected from 1.6 l of medium were washed and suspended in 1.6 ml of reduced mineral salt medium in sealed 2-ml glass vials, resulting in a 1000-fold concentration of the biomass. A 0.2 ml aliquot of the concentrated cell suspension was sacrificed to measure total protein with the Bradford assay [45].
Cell suspension assays were performed at room temperature in 20-ml glass vials f lushed with N 2 /CO 2 (80/20, v/v) and were sealed with Tef lon-lined butyl rubber stoppers held in place with aluminum crimps. The assay vials received a total of 0.9 ml reduced mineral salt medium, 0.1 ml of cell suspension, and increasing concentrations of substrates (i.e. MeOH, H 2, or acetate, with N 2 O as indicated in Tables S1-S7). For assay vials receiving H 2 as electron donor, the headspace was replaced with increasing volumes of premixed H 2 /CO 2 (4/1, v/v) to achieve H 2 concentrations ranging from 1.2 to 333 µM (Table S3). Small volumes (89-178 µl) of undiluted or 10-fold diluted (in N 2 ) N 2 O were directly injected into the 20-ml assay vials, what resulted in small pressure changes with negligible impact on the distribution of N 2 O between the aqueous phase and the headspace. All cell suspensions were freshly prepared following identical procedures to ensure consistency between independent experiments. Vials that received 0.1 ml of sterile mineral salt medium or 0.1 ml of heat-killed (i.e. autoclaved) cell suspension served as negative controls.
Following 10 min of equilibration, 0.1 ml of cell suspension was added to initiate the assays. Headspace samples (100 µl) were withdrawn with an air-tight syringe with a lock every 30 min over a 3-h incubation period and, for the kinetic experiments, CH 4 was analyzed using an Agilent 7890 GC series gas chromatograph equipped with a f lame ionization detector and a DB-624 capillary column (60 m length × 0.32 mm diameter, 1.8 µm film thickness). For each treatment at a fixed initial substrate concentration [S], an initial CH 4 production rate v, normalized to the amount of protein per vial in the unit of nmol CH 4 min -1 mg protein -1 , was determined. The determined rate data were fit into Michaelis-Menten competitive, noncompetitive, and uncompetitive inhibition models (Table S8) to determine the maximum CH 4 production rate V max , the half-velocity constant K m , and the inhibitory constant K I of N 2 O on CH 4 production from the different substrates. The best-fit inhibition model was chosen based on the highest coefficient of determination (R 2 ) and the lowest standard deviation of the residuals (Table S9). Each datum point on the Michaelis-Menten plots (Figure 5) represents a CH 4 production rate generated from at least four time points for one substrate concentration [S].
DNA extraction and PCR assays followed established procedures [46] and details are provided in the Supplemental Information. The purified DNA samples were processed and barcoded with primers 341F/785R targeting the V3/V4 region of the prokaryotic 16S rRNA gene [47] following established procedures [48]. The resulting sequence data were analyzed using the QIIME 2 v2021.4 environment [49]. The precise programs and settings are described in the Supplemental Information, and the QIIME 2 pipeline script and custom R file employed to parse results are available at https://github.com/rwmurdoch/methanogens_ and_N2O. The raw amplicon library reads were deposited in the Sequence Read Archive (SRA) under accessions SRR19782291 to SRR19782296.
Quantitative real-time PCR (qPCR) to enumerate archaeal 16S rRNA genes in M. barkeri and methanogenic mixed cultures followed established protocols using primer set Mtgen835F/918R and probe FAM-Mtgen831 (Table S10) [50]. Samples for enumeration of cell numbers were collected at the beginning and at the end of the prolonged growth experiments. Average growth yields of methanogens were calculated from the changes in 16S rRNA gene copy numbers in triplicate culture vessels divided by the total amounts of CH 4 produced over the same time period (Table S11). Reported growth yield (i.e. cells produced per µmol of CH 4 formed) used conversion factors of 3 and 2.5 16S rRNA gene copies per methanogen cell for M. barkeri [51] and the enrichment cultures [52], respectively. For comparison with theoretical [53] and reported values from the literature (Table 2), growth yields were also calculated as µg of dry biomass per µmol of CH 4 formed Downloaded from https://academic.oup.com/ismej/article/18/1/wrae027/7623399 by guest on 24 June 2024 making the following assumptions: an average methanogen cell has a volume of 2.5 µm 3 [54] with a density equal to water [55]. The dry cell biomass is 30% of the wet cell biomass [56], and 90% of the dry biomass represents organic material [55].
In the absence of N 2 O, MeOH-grown M. barkeri cultures consumed 895 ± 10 µmol of MeOH within a 6-day incubation period and produced 635 ± 34 µmol of CH 4 (Fig. 2A). This stoichiometry closely matched the expected CH 4 production based on Equation (1):
Growth yields of 5.3 × 10 6 ± 0.3 × 10 6 cells per µmol CH 4 were measured for cultures without N 2 O (Fig. 2B). By contrast, cultures that received 100 or 200 µM N 2 O produced negligible amounts of CH 4 during the initial 6-day incubation period without any apparent growth after 6 days. Following an 11day lag phase, cultures with 100 µM N 2 O started consuming MeOH, and 626 ± 20 µmol of CH 4 were produced following a 38-day incubation period, indicating that 100 µM N 2 O delayed, but did not prevent, CH 4 production from MeOH by M. barkeri (Fig. 2A). Although complete conversion of MeOH to CH 4 according to Equation (1) was achieved in the presence of 100 µM N 2 O over a prolonged 38-day incubation period, the growth yield decreased by 63.8% ± 7.8% compared to cultures without N 2 O, (i.e. 1.9 × 10 6 ± 0.4 x 10 6 vs. 5.3 × 10 6 ± 0.3 × 10 6 cells were produced per µmol of CH 4 formed) (Fig. 2B). In the presence of 200 µM N 2 O, only 55 ± 11 µmol of CH 4 were produced over a 38-day incubation period, and the growth yield decreased by over 80% to 0.9 × 10 6 ± 0.4 × 10 6 cells per µmol CH 4 , indicating a pronounced inhibitory effect of N 2 O on CH 4 production and growth of M. barkeri.
More pronounced N 2 O inhibition on CH 4 production and growth was observed in M. barkeri cultures that received H 2 as electron donor (i.e. hydrogenotrophic methanogenesis). In the absence of N 2 O, M. barkeri cultures, that received H 2 as electron donor, produced 318 ± 21 µmol of CH 4 from 1.24 mmol of H 2 over an 11-day incubation period, consistent with Equation (2):
In the absence of N 2 O, a growth yield of 0.8 × 10 6 ± 0.2 × 10 6 cells per µmol of CH 4 formed was measured (Fig. 2C and D). In the presence of 50 or 100 µM N 2 O, CH 4 production by M. barkeri cultures commenced after prolonged lag phases ranging from 13 to 17 days. At the end of the 38-day incubation period, M. barkeri cultures with 50 µM N 2 O and produced 311 ± 7 µmol of CH 4 . The M. barkeri cultures that had received 100 µM N 2 O only generated 17 ± 3.2 µmol of CH 4 , resulting in a 95% decrease in total CH 4 production compared to cultures without N 2 O. The average growth yield in cultures with 50 or 100 µM N 2 O decreased by ∼70% and 85% to 2.3 × 10 5 ± 0.3 × 10 5 and 1.2 × 10 5 ± 0.3 × 10 4 cells per µmol CH 4 formed, respectively, compared to cultures without N 2 O. The most pronounced N 2 O inhibition was observed in acetatefed M. barkeri cultures, and 20 µM N 2 O severely diminished CH 4 production (Fig. 2E). In the absence of N 2 O, M. barkeri produced 588 ± 24 µmol of CH 4 from 605 ± 17 µmol of acetate over a 38-day incubation period, consistent with Equation (3):
In the presence of 20 or 50 µM N 2 O, M. barkeri cultures only produced 21.3 ± 4.1 and 20.4 ± 3.8 µmol of CH 4 , respectively, a decline of over 96% in total CH 4 production compared to cultures without N 2 O over the 38-day incubation period. Acetate-grown M. barkeri cultures that had received 20 or 50 µM N 2 O generated 1.5 ± 0.2 × 10 6 and 1.0 ± 0.3 × 10 6 cells per µmol of CH 4 produced, decreases of 30.4% ± 3.3% and 52.5% ± 12.6%, respectively, compared to the average yield of 2.1 × 10 6 ± 0.7 × 10 6 cells per µmol CH 4 in cultures without N 2 O (Fig. 2F).
In all culture vessels with observed inhibition of methanogenic activity, N 2 O concentrations remained constant throughout the experiment, consistent with the absence of a "nos" operon on the genome of M. barkeri. Taken together, these results demonstrate that micromolar levels of N 2 O inhibit methanogenesis, reduce growth yields of this model methanogen, and further reveal that the inhibition is most pronounced for acetoclastic methanogenesis.
To examine the impact of N 2 O on mixed methanogen communities, growth and kinetic assays were performed with enrichment cultures derived from digester sludge. Microbial community analysis revealed the presence of diverse methanogen groups known to utilize acetate, H 2 /CO 2 , and MeOH as substrates, with distinct methanogen taxa prevalent under the different enrichment conditions (Tables S12 and S13). In cultures that received H 2 as electron donor, sequences representing the genus Methanobacterium and other unidentified members of the family Methanobacteriaceae dominated, whereas in acetate-fed cultures, Methanosarcina was the most abundant archaeal taxon. Methanomethylovorans was the most abundant archaeal genus in the MeOH-fed cultures, but sequences representing the genus Methanomassiliicoccus were also prevalent. 16S rRNA gene amplicons representing six families and four of the known eight orders of methanogens were represented in the examined mixed cultures derived from digester sludge (Fig. 3A and B; Table S12).
Without N 2 O addition, the mixed cultures produced 542.6 ± 23.3, 316.5 ± 25.4, and 589.2 ± 33.7 µmol of CH 4 from 0.9 mmol of MeOH, 1.24 mmol of H 2 , or 0.6 mmol of acetate, consistent with Equations ( 1)-(3). When cultures were amended with 10 or 30 µM N 2 O, CH 4 production in MeOH-, H 2 -, and acetate-fed methanogenic mixed cultures was substantially or completely inhibited (Fig. 4A, C, and E). Even over an extended 42-day incubation period, the presence 10 µM N 2 O still repressed the total CH 4 production in MeOH-, H 2 -, or acetate-grown mixed cultures by ∼60%, 80% and 50%, respectively, compared to incubations without N 2 O. With 30 µM N 2 O, only negligible amounts of CH 4 were detected in all incubations over a 42-day incubation period. Some N 2 O loss was observed in the mixed culture vessels that received H 2 /CO 2 or MeOH as substrates, with no more than 20% of the initial amount of N 2 O consumed. Taken together, these results demonstrate that N 2 O exerts a stronger inhibitory effect on methanogenesis in the mixed cultures harboring diverse methanogen populations than in axenic M. barkeri incubations.
Enumeration of total methanogens and total bacteria in the mixed cultures using qPCR revealed that N 2 O diminished methanogen growth yields (Fig. 4B, D, and F). The qPCR analysis further revealed significantly decreased ratios of methanogen-tobacterial 16S rRNA genes in all N 2 O-treated cultures (Fig. 3C-E), illustrating that N 2 O impacted the methanogen populations much more strongly than the bacterial populations. In the absence of N 2 O, the average growth yields of methanogens in the enrichment cultures with MeOH, H 2 , or acetate were 0.6 × 10 7 ± 0.1 × 10 7 , 0.6 × 10 5 ± 0.3 × 10 5 , and 1.0 × 10 6 ± 0.1 × 10 6 cells per µmol of CH 4 formed, respectively. In the presence of 10 µM N 2 O, the growth yields of methanogens declined by ∼90%, 60%, and 50% in enrichment cultures that received MeOH, H 2 , and acetate, respectively (Fig. 4B, D, and F). Only negligible CH 4 production and methanogen growth were measured in all mixed cultures with 30 µM N 2 O (Fig. 4B, D, and F). Taken together, the results of the mixed culture studies corroborate that N 2 O concentrations in the low micromolar range exhibit pronounced inhibitory effects on hydrogenotrophic, methylotrophic, and acetoclastic methanogenesis in microbial communities.
Whole-cell suspension assays using M. barkeri and the methanogenic mixed cultures were performed to quantitatively assess the inhibitory effects of N 2 O on CH 4 production from each methanogenic substrate (Table S1). The Michaelis-Menten single-substrate single-inhibitor model (R 2 > 0.95) best explained the trends of CH 4 production rates versus increasing substrate concentrations (Fig. 5). Among all assays with M. barkeri and the mixed cultures, maximum CH 4 production rates (i.e. V max values) of acetatefed cultures were most strongly affected by increasing N 2 O concentrations, followed by the H 2 -and MeOH-fed cultures (Figs 2 and 3).
In the absence of N 2 O, the V max values for CH 4 production in MeOH-, H 2 -, or acetate-amended M. barkeri cell suspension assays were 440.4 ± 10.2, 138.1 ± 6.9, and 40.8 ± 4.7 nmol CH 4 min -1 mg protein -1 , respectively (Fig. 5A, Table 1). The addition of N 2 O decreased the V max of CH 4 production in MeOH-, H 2 -, and acetate-fed M. barkeri cell suspension assays to different extents. Rate data determined in M. barkeri suspensions assays with MeOH fit the Michaelis-Menten inhibition model best. The V max values declined by ∼45% and 57% to 244.1 ± 38.4 and 188.8 ± 32.5 nmol CH 4 min -1 mg protein -1 , respectively, in the presence of 100 and 200 µM N 2 O. The determined inhibitory constant, K I , of N 2 O on methylotrophic CH 4 production was 130.9 ± 4.7 µM in M. barkeri cell suspensions (Table 1), indicating that N 2 O concentrations around 130 µM reduced the maximum CH 4 production rate (V max ) by 50%. More pronounced N 2 O inhibition was observed in M. barkeri whole-cell suspensions assays using H 2 as the electron donor for CO 2 reduction (Fig. 5B). The addition of 50 and 100 µM Downloaded from https://academic.oup.com/ismej/article/18/1/wrae027/7623399 by guest on 24 June 2024 N 2 O reduced the V max values in H 2 -fed M. barkeri cell suspension assays by ∼40% and 57% to 82.8 ± 29.5 and 59.9 ± 5.9 nmol of CH 4 min -1 mg protein -1 , respectively. The model simulation determined a K I value of 90.6 ± 10.8 µM N 2 O (Fig. 5B, Table 1), indicating a stronger inhibition of N 2 O on CH 4 production in H 2 -versus MeOH-amended M. barkeri cell suspension assays. The M. barkeri assays with acetate as substrate showed the most pronounced inhibition by N 2 O (Fig. 5C, Table 1). In assays amended with 20 and 40 µM N 2 O, the V max values decreased by 21% and 44% to 32.1 ± 4.2 and 22.8 ± 2.5 nmol of CH 4 min -1 mg protein -1 , respectively. From the best-fit inhibition model, a K I value of 24.8 ± 2.6 µM was determined for N 2 O inhibition of acetoclastic methanogenesis in M. barkeri cell suspensions. Collectively, the cell suspension assays corroborate strong inhibitory effects of N 2 O on methanogenesis, with the kinetics of acetoclastic methanogenesis being most impacted by N 2 O. Similar kinetic responses were observed for whole-cell suspension assays conducted with the methanogenic mixed cultures pregrown with the respective substrates (Fig. 5D-F). In the absence of N 2 O, the determined V max values for mixed culture cell suspension assays that received MeOH, H 2 , or acetate were 209.0 ± 4.4, 105.8 ± 3.7, and 22.8 ± 0.7 nmol CH 4 min -1 mg protein -1 , respectively (Table 1). Notably, the methanogenic mixed cultures were more sensitive to N 2 O than axenic M. barkeri cultures irrespective of the type of methanogenic substrate provided. In cell suspension assays that received MeOH, the presence of 50 and 100 µM N 2 O reduced the V max values by 37.4 ± 10.8% and 75.4 ± 21.4%, respectively, compared to assays without N 2 O. The inhibition model determined a K I value of 109.9 ± 6.8 µM for N 2 O inhibition on CH 4 production in cell suspension assays amended with MeOH (Fig. 5D, Table 1). In H 2amended cell suspension assays, the presence of 30 and 60 µM N 2 O decreased the V max values by 32.3 ± 9.8 and 65.4 ± 12.9%, respectively, compared to assays without N 2 O (Fig. 5E, Table 1). The best-fit inhibition model determined a K I value for N 2 O inhibition of 62.1 ± 6.4 µM for CH 4 production in assays that received H 2 as electron donor. Consistent with the observations made with M. barkeri, the most pronounced N 2 O inhibition on CH 4 production rates was observed in mixed culture cell suspension assays that received acetate as substrate (Fig. 5F). In Downloaded from https://academic.oup.com/ismej/article/18/1/wrae027/7623399 by guest on 24 June 2024 the presence of 10 and 30 µM N 2 O, the V max values in suspension assays with acetate decreased by 38 ± 7.8% and 74.9 ± 16.0%, respectively, compared to assays without N 2 O. Using the best-fit Michaelis-Menten inhibition model, a K I value of 17.7 ± 1.8 µM was determined for N 2 O inhibition of CH 4 production in cell suspension assays with acetate (Table 1).
Taken together, the experimental data demonstrate that dissolved
Microorganisms drive global C and N cycling and ultimately control CH 4 and N 2 O production, consumption, and thus emissions to the atmosphere [10,11,13]. Predictive climate models must consider the responses of microbial CH 4 and N 2 O production under environmental change scenarios [10,57]. Quantitative assessment of feedbacks that affect CH 4 production are crucial for refining greenhouse gas emission models. This study investigated the feedback effects between two important greenhouse gases, specifically the inhibitory effect of N 2 O on archaeal CH 4 production, to provide quantitative data that link environmental N 2 O concentrations with methanogenesis. The findings demonstrate that environmentally relevant, micromolar levels of N 2 O suppress CH 4 production and the growth of methanogens. The determined K I values reveal a concentration-dependent, progressively negative feedback by N 2 O on archaeal CH 4 formation rates and the total amounts of CH 4 produced and show that the strength of the inhibition is most pronounced for acetoclastic methanogenesis.
Assuming the current day partial pressure of 335 ppb N 2 O in the atmosphere, the theoretical concentration of N 2 O in air-equilibrated water should be around 7 nM; however, substantially elevated levels of dissolved N 2 O have been observed in groundwater and watersheds [33,42]. In areas impacted by agricultural activities (e.g. fertilizer application), dissolved groundwater N 2 O concentrations can exceed 100 µM [42]. Even in some remote, natural aquatic systems, such as ice-covered Antarctic lakes, N 2 O concentrations of up to 86 µM have been reported [58,59]. N 2 O is generated during N cycling and major formation processes include microbial denitrification, ammonia oxidation, and abiotic chemodenitrification [44]. Based on the physiology of the microorganisms (e.g. ammonia oxidizers are strict aerobes) and the thermodynamics of the processes (e.g. nitrate and nitrite reduction are associated with a greater change in Gibbs free energy than methanogenesis), one might argue that N 2 O formation and methanogenesis are physically separate processes, thus limiting the exposure and inhibitory effects of N 2 O on methanogens. Such redox stratification does occur; however, most environmental matrices, such as soils, are highly heterogenous and characterized by dynamic spatial and temporal gradients resulting in patchy distribution of redox processes. N 2 O is water-soluble and, depending on hydrology, can reach other redox zones [ 60]. Consequently, impacts of elevated N 2 O on various biogeochemical processes, including those associated with greenhouse gas emissions, are likely. Laboratory studies have reported inhibitory effects of nitrogen oxides (NOx), including N 2 O, on methanogenesis [34-37, 61, 62]; however, the available data are scarce and no uniform pattern has emerged that would support a quantitative relationship between N 2 O and microbial CH 4 production. Consequently, this negative feedback of N 2 O on CH 4 production has not been considered in greenhouse gas emission models.
The evaluation of N 2 O inhibition on CH 4 production from methanogenic substrates (i.e. MeOH, acetate, and H 2 /CO 2 ) with both the model methanogen M. barkeri and digester sludgederived mixed methanogenic cultures quantitatively links N 2 O concentrations with methanogen activity and growth. The growth experiments illustrate that micromolar N 2 O concentrations affect CH 4 production, and the whole-cell suspension assays and kinetic model simulations provide a plausible explanation for the inconsistent literature reports about N 2 O effects on methanogenesis. Specifically, acetoclastic methanogenesis was most sensitive to N 2 O (K I values of 18-25 µM), followed by hydrogenotrophic (K I 60-90 µM) and methylotrophic methanogenesis (K I 110-130 µM), indicating that the type of methanogenic substrate utilized affects the sensitivity of methanogens to N 2 O. N 2 O inhibition was significantly more pronounced in methanogenic enrichment cultures than in axenic M. barkeri cultures, and 30 µM N 2 O prevented CH 4 production and methanogen growth in the mixed culture experiments regardless of the type of methanogenic substrate utilized. Previous studies support that mixed methanogenic communities are more sensitive to N 2 O than commonly studied model methanogen isolates [34,35,62,63]. These observations suggest that the axenic methanogen cultures used to elucidate the biochemistry and genetics of methanogenesis may not serve as general models for other features of methanogen biology (e.g. N 2 O inhibition). The reasons for the reduced sensitivity of axenic versus mixed methanogen cultures to N 2 O may be a result of long-term adaptation of the isolates to laboratory cultivation, or not-yet-characterized microbe-microbe interaction networks render mixed cultures more susceptible to N 2 O inhibition. The enrichment cultures used to determine the K I values for N 2 O inhibition harbored diverse methanogen groups (Fig. 3A), and kinetic studies (e.g. determination of K I values for N 2 O inhibition) with representative isolates of the various lineages are warranted to capture the breadth of methanogen responses to N 2 O.
Taken together, the low µM range K I values of N 2 O that impact methanogenesis suggest major consequences of rising N 2 O concentrations for C cycling. Of note, the vast majority of the annual ∼1 billion metric tons of biogenic CH 4 is generated from acetateand H 2 -driven CO 2 reduction [16][17][18][19][20]64], the two processes with the lowest observed K I values for N 2 O inhibition. It is therefore reasonable to predict that elevated environmental N 2 O will impact CH 4 production and methanogen growth in ecosystems with high bioavailable C and N loads, such as wetlands, sediments, and permafrost soils.
Key enzyme systems involved in CH 4 production and energy conservation require cobamide prosthetic groups [65] (Fig. 1). The super-reduced Co(I) form of cobamides is susceptible to 1). The acetoclastic pathway involves two steps catalyzed by corrinoid-dependent enzyme systems, CODH/ACS and the MTR corrinoid enzyme complex [16,65]. (Fig. 1), both of which are targets for N 2 O inhibition. Distinct cobamide-dependent enzyme systems catalyze the formation of CH 3 -CoM in the hydrogenotrophic and the methylotrophic pathways [22,66]. Both the acetoclastic and hydrogenotrophic pathways depend on the MTR corrinoid enzyme complex to generate CH 3 -CoM and to conserve energy [22,27]. By contrast, methylotrophic methanogens can directly generate CH 3 -CoM without the energy-conserving MTR corrinoid enzyme complex from methylated compounds (e.g. MeOH) via a substrate-specific, corrinoid-dependent methyltransferase complex (i.e. MtaA, MtaB, and MtaC in the case of MeOH) [67]. The experimental efforts consistently demonstrated that CH 4 production from acetate and H 2 /CO 2 exhibited 7-and 3-fold higher sensitivities, respectively, to N 2 O than CH 4 production from MeOH. These findings suggest that N 2 O inhibition of methanogenesis is related to the oxidation of the super-reduced Co(I) cobamide, which is essential for the corrinoid enzyme complexes involved in the different methanogenesis pathways. While the differential susceptibilities of pathwayspecific corrinoid-dependent enzymatic steps can explain the distinct inhibitory effects of N 2 O on CH 4 production via the acetoclastic, hydrogenotrophic, and methylotrophic pathways, detailed enzymatic studies would be needed to assess the responses of individual enzyme systems to N 2 O. Kinetic data for the enzymatic regeneration of the super-reduced Co(I) state are lacking, and the rates of reduction may differ between enzymes and pathways, which could contribute to the observed physiological response of decreased CH 4 production. Recovery from N 2 O inhibition was not a focus, but the experimental data indicate partial recovery of methane formation in N 2 O-treated axenic and mixed cultures; however, the methanogen growth yields remained lower in N 2 O-treated cultures compared to controls without N 2 O even over the extended incubation period. Further studies are needed to generate mechanistic insights into modes of recovery from N 2 O inhibition. The presence of N 2 O significantly decreased growth yields or completely abolished growth in the examined methanogenic cultures (Table 2). The type of methanogenic substrate utilized determines the fraction of electrons available from electron donor oxidation directed toward cell synthesis and thus governs the growth yields of methanogens [53,68]. N 2 O affects corrinoiddependent enzymes involved in electron transfer (e.g. the MTR enzyme complex), and it is not surprising that N 2 O interferes with energy conservation in methanogens. Consistently, enumeration of methanogen 16S rRNA genes at the termination of all growth experiments illustrated that N 2 O not only negatively impacted CH 4 production but also methanogen growth yields. An alternate explanation for the reduced methanogen growth yields in the mixed cultures exposed to N 2 O could be competition for electron donor (e.g. H 2 ); however, the sequencing and the qPCR data do not support this hypothesis, and N 2 O inhibition explains the decline of methanogens and the changes of methanogento-bacteria ratios. The measured methanogen growth yield data in the absence of N 2 O were on par with reported experimental data and closely matched the theoretical values (i.e. yields calculated based on thermodynamics) (Table 2). One exception were the growth yields measured in H 2 /CO 2 -fed M. barkeri cultures, which were ∼10-fold lower than data reported in the literature. The most pronounced growth suppression was observed in the mixed methanogenic cultures, where 30 µM N 2 O was sufficient to prevent the growth of methanogenic archaea. Collectively, the data show that micromolar concentrations of N 2 O decrease or abolish CH 4 production, reduce methanogen growth yields, and exhibit progressively negative feedback on microbial CH 4 production. yield (µg organic matter per µmol CH 4 formed) Cultures Substrates N 2 O (µM) Measured a Predicted from thermodynamics b Range (references) M. barkeri MeOH 0 3.57 ± 0.32 5.2 3.3-6.4 ([71, 72]) MeOH 100 1.30 ± 0.28 MeOH 200 0.62 ± 0.03 H 2 0 0.53 ± 0.14 3.6 5.4-8.6 ([73]) H 2 50 0.15 ± 0.02 H 2 100 0.08 ± 0.02 Acetate 0 1.45 ± 0.46 2.1 1.4-5.7 ([74]) Acetate 20 1.02 ± 0.16 Acetate 50 0.70 ± 0.02 Mixed methanogenic cultures MeOH 0 4.30 ± 0.56 5.2 3.3-6.4 ([72]) MeOH 10 0.14 ± 0.05 MeOH 30 NA H 2 0 0.43 ± 0.05 3.6 1.3-7.2 ([14]) H 2 10 0.16 ± 0.01 H 2 30 NA Acetate 0 0.65 ± 0.05 2.1 1.4-5.7 ([23, 64, 75]) Acetate 10 0.31 ± 0.02 Acetate 30 NA
a Cell numbers were determined with qPCR. Because M. barkeri has three copies of the 16S rRNA gene, the qPCR results were divided by a factor of three to obtain cell numbers. Calculation of methanogen cell numbers in the mixed cultures assumed an average 16S rRNA gene content of 2.5. b Theoretical values calculated in this study based on thermodynamics and published information [53 ].
Microbial processes are strongly inf luenced by environmental factors and their responses to climate change vary both spatially and temporally [ 11,69]. To improve the predictive power of climate models and potentially justify the application of biotechnological approaches for managing greenhouse gas emissions, interactions and feedbacks between relevant biotic/abiotic processes must be understood and quantitatively captured [70]. Attempts have been made to include microbial data (i.e. biomass, enzyme, and growth kinetics) to improve climate models [70], but the incorporation of multifactorial, multidirectional, and often nonlinear biotic/abiotic feedbacks underlying the global CH 4 and N 2 O budgets is challenging and requires robust quantitative data. The determined K I values for N 2 O inhibition of methanogenesis reveal a relevant negative feedback effect on CH 4 emissions, and the new quantitative information generates opportunities to refine CH 4 emission models.