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. 2023 Mar 28;57(14):5655–5665. doi: 10.1021/acs.est.2c09457

Inhibition of Methylmercury and Methane Formation by Nitrous Oxide in Arctic Tundra Soil Microcosms

Lijie Zhang †,‡, Yongchao Yin §,∥,⊥, Yanchen Sun ∥,#, Xujun Liang †, David E Graham §, Eric M Pierce †, Frank E Löffler §,∥,⊥,#,∇,*, Baohua Gu †,∇,*
PMCID: PMC10100821  PMID: 36976621

Abstract

graphic file with name es2c09457_0006.jpg

Climate warming causes permafrost thaw predicted to increase toxic methylmercury (MeHg) and greenhouse gas [i.e., methane (CH4), carbon dioxide (CO2), and nitrous oxide (N2O)] formation. A microcosm incubation study with Arctic tundra soil over 145 days demonstrates that N2O at 0.1 and 1 mM markedly inhibited microbial MeHg formation, methanogenesis, and sulfate reduction, while it slightly promoted CO2 production. Microbial community analyses indicate that N2O decreased the relative abundances of methanogenic archaea and microbial clades implicated in sulfate reduction and MeHg formation. Following depletion of N2O, both MeHg formation and sulfate reduction rapidly resumed, whereas CH4 production remained low, suggesting that N2O affected susceptible microbial guilds differently. MeHg formation strongly coincided with sulfate reduction, supporting prior reports linking sulfate-reducing bacteria to MeHg formation in the Arctic soil. This research highlights complex biogeochemical interactions in governing MeHg and CH4 formation and lays the foundation for future mechanistic studies for improved predictive understanding of MeHg and greenhouse gas fluxes from thawing permafrost ecosystems.

Keywords: nitrous oxide, mercury methylation, methanogenesis, sulfate reduction, greenhouse gases, microbial community response, Arctic ecosystem

Short abstract

Nitrous oxide strongly inhibits methylmercury and methane formation in Arctic soil microcosms.

Introduction

Elevated levels of mercury (Hg), a global pollutant, are observed in Arctic soils due to long-distance transport of gaseous elemental Hg(0) and atmospheric deposition of Hg(II) following Hg(0) oxidation.1−3 It has been estimated that the Northern Hemisphere permafrost regions store 1656 ± 962 Gg Hg, of which 793 ± 461 Gg Hg is trapped in permafrost.3 Climate warming is causing permafrost thaw, which could significantly impact Hg biogeochemical transformations and cycling in the Arctic ecosystem.4−6 Experimental and modeling studies have shown that thawing permafrost drives the release and export of Hg from soils to open water bodies including rivers and lakes.4,5,7 A recent study reported that increasing temperatures could enhance microbial methylation of inorganic Hg(II) to methylmercury (MeHg), a potent neurotoxin, by an order of magnitude in Arctic soils.6 This enhanced microbial MeHg formation is thought to be associated with an increased availability of soil organic carbon (SOC) in thawing permafrost.6 Under anoxic conditions, certain sulfate-reducing bacteria (SRB), methanogens, and iron-reducing bacteria that possess the HgcAB protein complex consisting of a corrinoid prosthetic group and a ferredoxin have the potential to convert inorganic Hg(II) to MeHg.8

Permafrost contains about 50% of the total global SOC, which is a subject of intensive investigations due to its potential climate impact.9−12 Thawing exposes large amounts of previously frozen SOC, and enhanced microbial decomposition substantially increases greenhouse gas (CO2, CH4, N2O) emissions.13,14 It has been estimated that the atmospheric CH4 concentrations have increased by a factor of 2–3 since the 1700s.15 The emission of CH4 is of great concern as it is a powerful greenhouse gas with about 30 times greater warming potential than CO2.16 Additionally, permafrost soils store a large amount of organic nitrogen (N), and its decomposition generates nitrous oxide (N2O), another potent greenhouse gas.17−21 N2O formation is generally attributed to enhanced turnover of organic N producing ammonium and nitrate that, respectively, fuel nitrification and denitrification, the two dominant processes generating N2O in soils.17,20 Elevated N2O concentrations reaching tens to hundreds of micromolar were reported in Arctic soil porewaters.17,18 Since the warming potential of N2O is even greater than that of CH4,16 N2O emission represents another strong positive noncarbon feedback on global warming.

Microcosm, enrichment culture, and pure culture studies have reported inhibitory effects of N2O on methanogenesis.22−25 Two possible mechanisms have been proposed, including (1) the utilization of N2O as an energetically favorable electron acceptor thereby reducing electron flow to produce CH4 and (2) potential cytotoxic effects of N2O,26,27 presumably due to interference with Co(I) corrinoid-dependent enzyme systems required for CH4 formation.28,29 Support for the latter mechanism is provided by studies showing that the suppression of methanogenesis by N2O was not solely caused by changes in redox potential.24,26,27 Metabolic consequences of the toxic effects of N2O have been demonstrated including ceased corrinoid-dependent methionine biosynthesis and impaired organohalide respiration.30−32 Some early studies postulated that N2O suppresses certain methanogenic and sulfate-reducing populations,24,33 which are relevant contributors to MeHg formation. These previous observations raise an important yet unanswered question whether increased N2O formation during permafrost thaw has the potential to impair microbial Hg(II) methylation and methanogenesis in Arctic soils. Improved understanding of these poorly understood interactions could help predict toxic MeHg formation and greenhouse gas emissions from thawing Arctic ecosystems in response to climate warming.

To investigate the impact of N2O on biogeochemical processes leading to MeHg and greenhouse gas (i.e., CO2 and CH4) formation, circumneutral pH Arctic tundra soil was collected from the toe of a hillslope near Nome, Alaska. A series of laboratory soil microcosms with low and high concentrations of N2O were established, and MeHg, CH4, and CO2 formation, sulfate reduction, and changes in microbial community structure were monitored.

Materials and Methods

Soil Samples and Microcosm Incubations

A thawed Arctic active layer soil (10–40 cm below surface) was collected in the August 2018 field campaign from the Next Generation Ecosystem Experiment in the Arctic (NGEE-Arctic) Intensive Site 9 near mile marker 27 on Teller Road in Nome, Alaska. The soil was sealed in sterile Whirl-Pak bags, frozen during shipment to Oak Ridge National Laboratory, Oak Ridge, TN, and then stored at −20 °C until use. Detailed site information, soil collection, and preservation have been described elsewhere,9,34,35 and the basic geochemical properties of the soil are listed in Supporting Information (SI) Table S1.

To set up microcosms, the frozen soil was thawed and homogenized inside an anoxic glove chamber (Coy Laboratory Products, Grass Lake, MI) under ∼98% N2 and 2% H2 atmosphere. The homogenized soil (3.0 ± 0.1 g wet weight, equivalent to 1.18 g dry weight) was transferred into a series of acid-cleaned and autoclaved 26-mL glass serum bottles, which were sealed with butyl rubber stoppers held in place with aluminum crimps. Samples were then flushed with ultrapure N2 for 5 min and supplemented with N2O (99%, Sigma-Aldrich, St. Louis, MO), prefilled in an autoclaved sealed serum bottle. An airtight syringe with a stopcock was used to transfer N2O to microcosms to achieve an initial porewater N2O concentration of 0.1 mM and 1 mM, corresponding to 0.154 and 1.54 mmol of N2O g–1 dry weight (dw) soil, respectively. The soil porewater content was determined by drying pre-weighted homogenized wet soil (∼1 g) in triplicates at 60 °C and then calculating the mass difference between the initial soil and the soil after drying for at least 2 days until a constant mass was obtained. Aqueous phase N2O concentrations were estimated using published Henry’s Law constants36 based on the headspace volume (23 mL) and soil porewater content (Table S1). Microcosms were incubated at 8 °C to simulate the elevated air temperatures observed during the summer months (up to 17 °C in Nome, Alaska) and to accelerate microbial activities and reactions.34,35 N2O concentrations were monitored every 2–3 days for up to 80 days by taking 0.1 to 1 mL of headspace gas samples using an airtight syringe with a stopcock over the course of the incubation period. After each sampling event, N2O was supplemented to maintain the initial target concentrations of either 0.1 or 1 mM to compensate N2O consumption until day 44 (Figure S1). To observe whether the production of CH4 and MeHg could be restored following the depletion of N2O, no N2O was added after day 44. Replicate control microcosms received no N2O.

Chemical Analyses

Triplicate headspace samples from each treatment were taken and analyzed for N2O, CO2, and CH4 following incubation periods of 4, 8, 15, 28, 45, 56, 63, 80, 105, 115, 130, and 145 days. In addition, destructive sampling of soil from triplicate microcosms occurred after 0, 4, 8, 15, 28, 45, 115, 130, and 145 days of incubation to measure MeHg and water-extractable sulfate and nitrate concentrations in soil, except for samples treated with 1 mM N2O at 115 and 130 days due to limited quantities of soil samples. Headspace N2O was analyzed on an Agilent 3000A Micro-gas chromatograph (GC) equipped with a PLOT Q column and a thermal conductivity detector. Headspace CO2 and CH4 were measured with a gas chromatograph equipped with a methanizer and a flame ionization detector (SRI 8610C, SRI Instruments, Torrance, CA).9 Headspace gas concentrations were converted to total production (in μmol) using Henry’s law, as described.36 Multicomparisons of CH4 formation among treatments were tested by analysis of variance (ANOVA) and pairwise comparisons (Bonferroni test) following validation for normality,37 using the OriginPro analysis software (OriginLab, Northampton, MA) with a significance threshold of P = 0.05.

Following headspace gas measurements, microcosms were destructively sampled, as noted above, and the soils were homogenized in an anoxic glove chamber prior to chemical analyses. Total extractable MeHg was analyzed using established methods.6,38 Briefly, an aliquot of wet soil sample (0.35–0.5 g) was mixed with 10 mL of 1.5 M KBr in 5% (v/v) H2SO4 and 2 mL of 1 M CuSO4 in a 50-mL conical plastic centrifuge tube by shaking for 1 h. An isotope labeled Me200Hg internal standard was added to correct for any potential loss during extraction and sample distillation. Then, 10 mL of methylene chloride (99.99%, Fisher Scientific) was added, and samples were shaken for another 1 h to extract MeHg. The extraction was accomplished by collecting the methylene chloride extract using a glass pipet, combining it with 20 mL of Milli-Q water, and then purging with ultrapure N2 at 40 °C to remove methylene chloride. MeHg remaining in the aqueous phase was subsequently analyzed following the modified US EPA Method 1630, via distillation, ethylation, and measurements using an inductively coupled plasma mass spectrometer (ICP-MS, Elan DRC-e, PerkinElmer, Shelton, CT) as previously described.39−41 The ICP-MS was interfaced with automated MERX purge & trap and Hg speciation GC & pyrolysis systems. For quality assurance and quality control, a sediment reference material (ERM-CC580 with a certified MeHg value of 75 ± 4 ng g–1) was extracted and processed in parallel with the microcosm samples. The measured MeHg values in the reference material ranged from 70.2 to 79.5 ng g–1, and the recovery was within 100 ± 10%. Calibration samples with known amounts of ambient MeHg (a reference standard from Brooks Rand Instruments, Seattle, WA) and Me200Hg were also analyzed every 20 samples with a mean recovery of 100 ± 3.5%, as previously described.42,43 The detection limit of MeHg by ICP-MS measurement was about 1.5 pg.43,44

To determine water-extractable organic carbon (WEOC) and anions, ∼0.5 g of wet soil was extracted with 10 mL of Milli-Q water for 1.5 h on a rotary shaker. The soil suspension was centrifuged at 3000g, and the supernatant was filtered through a 0.2 μm PTFE membrane syringe filter (Pall Corporation, Port Washington, NY). Anions, including SO42– and NO3–, as well as organic acids were analyzed on a Dionex Integrion ion chromatograph (IC, Thermo Scientific) using established methods.45,46 WEOC was analyzed on a total carbon analyzer (TOC-L, Shimadzu, Japan) following sample acidification in 0.1% HCl and purging to remove any dissolved inorganic carbonates.

DNA Extraction, 16S rRNA Gene Amplicon Sequencing, and Microbial Community Composition

16S rRNA gene amplicon sequencing was performed following a 45-day incubation period. Genomic DNA was extracted from triplicate 0.2 g wet soil samples collected from the microcosms using the DNeasy PowerSoil Kit (Qiagen, Germantown, MD) following the manufacturer’s instructions. The 16S rRNA genes were amplified with primers F515/R806 targeting the V4 region.47 All samples were barcoded and processed at the University of Tennessee, Knoxville, Genomics Core Facility following established Illumina MiSeq procedures.48 The raw read sequence data were deposited to the NCBI Sequence Read Archive database under the accession number SRP385269.

Raw sequence data were analyzed using a QIIME 2 (v2021.4)49 pipeline employing Cutadapt to remove primers50 and DADA2 to denoise and generate amplicon sequence variants (ASVs).51 Resulting ASV sequences were taxonomically classified using the QIIME 2 naïve Bayes classifier, which was trained on the Silva v138 database52 and pre-trimmed with the F515/R806 primers. To minimize the bias caused by variation in sequencing depth and allow for comparison of sequencing results among samples, the sequences were normalized by minimum sample sequence numbers. Microbial community analyses, including α and β diversity indexes, and Unconstrained Principal Coordinates Analysis (PCoA) were performed and plotted in R (version 4.0.2). The sequence abundance analysis of microbial clades implicated in MeHg formation was based on the identified microbial taxa containing hgcA sequences from available genomic and metagenomic sequences.53 Groupwise Shannon and Chao1 diversity comparisons among treatments were tested by ANOVA in R (version 4.0.2), and pairwise abundance comparisons were performed by Student’s t test following validation for normality, using Microsoft Excel with a significance threshold of P = 0.05.

Results

N2O Inhibits MeHg Formation

The Arctic soil microcosm studies reveal that MeHg formation approximately doubled from an initial level of 0.42 ± 0.05 ng g–1 dry weight (dw) to 0.82 ± 0.05 ng g–1 dw on day 8, independent of the addition of N2O (Figure 1A). After 8 days, MeHg in the no-N2O control incubations continued to increase up to 2.23 ± 0.36 ng g–1 dw on day 45. In contrast, MeHg in soil samples treated with 0.1 and 1 mM N2O remained essentially unchanged and much lower than that in the control incubations lacking N2O (Figure 1A). The results suggest that following an initial MeHg production phase, the presence of N2O strongly inhibited the formation of MeHg in the soil microcosms. No significant differences in MeHg levels were observed between soil samples treated with 0.1 or 1 mM N2O over the 45-day incubation period, suggesting that the lower N2O concentration was sufficient to halt microbial methylation of Hg(II).

Figure 1.

Figure 1

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Changes of (A) methylmercury (MeHg) and (B) water-extractable sulfate in anoxic Arctic tundra soil microcosms incubated with and without N2O at 8 °C. Replicate microcosms received no N2O (control, black squares), 0.1 mM N2O (orange triangles), and 1 mM N2O (green circles). Shown are data averages from triplicate microcosms with error bars representing one standard deviation. The light gray shaded areas indicate the incubation periods when N2O was added every 2–3 days to restore the initial N2O levels, and no N2O was supplemented after day 44 (light blue shaded areas, see Figure S1). The arrow in (A) indicates when N2O was depleted and no longer detected. The detection limit for sulfate was 0.14 μmol g–1 dw (dashed line).

Following the cessation of N2O additions after day 44, MeHg formation in the low-N2O (0.1 mM) incubations was rapidly restored (Figure 1A). MeHg in the low-N2O incubations increased from 1.04 ± 0.14 ng g–1 dw at day 45 to 2.89 ± 0.05 ng g–1 dw at day 115, compared to from 2.27 ± 0.36 to 3.41 ± 0.08 ng g–1 dw in the control incubations without N2O during the same period. Similar increases in MeHg production were observed in the low-N2O and no-N2O control incubations from day 115 to day 145. In contrast, Hg(II) methylation did not recover in the high-N2O (1 mM) microcosms throughout the incubation period (Figure 1A). N2O concentrations decreased to ∼0.30 mM after 80 days in the high-N2O incubations (SI Figure S1). In the low-N2O microcosms, N2O was consumed within about 5 days following the addition of N2O (Figure S1) and no N2O was detected after 50 days of incubation (i.e., 6 days after the last N2O addition). These results indicate that N2O could strongly inhibit microbial Hg(II) methylation, but the inhibition was reversible and MeHg formation resumed once N2O was depleted.

N2O Impacts CO2 and CH4 Production

The cumulative CO2 production in microcosms that received 0.1 or 1 mM N2O was similar and reached 25.32 ± 2.58 μmol g–1 dw after 63 days of incubation (Figure 2A). About 30% lower CO2 production (18.80 ± 2.72 μmol g–1 dw) was observed in control incubations without N2O over the same incubation period (Figure 2A). The CO2 concentration in the high-N2O (1 mM) microcosms continued to be higher than that in the no-N2O control incubations and reached up to 40.49 ± 4.18 μmol g–1 dw at day 115, whereas no further increase was observed in the low-N2O (0.1 mM) microcosms whereby N2O was depleted after 50 days (Figure S1). Following 115 days of incubation, similar CO2 concentrations were observed in the low-N2O and the control microcosms without N2O (Figure 2A). These results suggest that N2O had a stimulatory effect on CO2 production.

Figure 2.

Figure 2

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Effects of N2O on CO2 (A) and CH4 (B) production in anoxic Arctic tundra soil microcosms incubated at 8 °C. Replicate microcosms received no N2O (control, black squares), 0.1 mM N2O (orange triangles), and 1 mM N2O (green circles). Data represent averages from triplicate incubations with error bars showing one standard deviation. The inset in (B) shows the cumulative CH4 production during the first 15 days, and significant differences between N2O-treated and control groups at each timepoint are marked as (*) (P < 0.05). The graphs are shaded to indicate the periods when N2O was added every 2–3 days to restore the initial N2O levels (gray shaded areas) and no N2O additions after day 44 (blue shaded areas).

Contrary to CO2 production, immediate and substantial inhibition of CH4 formation was observed in microcosms that received N2O (Figure 2B, inset). On day 4 (the first sampling time point), CH4 formation was 0.18 ± 0.01 μmol g–1 dw in the control microcosms without N2O but was only 0.09 ± 0.02 and 0.04 ± 0.001 μmol g–1 dw in the low-N2O and high-N2O incubations, respectively. CH4 formation in N2O-treated incubations remained low or nondetectable but increased continuously in control microcosms reaching up to 3.77 ± 0.24 μmol g–1 dw on day 63 and 20.35 ± 1.88 μmol g–1 dw on day 145. In the low-N2O microcosms, small amounts of CH4 were observed after 63 days, indicating the onset of methanogenesis following the complete consumption of N2O after 50 days (Figure S1). The CH4 concentration increased to 1.70 ± 0.39 μmol g–1 dw on day 115 and to 3.74 ± 0.51 μmol g–1 dw on day 145, although this amount was ∼5 times lower than what was observed in the control incubations (Figure 2B). In the high-N2O microcosms, CH4 formation was negligible throughout the 145-day incubation period. In these microcosms, N2O was not depleted, and concentrations exceeded 0.3 mM following the cessation of N2O addition on day 44 (Figure S1), indicating that N2O inhibited methanogenesis throughout the incubation period.

Effects of N2O on Sulfate and Nitrate Concentrations

At the beginning of the incubation (i.e., time zero), water-extractable sulfate was 0.53 ± 0.04 μmol g–1 dw (Figure 1B). By day 4, sulfate had been consumed to below the detection limit of ∼0.14 μmol g–1 dw in all microcosms. However, after 8 days sulfate in the microcosms amended with N2O increased, while sulfate remained below the detection limit in the control microcosms without N2O. In the high-N2O (1 mM) microcosms, the amount of sulfate increased to 1.14 ± 0.13 μmol g–1 dw on day 145, compared to 0.53 ± 0.04 μmol g–1 dw in the initial soil (Figure 1B). Sulfate increases were also observed in the low-N2O (0.1 mM) microcosms until day 45, followed by a decline to below the detection limit, which coincided with the depletion of N2O (i.e., after N2O additions stopped). Apparently, sulfate reduction activity in the low-N2O microcosms was restored (Figure 1B) when all N2O had been consumed (Figure S1A). Sulfate reduction was not apparent in the high-N2O microcosms (Figure 1B), as N2O remained at concentrations above 0.3 mM (Figure S1B). These results illustrate that N2O also inhibited sulfate reduction or increased sulfate formation. The inhibition of sulfate reduction was reversible upon the depletion of N2O, as was observed in the low-N2O microcosms (Figure 1B) and, in particular, phases of sulfate consumption coincided with MeHg formation (Figure 1A). In all microcosms, the amounts of nitrate remained close to or below the detection limit of 0.2 μmol g–1 dw over the entire incubation period (Figure S2), as most nitrate would have been denitrified or assimilated in these anoxic tundra soils.54,55

Microbial Community Responses to N2O

The microbial communities in the soils incubated without and with N2O addition for 45 days were subjected to 16S rRNA gene amplicon sequence analysis. The comparison of the microbial community profiles revealed moderate variations at the phylum level among treatments (Figure 3A), with a slight relative abundance increase of Proteobacteria (Pseudomonadota) and a decrease of Desulfobacterota in microcosms amended with N2O (Figure 3A). Members of the phylum Proteobacteria may utilize N2O as respiratory electron acceptor, consistent with N2O consumption in the microcosms (Figure S1). The phylum Desulfobacterota comprises SRB implicated in MeHg formation,53 and a relative abundance decrease in microcosms amended with N2O is consistent with the lack of sulfate consumption and MeHg formation (Figure 1).

Figure 3.

Figure 3

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Microbial community composition in the Arctic tundra soil microcosms after incubation without N2O (control), and under low-N2O (0.1 mM) and high-N2O (1 mM) conditions for 45 days. (A) Changes in relative phylum abundances in soil microcosms without N2O, and under low-N2O and high-N2O conditions. (B) Unconstrained principal coordinates analysis (PCoA) of microbial β diversity using an unweighted UniFrac metric with the percentages of variations explained by axis 1 and axis 2. Comparison of the α diversity with Shannon (C) and Chao1 (D) indexes, representing the richness and evenness, respectively, of the microbial community based on one-way ANOVA (*P < 0.05).

β diversity analyses by PCoA based on Bray–Curtis distance matrix of the relative abundances of ASVs demonstrated pronounced changes in microbial community structure in response to N2O, with the highest groupwise distance observed in the high-N2O versus no-N2O control treatments (Figure 3B). Furthermore, the α diversity measured by the Shannon and Chao1 indices significantly decreased (P < 0.05, Table S2) in both the low- and high-N2O microcosms compared to the control incubations without N2O (Figure 3C,D). No significant differences in α diversity were observed between the low- and high-N2O treatments, suggesting that 0.1 mM N2O was sufficient to trigger the observed microbial community responses. These analyses illustrate the impact of N2O on the microbial community structure, which reflects process-level changes, including the inhibition of MeHg formation, methanogenesis, and sulfate reduction in the Arctic tundra soil microcosms.

Additional pairwise comparative analyses show changes in sequence abundances, such as decreased relative abundances of Geobacteraceae sequences at the family level (Figure 4A), and uncultured Geobacteraceae and Geobacter sequences at the genus level (Figure 4B) in low-N2O and high-N2O microcosms compared to controls without N2O. These microbial taxa have been implicated in sulfate and ferric iron reduction and MeHg formation.53 Consistent with the phenotypic observations, the sum of the relative abundances of SRB and methanogen sequences significantly decreased in the low-N2O and high-N2O microcosms (P < 0.05). Thus, the 16S rRNA gene amplicon sequencing results are consistent with measured activities in the microcosms showing that Hg(II) methylation, sulfate reduction, and methanogenesis were inhibited by N2O (Figures 1 and 2).

Figure 4.

Figure 4

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Changes in relative abundances of 16S rRNA gene amplicon sequences representing microbial clades implicated in MeHg formation. Triplicate Arctic tundra soil microcosms were incubated without N2O, with low N2O (0.1 mM), and high N2O (1 mM) for 45 days. (A) Overall decrease of taxa implicated in MeHg formation at the family level in response to N2O exposure. The category “Others” comprises sequences representing SRB and methanogens with abundances <0.001%. (B) Relative abundance changes (indicated by the size of the circles) of specific taxa at the genus level including taxa without cultured representatives. The Silva database lists the improper name “Citrifermentans”, which was replaced with the validly described genus name Geomonas.56

Discussion

The Arctic soil microcosm study demonstrated that micromolar concentrations of N2O completely shut down MeHg formation, although the inhibition was not immediate but apparent after a lag phase of ∼8 days (Figure 1). MeHg formation coincided with sulfate reduction occurring during the first ∼8 days of incubation and both processes resumed in the low-N2O microcosms after depletion of N2O at ∼50 days. These observations corroborate a linkage between the processes of sulfate reduction and Hg(II) methylation, indicating that SRB played a role in Hg(II) methylation in the Arctic Tundra soil microcosms. The result is consistent with prior studies that established SRB as the dominant players in Hg(II) methylation in Arctic soils and freshwater sediments when sulfate is available.57,58 Therefore, the lack of MeHg formation in microcosms with N2O may be attributed to the inhibition of SRB, although the data cannot rule out the involvement of other microbial guilds [e.g., Fe(III)-reducing bacteria, methanogens].57,59 Fe(III) reduction was reported in these Arctic tundra soils,9 and the observation of abundant Fe(III)-reducing bacteria (e.g., Geobacter) (Figure 4B) suggests Fe(III) reduction activity in the microcosms.

N2O showed pronounced and immediate inhibition on CH4 production in the Arctic soil microcosms (Figure 2B), consistent with previous studies reporting that N2O impacts methanogenesis.22,23 Unlike Hg(II) methylation and sulfate reduction, CH4 production in microcosms with low N2O was not fully restored following the depletion of N2O, even over a prolonged 145-day incubation period. These results suggest that methanogens were minor contributors to Hg(II) methylation, again supporting that the bulk of the observed MeHg formation was due to SRB activity.

Varying responses of different microbes to N2O have been demonstrated,22,24,26,33 which could explain the initial formation of MeHg (Figure 1A) but immediate inhibition of CH4 production (Figure 2B, inset) in the soil microcosms amended with N2O. Varied susceptibility of SRB and methanogens to N2O inhibition may also explain why Hg(II) methylation and sulfate reduction recovered quickly (Figure 1), but methanogenesis did not (Figure 2B), upon depletion of N2O after 50 days. We speculate that one of the causes of the immediate inhibition of CH4 production by N2O could be due to direct enzymatic inhibition, as described in previous studies.26,32 N2O has a very strong impact on microbes that utilize corrinoid-dependent proteins in their energy metabolism, such as methanogens and organohalide-respiring bacteria.32,60 N2O can chemically react and oxidize the super-reduced Co(I) corrinoid prosthetic group and thus impair enzyme activity and metabolic functions of these organismal groups.28−31 Methanogens have no other means for conserving energy and strictly depend on corrinoid-dependent enzyme systems,61,62 so the impact of N2O on methanogenesis is strong and immediate. Organisms that use corrinoid-dependent proteins in anabolic pathways often have alternate means of acquiring essential metabolites, so there is inhibition, but at least some organisms, including many SRB, may have a workaround. Therefore, an immediate impact of N2O on MeHg formation was not apparent; however, this observation does not preclude inactivation of the Hg(II) methylation enzymes by N2O. The functional HgcAB protein complex hinges on a super-reduced Co(I) to catalyze Hg(II) methylation,8,63−65 but this process is not linked to energy conservation or anabolism, a possible explanation for the delayed inhibition by N2O observed in the microcosm experiments. Additional studies would be required to generate detailed understanding of the modes and biochemistry of N2O inhibition on corrinoid-dependent proteins and explain the variable responses of proteins to N2O.

Although direct evidence for these interactions is currently lacking, the inhibition of N2O on microbial processes that hinge on Co(I)-corrinoid-dependent enzyme systems is not unprecedented and has been observed in microbial systems, such as methionine synthase (MetH)-dependent methionine biosynthesis and bacterial organohalide respiration, a process that utilizes corrinoid-dependent reductive dehalogenases.32,66,67 All organisms require methionine, but MetH (corrinoid-dependent methionine synthase) and MetE (corrinoid-independent methionine synthase) have been found in bacteria and archaea for methionine biosynthesis. Some bacteria and archaea have developed mechanisms to compensate for the loss of function of MetH by upregulating MetE and cobalamin B12 scavenging pathways to compensate the inhibitory effect of N2O, whereas others strictly depend on functional MetH to synthesize methionine.68 Consistently, we found that N2O treatment selectively impacted the microbial community (Figures 3 and 4) and decreased the abundance of SRB and methanogens, presumably due to the inhibition of corrinoid-dependent enzyme systems involved in CH4 and MeHg formation. Our analyses show that the microbial community response to N2O generally correlated with the phenotypic observations [i.e., inhibited Hg(II) methylation, sulfate reduction, and methanogenesis]. For example, many known Hg(II) methylators belonging to the Desulfobacterota(69) and methanogenic archaea53,70 declined in relative abundances following N2O exposure. The decreases in relative abundances of methanogenic archaea and SRB suggest growth inhibition of these microorganisms by N2O.

Another potential cause of N2O inhibition may be attributed to N2O-reducers, which compete with methanogenic archaea and SRB for SOC-derived electron donors,25 thereby impeding the growth of these microbes and suppressing CH4 and MeHg formation. However, the Arctic soil used for microcosms setup has a high SOC content, and the WEOC content remained essentially unchanged from ∼38.6 μmol g–1 dw before incubation (Table S1) to 35–45 μmol g–1 dw in all the treatments over a 145-day incubation period. Sulfate reduction and Hg(II) methylation resumed when N2O was depleted in the low-N2O incubations, indicating sufficient electron donor was available. These findings suggest that electron donor-limiting conditions were not established in the microcosms during the incubation periods, and electron donor scarcity cannot explain the decreased CH4 and MeHg formation activity in the presence of N2O. Although N2O reduction is energetically more favorable than sulfate reduction, electron acceptor utilization is not solely determined by thermodynamics. In static microcosm incubations, the rates at which competing redox processes are co-occurring are not immediately controlled by thermodynamics but enzymatic electron transfer reactions to the respective electron acceptors.71 Further complicating matters are inhibitory interactions, such as the formation of sulfide from sulfate reduction, which precipitates Cu2+, a required co-factor for the assembly of functional N2O reductase.72,73 Future studies are again warranted to provide detailed mechanistic insights about how N2O affects electron flow through different redox processes.

Microcosms amended with N2O produced slightly more CO2 than control incubations without N2O, likely due to the utilization of N2O as a respiratory electron acceptor by bacteria possessing the nos operon.72−74 Thus, N2O may have promoted SOC mineralization (i.e., CO2 formation) in microcosms amended with N2O. In addition, less conversion of CO2 to CH4 is expected to occur due to the inhibitory effects of N2O on methanogenesis.75

Environmental Implications

Our findings illustrate that N2O strongly inhibits MeHg and CH4 production in Arctic permafrost, which is considered a critical zone environment for CH4, CO2, and N2O emissions and MeHg formation under climate warming scenarios. The observed inhibitory effects of N2O have implications for predicting neurotoxic MeHg formation and greenhouse gas emissions in Arctic ecosystems. MeHg formation is a great concern as Hg(II) typically enters food webs after conversion to MeHg, which bioaccumulates and biomagnifies along food chains.76 Elevated Hg concentrations have been reported in fish and marine mammals that are used as traditional food sources by indigenous populations in the Arctic, resulting in high levels of Hg in their blood.77,78 Therefore, understanding MeHg formation in permafrost soils is critical to assess health and environmental risks in the Arctic. Permafrost harbors fixed nitrogen in frozen SOC, and climate warming and permafrost thawing will increase nitrogen turnover and the production of N2O.17,18 N2O is generated by various biotic and abiotic processes under different geochemical conditions.79 N2O can distribute with the flow of water and reach geochemical zones where N2O formation is not prevalent, including zones of sulfate and/or CO2 reduction. Soil is highly heterogeneous, and microenvironments with different geochemistries can exist in proximity without strict spatial and/or temporal separation, and N2O can reach pockets dominated by sulfate reduction or methanogenesis. Therefore, warmer temperatures and permafrost thaw can promote MeHg formation,6 but the increased generation of N2O in thawing permafrost can inhibit, and potentially limit MeHg formation, as demonstrated in this study. The new information on MeHg production illustrates the complicated interplay of processes that ultimately determine how climate change will influence Hg biogeochemical transformation and the associated bioaccumulation and trophic transfer of MeHg. Better understanding of these processes can improve predictions of permafrost carbon and noncarbon feedbacks on the fate and transport of Hg in the Arctic ecosystem, although long-term field measurements and monitoring regimes will be needed.

The results of this study also have important implications for other soil ecosystems, as elevated N2O concentrations reaching 0.14 mM have been observed in watersheds impacted by agricultural activities.80 The effect of N2O on the formation of highly toxic MeHg has been overlooked so far. Due to increased use of synthetic nitrogen fertilizers in agricultural production and subsequent microbial nitrification/denitrification and/or chemodenitrification processes, agricultural lands are among the largest contributors to the global anthropogenic N2O budget. N2O is released to the atmosphere either through direct emission from the soil or indirect emission from contiguous aquatic compartments with nitrogen imported from agricultural lands by runoff and leaching processes.80,81 Previous studies have shown that nitrate suppressed MeHg production in paddy soils and in a Hg-contaminated lake, and the mechanism has been attributed to the suppression of SRB activities by nitrate.82−84 Our finding that N2O is a potent inhibitor of MeHg production can offer an alternative explanation as N2O is produced during nitrate turnover. We therefore suggest that future studies account for elevated N2O concentrations in soils, sediments, and water, and a new model is developed for improved prediction of the fate, transformation, and bioaccumulation of Hg in food webs.

Acknowledgments

The authors thank Xiangping Yin for technical assistance in MeHg analyses. This research was sponsored in part by the Office of Biological and Environmental Research within the Office of Science of the U.S. Department of Energy (DOE), as part of the Critical Interfaces Science Focus Area and the Next Generation Ecosystem Experiments (NGEE-Arctic) projects at the Oak Ridge National Laboratory (ORNL). The Department of Energy will provide public access to these results of federally sponsored research in accordance with the DOE Public Access Plan (http://energy.gov/downloads/doe-public-access-plan). ORNL is managed by UT-Battelle, LLC under Contract No. DE-AC05-00OR22725 with DOE. F.E.L. acknowledges funding through the Dimensions of Biodiversity program of the US National Science Foundation (award 1831599). Y.Y. acknowledges a graduate student fellowship from the China Scholarship Council. L.Z. acknowledges support from the start-up fund from the Department of Chemistry and Environmental Science at the New Jersey Institute of Technology (NJIT).

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.est.2c09457.

  • Changes in N2O concentrations during anoxic incubation of the Arctic soil (Figure S1); changes in the amounts of water-extractable nitrate in the Arctic tundra soil microcosms (Figure S2); basic geochemical properties of the original Arctic soil used in this study (Table S1); and Chao1 and Shannon indexes of tundra soil microbial community (Table S2) (PDF)

Author Present Address

◆ Department of Biology, Antimicrobial Discovery Center, Northeastern University, Boston, Massachusetts 02115, United States

Author Contributions

○ L.Z. and Y.Y. contribute equally.

The authors declare no competing financial interest.

Supplementary Material

es2c09457_si_001.pdf (235KB, pdf)

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