Simultaneous sulfide and methane oxidation by an extremophile

Nature Communications volume 14, Article number: 2974 (2023) Cite this article

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Hydrogen sulfide (H2S) and methane (CH4) are produced in anoxic environments through sulfate reduction and organic matter decomposition. Both gases diffuse upwards into oxic zones where aerobic methanotrophs mitigate CH4 emissions by oxidizing this potent greenhouse gas. Although methanotrophs in myriad environments encounter toxic H2S, it is virtually unknown how they are affected. Here, through extensive chemostat culturing we show that a single microorganism can oxidize CH4 and H2S simultaneously at equally high rates. By oxidizing H2S to elemental sulfur, the thermoacidophilic methanotroph Methylacidiphilum fumariolicum SolV alleviates the inhibitory effects of H2S on methanotrophy. Strain SolV adapts to increasing H2S by expressing a sulfide-insensitive ba3-type terminal oxidase and grows as chemolithoautotroph using H2S as sole energy source. Genomic surveys revealed putative sulfide-oxidizing enzymes in numerous methanotrophs, suggesting that H2S oxidation is much more widespread in methanotrophs than previously assumed, enabling them to connect carbon and sulfur cycles in novel ways.

Hydrogen sulfide (H2S) is the most reduced form of sulfur (S) and a potent energy and sulfur source, toxicant, and signaling molecule1,2,3. It is a weak acid that easily diffuses through membranes and inhibits various processes such as aerobic respiration by binding to cytochrome c oxidases. In addition, other metabolic processes that use copper- and iron-containing enzymes are severely inhibited by H2S1,4,5,6. Hence, microorganisms living in sulfide-rich environments require adequate mechanisms to detoxify H2S7,8. In a myriad of environments, such as wetlands, marine sediments, soil, wastewater treatment plants, lakes, paddy fields, landfills, and acidic geothermal environments, H2S is produced through sulfate (SO42−) reduction, mineralization of organic matter, and thermochemistry8,9,10,11,12,13,14,15,16,17,18.

Upon depletion of sulfate, organic matter is ultimately converted to methane (CH4) in oxygen-depleted ecosystems9,12,13,19,20,21. When both H2S and CH4 diffuse into the overlaying oxic zones, CH4 can be utilized as an energy source by aerobic methane-oxidizing bacteria, which are assumed to mitigate most emissions of this potent greenhouse gas22. Despite this effective methane biofilter, 548 to 736 Tg of CH4 is annually released into the atmosphere from various natural and anthropogenic sources23,24. Aerobic methanotrophs are part of various bacterial classes and families, including the ubiquitous Alpha- and Gammaproteobacteria16,25,26, Actinobacteria27 and the extremophilic Methylacidiphilaceae of the phylum Verrucomicrobia28,29,30,31. The latter are acidophilic bacteria that share a low pH optimum (2.0 − 3.5) and live between 35 and 60 °C26,31,32. All known verrucomicrobial methanotrophs have been isolated from geothermal habitats such as fumaroles and mudpots, from which large amounts of mostly thermogenic CH4 and H2S are emitted16,28,33,34,35. Geothermal environments are typically characterized by high H2S emissions and thus the verrucomicrobial methanotrophs isolated from these ecosystems are preeminent examples to study how methanotrophs are affected by H2S.

It is becoming increasingly clear that methanotrophs are metabolically versatile and able to use environmentally relevant energy sources such as H2, propane, ethane, acetate, acetone, 2-propanol, and acetol16,36,37,38. The ability to utilize various energy sources is highly beneficial in environments with heavily fluctuating gas emissions. Recently, it was demonstrated that pure cultures of the verrucomicrobial methanotroph Methylacidiphilum fumariolicum SolV can consume methanethiol (CH3SH), with the concomitant sub-stoichiometric formation of H2S, indicating that strain SolV partially metabolized toxic H2S39. Hereafter, an elegant study demonstrated that also proteobacterial methanotrophs can oxidize H2S40. The authors isolated the versatile alphaproteobacterium ‘Methylovirgula thiovorans’ strain HY1 from a South Korean peatland that could grow on thiosulfate (S2O32−), tetrathionate (S4O62−), elemental sulfur (S0), and a range of carbon compounds. However, strain HY1 cells grown on CH4 as sole energy source were not able to oxidize H2S, and H2S oxidation was only initiated and observed in cells grown in the presence of thiosulfate. In addition, growth on H2S was not studied. Considering recent observations, it is paramount to investigate whether microbes exist that can oxidize the environmentally relevant gases CH4 and H2S simultaneously, how methanotrophs cope with H2S and whether such methanotrophs can conserve energy and produce biomass using H2S as an energy source.

Here, through extensive chemostat cultivation, we show for the first time that a microorganism can oxidize CH4 and H2S simultaneously. M. fumariolicum SolV is inhibited by the presence of elevated H2S concentrations but H2S is rapidly oxidized to elemental sulfur (S0) as a detoxification mechanism to alleviate the inhibitory effect of H2S on CH4 oxidation. Strain SolV adapts to H2S with the upregulation of a Type III sulfide:quinone oxidoreductase (SQR) and an H2S-insensitive ba3-type cytochrome c oxidase, creating an electron transfer pathway from H2S to O2. Additionally, strain SolV incorporates 13CO2 using H2S as sole energy source. We propose that the H2S oxidation capacity of verrucomicrobial methanotrophs is essential to thrive in sulfur-rich acidic geothermal ecosystems. In addition, we found SQR in a plethora of proteobacterial methanotrophs of various environments. Considering CH4 and H2S coexist in a myriad of oxygen-limited ecosystems, H2S oxidation could be a trait present among many aerobic methanotrophs.

The detection of genes encoding putative sulfide:quinone oxidoreductases (SQRs) in the genomes of verrucomicrobial methanotrophs prompted us to investigate whether methanotrophs can oxidize and adapt to H2S16. Accordingly, a continuous culture of the thermoacidophilic aerobic methanotroph Methylacidiphilum fumariolicum SolV (running as chemostat at a dilution rate (D) of 0.016 h−1) was maintained with CH4 as energy source and CO2 as carbon source (non-adapted cells; Fig. 1a), up to a load of 39 μmol CH4 min−1 · g DW−1 (Table 1). For comparison, a distinct continuous cultivation system was designed (with identical CH4 load) to adapt cells to increasing loads of H2S (Supplementary Fig. 1). The cells growing in this chemostat simultaneously oxidized H2S and CH4 (sulfide-adapted cells; Fig. 1b), up to concurrent loads of 42 μmol H2S · min−1 · g DW−1 and of 38 μmol CH4 · min−1 · g DW−1 (Table 1), while the H2S concentration in the gas outlet remained below 2 nmol · L−1. Steady state continuous cultures of non-adapted and sulfide-adapted cells could be maintained for many generations (Fig. 1a, b). Accumulation of elemental sulfur (S0) over weeks of growth was evident, as increasing amounts of a yellow precipitate (irregular microscopic particles) developed and attached to the metal parts and walls of the chemostat (Supplementary Fig. 2). After a few weeks of operation with H2S it was identified as being over 99% pure sulfur and the amount could account for at least 80% of the sulfide converted over this period. Through microscopy, only minute amounts of sulfur particles in the liquid could be observed as opposed to bacterial cells. Both non-adapted and sulfide-adapted cultures were operated under low dissolved O2 concentrations (1% air saturation) to minimize chemical sulfide oxidation. The low O2 concentrations also resulted in expression of hydrogenase activity as observed previously41. Control incubations in membrane-inlet mass spectrometry (MIMS) experiments without cells showed negligible oxidation of sulfide at micromolar range concentrations.

a Continuous culture oxidizing methane. b Continuous culture simultaneously oxidizing high concentrations of CH4 and H2S. c Fed-batch culture showing increase in 13C-biomass with H2S as sole energy source. Data are presented as mean ± standard deviations (n = 3 technical replicates). Source data are provided as a Source Data file.

Verrucomicrobial methanotrophs possess the Calvin-Benson-Bassham cycle for CO2 fixation42, raising the question whether they can grow as chemolithoautotroph on CO2 with H2S as energy source. Accordingly, a fed-batch reactor was inoculated with a diluted culture (OD600 = 0.05) of the dual H2S-CH4 chemostat and H2S and 13CO2 were supplemented as the only energy and carbon source, while the CH4 supply was disconnected. Over time, the biomass of M. fumariolicum SolV cells became enriched in carbon-13 by incorporating 13CO2 into biomass (Fig. 1c). When the H2S load was increased, the percentage of 13C-biomass increased accordingly. Growth was evident, as an increase in 13C-biomass was accompanied with an increase in dry weight (Supplementary Fig. 3). By quantifying H2S in the gas inlet and outlet of the reactor, H2S conversion efficiencies of ~98–100% were determined throughout the whole incubation period.

H2S consumption rates and inhibitory effects of H2S on M. fumariolicum SolV cells were measured inside a liquid-filled chamber connected to a membrane-inlet mass spectrometer (MIMS), which allows for the real-time and concurrent quantification of multiple gases, while O2 was measured by a sensor spot. A maximum CH4 conversion rate of non-adapted cells of 200 ± 11 μmol CH4 · min−1 · g DW−1 was measured with a concomitant O2 consumption rate of 302 ± 9 μmol O2 · min−1 · g DW−1 (Table 1). In comparison, for the sulfide-adapted cells a maximum CH4 conversion rate and concomitant O2 consumption rate of 33 and 30% lower was measured, respectively. Similarly, the maximum methanol respiration rates of sulfide-adapted cells were 32% lower than measured for the non-adapted cells (Table 1). Taking the 1 mol O2 required for the activation of 1 mol CH4 into account, the maximum CH4 respiration rates of the non-adapted and sulfide-adapted cells were about 3-fold lower compared to maximum methanol respiration rates (Table 1), indicating the conversion of methane to methanol as the rate limiting step. In addition, presumably due to the low dO2 concentration in the continuous cultures, the non-adapted and sulfide-adapted cells expressed a high hydrogenase activity (Table 1), with a measured H2:O2 consumption ratio of ~1:0.35 as expected32,42. As was the case for CH4 and methanol respiration, the maximum H2 respiration rates of the sulfide-adapted cells were lower than those of the non-adapted cells (Table 1). Hence, the gain in increased H2S oxidation capacity in sulfide-adapted cells comes at the expense of the CH4, methanol and H2 conversion capacities.

Sulfide-adapted cells in the chemostat oxidized H2S to low, non-inhibitory concentrations (Fig. 1b), which is necessary since the CH4 oxidation capacity of non-adapted cells (as well as sulfide-adapted cells) was affected by an H2S concentration as low as 1 μM. CH4 oxidation was inhibited by about 25%, 70–85% and 95% in the presence of 2 μM, 4-5 μM and 10 μM H2S, respectively (Fig. 2). Inhibition of CH4 conversion appeared reversible, as when H2S was consumed or flushed out of short-term incubations, CH4 conversion and CO2 production resumed immediately at their previous rates. After longer periods (2 h) of inhibition by 10–20 μM H2S, CH4 conversion rates were 25–35% lower. Whether these lower rates were the result of inhibition of pMMO or other parts of the respiratory chain as well could not be concluded as methanol (CH3OH) conversion was impaired as well after such long H2S exposures.

H2S was kept at various stable concentrations (indicated at the bottom) by pulse-wise additions of H2S to the MIMS chamber. Numbers indicate CH4 consumption rates in μmol CH4 · min−1 · g DW−1. At 170 min the MIMS chamber has become anoxic, resulting in cessation of CH4 consumption. Source data are provided as a Source Data file.

High initial O2 consumption rates were measured when only H2S was administered to non-adapted cells in the MIMS chamber. Interestingly, these rates immediately and rapidly decreased ~15-fold within a few minutes to stable rates of 10 ± 1 μmol O2 · min−1 · g DW−1 (at 40–80 μM H2S and <10 μM O2). This rapid decrease in respiration rate indicated the presence of sulfide-sensitive terminal oxidases (SSTOs) that were quickly inactivated after the addition of H2S and at least one type of sulfide-insensitive terminal oxidase (SITO) responsible for the remaining low respiration rate43. The maximum reaction rate of the SITO (10 ± 1 μmol O2 · min−1 · g DW−1) is limited, as it constitutes only 3% of the maximum respiration rate of these non-adapted cells on methanol (Table 1). At 10-fold higher O2 concentrations (70–90 µM O2 and 30–80 μM H2S), the remaining respiration rate increased ~40% (Table 1), suggesting that O2 is competing with H2S for the active site of the SSTOs, thereby alleviating H2S inhibition. SITO activity was cyanide sensitive as 95% of the respiration rate was inhibited at 1 mM potassium cyanide. The sulfide-adapted cells oxidized H2S with maximum O2 consumption rates of 53 ± 4 μmol O2 · min−1 · g DW−1 (Table 1). As at 40–80 μM H2S the SSTOs were assumed to be completely inhibited, these values represent the rates of the SITO, which are more than five times higher compared to the non-adapted cells (Table 1). H2S is primarily converted to elemental sulfur (S0), as a H2S:O2 stoichiometry of 1:0.48 ( ± 0.005; n = 3) was determined after simultaneous quantification of H2S and O2 consumption, together with the visible production of S0 (Supplementary Fig. 7b).

Maximum conversion rates of H2S at non-inhibiting, low (sub-micromolar) concentrations in the MIMS chamber were difficult to perform due to its rapid consumption that resulted in a variable inhibition. Alternatively, the maximum H2S conversion rates were determined in the dual H2S-CH4 chemostat by gradually increasing the sulfide load to 156 μmol H2S · min−1 · g DW−1 over the course of a day while monitoring the outlet concentration (Table 1). The latter increased from 2 to 25 nmol · L−1 and therefore remained below a liquid concentration of 40 nM, which was considered not to affect pMMO (as measured through MIMS incubations). Nevertheless, CH4 conversion did decrease about 40% but remained stable for days. When in a similar way the chemostat was given only H2S while CH4 was disconnected, the same maximum H2S conversion rate of 156 μmol H2S · min−1 · g DW−1 was measured. As respiration is not the limiting factor in this setup (Table 1), this rate is considered the maximum H2S conversion rate, which is 1.5 times higher than the SITO activity can account for in these sulfide-adapted cells and made possible by the SSTOs that were only partially inhibited at these low sulfide concentrations. Similar rates were measured for sulfide-adapted cells in the MIMS chamber in the presence of 60–80 μM O2 (Table 1). Hence, at low H2S and/or high O2 concentrations, the cells demonstrate the highest sulfide conversion rates, as the SSTOs are only partially inhibited. Noticeably, the above measured maximum H2S conversion rate in the MIMS chamber exceeded that of the maximum CH4 conversion rate of the sulfide-adapted cells (Table 1).

Upon addition of methanol during respiration of 20–40 μM H2S by non-adapted cells in the MIMS chamber, H2S consumption ceased immediately (Fig. 3a) but the total respiration rate increased by ~40%. In contrast, H2S oxidation by sulfide-adapted cells (having five times higher SITO activity) continued at 43% of the rate when methanol was added (Fig. 3b), while the total respiration rate increased by ~25% (Supplementary Fig. 4). Hence, methanol and H2S were respired simultaneously and seem to compete for the same terminal oxidase. When sulfide (30 µM) was added to sulfide-adapted cells during methanol respiration, O2 consumption decreased ~3-fold (Supplementary Fig. 5). The remaining respiration rates (66 μmol O2 · min−1 · g DW−1) were higher than expected from the maximum (SITO-dependent) H2S respiration rate (53 ± 4 μmol O2 · min−1 · g DW−1), indicating that at least some methanol was still respired, which was confirmed by the fact that the CO2 production rate continued at 20–30% in the presence of sulfide. In contrast, at the same H2S concentrations, CH4 respiration had ceased almost completely (Fig. 2).

a Cessation of H2S consumption by non-adapted cells after the addition of methanol (final concentration 0.4 mM). b Inhibition of H2S consumption by sulfide-adapted cells after the addition of methanol (final concentration 5 mM). Numbers indicate consumption rates in μmol H2S · min−1 · g DW−1. The black horizontal line indicates a brief moment of anoxia to demonstrate H2S oxidation is dependent on O2. Source data are provided as a Source Data file.

Addition of H2S to non-adapted cells consuming H2 decreased the H2 consumption rate by 30%, while H2S consumption only started after complete depletion of H2 (Fig. 4). Furthermore, H2 respiration was up to two times higher than H2S respiration after all H2 had become depleted. Similarly, H2S oxidation at 20–40 μM H2S was reduced by about 80% upon addition of 200 μM formic acid (CHOOH) (Supplementary Fig. 6), while the total respiration rate increased by 15%. The observation that H2S is only oxidized after H2 (or methanol) has become depleted (Fig. 4) suggests competitive electron transfer pathways to the sulfide-insensitive terminal oxidase (SITO) due to its limited respiration capacity. Interestingly, when in a separate experiment 1.2 mM H2S was oxidized as sole energy source over ~3 h and O2 additions were stopped, H2S was produced under anoxic conditions (Supplementary Fig. 7). Conceivably, a sulfide-producing enzyme is being used by strain SolV, reducing the heretofore produced polysulfides and/or elemental sulfur (S0). In contrast, H2 was not consumed in the absence of polysulfides and/or elemental sulfur under anoxic conditions, indicating that these sulfur compounds and not sulfate present in the medium was used as electron acceptor. H2S production was stimulated up to 13 μmol H2S · min−1 · g DW−1 when H2 or methanol was present as electron donor.

Green numbers indicate H2 consumption rates in μmol · min−1 · g DW−1 before and after H2S addition, respectively. The red number and line indicate H2S consumption rate in μmol · min−1 · g DW−1 after depletion of H2. Source data are provided as a Source Data file.

H2S oxidation kinetics by sulfide-adapted cells were studied using a gas chromatograph, which has a lower detection limit than the MIMS. Starting at 3.5 μM H2S and 190 μM O2, an almost linear decrease down to 1 μM H2S was observed with a rate of 167–223 μmol H2S · min−1 · g DW−1 (Fig. 5a). These rates are slightly higher than the maximum H2S conversion rates measured in the MIMS chamber at 5–30 μM H2S and 60–80 μM O2 (Table 1), which could be explained by the higher O2 and low H2S concentrations present in the incubations used for GC measurements. Because O2 and H2S compete for the sulfide-sensitive terminal oxidase (SSTO), a low H2S and high O2 concentration alleviate SSTO inhibition, leading to higher H2S consumption rates. Michaelis-Menten modeling of H2S consumption resulted in an apparent affinity constant Ks of 0.32 μM H2S. However, the H2S traces below 1 μM H2S did not follow the predicted curve and remained slightly above it (Fig. 5a). When in the MIMS chamber O2 consumption was followed down to zero at H2S concentrations of 15–20 μM an apparent affinity constant Ks of 0.14 ± 0.01 μM O2 was determined that follows Michaelis-Menten kinetics (Fig. 5b). In the presence of 15 μM H2S, SITO was not inhibited as identical O2 consumption rates were measured after sequential addition of O2 (Supplementary Fig. 8). Assuming only one terminal oxidase type to be active under these conditions and H2S conversion not being the limiting factor, a Ks of 0.14 ± 0.01 μM O2 could be a reliable value for the sulfide-insensitive terminal oxidase.

a H2S oxidation measured through gas chromatography. Different blue shaded diamonds represent biological replicates (n = 3). The reaction was initiated by addition of cells after 33 min. b H2S respiration measured through a fiber-optic oxygen sensor spot in the MIMS chamber. Black lines indicate Michaelis-Menten curve fitting. The reaction was initiated by addition of cells at 0 min. Source data are provided as a Source Data file.

To assess how M. fumariolicum SolV cells adapt to H2S, mRNA from the dual H2S-CH4 chemostat (sulfide-adapted cells) and the CH4 chemostat (non-adapted cells), both in steady state, were extracted and gene expression was quantified (Table 2; Supplementary Fig. 9; Supplementary Data 1). In sulfide-adapted cells, the operon MFUM_v2_0219-21 was upregulated about 1.7-fold. The genes in this operon are annotated as NAD(FAD)-dependent dehydrogenase (MFUM_v2_0219), a protein homologous to the sulfur carrier protein TusA (MFUM_v2_0220) and a putative sulfur carrier protein DsrE2 (MFUM_v2_0221), respectively. A more detailed investigation revealed MFUM_v2_0219 to encode a type III sulfide:quinone oxidoreductase (SQR)44. Based on gene comparisons, a second gene (MFUM_v2_0138) might encode an SQR, although this gene was not significantly upregulated in the presence of H2S and transcribed to a much smaller degree than MFUM_v2_0219 in sulfide-adapted cells (Supplementary Data 1). Two genes (MFUM_v2_0873 and MFUM_v2_1149) were transcribed that might encode sulfur dioxygenases, which could putatively oxidize elemental sulfur to sulfite (SO32−) (Supplementary Data 1). In addition, the genes MFUM_v2_0942 and MFUM_v2_0943 were upregulated 2-fold and 8-fold (Table 2) and show high similarity to genes encoding the cytochrome c protein SorB and sulfite:cytochrome c oxidoreductase SorA of Thiobacillus novellus, respectively45. In sulfide-adapted cells, the putative sulfur dioxygenase (MFUM_v2_0873) was transcribed to a similar degree as SQR (MFUM_v2_0219). However, based on the stoichiometry of 1 H2S: 0.48 O2 ( ± 0.005; n = 3) quantified in the MIMS chamber, the conversion of elemental sulfur and polysulfides via sulfite to sulfate is thought to have a minor role under the tested conditions. In addition, the oxidation of H2S was never accompanied by a decrease in pH, which would have been the case if elemental sulfur had been oxidized further to thiosulfate, sulfite or sulfate. The operon MFUM_v2_1257-61 encodes a ba3-type cytochrome c oxidase that was upregulated ~5-fold in the presence of H2S, agreeing with the 5-fold higher SITO respiration rate in sulfide-adapted cells. Interestingly, the highest upregulated gene (15-fold) encodes an 89 kDa heptahaem cytochrome c protein of unknown function (MFUM_v2_1950), showing highest similarity to genes found in thermophilic sulfide-oxidizers. In the presence of H2S, several genes encoding enzymes involved in the biosynthesis of sulfide for production of sulfur-containing metabolites (e.g., cysteine, methionine and glutathione) were heavily downregulated (Table 2). In addition, the downregulation of genes involved in CH4 oxidation and subsequent electron transfer in the respiratory chain was observed (Table 2). This downregulation is in accordance with the measured decreased maximum methane conversion and respiration rates.

The observation that M. fumariolicum SolV possesses an SQR and the fact that CH4 and H2S coexist in a large variety of environments prompted us to investigate the presence of SQR in methanotrophs. Indeed, genes encoding putative SQRs are also widespread in proteobacterial methanotrophs of various environments such as lakes, wetlands, rhizosphere, ocean sediments, permafrost soil, paddy fields, wastewater treatment plants, alkaline soda lakes, landfills and groundwater aquifers (Supplementary Fig. 10). SQRs are classified into six different types based on their structure, and differ in their affinity for H2S and their physiological role in the cell44. Putative SQRs were detected in a large variety of proteobacterial genera in which a pMMO and/or sMMO was present, such as Crenothrix, Methylobacter, Methylocaldum, Methylocapsa, Methylococcus, Methylocystis, Methylohalobius, Methylomagnum, Methylomarinum, Methylomicrobium, Methylomonas, Methyloprofundus, Methylosinus, Methylospira, Methyloterricola, Methylotetracoccus, Methylotuvimicrobium and Methylovulum (Supplementary Fig. 10). In addition, the recently isolated alphaproteobacterium ‘Methylovirgula thiovorans’ strain HY1 encodes a type I SQR40. In contrast, verrucomicrobial methanotrophs possess genes encoding a type III SQR, comprising bacterial and archaeal SQRs of which the least is known44.

In this study, we show for the first time that a microorganism can oxidize CH4 and H2S simultaneously, and that a methanotroph can produce biomass from CO2 with H2S as sole energy source. We showed that oxidation of H2S is necessary because H2S inhibits both respiration and CH4 oxidation. Cells responded to the presence of H2S by upregulating a type III sulfide:quinone oxidoreductase (SQR) and a sulfide-insensitive ba3-type terminal oxidase (SITO). In addition, we provide evidence for an H2S detoxification mechanism in methanotrophs, which, according to genomic information and the co-occurrence of methane and sulfide in a myriad of environments, seems to be widespread.

Very little is known about the effect of H2S on methanotrophy. A methanotrophic consortium sampled from a landfill showed decreased methanotrophic activity in the presence of H2S46. In addition, CH4 oxidation by Methylocaldum sp. SAD2, isolated from a sulfide-rich anaerobic digester, was significantly inhibited (44–60% decrease in methanol production) in the presence of 0.1% H2S, but the mechanism was not explored47,48. ‘Methylovirgula thiovorans’ strain HY1A was recently shown to be able to consume various reduced sulfur compounds together with CH4, but simultaneous oxidation of CH4 and H2S could not be observed40. In the peatland where strain HY1A was isolated from, the H2S concentration was below the detection limit, suggesting that a vigorous H2S detoxification might not be necessary. In contrast, the geothermal environments where M. fumariolicum SolV and other verrucomicrobial methanotrophs reside, are characterized by high concentrations of H2S (from <50 ppm to 20000 ppm)28,35,49. Accordingly, the demonstrated ability to fix CO2 with H2S as sole energy source and efficiently oxidize H2S to S0 could be highly advantageous in such harsh systems. Considering that in the natural environment multiple substrates coexist, a mixotrophic lifestyle, in which CH4, H2 and H2S are utilized simultaneously is expected to be more beneficial32,50.

H2S is known to bind to metals such as copper and iron, which could lead to inhibition of the CH4 oxidation capacity of the copper-dependent pMMO and terminal oxidases involved in the reduction of O21,4,5,6,51,52. Interestingly, ‘Methylovirgula thiovorans’ strain HY1A only encodes an iron-dependent sMMO40, whereas M. fumariolicum SolV encodes three copper-dependent pMMOs16. The former strain does not simultaneously oxidize H2S and CH4, while the latter has a rapid H2S detoxification system to alleviate inhibition of methanotrophy. The extent to which a type of methane monooxygenase is inhibited by H2S could therefore influence the need for an H2S detoxification system. Because in M. fumariolicum SolV the gene encoding a type III SQR was upregulated in the presence of H2S, we propose that this enzyme is responsible for the observed oxidation of H2S to elemental sulfur. Indeed, type III SQRs were shown to couple the oxidation of H2S to the reduction of quinones in several archaea and bacteria53,54. In verrucomicrobial methanotrophs, three different types of terminal oxidases are found: an aa3-type, a ba3-type, and a cbb3-type16. Possessing multiple types of terminal oxidases allows a branched electron transport chain, which is highly advantageous in environments with fluctuating conditions and varying substrate and oxygen availability. Through respiration studies, we showed that M. fumariolicum SolV possesses one or more sulfide-sensitive terminal oxidases (SSTO) and at least one sulfide-insensitive terminal oxidase (SITO). Because a ba3-type terminal oxidase is strongly upregulated in cells growing at high H2S loads, we propose this specific enzyme complex to be the dedicated SITO in verrucomicrobial methanotrophs. Similarly, in sulfur-grown cells of Acidithiobacillus ferrooxidans this ba3-type oxidase was upregulated55. The highly upregulated heptahaem cytochrome c protein (MFUM_v2_1950) in M. fumariolicum SolV might be involved as electron carrier from SQR to the electron transport chain. This putative electron carrier could explain why H2S respiration still partially continues upon addition of methanol in the sulfide-adapted cells and not in the non-adapted cells. In the latter, the lack of this putative heptahaem electron carrier could be the limiting factor for H2S respiration, being overruled by the relatively large amounts of the electron carrier XoxGJ, mediating electron transfer from methanol to the terminal oxidase56. In contrast, in non-adapted cells the ratio in transcripts of the genes encoding XoxGJ and the putative heptahaem electron carrier is 27.6 compared to 1.2 in sulfide-adapted cells. Accordingly, the upregulation of the gene encoding the heptahaem electron carrier might enable sulfide respiration to occur concurrently with methanol oxidation, using the same terminal oxidase. H2S impedes both the SSTO, and the reaction catalysed by pMMO, as at 10 μM H2S the conversion of CH4 was almost completely inactivated while methanol, formate and H2 conversion could still proceed. The observed decrease in CH4 conversion in the chemostat at a maximum H2S load of 156 μmol H2S · min−1 · g DW−1 (liquid concentration <40 nM) was more than can be expected from our methane conversion inhibition studies and may indicate that a large portion of the respiratory chain is used for electrons generated by H2S oxidation, resulting in an overreduced Q-pool which prohibits proper functioning of alternative complex III (ACIII). Oxidation of H2S is needed to keep this molecule at low, non-inhibitory concentrations. Consequently, the electrons released from this oxidation need to be processed by the electron transport chain, leading to substrate competition during the simultaneous oxidation of multiple compounds such as H2S and CH4. Similarly, it was proposed that an overactive SQR in Rhodobacter capsulatus could lead to an overreduction of the quinone pool57. The upregulated ba3-type oxidase may alleviate this problem by oxidizing quinol and reducing the terminal electron acceptor O2. In Aquifex aeolicus, a related ba3-type oxidase was found in a supercomplex with SQR58. This terminal oxidase was shown to not only oxidize reduced cytochrome c, but also ubiquinol directly59. In strain SolV there may be an important role for the highly upregulated heptahaem protein as a dedicated electron shuttle between the quinone-accepting ACIII and the ba3-type oxidase. A branched electron transport chain with different terminal oxidases enables metabolic versatility and adaptations. For example, E. coli uses the proton-pumping bo3-type oxidase during growth but requires the sulfide-insensitive bd-type oxidases to keep growing in the presence of H2S60. Interestingly, two genes are present that could encode sulfur dioxygenases (MFUM_v2_0873 and MFUM_v2_1149) to further oxidize elemental sulfur. However, the measured stoichiometry of 1 H2S to 0.48 O2, the production of elemental sulfur and absence of acid production clearly show that H2S is not oxidized further to a significant extent. It remains to be investigated if methanotrophs can oxidize H2S further to sulfite and sulfate.

Cells of M. fumariolicum SolV were shown to rapidly oxidize H2S with a low apparent affinity constant (i.e., high affinity) below 1 μM H2S. The observed kinetic values are not surprising, since H2S already inhibits methanotrophy at such low concentrations. Through gas chromatography, an exact apparent affinity constant for whole cells could not be determined, as H2S consumption did not follow a typical Michaelis-Menten curve. A limitation of the respiratory capacity for H2S oxidation above about 1 μM may explain such deviation and could be resolved by purification of SQR. The observation that M. fumariolicum SolV reduced elemental sulfur or polysulfides to H2S in the presence of H2 or methanol is intriguing. ‘Methylovirgula thiovorans’ strain HY1A grown on thiosulfate increasingly produced an enzyme that resembles a protein known to possess sulfhydrogenase activity40. Interestingly, this enzyme clusters with the group 3b [NiFe] hydrogenase of M. fumariolicum SolV, thought to be involved in the production of NAD(P)H for CO2 fixation50. Indeed, it is proposed that these hydrogenases can have sulfhydrogenase activity, which could be a mechanism to dispose of reducing equivalents61,62. Hence, the group 3b [NiFe] hydrogenase of M. fumariolicum SolV might be responsible for the conversion of S0 to H2S. Culturing in chemostats again turned out to be a very powerful tool to investigate the metabolism of methanotrophs41,63. Through adaption, M. fumariolicum SolV was able to respire H2S at a rate five times higher than non-adapted cells, presumably due to the upregulation of SQR and the ba3-type terminal oxidase.

Verrucomicrobial methanotrophs that thrive in geothermal environments possess a clear mechanism to cope with H2S. Accordingly, SQR and a sulfide-insensitive terminal oxidase could enable these methanotrophs to thrive in H2S-rich environments. Indeed, pyrosequencing showed that Methylacidimicrobium-related 16 S rRNA gene sequences were abundantly present in the crown of concrete sewage pipes rich in CH4 and H2S64. Concerning proteobacterial methanotrophs, the effect of H2S warrants further investigation. Because aerobic methanotrophs live in environments in which H2S is often present, we propose that the mechanism of H2S detoxification is widespread in methanotrophs in various environments.

Methylacidiphilum fumariolicum SolV used in this study was isolated from a mud pot of the Solfatara near Naples, Italy28. The genome of this strain is publicly available and accessible at Genoscope [https://mage.genoscope.cns.fr/microscope/mage/viewer.php?O_id=1176], as well as at EMBL/NCBI (BioProject PRJEA85607; accession ERS14853105). This environment is characterized by large sulfide emissions, high temperatures and extremely low pH values. The growth medium was composed of 0.2 mM MgCl2, 0.2 mM CaCl2, 1 mM Na2SO4, 2 mM K2SO4, 7.5 mM (NH4)2SO4 and 1 mM NaH2PO4 and trace elements at final concentrations of 1 μM NiCl2, 1 μM CoCl2, 1 μM MoO4Na2, 1 μM ZnSO4, 1 μM CeCl3, 5 μM MnCl2, 5 μM FeSO4, 10 μM CuSO4 and 40–50 μM nitrilotriacetic acid (NTA). Cells were grown as methane-limited continuous culture at 55 °C as described before65, except that the pH was regulated at 2.5–3.0, that a small 400 mL chemostat was used with medium as described above and that H2 was not supplemented. The oxygen concentration was regulated at 1% air saturation. In addition, a second chemostat was operated under similar conditions, but to which H2S was added through an additional gas inlet (Supplementary Fig. 1). H2S was produced by mixing 100 mM anoxic Na2S and 210 mM HCl in a 50 mL bottle with a peristaltic pump. The argon/CO2 (95%/5%, v/v) gas stream to the reactor was led through this bottle. In order to determine the maximum H2S conversion rate of the chemostat, the cells were gradually exposed to higher H2S concentrations by regulating the peristaltic pump. The H2S concentrations in the gas inlet and gas outlet were determined using gas chromatography (described in the subsection: Batch incubations and gas chromatography). Because H2S was supplied through the gas inlet and therefore needs to be transferred to the liquid phase, the liquid H2S concentration will be close to or lower than its equilibrium concentration, which at 55 °C is 1.6 times the gas concentration (calculated from the Ostwald coefficient at 55 °C)66. In addition, to observe whether M. fumariolicum SolV can grow on H2S as sole energy source a fed-batch culture was operated in the same setup as the chemostat system. In this case the medium flow was stopped, and the argon/CO2 gas was changed for an argon only gas stream. At the same time equal amounts of a 13C-labeled bicarbonate solution (50 mM) and HCl solution (100 mM) were additionally added to the sulfide mixing bottle, creating a 13C-CO2 gas concentration of about 2%. 5 mL biomass samples from the fed-batch culture were collected by centrifugation over several days and the pellets were washed with acidified water (pH 3). Pellets were then resuspended in small amounts of acidified water and samples were subsequently pipetted into tin cups and dried overnight at 70 °C under vacuum. 13CO2 incorporation into biomass was assessed by measuring the 13/12C ratio using a Finnigan DeltaPlus isotope-ratio mass spectrometer (IR-MS) as described before42.

To accurately measure dissolved gases, membrane-inlet mass spectrometry (MIMS) was performed as described previously65, except that a 30 mL MIMS chamber was used. All rates were measured at 52 °C. The inserted probe consisted of a blunt end stainless steel tube (diameter 3 mm) that was perforated with 4–16 holes of 1 mm diameter. The holes were covered with silicon tubing (Silastic, 50VMQ Q7-4750 Dow Corning, supplied by Freudenberg Medical via VWR international, 1.96 mm outer diameter x 1.47 mm inner diameter). For easy mounting the silicon tubing was soaked briefly in hexane, which causes silicone to swell. The metal part was wetted with iso-propanol as lubricant. The probe was directly connected via a 1/8- or 1/16-inch stainless steel tube to the MS that was operated at 40 μA emission current. Medium with a pH equal to that of the culture (pH 2.5–3.0) added to the chamber was first flushed with 3% CO2 in argon gas after which the oxygen concentration was adjusted to the desired value by adding pure oxygen gas or air via the headspace. Mass 15 and 16 are both dominant masses for CH4 in the mass spectrometer, but mass 15 has a much lower background signal than mass 16 and was therefore chosen to measure CH4. Methane and hydrogen (mass 2) were added as a gas in the headspace or, in the case of calibration, from a saturated stock solution. These stock solutions were prepared in a closed bottle with water at room temperature and a headspace of pure gas with known pressure. For the solubility in water 1.47 mM and 0.80 mM were taken for methane and hydrogen, respectively (at 22 °C and 1 bar)66. When CO2 production rates were to be measured, 13C-bicarbonate and equimolar amounts of sulfuric acid were added after flushing with argon. In this way the simultaneously occurring CO2 fixation is mainly from 13C-labeled CO2 (mass 45), leading to less interference with measurement of CO2 production. At the start, unlabeled CO2 (mass 44) was very low, and its increase reflected almost exclusively CO2 production from unlabeled methane or methanol.

The stoichiometry of H2S oxidation was determined through pulse-wise additions of a sulfide stock solution and O2 (as tiny gas bubbles with a syringe) in order to keep concentrations low at 1–20 μM H2S and 0–5 μM O2. In total, 0.7–1.4 mM of Na2S was added over a period of 1.5–3 h. During this experiment, equimolar amounts of a 200 mM sulfuric acid stock solution were added simultaneously to limit the pH change within 0.2 units. The oxygen concentration was simultaneously measured in the MIMS chamber by means of a fiber-optic oxygen sensor spot (TROXSP5, PyroScience, Aachen, Germany) that was glued on the inside of the chamber. These spots could measure down to about 20 nM oxygen, which is much lower than can be measured with the mass 32 signal of MIMS.

To determine kinetic parameters of H2S oxidation by sulfide-adapted cells, batch incubations were performed in 120 mL serum bottles containing 10 mL medium without any trace elements. Trace elements were omitted to minimize the effect of abiotic sulfide oxidation. The bottles were closed with butyl rubber stoppers. Incubations were performed at 55 °C and 350 rpm. H2S was prepared by mixing Na2S with HCl in a closed bottle. A volume headspace was taken and injected into 120 mL serum bottles and equilibrated for 30 min before initiating the assay by addition of cells. H2S was measured by injecting 100 μL of the headspace of the bottles with a Hamilton glass syringe into a GC (7890B GC systems Agilent technologies, Santa Clara, USA) equipped with a Carbopack BHT100 glass column (2 m, ID 2 mm) and a flame photometric detector (FPD). The areas obtained were used to calculate H2S amounts using calibration standard curves with H2S. Briefly, 400 μL of a 25 mM Na2S stock (sodium sulfide nonahydrate, purity >98%, Sigma-Aldrich) was acidified with 2 mL 0.5 M HCl in a 574 mL bottle creating a headspace concentration of 17.4 nmol · mL−1. Small volumes of the headspace were subsequently added to a 1162 mL bottle to create various H2S concentrations to be injected (0.1 mL) into the GC for calibration. The calibration curve ranged from ~1 nmol · L−1 to 1 μmol · L−1 H2S.

For each replicate, 10 mL was sampled from the chemostat, and cells were immediately pelleted for 3 min at 15,000 × g, snap-frozen in liquid nitrogen and stored at –80 °C. Cells were harvested from cultures in steady state, which corresponds to constant parameters over at least 5 reactor volume changes. Total RNA was isolated using the RiboPure™ RNA Purification Kit for bacteria (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer's protocol. Ribosomal RNA was removed from the total RNA samples to enrich for mRNA using the MICROBExpress™ Bacterial mRNA Enrichment Kit (Thermo Fisher Scientific) according to the manufacturer's protocol. The Qubit™ RNA HS Assay Kit (Thermo Fisher Scientific) and the Agilent RNA 6000 Nano Kit (Agilent Technologies, Waldbronn, Germany) and protocols were used for the quantitative and qualitative analysis of the extracted total RNA and enriched mRNA. The latter was used for library preparation by using the TruSeq Stranded mRNA Reference Guide (Illumina, San Diego, CA, USA) according to the manufacturer's protocol. For quantitative and qualitative assessment of the synthesized cDNA, the Qubit™ dsDNA HS Kit (Thermo Fisher Scientific) and the Agilent High Sensitivity DNA kit (Agilent Technologies) and protocols were used. Transcriptome reads were checked for quality using FastQC67 and subsequently trimmed 10 base pairs at the 5' end and 5 base pairs at the 3' end of each read. Reads were mapped against the M. fumariolicum SolV complete genome (accession number LM997411)68 using Bowtie269. The remainder of the analysis and the production of images was performed in version 4.0.2 of the R environment70. The mapped read counts per gene were determined using Rsubread71 and fold change and dispersion were estimated using DEseq272. Before doing any statistics, principal component analysis on the top 1000 genes by variance of each sample was performed to check whether samples within the same condition were both similar to each sample part of the same condition, and dissimilar to any other sample. For differential expression, a Wald test was employed by DEseq2 to calculate adjusted p-values. Differences in counts were considered to be significant if the basemean was >4, the log2-fold change was higher than [0.58] and the adjusted p-value was ≤0.05. For easy comparisons between samples, TPM (Transcripts Per Kilobase Million) values were calculated.

The total organic carbon (TOC) concentrations of the cultures were determined using a TOC-L CPH/CPN analyzer (Shimadzu, Duisburg, Germany). Samples were diluted three times in Milli-Q water before measurements and subsequently sparged for 20 min with ozone while stirring to remove CO2 from the liquid. Acidification of the solutions was not needed due to the low pH of the samples. An optical density of 1 measured at 600 nm is equivalent to ~450 mg dry weight (DW) per litre.

All available genome sequences of known methylotrophs from the orders Methylococcales (Gammaproteobacteria) and Methylacidiphilales (Verrucomicrobia), the families Methylocystaceae and Beijerinckia (Alphaproteobacteria), and the genus Methylomirabilis were retrieved from the NCBI database. Genomes of methanotrophs were selected by blasting amino acid sequences of PmoA from Methylococcus capsulatus (SwissProt accession Q607G3) and sMMO from Methylosinus acidophilus (NCBI accession AAY83388.1) with an e-value threshold of 10−3 and a %-id threshold of >30%. Genomes containing a methane monooxygenase sequence were subsequently mined for putative SQR sequences by blasting a representative sequence of each of the SQR subtypes as defined by previous research44: type I, WP_010961392.1; type II, WP_011001489.1; type III, WP_009059890.1; type IV, WP_011372252.1; type V, WP_012502121.1; type VI, WP_011439951.1. Putative SQR sequences were aligned with those in the phylogenetic tree of44 using Muscle 3.8.155173 with default settings. A maximum-likelihood phylogenetic tree with 500 bootstrap replicates was constructed using RAxML 8.2.1074 using the rapid bootstrapping option and the LG amino acid substitution model75. The final tree was visualized using MEGA7 and the clade of flavocytochrome c sulfide dehydrogenase (FCSD) sequences was used as outgroup.

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

The RNA sequencing data in this study have been deposited in the NCBI database under accession number PRJNA766544. The genome of Methylacidiphilum fumariolicum SolV has been deposited in the NCBI database under accession number ERS14853105. Supplementary Data 1 and the Source Data file are also available on figshare (https://doi.org/10.6084/m9.figshare.22779005). Source data are provided with this paper.

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R.A.S., S.S.M., T.B., and H.J.M.O.d.C. were supported by the European Research Council (ERC Advanced Grant project VOLCANO 669371), M.S.M.J. was supported by the European Research Council (ERC Advanced Grant project EcoMoM 339880), W.V. was supported by the Netherlands Organisation for Scientific Research (NWO) grant VI.Vidi.192.001 and S.H.P. was supported by the Netherlands Organisation for Scientific Research (NWO) grant ALWOP.308. We want to thank Dr. Marianne Guiral and Dr. Frauke Baymann (CNRS, Aix-Marseille University, Marseille, France) for fruitful discussions. The LABGeM (CEA/Genoscope & CNRS UMR8030), the France Génomique and French Bioinformatics Institute national infrastructures (funded as part of Investissement d’Avenir program managed by Agence Nationale pour la Recherche, contracts ANR-10-INBS-09 and ANR-11-INBS-0013) are acknowledged for support within the MicroScope annotation platform.

Rob A. Schmitz

Present address: Institute of Biogeochemistry and Pollutant Dynamics, Department of Environmental Systems Science, ETH Zurich, 8092, Zurich, Switzerland

Department of Microbiology, Radboud Institute for Biological and Environmental Sciences, Radboud University, Heyendaalseweg 135, 6525AJ, Nijmegen, The Netherlands

Rob A. Schmitz, Stijn H. Peeters, Sepehr S. Mohammadi, Tom Berben, Timo van Erven, Carmen A. Iosif, Theo van Alen, Wouter Versantvoort, Mike S. M. Jetten, Huub J. M. Op den Camp & Arjan Pol

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R.A.S., S.S.M., A.P. and H.J.M.O.d.C. designed the project and experiments. R.A.S., S.S.M., T.v.E., C.A.I., T.v.A., W.V. and A.P. conducted the experiments. T.B. conducted phylogenetic analyses. R.A.S., S.H.P., S.S.M., T.v.E., C.A.I. and A.P. performed data analyses. R.A.S., M.S.M.J., H.J.M.O.d.C. and A.P. wrote the manuscript. R.A.S., M.S.M.J., H.J.M.O.d.C. and A.P. supervised the research.

Correspondence to Huub J. M. Op den Camp.

The authors declare no competing interests.

Nature Communications thanks Christiane Dahl and the other, anonymous, reviewers for their contribution to the peer review of this work. A peer review file is available.

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Schmitz, R.A., Peeters, S.H., Mohammadi, S.S. et al. Simultaneous sulfide and methane oxidation by an extremophile. Nat Commun 14, 2974 (2023). https://doi.org/10.1038/s41467-023-38699-9

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Received: 04 January 2023

Accepted: 11 May 2023

Published: 23 May 2023

DOI: https://doi.org/10.1038/s41467-023-38699-9

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Simultaneous sulfide and methane oxidation by an extremophile