Metabolic Basis for the Microbial Oxidation of Atmospheric Methane

doi:10.21203/rs.3.rs-3461927/v1

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Metabolic Basis for the Microbial Oxidation of Atmospheric Methane

https://doi.org/10.21203/rs.3.rs-3461927/v1

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published 16 May, 2024

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Atmospheric methane oxidizing bacteria (atmMOB) constitute the sole biological sink for atmospheric methane and have been discovered worldwide over the past decades. Still, insufficient knowledge about the metabolic basis of atmMOB, caused by the lack of pure cultures, limits our ability to manage, study, and exploit the atmospheric methane sink and thus to fight the 21st century methane surge. Here we combine filter cultivation, trace gas oxidation, ¹⁵N₂-incorporation experiments, and comparative proteomics, to assess the potential of seven methanotrophic species to grow on atmospheric methane. Four species, three of which are outside the canonical atmMOB group USCα, enduringly oxidized atmospheric methane, hydrogen, and carbon monoxide with distinct substrate preferences over a 12-month growth period "on air". Despite this mixotrophy and high specific affinities for methane, the estimated energy yields of the atmMOB were substantially lower than previously assumed necessary for cellular maintenance, contradicting the basic energy premise for atmMOB. Comparative proteomics indicate major physiological adjustments to grow “on air” as the atmMOB allocated their proteomes to decrease energy intensive processes, including biosynthesis, and increase investments into trace gases oxidation. Our work outlines the metabolic basis of atmMOB, microorganisms that exploit the atmosphere as energy and carbon source while mitigating the potent greenhouse gas methane.

Biological sciences/Biochemistry/Proteomics

Biological sciences/Biophysics/Bioenergetics

Biological sciences/Physiology/Metabolism

Biological sciences/Microbiology/Bacteria/Bacterial physiology

Biological sciences/Ecology/Ecosystem services

During the first two decades after emission to the atmosphere, methane (CH₄) is a greenhouse gas 80 times more potent than carbon dioxide (CO₂)^1,2. Since 2007, the atmospheric CH₄ concentration (1905 p.p.b.v. in July 2022 https://gml.noaa.gov/ccgg/trends_ch4/), that is responsible for approximately 20% of the direct radiative forcing, has been increasing rapidly¹. The CH₄ increase further accelerated in 2014 and is linked to several causes: A decline in the atmospheric concentration of hydroxyl radicals (OH) which is the main sink of atmospheric CH₄ as OH oxidize CH₄ in the atmosphere^3–5, anthropogenic emissions from fossil fuel, agricultural, and waste sources⁶, and increased microbial CH₄ production in wetlands which suggests that current increases are also driven by feedback responses to global warming⁷. Atmospheric CH₄ oxidizing bacteria (atmMOB), a subgroup of aerobic methanotrophs, that oxidize CH₄ at its atmospheric trace concentration are the only biological sink of atmospheric CH₄. Compared to the OH sink (~ 500 Tg), the biological sink is rather small as it removes approximately 30 Tg (11–49 Tg) CH₄ from the atmosphere every year⁴. However, the biological sink, in contrast to the declining OH, has the potential to grow with increasing CH₄ concentrations. Additionally, it is within reach of management practices devised to maximize its natural potential and to harness it for CH₄ removal^8,9. Yet, substantial uncertainties concerning the size of the biological sink, and the ecology and metabolic basis of atmMOB, caused by the lack of atmMOB in pure culture, impedes our ability to study, manage, and exploit the sink⁸. In the following, by screening seven methanotrophic species, we outline the metabolic basis that enables atmMOB to overcome the apparent energetic limitations inherent to oxidation of atmospheric CH₄ and to serve as atmospheric CH₄ sink.

In 1992, ten years after Harriss et al. reported the first indications of atmospheric CH₄ oxidation by microorganisms¹⁰, Bender and Conrad concluded from biphasic CH₄ oxidation kinetics of soils that an unknown group of methanotrophs might be responsible for atmospheric CH₄ oxidation¹¹. Since then, two major, yet only partially answered questions, have shaped most of the research within this field: Which methanotrophs are responsible for oxidation of atmospheric CH₄? How can these organisms survive and grow with apparent energetic limitations inherent to the oxidation of the low atmospheric CH₄ concentrations?

Isotopic labeling studies revealed that members of Alpha- and Gammaproteobacteria contributed to atmospheric CH₄ oxidation and assigned them to the upland soil clusters alpha and gamma (USCα and USCγ)^12–14. Several environmental and ecological studies have ascribed atmospheric CH₄ oxidation mainly to these two clusters^15–18. However, over the years, studies targeting methanotrophs in upland soils have also reported the presence of alphaproteobacterial methanotrophs outside the USCα^12,19–22. These observations suggest that also “conventional” methanotrophs (methanotrophs assumed to grow only at high CH₄ concentrations), from genera like Methylocapsa, Methylosinus and Methylocystis, could contribute to the atmospheric CH₄ sink.

Dunfield²³ summarized three potential lifestyles of atmMOB that might enable cellular maintenance and growth at the low CH₄ concentrations in air and the associated energy limitation: (i) Flush feeding on high CH₄ concentrations generated periodically in deeper soil layers, in addition to atmospheric CH₄ oxidation; (ii) An oligotrophic lifestyle based on atmospheric CH₄ as sole carbon and energy source; (iii) A mixotrophic lifestyle to utilize other substrates for energy conservation in addition to CH₄.

Flush feeding (i) is supported by the declining potential of methanotrophs to oxidize atmospheric CH₄ after several months of CH₄ starvation^23,24. A study on conventional methanotrophs in rice paddy soils provided the first experimental evidence for flush feeding, where the methanotrophs regained the ability to oxidize atmospheric CH₄ after exposure to high CH₄ concentrations²⁵. A high specific affinity (${a}_{A}^{0}$) for CH₄ has been suggested as the key trait of oligotrophic atmMOB (ii) to enable growth with air as their only energy and carbon source^23,26. This is based on the assumption that cellular energy requirements for maintenance are 4.5 kJ C-mol^− 1 h^− 1 (at 25°C)²⁷. Thus, to survive, oligotrophic atmMOB presumably need an atmospheric CH₄ oxidation rate high enough to meet these energy requirements. Such a rate can be achieved by the combination of a high affinity for CH₄, reflected in a low half saturation constant (K_m), and a high maximum CH₄ oxidation rate (V_max), the fraction of V_max and K_m being referred to as ${a}_{A}^{0}$ ²⁸. This theory is supported by low K_m values for CH₄ found in several soils¹¹. However, in a later study the cell specific CH₄ oxidation of USCα members was estimated to be 2.9 to 40 times lower than the presumed rate needed for cellular maintenance²⁹. Therefore, the authors considered that a mixotrophic lifestyle (iii) could be the basis for atmospheric CH₄ oxidation. In line with this, a recent study reported the simultaneous oxidation of atmospheric hydrogen (H₂), carbon monoxide (CO), and CH₄ by Methylocapsa gorgona MG08, the first known methanotroph and USCα member in pure culture that can grow “on air” (with air as sole energy and carbon source)^30,31. Despite its mixotrophic lifestyle, M. gorgona MG08 does not conserve enough energy (0.38 kJ C-mol^− 1 h^− 1) to cover the 2.8 kJ C-mol^− 1 h^− 1 theoretically required to support maintenance at 20°C (2.8 kJ C-mol^− 1 h^− 1 at 20°C correspond to 4.5 kJ C-mol^− 1 h^− 1 at 25°C)^27,30, questioning whether this maintenance energy value and thus a high ${a}_{A}^{0}$ is a relevant benchmark for the physiological capabilities of trace gas oxidizing bacteria. While the isolation of M. gorgona MG08 is a key advancement in understanding atmMOB, our knowledge of the metabolic basis allowing atmMOB to grow on atmospheric CH₄, despite the apparent energetic limitation, remains incomplete as we lack physiological data from other atmMOB. On this background and to answer if atmospheric CH₄ oxidation is not only restricted to members of the USCα and USCγ, we used filter cultivation to screen the six alphaproteobacterial, methanotrophic strains Methylocapsa gorgona MG08, Methylosinus trichosporium OB3b, Methylocystis rosea SV97, Methylocapsa aurea KYG, Methylocapsa acidiphila B2, and Methylocapsa palsarum NE2^31–36 and the gammaproteobacterial strain Methylobacter tundripaludum SV96 for their ability to grow on air. To outline the metabolic basis that enables the strains confirmed as atmMOB to live on air, we applied trace gas oxidation experiments, estimated their energy yield from the oxidation of trace gases in air, used comparative proteomics to investigate how atmMOB allocate their proteome when exposed to air, determined the ${a}_{A}^{0}$ for CH₄, and evaluated the ability of atmMOB to fix dinitrogen (N₂) when grown on air, using nanoscale secondary ion mass spectrometry (NanoSIMS) based ¹⁵N₂-incorporation measurements.

Colony formation, trace gas oxidation, and cellular energy yield on air

To test the ability of the seven selected methanotrophs to grow on air, we incubated each strain on filters floating on carbon free medium with ambient air as the sole carbon and energy source, hereafter referred to as filter culture. Microscopy demonstrated that all strains except Methylobacter tundripaludum SV96, the gammaproteobacterial strain, formed colonies during six months of incubation (Fig. 1A). To test the activity of the six months old strains, we performed trace gas oxidation experiments (Fig. 1A, 1B). During these experiments, filter cultures were floating on mineral medium in bottles with trace concentrations of CH₄, H_2, and CO in the headspace. The strains, M. aurea KYG, M. gorgona MG08, M. palsarum NE2, and M. rosea SV97, hereafter referred to as atmMOB, oxidized CH₄, CO, and H₂, or CH₄ and CO to sub-atmospheric concentrations showing strain-specific oxidation patterns (Fig. 1A, Supplementary Fig. 1). M. aurea KYG oxidized CO at the highest rate, followed by CH₄ and H₂. M. gorgona MG08 oxidized H₂ at the highest rate, followed by CO and CH₄. M. rosea SV97 oxidized CO at the highest rate, followed by CH₄ and H₂. M. palsarum NE2 oxidized CH₄ and CO at similar rates but did not oxidize H₂. The gas oxidation patterns were similar after 12 months of incubation with air (Supplementary Fig. 1), confirming both the substrate preferences of these strains and their ability to live and grow for extended periods of time with air as the sole energy and carbon source. This is in line with the previous observation of sustained growth on air for 12 months by M. gorgona MG08³¹. The observed oxidation of at least one atmospheric trace gas in addition to CH₄ demonstrates that all four strains are mixotrophic, matching the proposition by Dunfield that mixotrophy could be a physiological basis for atmMOB²³. Additionally, the strain-specific trace gas uptake patterns demonstrate substantial metabolic differences between these four strains from two different genera, and therefore indicate a diverse metabolic basis for the biological oxidation of atmospheric CH₄. These results also demonstrate that the enduring oxidation of atmospheric CH₄ after 12 months of growth on air is not restricted to members of the USCα and USCγ, since M. aurea KYG, M. palsarum NE2, and M. rosea SV97 do not cluster within either of these two lineages (Fig. 2).

Despite formation of colonies when exposed to air only, M. acidiphila B2 and M. trichosporium OB3b did not oxidize trace gases after six months of incubation (Fig. 1A). The ability of these two strains to produce polyhydroxyalkanoates (PHA) as storage compounds^34,37 might serve as an explanation for the initial colony formation. Possibly, the two strains accumulated PHA during pre-cultivation at 20% CH₄, and then sustained their growth on air by utilizing PHA and atmospheric CH₄ as energy and carbon sources until the depletion of their PHA storage. A study showing that both strains could not stay active at low CH₄ concentrations²¹ supports our observations, but further studies are needed to clarify whether the initial colony formation is based on PHA storages.

Next, we asked whether atmMOB can obtain enough energy from growth on air to meet the basic maintenance energy of 2.8 kJ C-mol^− 1 h^− 1 at 20°C postulated previously²⁶. To calculate the strain-specific energy yields in C-mol, we first estimated the cellular energy yields based on the measured CH₄, H₂, and CO oxidation rates. M. aurea KYG yielded 6.89 x 10^− 12 J cell^− 1 h^− 1 (n = 5, SD = 3.22 x 10^− 12), M. gorgona MG08 2.21 x 10^− 12 J cell^− 1 h^− 1 (n = 5, SD = 4.36 x 10^− 13), M. palsarum NE2 3.09 x 10^− 12 J cell^− 1 h^− 1 (n = 5, SD = 2.90 x 10^− 13), and M. rosea SV97 3.879 x 10^− 12 J cell^− 1 h^− 1 (n = 5, SD = 1.57 x 10^− 12) by the oxidation of either two or three trace gases in air (Fig. 1B). Due to the higher free energy potential and atmospheric concentration of CH₄ compared to atmospheric H₂ and CO, the energy estimates predict CH₄ as the major energy source for all four strains. Our estimates derive from the Gibbs free energy change of the following reactions at atmospheric conditions: ${CH}_{4}+2{O}_{2}\to 2{H}_{2}O+C{O}_{2}$, ${2H}_{2}+{O}_{2}\to 2{H}_{2}O$, and $2CO+{O}_{2}\to 2C{O}_{2}$ that amount to -797.4 kJ mol⁻¹, -236.8 kJ mol⁻¹, and − 199.9 kJ mol⁻¹, respectively. However, these numbers do not account for the energy required for activation of CH₄ by the pMMO or production cost of the enzymes involved in energy conservation from the gases. While the oxidation of H₂ and CO is catalyzed by one enzyme⁴⁰, the oxidation of CH₄ to CO₂ by atmMOB involves at least seven enzymes (Fig. 4). Thus, due to the larger investments required for energy conservation from CH₄, the gases CO and H₂ might play more important roles as energy sources for growth on trace gases in air than indicated by the energy calculations alone. Considering the cellular dry masses and carbon content (Supplementary table S3) of M. gorgona MG08, M. aurea KYG, M. palsarum NE2, and M. rosea SV97, the energy yields per cell and hour translates to 0.40 kJ C-mol⁻¹ h⁻¹ (SD = 0.08 kJ C-mol^− 1 h⁻¹), 0.71 kJ C-mol^− 1 h^− 1 (SD = 0.33 kJ C-mol^− 1 h^− 1), 0.38 kJ C-mol^− 1 h^− 1 (SD = 0.04 kJ C-mol^− 1 h^− 1), and 0.65 kJ C-mol^− 1 h^− 1 (SD = 0.26 kJ C-mol^− 1 h^− 1), at 20°C, respectively (Fig. 3). These estimated energy yields for M. gorgona MG08, M. aurea KYG, M. palsarum NE2, and M. rosea SV97 are 3.9–7.4 times lower than the reported average energy requirements for cellular maintenance in aerobic bacteria (2.8 kJ C-mol^− 1 h^− 1 at 20°C correspond to 4.5 kJ C-mol^− 1 h^− 1 at 25°C)^27,30. Thus, our observations contradict the basic energy premise for the lifestyle of atmospheric CH₄ oxidizing bacteria. Our energy estimations correspond to the similarly low energy estimates previously reported for M. gorgona MG08³⁰ and that of acetogenic and methanogenic microorganisms (0.2 kJ C-mol^− 1 h^− 1 at 37°C)⁴¹.

Comparative proteomics

We further investigated the cellular adjustments required for life on air by M. gorgona MG08, M. rosea SV97, and M. palsarum NE2 by comparing the proteomes of the three strains when exposed to air (~ 1.9 p.p.m.v. CH₄) and when exposed to high CH₄ concentrations (~ 1000 p.p.m.v. CH₄) in air. Correspondence analyses (CA) of relative protein abundances at the two CH₄ concentrations revealed a clear difference in proteome allocation. This is shown by the separation of samples from the two conditions along the first CA dimension, which accounted for 79.3%, 76.5%, and 69.6% of the total inertia for M. gorgona MG08, palsarum NE2, and M. rosea SV97, respectively (Supplementary Fig. 2). Thus, the CA indicate that the largest shifts in the proteomes occurred as responses to changes in CH₄ concentration. To identify which proteins contributed most to these shifts, we identified the proteins (top 10%) with the largest contribution to the first-dimension inertias of the CA and plotted their abundances across the two conditions (Supplementary Fig. 3). We found major protein expression shifts within a variety of functional categories (based on hierarchical EggNOG⁴² annotations), but for all three strains, a large proportion was related to core metabolisms, including transcription, translation, growth, energy metabolism, and amino acid and carbohydrate transport. A prominent trend observed for all three strains were protein abundance shifts associated with the categories "Cell cycle control, cell division, chromosome partitioning" and "Cell wall/membrane/envelope biogenesis” showing a lower relative abundance, hereafter referred to as “downregulation”, at atmospheric CH₄ compared to the 1000 p.p.m.v. CH₄ treatment. The observed pattern suggests that atmMOB lower the allocation of resources for growth when CH₄ availability is low, which is in line with previously reported differences in colony growth over time when comparing incubations on atmospheric and 1000 p.p.m.v. CH₄ concentrations²⁸. We observed major shifts in protein abundances assigned to the categories “Energy production and conversion” and “Carbohydrate transport and metabolism", including proteins involved in trace gas oxidation and carbon assimilation (Supplementary tables S12-14). These shifts suggest that the three atmMOB strains allocate proteome investments towards catabolic processes, and away from anabolic processes, when living on trace concentrations of CH₄. Based on that and the different trace gas oxidation patterns, we further continued to investigate the relative abundance of proteins involved in trace gas oxidation, carbon assimilation via the serine cycle (Fig. 4), and the electron transport chain (Supplementary Fig. 4).

Comparative proteomics: Trace gas oxidation

The differences in enzyme expression patterns related to trace gas oxidation indicate that the three strains, M. gorgona MG08 and M. rosea SV97 and M. palsarum NE2 employed different metabolic strategies to grow on air. All three strains expressed a particulate methane monooxygenase (pMMO) that catalyzes the hydroxylation of CH₄ to methanol (CH₃OH). M. gorgona MG08 and M. rosea SV97 contained higher relative abundances, hereafter referred to as “upregulation”, of pMMO at 1000 p.p.m.v. CH₄, whereas M. palsarum NE2 upregulated pMMO at atmospheric CH₄ concentrations. Furthermore, both M. gorgona MG08 and M. rosea SV97 upregulated a high affinity [NiFe] hydrogenase (Hhy), and M. gorgona MG08 a molybdenum-dependent carbon monoxide dehydrogenase (CODH), when exposed to air. These differences in enzyme expression patterns demonstrate different metabolic strategies to grow on air: M. rosea SV97 and M. gorgona MG08 compensate for energy limitation by upregulating enzymes for energy conservation from H₂ or H₂ and CO, while M. palsarum NE2 compensates for the limitation by upregulating pMMO for CH₄ consumption. The upregulation of pMMO, Hhy, or CODH in the three strains suggests that increases in the ${a}_{A}^{0}$ for trace gases are an adaptation to growth on air.

Despite the ability of M. rosea SV97 and M. palsarum NE2 to oxidize CO (Fig. 1A), we were not able to determine the responsible protein complex(es) or the corresponding gene expression responses. In M. rosea SV97, candidate genes for CODH were identified by blasting the genome sequences against a non-redundant CODH sequence database created by using the Identical Protein Groups resource⁴³ (Supplementary table S11). However, not all the potential CODH subunits necessary to form a functional CODH were detected in the proteomes. The same blast-based approach did not result in potential candidate genes that could encode a functioning CODH in M. palsarum NE2 (Supplementary table S10). Thus, our results indicate that distantly related or previously undiscovered enzymes catalyze atmospheric CO oxidation in M. palsarum NE2.

The expression of the methanol dehydrogenase (MDH), which catalyzes the second step in CH₄ oxidation, namely the oxidation of methanol (CH₃OH) to formaldehyde (CH₂O), matched the pMMO expression patterns for all three strains. This indicates a close interaction between these two enzymes as previously suggested for Methylococcus capsulatus⁴⁴. MDHs were upregulated at high CH₄ concentrations by M. gorgona MG08 and M. rosea SV97, but not by M. palsarum NE2 (Fig. 4). Only the putative lanthanide-dependent methanol dehydrogenase (XoxF) of M. palsarum NE2 did not follow the pMMO pattern.

Formaldehyde is a key intermediate of both catabolism and anabolism in methanotrophs. The enzymes involved in the catabolic oxidation of formaldehyde via methylene-tetrahydromethanopterin (-H₄MPT), methenyl-H₄MPT, and formyl-H₄MPT to formate (CHOO⁻), were upregulated in the high CH₄ treatment of the three strains (Fig. 4). The toxic and highly reactive formaldehyde condenses spontaneously to methylene-H₄MPT⁴⁵ and methylenetetrahydrofolate (H₄F)⁴⁶ and thus, higher concentrations of formaldehyde activating enzymes (Fae) may not be required for the oxidation of formaldehyde during growth on air. At high CH₄ concentrations, however, the upregulation of Fae and downstream enzymes catalyzing the further oxidation to formate could represent a cellular detoxification mechanism to avoid high cellular formaldehyde concentrations as well as increasing the rate of NADH generation⁴⁷.

Two different formate dehydrogenases (Fdh A and Fdh AB) that catalyze the energy-conserving oxidation of formate to CO₂ were expressed by M. gorgona MG08 and M. palsarum NE2 (Fig. 4). The metal-free and potentially irreversible NAD⁺-dependent Fdh A^31,48 was upregulated at atmospheric CH₄ concentrations in M. gorgona MG08 and M. palsarum NE2. Possibly, the upregulation of a one-directional enzyme minimizes back-flow of CO₂ to formate at high CO₂ concentrations and thus prevents energy loss by CO₂ reduction. Alternatively, a high affinity for formate could explain the upregulation of the Fdh A at atmospheric concentrations, as it would allow an efficient conversion of formate at very low intracellular concentrations. However, low-molecular, metal-free Fdhs are assumed to have a low affinity for formate⁴⁹. Fdh AB, which was expressed by all three strains, is molybdopterin-dependent and homologous to the reversible Fdh found in Rhodobacter capsulatus⁵⁰. It catalyzes, in addition to formate oxidation, the reduction of CO₂ to formate^50,51. The Fdh AB was upregulated in all three strains at high CH₄ concentration. Under these conditions, when excess reducing power is available, the Fdh AB might enable the reduction of CO₂ for carbon assimilation via the reductive glycine pathway and serine cycle, but further investigations are needed to test whether CO₂ reduction to formate truly occurs in atmMOB.

Comparative proteomics: Carbon assimilation

Carbon assimilation was downregulated in all three strains at atmospheric CH₄ concentrations. In the three strains, formaldehyde can condense with H₄F to methylene-H₄F and then be further assimilated through the glycine cleavage system and serine pathway. Additionally, formaldehyde can be first oxidized via the H₄MPT-mediated pathway to formate and then enter, instead of being oxidize to CO₂, the H₄F-mediated reductive pathway to methylene-H₄F (Fig. 4). The increased expression of enzymes involved in the reduction of formate via formyl-H₄F and methenyl-H₄F to methylene-H₄F in the high CH₄ treatment indicates an enhanced investment into carbon assimilation. Alternatively, since formaldehyde spontaneously condenses with H₄F to methylene-H₄F, the upregulated enzymes for the H₄F-mediated formaldehyde oxidation could also contribute to formate formation. However, experiments with Methylobacterium extorquens demonstrated that formaldehyde oxidation occurred through its H₄MPT-mediated pathway while the reductive pathway via formate to methylene-H₄F represented the major assimilatory flux^47,52. Therefore, we consider this as the most likely explanation for the expression patterns in M. gorgona MG08, M. rosea SV97, and M. palsarum NE2.

The glycine cleavage/synthase system (GCS)^53,54, consisting of four proteins, catalyzes the oxidative cleavage of glycine to NADH, NH₃, CO₂, and a methylene group. It also catalyzes the reverse reaction, the synthesis of glycine from NADH, NH₃, CO₂, and methylene-H₄F. The upregulation of the GCS by M. rosea SV97 and M. palsarum NE2 at high CH₄ concentrations suggests an increased synthase activity and thus increased carbon assimilation, corresponding to the overall upregulation of the reductive glycine pathway (Fig. 4). The increased expression of the GCS by M. gorgona MG08 at atmospheric CH₄ concentrations is less intuitive and we lack a plausible explanation for this upregulation.

As the next step in carbon assimilation, the bidirectional serine hydroxymethyltransferase (GlyA) condenses glycine with methylene-H₄F to serine, representing the first step of the serine cycle (Fig. 4). The enzymes involved in the serine cycle were mainly upregulated by the strains when exposed to high CH₄ concentrations, further indicating more investment into carbon assimilation at this condition.

Comparative proteomics: Electron transport chain

The relative abundances of protein complexes involved in the electron transport chain for ATP synthesis was lower in atmospheric CH₄ compared to the high CH₄ treatment (Supplementary Fig. 4). The NADH-quinone oxidoreductase, cytochrome c oxidase, and the ATP synthase were all highly expressed at high CH₄ concentrations indicating an increased investment into energy conservation at high substrate supply. Only the ubiquinol-cytochrome c reductase expression did not follow a clear pattern. The expression patterns for electron transport matches the patterns for carbon assimilation (Supplementary Fig. and Fig. 4). We propose that to overcome energy limitations when exposed to air, all three strains upregulate the expression of enzymes for the oxidation of at least one trace gas to maximize uptake rates and energy yield, while investments in energy conservation and energy-intensive carbon assimilation are reduced. This reduced investment in assimilation is in line with the low concentrations of the trace gases, low uptake rates, and overall slow growth of atmMOB when incubated with air as energy and carbon source³¹.

Specific affinity

The ${a}_{A}^{0}$ for CH₄, expressed as the fraction of V_max(app) and K_m(app), directly represents the capacity a methanotroph to oxidize CH₄ at low concentrations. Here, we show that the specific affinities of the atmMOB M. gorgona MG08 and M. palsarum NE2 are the highest measured for CH₄.

To test if the pMMO upregulation in M. palsarum NE2 and downregulation in M. gorgona MG08 at atmospheric CH₄ concentrations translate into a higher ${a}_{A}^{0}$ for CH₄ by M. palsarum NE2 compared to M gorgona MG08, we measured the ${a}_{A}^{0}$ of the two strains when grown on air. To avoid increases in the apparent half saturation constants K_m(app)⁵⁵ at high CH₄ concentrations, we pre-incubated the cultures on filters floating on carbon-free medium for five months with air as sole energy and carbon source.

By measuring the CH₄ oxidation per cell and hour at different CH₄ concentrations, we estimated a K_m(app) of 48.54 nM CH₄ and a V_max(app) of 4.91 x 10^− 8 nmol cell^− 1 h^− 1 resulting in a ${a}_{A}^{0}$ of 1.01 x 10^− 9 L cell^− 1 h^− 1 for M. gorgona MG08 (Supplementary Fig. 5). A K_m(app) of 48.54 nM CH₄ is within the range of the K_m(app) values measured for fresh oxic soils reported by Bender and Conrad (30–51 nM)¹¹, which led to the theory that atmMOB are oligotrophs with a high affinity for CH₄. Our observation is the first observation of a methanotroph in pure culture showing a K_m(app) that is in the range of the low K_m(app) values measured in upland soils^23,56. Our ${a}_{A}^{0}$ estimate for M. gorgona MG08 (1.01 x 10^− 9 L cell^− 1 h^− 1) is approximately five times higher than the ${a}_{A}^{0}$ reported by Tveit et al. 1.95 ×10^− 10 L cell^− 1 h^{− 1 31}. However, the K_m(app) and V_max(app) reported by Tveit et al. amount 4905 nM and 95.4 x 10^− 8 nmol cell^− 1 h^− 1, respectively, values approximately 100 and 20 times higher than in the current study. These differences might derive from the use of liquid cultures pre-incubated at 20% CH₄ by Tveit et al. Such a high CH₄ concentration could have influenced the cellular pMMO concentration as indicated by the pMMO upregulation at 1000 p.p.m.v. CH₄ described above (Fig. 4). In this study, we have used filter cultures acclimated to atmospheric CH₄ concentrations, resulting in lower cellular pMMO concentrations in M. gorgona MG08, when compared to 1000 p.p.m.v. (Fig. 4), and thus possibly leading to the lower K_m(app) and V_max(app) for CH₄ observed. Dunfield and Conrad reported a similar alteration of K_m(app) and V_max(app) for Methylocystis strain LR1 after comparing starved cells to cells exposed to 10% CH₄ while the ${a}_{A}^{0}$ was more constant⁵⁵.

The K_m(app) of M. palsarum NE2 was 402.08 nM CH₄, which is approximately 8 times higher than the K_m(app) of M. gorgona MG08. Despite this higher K_m(app), the ${a}_{A}^{0}$ of M. palsarum NE2 was 3.30 x 10^− 9 L cell^− 1 h^− 1, three times higher than estimated for M. gorgona MG08. This high ${a}_{A}^{0}$ derives from its substantially higher V_max(app) of 133 x 10^− 8 nmol cell^− 1 h^− 1 (Supplementary Fig. 5). An apparent affinity of 402.08 nM CH₄ is not considered as high affinity ^26,40. Thus, since both strains grow on air, a high apparent affinity for CH₄ is not a prerequisite for this lifestyle. This is also reflected in the CH₄ oxidation rate of M. palsarum NE2 at atmospheric concentrations, which surpassed the rate of M. gorgona MG08, despite having a lower apparent affinity for CH₄ (Supplementary Fig. 5). The ${a}_{A}^{0}$ of M gorgona MG08 and M. palsarum NE2 are approximately equally high and three times higher, respectively, than the recently reported ${a}_{A}^{0}$ of Methylotuvimicrobium buryatense 5GB1C⁵⁷ and 30 and 100 times higher, respectively, than the ${a}_{A}^{0}$ of Methylocystis sp. SC2 (3.4 × 10^− 11 L cell^− 1 h^− 1) ²⁴, the MOB with the fourth highest ${a}_{A}^{0}$ for CH₄ known so far. However, the different experimental setups and CH₄ concentrations used to determine ${a}_{A}^{0}$ might render the comparisons invalid^24,57. Nevertheless, our results indicate that⁵⁸ the specific affinity (${a}_{A}^{0})$, rather than the affinity (K_m(app)), is the suitable measure to determine the capacity of methanotrophs to oxidize atmospheric CH₄, as previously suggested for oligotrophic substrate uptake at low concentrations by Button⁵⁸,. Thus, a high specific affinity for CH₄ and not a high affinity is the key prerequisite for methanotrophs to grow on air.

N₂-fixation at atmospheric trace gas concentrations

The four atmMOB encode all genes required for dinitrogen (N₂) fixation³¹ and grow in nitrogen-free medium at high CH₄ concentrations^32,33,35,59. To test the potential for N₂-fixation during growth on air, we incubated filter cultures of M. gorgona MG08, M. palsarum NE2, M. rosea SV97, and M. aurea KYG with air as the sole energy, carbon, and nitrogen source. The colony formation of all four strains after three months demonstrates growth in the absence of bioavailable nitrogen sources in the medium (Fig. 5A). This suggests that all four strains can either cover their nitrogen requirements by the fixation of N₂ or by the utilization of atmospheric reactive nitrogen during growth on air. Ammonia concentrations in air, for example, have been observed to range between 0.2 and 24 p.p.b.v.⁶⁰.

To test for N₂ fixation during growth on air by M. gorgona MG08, we incubated filter cultures in air enriched with ¹⁵N-N₂ (~ 23 atom % (at%) of the total N₂) and CH₄, H₂, and CO concentrations fluctuating between 0.03 and 3 p.p.m.v. for two months. Afterwards, we measured cellular ¹⁵N₂ fixation using NanoSIMS (Fig. 4C). The cellular ¹⁵N ranged from 2.99 at% to 61.89 at% with an average of 30.79 at% (SD = 7.82 at%) (Supplementary table S15) while the control without ¹⁵N₂ enrichment averaged at 0.37 at% (SD = 0.04) (Supplementary table S16). Since the ¹⁵N₂ in the headspace amounted to approximately 23 at% of the total N₂ during incubation, the ¹⁵N average of the cells should not have amounted to more than 23 at%. The high values might issue from bioavailable ¹⁵N-species contaminating the ¹⁵N₂ gas used for incubation⁶¹. Purity tests of the ¹⁵N₂ gas prior to the ¹⁵N₂ fixation experiments revealed only a minor NO_x contamination of 101.59 (nmol ml^− 1) with a very low ¹⁵N fraction of 0.0014 at% and ammonia levels below detection limit (1 nmol ml^− 1). However, even trace amounts of ¹⁵N ammonia contaminating the ¹⁵N₂ are sufficient to cause high cellular ¹⁵N-enrichments given the low amount of biomass of the filter cultures and the long incubation times. Thus, to verify N₂ fixation by atmMOB, extremely pure ¹⁵N₂ is required. However, we are currently not able to detect sufficiently low concentrations of ammonia or other potential reactive nitrogen contaminants to exclude that the high cellular ¹⁵N has originated from other sources than ¹⁵N₂. Considering the growth of all four strains on nitrogen-free medium and the enduring trace gas oxidation of M. gorgona MG08, the atmMOB are either extremely oligotrophic and can cover their nitrogen requirements solely by deposited reactive nitrogen, or they fix N₂ when growing at atmospheric concentrations of CH₄, H₂, and CO. Either way, atmMOB seem to harness the required nitrogen, like carbon and energy, directly from the air.

Enduring oxidation of atmospheric CH₄ is not restricted to members of the clades USCα and γ, but more widespread than previously assumed. Our data shows that former liquid culture-based cultivation approaches lead to an underestimation of the true potential of methanotrophic pure cultures to oxidize atmospheric CH₄. The appearance of atmMOB outside the USCα and γ revises our understanding of the biological atmospheric CH₄ sink and should be considered in future studies. The strain specific oxidation patterns and the estimations of the ${a}_{A}^{0}$ for CH₄ demonstrate that both the mixotrophic oxidation of atmospheric trace gases and a high ${a}_{A}^{0}$ for CH₄ are key to obtain sufficient energy for growth on air. The estimated energy requirements for growth of the four atmMOB are substantially lower than the maintenance energy value used as basic premise for an oligotrophic lifestyle of MOB. Additionally, atmMOB seem to cover not only their energy and carbon but also their nitrogen requirements from the atmosphere. This opens a new perspective on physiological limitations of atmospheric trace gas oxidizers and suggests that atmMOB may carry an ideal set of properties needed for pioneering species to initiate primary succession in unfavorable environments. Additionally, the high ${a}_{A}^{0}$ for CH₄ enables atmMOB to utilize trace concentrations of CH₄ as energy and carbon source for growth while being extremely oligotrophic. This bears the potential for cost-effective and efficient biofiltration of anthropogenic emissions containing CH₄ concentrations far below the lower explosive limit. The common metabolic strategy to grow on air seems to be the downregulation of enzymes involved in energy-intensive processes combined with the upregulation of enzymes for the oxidation of trace gases. However, the differing expression patterns of enzymes for trace gas oxidation and the strain-specific trace gas oxidation patterns indicate that a diverse metabolic repertoire has evolved to enable life on air.

Acknowledgements

We thank Alena Didriksen from the University of Tromsø for her involvement in cell quantification, Toril Anne Grønset and Jack-Ansgar Bruun from PRiME for their contributions to the comparative proteomics analysis and Cornelia Rottensteiner, Judith Prommer, and Margarete Watzka from the University of Vienna for the carbon content analysis and ¹⁵N₂ gas purity test.

Funding

This study was supported by the Research Council of Norway FRIPRO Young Researcher Grant 315129, Living on Air, and Tromsø Research Foundation starting grant project Cells in the Cold 17_SG_ATT to ATT. Mass spectrometry-based proteomic analyses were performed by UiT Proteomics and Metabolomics Core Facility (PRiME). This facility is a member of the National Network of Advanced Proteomics Infrastructure (NAPI), which is funded by the Research Council of Norway INFRASTRUKTUR-program (project number: 295910).

Authors contributions

T.S., M.M.S. and A.T.T. conceived the study. T.S., A.G.H., J.B., H.S., A.Sch., B.R., O.S., and A.T.T. performed experiments. T.S., H.S., A.Sch., A.S., A.R. and A.T.T. analyzed data. T.S. and A.T.T. created the figures. T.S., H.S., A.Sch., B.R., A.R., A.S., M.P, M.M.S, and A.T.T. contributed to methods. T.S. and A.T.T. wrote the manuscript with inputs from all authors.

Competing interests

The authors declare no conflict of interest.

Data availability

The proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE⁶² partner repository with the dataset identifier PXD046190. The data used in this study are provided in the excel file “Supplementary tables”.

Filter-based cultivation with air as the sole carbon and energy source

The strains were pre-incubated in liquid cultures in 100 mL serum bottles containing 10 mL of 10x diluted, EDTA-free NMS medium (DSMZ medium 921 with 10x the iron concentration and 1 µM lanthanum) and a headspace of 20% CH₄ in air. The Fe-EDTA stated in the original DSMZ recipe was substituted with FeSO₄. Depending on the strain, the medium was adjusted to a pH of 6.8 (Methylocapsa gorgona MG08, Methylosinus trichosporium OB3b, Methylocystis rosea SV97, Methylobacter tundripaludum SV96) or 5.8 (Methylocapsa aurea KYG, Methylocapsa acidiphila B2, and Methylocapsa palsarum NE2). The serum bottles were sealed using butyl rubber stoppers (Chromacol 20-B3P, Thermo Fisher Scientific, Waltham, Massachusetts, USA) and crimp caps, and incubated at 20°C in dark, until the strains were in an exponential growth phase. Strain purity was controlled routinely via microscopy and by confirming the absence of heterotrophic growth on agar plates with a rich medium containing tryptone, yeast extract, and glucose³⁰. After reaching exponential growth phase, the strains were diluted with the described medium and filtered on 25 mm polycarbonate (PC) filters (Whatman 10417006, Cytiva, Massachusetts, Marlborough, USA) using a filtration manifold (EQU-FM-10X20-SET, DHI, Hørsholm, Denmark)²⁸. The final cell density on the filters amounted approximately 20 cells per photo area (630x magnification). The filters were transferred into Petri dishes containing 10x diluted, EDTA-free NMS medium (DSMZ medium 921 with 10x the iron concentration and 1 µM lanthanum) and left floating on the medium with the cells facing upwards. The cultures were incubated at 20°C in dark, with air as the sole energy and carbon source, for at least three months until initiation of experiments. All cultivation steps were carried out under sterile conditions. Before experiments, colony formation and microcolony morphology of the strains was routinely checked via microscopy as previously described³¹.

Trace gas oxidation experiments

For all trace gas oxidation experiments, filter cultures of the respective strains, pre-incubated with air as the sole carbon and energy source, were transferred into 250 mL glass bottles containing 50 mL 1:10 EDTA-free NMS medium (DSMZ medium 921 with 10x the iron concentration and 1 µM lanthanum). The bottles were capped with Safety-Caps (JR-S-11011, VICI AG International, Schenkon, Switzerland). To create a defined atmosphere within the bottles, the headspace of each bottle was flushed for 10 minutes with high-purity synthetic air containing 400 p.p.m.v. CO₂ (HiQ, AGA, Sweden) using a gassing manifold. The headspace pressure was adjusted to approximately 1.05 bar. Then, 1 mL 1000 p.p.m.v. CH₄ in N₂ (HiQ, AGA, Sweden), 1 mL 1000 p.p.m.v. H₂ in N₂ (HiQ, AGA, Sweden), and 1 mL 1000 p.p.m.v. CO in N₂ (HiQ, AGA, Sweden) were added to the headspace using a gas tight syringe. To assess the oxidation rate of the respective strains, the cultures were incubated at 20°C and the change in CH₄, H₂, and CO concentrations within bottle headspaces were measured. For each measurement, 2 mL of the gas in the headspace were sampled using a gas tight syringe. The gas samples were analyzed with a gas chromatograph (ThermoScientific Trace 1300, Thermo Fisher Scientific) equipped with a sample loop, a Hayesep column (SU12875, RESTEK, Bellefonte, Pennsylvania, USA), a Molsieve 5A column (PKC17080, RESTEK), a pulsed discharge detector, and a flame ionization detector. A high-quality gas containing 2.5 p.p.m.v. CH₄, 2.5 p.p.m.v. H₂, and 2.5 p.p.m.v. CO in N₂ served as standard (HiQ, AGA, Sweden). To determine the gas concentrations in the bottle headspaces, standard curves were created every day of measurement. The headspace pressure was measured at the end of the experiment using a manometer (LEO1, Keller, Winterthur, Switzerland). The mass of each trace gas was calculated by applying the ideal gas law and adjusted for changes in pressure caused by gas removal during measurement.

Cell quantification

For quantification, cells from filters were processed as follows: The filters were placed on 150 µL 25x SYBR green I (10463252, Thermo Fisher Scientific) with cells facing upwards and incubated for 10 minutes in dark. Next, the cells were washed twice by letting the filters float on milliQ water for 5 minutes in dark. To remove the cells from the filters, the filters were transferred into 5 mL tubes (0030122321, Eppendorf, Hamburg, Germany) containing 2 mL 10x diluted solution 1 (DSMZ medium 921), so that the filters sticked to the tube wall and the cells faced inwards⁶³. The tubes were vortexed for 10 minutes at medium speed. After vortexing, remaining cells on the filters were washed off and collected by rinsing the filters with 1 mL solution 1 (DSMZ medium 921). The washed filters were dried in dark, and the cell removal controlled via fluorescence microscopy. The 3 mL cell suspension resulting from the cell removal was spiked with 50 µL absolute counting beads (Invitrogen^tm CountBright^tm Plus, Thermo Fisher Scientific). After, the cells in suspension were immediately counted using a flow cytometer (BD FACSAria III, BD Biosciences, Franklin Lakes, New Jersey, USA). The flow cytometer was set up to capture SYBR Green I (excitation = 498 nm, emission = 522 nm) in the green channel and the Invitrogen^tm CountBright^tm Plus Absolute Counting Beads (excitation = 350–810 nm, emission = 385–860 nm) in the blue channel. Unstained cells of M. gorgona MG08, M. palsarum NE2, M. rosea SV97, and M. aurea KYG were used to define the autofluorescence threshold in the different channels. Size beads (NPPS-4K, Spherotech, Lake Forest, Illinois, USA) were used to draw the forward scatter gate covering events between 0.58 and 4 µm. SYBR green I stained cells and absolute counting beads were counted to determine the cell number within filter cultures, using the following equation:${x}_{filter}=\frac{{b}_{abs}}{{b}_{c}} \times {x}_{c}$

Where ${x}_{filter}$ is the absolute cell number of the filter culture, ${b}_{abs}$ the number of counting beads added, ${b}_{c}$ the number of counting beads counted, and ${x}_{c}$ the number of SYBR green I stained cells counted.

Cell-specific oxidation rates and energy calculation

The gas-specific oxidation rates of the different strains at atmospheric CH₄, H₂, and CO concentrations and pseudo first order kinetics were calculated and tested as described by Tveit et al³⁰. The oxidation rate per cell was calculated by dividing the oxidation rate of the filter culture by the corresponding cell number. The Gibbs free energy changes (${\varDelta }_{r}G$) for the following reactions: ${CH}_{4}+2{O}_{2}\to 2{H}_{2}O+C{O}_{2}$, ${2H}_{2}+{O}_{2}\to 2{H}_{2}O$, $2CO+{O}_{2}\to 2C{O}_{2}$, at 20°C, 1.013 bar absolute, and atmospheric concentrations of CH₄ (1.87 p.p.m.v.), H₂ (0.5 p.p.m.v.), and CO (0.2 p.p.m.v.) amount to -797.4 kJ mol⁻¹, -236.8 kJ mol⁻¹, and − 199.9 kJ mol⁻¹ respectively. The values are based on the values for Gibbs free energy of formation found in literature^30,64 and the following equation:

${\varDelta }_{r}G= {\varDelta }_{r}{G}^{^\circ }+RTln{Q}_{r}$

where ${\varDelta }_{r}{G}^{^\circ }$ is the Gibbs free energy change at standard conditions, $R$ the gas constant, $T$ the temperature, $ln$ the natural logarithm, and ${Q}_{r}$ the reaction quotient. The estimates of energy yield per cell were obtained from the trace gas oxidation per cell and the ${\varDelta }_{r}G$. The energy yield from trace gas oxidation per mol of biomass and hour (kJ C-mol⁻¹ hour⁻¹) was estimated including the strain specific carbon content and dry weight as previously shown²⁴.

Cellular dry mass and carbon content estimations

The strain-specific cellular dry mass was determined by measuring single-cell buoyant mass distributions of the strains in H₂O-based and deuterium oxide (D₂O)-based solutions of phosphate saline buffer (PBS) using a suspended microchannel resonator (SMR)^65,66. To do so, filter cultures of M. gorgona MG08, M. rosea SV97, M. palsarum NE2, and M. aurea KYG were incubated on 10x diluted, EDTA-free NMS medium (DSMZ medium 921 with 10x the iron concentration and 1 µM lanthanum) under an atmosphere of 1000 p.p.m.v. CH₄ in air for three weeks. After, cells of filter cultures were fixed by incubating the filters on 150 µl formaldehyde in 1×PBS (4% w/v) for an hour at room temperature. The cells were washed twice by letting the filters float on water for 5 minutes. The fixed cells were harvested as described in “Cell quantification”. The resulting cell suspension was stored at 4°C until measurements of the buoyant cell mass. For the measurements, two aliquots with the same volume of each cell suspension were created. The water of the aliquots was replaced by H₂O-based (1x PBS in H₂O) and D₂O-based (1x PBS in 9:1 D₂O:H₂O) solutions of known density (1.0043 g cm^− 3 and 1.1033 g cm^− 3 at 20°C, respectively). To do so the aliquots were dried in a vacuum concentrator at 37°C. After, the cells of one of the two aliquots were resuspended in 50 µl of the H₂O-based solution. The cells of the second aliquot were resuspended in 50 µl of the D₂O-based solution. The buoyant mass of the cells in the aliquots were measured with a SMR (LifeScale, Affinity Biosensors, Santa Barbara, California, USA). The precision and accuracy of the SMR was verified by creating calibration curves using NIST-certified polystyrene beads (ThermoFisher Scientific) as performed previously⁶⁷.

The buoyant mass data were exported from the LifeScale instrument and further analyzed using the Python 3⁶⁸ packages Pandas⁶⁹, Matplotlib⁷⁰, and Seaborn^68,71 (Supplementary Fig. 8). The dry mass of the strains was calculated as described⁶⁵ using the median of the single-cell buoyant mass distributions in H₂O-based and D₂O-based solutions and the following equation:

$${m}_{dry}= \frac{{\rho }_{{D}_{2}O} \times {m}_{{b.H}_{2}O}-{\rho }_{{H}_{2}O} \times {m}_{{b.d}_{2}O} }{{\rho }_{{D}_{2}O}- {\rho }_{{H}_{2}O}}$$

Where ${m}_{dry}$ is the dry mass, ${\rho }_{{D}_{2}O}$ the density of the D₂O-based solution, ${m}_{{b.H}_{2}O}$ the cell’s buoyant mass in the H₂O-based solution, ${\rho }_{{H}_{2}O}$ the density of the H₂O-based solution, and ${m}_{{b.d}_{2}O}$ the cell’s buoyant mass in the D₂O-based solution.

The cellular carbon contents of the atmMOB were analyzed using an elemental analyzer coupled to an isotope ratio mass spectrometer (EA-IRMS; EA1110 coupled via a ConFlo III interface to a DeltaPLUS IRMS, Thermo Fisher Scientific). Biomass for the carbon content analysis was derived from stirred-tank bioreactor (DASbox® Mini Bioreactor System, Eppendorf) cultures. The cultures were grown in 10x diluted, EDTA-free NMS medium (DSMZ medium 921 with 10x the iron concentration and 1 µM lanthanum) at 80 rpm (marine impeller), 20°C, pH 6.8 (CO₂ controlled) and a gassing rate of 0.24 vessel volumes per minute (6000 p.p.m.v. CH₄ in air) using a microsparger (78530205, Eppendorf). After harvest, cultures were washed three times with milliQ water and lyophilized.

Comparative proteomics

For the atmospheric CH₄ treatment, filter-cultures of M. gorgona MG08, M. rosea SV97 and M. palsarum NE2 were incubated with air as the only carbon and energy source for seven months. After, the cells on filters were harvested as described in “Cell quantification”, lyophilized, and stored at -80°C until further processing. The 1000 p.p.m.v. CH₄ treatment of the respective strains was processed the same way as the atmospheric CH₄ treatment with the difference that filter cultures had been exposed to approximately 1000 p.p.m.v. CH₄ in air for two weeks before harvest. Samples of lyophilized cells were lysed by sonication in 20 µL buffer containing 4 M urea, 2.5% sodium deoxycholate (SDC) and 100 mM triethylammonium bicarbonate (TEAB). Samples were sonicated for 25 cycles (1 min on, 30 sec off) with maximum amplitude in a cup horn sonicator with a recirculating chiller (Cup horn: model 413C2, Qsonica, Newtwon, Connecticut, USA; Sonicator: Fisherbrandtm FB705, Thermo Fisher Scientific; Recirculating chiller: model 4905 Qsonica). Then, disulfide bridges were reduced with 1,4-dithiothreitol (DTT) at a final concentration of 5 mM and incubation at 54°C for 30 min. Cysteines were alkylated with 15 mM iodoacetamide (IAA) and incubated for 30 min at room temperature in dark. To remove excess IAA, DTT solution corresponding to a final concentration of 5 mM was added. Calcium chloride solution (final concentration of 1 mM) and 1 µg Lysyl Endopeptidase (125–05061, FUJIFILM Wako Chemicals Europe, Neuss, Germany) were added to the samples and incubated for 5 hours under gentle agitation at 37°C for enzymatic digestion. After, samples were diluted with a buffer containing 100 mM triethylammonium bicarbonate (TEAB) and 1 mM CaCl₂ to lower the urea and sodium deoxycholate (SDC) concentration to 1 M and 0.65% v/v, respectively, resulting in a final sample volume of 80 µL. For digestion, 2 µg trypsin (V511A, Promega, Wisconsin, USA) were added and the samples incubated on a gently agitated shaker at 37°C for 16 hours. After digestion, SDC was precipitated by adding 50% formic acid to the sample (final concentration of 2.5% v/v). Samples were then incubated for 10 min and centrifuged at 16200 rcf for 15 min. Supernatants containing peptides were transferred to low-protein-binding tubes. The peptides were concentrated and cleaned up using DPX C18 pipette tips (DPX Technologies, XTR tips 10 mg C18AQ 300Å) on a Tecan Fluent pipetting robot (Tecan Group Ltd., Männedorf, Switzerland). Purified peptide samples were dried in a vacuum concentrator and dissolved in 12 µL 0.1% formic acid. Peptide concentrations were measured on a spectrophotometer (Nanodrop ONE, Thermo Fisher Scientific) at 205 nm. 0.25 µg peptides per sample were loaded onto a liquid chromatograph (EASY-nLC1200, Thermo Fisher Scientific) equipped with an EASY-Spray column (C18, 2µm, 100 Å, 50µm, 50 cm). Peptides were fractionated using a 5–80% acetonitrile gradient in 0.1% formic acid over 120 min at a flow rate of 300 nL min^− 1. The separated peptides were analyzed using a mass spectrometer (Orbitrap Exploris 480, Thermo Fisher Scientific). Data was collected in data dependent mode using a Top40 method. Annotated genomes of the three strains downloaded from MicroScope⁷² served as databases for the CHIMERYS-based data search using Proteome Discoverer 3.0. Normalized abundances (scaling mode: On All Average) of proteins were further processed via Perseus⁷³. Normal distribution of Log2 fold transformed data was visually screened using histograms. Proteins were filtered using a threshold of at least three valid values in at least one treatment (n = 4 per treatment). Proteins that passed the filtering were included in downstream analyses. Missing values were imputed from normal distribution using default settings (width = 0.3, down shift = 1.8). The Pearson correlation coefficient ranged from ρ = 0.918 to ρ = 0.989 between replicates and from 0.68 to 0.799 between replicates of the different treatments. Imputed protein abundance results of the 1.9 p.p.m.v. and 1000 p.p.m.v. treatments were used for correspondence analysis (CA). CA was conducted in R⁷⁴ using the “ca” function of R package “ca”⁷⁵. The top 10% of proteins that contributed most to the inertia of the CA’s first dimension were extracted using the function “get_ca_row” of the R package “factoextra”⁷⁶. To map the abundances and the hierarchical EggNOG⁴² annotations of the top 10% proteins, the EggNOG annotations of the three strains were downloaded from MicroScope. For the in-depth analysis of trace gas oxidation, carbon assimilation, and the electron transport chain, the differences between treatments were tested using a two-sided t-test (s0 = 2) and permutation-based false discovery rate (FDR = 0.1). From the z-score normalized results, the proteins involved in trace gas oxidation, carbon assimilation via the serine cycle, and the electron transport chain were selected for further analysis using Python 3⁶⁸. The core carbon and energy metabolism of M. rosea SV97 and M. palsarum NE2 were reconstructed manually using MicroScope annotations, the published metabolism entries of M. gorgona MG08³¹, KEGG⁷⁷, and protein BLAST searches⁷⁸. To conduct a thorough screening for putative carbon monoxide dehydrogenase subunits a blast-searchable database was constructed out of all sequences retrieved from the NCBI Identical Protein Groups resource⁴³ (accessed May, 30th 2023) using the search term “carbon monoxide dehydrogenase” (54076 amino acid sequence entries). Entries with an amino acid sequence length < 30 were removed from the database prior to the blastp search. The annotated genomes of M. rosea SV97 and M. palsarum NE2, downloaded from MicroScope, were used as blastp queries (default settings, -outfmt 6 including slen). After a pre-filtering removing hits with an E-value > 1 and an alignment length < 50% of the aligned database entry, the blastp output tables were manually evaluated (Supplementary table S10-11).

Specific affinity

Filter cultures of M. gorgona MG08 and M. palsarum NE2 were pre-incubated on 10x diluted, EDTA-free NMS medium (DSMZ medium 921 with 10x the iron concentration and 1 µM lanthanum) for five months with air as sole energy and carbon source. The filter cultures were transferred into 250 mL glass bottles containing 250 mL of the mentioned medium. The glass bottles were capped with Safety-Caps (JR-S-11011, VICI AG International). For each strain, the CH₄ concentrations in the 50 mL headspace were adjusted to approximately 1.9, 14, 30, 70, and 175 p.p.m.v. by adding 0–2.5 mL 1000 or 5000 p.p.m.v. CH₄ in N₂ (HiQ, Linde, Sweden). The headspace pressure of all bottles was adjusted to approximately 1.1 bar using air. After, the strains were incubated for 48 hours at 20°C. The change in CH₄ was measured at incubation start, after 24 hours and 48 hours as described above. Standard curves were created using 2.5 and 50 p.p.m.v. CH₄ in N₂ (HiQ, AGA, Sweden). At the end of the oxidation experiment, cells in filter cultures were quantified as described above. The change in mass of CH₄ in the headspace was calculated by applying the ideal gas law and adjusted for changes in pressure caused by gas removal during measurement. The mass of dissolved CH₄ at different CH₄ partial pressures in the headspace was calculated by applying the Henry’s law solubility constant for CH₄ at 20°C. The Michaelis-Menten CH₄ oxidation kinetics were modelled using the “nls” function of the “nlstools” R package⁷⁹, specifying the “michaelis” model and providing start values for K_m(app) and V_max(app). The ${a}_{A}^{0}$ was calculated by dividing V_max by K_m.

N₂ fixation at atmospheric trace gas concentrations

Filter cultures of M. gorgona MG08, M. palsarum NE2, M. rosea SV97, and M. aurea KYG were pre-incubated on 10x diluted, EDTA-free, and potassium nitrate (KNO₃)-free NMS medium (DSMZ medium 921 with 10x the iron concentration and 1 µM lanthanum) with air as sole energy, carbon, and nitrogen source. After three months, colony formation was checked via microscopy³¹. After 12 months, the activity of M. gorgona MG08 was measured as described in the section “Trace gas oxidation experiments” with the only difference that the KNO₃-free medium was used.

For the detection of ¹⁵N₂ fixation via NanoSIMS, filter cultures were pre-incubated on 10x diluted, EDTA-free, and potassium nitrate (KNO₃)-free NMS medium with air as sole energy, carbon source, and nitrogen source. After three months, the filters were transferred into 250 ml glass bottles containing 50ml of the KNO₃-free NMS medium. The bottles were capped with Safety-Caps (JR-S-11011, VICI AG International) and the headspace of each bottle was flushed for five minutes with compressed air using a gassing manifold. Afterwards, 50 ml of 98 + at% ¹⁵N₂ gas (NLM-363-1-LB, Cambridge Isotopes Laboratories, Tweksbury, Massachusetts, USA) were added using a gas syringe so that the ¹⁵N-N₂ in headspace atmosphere amounted approximately 23 at% of the total N₂. The headspace pressure was adjusted to 1.05 bar and 0.5 mL 1000 p.p.m.v. CH₄ in N₂ (HiQ, AGA, Sweden), 1 mL 1000 p.p.m.v. H₂ in N₂ (HiQ, AGA, Sweden), and 1 mL 1000 p.p.m.v. CO in N₂ (HiQ, AGA, Sweden) were added to the headspace using a gas syringe. To account for the natural abundance of ¹⁵N-N₂, a control without the addition of ¹⁵N₂ gas was prepared accordingly. The filter cultures were incubated at 20°C for two months. The headspace concentration of CH₄, H₂, and CO during incubation was measured once a week as mentioned in the “Trace gas oxidation experiments” section and replenished if the concentration of CH₄, H₂, and CO dropped below 1.9 p.p.m.v., 0.5 p.p.m.v., 0.2 p.p.m.v., respectively.

After two months, the cells of the filter cultures were fixed by incubating the filters on 150 µl formaldehyde in 1×PBS (4% w/v) for an hour at room temperature. The cells were washed twice by letting the filters float on water for 5 minutes. The fixed cells were harvested as described in “Cell quantification”, lyophilized, and stored at -80°C until further processing. For NanoSIMS analysis the lyophilized cells were resuspended in 20 µl MilliQ water. 10 µl of the cell suspensions were deposited on antimony-doped silicon wafer platelets (7.1 × 7.1 × 0.75 mm, Active Business Company, Germany) and dried in air. The following two samples were prepared accordingly for NanoSIMS analysis: i) The unlabeled M. gorgona MG08 cells, serving as control for the natural abundance of ¹⁵N₂; and ii) the M. gorgona MG08 cells incubated with ¹⁵N₂ gas and trace concentrations of CH₄, H₂, and CO that were analyzed to see whether the strain is capable of fixing atmospheric nitrogen during growth with air as sole energy and carbon source.

The NanoSIMS measurements were performed on a NanoSIMS 50 l (Cameca, Gennevilliers, France) at the Large-Instrument Facility for Advanced Isotope Research at the University of Vienna. Before the data acquisition, analysis areas were preconditioned in situ by rastering of a high-intensity, defocused Cs⁺ ion beam in the following sequence of high and extreme low ion impact energies (HE/16 keV and EXLIE/50 eV, respectively): HE at 25 pA beam current to a Cs + fluence of 5.0E14 ions cm^− 2; EXLIE at 400 pA beam current to a fluence of 5.0E16 ions cm^− 2; and HE at 25 pA to a fluence of 5.0E14 ions cm^− 2. Data were acquired as multilayer image stacks by repeated scanning of a finely focused Cs⁺ primary ion beam (c. 80 nm probe size at approx. 2 pA beam current) over areas between 34 × 34 and 72 × 72 µm² at 512 × 512-pixel image resolution and a primary ion beam dwell time of 5 ms pixel^− 1.

NanoSIMS images were generated and analyzed with the OpenMIMS plugin⁸⁰ in the image processing package Fiji⁸¹. All images were auto-tracked for compensation of primary ion beam and/or sample stage drift, and secondary ion signal intensities were corrected for detector dead-time and quasi-simultaneous arrival (QSA) of secondary ions, utilizing sensitivity factors (‘beta’ values) of 1.06, and 1.05 for C₂⁻, and CN⁻ ions, respectively. Regions of interest (ROIs) were defined in the ¹²C⁻-ion image where each ROI corresponded to an individual cell. Cells touching the border of the image were omitted from the selection.

Acquisition cycles of all three analysis areas were reduced to 27 each to improve comparability among the measurements. Regions of interest were analyzed for their ¹⁵N content by calculating the average value across acquisition cycles per analysis area, referred to by the ¹⁵N/(¹⁴N + ¹⁵N) isotope fraction designated as at% ¹⁵N, which was calculated from the ¹²CN⁻ signal intensities via:

$$at\% {}^{15}N= \frac{{}^{12}C{}^{15}{N}^{-}}{{}^{12}C{}^{15}{N}^{-}+{}^{12}C{}^{14}{N}^{-}}$$

The natural abundance of ¹⁵N in cellular biomass was inferred from unlabeled cells yielding 0.369 ± 0.043 at% (mean ± 1 SD).

Plotting

Plots were created using the Python 3⁶⁸ packages Pandas⁶⁹, Matplotlib⁷⁰, and Seaborn⁷¹ and R packages ggplot2⁸². Figures were finalized using Adobe Illustrator 2023.

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Metabolic Basis for the Microbial Oxidation of Atmospheric Methane

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