Observations of trace methane (CH4) in the Martian atmosphere are significant to the astrobiology community given the overwhelming contribution of biological methanogenesis to atmospheric CH4 on Earth. Previous studies have shown that methanogenic Archaea can generate CH4 when incubated with perchlorates, highly oxidizing chaotropic salts which have been found across the Martian surface. However, the regulatory mechanisms behind this remain completely unexplored. In this study we performed comparative transcriptomics on the methanogen Methanosarcina barkeri, which was incubated at 30˚C and 0˚C with 10–20 mM calcium-, magnesium-, or sodium perchlorate. Consistent with prior studies, we observed decreased CH4 production and apparent perchlorate reduction, with the latter process proceeding by heretofore essentially unknown mechanisms. Transcriptomic responses of M. barkeri to perchlorates include up-regulation of osmoprotectant transporters and selection against redox-sensitive amino acids. Increased expression of methylamine methanogenesis genes suggest competition for H2 with perchlorate reduction, which we propose is catalyzed by up-regulated molybdenum-containing enzymes and maintained by siphoning diffused H2 from energy-conserving hydrogenases. Methanogenesis regulatory patterns suggest Mars’ freezing temperatures alone pose greater constraints to CH4 production than perchlorates. These findings increase our understanding of methanogen survival in extreme environments and confers continued consideration of a potential biological contribution to Martian CH4.
The story of Martian atmospheric methane (CH4) remains enigmatic and under intense debate. In the past 15 years, a growing body of evidence has unfolded to suggest episodic appearances (and disappearances) of ppbv-level CH41,2,3,4,5,6,7. Myriad abiotic mechanisms have been suggested as potential CH4 sources, including cometary impacts8, UV degradation of meteoritic and interplanetary dust particle organics9,10,11, and Fischer–Tropsch-type synthesis in the subsurface coupled to either serpentinization of ultramafic silicates12,13 or radiolysis of H2O14, which subsequently releases CH4 to the surface through seeps and salinity-induced hydrate dissociation15 (Fig. 1). On Earth, however, nearly 70% of CH4 is of biological origin, generated by methanogenic Archaea16. This has led to an extensive debate considering the biological origin of Martian CH4. Understandably, the quest to comprehend the nature of CH4 cycling on Mars is a fervent one, as it may be the most conspicuous biosignature detected on Mars to date. In the absence of returned Martian samples or stable isotopic data of Martian atmospheric CH4, presently the best approach to constrain this debate is to experimentally test the ability of methanogenic Archaea to metabolize under Martian conditions.
Methanogens are not only among the most deeply rooted microorganisms in the tree of life, but they have also proliferated into nearly every habitable anaerobic environment and possess conserved adaptions for growth and survival under exposure to prolonged desiccation17,18,19, high salinity20,21,22,23,24, strong oxidants25,26, and extremes in temperature, pH, and pressure27,28,29,30,31. Thus, they have been the subject of extensive study to infer how hostile conditions simulating modern Mars may allow—or inhibit—biological methanogenesis17,19,25,26,31,32,33,34.
Perchlorate salts are highly soluble, chaotropic compounds of primarily human-manufactured origin on Earth35, with naturally occurring deposits mostly limited to evaporites in hyper-arid regions such as the Atacama Desert36 and the Antarctic Dry Valleys37. On Mars, however, perchlorates appear to be pervasive38,39,40,41,42,43,44. Perchlorates are of great interest to astrobiology—particularly, Martian habitability studies—for their hygroscopicity and low eutectic temperatures, which allows for the formation of stable liquid water brines at temperatures as low as -74.6˚C and 55% relative humidity45,46,47,48,49,50. Previous work has reported decreased CH4 production in methanogenic cultures supplemented with increasing concentrations of perchlorate salts25,26. This suggests that this aspect of the Martian environment may be debilitating to any potential biological methanogenesis that could occur under favorable conditions, but the mechanisms resulting in this apparent inhibition in analogue experiments have not been identified. Here we utilize transcriptomics to evaluate regulatory responses of the methanogenic archaeon Methanosarcina barkeri strain MS during a 28-day exposure to high concentrations of sodium-, magnesium-, and calcium perchlorate salts at 30 °C and 0 °C to assess if perchlorate salts are inhibitory to methanogenesis—and if so, how—in order to appraise the potential for methanogens’ survival on Mars.
Transcriptome assembly statistics
M. barkeri possesses the second largest described genome amongst the methanogenic Archaea51. This genome comprises a 4.53 megabase (Mb) circular chromosome and a 40 kilobase (kb) plasmid, which collectively encode 3,760 genes, 3,470 of which are protein-encoding coding sequence (CDS) regions (3.17 Mb). RNA sequencing yielded a total of 1,500,716,043 quality paired end reads across 24 libraries (8 conditions × 3 replicates/condition) with a mean Phred (sequence quality) score of 36. On average 1.6 ± 0.7% of 30 °C and 1.5 ± 0.5% of 0 °C quality-filtered reads mapped back to CDS regions (n = 12 libraries/temperature condition; Table S1 consistent with expectations that mRNA typically comprises 1–5% of total RNA in prokaryotic cells52. Average fragment counts per million mapped reads (FPM) are organized by gene position in Table S2 and visualized in Figs. S1–S2.
Methanogenesis and associated regulatory responses
At 30 °C, the addition of Ca(ClO4)2, Mg(ClO4)2, and Na(ClO4) reduced total net CH4 production by 48%, 32%, and 24%, respectively, relative to the perchlorate-free control (Fig. 2). Significant reduction in CH4 production rates were observed across all conditions at 0 °C with respect to their 30 °C counterparts, and each perchlorate condition at 0 °C showed a statistically significant decrease in CH4 for at least one time point relative to the 0 °C control (Fig. 2). No CH4 production was observed in the media blank controls (data not shown). Cultures were monitored via optical density measurements at 600 nm (OD600), but perchlorate-amended media experienced precipitate formation which made obtaining reliable growth data difficult (Fig. S3).
When incubated at 0 °C, the perchlorate-free control demonstrated significant up-regulation of several genes in the hydrogenotrophic pathway relative to the 30 °C perchlorate-free control (Table 1, Fig. 3), including log2-fold changes (LFC) in molybdenum (Mo)-formylmethanofuran dehydrogenase subunit B (fmdB), methenyl-tetrahydrosarcinapterin (H4SPT) cyclohydrolase (mch), and periplasmic heterodisulphide reductase (hdrDE). Likewise, significant down-regulation was observed in the sodium ion (Na+) transporter methyl-H4SPT:coenzyme M methyltransferase complex (mtrA), as well as the F420-reducing subunit of the periplasmic energy conserving hydrogenase (echF). All subunits of both coenzyme F420 hydrogenases (frhαβγ) were significantly up-regulated in perchlorate-free 0 °C incubations (Table 1, Fig. 3). In the perchlorate-treated incubations at 0 °C, we only observed down-regulation of frhαβγ in the presence of Ca(ClO4)2 with respect to the 0 °C perchlorate-free control (Table 2, Fig. 3).
At 30 °C, methanogenesis-associated regulatory responses were shared across all perchlorate conditions (Table 1). This included up-regulation of B, C, and D subunits of (Mo)-formylmethanofuran dehydrogenase, fmd, which catalyzes the CO2 reduction step of the hydrogenotrophic pathway (Fig. 3). In the presence of both Mg(ClO4)2 and Na(ClO4), one copy of (Mo)-fmdE exhibited significant up-regulation, while the other was significantly down-regulated (Table 1). Unique to the Mg(ClO4)2 incubations was the up-regulation of tungsten (W)-fmdA. In contrast, perchlorate-supplemented treatments incubated at 0 °C demonstrated statistically significant down-regulation of Mo-fmd genes, but up-regulation of the (W)-fmd operon (Table 2).
Energy conserving hydrogenase subunit F, echF, which supplies reduced ferredoxin to fmd, was down-regulated in the presence of Mg(ClO4)2 and Na(ClO4) at 30 °C, whereas most subunits of ech were significantly up-regulated in 30 °C Ca(ClO4)2 conditions (Fig. 3). Among other hydrogenases demonstrating significant up-regulation was methanophenazine-dependent hydrogenase, vht, specifically, the large subunit vhtA and cytochrome b subunit vhtC (Fig. 3). At 0 °C, ech hydrogenases were not differentially regulated in the presence of perchlorates relative to the perchlorate-free control (Fig. 3).
Despite H2 being the only reducing equivalent provided for the production of CH4 in our incubations, the addition of Ca(ClO4)2, Mg(ClO4)2, and Na(ClO4) resulted in significant up-regulation of all genes in the mono-, di-, and trimethylamine pathways and associated membrane permeases in the 30 °C perchlorate treatments (Fig. 3, Tables S7, S9, S11).
The terminal step encoding methyl-coenzyme M (CH3-CoM) reductase subunit alpha (mcrA) was down-regulated at 30 °C in Mg(ClO4)2 and Na(ClO4) enrichments (Fig. 3, Table 1). Although this reduction in expression is consistent with the decreased CH4 production rates observed in these treatments (Fig. 2), the Ca(ClO4)2-amended M. barkeri MS, which generated the least CH4, showed no significant difference in expression of the mcr complex relative to the 30 °C perchlorate-free control (Fig. 3, Table 1). Furthermore, no elements of mcr were significantly differentially expressed at 0 °C in the perchlorate-free control relative to the 30 °C perchlorate-free control (Fig. 3) despite its far lower CH4 production rate (Fig. 2). Therefore, the decreased expression of mcrA does not appear to be sufficient to explain the associated decrease in CH4 production rates with perchlorate exposure at 30 °C or incubation at 0 °C.
The carbon monoxide dehydrogenase/acetyl-CoA synthase complex (CODH/ACS), which plays a key role in both energy conversation and carbon fixation via the Wood-Ljungdahl pathway, demonstrated significant regulatory changes as a function of perchlorate exposure and temperature. The cooS subunit of CODH, which reversibly converts CO and CO2, was up-regulated in 30 °C Mg(ClO4)2 and Na(ClO4) treatments (Fig. 3, Table 1). Carbon monoxide dehydrogenase subunit epsilon (cdhε), which recycles ferredoxin in the reversible conversion between CO and CO2, was up-regulated with Ca(ClO4)2 at 30 °C (Fig. 3, Table 1), but was down-regulated in the 0 °C Mg(ClO4)2 and Na(ClO4) treatments (Fig. 3, Table 2). The 30 °C Ca(ClO4)2 treatment also demonstrated significant up-regulation of 5-H4SPT:corrinoid Fe-S protein methyltransferase (cdhγ), which plays a key role in the generation H4SPT and acetyl-CoA for biomass synthesis in the Wood-Ljungdahl pathway (Fig. 3, Table 1). Both copies of this gene were down-regulated at 0 °C in the Mg(ClO4)2 and Na(ClO4) treatments (Fig. 3, Table 2).
Both alpha and beta chains of ACS, respectively encoded by cdhα and cdhβ, were down-regulated in the 0 °C Na(ClO4) treatment with respect to the 0 °C perchlorate-free control, while only cdhα was down-regulated in the 0 °C Mg(ClO4)2 treatment (Fig. 3, Table 2). Cdhδ, which encodes an iron-sulfur corrinoid protein, was also down-regulated at 0 °C in Mg(ClO4)2 and Na(ClO4)-supplemented incubations (Fig. 3, Table 2). We observed no significant differential expression of CODH/ACS complex genes in the 0 °C Ca(ClO4)2 treatment (Fig. 3).
Concurrent up-regulation of ammonium transporters, Mo-nitrogenases, and P-II repressors
The most substantial up-regulation patterns were observed in genes relating to nitrogen cycling in perchlorate-incubated treatments. The 30 °C Mg(ClO4)2 and Na(ClO4)-amended conditions showed positive differential expression of 6 genes in the (Mo)-nitrogenase (hereafter (Mo)-Nase) complex (Fig. 4) including nifH, the MoFe-dinitrogen reductase that is responsible for electron transfer to the α2β2 N2 binding site (encoded by nifD and nifK, respectively) via ATP hydrolysis, as well as the P-II regulatory repressors, nifI, which shut off N2 fixation when ammonia is bioavailable53,54,55,56. Biosynthesis and assembly proteins for (Mo)-Nase, nifE and nifN, were also upregulated at 30 °C with Mg(ClO4)2 and Na(ClO4). At 30 °C, only nifH demonstrated significant up-regulation in both the Ca(ClO4)2 and perchlorate-free control, but the 0 °C Ca(ClO4)2 treatment also showed significant up-regulation of nifI and nifE (Fig. 4).
We observed substantial up-regulation of ammonium transporters (amt) in the 30 °C perchlorate treatments, but no significant differences in amt expression were observed in any 0 °C treatments (Fig. 4). Ammonium measurements confirmed that bioavailable fixed N remained replete (mM-level concentration) across all conditions for the duration of the incubation (Table S3).
Up-regulation of osmostress response genes
In the 30 °C Mg(ClO4)2 treatment, the complete operon for osmostress protectants uptake A, opuA, observed positive LFC of 0.64 ± 0.17 (opuAA, P < 0.001), 0.73 ± 0.20 (opuAB, P < 0.001), and 0.66 ± 0.30 (opuAC, P = 0.02). In the 30 °C Na(ClO4) treatment, significant up-regulation was also observed for opuAA (LFC = 0.54 ± 0.17, P = 4 \(\times\) 10–3) and opuAB (LFC = 0.59 ± 0.20, P = 7 \(\times\) 10–3). OpuA is responsible for the uptake of extracellular glycine betaine, belonging to a family of ABC transporters that hydrolyze ATP to import glycine betaine and other osmoprotectants such as proline57,58,59. The opu family also includes uptake systems for choline, a glycine betaine precursor59, but M. barkeri lacks the cellular machinery for de novo glycine betaine synthesis60. Relative to the 0 °C perchlorate-free control, significant down-regulation of opuAA in 0 °C Mg(ClO4)2 (LFC = − 0.94 ± 0.30, P = 8.02 \(\times\) 10–3) and Na(ClO4) (LFC = − 0.69 ± 0.31, P = 3.14 \(\times\) 10–2) treatments was observed. OpuAB was also down-regulated in the 0 °C Mg(ClO4)2 treatment (LFC = − 1.03 ± 0.38, P = 1.56 \(\times\) 10–2).
Evidence for osmotic stress can also be observed in the regulation of cell surface protein synthesis. Methanochondroitin is the primary constituent of the extracellular matrix that clumps M. barkeri cells into multicellular aggregates under optimal growth conditions61. Glucuronic acid, generated from glucose degradation via UDP-glucose dehydrogenase (UGDH), is a major component of methanochondroitin61,62. We observed significant down-regulation of UGDH under Mg(ClO4)2 and Na(ClO4) conditions at 30 °C (LFCMg = − 0.55 ± 0.17, P = 3 \(\times\) 10–3; LFCNa = − 0.51 ± 0.17, P = 7 \(\times\) 10–3).
Down-regulation of sulfur-containing amino acids
In addition to the 20 common amino acids, M. barkeri also encodes a 21st residue, pyrrolysine, via the ‘amber’ stop codon UAG63. A comparison of the genes encoding amino acid synthesis proteins showed large negative log2-fold changes at 30 °C in Mg(ClO4)2 and Na(ClO4)-amended treatments with respect to cysteine-producing proteins cysteine synthase (cysK) and serine acetyltransferase (cysE) (Fig. 5a). Further examination of complete amino acid metabolic pathways (Figs. S4–S16) revealed that this pattern of substantial gene down-regulation was characteristic of not only cysteine, but also the other sulfur-containing amino acid methionine (Fig. 5b).
In this study we took our inspiration from nearly two decades of research into Martian methane and associated astrobiology analogue studies to investigate how methanogens may survive under challenging conditions analogous of the Martian shallow subsurface, specifically under freezing temperatures and in the presence of chaotropic perchlorate salts. We tracked CH4 production and changes in global gene expression in M. barkeri MS via RNA-Seq following 28 days’ incubation at 30 °C or 0 °C, with and without 10 mM of dissolved Na-perchlorate and 20 mM of dissolved Mg- or Ca-perchlorate. In accordance with prior findings25,26 we observed quantifiable but inhibited methanogenesis in cultures supplemented with perchlorates (Fig. 2). For the first time we present transcriptomic evidence of the underlying biochemistry. Regulatory responses point to unambiguous shifts in amino acid synthesis and recycling, as well as mechanisms to combat increased osmotic stress—not unexpected reactions for an obligate anaerobe exposed to strongly oxidizing and chaotropic perchlorate salts. A surprising regulatory switch was observed in the methanogenesis pathway, with a significant up-regulation of methylamine-utilizing genes, despite these substrates not being present in the media ab initio. Metalloenzymes with molybdenum active sites, including (Mo)-formylmethanofuran dehydrogenase and (Mo)-Nase were among the most significantly up-regulated genes in perchlorate-amended incubations at 30 °C. We contextualize these revelations with a precedent of perchlorate reduction in methanogenic cultures25,26, offering new insight into Mo active sites as hydrogenation catalysts for these reactions to proceed.
Evidence for selection against redox-sensitive amino acids under temperate conditions
Cysteine and methionine are exceptionally sensitive to oxidation by reactive radical species64. Both residues have shown strong binding affinities to both perchlorate and perchloric acid, resulting in the oxidation of methionine to methionine sulfoxide and cysteine to sulfonic acid65. The susceptibility of cysteine and methionine to react with perchlorate risks degradation of protein structure and function. The extensive and substantial down-regulation of cysteine and methionine metabolic pathways we observed in the presence of Na- and Mg-perchlorates at 30 °C (Fig. 5) suggests a concerted effort by M. barkeri to reduce the synthesis and repair of these residues in the presence of these perchlorate species. A dearth of significant differential expression of amino acid-recycling genes (including those specific for cysteine and methionine) in both 0 °C and 0 °C perchlorate treatments demonstrates that freezing conditions are not sufficient to confer a decrease in transcriptional activity. It is clear that the synthesis and processing of essential amino acids in M. barkeri must proceed despite the metabolic stresses of frigid temperatures and strong oxidants.
A role for glycine betaine in cryo- and osmoprotection, and as a potential methylamine precursor
The regulatory patterns of enzyme complexes protecting against osmotic stress like opuAABC coincide with temperature and redox conditions. This operon is up-regulated at 30 °C with Na(ClO4) and Mg(ClO4)2, implying an increased need for cellular machinery to import glycine betaine to combat osmotic stress. In addition to being an osmoprotectant, glycine betaine is also cryoprotective66. While perchlorate salts are highly oxidizing, they also depress the freezing point of water. The down-regulation of opuAB in Mg(ClO4)2-amended cultures at 0 °C might suggest a decreased cold shock response in M. barkeri, wherein glycine betaine uptake for its cryoprotective properties was not needed under freezing conditions.
The up-regulation of methylamine-specific methanogenesis pathway genes in all three 30 °C perchlorate treatments was unexpected. Methylamines were not present in the media ab initio and ion chromatography mass spectrometry did not reveal the presence of monomethylamine in assessed samples (Table S3)—though if samples had ppb-levels of CH3NH2, it was diluted below the instrument’s detection limit (see Materials and Methods). Free energy yields of methylamine methanogenesis reactions at 30 °C range from -91 to -143 kJ/mol CH4—significantly less than the − 158 kJ/mol CH4 of the hydrogenotrophic pathway (Table S4). It has been suggested that glycine betaine may be a potential precursor of trimethylamine (TMA)67,68. The observed up-regulation of monomethylamine and dimethylamine permeases mtmP and mtbP (Fig. 3) imply possible interactions between perchlorates and complex media constituents (e.g., perhaps yeast extract or casitone) that might yield methylaminated species for utilization by M. barkeri, though we acknowledge such interpretations are presently speculative as our ion chromatography analyses were not sensitive to the detection of dimethylamine, trimethylamine, or any other methylaminated species.
Up-regulation of Mo-containing enzymes suggest mechanism for H2-dependent perchlorate reduction
Fixed nitrogen was replete under initial incubation conditions (9.3 mM NH4Cl plus additional N from complex ingredients like yeast extract and casitone). The significant up-regulation of ammonium transporters (amt, Fig. 4), methylamine permeases (mtmP and mtbP, Fig. 3), and P-II repressors (nifI, Fig. 4) in 30 °C perchlorate treatments were consistent with observations of mM-level NH4+ (Table S3) at the time of sample preservation for RNA-Seq. In M. barkeri, NH4+ concentrations as low as 10 µM have been shown to be inhibitory to N2 fixation53,54,55,56. It is therefore surprising to observe a simultaneous and substantial up-regulation of nitrogenase genes, an overwhelming proportion of which being associated with the molybdenum isoform, (e.g., nifDEHK, Fig. 4). Nitrogen fixation is an energetically expensive process, consuming at least 16 (and, by one calculation for methanogens69, perhaps more than 50) moles of ATP per mole of N2 fixed70. The significant increase in expression of the nifI repressors in the 30 °C Mg(ClO4)2 and Na(ClO4)-amended treatments suggest a concerted effort to dedicate cellular energy towards signaling the shut off of Mo-Nase activity. What is the purpose of this transcriptional arms race?
Microbial perchlorate reduction has been identified throughout the Proteobacteria35,71,72, as well as halophilic73 and hyperthermophilic Archaea74. The substrate-specific and Mo-containing perchlorate reductase (pcrAB), which reduces perchlorate to chlorite (ClO2-), and chlorite dismutase (cld), which dismutates ClO2- into chloride (Cl-) and molecular oxygen (O2), have been isolated and purified35, but homologues of these genes have not been found in the M. barkeri genome. However, chlorate- and perchlorate-binding affinities have been documented in structurally similar α- and β-subunits of other Mo-containing DMSO family reductases35,75,76. A plethora of inorganic and organic electron donors have also been implicated in microbial perchlorate reduction35,77,78,79 including H2, sulphide (S2-), CH4, and yeast extract, which are available constituents in the incubations analyzed in this study. The biochemistry of microbial perchlorate reduction is complex and not yet fully understood. Nonetheless, the following environmental conditions must be met in order for perchlorate reduction to proceed: (1) dissolved O2 concentrations must be < 2 mg L−1; (2) NO3- must be completely consumed; and (3) molybdenum must be bioavailable as molybdate, MoO42−, to synthesize Mo cofactors35,80 in perchlorate-reducing metalloenzymes. All three conditions were coincidentally satisfied in this study’s incubation conditions.
Perchlorate reduction has been observed in sterile methanogenic media under an 80:20 H2:CO2 atmosphere25,26 and apparently exacerbated in media with the psychrotolerant methanogen Methanobacterium arcticum M2, leading to the suggestion that perchlorate could be utilized as a possible electron acceptor in a reversal of methanogenesis – the anaerobic oxidation of methane, AOM25 (Eq. 1). To date these interpretations remain unverified. In this study, we similarly observed a significant decrease in perchlorate concentrations in media containing M. barkeri – a loss of 17.4% for Mg(ClO4)2, 26.8% for Ca(ClO4)2, and 58.9% of Na(ClO4) by mass (Table S3). Anion measurements in the perchlorate-amended incubations did yield an increase in total Cl-, but only enough to account for 35.5% Mg(ClO4)2, 5.6% Ca(ClO4)2, and 44.8% of Na(ClO4) apparently reduced (Table S3). We note that due to its co-elution with Cl-, we were not able to ascertain the presence of chlorate (ClO3-). Chlorite (ClO2-) was not detected in any samples (Table S3). Additional resources and measurements in a future study are necessary to quantify ClO3- and other potential intermediates (e.g. chlorinated organics) to fully assess the fate of ClO4-.
Sodium sulphide, Na2S, a common reducing agent in anaerobic media, has been previously ruled out as a significant contributor to abiotic perchlorate reduction in methanogenic media (medium MSH in Kral et al.)26. Yeast extract is a complex and undefined carbon source, and its inclusion in sterile MSH media has been associated with increased rates of perchlorate reduction compared to a minimal salts medium (medium MM in Kral et al.)26. In this study, preliminary measurements of perchlorate concentrations in sterile DSMZ 120a media failed to detect conclusive perchorate reduction (Table S3). More work is necessary to fully explore the composition and role of yeast extract as a potential reducing agent.
The free energy yield of H2 oxidation coupled to perchlorate reduction (Eq. 2) is substantial (∆G˚' = − 289 kJ/mol H2) but the reaction is kinetically sluggish at ambient conditions and requires a metallic catalyst to overcome its large activation energy81.
Nickel (Ni) has long been known as an excellent hydrogenation catalyst82 and is an essential cofactor for hydrogenase activity in methanogens83, but an essential role in microbial perchlorate reduction has never been suggested and it is not obvious from our transcriptomic data that perchlorates elicit a universal response by Ni active sites in hydrogenases (e.g. echE, vhtA, and frhα in Fig. 3). Instead, given perchlorate reduction’s known Mo dependency35,84 and the additional empirical evidence presented here, we cannot ignore the most parsimonious explanation: the substantial up-regulation of (Mo)-Nase (Fig. 4) and (Mo)-formylmethanofuran dehydrogenase (Fig. 3, Table 1) at 30 °C must be associated with perchlorate reduction. Certainly, a thorough investigation is warranted in order to elucidate finer details of the biochemistry.
The methylamine methanogenesis pathway as a mechanism for energy conservation
In methylamine methanogenesis, H2 is recycled by the partial reversal of the methanogenesis pathway, generated via the oxidation of F420H2 by frhα and the oxidation of ferredoxin by echF (green arrows in Figs. 3). H2 generated by the oxidation of F420H2 and ferredoxin diffuses across the membrane to be oxidized by vhtA in an energy conserving scheme to recycle methanophenazine (MP)85,86,87,88. Ferredoxin oxidation by echF also results in the translocation of 2H+ through ech, contributing to the production of a proton gradient (high outside the cell)88. Evidence for the generation of this proton gradient might be indicated in our incubations based on drops in pH (up to 0.77 pH units) (Table S5). Based on up-regulation of the genes for methylamine methanogenesis, we attribute the differential expression of ech, vht, and frh hydrogenases (Fig. 3) and ferredoxin (Tables S10, S12, S14) as putative sources of a proton motive force in the 30 °C Ca(ClO4)2-supplemented incubations.
We theorize that the thermodynamic spontaneity of H2-dependent perchlorate reduction (in the presence of a Mo catalyst) results in it outcompeting the endergonic first step of hydrogenotrophic methanogenesis (Eq. 3; ∆G˚' = + 16 kJ/mol H2)89 (Fig. 6a).
Likewise, we imagine H2 intermediates generated during energy conserving steps of methylamine methanogenesis are siphoned off to reduce perchlorate in lieu of reducing MP via vht (∆G˚' = − 289 kJ/mol H2 for perchlorate reduction90 vs. ∆G˚' = − 50 kJ/mol H2 for MP reduction91,92) (Fig. 6b). Such interpretations warrant further investigation, but do fit within previously reported narratives of observed decreases in CH4 production in perchlorate-supplemented methanogenic cultures25,26.
Do all perchlorates trigger universal regulatory responses?
Calcium-, magnesium- and sodium perchlorate were investigated in this study as they have been detected in active Recurring Slope Lineae (RSL) on Mars43. Until now, studies investigating methanogen survival in high concentrations of perchlorate salts have been unable to ascertain the specific metabolic responses behind observed decreases in CH4 production, and whether these metabolic responses are universal regardless of the perchlorate’s constituent cation. Our work demonstrates that, for M. barkeri MS transcriptomes assembled after 28 days of incubation, methanogenesis regulatory patterns are consistent across all three perchlorate conditions with methylamine gene utilization and a handful of (Mo)-fmd subunits at 30 °C (Fig. 3). Otherwise, global expression patterns in Ca(ClO4)2 treatments were somewhat distinct from Na(ClO4) and Mg(ClO4)2 conditions, which, on the whole, exhibited similar transcriptomic responses at both temperatures (Figs. 4, 5; Tables 1, 2). Transcriptomes from additional time points can help clarify whether these discrepancies in expression are reflective of distinct metabolic responses to Ca-perchlorate versus Mg- and Na-perchlorate, or simply different stages of metabolism following prolonged exposure to perchlorates in general. We infer the latter is more likely given the novel and ubiquitous response of genes in the methylamine methanogenesis pathway.
Implications for astrobiology and Martian methane
These findings better constrain our growing understanding of how microbial life responds to prolonged exposure under extreme conditions characterized by strong oxidants, freezing temperatures, and osmotic stress—e.g., a Mars Special Region93 potentially conducive to microbial habitability. Notably, our study shows that major metabolic disruption by perchlorates at 30 °C is not reflected to the same degree at 0 °C, which is more appropriately representative of the conditions at Mars Special Regions93. This finding offers new perspectives to contextualize observations of diffuse Martian CH4 emissions, particularly in situ measurements made by the Curiosity rover5,6. This work offers an in-depth glimpse into the remarkable adaptability of methanogens to survive under oxidative stresses that mimic the challenging conditions of the Martian subsurface. The inferences made from this study provide many exciting opportunities for further research to better understand methanogen ecophysiology under extreme stress that may offer terrestrial insight into the enigmatic Martian methane story.
Materials and methods
Materials and culture conditions
Methanosarcina barkeri wild-type strain MS (ATCC 51582) was grown hydrogenotrophically (15 mL of 80:20 H2:CO2 headspace pressurized to 1.5 bar) in anaerobic balch tubes (Chemglass Life Sciences LLC, Vineland, NJ USA) containing 9 mL DSMZ medium 120a, 10 mg/L EDTA (pH 7.0 to 7.2)94, supplemented with trace element solution SL-1095 and vitamin supplement MD-VS (ATCC, Manassas, VA USA). Media assaying for perchlorate tolerance were augmented to final concentrations of 10 mM NaClO4 and 20 mM Mg(ClO4)2 or Ca(ClO4)2. Balch tubes were inoculated with 1 mL cell suspensions containing ~ 4 × 107 cells and incubated in batches of 9 biological replicates plus one autoclave-sterilized media blank for 28 days (Fig. S17). For each experimental treatment (perchlorate-free control, NaClO4, Mg(ClO4)2, or Ca(ClO4)2), optical density measurements (OD) and CH4 production were monitored weekly in parallel experiments incubated at 30 °C or 0 °C (Fig. 2).
OD measurements were performed spectrophotometrically at 600 nm using a Genesys 30 Visible Spectrometer (Thermo-Scientific Corp., Madison, WI USA). At the end of the incubation experiment, direct cell counts were performed on enrichment aliquots fluorescently stained with Acridine Orange (AO) dye (catalogue #318,337, Sigma-Aldrich Chemical, Co., Milwaukee, WI USA) (50 µM final concentration) following an established procedure96. Stained cells were enumerated in a Spotlite hemocytometer counting chamber (McGaw Park, IL USA) with a Zeiss Axioskope 40 epifluorescence microscope (ZEISS Microscopy, Jena, Germany) at 600 × magnification using an AO filter cube set (excitation 470/20 nm; emission > 510 nm).
Headspace gas samples were collected via gas-tight syringes (catalogue #24886, Restek U.S., Bellefonte, PA USA) to measure CH4 using a Trace 1310 gas chromatograph equipped with a flame-ionizing detector (GC-FID) (ThermoFisher Scientific, Waltham, MA USA). H2, O2, and N2 was were measured using a Peak Performer 1 series gas chromatograph coupled to a thermal conductivity detector (GC-TCD; Peak Laboratories, Mountain View, CA USA) and CO2 and CH4 were measured using an FID coupled to a methanizer (GC-FID; Peak Laboratories, Mountain View, CA USA). To avoid injecting atmospheric O2 and residual carrier-over of trace gases within the syringes between samples, all syringes were flushed three times with ultra-high purity argon gas between samples.
Single experimental replicates from each condition were reserved for measurement of anions (ClO4−, ClO2−, F−, Cl−, SO42−, NO2−, NO3−, HPO42−), major cations (Na+, Mg2+, Ca2+, K+), ammonium (NH4+), and monomethylamine (CH3NH2). Media was diluted 500 × in milliQ H2O and compositional analysis was performed by ion chromatography on a Dionex ICS-5000 + Capillary HPIC system (ThermoFisher Scientific, Waltham, MA USA). Perchlorate (ClO4−) and chlorite (ClO2−) were separated from other anions using a Dionex IonPac AS16 column (ThermoFisher Scientific, Waltham, MA USA). We note that chlorate (ClO3-) coeluted with Cl- and could not be distinguished. TraceCERT perchlorate standard (catalogue # 76462, Sigma-Aldrich Chemical, Co., Milwaukee, WI USA) and Dionex Combined Seven Anion II (catalogue # 057590, ThermoFisher Scientific, Waltham, MA USA) were used for standardization. Major cations, NH4+, and CH3NH2 were measured by diluting the media 100 × in milliQ H2O on the same ion chromatograph fitted with a CS16 column (ThermoFisher Scientific, Waltham, MA USA), and was standardized by the Dionex Combined Six Cation solution (catalogue #046070, ThermoFisher Scientific, Waltham, MA USA) and monomethylamine solution 40% by wt. in H2O (Sigma-Aldrich catalogue # 426466).
Compositional analyses of cations, anions, and headspace gas is available for reference in Table S3.
RNA isolation and purification
After 28 days’ incubation, culture and blank volume contents were briefly vortexed and aseptically transferred into sterile 15 mL Falcon tubes (Corning Inc., Corning, NY USA) at incubation temperature inside a Coy anaerobic chamber (Coy Laboratory Products Inc., Grass Lake, MI USA). Tubes were centrifuged at 3,000 × g for 1 min using an IEC Centra CL2 centrifuge (Thermo Electron Company, Milford, MA USA) to pellet cells. Media was poured off for pH and electrical conductivity measurements, leaving behind 1 mL. To ensure quantifiable RNA yields downstream, the nine biological replicates from each condition were consolidated into three sets of three samples each for extraction (3 mL pelleted cell suspension/tube). RNAlater solution (ThermoFisher Scientific, Waltham, MA USA) was added to a final volume of ~ 12 mL. Samples were left to equilibrate at incubation temperature for 3 h, transferred to 4 °C for 24 h, and then stored at -80 °C until overnight shipment on dry ice to Princeton University for RNA extraction.
Samples preserved at -80 ̊C in RNAlater were thawed on ice in a sealed container before contents were transferred to 50 mL Falcon® tubes. An equal volume (12 mL) of nuclease-free water (Qiagen, Hilden, Germany) was added to RNAlater-preserved samples, briefly vortexed, and centrifuged at 5000 gx for 10 min using a Sorvall Legend XI centrifuge (ThermoFisher Scientific, Waltham, MA USA). The supernatant was subsequently discarded, and RNA was extracted following a modified protocol from a Zymo Quick-RNA Miniprep Plus Kit (Zymo Research, Irvine, CA USA). RNA lysis buffer and nuclease-free water were added to each sample in a 5:1 ratio, and sterile 0.7 mm garnet bashing beads (Qiagen, Hilden, Germany) were added to facilitate mechanical lysis during subsequent vortexing. Samples were then vortexed for 1 min and centrifuged at 10,000 × g at 4 °C for 1 min using an Eppendorf 5810R (Eppendorf, Hamburg, Germany) to pellet cell debris. The supernatant containing total nucleic acids was transferred to a yellow Spin-Away column (Zymo Research, Irvine, CA USA) fitted in a 2 ml collection tube. Samples were centrifuged at 10,000 × g for 1 min using an AccuSpin Micro 17 (ThermoFisher Scientific, Waltham, MA USA) to separate out genomic DNA. Following centrifugation, the Spin-Away filter was discarded and the filtrate was collected from the column for RNA purification.
Total nucleic acids were then precipitated by adding 1 mL of chilled absolute ethanol. Pellets were mixed by pipetting and then transferred to green Zymo-Spin IIICG column filters fitted in clean collection tubes. Samples were centrifuged at 10,000 × g for 30 s to collect precipitated nucleic acids on the column, subsequently treated with 400 µL RNA wash buffer, and centrifuged for 30 s at 10,000 gx. The wash buffer was discarded and columns were treated with 80 µL DNAse I reaction mixture (per 80 µL: 5 µL DNAse 1 [1 U/µL], 8 µL 10X DNAse I reaction buffer [Zymo Research, Irvine, CA USA], 3 µL nuclease-free water, 64 µL RNA wash buffer [Zymo Research, Irvine, CA USA]) to degrade trace genomic DNA and left to incubate on ice in the dark for 15 min. DNAse-treated samples were then centrifuged at 10,000 × g for 30 s. The reaction buffers were discarded, column filters were washed three times with 400, 700, and 400 µl of RNA Prep Buffer (Zymo Research, Irvine, CA USA), centrifuging twice for 30 s at 10,000 × g and at 16,000 × g for 2 min for the final wash. Total RNA was then eluted into sterile, nuclease-free PCR tubes on ice using nuclease-free water pre-heated to 95 ̊C. Total RNA was quantified using a Qubit hs RNA assay kit coupled to a Qubit 2.0 analyzer (ThermoFisher Scientific, Waltham, MA USA) following the manufacturer’s protocol. RNA quality was subsequently assessed using a 2100 Bioanalyzer (Agilent Technologies Inc., Santa Clara, CA USA). RNA samples were kept at -20 °C until library preparation and sequencing.
All RNA extraction steps were performed using nuclease-free pipette tips (Corning Inc., Corning, NY USA) in a UV-sterilized laminar flow hood. All surfaces, pipettes, and gloves were wiped down at each step with RNaseZap solution (ThermoFisher Scientific, Waltham, MA USA) to minimize potential RNase contamination. Parallel extraction blanks of extraction kit reagents and blank growth media co-incubated with M. barkeri enrichment cultures were performed to ensure cleanliness of the extraction procedure and sterility of uninoculated growth media.
RNA library prep and RNA-Seq
Ribosomal RNA was depleted from total RNA using a Ribo-Zero Bacterial rRNA Removal Kit (Illumina, Inc., San Diego, CA USA) following the manufacturer’s instructions and using the provided universal probe sequence. One-directional library prep was performed for each treatment and its constituent 3 sequencing replicates using the Nextera DNA Flex Library Prep Kit (Illumina, Inc., San Diego, CA USA) on the automated Apollo 324 system (Takara Bio USA, Inc., Mountain View, CA, USA). RNA-Seq was carried out for 318 cycles on two lanes of a NovaSeq SP Flowcell (Illumina, Inc., San Diego, CA USA) (1 × 150 bp) at the Princeton University genomics core facility.
Annotation and comparative transcriptomics
Quality filtering of single-end reads was performed using fastp v.0.12.697, removing reads < 50 nt, containing > 1 Ns, Phred quality scores < 30, and sample barcode sequences. Using Bowtie2 v.2.3.298, retained, quality-filtered reads from each experiment were mapped to coding sequence (CDS) regions subset from the complete M. barkeri MS reference genome obtained from NCBI GenBank (NZ_CP009528.1). CDS-mapped reads were then sorted, indexed, and processed for extraction from the sequenced transcriptome using Samtools v.1.599 and BEDTools v.2.17.0100. Gene annotation was performed using NCBI BLASTn v.2.2.29 + 101 against a reference M. barkeri MS CDS assembly database generated using option -makeblastdb. Protein assignment was determined as the entry with the greatest sequence identity alignment with the query sequence, the lowest E-value, and largest bit score. Differential expression analysis of investigated treatments relative to the 30 °C and 0 °C perchlorate-free controls, within-group (i.e. biological replicate) variance estimation, and FPM were performed using DESeq2102. Metabolic pathway involvement of identified genes was determined by referencing the Kyoto Encyclopedia of Genes and Genomes (KEGG)103.
RNA-Seq data is available at NCBI GenBank under accession number PRJNA635445 (Sequence Read Archive SRP265010). The source code of all transcriptomic analyses presented in this manuscript are available upon request.
Krasnopolsky, V. A., Maillard, J. P. & Owen, T. C. Detection of methane in the martian atmosphere: evidence for life?. Icarus 172, 537–547 (2004).
Formisano, V., Atreya, S., Encrenaz, T., Ignatiev, N. & Giuranna, M. Detection of methane in the atmosphere of mars. Science 306, 1758–1761 (2004).
Geminale, A., Formisano, V. & Giuranna, M. Methane in Martian atmosphere: average spatial, diurnal, and seasonal behaviour. Planet. Space Sci. 56, 1194–1203 (2008).
Mumma, M. J. et al. Strong release of methane on mars in northern summer 2003. Science 323, 1041–1045 (2009).
Webster, C. R. et al. Mars methane detection and variability at Gale crater. Science 347, 415–417 (2015).
Webster, C. R. et al. Background levels of methane in Mars’ atmosphere show strong seasonal variations. Science 360, 1093–1096 (2018).
Korablev, O. et al. No detection of methane on Mars from early ExoMars Trace Gas Orbiter observations. Nature 568, 517–520 (2019).
Fries, M. et al. A cometary origin for martian atmospheric methane. Geochem. Perspect. Lett. 2, 10–23 (2016).
Keppler, F. et al. Ultraviolet-radiation-induced methane emissions from meteorites and the Martian atmosphere. Nature 486, 93–96 (2012).
Moores, J. E. & Schuerger, A. C. UV degradation of accreted organics on Mars: IDP longevity, surface reservoir of organics, and relevance to the detection of methane in the atmosphere. J. Geophys. Res. Planets 117, E8 (2012).
Schuerger, A. C., Moores, J. E., Clausen, C. A., Barlow, N. G. & Britt, D. T. Methane from UV-irradiated carbonaceous chondrites under simulated Martian conditions. J. Geophys. Res. Planets 117, E8 (2012).
Etiope, G., Ehlmann, B. L. & Schoell, M. Low temperature production and exhalation of methane from serpentinized rocks on Earth: a potential analog for methane production on Mars. Icarus 224, 276–285 (2013).
Oehler, D. Z. & Etiope, G. Methane seepage on mars: where to look and why. Astrobiology 17, 1233–1264 (2017).
Onstott, T. C. et al. Martian CH 4: sources, flux, and detection. Astrobiology 6, 377–395 (2006).
Elwood Madden, M. E., Ulrich, S. M., Onstott, T. C. & Phelps, T. J. Salinity-induced hydrate dissociation: A mechanism for recent CH4 release on Mars. Geophys. Res. Lett. https://doi.org/10.1029/2006GL029156 (2007).
Conrad, R. The global methane cycle: recent advances in understanding the microbial processes involved. Environ. Microbiol. Rep. 1, 285–292 (2009).
Kendrick, M. G. & Kral, T. A. Survival of methanogens during desiccation: implications for life on mars. Astrobiology 6, 546–551 (2006).
Anderson, K. L., Apolinario, E. E. & Sowers, K. R. Desiccation as a long-term survival mechanism for the archaeon Methanosarcina barkeri. Appl. Environ. Microbiol. 78, 1473–1479 (2012).
Kral, T. A. & Altheide, S. T. Methanogen survival following exposure to desiccation, low pressure and martian regolith analogs. Planet. Space Sci. 89, 167–171 (2013).
Sowers, K. R. & Gunsalus, R. P. Adaptation for growth at various saline concentrations by the archaebacterium Methanosarcina thermophila. J. Bacteriol. 170, 998–1002 (1988).
Maestrojuan, G. M. et al. Taxonomy and halotolerance of mesophilic methanosarcina strains, assignment of strains to species, and synonymy of methanosarcina mazei and methanosarcina frisia. Int. J. Syst. Bacteriol. 42, 561–567 (1992).
Sowers, K. R., Boone, J. E. & Gunsalus, R. P. Disaggregation of methanosarcina spp and growth as single cells at elevated osmolarity. Appl. Environ. Microbiol. 59, 3832–3839 (1993).
Sowers, K. R. & Gunsalus, R. P. Halotolerance in methanosarcina spp: Role of N(sup(epsilon))-Acetyl-(beta)-Lysine, (alpha)-Glutamate, Glycine Betaine, and K(sup+) as Compatible Solutes for Osmotic Adaptation. Appl. Environ. Microbiol. 61, 4382–4388 (1995).
Roessler, M. et al. Identification of a salt-induced primary transporter for glycine betaine in the methanogen methanosarcina mazei go1. Appl. Environ. Microbiol. 68, 2133–2139 (2002).
Shcherbakova, V., Oshurkova, V. & Yoshimura, Y. The effects of perchlorates on the permafrost methanogens: implication for autotrophic life on mars. Microorganisms 3, 518–534 (2015).
Kral, T. A. et al. Sensitivity and adaptability of methanogens to perchlorates: Implications for life on Mars. Planet. Space Sci. 120, 87–95 (2016).
Rivkina, E. M., Laurinavichus, K. S., Gilichinsky, D. A. & Shcherbakova, V. A. Methane generation in permafrost sediments. Dokl. Biol. Sci. https://doi.org/10.1023/A:1015366613580 (2002).
Rivkina, E. et al. Microbial life in permafrost. Adv. Sp. Res. 33, 1215–1221 (2004).
Rivkina, E. et al. Biogeochemistry of methane and methanogenic archaea in permafrost. FEMS Microbiol. Ecol. 61, 1–15 (2007).
Takai, K. et al. Cell proliferation at 122 degrees C and isotopically heavy CH4 production by a hyperthermophilic methanogen under high-pressure cultivation. Proc. Natl. Acad. Sci. U. S. A. 105, 10949–10954 (2008).
Sinha, N., Nepal, S., Kral, T. & Kumar, P. Survivability and growth kinetics of methanogenic archaea at various pHs and pressures: implications for deep subsurface life on Mars. Planet. Space Sci. 136, 15–24 (2017).
Chastain, B. K. & Kral, T. A. Approaching mars-like geochemical conditions in the laboratory: omission of artificial buffers and reductants in a study of biogenic methane production on a Smectite clay. Astrobiology 10, 889–897 (2010).
Kral, T. A., Altheide, T. S., Lueders, A. E. & Schuerger, A. C. Low pressure and desiccation effects on methanogens: Implications for life on Mars. Planet. Space Sci. 59, 264–270 (2011).
Mickol, R. L. & Kral, T. A. Low pressure tolerance by methanogens in an aqueous environment: implications for subsurface life on mars. Orig. Life Evol. Biosph. 47, 511–532 (2017).
Coates, J. D. & Achenbach, L. A. Microbial perchlorate reduction: rocket-fuelled metabolism. Nat. Rev. Microbiol. 2, 569–580 (2004).
Ericksen, G. E. The Chilean Nitrate Deposits: The origin of the Chilean nitrate deposits, which contain a unique group of saline minerals, has provoked lively discussion for more than 100 years. Am. Sci. 71, 366–374 (1983).
Kounaves, S. P. et al. Discovery of natural perchlorate in the antarctic dry valleys and its global implications. Environ. Sci. Technol. 44, 2360–2364 (2010).
Hecht, M. H. et al. Detection of perchlorate and the soluble chemistry of Martian soil at the phoenix lander site. Science 325, 64–67 (2009).
Navarro-González, R., Vargas, E., de la Rosa, J., Raga, A. C. & McKay, C. P. Reanalysis of the Viking results suggests perchlorate and organics at midlatitudes on Mars. J. Geophys. Res. 115, E12010 (2010).
Glavin, D. P. et al. Evidence for perchlorates and the origin of chlorinated hydrocarbons detected by SAM at the Rocknest aeolian deposit in Gale Crater. J. Geophys. Res. Planets 118, 1955–1973 (2013).
Kounaves, S. P. et al. Identification of the perchlorate parent salts at the Phoenix Mars landing site and possible implications. Icarus 232, 226–231 (2014).
Kounaves, S. P., Carrier, B. L., O’Neil, G. D., Stroble, S. T. & Claire, M. W. Evidence of martian perchlorate, chlorate, and nitrate in Mars meteorite EETA79001: Implications for oxidants and organics. Icarus 229, 206–213 (2014).
Ojha, L. et al. Spectral evidence for hydrated salts in recurring slope lineae on Mars. Nat. Geosci. https://doi.org/10.1038/ngeo2546 (2015).
Clark, B. C. & Kounaves, S. P. Evidence for the distribution of perchlorates on Mars. Int. J. Astrobiol. 15, 311–318 (2016).
Pestova, O. N., Myund, L. A., Khripun, M. K. & Prigaro, A. V. Polythermal study of the systems M(ClO4)2–H2O (M2+ = Mg2+, Ca2+, Sr2+, Ba2+). Russ. J. Appl. Chem. 78, 409–413 (2005).
Chevrier, V. F., Hanley, J. & Altheide, T. S. Stability of perchlorate hydrates and their liquid solutions at the Phoenix landing site Mars. Geophys. Res. Lett. 36, L10202 (2009).
Marion, G. M., Catling, D. C., Zahnle, K. J. & Claire, M. W. Modeling aqueous perchlorate chemistries with applications to Mars. Icarus 207, 675–685 (2010).
Stillman, D. E. & Grimm, R. E. Dielectric signatures of adsorbed and salty liquid water at the Phoenix landing site Mars. J. Geophys. Res. 116, E09005 (2011).
Toner, J. D., Catling, D. C. & Light, B. The formation of supercooled brines, viscous liquids, and low-temperature perchlorate glasses in aqueous solutions relevant to Mars. Icarus 233, 36–47 (2014).
Nikolakakos, G. & Whiteway, J. A. Laboratory investigation of perchlorate deliquescence at the surface of Mars with a Raman scattering lidar. Geophys. Res. Lett. 42, 7899–7906 (2015).
Maeder, D. L. et al. The Methanosarcina barkeri Genome: Comparative Analysis with Methanosarcina acetivorans and Methanosarcina mazei Reveals Extensive Rearrangement within Methanosarcinal Genomes. J. Bacteriol. 188, 7922–7931 (2006).
Sorek, R. & Cossart, P. Prokaryotic transcriptomics: a new view on regulation, physiology and pathogenicity. Nat. Rev. Genet. 11, 9–16 (2010).
Lobo, A. L. & Zinder, S. H. Diazotrophy and Nitrogenase Activity in the Archaebacterium Methanosarcina barkeri 227. Appl. Environ. Microbiol. 54, 1656–1661 (1988).
Lobo, A. L. & Zinder, S. H. Nitrogenase in the archaebacterium Methanosarcina barkeri 227. J. Bacteriol. 172, 6789–6796 (1990).
Kessler, P. S. & Leigh, J. A. Genetics of nitrogen regulation in Methanococcus maripaludis. Genetics 152, 1343–1351 (1999).
Kessler, P. S., Daniel, C. & Leigh, J. A. Ammonia Switch-Off of Nitrogen Fixation in the Methanogenic Archaeon Methanococcus maripaludis: Mechanistic Features and Requirement for the Novel GlnB Homologues, NifI1 and NifI2. J. Bacteriol. 183, 882–889 (2001).
Kempf, B. & Bremer, E. OpuA, an osmotically regulated binding protein-dependent transport system for the osmoprotectant glycine betaine in bacillus subtilis. J. Biol. Chem. 270, 16701–16713 (1995).
Kempf, B. & Bremer, E. Uptake and synthesis of compatible solutes as microbial stress responses to high-osmolality environments. Arch. Microbiol. 170, 319–330 (1998).
Hoffmann, T. & Bremer, E. Guardians in a stressful world: the Opu family of compatible solute transporters from Bacillus subtilis. Biol. Chem. 398, 193–214 (2017).
Hippe, H., Caspari, D., Fiebig, K. & Gottschalk, G. Utilization of trimethylamine and other N-methyl compounds for growth and methane formation by Methanosarcina barkeri. Proc. Natl. Acad. Sci. 76, 494–498 (1979).
Kreisl, P. & Kandler, O. Chemical structure of the cell wall polymer of methanosarcina. Syst. Appl. Microbiol. 7, 293–299 (1986).
Jarrell, K. F., Jones, G. M., Kandiba, L., Nair, D. B. & Eichler, J. S-layer glycoproteins and flagellins: reporters of archaeal posttranslational modifications. Archaea 2010, 1–13 (2010).
Srinivasan, G. Pyrrolysine encoded by UAG in archaea: charging of a UAG-decoding specialized tRNA. Science 296, 1459–1462 (2002).
Bin, P., Huang, R. & Zhou, X. Oxidation resistance of the sulfur amino acids: methionine and cysteine. Biomed Res. Int. 2017, 1–6 (2017).
Armesto, X. L., Canle, L. M., Fernández, M. I., Garcı́a, M. V. & Santaballa, J. A. First steps in the oxidation of sulfur-containing amino acids by hypohalogenation: very fast generation of intermediate sulfenyl halides and halosulfonium cations. Tetrahedron 56, 1103–1109 (2000).
Casanueva, A., Tuffin, M., Cary, C. & Cowan, D. A. Molecular adaptations to psychrophily: the impact of ‘omic’ technologies. Trends Microbiol. 18, 374–381 (2010).
Oren, A. Formation and breakdown of glycine betaine and trimethylamine in hypersaline environments. Antonie Van Leeuwenhoek 58, 291–298 (1990).
Seibel, B. A. & Walsh, P. J. Trimethylamine oxide accumulation in marine animals: relationship to acylglycerol storage. J. Exp. Biol. 205, 297–306 (2002).
Lobo, A. L. & Zinder, S. H. Nitrogen fixation by methanogenic bacteria. in Biological Nitrogen Fixation (eds. Stacey, G., Burris, R. H. & Evans, H. J.) 191–211 (Chapman and Hall, 1992).
Sohm, J. A., Webb, E. A. & Capone, D. G. Emerging patterns of marine nitrogen fixation. Nat. Rev. Microbiol. 9, 499–508 (2011).
Bardiya, N. & Bae, J.-H. Dissimilatory perchlorate reduction: A review. Microbiol. Res. 166, 237–254 (2011).
Barnum, T. P. et al. Genome-resolved metagenomics identifies genetic mobility, metabolic interactions, and unexpected diversity in perchlorate-reducing communities. ISME J. 12, 1568–1581 (2018).
Oren, A., Elevi, B. R. & Mana, L. Perchlorate and halophilic prokaryotes: implications for possible halophilic life on Mars. Extremophiles 18, 75–80 (2014).
Liebensteiner, M. G., Pinkse, M. W. H., Schaap, P. J., Stams, A. J. M. & Lomans, B. P. Archaeal (Per)Chlorate reduction at high temperature: an interplay of biotic and abiotic reactions. Science 340, 85–87 (2013).
Bender, K. S. et al. Identification, characterization, and classification of genes encoding perchlorate reductase. J. Bacteriol. 187, 5090–5096 (2005).
Youngblut, M. D. et al. Perchlorate reductase is distinguished by active site aromatic gate residues. J. Biol. Chem. 291, 9190–9202 (2016).
Okeke, B. C., Giblin, T. & Frankenberger, W. T. Reduction of perchlorate and nitrate by salt tolerant bacteria. Environ. Pollut. https://doi.org/10.1016/S0269-7491(01)00288-3 (2002).
He, L. et al. Biological perchlorate reduction: which electron donor we can choose?. Environ. Sci. Pollut. Res. 26, 16906–16922 (2019).
Xie, T. et al. Perchlorate bioreduction linked to methane oxidation in a membrane biofilm reactor: performance and microbial community structure. J. Hazard. Mater. https://doi.org/10.1016/j.jhazmat.2018.06.011 (2018).
Chaudhuri, S. K., O’Connor, S. M., Gustavson, R. L., Achenbach, L. A. & Coates, J. D. Environmental factors that control microbial perchlorate reduction. Appl. Environ. Microbiol. https://doi.org/10.1128/AEM.68.9.4425-4430.2002 (2002).
Abu-Omar, M. M. Effective and catalytic reduction of perchlorate by atom transfer-reaction kinetics and mechanisms. Comments Inorg. Chem. 24, 15–37 (2003).
Adkins, H. & Cramer, H. I. The use of nickel as a catalyst for hydrogenation. J. Am. Chem. Soc. 52, 4349–4358 (1930).
Thauer, R. K. et al. Hydrogenases from methanogenic archaea, nickel, a novel cofactor, and H2 storage. Annu. Rev. Biochem. 79, 507–536 (2010).
Zhang, H., Bruns, M. A. & Logan, B. E. Perchlorate reduction by a novel chemolithoautotrophic, hydrogen-oxidizing bacterium. Environ. Microbiol. https://doi.org/10.1046/j.1462-2920.2002.00338.x (2002).
Ide, T., Bäumer, S. & Deppenmeier, U. Energy conservation by the H2: heterodisulfide oxidoreductase from methanosarcina mazei Gö1: identification of two proton-translocating segments. J. Bacteriol. 181, 4076–4080 (1999).
Deppenmeier, U. The membrane-bound electron transport system of methanosarcina species. J. Bioenerg. Biomembr. 36, 55–64 (2004).
Meuer, J., Kuettner, H. C., Zhang, J. K., Hedderich, R. & Metcalf, W. W. Genetic analysis of the archaeon Methanosarcina barkeri Fusaro reveals a central role for Ech hydrogenase and ferredoxin in methanogenesis and carbon fixation. Proc. Natl. Acad. Sci. 99, 5632–5637 (2002).
Kulkarni, G., Mand, T. D. & Metcalf, W. W. Energy Conservation via Hydrogen Cycling in the Methanogenic Archaeon Methanosarcina barkeri. MBio 9, (2018).
Bobik, T. Formyl-methanofuran synthesis in Methanobacterium thermoautotrophicum. FEMS Microbiol. Lett. 87, 323–326 (1990).
Wang, D. M., Shah, S. I., Chen, J. G. & Huang, C. P. Catalytic reduction of perchlorate by H2 gas in dilute aqueous solutions. Sep. Purif. Technol. 60, 14–21 (2008).
Thauer, R. K., Kaster, A.-K., Seedorf, H., Buckel, W. & Hedderich, R. Methanogenic archaea: ecologically relevant differences in energy conservation. Nat. Rev. Microbiol. 6, 579–591 (2008).
Mand, T. D. & Metcalf, W. W. Energy Conservation and Hydrogenase Function in Methanogenic Archaea, in Particular the Genus Methanosarcina. Microbiol. Mol. Biol. Rev. 83, (2019).
Rummel, J. D. et al. A new analysis of mars “special regions”: findings of the second MEPAG special regions science analysis group (SR-SAG2). Astrobiology 14, 887–968 (2014).
Bryant, M. P. & Boone, D. R. Emended description of strain MST(DSM 800T), the type strain of methanosarcina barkeri. Int. J. Syst. Bacteriol. 37, 169–170 (1987).
Widdel, F., Kohring, G.-W. & Mayer, F. Studies on dissimilatory sulfate-reducing bacteria that decompose fatty acids. Arch. Microbiol. 134, 286–294 (1983).
Francisco, D. E., Mah, R. A. & Rabin, A. C. Acridine orange-epifluorescence technique for counting bacteria in natural waters. Trans. Am. Microsc. Soc. 92, 416 (1973).
Chen, S., Zhou, Y., Chen, Y. & Gu, J. fastp: an ultra-fast all-in-one FASTQ preprocessor. Bioinformatics 34, i884–i890 (2018).
Langmead, B. & Salzberg, S. L. Fast gapped-read alignment with Bowtie 2. Nat. Methods 9, 357–359 (2012).
Li, H. et al. The sequence alignment/map format and SAMtools. Bioinformatics 25, 2078–2079 (2009).
Quinlan, A. R. & Hall, I. M. BEDTools: a flexible suite of utilities for comparing genomic features. Bioinformatics 26, 841–842 (2010).
Camacho, C. et al. BLAST+: architecture and applications. BMC Bioinformatics 10, 421 (2009).
Love, M., Anders, S. & Huber, W. Differential analysis of count data–the DESeq2 package. Genome Biol. 15, 10–1186 (2014).
Ogata, H. et al. KEGG: kyoto encyclopedia of genes and genomes. Nucleic Acids Res. 27, 29–34 (1999).
This research was supported by NASA Exobiology Grant NNX17AK87G to ACS, TCO, and RLH. RLH was also supported by National Science Foundation grant DGE-1148900. YT was funded by the Princeton Environmental Institute as part of the 2019 PEI summer internship program. The authors thank Dr. Renxing Liang for obtaining cation measurements and the staff of the Princeton University genomics core facility for performing library prep and RNA-Seq. A special thank you goes to Dr. Xinning Zhang, Katja Luxem, and Dr. Peter Girguis for sharing thoughtful insights which helped inform interpretations made in this manuscript.
The authors declare no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Harris, R.L., Schuerger, A.C., Wang, W. et al. Transcriptional response to prolonged perchlorate exposure in the methanogen Methanosarcina barkeri and implications for Martian habitability. Sci Rep 11, 12336 (2021). https://doi.org/10.1038/s41598-021-91882-0