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Methanogenesis in the presence of oxygenic photosynthetic bacteria may contribute to …

Abstract

Accumulating evidences are challenging the paradigm that methane in surface water primarily stems from the anaerobic transformation of organic matters. Yet, the contribution of oxygenic photosynthetic bacteria, a dominant species in surface water, to methane production remains unclear. Here we show methanogenesis triggered by the interaction between oxygenic photosynthetic bacteria and anaerobic methanogenic archaea. By introducing cyanobacterium Synechocystis PCC6803 and methanogenic archaea Methanosarcina barkeri with the redox cycling of iron, CH4 production was induced in coculture biofilms through both syntrophic methanogenesis (under anoxic conditions in darkness) and abiotic methanogenesis (under oxic conditions in illumination) during the periodic dark-light cycles. We have further demonstrated CH4 production by other model oxygenic photosynthetic bacteria from various phyla, in conjunction with different anaerobic methanogenic archaea exhibiting diverse energy conservation modes, as well as various common Fe-species. These findings have revealed an unexpected link between oxygenic photosynthesis and methanogenesis and would advance our understanding of photosynthetic bacteria’s ecological role in the global CH4 cycle. Such light-driven methanogenesis may be widely present in nature.

Introduction

Atmospheric methane (CH4), one of the most important greenhouse gases, reached an exceptionally high concentration of 1912 part per billion in 20221,2. This calls for an immediate action to understand and address CH4 emission problems. Freshwater ecosystems such as rivers, streams, lakes, oceans, and wetlands, play a vital role in contributing to the global atmospheric CH4 budget3. It is widely recognized that CH4 in freshwater ecosystems is primarily produced via the transformation of organic matters in anoxic profundal and littoral sediments4,5. Nevertheless, despite the limited exchange between the oxic surface layers of freshwater ecosystems and sediments due to the deep water columns, a prevalent CH4 supersaturation was observed6. This unexpected phenomenon, also known as the methane paradox wherein methane concentrations exceed atmospheric equilibrium values, suggests the existence of a significant CH4 production process that has yet to be defined.

Photosynthetic bacteria hold a dominant presence in the surface layers of freshwater ecosystems and exhibit excellent phototactic motility and versatile metabolic patterns7. Their interaction with other coexisting microorganisms significantly influences the biogeochemical cycle of elements via harnessing solar light as an energy source. The correlation between photosynthetic bacteria and CH4 production under illumination has been reported previously8,9, but the underlying mechanisms are yet to be elucidated. It is likely that anoxygenic photosynthetic bacteria act as photosensitizers, driving the CO2-to-CH4 conversion with anaerobic methanogenic archaea when being cocultured in an anoxic layer10. The role of oxygenic photosynthetic bacteria in the context of CH4 supersaturation is largely unknown. This oversight arises from the traditional belief that methanogenic archaea are highly sensitive to oxygen exposure and can only thrive in highly reduced, anoxic environments11. However, the coexistence of oxygenic photosynthetic bacteria and anaerobic methanogenic archaea occurs in various natural habitats, such as microbial mats, soil crusts, and aerobic epilimnion of an oligotrophic lake12,13,14. The in situ detection of the close attachment between methanogenic archaea and photosynthetic bacteria in these oxygenated and methane-rich environment, along with the finding that methanogens can survive oxygen exposure15, suggested their potential interactions through direct nutrient exchange or signal transduction16,17. Thus, a comprehensive understanding of photosynthetically regulated CH4 production is of ecological and biogeochemical importance, and will offer valuable insights into global CH4 cycle with implications for climate change.

Here, we demonstrated the methanogenesis involved in the coculture of Cyanobacterium Synechocystis sp. strain PCC6803 (hereafter PCC6803) and Methanosarcina barkeri (hereafter M. b). PCC6803 is a model oxygenic photosynthetic bacterium that can perform solar energy conversion of water and CO2 to carbohydrates and oxygen. In the absence of light, the produced carbohydrates are metabolized to generate CO2 and ATP through a respiratory system, creating an anoxic microenvironment suitable for microbial methanogenesis18. Meanwhile, M. b as a model methanogen was chosen owing to its widespread environmental presence with physiological and metabolic diversity19. It is reported that iron exists in many open water systems and is quantitatively the most important trace metal in photosynthetic bacteria20. Over 99% of the dissolved Fe pool is complexed by organic ligands21. Therefore, Fe-ethylenediaminetetraacetic acid (Fe-EDTA) was selected as a typical iron species in this study due to its stability and solubility in aqueous solutions. The results showed that CH4 production by the interaction of oxygenic photosynthetic bacteria and anaerobic methanogenic archaea was significantly enhanced through the redox cycling of Fe-ethylenediaminetetraacetic acid (Fe-EDTA), involving both syntrophic methanogenesis and abiotic methanogenesis during the periodic dark-light cycles (Fig. 1). Specifically, in darkness, the organics and H2 produced by PCC6803 during dark fermentation were utilized as carbon sources and reducing equivalents by M. b for syntrophic methanogenesis under anoxic conditions. The significantly lowered hydrogen pressure by M. b, in turn, created more thermodynamically favorable conditions for PCC6803. In contrast, in illumination, the photosynthesized organic compounds and intermediate products by PCC6803 served as potential methyl donors (-CH3). Meanwhile, the simultaneously produced O2 stimulated reactive oxygen species (ROS) production by M. b. Along with the Fenton reaction with Fe-EDTA, various methyl donors were oxidized by ROS to form methyl radicals (•CH3) as intermediates that eventually resulted in abiotic methanogenesis under oxic conditions. Further studies indicated that other model oxygenic photosynthetic bacteria and anaerobic methanogenic archaea were also able to conduct this light-driven methanogenesis process. These findings not only unveil an unexpected link between oxygenic photosynthesis and methanogenesis, but also advance our understanding of the ecological role of photosynthetic bacteria in the global CH4 cycle.

Fig. 1: Schematic illustration of CH4 production in the presence of oxygenic photosynthetic bacteria.
figure 1

a Syntrophic methanogenesis under anoxic conditions in darkness. b Abiotic methanogenesis under oxic conditions in illumination.

Results

Light-driven methanogenesis with PCC6803 and M. b

PCC6803 and M. b were cocultured in a defined medium (Supplementary Table 1), where CO2 served as the only electron acceptor. The visible light LEDs (12 ± 0.6 W m-2) over the wavelength range of 380–800 nm was used as a simulated sunlight source (Supplementary Fig. 1). The experiment was conducted under a light-dark cycle of 4 hours of light and 20 hours of darkness at 35 oC but some tests were performed with a light-dark cycle of 12 h-12 h that simulates a full day. Compared to the pure M. b control that had a negligible CH4 production under the same condition, the coculture of PCC6803 and M. b achieved a higher CH4 yield of 1.0 ± 0.1 µmol. The addition of Fe-EDTA (hereafter PCC6803-M. b-Fe-EDTA) further enhanced the yield to 2.5 ± 0.5 µmol (Fig. 2a), with a linear correlation between the CH4 yield and the Fe-EDTA concentration (Supplementary Fig. 2). The single-factor experiments showed that the removal of any following components: PCC6803, M. b, Fe-EDTA or light, would result in a significant declination of methanogenesis performance, demonstrating the importance of each component in this process (Fig. 2a, Supplementary Fig. 3). Notably, the CH4 yield with PCC6803-M. b-Fe-EDTA increased both under illumination and in darkness (Fig. 2b), different from the light-dependent CH4 production with anoxygenic photosynthetic bacteria and methanogenic archaea that was only activated by light but inhibited in darkness10. This difference might be attributed to the production of O2 by PCC6803 with H2O as electron donors under illumination, thereby stimulating the generation of ROS by M. b. These ROS, in turn, could oxidize organic matters to create potential carbon sources for biotic methanogenesis with M. b in the dark (see detailed discussion below). The CH4 production rate with PCC6803-M. b-Fe-EDTA continuously increased during the three successive cycles (Fig. 2c), contributing to the rapid growth and formation of a stable syntrophic coculture for methanogenesis (Supplementary Fig. 4).

Fig. 2: Light-driven methanogenesis in a light-dark cycle.
figure 2

a CH4 yields by PCC6803-M. b-Fe-EDTA and controls. b Typical time course of CH4 yield in the first 4 days. c CH4 yields and yield rates during the three successive 18-day cycles. d Mass spectrometry of headspace gases with 13C-labeled NaHCO3 as a sole carbon source. e Periodic variation of dissolved oxygen (DO) concentration. f Evaluation of the gene copy numbers of mcrA and cpcG. The gray color in b and e presents the dark period during the light-driven methanogenesis under a light-dark cycle of 4 h-20 h. Data are presented as mean values ± SD derived from n = 3 independent experiments. Statistical analysis was conducted with paired two-tailed t tests, and different letters represent statistically significant difference (P < 0.05) in different groups. All P values are provided in the source data. Source data are provided as a Source Data file.

To understand the source of CH4 produced by PCC6803-M. b-Fe-EDTA, we performed isotopic labeling experiments. The selected m/z = 45 (13CO2) and 17 (13CH4) were detected when 13C-labeled NaHCO3 was used as the sole carbon source (Fig. 2d). This provided direct evidence that the produced CH4 was derived from the CO2 conversion. Unexpectedly, the relatively weak signals at m/z = 44 (12CO2) and 16 (12CH4) were also detected, indicating the existence of other CH4 production pathways that might utilize the carbon sources synthesized during the initial cultivation process, such as biomass and oxidation intermediates (see detailed discussion below). A noticeable periodic variation in the dissolved oxygen (DO) concentration was observed in the coculture during the light-dark cycles (Fig. 2e), likely attributed to the alternate photosynthetic oxygen evolution and respiratory oxygen consumption22. The addition of Fe-EDTA would reduce the oxygen concentration under illumination and expedite the oxygen consumption in darkness, through the oxygen oxidation with Fe(II) chelated by EDTA, resulting in an anaerobic microenvironment for the growth and metabolism of M. b. This hypothesis was confirmed by the increased copy number of mcrA gene, which is ubiquitous in M. b during the methanogenesis with PCC6803-M. b-Fe-EDTA (Fig. 2f), along with a simultaneous rise in the copy number of cpcG that referred to as rod-core linker genes in PCC 6803. However, the growth of PCC6803 led to an enhanced photosynthetic oxygen evolution, elevating DO accumulation in the light and extending the time required to establish an anaerobic microenvironment for biotic methanogenesis in darkness (Fig. 2e). As a result, the rate of CH4 production with PCC6803-M. b-Fe-EDTA slowed down somewhat after day 2 but persisted (Fig. 2b).

Syntrophic methanogenesis by organic degradation and CO2 reduction

The close contact between different microorganisms can enhance the interspecies exchange of matter and energy to shape specific communities16. As shown in Fig. 3a, PCC6803 and M. b were connected to form dense PCC6803-M. b biofilm at the bottom of the culture bottles after 10 days of coculturing. Both optical microscopy images (Fig. 3b) and fluorescence in situ hybridization (FISH) images (Fig. 3c) showed the aggregation of PCC6803 and M. b cells in the biofilm. An increase in the biofilm thickness from 30 μm on day 0 to 45 μm on day 10 was observed by confocal laser scanning microscopy (CLSM) (Supplementary Fig. 5), demonstrating the growth of both microbes in the coculture biofilms.

Fig. 3: Characterization of coculture biofilm and produced organic substances.
figure 3

a Dense biofilm in culture bottle (the inset image shows the picture of culture bottle). Optical microscopy image (b) and FISH image (c) with M. b (green-fluorescing probe) and PCC6803 (red, autofluorescence); representative of 10 images. d Schematic illustration for biotic methanogenesis driven by intermediates with PCC6803-M. b-Fe-EDTA. e Possible organic substances generated via ROS oxidation. gCOSY (f) and gHMBC (g) superimposed aqueous 600-MHz NMR spectra of supernatants after the syntrophic coculture for 6 days (black), 0.1 M lactate (green), 0.1 M acetate (red), 0.1 M pyruvate (blue), and 0.1 M methanol (orange yellow). The red circled peak in (g) is assigned to acetate, the blue circled peak is assigned to pyruvate, and the orange yellow circled peak is assigned to methanol. h Variation of acetate concentration under a light-dark cycle of 4 h–20 h. The gray color presents the dark period. Data are presented as mean values ± SD derived from n = 3 independent experiments. Scale bars: 1 μm in (a), 10 μm in (b), and 10 μm in (c). Source data are provided as a Source Data file.

PCC6803 can synthesize carbohydrates via photosynthesis, and then degrade these carbohydrates through a respiration process in darkness, excreting a variety of organic matters23. These organic matters, along with their intermediates produced via the oxidative degradation by the concomitant reactive oxygen species (ROS), would serve as the potential carbon sources for biotic methanogenesis with M. b (Fig. 3d). Lactate, pyruvate and acetate were detected by the 1H nuclear magnetic resonance (NMR) spectra (Fig. 3e). These findings were further confirmed by two-dimensional NMR spectra, which provide the crucial through-bond correlations existing between the coupled protons [two-dimensional gradient-selected homonuclear correlation spectroscopy (gCOSY) in Fig. 3e] and between protons and carbons via multiple-bond correlations [gradient-selected heteronuclear multiple bond correlation (gHMBC) in Fig. 3f] for each compound24. The 1H and 13C chemical shifts of lactate, pyruvate, methanol, and acetate correspond well to values from known library spectra (PubChem Database), such as the chemical shifts of 1H (1.96 ppm) and 13C (176.5 ppm) in acetate, confirming the presence of these compounds in aqueous solution during syntrophic methanogenesis. Although pyruvate was reported to be the sole carbon and energy source for the growth of M. b25, this species is more efficient in converting acetate to CH4 (CH3COOH → CH4 + CO2) due to less energy requirement for acetate activation26, resulting in a lower acetate yield with PCC6803-M. b-Fe-EDTA than that with pure PCC6803 (Fig. 3h).

In addition to organic substances, H2 was also detected in the headspace with PCC6803-M. b-Fe-EDTA (Supplementary Fig. 6). These H2 molecules could originate from the direct (proton reduction with hydrogenase) and indirect (hydrogen release from the carbohydrate degradation) biophotolysis, and serve as electron donors for CH4 production via a hydrogenotrophic pathway (CO2 + 4H2 → CH4 + 2H2O)27. Compared with pure PCC6803, a lower H2 yield was detected with PCC6803-M. b-Fe-EDTA after 18 days of coculture (Supplementary Fig. 6), which is probably because that H2 produced by PCC6803 was used as electron donors for methanogenesis. To confirm this argument, sodium 2-bromoethanesulfonate (SBES) was added to the coculture medium to inhibit hydrogenases in methanogenic archaea. More H2 residue and a lower CH4 yield indicated that the hydrogenotrophic pathway was existed but inhibited by SBES in PCC6803-M. b-Fe-EDTA (Supplementary Fig. 7). However, it should be noted that PCC6803 contains a single [NiFe]-hydrogenase, HoxEFUYH, which is involved in fermentative hydrogen production as well as working as an electron valve when photosynthesis resumes under anaerobic conditions28. Notably, HoxEFUYH works bidirectionally with a bias to proton reduction rather than hydrogen oxidation29. Therefore, an increase in hydrogen pressure due to hydrogen accumulation during dark fermentation might result in a significant decrease in hydrogenase activity, thereby influencing the growth and metabolism of PCC680330. This inference was confirmed by the lower chlorophyll concentration, quantum yield of PSII primary photochemical reactions (Fv/Fm), and copy number of cpcG in bare PCC6803, along with a higher H2 concentration compared with syntrophic methanogenesis (Supplementary Fig. 8). The superior activity of PCC6803 during syntrophic methanogenesis were attributed to the versatile metabolic pathways of M. b, which significantly lowered hydrogen pressure via CO2/H2 methanogenesis (Supplementary Fig. 7), and created more thermodynamically favorable conditions for the dark fermentation of PCC6803.

A distinct photocurrent was also observed under illumination during syntrophic methanogenesis (Supplementary Fig. 9a), indicating that PCC6803 can release electrons extracellularly. Although M. b served as electron acceptors capable of accepting photoelectrons from PCC6803 for CO2 reduction (CO2 + 8e + 8H+ → CH4 + 2H2O)31,32, the syntrophic methanogenesis via a direct interspecies electron transfer (DIET) pathway was less likely to occur with PCC6803-M. b-Fe-EDTA under illumination. This was because that the growth and metabolism of M. b is highly sensitive to the oxygen exposure during the photosynthetic oxygen evolution. Besides the extracellular electron transfer for Fe(III) reduction, the light-induced electrons by PCC6803 would form excited triplet state of chlorophyll in the photosystem II reaction center, which then interacts with molecular oxygen for singlet oxygen (1O2) production33, eventually leading to the production of other ROS for abiotic methanogenesis (see detailed discussion below). In contrast, previous studies have shown that photosynthetic microorganisms can also generate an electrical current exclusively in darkness, using illumination as a recharge stage34,35. To validate this, two-chamber H-cells were constructed, with PCC6803 and M. b separately inoculated into each chamber and then electrically connected by an external circuit, to mitigate the influence of direct electron exchange between M. b and PCC6803 on the photoelectron measurement. Similar results were observed that a continuous current was recorded during its dark fermentation period, albeit with lesser intensity compared to that in the light (Supplementary Fig. 9b). This finding indicates the potential DIET between PCC6803 and M. b via an electric syntrophic coculture in darkness.

Electron flow and energy metabolisms at the genetic level

Syntrophic methanogenesis with PCC6803-M. b-Fe-EDTA was further confirmed by transcriptomic analyses (Fig. 4). The transcript levels of PSII and cytochrome b6f complex (cytb6f) exhibited similar oscillation patterns that were highly upregulated during the light period. With plastocyanin and cytc553 in the thylakoid lumen as alternate donors, the produced electrons would further transfer from cytb6f to PSI, resulting a higher expression of genes encoding PSI. Meanwhile, ATP synthase (ATPase) and ferredoxin-NADP+ reductase (FNR) were upregulated during the light period and downregulated during the dark period. The upregulation during the light period was expected, as all transcripts associated with photosynthetic electron transport and ATP synthesis were upregulated during this phase. In addition, most of the CO2 fixation machinery was highly active in the light period. For example, the key genes for the Calvin cycle, such as RubisCO (rbc), phosphoribulokinase (prk) and glyceraldehyde-3-phosphate dehydrogenase (gap)36, were significantly upregulated to ensure the efficient CO2 fixation for producing carbohydrates (Fig. 4, Supplementary Fig. 10). The accumulated carbohydrates, acting as storage molecules, then serve as carbon sources for microbial metabolism. The oxidation of carbohydrates was further confirmed by the upregulated transcript level of genes encoding the tricarboxylic acid (TCA) cycle, such as citrate synthase (glt), isocitrate dehydrogenase (icd), synthetase (suc) and malate dehydrogenase (mdh) (Fig. 4, Supplementary Fig. 11). Likewise, genes related to the biosynthesis of lactate and acetate were significantly upregulated, which could be oxidized by ROS and used as substrates by M. b. Remarkably, the iron uptake and transport (ferric uptake (Fut) and ferrous iron transport (Feo) systems) were also enhanced with higher gene expression, particularly in the light period, thereby facilitating the Fe redox process during syntrophic methanogenesis.

Fig. 4: Mechanisms of biotic methanogenesis with PCC6803-M. b-Fe-EDTA as revealed by transcriptomic analyses.
figure 4

Genes involved in processes include those that encode (1) Calvin cycle; (2) 2;3-bisphosphoglycerate-dependent phosphoglycerate mutase Pgm; (3) enolase Eno; (4) Pyruvate kinase Pyk; (5) Phosphoenolpyruvate carboxylase Ppc; (6) TCA cycle; (7) malate dehydrogenase Mae; (8) D/L-lactate dehydrogenase D/L-LDH; (9) pyruvate-ferredoxin oxidoreductase Por; (10) acetate kinase Ack; (11) phosphate acetyltransferase Pta; (12) CO dehydrogenase/acetyl-CoA synthase Codh/Acs; (13) formylmethanofuran dehydrogenases Fmd; (14) formylmethanofuran-tetrahydromethanopterin N-formyltransferase Ftr; (15) methenyltetrahydrome-thanopterin cyclohydrolase Mch; (16) methylenetetrahydromethanopterin dehydrogenase Mtd; (17) methylenetetrahydromethanopterin reductase Mer; (18) methyltransferase subunit Mtr; (19) methyl-CoM reductase Mcr. (20) ferric uptake (Fut) and ferrous iron transport (Feo) systems. PSII photosystem II, Cyt b6f cytochrome b6f complex, PSI photosystem I, FNR ferredoxin–NADP+ reductase, NADH nicotinamide adenine dinucleotide phosphate, ATPase ATP synthase, DHAP dihydroxyacetone phosphate, RuBR ribulose-1,5-biphosphate, G3p glyceraldehyde 3-phosphate, Pep phosphoenolpyruvate, OAA oxaloacetate, AcCoA acetyl coenzyme A, Acetyl-P acetylphosphate, Fpo membrane-bound F420H2 dehydrogenase, Hdr heterodisulfide reductase, Hox bidirectional hydrogenase complex protein. Source data are provided as a Source Data file.

In darkness, genes responsible for the complete pathways for oxidation of organic compounds (i.e., acetate and pyruvate) and CO2 reduction were also highly expressed, suggesting the existence of multiple CH4 production pathways. For instance, the significantly increased activity of pyruvate-ferredoxin oxidoreductase (Por) could oxidize pyruvate to AcCoA and CO2, and the subsequent conversion of AcCoA to CH4 would occur via the native aceticlastic pathway37. Notably, the transcriptional levels of genes encoding functional hydrogenases for H2 production and consumption (e.g., energy-converting [NiFe]-hydrogenase (Ech), methanophenazine-reducing [NiFe]-hydrogenase (Vht) and F420H2 dehydrogenase (Fpo))38 were upregulated, implying that the CO2 reduction was conducted via H2 transfer during the dark period.

Abiotic methanogenesis via the oxidation of methyl donors by ROS

Substantial CH4 production during the light period was observed with PCC6803-M. b-Fe-EDTA (Fig. 2b). However, deletional control experiment revealed that PCC6803 produced almost no CH4 under illumination without the presence of other factors (Supplementary Fig. 3), suggesting the potential existence of an abiotic CH4 production process. ROS are vital cellular metabolic products found in all living organisms. They are responsible for oxidizing the methyl groups (-CH3) of methyl donors and can play a crucial role in modulating chemical CH4 formation (Fig. 5a). Electron paramagnetic resonance (EPR) spectra revealed clear signals of 5,5-dimethyl-1-pyrroline-N-oxide (DMPO)/•OH, DMPO/superoxide anion radicals (•O2) and 2,2,6,6-tetramethylpiperidine-1-oxyl (TEMPO)/1O2 from PCC6803-M. b-Fe-EDTA (Fig. 5b, Supplementary Fig. 12). In addition, H2O2 was produced as shown in the ultraviolet–visible (UV–Vis) absorption spectra (Fig. 5c). The concentrations of these ROS, particularly •OH and H2O2, significantly increased under light illumination (Fig. 5d), likely attributed to the produced O2 by PCC6803 during the photosynthetic oxygen evolution that significantly stimulated the ROS production by M. b. This inference was confirmed through the ROS production experiments with M. b under varying O2 concentrations. It was found that higher O2 concentrations led to the production of more ROS, such as •OH and H2O2 (Supplementary Fig. 13). Stable isotope analysis with 18O2 further confirmed that the produced ROS stemmed from O2 reduction, evidenced by the observed 5,5-dimethyl-1-pyrroline-N-oxide (DMPO)-18OH (m/z = 132.09, Supplementary Fig. 14). Previous studies have shown that when anaerobic cells were exposed to oxygen-rich environments, molecular O2 adventitiously abstracted electrons from the reduced flavins or metal centers of some redox enzymes, resulting in the ROS formation (O2 → •O2 → H2O2 → •OH)39,40,41. As these events relied on collision frequency, the ROS production rate was directly proportional to the O2 concentration42. Subsequently, the produced ROS were inadvertently released extracellularly43. Meanwhile, ROS could also be produced by PCC6803 during the photochemical energy conversion for bioenergetic production44. These ROS could be further used for -CH3 and Fe(II) oxidization. Particularly, the light-driven H2O2 may also interact with EDTA chelated-Fe(II) for •OH production via Fenton reaction (Fe(II) + H2O2 → Fe(III) + •OH), leading to an increasing Fe(III) concentration under illumination in the first light-dark cycle (Fig. 5e). The produced Fe(III) through the oxidation of ROS and O2 could be reduced by PCC6803 (Supplementary Fig. 15), either through intracellular metabolism or extracellular electron transfer, and finally established an effective Fe(III)/Fe(II) redox cycle.

Fig. 5: Abiotic methanogenesis with PCC6803-M. b-Fe-EDTA.
figure 5

a Illustration of abiotic CH4 production with PCC6803-M. b-Fe-EDTA. b EPR spectra of •CH3, •OH, and •O2. c UV/Vis absorption spectra for H2O2 with o-tolidine as the peroxide indicator. d − e Typical time course of the concentrations of H2O2 and •OH (d), and Fe(II) and Fe(III) (e). The gray color presents the dark period. f Characterization of DMS and DMSP as methyl donors (the inset image shows the mass spectrometry of DMS). Data are presented as mean values ± SD derived from n = 3 independent experiments. Source data are provided as a Source Data file.

EPR spectra also revealed the prominent signals of DMPO/•CH3 (Fig. 5b). The potential sources of methyl donors for •CH3 production with PCC6803-M. b-Fe-EDTA were diverse. On one hand, PCC6803 could synthesized organic compounds containing sulfur-bonded methyl groups, such as pyruvate and ethanol (Fig. 3g). Meanwhile, the existence of DMS was confirmed by gas chromatography-mass spectrometry (GC-MS) with an ion signal at m/z = 62, along with the retention time of 8.7 min in 400 MHz 1H nuclear magnetic resonance (NMR) spectra (Fig. 5f). The DMS production might be partly attributed to the ROS oxidation of DMSP, because the DMSP was confirmed by the increasing intensity of DMS after alkali treatment for 12 h via the DMSP-to-DMS conversion45. On the other hand, the possible release of the intracellular metabolites by M. b, such as 2-(methylthio)ethanesulfonic acid (CH3-S-CoM), might also serve as potential sources of methyl donors. These methyl donors could be oxidized by ROS for •CH3 formation, ultimately leading to abiotic CH4 production46,47. The role of ROS was further examined by the scavenger trapping tests, which revealed that the methanogenesis process was significantly inhibited after the addition of ROS quenching reagents (Supplementary Fig. 16). The CH4 yield with PCC6803-M. b-Fe-EDTA was decreased from 2.5 µmol to 0.5 µmol with the •OH scavenger of tert-butyl alcohol (TBA, 10 mmol-1). The results were confirmed by previous studies in which abiotic CH4 production occurred with highly reactive •OH generated not only through Fenton reaction48,49 but also through ubiquitous non-Fenton chemistry reactions driven by diversified external fields50. Because •OH is involved in both the organic degradation and -CH3 oxidization, the difference in the CH4 yield with and without the •OH scavenger suggested that the contribution of abiotic methanogenesis (i.e., methyl donors-to-CH4 conversion) to total CH4 production would be around 65.4%, and the remaining 34.6% of CH4 came from syntrophic methanogenesis (i.e., organics/CO2-to-CH4 conversion). In addition, the ROS-induced •CH3 could also combine with O2 under illumination47, contributing to the formation of CH3OH as a carbon source for biotic methanogenesis in darkness.

Notably, excessive ROS accumulation has been reported to induce oxidative stress, causing damage to cellular components such as DNA, proteins, and lipids, ultimately inhibiting microbial growth and survival51. However, ROS were effectively consumed during the light-driven methanogenesis for the oxidation of -CH3 and organic compounds, as well as Fe(II) oxidation, thereby suppressing their accumulation, particularly in the dark (as shown in Fig. 5d). Additionally, the detoxification systems in both microorganisms, including superoxide dismutase, catalases, and peroxidases, were found to be significantly upregulated (Supplementary Fig. 17), effectively alleviating the potential oxidative stress and toxicity for M. b and PCC6803 during syntrophic methanogenesis.

Universality of the light-driven methanogenesis in nature

The investigation of the light-driven methanogenesis was conducted to other oxygenic photosynthetic bacteria and anaerobic methanogenic archaea (Fig. 6). In addition to PCC6803 (Cyanophyta), Tribonema minus (Xanthophyta), Euglena gracilis (Euglenophyta) and Chlorella sp. (Chlorophyta) from various phyla also produced CH4 with M. b. Furthermore, similar methanogenesis was observed when M. b was replaced with Methanobacterium bryantii (hydrogenotrophic methanogen), Methanococcoides orientis (methylotrophic methanogen), or Methanosphaera stadtmaniae (aceticlastic methanogen). These results suggest that the light-driven methanogenesis process with cocultures is not dependent on the energy conservation types of methanogenic archaea. We also found that various common Fe-species, such as zero-valent iron, ferric citrate and ferric oxalate, could enhance the performance of light-driven methanogenesis. Therefore, these results have provided strong evidence that the methanogenesis by oxygenic photosynthetic bacteria and anaerobic methanogenic archaea may be a prevalent occurrence in nature.

Fig. 6: Common light-driven methanogenesis by photosynthetic bacteria and methanogenic archaea.
figure 6

a Simplified tree of influence factors on light-driven methanogenesis. b Light-driven methanogenesis by different oxygenic photosynthetic bacteria, anaerobic methanogenic archaea, and Fe-species (zero-valent iron (ZVI), Fe3O4 and FeCl2). C. sp. represents Chlorella sp. (Chlorophyta), E. sp. represents Euglena gracilis (Euglenophyta), and T. m represents Tribonema minus (Xanthophyta). M. b represents Methanosarcina barkeri, M. br represents Methanobacterium bryantii, M. o represents Methanococcoides orientis, M. s represents Methanosphaera stadtmaniae. Data are presented as mean values ± SD derived from n = 3 independent experiments. Statistical analysis was conducted with paired two-tailed t tests: *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001. All P values are provided in the source data. Source data are provided as a Source Data file.

Discussion

Unlike the previous studies that CH4 production by oxygenic photosynthetic bacteria could be progressed by the demethylation of methylphosphonates or the conversion of fixed inorganic carbon into CH452,53, this work has elucidated an unappreciated but potentially widespread pathway for CH4 production. The alternating phases of photosynthetic oxygen evolution (oxic) and respiratory oxygen consumption (anoxic) are essential for methanogenesis, which could be achieved under varying light times, even a light-dark cycle of 12 h-12 h that simulates a full day (Supplementary Fig. 18). The light-driven methanogenesis experiments were also conducted on the roof of the Research Center for Water Resources and Security Building at Fujian Agriculture and Forestry University in Fuzhou, China (latitude: 26.05 oN, longitude: 119.14 oE) under natural sunlight (from 08:00 to 20:00 with an average solar heat flux of ~0.5 kW m−2), with ambient temperatures ranging between 25 °C and 37 °C. A similar methanogenesis process was also observed (Supplementary Fig. 19). In conclusion, besides the abiotic methanogenesis under illumination, there exists a co-evolved, specific interaction during syntrophic methanogenesis by oxygenic photosynthetic bacteria and anaerobic methanogenic archaea in darkness. Specifically, M. b, in the absence of other cell types except PCC6803, were benefiting from photosynthetic organic matter production. It was estimated that 5.9% of gross primary production was diverted to CH4 formation. Meanwhile, due to the CO2/H2 methanogenesis of M. b, PCC6803 were benefiting from the lowered hydrogen pressure, creating more thermodynamically favorable conditions for the dark fermentation of PCC6803.

Oxygenic photosynthesis has been recognized as the most important metabolic innovation on Earth, enabling life to harness energy and reducing power directly from sunlight and water, thus liberating it from the constraints of geochemically derived reductants54. Consequently, these diverse and intriguing oxygenic photosynthetic bacteria contain considerable metabolic flexibility, utilizing numerous unconventional central carbon metabolic pathways and novel enzymes for autotrophic, mixotrophic, and heterotrophic growth, tailored to their specific ecological niches22,55. Considering the extensive coexistence and interaction of diverse microbial species in natural and engineered ecosystems56, along with ferruginous environment on Earth (e.g., oceans with abundant Fe(II) and Fe(III)-carboxylate complexes), syntrophic methanogenesis by oxygenic photosynthetic bacteria and anaerobic methanogenic archaea creates more thermodynamically favorable conditions for both microorganisms. Thus, this light-driven methanogenesis process, involved both syntrophic methanogenesis (under anoxic conditions in darkness) and abiotic methanogenesis (under oxic conditions in illumination) during the periodic dark-light cycles, surpasses the conventional methane production pathways (i.e., acetoclastic methanogenesis and hydrogenotrophic methanogenesis), and potentially making a more significant contribution to the global CH4 cycle. The inference was supported by the correlation between CH4 supersaturation and photosynthesis8,9,57. Various potential mechanisms for CH4 production by phototrophic microorganisms having been extensively investigated, including the photosynthesis-driven metabolism9,53 and ROS-driven demethylation of methyl donors46,47,48,49. Our study innovatively demonstrated the synergistic interaction between these two mechanisms, along with the Fe redox cycles. However, the existence of such methanogenesis by oxygenic photosynthetic bacteria and anaerobic methanogenic archaea in the natural environments requires further validation with multiple complementary approaches, including the evaluation of in situ CH4 profiles and microbial composition, incubation experiments with freshwater microbial cultures using NaH13CO3 as a supplementation carbon source, and the assessment of the exact contribution of both abiotic and biotic pathways. Meanwhile, recent studies have indicated the potential importance of various metal elements in the evolution of oxygenic photosynthesis, such as manganese58. Therefore, the potential involvement of other metal elements in such light-driven methanogenesis warrants further evaluation.

Methods

Light-driven methanogenesis experiments

PCC6803 was purchased from the China General Microbiological Culture Collection Center, and cultured in BG11 medium under illumination by visible light LEDs (12 ± 0.6 W m-2). M. b MS (DSM 800) was purchased from the German Collection of Microorganisms and Cell Cultures, and cultured in heterotrophic medium modified from DSM311b medium31,32. When PCC6803 and M. b grew to their late exponential stages, cells were collected and washed with 0.9% NaCl solution for three times by centrifuging at 5000 × g for 20 min at 4 °C (Eppendorf AG 5811, Hamburg, Germany). Then, the washed cells were used to initiate coculture by inoculating PCC6803 and M. b cells in 200 mL of autotrophic medium with different concentrations of sterile Fe-EDTA. The suspension was sparged with sterile N2/CO2 (80/20, vol/vol), creating an initial anaerobic environment with CO2 as the sole carbon source, and incubated without illumination for 24 h to promote heterotrophic respiration with the residual carbon sources from the initial microbial cultivation. After that, the PCC6803-M. b-Fe-EDTA coculture was cultivated for CH4 production under a light-dark cycle of 4 h-20 h at 35 oC using visible light LEDs (12 ± 0.6 W m-2) as light sources. For comparison, a series of single-factor experiments were conducted, by removing M. b, PCC6803, Fe-EDTA or light. The coculture in suspension were harvested after 18 days of light-driven methanogenesis, and the feedstocks were then replaced with the fresh sterilized autotrophic medium to start a new cycle (for a total of three cycles). To evaluate the origin of the produced CH4 with PCC6803-M. b-Fe-EDTA, isotopic labeling experiments were conducted with 13C-labeled NaHCO3 as a sole carbon source. Sodium 2-bromoethanesulfonate (SBES) as a hydrogenase inhibitor was added to evaluate the contribution of H2 to the light-driven methanogenesis. ROS production experiments with M. b under varying O2 concentrations (0.0%, 0.5% and 1.0%) were conducted to evaluate the role of M. b in abiotic CH4 production under illumination. Meanwhile, stable isotope experiments using 18O-labeled O2 were carried out to further determine the origin of the produced •OH. The DMPO-18OH, formed through the interaction between DMPO and •18OH, was identified using an ultrahigh-performance liquid chromatography-triple quadrupole mass spectrometer (LC‒MS, TSQ Endura, Thermo Fisher, USA)58.

To evaluate the universality of such light-driven methanogenesis in nature, several other oxygenic photosynthetic bacteria and anaerobic methanogenic archaea were selected. Oxygenic photosynthetic bacteria Tribonema minus (FACHB-2214, medium BG-11), Euglena sp. (FACHB-1862, Medium HUT) and Chlorella sp. (FACHB-5, medium BG-11) were obtained from the collection of Freshwater Algae Culture Collection at the Institute of Hydrobiology (FACHB), China. Anaerobic methanogenic archaea Methanobacterium bryantii (ATCC33272, DSM medium 1523) and Methanosphaera stadtmaniae (CCAM456, DSM medium 322) were purchase from Biogas Institute of Ministry of Agriculture and Rural Affairs, China. The initial inoculum of Methanococcoides orientis (PRJNA718391, DSM medium 141c) was graciously obtained from the laboratory of Prof. Guangyu Li in Third Institute of Oceanography, Ministry of Natural Resources, China. All culture and sampling manipulations were performed using the sterile technique.

Microscopy and fluorescent in situ hybridization

An optical microscope (Nikon Eclipse E200, Japan) was used to examine the cocultured cells grown at their logarithmic phase. Fluorescent in situ hybridization was conducted with PCC6803-M. b cocultured cells that were first fixed in premixed paraformaldehyde (1%) and glutaraldehyde (0.5%), and then dehydrated in a graded ethanol series (30%, 50%, 70%, 80% and 100% for 3 min at 4 °C). Next, the samples were incubated in a UVP HL-2000 HybriLinker hybridization oven at room temperature for 2 h with PCC6803 (red, autofluorescence) and green-fluorescing probes specific for M. b (5’-(FAM)GTGCTCCCCCGCCAATTCCT-3’). Ultimately, the obtained samples were washed for 30 min in washing buffer, rinsed with Milli-Q water, and then visualized with a Carl Zeiss LSM 880 confocal laser scanning microscope. In addition, to investigate the variation of biofilm thickness with PCC6803-M. b-Fe-EDTA during light-driven methanogenesis, a graphite plate was positioned on the bottom of the culture bottle. Then the formed biofilm was stained by a LIVE/DEAD BacLight Bacterial Viability Kit (Invitrogen, CA), and examined with a Carl Zeiss LSM 880 confocal laser scanning microscope.

Quantitative analysis of mcrA and cpcG genes

RT-PCR was performed for the quantitative analysis of mcrA gene in M. b and cpcG gene in PCC6803 via a Roche LightCycler 480 System (Roche Applied Science, Penzberg, Germany). mcrA gene was amplified using the primer pair mcrAFor (5ʹ-GGYGGTGTMGGDTTCACMCARTA-3ʹ) and mcrARev (5ʹ- CGTTCATBGCGTAGTTVGGRTAGT-3ʹ). cpcG gene was amplified using the primer pair cpcGFor (5ʹ-GTCGGGAAGCGGGTGA-3ʹ) and cpcGRev (5ʹ-TTGGCGGCAGGGTTGA-3ʹ). The detailed procedures for RNA collection and quantitative RT-PCR quantification could be found in the Supporting Information59.

Transcriptomic analysis

PCC6803-M. b cocultured cells from triplicate experiments were collected by centrifugation at 12,000 × g for 2 min at 4 °C (Eppendorf AG 5811, Hamburg, Germany). Total RNA was extracted RNAprep pure cell/bacteria Kit (TIANGEN, Beijing, China), and the RNA integrity was assessed using the Agilent RNA Nano 6000 Assay Kit of the Bioanalyzer 2100 system. Library preparation for strand-specific transcriptome sequencing was generated using NEBNext Ultra II Directional RNA Library Prep Kit following the manufacturer’s recommendations. All the raw sequencing data were quality-checked, and the clean reads were obtained by removing sequencing adapters, trimmed ambiguous bases (N) from the start and end, and other low-quality reads. The remaining reads were subsequently used to map against the published genome of PCC6803 (GCF_018845095.1) and M. b (GCF_000970025.1).

Analytical techniques

The concentrations of CH4 and H2 were determined using a Shimadzu Gas Chromatograph (GC-2014, Shimadzu, Japan) equipped with both flame ionization detector (FID) and thermal conductivity detector (TCD), as well as a Porapak Q column (3.00 mm ID, 5.0 m long). Nitrogen gas (purity >99.995%) with a flow rate of 30 mL min-1 was used as the carrier gas. The injector port and detector temperatures were set at 100 °C and 250 °C, respectively. The injection volume was 100 μL, with the detection limits of 0.1 ppm for CH4 and 5 ppm for H2. The gas products during the isotopic labeling experiments were determined by a Shimadzu GC-2010 equipped with a Shimadzu AOC-20i auto sampler system, and interfaced with a Shimadzu QP 2010S mass spectrometer (Shimadzu, Japan) (DB-5 capillary column (30 m×0.25 mm×0.25μm), Helium (99.999%) as carrier gas with a flow rate of 1.2 mL min-1). The DO concentration was measured using UNISENSE OX-NP oxygen needle sensor with a detection limit of 0.3 μM, which was calibrated according to the manufacturer’s instructions.

The photocurrents (I-t) were initially measured in a single-chamber microbial fuel cell (MFC) by applying an external potential bias of -0.5 V. Carbon cloth with a size of 3 cm × 3 cm was used as the working electrodes, and autotrophic medium with EDTA-Fe was employed as electrolyte. The cell had a liquid volume of 200 mL and a headspace volume of 150 mL. Data were recorded every 1 min by a data acquisition system (model 2700, Keithley Instruments, Ohio, USA). In addition, photocurrent measurement was also conducted under identical condition, but using two-chamber H-cells instead of the single-chamber MFC10,60. A proton exchange membrane (Nafion 117, DuPont Co., USA) was used to separate the anodic and cathodic chambers, along with autotrophic medium as electrolyte. To examine the compositions of organic substances produced with PCC6803-M. b, the coculture cells were centrifuged at 15,000 × g for 20 min at 4 °C (Eppendorf AG 5811, Hamburg, Germany), and then the supernatant was characterized by the Varian INOVA 600-MHz NMR spectroscopy via 1H NMR, gCOSY and gHMBC. The concentration of acetate was monitored by a Shimadzu Nexis GC-2030 equipped with an FID detector and a 10 m × 0.53 mm HP-FFAP fused-silica capillary column. Nitrogen was used as the carrier gas with a flow rate of 1.0 mL min-1. The reactive species were characterized using a Bruker A300-10/12 EPR spectrometer, in which •CH3 and •OH were captured with DMPO while •O2 was captured with TEMPO. The concentrations of •OH and H2O2 were measured by a terephthalic acid method and the Shimadzu UV-2600 UV–Vis spectroscopy by adding o-tolidine as the peroxide indicator, respectively. Aqueous Fe(II) concentration was determined using the ferrozine method at a wavelength of 562 nm with the Shimadzu UV-2600 UV-Vis spectroscopy46. Total iron (Fetotal) was measured after the reduction of Fe(III) to Fe(II) with hydroxylamine-HCl. The Fe(III) concentration was subsequently calculated as the difference between the Fetotal and Fe(II) concentrations61. DMSP measurement was made by cleaving DMSP into DMS with strong alkali and quantifying DMS by gas chromatography-mass spectrometry (GCMS-QP2020 NX, Shimadzu, Japan)62.

Cells were harvested by centrifugation at 5000 × g for 10 min (Eppendorf AG 5404, Hamburg, Germany). The pellet was resuspended in 1 mL of 90 % (v/v) acetone and remained in the dark and at 4 °C for 1 h or until no green pigment was visible. Subsequently, the samples were centrifuged again at 5000 × g for 10 min to separate the water soluble phase and the cell fragments (pellet) from the acetone extract (supernatant).

The quantification of Chl a was conducted with a Varian Cary Eclipse (now Agilent, USA) spectrofluorometer with the following equation63:

$${{{{{rm{Chl}}}}}}, {{{{{rm{a}}}}}}, ({{{{{rm{mu }}}}}}{{{{{rm{g}}}}}},{{{{{{rm{mL}}}}}}}^{-1})=(11.93times {A}_{664})-(1.93times {A}_{647})$$
(1)

Where A is the absorbance at different wavelengths (nm).

Cells were collected in a 1.5 mL tube and kept in the dark for 20 min. Then the photosynthetic efficiency was measured with an AquaPen-C AP-C 100 (Photon Systems Instruments, Brno, Czech Republic). The maximum quantum yield at PSII was measured in dark-adapted cells by providing a blue light (455 nm) to excite chlorophyll.

Fv/Fm was calculated according to the following equation63:

$${F}_{{{{{{rm{v}}}}}}}/{F}_{{{{{{rm{m}}}}}}}=({F}_{{{{{{rm{m}}}}}}}-{F}_{0})/{F}_{m}$$
(2)

Where Fm is the maximal fluorescence yield observed in dark-adapted cells after stimulation with a saturating light pulse (3000 μmol photons m-2 s-1); F0 is the fluorescence yield measured in dark-adapted sample when all PSII reaction centers are open and it was measured with a measuring light (0.045 μmol photons m-2 s-1).

The conversion efficiency of photosynthate to methane (i.e., percentage of gross primary production diverted to CH4 formation) was calculated based on the reduced concentration of inorganic carbon and CH4 yield.

Reporting summary

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

Data availability

The data supporting the findings of this study are available within the paper and its supplementary information. The RNA-seq data generated in this study have been deposited in the NCBI Trace Archive database under accession code PRJNA1114667. Source data are provided with this paper.

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Acknowledgements

This work was supported by the National Science Fund for Distinguished Young Scholars (41925028), the National Science Fund for Excellent Young Scholars (42322706), the National Natural Science Foundation of China (92251301), and a faculty startup fund from Washington University in St. Louis.

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J.Y., Z.H. and S.Z. conceived the idea and designed the experiments. M.Z., M.H., G.R., A.H. and C.Y. performed the experiments. J.Y., M.Z., and M.H. analyzed the data. J.Y., D.Z., Z.H. and S.Z. wrote and revised the manuscript. All the authors contributed to the interpretation of the data and preparation of the manuscript.

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Zhen He or Shungui Zhou.

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Nature Communications thanks Zhaoji Zhang, Gefu Zhu and the other, anonymous, reviewer for their contribution to the peer review of this work. A peer review file is available.

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Ye, J., Zhuang, M., Hong, M. et al. Methanogenesis in the presence of oxygenic photosynthetic bacteria may contribute to global methane cycle.
Nat Commun 15, 5682 (2024). https://doi.org/10.1038/s41467-024-50108-3

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  • Received: 20 January 2024

  • Accepted: 01 July 2024

  • Published: 06 July 2024

  • DOI: https://doi.org/10.1038/s41467-024-50108-3

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