# H2-dependent formate production by hyperthermophilic Thermococcales: an alternative to sulfur reduction for reducing-equivalents disposal

Sébastien Le Guellec, Elodie Leroy, Damien Courtine, Anne Godfroy, Erwan Roussel

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## To cite this version:

Sébastien Le Guellec, Elodie Leroy, Damien Courtine, Anne Godfroy, Erwan Roussel. H2-dependent formate production by hyperthermophilic Thermococcales: an alternative to sulfur reduction for reducing-equivalents disposal. The International Society of Microbiological Ecology Journal, 2021, 10.1038/s41396-021-01020-x. http://hal-03263886

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# H₂-dependent formate production by hyperthermophilic Thermococcales: an alternative to sulfur reduction for reducing-equivalents disposal

Sébastien Le Guellec<sup>1</sup> · Elodie Leroy<sup>1</sup> · Damien Courtine<sup>1</sup> · Anne Godfroy<sup>1</sup> · Erwan G. Roussel<sup>1</sup>

Received: 24 November 2020 / Revised: 13 May 2021 / Accepted: 20 May 2021 / Published online: 4 June 2021
© The Author(s), under exclusive licence to International Society for Microbial Ecology 2021

## Abstract

Removal of reducing equivalents is an essential catabolic process for all microorganisms to maintain their internal redox balance. The electron disposal by chemoorganotrophic *Thermococcales* generates H₂ by proton reduction or H₂S in presence of S⁰. Although in the absence of S⁰ growth of these (hyper)thermopiles was previously described to be H₂-limited, it remains unclear how *Thermococcales* could be present in H₂-rich S⁰-depleted habitats. Here, we report that 12 of the 47 strains tested, distributed among all three orders of *Thermococcales*, could grow without S⁰ at 0.8 mM dissolved H₂ and that tolerance to H₂ was always associated with formate production. Two conserved gene clusters coding for a formate hydrogenlyase (FHL) and a putative formate dehydrogenase-NAD(P)H-oxidoreductase were only present in H₂-dependent formate producers, and were both systematically associated with a formate dehydrogenase and a formate transporter. As the reaction involved in this alternative pathway for disposal of reducing equivalents was close to thermodynamic equilibrium, it was strongly controlled by the substrates-products concentration ratio even in the presence of S⁰. Moreover, experimental data and thermodynamic modelling also demonstrated that H₂-dependent CO₂ reduction to formate could occur within a large temperature range in contrasted hydrothermal systems, suggesting it could also provide an adaptive advantage.

## Introduction

*Thermococcales* is one of the most ubiquitous hyperthermophilic *Archaea* orders found in hydrothermal ecosystems colonising a wide range of ecological niches such as deep-sea hydrothermal vents or oil reservoirs [1]. *Thermococcales* order is divided into three genera *Thermococcus* [2], *Pyrococcus* [3] and *Palaecoccus* [4] and comprises forty-two type strains and numerous isolates in laboratory and culture collections. They are described as obligate anaerobes and organotrophs utilising peptides or carbohydrates and producing acetate, carbon dioxide (CO₂) and hydrogen sulphide (H₂S)/dihydrogen (H₂) as major end products. Moreover,

some carboxidotrophic *Thermococcales* can also oxidise carbon monoxide (CO) to H₂ and CO₂ [5, 6].

Reducing equivalents are disposed either by reduction of elemental sulfur (S⁰) to H₂S or in its absence by proton reduction to H₂. In both cases, an electrochemical sodium ion gradient is generated driving the synthesis of ATP by a Na⁺-dependent ATP synthase [7, 8]. The sulfur-dependent metabolic switch is controlled by the transcription factor SurR in response to the S⁰ availability [9–11]. Thus, in the presence of S⁰ or polysulphide (e.g., [12]), a membrane-bound sulfane reductase complex (MBS, previously described as MBX) is transcribed leading to the generation of H₂S [13], whereas transcription of a membrane-bound ferredoxin (Fd)-oxidising hydrogenase complex (MBH) responsible for H₂ production is not induced [14, 15]. Inversely, in the absence of S⁰ only the transcription of the Mbh complex is activated. In this case and without interspecies H₂ transfer [16], endogenous or environmental H₂ accumulation inhibits growth of *Thermococcales* by blocking the recycling of the reduced ferredoxin pool [17, 18]. However, some *Thermococcales* have been shown to tolerate H₂ in the absence of S⁰ as several have been isolated in the presence of H₂ (e.g., *Thermococcus piezophilus* CDGS<sup>T</sup> [19]).

**Supplementary information** The online version contains supplementary material available at https://doi.org/10.1038/s41396-021-01020-x.

☒ Erwan G. Roussel
Erwan.Roussel@ifremer.fr

¹ Ifremer, Univ Brest, CNRS, Laboratoire de Microbiologie des Environnements Extrêmes, Plouzané, France
A recent transcriptomic study suggests that *Thermococcus paralvinellae* ES1T could dispose of the excess electrons through H₂ oxidation to formate using Formate hydrogenlyase (FHL) complex encoded by the *fdh-mfh-mnh* cluster [20]. Although FHL was initially described as responsible for formate oxidation to H₂ and CO₂ (HCOO⁻ + H₂O → H₂ + HCO₃⁻) coupled to ATP synthesis for *Thermococcus onnurineus* NA1 in pure culture conditions [21–23], it remains unclear whether the cells only rely on formate or if the relatively high amounts of yeast extract present in the medium could also sustain the anabolism [24]. However, some formate dehydrogenases (FDH) are also known to catalyse both reactions, oxidation of formate and reduction of CO₂ [25–27].

Owing to the small Gibbs free energy change in standard conditions (ΔG° = + 1.3 kJ mol⁻¹), the direction of the reaction is primarily thermodynamically controlled by the concentration of reagents. However, it still remains unclear what reaction is catalysed in situ by *Thermococcales* [20]. The aim of this study is to characterise the reaction catalysed by formate dehydrogenase (H₂-dependent formate production and/or formate oxidation) in hyperthermophilic *Thermococcales* and to determine its in situ physiological role.

# Materials and methods

## Strain collections

All *Thermococcales* strains used in this study were obtained from the Laboratory of Microbiology of the Extreme Environments microorganisms collection (Ifremer, Plouzané, France), the Université de Bretagne Occidentale Culture Collection (UBOCC, Plouzané, France), the German Collection of Microorganisms and Cell Cultures (DSMZ, Braunschweig, Germany) and the Japan Collection of Microorganisms (JCM, Wako, Japan) (for details, see Table S1). *Thermococcus nautili* 30-1 T was kindly provided by Dr. A. Gorlas (Institut de Biologie Intégrative de la Cellule, Orsay, France). *Thermococcus* sp. MF15 was isolated from samples collected from the Snake Pit hydrothermal field during cruise BICOSE 2014 [28].

## Culture conditions

*Thermococcales* were grown under anaerobic conditions in an artificial seawater medium (ASW) containing the following components in g L⁻¹: NaCl (27.2), MgCl₂·6H₂O (10.0), CaCl₂·2H₂O (1.5), KCl (0.66), KBr (0.1), H₃BO₃ (0.025), SrCl₂·6H₂O (0.04), NH₄Cl (0.021), KH₂PO₄ (0.0054) and NaF (0.003). The medium was supplemented with 1 mL L⁻¹ of trace-element solution SL 10 [29] and 0.2 mL L⁻¹ of selenite tungstate solution [30]. After

autoclaving at 121 °C during 60 min, the medium was cooled under an atmosphere of N₂/CO₂ (80:20, v/v; 30 kPa), prior to the addition of sodium bicarbonate solution from a sterile stock solution at the desired concentration. The medium was reduced with 1.2 mL L⁻¹ of sodium sulphide (Na₂S·9H₂O) at 1 M and the pH was adjusted at 6.8 by the addition of sterile HCl from 1 M sterile stock solution. Then, 50 mL medium was transferred under nitrogen flow into 120 mL serum bottles hermetically sealed with butyl rubber septa.

Colloidal sulfur (Sigma Aldrich, St. Louis, MO, USA) and yeast extract (Fisher Scientific, Hampton, NH, USA) were added at a constant volume ratio of 1% (v/v) in serum bottles at a desired final concentration from sterile stock solutions. Serum bottles were flushed and pressurised (200 kPa at 80 °C) with the desired gas phase consisting of H₂/N₂/CO₂ (CO₂ was kept constant in the gas phase at 20%).

Unless otherwise indicated, each experiment was carried out in triplicate and incubated at 80 °C (optimal growth temperature of *T. onnurineus* NA1 [31]) in unshaken static conditions. Inoculation was always performed at 1% (v/v) from exponential phase cell cultivated on ASW with 2 g L⁻¹ yeast extract and with N₂/CO₂ (80:20; v/v; 200 kPa) as gas phase. Cells were observed under an Olympus BX60 phase-contrast microscope (Olympus Corporation, Tokyo, Japan). Cell quantification was assessed by direct cell counting using a Thoma cell counting chamber (depth 0.02 mm).

## Volatile fatty-acid analysis

Formate and acetate concentrations were determined using a Dionex ICS-2000 Reagent-Free Ion Chromatography System equipped with an AS50 autosampler (Thermo Fisher Scientific, Waltham, MA, USA) as described by Roussel et al. [32]. Chromatographic separation was conducted on two Ionpac AS15 columns (4 × 250 mm) at 30 °C and the determination of species was carried out using an anion self-regenerating suppressor (ASRS 300 4-mm) unit in combination with a DS6 heated conductivity cell (35 °C). The gradient programme was as follows: 8 mM KOH (29.9 min), increase 28.5 mM KOH min⁻¹ to 65 mM (30.1 min), decrease 57 mM KOH min⁻¹ to 8 mM (9 min). Prior to ion chromatographic analysis, 0.5 mL of culture was sampled and then centrifuged (15 min at 12,000 × g at room temperature). The supernatant was diluted (1:10, v/v) in ultrapure water (Millipore, Billerica, MA, USA).

## ¹⁴C-radiolabelled formate measurements

H₂-dependent formate production from bicarbonate was measured using ¹⁴C-radiolabelled substrate. Culture conditions were identical as previously described. *T*.
*onnurineus* NA1 was cultivated in 50 mL ASW medium supplemented with 0.2 g L⁻¹ of yeast extract and 30 mM bicarbonate, without S⁰ and pressurised with H₂/CO₂ or N₂/CO₂ (80:20, v/v; 200 kPa). [¹⁴C]Na-HCO₃⁻ with a specific activity of 59.0 mCi mmol⁻¹ was purchased from Perkin Elmer (Waltham, MA, USA). Cultures were amended with 28 µL of [¹⁴C]Na-HCO₃⁻ to a final activity of 96 kBq using a Hamilton syringe. Abiotic controls were not inoculated and negative control experiments were performed by autoclaving inoculated cultures (20 min at 121 °C). Each activity measurement for each time point was performed in triplicate. Production of ¹⁴C-formate was measured by collection of the ¹⁴C-formate fraction using a Dionex ICS-2000 Ion Chromatography System equipped with an AFC-3000 Automated Fraction Collector (Thermo Fisher Scientific, Waltham, MA, USA). All analytical conditions were identical to those described previously for anion analysis. Samples were transferred to OptiPhase HiSafe 3 scintillation cocktail (Perkin Elmer, Waltham, MA, USA) and quantification of ¹⁴C-formate was determined by liquid scintillation counting with a Tri-Carb 2910TR liquid scintillation counter (Perkin Elmer, Waltham, MA, USA).

## Phylogenomic analysis

A phylogenomic tree was constructed with all *Thermococcales* genomes retrieved from the NCBI genome database (https://www.ncbi.nlm.nih.gov/genome) using a set of 59 conserved phylogenetic marker proteins (Table S2) [33]. Sequences from three *Methanococcus* genomes (*M. aeolicus*, NC_009635.1; *M. vannielii*, NC_009634.1 and *M. voltae*, NC_014222.1) were used as out-group. For each protein, sequences were aligned using MAFFT v7.055b (parameter ‘linsi’) [34] and alignments were trimmed using BMGE v1.12 (default parameters) [35]. All alignments were then concatenated, and conserved positions across all genomes were removed with Seaview v4.6 [36]. The phylogenomic tree was computed with PhyML v3 [37] on the ATGC Montpellier Bioinformatics webserver (http://www.atgc-montpellier.fr). Model selection was done with SMS [38] using Akaike information criterion. The best model used for the tree was ‘LG’ [39].

Comparative genomic analysis was performed using EDGAR 1.3 software framework (http://edgar.computational.bio.uni-giessen.de) [40]. Homology searches were conducted with BLASTp using default settings (http://blast.ncbi.nlm.nih.gov/Blast.cgi). Studies of synteny of genes were performed using the SyntTax software (http://archaea.u-psud.fr/synttax) using default parameters [41].

## Thermodynamic calculations

Standard Gibbs free energies (ΔG⁰) of the reaction of formate production at 25 °C:

$$H_2(g) + HCO_3^-(aq) \leftrightarrow HCOO^-(aq) + H_2O(l)$$

were calculated according to:

$$\Delta G^0 = \Delta fH^\circ - T \cdot \Delta S^\circ$$

Where ΔfH° is the standard enthalpy of formation, T the absolute temperature (K) and ΔS° the standard entropy change. Thermodynamic data of reactants and products for species were obtained from the NBS tables of chemical thermodynamic properties [42]. Bicarbonate and formate were assumed as dissolved species. In order to strengthen the demonstration, H₂ was assumed as a gaseous substrate, as the Gibbs free energies of the reaction under non-standard conditions (ΔG') for CO₂ reduction to formate were less favourable than assuming H₂ as dissolved.

The Gibbs free energies of reactions under non-standard conditions (ΔG') were calculated according to

$$\Delta G' = \Delta G^0 + RT \cdot \ln Q_r$$

Where Q_r is the reaction quotient (expressed as concentrations) considering activities almost equal to concentrations based on Amend and LaRowe 2019 [43] (see Supplementary Methods section), R the gas constant (8.314 × 10⁻³ kJ mol⁻¹ K⁻¹) and T the absolute temperature (K).

## Arrhenius parameters

An estimation of the temperature dependence of each studied metabolic process could be obtained by calculating the activation energy (Ea) and the Q₁₀ factor from Arrhenius plots [44]. The Arrhenius profiles were obtained by plotting the natural logarithm of each maximum rate for each incubation temperature versus the inverse of temperature. The activation energy for each metabolic process was calculated from the following equation:

$$\ln(k) = \ln(A) + \left( \frac{-Ea}{R} \cdot \frac{1}{T} \right)$$

Where Ea is the activation energy (kJ mol⁻¹), k the reaction rate (µmol cm⁻³ day⁻¹), A the Arrhenius constant, R the gas constant (8.314 × 10⁻³ kJ mol⁻¹ K⁻¹), and T the absolute temperature (K).

Q₁₀ is the factor by which the rate of reaction increases with a temperature increase of 10 °C. The selected
temperature range in this study was between 75 °C and 85 °C. $Q_{10}$ was calculated using the following equation:

$$Q_{10} = \exp\left[\frac{Ea \cdot 10}{RT(T + 10)}\right]$$

## Dissolved H₂ concentration determination

Dissolved H₂ concentrations estimated as if they were at equilibrium with the overlying gas phase were calculated according to the Bunsen gas solubility coefficient [45, 46] and Henry's Law:

$$[H_2]_{(aq)} = \frac{P_{H_2} \times \beta}{R \times T}$$

Where $[H_2]_{(aq)}$ is the dissolved H₂ concentration (mol L⁻¹), $P_{H_2}$ the partial pressure of H₂ in the headspace (atm) at the temperature of measurement, $\beta$ the Bunsen solubility coefficient (0.015022), T the temperature of measurement (353.15 K) and R the gas constant (0.08206 atm L mol⁻¹ K⁻¹).

The Bunsen solubility coefficient ($\beta$) is expressed as cm³ of gas STP per cm³ of water at the temperature of measurement when the partial pressure of gas (gas volume corrected to STP, 0 °C and 1 atm) above the water is 1 atm. According to Weiss [46], it can be expressed as a function of temperature and salinity as follows:

$$\ln \beta = A_1 + A_2 \left(\frac{100}{T}\right) + A_3 \ln \left(\frac{T}{100}\right) + S_{\%e} \left[ B_1 + B_2 \left(\frac{T}{100}\right) + B_3 \left(\left(\frac{T}{100}\right)^2\right) \right]$$

where the A's and B's are constants from Wiesenburg and Guinasso [45] (for this study: $A_1 = -47,8948$; $A_2 = 65,0368$; $A_3 = 20,1709$; $B_1 = -0,082225$; $B_2 = 0,049564$ and $B_3 = -0,0078689$), T is temperature of measurement (353.15 K) and $S_{\%e}$ is salinity (31,860 g kg⁻¹).

Unless otherwise indicated, serum bottles were pressurised with 200 kPa of H₂/CO₂ (80:20, v/v) at 80 °C, which correspond to a calculated dissolved H₂ concentration of 0.8 mM.

## Results and discussion

### Growth in the absence of S⁰ and distribution of H₂-dependent formate production among *Thermococcales*

In order to investigate the inhibition of cell growth by H₂ in the absence of S⁰, 47 *Thermococcales* strains were tested in the presence of H₂ in gas phase (80% at 200 kPa; dissolved H₂ concentration 0.8 mM) (Table S1). Twelve

strains (26%) that presented significant growth were systematically associated with production of formate (1–12 mM), whereas 35 strains (74%) did not show either of these traits (Fig. 1). The ability to produce formate was evenly distributed across *Thermococcales* order (9 *Thermococcus*, 2 *Pyrococcus* and 1 *Palaeococcus*) with no clear distribution pattern. However, this distribution might be biased as solely relying on the study of species mostly isolated on S⁰ rich media. Growth temperatures were also distributed over a large range (50–112 °C) with an average optimal growth temperature of 86 °C, suggesting H₂-dependent formate production could occur over a wide range of microhabitats.

Mainly based on Fiala's pioneering work on *Pyrococcus furiosus* Vc1T [3, 17], growth of *Thermococcales* is usually described as being inhibited by H₂ accumulation in the absence of S⁰. However, our study shows that at least more than a quarter of *Thermococcales* strains tested were tolerant to H₂ in the absence of S⁰ and this proportion could be underestimated as most strains capable of formate production were isolated from environments presenting relatively high H₂ concentrations. For example, *T. piezophilus* CDGS<sup>T</sup> was isolated from the Piccard hydrothermal vent field (4964 m) that exhibits the highest H₂ concentration measured so far (19.9 mM) in high-temperature hydrothermal fluid [19, 47]. Moreover, with the exception of *T. kodakarensis* KOD1T isolated from a shallow solfataro on the shore of Kodakara Island, all these strains were also isolated among a wide range of deep-sea hydrothermal vent fields that are characterized by relatively low S⁰ (< 33 µM) and polysulfide (0.27 µM) concentrations [48–50], suggesting H₂-dependent formate production would also provide an adaptive advantage.

## Genomic comparison

### Identification and distribution of gene clusters

A genomic comparison was performed between genomes of strains exhibiting H₂-dependent formate production (n = 12) and those for which formate production was not detected (n = 22; Table S1). All formate producers had exclusively one and/or two complete specific gene clusters coding for an FHL1 and a putative formate dehydrogenase-NAD(P)H-oxidoreductase (Fdh-Nor), both annotated *fdh-mfh-mnh* and *fdh-nor*, respectively. These two clusters were composed of genes encoding formate dehydrogenase (*fdh*) and formate transporter (Fig. 2a), which seemed essential for H₂-dependent formate production.

The FHL1 cluster was previously described for *T. onnurineus* NA1 as ‘the formate dehydrogenase operon’ responsible for formate oxidation in the absence of H₂ in the culture at formate concentrations exceeding 1100 µM
**Fig. 1** Phylogenomic tree (based on 59 conserved archaeal marker proteins) representing H₂-tolerant *Thermococcales* capable of formate production in the absence of S⁰. *Thermococcales* showing growth and production of formate in the presence of H₂ after 96 h of incubation are shown in bold and red, whereas H₂-sensitive strains are in blue. ASW medium contained: 30 mM bicarbonate, no S⁰, 2 g L⁻¹ yeast extract and H₂/CO₂ in headspace gas (80:20, v/v; 200 kPa). Incubation temperature was 80 °C for *Thermococcus* and *Palaeococcus* and 95 °C for *Pyrococcus*. Nodes with a bootstrap value >95% were marked in black solid dots. A black solid symbol indicated the presence in genomes of specific gene clusters: triangles for 'FHL1', circles for 'FHL2', diamonds for 'FHL3' and squares for 'Fdh-Nor'. GenBank accession numbers are given in brackets. Scale bar represents 0.01 substitutions per nucleotide position.

<figure><img src="image_3.png" /></figure>

[21–23, 51]. This 17.1 kbp operon encompasses 18 genes coding for four subunit modules: a formate dehydrogenase (two genes, *fdh*), a group 4 membrane-bound [NiFe]-hydrogenase (eight genes, *mfh*) related to Complex I (NADH:ubiquinone oxidoreductase), a formate transporter (one gene) and a Na⁺/H⁺ antiporter (seven genes, *mnh*) [21] (Fig. 2a). However, here we identified a correlation between the presence of the FHL1 cluster and H₂-dependent formate production that disposes of reducing equivalents as previously suggested by Topçuoğlu and colleagues [20]. Moreover, further in silico genome analysis of *Thermococcales* also revealed the presence of numerous homologous nonspecific gene clusters (FHL2 and 3) in some strain genomes regardless of their capacity to produce formate (Fig. 2a and S1). Although these two gene clusters encoded homologous complexes of FHL1, proteins of the FHL2 were also quite divergent (average amino-acid sequence similarity of 39% with FHL1 of *T. onnurineus* NA1 TON_0541-43) and the gene coding for the formate transporter was systematically missing, suggesting again it was essential for the formate metabolism. In addition, the end of the FHL2 was also followed by a set of three genes (*nor*), encoding a putative cytoplasmic NAD(P)H-

oxidoreductase, that were similar to the ones found in *fdh-nor* cluster (average amino-acid sequence similarity of 71% with the *fdh-nor* of *T. onnurineus* NA1 TON_0541-43, Fig. S1). Regarding the FHL3 cluster, it was partially present in fourteen strains as genes encoding Na⁺/H⁺ antiporter were specifically missing for seven strains (Figs. 1 and 2a), which formed a tight phylogenetical cluster, suggesting their loss was related to a common ancestor. Moreover, an exclusive TetR/AcrR family transcriptional regulator was also present upstream of this cluster. Five of the complete FHL3-possessing strains could produce formate and also had the FHL1 cluster, suggesting that FHL3 had a different physiological function than FHL1.

Additional to the FHL1, a new cluster was also exclusively found in *Thermococcales* capable of H₂-dependent formate production (Figs. 1 and 2). This 5.5 kbp cluster was composed of three functional modules: a formate transporter (one gene), a formate dehydrogenase (two genes, *fdh*) and a putative cytoplasmic NAD(P)H-oxidoreductase (three genes, *nor*). Initially, the three-gene *nor* module was inferred to code for a putative glutamate synthase. Moreover, based on the cluster structures the Fdh-Nor cluster could be divided into two subgroup, Fdh-Nor1 and
<figure><img src="image_4.png" /><figcaption>**Fig. 2** Comparison of formate dehydrogenase containing gene clusters from 35 *Thermococcales* genomes sorted according to their H₂-dependent formate production. **a** Heat map showing similarity (blastp score) of each amino-acid (AA) sequence for *Thermococcales* compared with those of gene clusters from *T. onnurineus* NA1, which are represented above heat map (TON_1563-1580 for FHL1 and FHL2, TON_0282-0264 for FHL3 and TON_0538-0543 for Fdh-Nor). Grey squares indicate that no corresponding proteins</figcaption></figure>

were found. Gene clusters of *Thermococcales* that perform H₂-dependent formate production are indicated in red. GenBank accession numbers are given in brackets. **b** Formate production after 96 h of incubation in an ASW medium contained: 30 mM bicarbonate, H₂/CO₂ (80:20, v/v; 200 kPa) and 0.2 g L⁻¹ yeast extract. *Thermococcus* and *Palaeococcus* were incubated at 80 °C and *Pyrococcus* at 95 °C. *Pyrococcus yayanosii* Ch1T and *T. sp.* P6 were not tested.

Fdh-Nor2 (Fig. S1). However, as several studies suggest, this module could in fact code for a putative cytoplasmic NAD(P)H-oxidoreductase [52–54] and therefore also represents a new pathway to discard electrons and dispose of reducing equivalents.

## Putative evolutionary history

Although FHL-like complexes are found throughout the *Archaea* and *Bacteria* domains [55], these FHL-like clusters were almost specific to *Thermococcales* as the only known exception was found in *Thermofilum pendens* Hrk5T (Fig. S1), a *Crenarchaeota* also capable of sulfur respiration of peptides [56], which might have acquired this cluster through horizontal gene transfer. Among *Thermococcales*, the presence of multiple similar gene clusters reinforces the

hypothesis that they could be remnants from a common ancestral cluster [57]. However, their evolutionary history remains unclear, and whether these genes were obtain by genetic transfer or lost through genetic drift [20], especially that the distribution of *Thermococcus* lineages may not follow a simple model of allopatric speciation [58]. Both FHL1 and Fdh-Nor gene clusters only co-occurred in the genomes of three formate producers (*T. onnurineus* NA1, *T. piezophilus* CDGS<sup>T</sup> and *T. sp.* MF15) that formed a tight phylogenomic cluster (Figs. 1 and 2a). These strains also showed a highly conserved (>95% amino-acid sequence similarity) gene cluster FHL3 (Fig. 2a). Moreover, sequence analysis of Fdh catalytic subunit and formate transporter showed two monophyletic groups (FHL1/Fdh-Nor1 and Fdh-Nor2, Fig. S2), suggesting a common evolutionary history as the current distribution of these two gene clusters
could be owing to initial horizontal transfers followed by a
vertical inheritance.

Eleven *Thermococcales* also had two similar complexes to the FHL and Fdh-Nor with a carbon monoxide dehydrogenase as oxidoreductase instead of formate dehydrogenase (COdh-Mfh-Mnh and COdh-Nor) (Fig. S3). The COdh-Mfh-Mnh complex was previously described as involved in carboxidotrophic hydrogenogenic growth of *Thermococcus* (CO + H₂O → H₂ + CO₂) [59], whereas the COdh-Nor complex seems to be involved in carboxidotrophic growth coupled to S⁰ reduction in *Thermococcus gammatolerans* EJ3T [51]. Moreover, three hydrogenogenic CO-oxidising *Thermococcus* (*T. barophilus* Ch5, *T. onnurineus* NA1 and *T. paralvinellae* ES1T) have also the FHL1 cluster, which allows them to remove the H₂ and CO₂ produced through formate production, as *T. onnurineus* NA1 produced 5 mM of formate after 110 hours in the presence of CO in the gas phase (CO/N₂/CO₂, 50:30:20, v/v; 200 kPa). This suggests that H₂-dependent formate production could also act as a pathway for reducing-equivalents disposal in carboxidotrophic growth.

Function of the formate dehydrogenase, formate
transporter and hydrogenases of the fhl1 and fdh-nor
clusters

All these gene clusters probably have different functions
[57, 60], as suggested for *T. onnurineus* NA1, which possess
three clusters including the *fhl1* and *fdh-nor* clusters. Gene
expression of these different clusters varied according to the
CO, formate and starch concentrations [21, 61–63], suggest-
ing enzymes coded by these clusters could be specifically
expressed in different environmental conditions. Moreover,
although all *Thermococcales* with *fhl1* or *fdh-nor* clusters
were tolerant to H₂ (Figs. 1 and 2), strains with the *fhl1* cluster
had always produced more formate after 96 h (<11.9 mM for
*T. barophilus* Ch5) than the four strains with only *fdh-nor*
cluster (<5.3 mM for *Thermococcus cleftensis* CL1T). How-
ever, production for the three strains with both clusters was
not enhanced (from 7.5 to 9.5 mM) (Fig. 2). This discrepancy
could be the consequence of the different pathways used by
FHL1 and Fdh-Nor complexes to reduce CO₂ to formate.

In both FHL1 and Fdh-Nor, formate dehydrogenase (Fdh, 2 genes) and formate transporter (1 gene) proteins were conserved among strains as described in the previous section (Fig. 2 and S2). The Fdh catalyses the reduction of CO₂ as it was shown that the deletion of the catalytic subunit from the Fdh (TK_2076) of *fdh-nor* cluster in *T. kodakarensis* KOD1T, disables formate production [64]. Moreover, H₂-dependent formate production was shown to be reversible in *T. onnurineus* NA1, *T. cleftensis* CL1T and *Palaeococcus pacificus* DY20341T depending on the ratio of formate and H₂ strongly suggesting that the reaction

catalysed by FHL1 and Fdh-Nor complexes was bidirec-
tional (Table S3).

The protein responsible for formate secretion was only found in *Thermococcales* and *Thermofilum pendens* Hrk5T (Tpen_0191, Fig. S2) and therefore seems essential to H₂-dependent formate production. Protein TON_1573 was described as a formate transporter in *T. onnurineus* NA1 [21] that could be a formate-specific channel [55, 65]. Formate channels belong to the formate-nitrite transporters family and although functional mechanism of its members remains unclear, several studies suggested they function as specific bidirectional formate channels [66–68].

Besides the formate dehydrogenase and formate trans-
porter, H₂-dependent formate production requires also a
transfer, of the electrons generated through H₂ oxidation, to
Fdh (Fig. 3). This could be performed by the Mfh complex
(eight genes, 9.0 kbp) of the *fhl1* cluster or/and by the
putative Nor complex (three genes, 1.9 kbp) of the *fdh-nor*
cluster (Fig. 3). The function of this putative Nor complex,
initially described as a glutamate synthase [69], remains
controversial, and as several studies also suggest this
complex could be an oxidoreductase [52–54]. Expression of
the *fdh-nor* cluster in *T. onnurineus* NA1 is always corre-
lated with the *sulf1* cluster encoding a NADPH-dependent
cytosolic hydrogenase [61, 62, 70]. Hence, Fdh-Nor puta-
tive enzymatic complex could oxidise NADPH produced by
the *Sulf1* hydrogenase that reduces NADP⁺ from H₂ oxi-
dation and then transfers electrons to the Fdh for CO₂
reduction to formate (Fig. 3). Compared with the direct
transfer of electrons from the Mfh complex to the Fdh, this
different pathway encompasses additional intermediates that
could lead to a lower yield in formate synthesis. This might
explain why strains with only Fdh-Nor complex produce on
average 25% less formate than strains with only FHL1
(Fig. 2).

H₂-dependent formate production: an alternative
mechanism to S⁰ reduction

Two different strains (MPT and Ch5), both affiliated to
*Thermococcus barophilus* [71, 72] had a different tolerance
to H₂ in the absence of S⁰ (Figs. 1 and 4). *T. barophilus* Ch5
presented growth with H₂, whereas *T. barophilus* MPT was
unable to grow at dissolved H₂ concentrations exceeding
0.8 µM. In order to understand why their tolerances to H₂ in
the absence of S⁰ were so different, the effect of different H₂
concentrations on growth of these two *T. barophilus* strains
was compared with or without S⁰ (Fig. 4). In the presence of
S⁰, cell growth of both strains after 24 hours of incubation
was not affected by H₂ (Fig. 4a) as cell concentrations
remained relatively constant at 3 × 10⁸ cells mL⁻¹ regard-
less of the initial H₂ concentration. In the absence of S⁰,
increasing H₂ partial pressures led to a total inhibition of
<figure><img src="image_5.png" /></figure>

**Fig. 3** Schematic representation of the metabolism of *Thermococcales* strains producing formate in the presence of H₂. Red and blue indicate proteins expressed prominently during growth in the presence or in absence of sulfur, respectively. Other proteins are designated by green and orange text. **a** Genetic control by sulfur (adapted from [10]); **b** Catabolism (transport and utilisation of carbohydrates and peptides); **c** Energy generation & redox balance. Most proteins indicated in this illustration are shared by the majority of *Thermococcales*. BRE, transcription factor B recognition element, EM Embden-Meyerhof, Fd<sub>ox</sub> oxidised ferredoxin, Fd<sub>red</sub> reduced ferredoxin,

fdh formate dehydrogenase, fhl formate hydrogenlyase, mbh membrane-bound [NiFe]-hydrogenases, mbs membrane-bound sulfane reductase (previously known as ‘mbx’ membrane-bound complex); MFH membrane-bound formate-dependent hydrogenase, MRP multiple resistance and pH antiporter, NAD(P) nicotinamide adenine dinucleotide (phosphate), nfn NADH-dependent ferredoxin NADP⁺ oxidoreductase, nor NAD(P)H-oxidoreductase, nsr CoA-dependent NAD(P)H:sulfur oxidoreductase, SULF sulfhydrogenase; SurR sulfur response regulator. Diagram modified from [1, 20, 81, 82].

growth for strain MP<sup>T</sup> at 0.8 mM of dissolved H<sub>2</sub> whereas growth of strain Ch5 was not affected by the increase in H<sub>2</sub> concentrations at 30 mM of bicarbonate (Fig. 4b). Furthermore, formate production was only observed for strain Ch5 (up to 5.7 mM) (Fig. 4d) and was correlated to the H<sub>2</sub> concentration (R<sup>2</sup> = 0.99, p < 0.00001). Although H<sub>2</sub>-dependent formate production provided tolerance to H<sub>2</sub>, this mechanism seemed not as efficient at 0.8 µM dissolved H<sub>2</sub> compared to S<sup>0</sup> reduction as there were on average 3.2-fold less cells for strain Ch5 in the absence of S<sup>0</sup> (Figs. 4a and 4b). So even in the absence of H<sub>2</sub> the presence of S<sup>0</sup> stimulated cell growth of strains MP<sup>T</sup> and Ch5, suggesting that the yield and/or the kinetics of reducing-equivalents disposal were higher with S<sup>0</sup> than through H<sub>2</sub>-dependent formate production.

In addition, CO₂ concentrations also had an effect on growth of both strains. CO₂ occurs at several levels of the

catabolism of *Thermococcales* in peptides and pyruvate catabolism (e.g., [1]) but also in H₂-dependent formate production [20, 64]. In the presence of S⁰ and regardless of the concentration of H₂, variation of bicarbonate concentrations did not notably affect the growth of these two strains (Fig. 4a), although acetate production systematically decreased as bicarbonate concentrations increased (Figs. 4f and 4e), strongly suggesting that the accumulation of CO₂ controlled thermodynamically the catabolism activity. In the absence of S⁰, although growth of strain Ch5 (i.e., H₂-dependent formate producer) was also not affected by increasing H₂ concentrations at the highest bicarbonate concentration (30 mM), strain Ch5 growth decreased 2.8-fold (at 0.8 mM of dissolved H₂) simultaneously with the formate production (10.3-fold) as the bicarbonate concentration decreased from 30 to 2 mM (i.e., seawater concentration) (Figs. 4b and 4d). In contrast to strain Ch5, the increase of
<figure><img src="image_6.png" /><figcaption>**Fig. 4** Comparison of the effect of dissolved H₂ on growth, formate and acetate production after 48 h incubation of two strains of *Thermococcus barophilus* (MP<sup>T</sup> and Ch5). In the presence (a, c and e) or in the absence (b, c and f) of 3 g L<sup>-1</sup> of S<sup>0</sup> and at 2 or 30 mM of bicarbonate (empty dot and solid square, respectively). a, b Cell concentration. Dashed lines represent initial cell concentration before incubation. c, d Formate concentration. e, f Acetate concentration. T.</figcaption></figure>

bicarbonate concentrations seemed to limit the growth of strain MP<sup>T</sup> in the absence of S<sup>0</sup>, especially between 0.1 and 0.6 mM of dissolved H<sub>2</sub> (in average 3.5-fold decrease, Fig. 4b). This strongly suggests that not only H<sub>2</sub>-dependent formate production coupled to CO<sub>2</sub> reduction controlled the tolerance of strain Ch5 to H<sub>2</sub> in the absence of S<sup>0</sup> but that CO<sub>2</sub> also enhanced strain Ch5 growth by improving H<sub>2</sub> removal.

## Characterisation of H₂-dependent formate production

Formate oxidation (HCOO⁻ + H₂O → H₂ + HCO₃⁻) catalysed by formate dehydrogenase of the FHL1 complex is well studied in *Thermococcales* and more specifically in *T. onnurineus* NA1 [21–23, 62, 73]. However, the reverse reaction (i.e., H₂-dependent formate production) remains poorly characterised [20]. In order to confirm that bicarbonate reduction coupled to H₂ oxidation was responsible for the formate production in *Thermococcales*, *T. onnurineus* NA1

*barophilus* Ch5 (red) can perform H₂-dependent formate whereas *T. barophilus* MP<sup>T</sup> (blue) cannot. Cell concentrations and anion concentrations (formate and acetate) were measured after 24 and 48 h incubation, respectively at 80 °C in an ASW medium containing 2 g L<sup>-1</sup> yeast extract. Effect of H₂ concentrations were examined in a range of 0–80% (v/v; 200 kPa; 0–0.8 mM dissolved H₂) with CO₂ concentrations kept at 20% (v/v). Error bars show standard error (n = 3).

was incubated with <sup>14</sup>C-radiolabelled bicarbonate in the presence and in absence of H₂ (Fig. 5a). There was no <sup>14</sup>C-formate production neither in the abiotic control nor in the negative control (inoculated then autoclaved before incubation) and <sup>14</sup>C-formate production was only detected with viable cells in the presence of H₂, demonstrating that formate production was the result of H₂-oxidising bicarbonate reduction. In the presence of 0.8 mM of dissolved H₂ and 30 mM of bicarbonate, the maximum rate of formate production was in the same range (0.2 µmol cm<sup>-3</sup> h<sup>-1</sup>, 1.09 pmol cell<sup>-1</sup> h<sup>-1</sup>, 56 µmol hour<sup>-1</sup> mg<sup>-1</sup> of cells wet weight) as those obtained at 30 °C with *Acetobacterium woodii* (≈36 µmol hour<sup>-1</sup> mg<sup>-1</sup> of cells wet weight) and at 60 °C with *Thermoanaerobacter kivui* (≈115 µmol hour<sup>-1</sup> mg<sup>-1</sup> of cells wet weight) [26, 74]. As for *A. woodii*, formate accumulation increased in time and reached a plateau after 96 h (Fig. 5a), suggesting the reaction could have reached thermodynamic equilibrium.

In order to explore the possible thermodynamic control of the reaction, formate production was characterised for a range
<figure><img src="image_7.png" /><figcaption>**Fig. 5** Characterisation of H₂-dependent formate production of *T. onnurineus* NA1 at 80 °C. **a** Production of ¹⁴C-formate from ¹⁴C-bicarbonate (96 kBq) as a function of time. ASW medium contained: 30 mM bicarbonate, 0.2 g L⁻¹ yeast extract, H₂/CO₂ or N₂/CO₂ headspace (80:20, v/v; 200 kPa) and no S⁰ was added. Abiotic control was not inoculated. Negative control was autoclaved after inoculation (20 min at 121 °C). **b** Effect of dissolved H₂ (at 200 kPa) (blue diamonds) and bicarbonate (red spares) concentrations on formate production after 48 h incubation. ASW medium contained: 30 mM</figcaption></figure>

of substrate and product concentrations. Formate concentrations produced by *T. onnurineus* increased with increasing substrate concentrations as the formate production was positively and linearly correlated to dissolved H₂ and bicarbonate concentrations (R² ≥ 0.98, *p* < 0.0001, Fig. 5b). Levels of formate production did not reach a plateau even at the highest H₂ and bicarbonate concentrations tested (0.8 and 30 mM, respectively), suggesting that formate production per cell could be enhanced at higher dissolved H₂ concentrations. In culture, formate accumulation was the most limiting factor as formate production was negatively correlated with the initial concentration of formate (R² = 0.99, *P* < 0.001, Fig. 5c) and was inhibited at formate concentrations >9.9 mM in the presence of 0.8 mM of dissolved H₂ and 30 mM of bicarbonate (Fig. 5c). Interestingly, with bicarbonate in the range of seawater concentrations (i.e., 2 mM) and with moderate concentrations of dissolved H₂, compared with those usually detected in deep-sea hydrothermal fluids (i.e., 0.8 mM), formate production was inhibited at the higher formate concentrations (1077 µM, Fig. S4) in regard to the highest levels detected in these

bicarbonate for H₂ experiments, H₂/CO₂ (80:20, v/v; 200 kPa) for bicarbonate experiments, all conditions 0.2 g L⁻¹ yeast extract and no S⁰ was added. **c** Effect of initial formate concentration on formate production after 192 h incubation. ASW medium contained: 30 mM bicarbonate, H₂/CO₂ (80:20, v/v; 200 kPa), 0.2 g L⁻¹ yeast extract and no S⁰ was added. **d** Effect of S⁰ on formate (red squares) and acetate (blue triangles) productions after 384 h incubation. ASW medium contained: 30 mM bicarbonate, H₂/CO₂ (80:20, v/v; 200 kPa) and 2 g L⁻¹ yeast extract. Error bars represent standard error (n = 3).

environments (e.g., 158 µmol kg⁻¹ of hydrothermal fluid for Lost City) [75]. Hence, although thermodynamic constraints strongly controlled H₂-dependent formate production, our experimental data suggest that environmental formate concentrations should not limit formate production in most of the hydrothermal habitats described so far.

In addition, effects of S⁰ were also assessed as H₂-dependent formate production is an alternative pathway to S⁰ reduction for reducing-equivalents disposal. An increase in S⁰ concentrations stimulated *T. onnurineus* NA1 catabolic activity as acetate, one of the main final fermentation products of *Thermococcales* [1], increased simultaneously levelling out towards a maximum over 1 g L⁻¹ (31.2 mM) of S⁰ (Fig. 5d). This S⁰ threshold over which the catabolism was not further stimulated could either suggest that electron donors or accessibility to S⁰ became limited (especially that cultures were not shaken), or that the reaction reached the maximal velocity or that the SurR system was saturated [11]. However, maximum S⁰ concentrations measured in situ were three orders of magnitude lower than the lowest
<figure><img src="image_8.png" /></figure>

**Fig. 6** The direction of the reaction of CO₂ reduction depends primarily on substrate concentration. 2D-contour plot representing the thermodynamic modelling of the CO₂ reduction to formate for a range of concentrations of formate and H₂. Gibbs free energy of the reaction (H₂(g) + HCO₃⁻(aq) → HCOO⁻(aq) + H₂O(l)) was calculated for 80 °C and 2 mM of bicarbonate. As H₂ was assumed as a gaseous substrate for ΔG' calculations, H₂ partial pressures were converted to dissolved concentrations to compare with all the experimental and in situ data. The blue area indicates conditions for which formate oxidation occurs, whereas the red area indicates conditions for which CO₂ reduction to formate occurs. The black solid line and dashed lines define points of the chemical equilibrium of the reaction (ΔG' = 0 kJ mol⁻¹) from

thermodynamic modelling for three different bicarbonate concentrations, 2.0 mM, 0.2 mM, and 20.0 mM, respectively. The Green dashed line defines the experimental in vivo point of the chemical equilibrium of the reaction measured for *T. onnurineus* NA1 in culture medium after 96 h of incubation at 80 °C (see details Fig. S4). Effect of H₂ concentrations were examined in a range of 0–80% (v/v; 200 kPa; 0–0.8 mM dissolved H₂) with CO₂ concentrations kept at 20% (v/v). The effect of formate concentrations was examined in a range of 0 to 1 mM. ASW medium contained: 2 mM bicarbonate, 0.2 g L⁻¹ yeast extract and no S⁰ was added. Grey squares represent environmental concentrations measured for seawater and six hydrothermal vent fields (data available in Table S4).

S⁰ concentration over which the catabolism was not further stimulated [48, 50, 76]. At this in situ S⁰ concentration (<33 µM) [48–50], *T. onnurineus* NA1 would theoretically only exhibit 21% of the maximum acetate production, which is only 0.07% more than without S⁰, suggesting that the use by *Thermococcales* of the S⁰ reduction pathway could be limited in hydrothermal ecosystems as disposal of reducing equivalents through this pathway would be negligible compared to H₂-dependent formate production. Moreover, conversely to the S⁰ reduction pathway, formate production was inhibited by increasing S⁰ concentration especially at higher bicarbonate concentrations (Figs. 5d, 4c, d). This result was also consistent with the inhibition of the formate oxidation catabolic reaction [21] and with the fact that H₂-dependent formate production is also regulated by the accessibility to S⁰. Thus, *Thermococcales* probably save energy by only keeping one active electron discard pathway, depending mainly on the environmental sulfur availability. However, at in situ S⁰ concentrations and in the presence of H₂, it remains unclear if both electron discard pathways to dispose of reducing equivalents remain active.

## In situ reduction of carbon dioxide to formate

It has been demonstrated that *Thermococcales* conserved energy from formate oxidation at high formate concentrations (1.1–150 mM) in pure culture [21, 22], although it remains unclear whether cell growth solely relies on formate oxidation or if the yeast extract present in the medium also contributes to growth [24]. Moreover, in most hydrothermal environments, where levels of available S⁰ are not sufficient to efficiently dispose of reducing equivalents, it also remains uncertain whether formate oxidation occurs in situ as suggested in some studies [20, 22, 51]. Although the Gibbs free energy (ΔG⁰) of the H₂-dependent CO₂ reduction to formate is almost at equilibrium (−1.5 kJ mol⁻¹) in standard conditions considering H₂ as gas [77], it can reach −19 kJ mol⁻¹ if H₂ is considered as dissolved in the aqueous phase.

Here, we show using thermodynamic modelling and experimental data from cultures of *T. onnurineus* NA1 for a wide range of H₂ (assumed as gas or dissolved) and formate concentrations representative of those found in deep-sea hydrothermal systems [75, 78, 79] (Table S4), that only H₂-
dependent CO₂ reduction to formate occurred in these conditions (Fig. 6, S4–6), and even at hydrothermal sites for which formate oxidation was the most favourable. For example, using bicarbonate concentration similar to that of seawater (i.e. 2 mM) and in situ hydrothermal fluid H₂ concentrations for Lost City and Von Damm, the only known hydrothermal vent fields at which formate concentrations exceed 100 µM [75, 79], CO₂ reduction to formate was still thermodynamically more favourable (ΔG' between –13.4 and –21.4 kJ mol⁻¹) (Fig. 6). As formate oxidation by *Thermococcales* is probably encountered rarely in hydrothermal environments, the FHL1 and Fdh-Nor complexes mainly perform H₂-dependent CO₂ reduction to formate as an alternative pathway to sulfur reduction for electrons disposal. Moreover, H₂-dependent formate rates remain elevated beyond at least 5 °C over and below the temperature range for growth of *T. onnurineus* NA1, suggesting that the reaction could be maintained in cooler ecological niches (Ea = 28,6 kJ mol⁻¹; Q₁₀ (75;85 °C) = 1.318; Fig. S7). Given the ubiquity of *Thermococcales* in marine ecosystems (e.g. [80]), chemolithotrophic H₂-dependent formate production could play a role in the distribution of these microbial communities and on larger scales impact the organic carbon balance and the carbon cycle in the deep ocean.

**Acknowledgements** The authors thank Christophe Brandily, Françoise Lesongeur and Nadège Quintin for technical support, and Yves Fouquet, Karine Alain, Mohamed Jebbar, Jordan Hartunians and Jean-Pierre Donval for useful discussions. We are much indebted to anonymous referees for their helpful comments. We also thank Cécile Cathalot for providing hydrothermal fluid samples and Marie-Anne Cambon-Bonavita, chief scientist of BICOSE 1 and 2 cruises. All crew members and the Scientific Party were crucial in this effort, especially the ROV 'Victor 6000' and DSV 'Nautile' crews for the sampling efforts. Sébastien Le Guellec was supported by the 'Laboratoire d'Excellence' LabexMER (ANR-10-LABX-19) and co-funded by a grant from Ifremer and the Regional Council of Brittany. Elodie Leroy was supported by a grant from Ifremer.

## Compliance with ethical standards

**Conflict of interest** The authors declare no competing interest.

**Publisher's note** Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

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