Archaeal protein containing domain of unknown function 2193 undergoes oligomeric reconfiguration upon iron–sulfur cluster binding

Abstract

Methanogenic archaea are particularly rich in iron–sulfur proteins, yet their roles remain largely enigmatic. Here, we characterized a Methanococcus voltae (Mvo) protein from the domain of unknown function (DUF) 2193 family, a group of proteins present primarily in archaea and characterized by a conserved cysteine‐rich C‐terminal motif. MvoDUF2193 was heterologously expressed and characterized by a range of spectroscopic and analytical methods. The results demonstrate that MvoDUF2193 binds a single [4Fe–4S] cluster per subunit and that cluster occupancy regulates the transition from an apo tetramer to a [4Fe–4S] monomeric form. We hypothesize that MvoDUF2193 serves a regulatory role in the cell, mediated by [Fe–S] cluster binding and changes in oligomeric state.

This work presents the characterization of MvoDUF2193, a Methanococcus voltae (Mvo) protein from the domain of unknown function (DUF) 2193 family. We demonstrate that MvoDUF2193 binds a single [4Fe–4S] cluster per subunit and that cluster occupancy regulates the transition from an apo tetramer to a [4Fe–4S] monomeric form. This structural transition is hypothesized to regulate function.

Abbreviations

CW, continuous wave

DNA, deoxyribonucleic acid

DTT, dithiothreitol

DUF, domain of unknown function

E. coli , Escherichia coli

EDTA, ethylenediaminetetraacetic acid

EPR, electron paramagnetic resonance

Fe, iron

FeS ₂ , pyrite

GEE, glycine ethyl ester

IAA, iodoacetamide

IMS, ion mobility mass spectrometry

kDa, kilodalton

LCMS, liquid chromatography mass spectrometry

MIP, major intrinsic protein

Mvo , Methanococcus voltae

Na‐DT, sodium dithionite

NiNTA, nickel nitrilotriacetic acid

NMS, native mass spectrometry

S, sulfur

Cells from all domains of life require iron (Fe) and sulfur (S) for diverse biological processes, ranging from respiration and central metabolism to gene regulation [1, 2, 3]. Biological iron–sulfur (Fe–S) clusters, commonly found as [2Fe–2S] or [4Fe–4S] clusters, are often bound to proteins through cysteine thiolates and serve as cofactors, regulators, structural elements, and sensors [3]. Although [Fe–S] proteins have been the focus of intense study for many years, fundamental questions about the acquisition and biogenesis of these clusters remain. These questions are particularly puzzling for organisms that live in anaerobic environments, where the predominant oxidation states are Fe(II) and S²⁻, which have the propensity to form insoluble Fe–S minerals [4, 5, 6].

Methanogenic archaea (methanogens) are a diverse group of microbes that produce methane through a form of anaerobic respiration termed methanogenesis [7]. [Fe–S] proteins play key roles in methanogens, catalyzing oxidation–reduction reactions, assisting in electron transfer, and storing Fe and S, as well as in other essential biological processes [8]. In a study of over 300 methanogen genomes, ~ 5–8% of the protein‐coding genes were shown to code for [Fe–S] proteins [9]. Previous reports have shown that methanogens have a greater dependence on [Fe–S] proteins than most aerobic organisms; for example, Methanococcus maripaludis was shown to contain ~ 15‐fold more [Fe–S] clusters per mg of protein than Escherichia coli cells [10]. This increased dependence on [Fe–S] clusters in methanogens was initially surprising because Fe and S were presumed to be biologically unavailable in early anoxic environments due to their propensity to mineralize [11, 12, 13, 14, 15]. Nonetheless, methanogens were able to survive and diversify in those early anoxic environments, raising questions as to how the ‘unavailable’ [Fe–S] cluster components were acquired.

Recent work has demonstrated that two representative methanogens, Methanococcus voltae strain A3 (Mvo) and Methanosarcina barkeri MS, can reductively dissolve pyrite (FeS₂) [16]. These organisms use the released Fe and S, which are speculated to form aqueous [Fe–S] clusters up to (FeS)₁₅₀ in size, to meet biosynthetic demands [17]. To gain insights into the processes of Fe and S acquisition, trafficking, and storage following pyritic growth, shotgun proteomics was used to compare expression levels of Mvo proteins grown on Fe and S provided as aqueous Fe(II) and HS⁻ or FeS₂ [8, 18]. While a variety of functional classes were represented by proteins that were more abundant in FeS₂ growth conditions, many of these proteins lack functional annotation. One protein that stood out due to its large upregulation (13‐fold) under the FeS₂ growth condition was a member of the domain of unknown function (DUF) 2193 protein family, which we termed MvoDUF2193.

The DUF2193 family of archaeal proteins is noted for containing a cysteine‐rich domain, which we presumed to bind metals, yet these proteins lack further functional description. In this work, we cloned, heterologously expressed, and purified MvoDUF2193 to study and characterize it in vitro. We determined its metal‐binding properties using atomic absorption, UV–Vis, electron paramagnetic resonance (EPR), and Mössbauer spectroscopies and demonstrated that this protein binds a [4Fe–4S] cluster. By utilizing differential labeling and native mass spectrometry techniques, we further investigated the oligomeric state, redox status, and cofactor content of MvoDUF2193. Finally, we used a variety of computational tools to investigate known network partners (STRING), sequence/structure‐based function prediction (DeepFRI), and allosteric interactions (AlloSigMa) to gain insights into its intramolecular regulation and biological function. Our research shows that MvoDuf2193 exists in an equilibrium between oxidized and reduced states, and monomeric and tetrameric stoichiometries, an equilibrium that is dependent on [Fe–S] cluster binding. These investigations shed light on potential biological functions and provide a framework for future investigations.

Materials and methods

Target identification

Using results from Payne et al. [18], the 1270 proteins identified in the shotgun proteomics experiments were analyzed for potential targets of interest. Specifically, proteins that were upregulated in the FeS₂ growth conditions (> 2‐fold increase in FeS₂ vs. aqueous Fe/S growth conditions and had a P‐value < 0.05) were examined, which resulted in a list of 308 proteins. This list was further narrowed down by looking for potential metal‐binding motifs using Metalweb and Motif2 [19, 20]. MvoDUF2193 emerged as a target because it contained a cysteine motif, had a 13‐fold change in abundance between the two conditions, and was conserved in methanogens (as detailed below).

Conservation of MvoDUF2193 in methanogens

The InterPro database was searched for the DUF2193 family [21]. Information on protein sequences, taxonomic distribution, and predicted structure was used to make conclusions about the conservation of DUF2193 family members in methanogens. Additionally, publicly available genomes used in Johnson et al. [9] were compiled from the Joint Genome Institute Integrated Microbial Genomes and Microbiomes (JGI‐IMG/MER; https://img.jgi.doe.gov/cgi‐bin/mer/main.cgi) to assess genomic regions. These genomes were compiled, searched for MvoDUF2193 homologs, and then manually analyzed for porin genes located in the same open reading frame as the DUF2193 gene.

Cloning of MvoDUF gene

The gene corresponding to MvoDUF2193 was E. coli optimized, synthesized, and cloned into the bacterial expression plasmid pET19b by GenScript (Piscataway, NJ, USA) between the NdeI and BamHI restriction sites. It contains an N‐terminal deca‐His tag and an enterokinase cleavage site. The nucleotide and amino acid sequences are provided in the Supporting Information.

Expression, purification, and reconstitution of MvoDUF2193

The MvoDUF2193 construct was transformed into BL21 DE3 pLyS Rosetta E. coli cells and plated on 100 μg·mL⁻¹ carbenicillin plates. A single colony was picked and grown in an overnight culture of LB broth, which was then used to inoculate cell culture growths in phosphate‐buffered LB media (9 L in 2.8 L Fernbach flasks). Cultures were supplemented with carbenicillin (100 μg·mL⁻¹) and maintained at 37 °C with 200 r.p.m. shaking. At an OD₆₀₀ of 0.3, cells were cold shocked for 2 h at 4 °C. Following the cold shock, cells were induced with 1 mm IPTG, followed by an overnight incubation at 16 °C with 200 r.p.m. shaking. The cell pellets were harvested the next day through centrifugation and stored at −80 °C.

To isolate MvoDUF2193, all lysis and purification steps were performed in an anaerobic Coy (Grass Lake, Michigan, USA) vinyl glovebox. At a ratio of ~ 2 mL of buffer per 1 g cell pellet, lysis was performed on ice via sonication (5 s pulses, 15 s rest, 40% amplitude, 1 min 15 s total pulse time) in Buffer A (75 mm Tris pH 8, 300 mm KCl, 5% glycerol) in the presence of 1% Triton X‐100 (w/v), MgCl₂ (25 mg·mL⁻¹), lysozyme (0.2 mg·mL⁻¹), and DNaseI (0.2 mg·mL⁻¹) which was added fresh. After centrifugation (27 200g, 30 min, 4 °C), the clarified lysates were purified via a GE Healthcare (Chicago, IL, USA) AKTA FPLC on a 5 mL HisTrap High Performance (Cytiva, Logan, UT, USA) column using the following step gradient: (a) 0% Buffer B (75 mm Tris pH 8, 300 mm KCl, 5% glycerol, 500 mm Imidazole) wash for 5 column volumes (CV); (b) 20% Buffer B to wash non‐specific binding proteins off the column; (c) 80% Buffer B to elute MvoDUF2193. Eluted protein was concentrated with an Amicon Ultra‐15 centrifugal filter (30 kDa MWCO), then desalted into Buffer A using a disposable PD10 desalting column (Sigma, Burlington, MA, USA). The purified protein was analyzed by SDS/PAGE to check for purity. Aliquots of the purified concentrated protein were flash‐frozen in liquid nitrogen, then stored at −80 °C. Following purification, MvoDUF2193 was reconstituted at room temperature to increase iron–sulfur cluster content. Purified MvoDUF2193 (50 μm) was incubated with DTT (10 mm), ferrous ammonium sulfate (0.3 mm), and Li₂S (0.3 mm) for 3 h in an anaerobic chamber. Following reconstitution, the solution was centrifuged, desalted on a PD10 column with Buffer A, and subsequently concentrated to ~100 μm. Iron quantitation and protein concentration analyses were accomplished via flame atomic absorption spectroscopy and Bradford assay, respectively. The sulfide content was assayed using Beinhert's method [22].

Size exclusion chromatography

Purified MvoDuf2193 or Gel Filtration Standards (Biorad, Hercules, CA, USA) were loaded onto a HiPrep 16/60 Sephacryl S‐200 High Resolution column (Cytiva, Logan, Utah, USA), equilibrated in 75 mm Tris/Cl pH 8, 100 mm KCl, and eluted at a flow rate of 0.5 mL·min⁻¹.

Investigation of MvoDUF2193 oligomeric state

Native mass spectrometry

The oligomeric state of MvoDUF2193 was investigated using a native mass spectrometry protocol. All experiments were conducted on a SYNAPT G2‐Si instrument (Waters, Framingham, MA, USA) as described previously [23, 24]. Briefly, the reconstituted MvoDUF2193 sample was washed five times with 200 mm ammonium acetate, pH 7.5 (Sigma) using 10 kDa molecular weight cutoff spin filters (Pall Corporation, Port Washington, NY, USA) and then infused from in‐house prepared gold‐coated borosilicate glass capillaries to the electrospray source at a protein concentration of 5 μm and a flow rate of ~ 90 nL·min⁻¹. The instrument was tuned to enhance performance in the high mass‐to‐charge range with the following settings: source temperature 30 °C, capillary voltage 1.2 kV, trap bias voltage 16 V, and argon flow in collision cell (trap) 7 mL·min⁻¹. Transfer collision energy was held at 10 eV while trap energy was varied between 10 and 200 eV. Data analysis was performed in masslynx software version 4.1 (Waters).

Mass photometry

The oligomeric state of MvoDUF2193 was also assessed by mass photometry using the Two^MP mass photometer instrument (Refeyn, Watertown, MA, USA) following the manufacturer's instructions. Reconstituted MvoDUF2193 at a concentration of 270 μm was incubated in the presence and absence of 27 mm EDTA on ice for 1 h. Resulting samples were diluted with 75 mm Tris pH 8, 100 mm KCl, 5% glycerol to a final concentration of 10 nm, and a 20 μL sample volume was deposited on the instrument objective slide for measurement. The instrument was calibrated by interpolating the ratiometric contrast signal against the molecular weight of beta‐amylase (56, 112, 224 kDa for monomer, dimer, and tetramer, respectively), diluted to the same molar range in the sample buffer.

Protein modeling

To generate a model of MvoDUF2193, the amino acid sequence of the protein (Uniprot accession: D7DSA4) was submitted to alphafold2 (v2.3.2) on the Cosmic [3] web server [25, 26]. Additionally, a tetramer model was generated with the alphafold2 multimer function using the ‘Full_dbs’ database and multimer parameters.

Investigation of MvoDUF2193 cofactor

UV–vis spectroscopy

UV–vis spectra were recorded using an Agilent Cary 60 UV–visible spectrophotometer scanning from 1000 to 200 at 600 nm·min⁻¹. Samples were measured in Spectrosil Quartz cuvettes (Starna Cells, Atascadero, CA, USA) with a 10 mm path length.

Electron paramagnetic resonance spectroscopy

At room temperature in an anaerobic chamber, a solution of 90 μm MvoDUF2193 was reduced with sodium dithionite (0.9 mm) for 10 min in buffer (75 mm Tris pH 8, 100 mm KCl, 5% glycerol). The solution was then transferred to an X‐band EPR tube (Wilmad LabGlass, Vineland, NJ, USA, 4 mm OD) and frozen in liquid nitrogen. The X‐band continuous wave spectroscopy experiments were conducted at Montana State University using a Bruker EMX EPR spectrometer equipped with a Bruker/Cold Edge (Sumitomo Cryogenics, Allentown, PA, USA) 10 K waveguide cryogen‐free system with an Oxford Mercury iTC controller unit and helium Stinger recirculating unit (Sumitomo Cryogenics, Cold Edge Technologies, Allentown, PA, USA). Spectra were recorded at 12–30 K at 9.38 GHz using 10.0 G modulation amplitude and 1 mW power.

Mössbauer spectroscopy

To prepare MvoDUF2193 for Mössbauer spectroscopy, an overnight culture was grown in LB media, which was used to inoculate 1 L of minimal media. Labeled iron was added at the same time as induction, and then, the cells grew overnight at 16 °C. Harvesting, lysis, and purification were the same as described above. The reconstitution was done the same as described above, but with ⁵⁷Fe. Samples were transferred to Delrin Mössbauer cups and frozen in liquid nitrogen for storage. Spectra were collected with home‐built spectrometers using Janis Research Super‐Varitemp dewars.

Native mass spectrometry

The [4Fe–4S] cofactor content of reconstituted MvoDUF2193 was performed as described above for native mass spectrometry. By modulating the activation energy applied during ion transmission (trap energy), it was possible to detect several oligomeric states of reconstituted MvoDUF2193. Cofactor occupancy was estimated based on the homotetramer signal, which was the most abundant oligomeric form.

Investigation of MvoDUF2193 conformational flexibility

Ion mobility mass spectrometry

The conformational ensemble of MvoDUF2193 was probed using ion mobility mass spectrometry (IMS). As‐purified and reconstituted MvoDUF2193 samples were diluted to 5 μm and washed four times with 200 mm ammonium acetate pH 8 over a 10 kDa molecular weight cutoff spin filters (Pall Corporation). The reconstituted MvoDUF2193 sample was split and subsequently incubated with 10 mm EDTA (final) or 200 mm ammonium acetate pH 8 for 1 h on ice and then infused with an ionization source. The as‐purified MvoDUF2193 sample was diluted to 2.5 μm with 200 mm ammonium acetate pH 8 to match the protein concentration in all experiments. IMS data for all conditions were recorded on the same day and with the same instrument settings to enable direct comparisons of drift time values (dt). The following settings were used: source temperature 30 °C, capillary voltage 1.3 kV, trap bias voltage 60 V, and argon flow in collision cell (trap) 7 mL·min⁻¹. Wave velocity and height were, respectively, 300 m·s⁻¹ and 7 V in the trap, 800 m·s⁻¹ and 40 V in the IMS cell, and 64 m·s⁻¹ and 4.2 V in the transfer. IMS gas was nitrogen with a flow of 60 mL·min⁻¹. The complex activation was performed by ramping the collision voltage in the ion trap from 10 to 200 eV in steps of 10 eV while keeping the voltage constant (10 eV) in the transfer region. Data processing was performed in driftscope v 2.9 (Waters) and masslynx v 4.1 (Waters).

Structure‐based computational analyses

The tetramer model generated by alphafold2 multimer was submitted to AlloSigMA for the calculation of allosteric free energy calculations [27]. To mimic an [Fe–S] cluster binding, Cys 477, Cys 480, Cys 484, and Cys 493 in a single subunit were up‐mutated. Up‐mutations (replacement of the selected residue with a bulky amino acid) cause a strengthening of the local interaction network. In addition, the multimer model was submitted to MoleOnline to predict pores in the structure [28].

Investigation of MvoDUF2193 redox status and [Fe–S] cluster coordination site environment

The MvoDUF2193 redox status and the cofactor coordination site environment were probed by chemical modification of Cys (sulfhydryl group) and Glu/Asp (carboxyl group) residues using a protocol adopted from Boyer et al. [29]. This investigation was carried out at both the intact protein level (redox status) and peptide level (cofactor environment). To elucidate the number of (a) free Cys residues (reduced Cys), (b) Cys residues involved in inter‐ or intra‐disulfide bond formation (oxidized Cys) and (c) Cys residues engaged in [Fe–S] cluster coordination, the reconstituted MvoDUF2193 at a final concentration of 5 μm was alkylated with iodoacetamide (IAA, Sigma) without or after treatment with reducing (dithiothreitol (DTT, Sigma)) or chelating (ethylenediaminetetraacetic acid, (EDTA, Sigma)) reagents. To minimize variability between samples, control and labeling reactions were carried out in 175 mm Tris pH 7 in the dark and at room temperature, regardless of the presence/absence of alkylating reagent. In this experiment, the following samples were generated: (a) untreated MvoDUF2193; (b) MvoDUF2193 incubated with 55 mm IAA for 30 min; (c) MvoDUF2193 incubated with 10 mm DTT for 30 min; (d) MvoDUF2193 incubated with 10 mm DTT for 30 min then with 55 mm IAA for 30 min; (e) MvoDUF2193 incubated with 6.6 mm EDTA for 30 min; (f) MvoDUF2193 incubated with 6.6 mm EDTA for 30 min then with 55 mm IAA for 30 min.

Differentially labeled MvoDUF2193 samples were analyzed on a 1290 ultrahigh pressure series chromatography stack (Agilent Technologies, Santa Clara, CA, USA) coupled directly to an electrospray‐time of a flight mass spectrometer (Micro‐TOF, Bruker Daltonics, Billerica, MA, USA). Before infusion to the ionization source, samples were separated on a reverse phase column (Onyx monolithic C18, 100 × 2 mm; Phenomenex, Torrance, CA, USA) at 50 °C using a flow rate of 600 μL·min⁻¹ under the following conditions: 1 min, 10% B; 1–7 min, 10–70% B; 7–8 min, 70–90% B; 8–9 min, 90–95% B; 9–10 min, 95–10% B; 10–12 min, 10% B. Mobile phase solvent A and solvent B were 0.1% formic acid (FA, Sigma) in water (Thermo‐Fisher) and 0.1% FA in acetonitrile (Thermo‐Fisher, Waltham, MA, USA), respectively. Electrospray conditions were as follows: nebulizer at 6.0 bar, drying gas at a flow rate of 6.0 L·min⁻¹, drying temperature at 200 °C, capillary voltage at 4.5 kV, capillary exit voltage at 150 V, skimmer 1 set at 60 V, skimmer 2 set at 22 V, set at hexapole RF 350 V, and lens transfer at 87 μs. Data were collected in positive mode at 1 Hz over the 200 to 3000 m/z scan range. Data processing and analysis were done in the Bruker Data Analysis package 4.2. Charge deconvolution was performed using a maximum entropy algorithm for H⁺ adducts only and 0.1 m/z data point spacing. The low mass end was defined by the theoretical mass of the MvoDUF2193 monomer (calculated based on MvoDUF2193 amino acid sequence) while the high mass end was defined as 3.3× of the MvoDUF2193 monomer. The number of incorporated IAA labels was established based on additional mass in the increments of 57.02 m/z for the carbamidomethyl moiety.

To identify residues involved in coordinating the Fe‐S cluster, the reconstituted MvoDUF2193 was labeled with glycine ethyl ester (GEE, Sigma) in the presence of 1‐ethyl‐3‐(3‐dimethylaminopropyl)carbodiimide (EDC, Thermo‐Fisher) or with IAA. In the GEE/EDC protocol, MvoDUF2193 samples at a final concentration of 10 μm were incubated with 1 mm EDTA or 3.5 mm Tris pH 7 buffer for 30 min at room temperature and then treated with 5.7 mm EDC and 200 mm GEE for 15 min. In the IAA protocol, MvoDUF2193 samples at a final concentration of 10 μm were first incubated with 100 mm DTT or 1 mm EDTA or 175 mm Tris pH 7 for 30 min at room temperature and then treated with 55 mm IAA for 30 min in the dark at room temperature. In both protocols, control and labeling reactions were quenched by mixing with SDS/PAGE loading buffer and boiling for 5 min. All stock reagents were prepared immediately before the labeling reaction: DTT, IAA, and EDTA were dissolved in 175 mm Tris pH 7, while EDC and GEE were dissolved in 3.5 mm Tris pH 7.

The GEE‐ and IAA‐labeled MvoDUF2193 samples were separated on SDS/PAGE (4–20% linear gradient mini‐gel, Bio‐Rad), and the gel was stained with Coomassie brilliant blue (Thermo‐Fisher). The major protein bands were excised, briefly destained with 50% acetonitrile in 50 mm ammonium bicarbonate (Sigma) and subjected to in‐gel trypsin digestion by addition of 100 ng porcine sequencing grade modified trypsin (Promega, Madison, WI, USA) in 50 mm ammonium bicarbonate. After overnight incubation at 37 °C, peptide products were acidified in 0.1% FA (Pierce, Dallas, TX, USA). Tryptic peptides were separated by reverse phase XSelect CSH C18 2.5‐μm resin (Waters) on an in‐line 150 × 0.075 mm column using an UltiMate 3000 RSLCnano system (Thermo). Peptides were eluted using a 60 min gradient from 2% to 35% B (solution A: 0.1% FA, 0.5% acetonitrile; solution B: 0.1% FA, 99.9% acetonitrile). Eluted peptides were ionized by electrospray (2.4 kV) followed by analysis on an Orbitrap Eclipse Tribrid mass spectrometer (Thermo). MS data was acquired using the FTMS analyzer in profile mode at a resolution of 120 000 over a range of 375 to 1400 m/z with advanced peak determination. Following HCD activation, MS/MS data were acquired using the ion trap analyzer in centroid mode and normal mass range with a normalized collision energy of 30%. Peptides were identified using maxquant version 2.1.3.0 with a parent ion tolerance of 3 ppm and a fragment ion tolerance of 0.5 Da assuming the digestion enzyme trypsin (max two missed cleavages) [30]. Oxidation of methionine, acetyl of the N terminus, carbamidomethyl of cysteine, aspartic acid and glutamic acid, carbamidomethyl (M) of methionine, and GEE (DE) of aspartic acid and glutamic acid were specified in maxquant as variable modifications. scaffold q + s version 5.0.1 (Proteome Software Inc., Portland, OR, USA) was used to verify MS/MS‐based peptide and protein identifications. Peptide identifications were accepted if they could be established at greater than 5.0% probability to achieve an FDR less than 1.0% by the Peptide Prophet algorithm with Scaffold delta‐mass correction [31]. Only highly confident (probability score > 80%) peptides with ion intensities greater than 1 × 10⁷ counts were considered while searching for unique peptides between sample conditions.

[Fe–S] cluster stability

The individual reactions of the differential labeling experiment (chelation with EDTA, reduction with DTT, and alkylation IAA) were repeated with reconstituted MvoDuf2193 in 50 μL total volume. Following these reactions, the experiments and an unreacted sample of reconstituted MvoDuf2193 were exposed to oxygen and washed with 150 μL of 200 mm ammonium acetate pH 8. Washed samples were then filtered over a 3 kDa MWCO filter (Pall) until a volume of 50 μL remained above the filter. This was repeated several times over several hours. The retentate was then diluted to match the volume of each flow through. 300 μL of each sample's retentate and filtrate was acid digested with HNO₃ (20% w/v) with heat and pressure using the ‘20 fast’ cycle in a Steris‐Amsco SV‐120 autoclave. Acid‐digested samples were diluted to a volume of 3 mL with a solution of 2% HNO₃ and 0.5% HCl. Samples were then analyzed with an Agilent 7800 ICP‐MS equipped with an Agilent SPS 4 autosampler. Metal concentrations were determined against standard curves generated from serial dilutions of an environmental calibration standard mix (CPI International, Santa Rosa, CA, USA). An internal standard mix (Agilent, Santa Clara, CA, USA) was introduced with a T‐junction immediately before the ICP‐MS nebulizer to account for instrument drift and matrix effects.

MvoDUF2193 structure‐based function prediction

The potential function of MvoDUF2193 was predicted using two approaches: (a) sequence‐based function prediction with DeepFRI, a graph convolutional network which utilizes sequence features extracted from a protein language model and protein structures for function prediction and (b) protein–protein interaction network with STRING v.12.0 which derives information from five main sources: genomic context prediction, high‐throughput laboratory experiments, co‐expression, text mining, and previous knowledge in databases [32, 33]. MvoDUF2193 protein information was submitted for full STRING network (functional) analysis. The minimum required interaction score (confidence) was set to 0.4 and the maximum number of interactors in the 1st shell was set to no more than 50. The results considered for enrichment analysis had FDR ≤ 0.05.

Results

Identification of MvoDUF2193 as a target of interest

MvoDUF2193 is the protein product of Mvol_0354, which is proximal to a major intrinsic protein (MIP) family channel gene in Mvo (Fig. 1A). MIPs are transmembrane proteins that passively transport small polar molecules across membranes [36]. Interestingly, when the genomes of methanogen strains containing a DUF2193 gene were examined, 91% of the MvoDUF2193 homologs were proximal to a porin, a MIP family channel gene, an aquaporin, or a glycerol uptake facilitator protein (Table S1). Due to this genomic location and significant upregulation in FeS₂ growths, we hypothesized that MvoDUF2193 is involved in the acquisition and/or trafficking of [Fe–S] clusters. In addition, functional network analysis indicated several protein partners of MvoDUF2193 that contain FMN‐binding and rubredoxin‐like domains (Fig. S1, Table S2). Some of these proteins are known to be involved in the methanogen electron transport chain, mitigation of oxidative stress, and methane metabolism pathways. Therefore, MvoDUF2193 can potentially play a complex regulatory role in these processes.

Fig. 1.

Overview of the DUF2193 family. (A) Location of the MvoDUF2193 gene in the M. voltae A3 genome. (B) Taxonomic distribution of DUF 2193 family members. Taxons from archaea contain a patterned fill. (C) AlphaFold predicted model of MvoDUF2193 monomer [25, 26]. Cysteine residues are shown in purple and yellow, with key cysteine residues discussed in the text labeled with their number, and the conserved C‐terminal tail is shown in green. (D) Sequence logo of methanogen DUF2193 family members in the region of the conserved CX₂CX₆DX₂(H/C)X₂C motif, with the MvoDUF2193 residue numbering provided for the conserved cysteines [34, 35].

The DUF2193 family consists of 340 proteins that have no annotated function, according to its Interpro entry. Taxonomic distribution of this family reveals that 82% of family members are found in archaea, but homologs also exist in other domains of life (Fig. 1B). DUF2193 proteins are predicted to be predominantly composed of alpha helices, with an unstructured C‐terminus, as illustrated in the AlphaFold model of MvoDUF2193 (Fig. 1C). An additional unifying feature of this family is a conserved CX₂CX₆DX₂(H/C)X₂C motif in the C‐terminal tail (Fig. 1D). We hypothesized that the cysteines in this motif could serve as a binding site for labile [Fe–S] clusters.

Expression, purification, and initial characterization of MvoDUF2193

To gain insights into the function of MvoDUF2193, the Mvol_0354 gene was codon‐optimized for E. coli expression and cloned into a pET19b vector with an N‐terminal 10x‐histidine tag. The protein was overexpressed in E. coli BL21(DE3) cells and purified anaerobically using NiNTA affinity chromatography. We observed that the protein migrated according to its expected molecular mass of 59 kDa using denaturing gel electrophoresis (Fig. 2A) and appeared to be primarily tetrameric by size exclusion chromatography (Fig. S2). Although the protein was largely pure, we observed that MvoDUF2193 co‐purified with several E. coli proteins. Co‐purified protein bands were excised, digested with trypsin, and identified with tandem mass spectrometry. Interestingly, the analysis revealed the presence of DNA binding (rpoA, P0A7Z4; rpoB, P0A8V2), RNA binding (hfq, P0A6X3), transcription regulation (crp, P0ACJ8), and ribosomal proteins. Computational analysis using DeepFRI predicts that MvoDUF2193 may be involved in nucleotide metabolic processes (Fig. S3). This prediction supports the hypothesis that the endogenous biological function of this protein could involve nucleic acid binding.

Fig. 2.

Purity and cofactor content of MvoDUF2193. (A) SDS/PAGE gel of purified MvoDUF2193, with MW markers as shown in kDa. (B) Images of as‐purified (top) and reconstituted (bottom) MvoDUF2193 samples. (C) UV–Vis spectra of MvoDUF2193 as‐purified (black) and reconstituted (red). Shown are representative results from n = 4.

Due to the relatively high number of cysteines in the MvoDUF2193 sequence and the observed upregulation in the presence of FeS₂, we tested if this protein coordinated Fe. Using atomic absorption spectroscopy, we determined that following heterologous expression and purification, MvoDUF2193 contained ~ 1 Fe/monomer. To evaluate whether the protein could bind [Fe–S] clusters, we subjected MvoDUF2193 to Fe/S reconstitution conditions by adding ferrous ammonium sulfate, lithium sulfide, and dithiothreitol (DTT). Following reconstitution and desalting, the color of the solution changed from pale tan to dark brown, and the measured Fe content increased to 4.2 Fe/monomer (Fig. 2B). The S content of the reconstituted protein was measured to be 3.4 S/monomer following Beinert's method [22]. The protein was also analyzed using UV–Vis spectroscopy, and following reconstitution, we observed an increased absorbance in the visible region with a broad spectral feature at 420 nm (Fig. 2C). This absorbance is consistent with a [4Fe–4S]²⁺ cluster and, combined with the Fe and S stoichiometries, suggests that MvoDuf2193 coordinates a single [4Fe–4S] cluster.

Spectroscopic characterization of Fe‐S species in MvoDUF2193

Following the UV–Vis spectroscopic analysis, electron paramagnetic resonance (EPR) and Mössbauer spectroscopic studies were carried out to further characterize the Fe–S species bound to reconstituted MvoDUF2193. EPR samples of reconstituted MvoDUF2193 were prepared anaerobically and reduced with sodium dithionite (Na‐DT). The nearly axial EPR signal observed at 12 K was characteristic of a reduced [4Fe–4S]⁺ cluster and was well simulated with g‐values of (1.871, 1.915, and 2.063) (Fig. 3A). The signal intensity decreased with increasing temperatures between 12 K and 30 K, further supporting its assignment as a [4Fe–4S]⁺ cluster (Fig. S4).

Fig. 3.

Spectroscopic characterization of MvoDUF2193. (A) Continuous wave (CW) X‐band EPR spectrum (black) and simulation (red) of reconstituted MvoDUF2193 treated with Na‐DT. Simulations were generated with EasySpin [37]. Samples were collected at 12 K with a microwave frequency of 9.375 GHz and a modulation amplitude of 10 G. (B) Mössbauer spectrum of MvoDUF2193. Shown are representative results from n = 3 (EPR) and n = 2 (Mössbauer).

Mössbauer spectroscopy was carried out on MvoDUF2193 grown and reconstituted in the presence of ⁵⁷Fe. The data collected at 4.2 K and 0.1 T revealed that the major species observed are [4Fe–4S]²⁺ clusters (~ 60–70% of total Fe) with an isomer shift of δ = 0.44 mm·s⁻¹ and quadrupole splitting of ΔE _q = 1.12 mm·s⁻¹, parameters typical of a [4Fe–4S]²⁺ cluster (Fig. 3B). Additional broad magnetic features (~ 30–40% of total iron) were observed, which we speculate come from adventitiously bound Fe or [Fe–S] clusters with unknown structures, most likely arising from excess Fe–S present during the reconstitution. The combined observations from the calculated Fe and S content, as well as UV–Vis, EPR, and Mössbauer spectroscopies demonstrate that MvoDUF2193 binds a single redox‐active [4Fe–4S] cluster.

Oligomeric state of MvoDUF2193 is altered by [Fe–S] cluster binding

As noted above, as‐isolated MvoDuf2193 is tetrameric, with broad overlapping peaks in the SEC chromatogram suggesting some structural heterogeneity (Fig. S2). Reconstituted MvoDUF2193 was analyzed by native mass spectrometry (NMS) to examine the oligomeric state and further evaluate [4Fe–4S] cluster binding (Fig. 4A,B). Before proceeding, we determined that the [Fe–S] cluster remained stable and protein‐bound for at least a couple of hours in aerobic conditions, which include sample preparation and the duration of the experiment (Fig. S5). In NMS experiments, the monomeric, dimeric, and tetrameric stoichiometries were observed, with the tetrameric form being the most abundant. Both holo‐ and apo tetramer species were observed, and the deconvoluted mass confirmed the presence of four [4Fe–4S] cofactors in the holo‐tetramer, with an apo/holo ratio of 4.5/5.5. Mass photometry (MP) was pursued as a complementary, solution‐based technique with higher resolution than SEC (Fig. 4C). Similar to the NMS results, we observed both monomeric and tetrameric populations for the reconstituted protein; however, in MP, the monomer was the dominant population. When the protein was stripped of Fe using ethylenediaminetetraacetic acid (EDTA), the majority of the protein was observed to be in the tetrameric state. The differences in the relative amounts of monomer and tetramer observed by NMS (Fig. 4A) and MP (Fig. 4C) can be rationalized based on the different physical basis of the two techniques—dilute solution phase for mass photometry vs. gas phase for NMS. In NMS, as the sample is ionized into the gas phase, the local concentration of the protein increases, which can enhance protein–protein interactions, creating a bias for the tetrameric MvoDUF2193. As such, we reason that NMS data provides useful semi‐quantitation between apo‐ and holo‐tetrameric populations, but MP of dilute solutions provides better insight into the protein oligomeric state.

Fig. 4.

Reconstitution alters the oligomeric state of MvoDUF2193. (A) Native mass spectrometry analysis of reconstituted MvoDuf2193 shows multiple oligomeric forms. (B) Native mass spectrometry shows that the reconstituted MvoDUF2193 tetramer binds four [4Fe–4S] clusters. (C) Mass photometry of reconstituted (left, blue) and EDTA‐treated MvoDuf2193 (right, tan) shows that EDTA treatment shifts the equilibrium toward the tetrameric form. (D) AlphaFold model of MvoDUF2193 tetramer complex. C‐terminal cysteines potentially involved in [Fe–S] cluster binding are shown in pink and reside at the subunit‐subunit interface. Shown are representative results from n = 3.

Since NMS, mass photometry, and SEC all indicated the presence of tetrameric MvoDUF2193, we turned to AlphaFold Multimer and generated a model of the tetrameric complex (Fig. 4D) [38]. The resulting model was of high quality, with 96% of residues at high confidence (predicted local distance difference test (pLDDT) score > 90%) and 0% of residues of low confidence (below 50% pLDDT score) (Fig. S6). Intriguingly, the C‐terminal cysteines that we proposed to be involved in cofactor binding were located at the protein–protein interfaces in this model (Fig. 4D). Based on the predicted location of the [4Fe–4S] cluster within the monomer, combined with the predicted tetramer structure, we reasoned that metal binding influences the MvoDUF2193 oligomeric state.

Presence of [Fe–S] clusters impact the MvoDUF2193 conformational ensemble

Since the presence of the [Fe–S] cofactor impacts the oligomeric state of MvoDUF2193, we further investigated the structural changes observed upon cluster binding. We first used ion mobility mass spectrometry (IMS), which can detect conformational changes of proteins and protein complexes ranging from large movements of entire domain(s) (rigid body motions) to very subtle changes in the protein structure. In a typical IMS experiment, the time required for the biomolecule to progress through the ion mobility cell is measured, which depends on the molecule's mass, charge, and shape [39]. Therefore, small, well‐folded proteins travel faster than large complexes or denatured proteins. The MvoDUF2193 complex was investigated under three different conditions: (a) as purified, (b) reconstituted, and (c) stripped of Fe using EDTA. The tetramer in all conditions showed multiple conformations (Fig. 5A). The dominant conformation in the as‐purified condition was the largest (longest drift time) conformation observed. The addition of [Fe–S] shifted the equilibrium toward the smaller (shortest drift time) conformation. Stripping [Fe–S] clusters from the complex with EDTA restored the dominant conformation observed to the as‐purified condition. Together, these results indicate that when the MvoDuf2193 tetramer is in the holo, [4Fe–4S] cluster‐bound state, it adopts a more compact state than the as‐purified or Fe‐stripped proteins, both of which have low iron content. This trend was reversed for the monomeric species, with the as‐purified and stripped monomers having a smaller dominant conformation, while the reconstituted monomer had a larger dominant conformation (Fig. 5B). Overall, the IMS data show that Fe binding to MvoDuf2193 alters the conformational ensemble.

Fig. 5.

Impact of [Fe–S] cofactor binding on MvoDUF2193 protein conformation. Ion mobility mass spectrometry (IMS) was used to evaluate the conformational ensemble of MvoDUF2193 under three conditions: as purified, reconstituted, and reconstituted then stripped with EDTA. (A) Charge state distribution for tetrameric and (B) monomeric forms. The ion mobility drift time (red insets) points to the presence of a conformational ensemble that can be regulated by the presence of [Fe–S] cluster. Shown are representative results from n = 3.

To provide further insight into the effects of [Fe–S] cluster binding on the MvoDuf2193 complex, we used a structure‐based statistical mechanical model of allostery (AlloSigMA server), which has been used in other studies to investigate allosteric effects upon ligand binding [40, 41]. This tool calculates changes to the allosteric free energy under a given set of ‘mutations’ at single residue resolution [27]. Mutations can either be ‘up’ or ‘down’ depending on the allosteric question being investigated. An ‘up‐mutation’ increases the strength of interactions of the mutated residues, while a ‘down‐mutation’ destabilizes the mutated residues. AlloSigMA then calculates the changes in allosteric free energy across the protein complex caused by these defined mutations. Based on our tetrameric model, the proposed cluster‐binding Cys residues are rather buried at protein–protein interfaces (Fig. 4), where [4Fe–4S] cluster‐binding would be expected to stabilize and favor the proximity of the four cysteine residues. To mimic the effect of [4Fe–4S] cluster binding on the tetramer, four cysteines in the C‐terminal tail (Cys 477, 480, 484, 493) in a single subunit were computationally up‐mutated to strengthen the protein–protein interaction network near the binding site. Although this replacement resulted in large changes to protein dynamics across all subunits, the induced changes varied within each subunit (Fig. S7). These results indicate that, even though each subunit has a similar fold, all four subunits are dynamically quasi‐equivalent. For example, in the subunit harboring the mutations, a prominent decrease in dynamics was predicted across the entire subunit. Conversely, the other subunits showed more diverse changes: the largest destabilizing effect is observed primarily on the complex surface, distal from the core predicted to contain the [Fe–S] cluster binding site. The exception to this was in one subunit, which showed a large decrease in dynamics in residues 386–400. This region is in close proximity (less than 5 Å) to the mutation site on the mutated subunit. These findings, taken in context with the IMS data (Fig. 5), imply that changes in the C‐terminus in one subunit result in long‐range and widespread changes to the protein dynamics of the tetramer. This suggests that complex formation is allosterically driven in the presence of the [4Fe–4S] cluster by altering the MvoDUF2193 conformational ensemble.

Investigation of cysteine redox status in MvoDUF2193

The amino acid sequence of MvoDUF2193 includes 12 cysteines, three of which are strictly conserved (Fig. 1C,D). Cysteine residues are particularly interesting as they can be involved in metal coordination as well as inter‐ and intra‐subunit disulfide bond formation. To provide further insight into cysteine redox status and metal coordination, we analyzed the conservation scores and location of the cysteine residues in the structural model (Fig. 1C,D). Cysteines 477, 480, and 493 are conserved in all DUF2193 family members, while Cys 484 is specific to MvoDUF2193. The spatial proximity of cysteines 477, 480, 484, and 493 from our model suggests that they could be involved in [4Fe–4S] cluster coordination (Fig. 1C). This was further supported by the structure‐based prediction tool, DeepFRI, which showed a high likelihood for metal ion binding in this region (Fig. S8) [32]. Additionally, it is predicted that Cys 139 and 372 could be involved in disulfide bonds due to their proximity (less than 5 Å) and evidence from differential labeling experiments discussed below. Two other cysteine residues, Cys 270 and Cys 447, could also be involved in disulfide bond formation based on their proximity. These residues, along with Cys 372 and Cys 139, line the wide channel within the MvoDUF2193 monomer (Fig. S9). Considering the dynamic flexibility of the unstructured regions of the channel and the proximity of the cysteine residues, these observations suggest that a disulfide bond network may exist between these residues, which could work in unison to open and close the channel.

The redox status of the cysteine residues was evaluated by labeling reconstituted MvoDUF2193 with iodoacetamide (IAA) following disulfide bond reduction with DTT, or metal‐stripping with EDTA (Fig. 6A). As a reference, we used reconstituted MvoDUF2193 labeled with IAA without any DTT/EDTA treatment. The labeling, reduction, or metal‐stripping impact on protein‐cluster binding stability was evaluated using inductively‐coupled plasma ‐ mass spectrometry (ICP‐MS, Fig. S5), revealing that labeling with IAA or reduction with DTT has minimal impact on cluster binding stability while chelation with EDTA removed around 55% of bound iron from the MvoDuf2193 protein sample. Liquid chromatography mass spectrometry (LCMS) was used on the intact protein level to assess the IAA label incorporation. Since this analysis was conducted under denaturing conditions, we expected to see a wide distribution of charge states due to protein unfolding, as observed in our IAA‐labeled reference (Fig. 6B). Interestingly, the charge states distribution appeared to have at least two maxima (outlined with purple and green dashed lines), and deconvolution of the data revealed the presence of MvoDuf2193 proteins that differed by 2 Daltons. We attributed this mass difference to the absence or presence of a single disulfide bond, which we further confirmed by comparing labeling patterns with DTT reduced sample (8 labels incorporated in DTT condition vs. 6 and 8 labels incorporated in the reference). This disulfide bond was intra‐subunit since we did not detect any denatured oligomeric forms of MvoDuf2193 in any sample. Overall, these results indicate a mixture of protein redox states, in which one population harbors a single disulfide bond and the other does not. To investigate cysteine residues involved in cluster coordination, we treated MvoDuf2193 with EDTA to remove the iron–sulfur cluster prior to IAA labeling. We again detected two populations, however, the labeling pattern was different in comparison to the reference (6 and 9 labels incorporated in EDTA condition vs. 6 and 8 labels incorporated in reference), indicating that one additional cysteine is labeled upon EDTA treatment. Although one would expect 3 or 4 cysteine residues to be ‘revealed’ upon removal of the [4Fe–4S] cluster with EDTA (depending on whether it was coordinated with three or four cysteine residues), the data in Fig. S5 reveals that EDTA treatment does not completely remove the cluster, consistent with fewer than 3 or 4 additional cysteines being labeled. In addition, it is possible that cluster removal with EDTA could be accompanied by formation of a disulfide bond between the newly freed cysteine thiolates, or altering the conformation of other cysteines into inaccessible regions, which would reduce the number of free cysteines labeled after EDTA treatment. The fact that we were unable to detect all 12 cysteine residues being labeled in any of the conditions suggests that some of the residues must be buried deep in the protein core, which is in agreement with our MvoDuf2193 tetrameric model (Fig. 4D and Fig. S10).

Fig. 6.

Cysteine labeling in MvoDUF2193. (A) Schematic illustrating the differential labeling workflow (B) Mass spectrum of reconstituted MvoDUF2193 (monomer) collected in denaturing conditions. The green dashed line shows a charge state envelope for a protein with a single disulfide bond. The charge state envelope indicated by the purple dashed line is the reduced form. The inset table indicates the number of carbamidomethyl moieties incorporated after labeling with IAA. Shown are representative results from n = 3.

The spectroscopic and intact protein‐level mass spectrometry analyses described above show that MvoDUF2193 binds a [4Fe–4S] cluster and may harbor disulfide bonds under non‐reducing conditions. To gain insight into the specific cysteine residues that coordinate the [4Fe–4S] cluster or are involved in a disulfide bond, we pursued peptide‐level mass spectrometry analysis following a differential labeling approach. Combinations of IAA (which labels thiols), glycine ethyl ester (GEE) (which labels glutamate and aspartate residues), DTT (which reduces disulfide bonds), and EDTA (which chelates metal ions) were used as chemical probes of MvoDUF2193 (Fig. 6B). Following treatments, samples were separated on an SDS/PAGE gel, and selected protein bands were subjected to LCMS analysis. Although the purified and reconstituted MvoDUF migrated as a single band in denaturing conditions on SDS/PAGE (Fig. 2), we observed that samples treated with IAA exhibited two bands centered around the expected MvoDUF2193 molecular weight (Fig. S11). These two bands were compared with the corresponding bands from differentially treated samples to determine the cysteine redox and bonding status with and without [Fe–S]. For example, the upper band of the IAA‐treated sample was compared to the upper band of the IAA‐ and EDTA‐treated sample. Within each comparison, we searched for ‘unique’ peptides, that is, peptides that were labeled in only one condition and not the other. Specifically, we looked for modified peptides with a probability, or confidence level, of 80% or greater and with a peptide ion intensity larger than 1 × 10⁷ counts. This analysis revealed no unique peptides, which is consistent with labeling of several populations of reduced/oxidized and [Fe–S] cluster‐bound/unbound protein in equilibria.

Discussion

Iron and sulfur play critical roles as inorganic biological cofactors in all domains of life and have been shown to have diverse roles in methanogens. Recent reports have demonstrated the remarkable ability of methanogens to reductively dissolve pyrite (FeS₂) in order to meet biosynthetic demands; however, the process of iron–sulfur cluster acquisition and assimilation remains enigmatic [8, 16]. During the growth of Methanococcus voltae on pyrite, a number of proteins, including MvoDUF2193, are observed to be significantly more abundant when compared to growth on soluble sources of iron and sulfide. Here, we characterized MvoDUF2193, a methanogenic Fe‐binding protein that until now was a hypothetical metalloprotein. The DUF2193 gene is conserved across a wide variety of methanogens and is located proximal to a porin‐encoding gene in the majority of these strains, suggesting a role in iron acquisition and/or trafficking. Although a recent transposon dataset examining Methanococcus maripaludis grown in a variety of growth conditions did not find the MvoDUF2193 homolog (MMP_RS04980) as being essential under any of their growth conditions, and another previously published study examining the essentiality of Methanococcus maripaludis genes determined that both the M. maripaludis DUF2193 homolog and the proximal porin were non‐essential, the second study concluded that these genes were advantageous for the organism's growth [42, 43]. These results suggest that these proteins may be involved in the transport of essential nutrients [43]. DUF2193 family proteins harbor a conserved cysteine motif, which allows them to bind a single [4Fe–4S] cluster (Figs 1, 2, 3). Our biophysical characterization indicates that the oligomeric state of this protein is influenced by Fe‐loading: in the apo state, MvoDUF2193 exists primarily as a tetramer, while in the holo state, it shows substantial amounts of both monomer and tetramer (Fig. 4C). For both monomer and tetramer states, we observed that the protein can exist in oxidized and reduced forms, due to one or more intra‐subunit disulfide bonds (Fig. 6). Although further work is needed to establish the intracellular protein and cluster states, as well as the protein function, these observations have led us to propose a model of Duf2193 existing in a dynamic equilibrium between oligomeric states that is influenced by the [Fe–S] cluster binding and redox state of the protein (Fig. 7), suggesting the possibility that MvoDUF2193 is an iron–sulfur carrier that may also have a sensing or regulatory role.

Fig. 7.

Model of DUF2193 regulation between monomeric and tetrameric forms. MvoDUF2193 exists in an equilibrium between a monomer and a tetramer. Both the monomeric and tetrameric forms can contain a single intra‐subunit disulfide bond and a [4Fe–4S] cluster. In the monomeric form, the equilibrium favors the [4Fe–4S] cluster‐bound form, while when the protein exists as a tetramer, the apo form is favored.

To date, there are still a limited number of documented biological sensors that rely on changes in [Fe–S] clusters and oligomeric state to regulate redox sensing and activity. However, the number of proteins using this mechanism is rising in number and biological significance. A well‐studied example is fumarate and nitrate reduction regulator (FNR) protein, which controls the transition between anaerobic and aerobic respiration in E. coli depending on its oligomeric state [44, 45, 46, 47]. In the absence of oxygen, FNR coordinates a [4Fe–4S]²⁺ cluster and forms a dimer, which specifically binds DNA. In the presence of oxygen, the cluster is degraded, the protein dimer falls apart, and the ability to bind DNA is lost. Iron–sulfur cluster regulators (IscRs) serve as sensors of intracellular [Fe–S] cluster levels, depending on whether the protein is in the apo or holo state [48]. Our evidence suggests that similar to FNR, the oligomeric state of MvoDUF2193 is responsive to cluster coordination, with [Fe–S] cluster binding and redox state shifting the protein oligomeric equilibrium. Our in‐depth investigation suggests that DUF2193 family members are iron–sulfur binding proteins that may serve as regulators in methanogenic cells.

Author contributions

EMD—designed study, collected and analyzed data, drafted paper. JL—designed study, collected and analyzed data, contributed to paper writing. MT‐L—collected and analyzed data. JX—collected and analyzed data. YG—analyzed data and oversight of experimentation. WEB—oversight and design of project. BB—oversight and design of project, funding acquisition. JBB—design of project, oversight, writing, and editing of paper, funding acquisition.

Supporting information

Fig. S1. Functional protein network of MvoDuf2193 based on STRING analysis.

Fig. S2. DeepFRI biological process predictions made using MvoDUF2193 tetramer model.

Fig. S3. Temperature‐dependent EPR of reduced MvoDUF2193.

Fig. S4. Duf2193 has an oxygen‐stable Fe‐S cofactor.

Fig. S5. alphafold Multimer yields a high‐quality model of Duf2193 tetramer.

Fig. S6. Binding of [4Fe‐4S] is predicted to cause allosteric changes in dynamics.

Fig. S7. SDS/PAGE gel of MvoDUF2193 used for in‐gel digest prior to LC–MS/MS.

Fig. S8. DeepFRI molecular function analysis of MvoDuf2193 tetramer structure.

Fig. S9. Four cysteine residues that line a pore through the protein complex are potential disulfide bond locations.

Fig. S10. Structural model of the MvolDUF2193 tetramer.

Fig. S11. SDS/PAGE gel of IAA‐ and GEE‐labeled MvoDUF2193.

Table S1. DUF2193 family member genes and proximal porins.

Table S2. Interaction scores from STRING analysis.

Data accessibility

The data that support the findings of this study are openly available in OSF at http://doi.org/10.17605/OSF.IO/9AVCS.