# Cobalamin-mediated protection of Faecalibacterium duncaniae against oxidative stress: Insights from proteomic and membrane fatty acid profiles

Maria Alejandra de Angel Fontalvo, Simon Ménard, Rime Chebbo, Jasmina Vidic, Alban Amoros, Christine Péchoux, Lydie Oliveira Correia, Sébastien Dupont, Florence Dubois-Brissonnet, Laurent Beney, et al.

► To cite this version:

Maria Alejandra de Angel Fontalvo, Simon Ménard, Rime Chebbo, Jasmina Vidic, Alban Amoros, et al. Cobalamin-mediated protection of Faecalibacterium duncaniae against oxidative stress: Insights from proteomic and membrane fatty acid profiles. Microbiological Research, 2026, 306, pp.128455. 10.1016/j.micres.2026.128455. hal-05478791

HAL Id: hal-05478791

https://institut-agro-dijon.hal.science/hal-05478791v1

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# Cobalamin-mediated protection of *Faecalibacterium duncaniae* against oxidative stress: Insights from proteomic and membrane fatty acid profiles

Maria Alejandra de Angel Fontalvo<sup>a</sup>, Simon Ménard<sup>a</sup>, Rime Chebbo<sup>a</sup>, Jasmina Vidic<sup>a</sup>, Alban Amoros<sup>a</sup>, Christine Péchoux<sup>b</sup>, Lydie Oliveira Correia<sup>a</sup>, Sébastien Dupont<sup>c</sup>, Florence Dubois-Brissonnet<sup>a</sup>, Laurent Beney<sup>c</sup>, Bonastre Oliete<sup>c</sup>, Jean-Marc Chatel<sup>a</sup>, Sandrine Auger<sup>a, *d</sup>

## ARTICLE INFO

### Keywords:

* F. duncaniae

* Oxidative stress

* Proteomics

* Cobalamin

* Fatty acid

* Plasmalogen

## ABSTRACT

*Faecalibacterium* species are keystone commensals of the human gut, contributing to intestinal homeostasis, immune modulation, and epithelial health. However, their extreme sensitivity to oxygen and reactive oxygen species renders them highly vulnerable during inflammatory conditions, severely limiting their therapeutic application. Understanding the molecular mechanisms underlying their oxidative stress responses is therefore critical for harnessing these bacteria as next-generation probiotics to restore gut health. In this study, we investigated oxidative stress responses in *Faecalibacterium duncaniae* A2-165 using comprehensive proteomic and membrane fatty acid profiling. We demonstrated that increasing hydrogen peroxide (H₂O₂) concentrations extend the lag phase of growth and affect survival during the first hour of exposure, notably altering the redox potential. Exposure to H₂O₂ triggered a remodeling of the proteome, including detoxification systems, metal transporters, DNA repair systems, transcriptional regulators, and enzymes involved in cobalamin biosynthesis. Complementary RT-qPCR analyses revealed coordinated and time-dependent transcriptional activation of genes involved in oxidative stress response. Remarkably, cobalamin supplementation enhanced bacterial growth, mitigated H₂O₂-induced stress, and lowered superoxide levels in *F. duncaniae*, highlighting its direct antioxidant activity. By analyzing membrane fatty acid profiles, we showed that cobalamin preserves membrane fluidity, counteracting oxidative stress induced by H₂O₂ in *F. duncaniae*. These findings reveal the multifaceted strategies employed by *F. duncaniae* to withstand oxidative stress and provide a foundation for future efforts to optimize its production at industrial scales and its therapeutic potential as a next-generation probiotic.

## 1. Introduction

*Faecalibacterium* species are among the most prevalent and abundant strictly anaerobic bacteria in the healthy human gut microbiota (Piquer-Esteban et al., 2022) and play a central role in maintaining intestinal homeostasis and modulating immune responses (Martín et al., 2023). As butyrate producers, they provide energy to colonocytes, maintain the Th17/Treg balance, stimulate the production of anti-inflammatory cytokines such as interleukin (IL)-10, and inhibit pro-inflammatory mediators including IL-12 and interferon-γ (Lenoir et al., 2020; Sokol et al., 2008; Zhou et al., 2018). Additionally, bioactive

molecules such as the Microbial Anti-inflammatory Molecule (MAM) further contribute to their anti-inflammatory effects by inhibiting NF-κB signaling in intestinal epithelial cells (Auger et al., 2022a; Quévrain et al., 2016; Vilela Rodrigues et al., 2025). Certain strains can also prime IL-10-secreting T cells through dendritic cell modulation, highlighting their multifaceted role in mucosal immune tolerance (Alameddine et al., 2019).

The abundance and function of *Faecalibacterium* are strongly influenced by host health and colonic environmental conditions. Reductions in these bacteria have been consistently associated with metabolic and inflammatory disorders, including type 2 diabetes, colorectal cancer,

<footnote><sup>a</sup> Université Paris-Saclay, INRAE, AgroParisTech, MICALIS, Jouy-en-Josas 78350, France</footnote>

<footnote><sup>b</sup> Université Paris-Saclay, INRAE, AgroParisTech, GABI, Jouy-en-Josas 78350, France</footnote>

<footnote><sup>c</sup> Université Bourgogne Europe, Institut Agro, INRAE, UMR PAM, Dijon F-21000, France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: sandrine.auger@inrae.fr (S. Auger).</footnote>
irritable bowel syndrome, and inflammatory bowel disease (IBD) (Martín et al., 2023; Mclellan et al., 2025; Pandey et al., 2024). IBD, in particular, represents a major public health challenge, affecting millions of people worldwide. Current therapies mainly target inflammation rather than restoring microbial balance, highlighting the need for interventions that prevent disease recurrence and promote gut homeostasis. Oral administration of beneficial bacteria is a promising strategy, with restoration of *Faecalibacterium* populations emerging as a key target (Martín et al., 2023).

The sensitivity of *Faecalibacterium* to oxygen (O₂) and reactive oxygen species (ROS) limits their large-scale cultivation and oral delivery. *Faecalibacterium* species are widely classified as extremely oxygen-sensitive and survive only a few minutes of exposure to atmospheric oxygen (Botin et al., 2023; Duncan et al., 2002). In healthy individuals, an O₂ gradient exists along the gastrointestinal tract, decreasing from the stomach to the colon and radially toward the submucosa, where anoxia occurs at the luminal center (Kint et al., 2022). This gradient is crucial for maintaining strict anaerobiosis, and its disruption promotes dysbiosis by favoring facultative anaerobes such as *Enterobacteriaceae* while reducing strict anaerobes including *Faecalibacterium* (Rigottier-Gois, 2013; Zhou et al., 2018). During inflammation, immune cell recruitment increases ROS production, further exacerbating oxidative stress and disrupting microbial balance (Karhausen et al., 2004; Matthews et al., 2019; Muro et al., 2024; Sahoo et al., 2023).

Understanding how *Faecalibacterium* responds to oxidative stress is therefore critical for the development of effective probiotic formulations. Despite their vulnerability, these bacteria possess protective mechanisms, including antioxidants such as cysteine and riboflavin, as well as inducible O₂/ROS-detoxification systems that maintain redox balance (Botin et al., 2023; Khan et al., 2012). Recent studies suggest that O₂-tolerant variants of *F. prausnitzii* can survive O₂ exposure without losing their beneficial properties (Khan et al., 2023), offering potential avenues for clinical translation. Nevertheless, the molecular mechanisms underlying oxidative stress responses remain poorly understood, partly due to the lack of genetic tools for functional studies.

To address this gap, the present study investigates oxidative stress responses in the model strain *Faecalibacterium duncaniae* A2-165. To simulate acute oxidative bursts, *F. duncaniae* cells were exposed to varying concentrations of hydrogen peroxide (H₂O₂), mimicking the rapid release of ROS by immune cells into the intestinal lumen (Burgueño et al., 2019). To elucidate the mechanisms underlying the oxidative stress response in *F. duncaniae*, we performed proteomic analyses and membrane fatty acid profiling following H₂O₂ exposure. Taken together, this integrated approach provides a detailed characterization of oxidative stress responses in *F. duncaniae* A2-165 under controlled experimental conditions. Our results provide new insights into how this strict anaerobe responds to oxidative stress at a functional level, an aspect that has been far less explored than in aerobic or facultative anaerobic bacteria.

# 2. Materials and methods

## 2.1. Bacterial strain and growth conditions

The bacterial strain used in this work was *F. duncaniae* A2-165 (DSM 17677), originally isolated from human fecal material (Duncan et al., 2002). Cultivation was performed in a nutrient-rich, complex medium optimized to support the growth requirements of fastidious *Faecalibacterium* species (Beney et al., 2022). The formulation of this medium included 37 g/L plant-derived peptone (CAS-No. 100209-45-8, Organotechnie), 5 g/L selected yeast extract, 0.5 g/L L-cysteine, 6 g/L maltose, and 2.7 g/L sodium acetate trihydrate, 0.001 g/L pantothenic acid, 0.01 g/L biotin (B8), and 0.002 g/L pyridoxal hydrochloride. The medium was solidified via the addition of 15 g/L agar. Cultures were grown in a Jacomex chamber under anaerobic conditions (5 % H₂, 5 % CO₂, and 90 % N₂). During the experiments, strain A2-165 was cultured

in 15-mL or 50-mL centrifuge tubes (polypropylene; Sarstedt AG & Co).

## 2.2. Survival assays and growth in the presence of H₂O₂

After cultivation on agar plates, single colonies were inoculated into liquid medium and incubated at 37 °C. Overnight pre-cultures were used to inoculate 10 mL cultures at an initial optical density at 600 nm (OD₆₀₀) of 0.1, corresponding to ~10⁵ bacteria/mL. Oxidative stress was induced by adding hydrogen peroxide (H₂O₂; Thermoscientific) to final concentrations of 440, 880, 1320, 1760, 2640, 3520, or 4400 µM. The stability of H₂O₂ in the culture medium was assessed using the Amplex Red assay (Thermofisher), showing that H₂O₂ remained stable for up to 24 h in the used conditions (Supplementary Fig. S1). Aliquots were collected before stress induction (T0), and 1 or 3 h after H₂O₂ addition (T1h and T3h). Serial dilutions were prepared in a 96-well plate down to 10⁻⁴. From each dilution, 10 µL were spotted onto agar plates. Plates were tilted to facilitate droplet spreading and incubated at 37 °C for 48 h. Colony-forming units (CFU/mL) were then determined. Six independent experiments were performed.

For growth kinetics, overnight pre-cultures were used to inoculate 10 mL cultures at an initial OD₆₀₀ of 0.1. H₂O₂ was added at the desired final concentration. For each condition, 2 mL were transferred into 12-well deepwell polypropylene plates (Sarstedt AG & Co, Ref: 83.3921.500). Growth was monitored at 37 °C for 24 h under gentle agitation using a plate reader (Cerillo Stratus). When necessary, cyanocobalamin (Duchefa Biochemie) was added to a final concentration of 0.01 g/L. Experiments were conducted in triplicate with four independent biological replicates.

### 2.2.1. Transmission electron microscopy

Bacterial samples were fixed in the anaerobic chamber using 2 % glutaraldehyde prepared in 0.1 M sodium cacodylate buffer (pH 7.2) for 1 h at room temperature. Following fixation, cells were contrasted with 0.5 % Oolong tea extract in cacodylate buffer, then post-fixed with 1 % osmium tetroxide supplemented with 1.5 % potassium ferrocyanide. Samples were subsequently dehydrated through a graded ethanol series and embedded in Epon resin (Delta Microscopies). Ultrathin sections (~70 nm) were mounted on 200-mesh copper grids and counterstained with lead citrate. Observations were carried out using a Hitachi HT7700 transmission electron microscope operated at 80 kV, and images were captured with a CCD camera (Advanced Microscopy Techniques) in the MIMA2 platform (https://www6.jouy.inrae.fr/mima2/).

## 2.3. Redox potential measurement

Redox potential was monitored using a redox probe (Mettler Toledo, Switzerland). Baseline measurement was performed in the culture medium to establish the reference value. H₂O₂ was subsequently added to the medium at final concentrations of 440 µM, 880 µM, or 1320 µM, and redox potential was continuously recorded during 80 min.

## 2.4. Quantification of O₂⁻ free radicals

An assay with 2,3-bis (2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide (XTT; Sigma) was performed to measure the bacterial production of superoxide radical ions (O₂⁻). O₂⁻ production was monitored by following the formation of XTT formazan at 470 nm. XTT was dissolved in phosphate-buffered saline (PBS; pH 7) and added to the bacterial culture to a final concentration of 0.4 mM. Absorbance at 470 nm was monitored over time using an Infinite M200 luminescence reader (Tecan, Germany). No light adsorption was observed for the culture medium alone.

## 2.5. Protein extraction and preparation for mass spectrometry analysis

Bacterial cultures were performed as described above. Each
condition was tested in four independent biological replicates. Aliquots
were collected prior to stress induction (T0) and at 30 min (T30) or
60 min (T60) following the addition of H₂O₂ at a final concentration of
440 µM. Cells were harvested by centrifugation at 4500 rpm for 3 min at
4 °C, and the supernatants were discarded. Bacterial pellets were stored
at -20 °C until further processing. For protein extraction, pellets were
resuspended in lysis buffer (50 mM Tris-HCl, pH 8.0) supplemented with
protease inhibitor cocktail (50 µL/mL; Roche). Cell disruption was
performed by sonication on ice (six cycles of 10 s at 20 % amplitude).
The lysates were centrifuged at 13,000 × g for 15 min at 4 °C. The
resulting supernatants were centrifuged at 15,000 × g for 2 h at 4 °C to
separate the soluble protein fraction from insoluble material. The final
supernatants, corresponding to the cytoplasmic protein-enriched frac-
tion, were collected. Protein concentrations were quantified using the
Bradford assay. For electrophoretic separation, 10 µg of proteins were
loaded onto a NuPAGE 4–12 % Bis-Tris gel (Novex) and subjected to a
short migration, allowing proteins to migrate 0.5 cm. Each lane was
excised and sent to PAPPSO plateform facilities (http://pappso.inrae.
fr/) for proteomic analysis. Briefly, disulfide bridges were reduced by
10 mM DTT for 1 h at 56 °C and resulting reduced cysteine were alky-
lated by 50 mM iodoacetamide for 1 h at room temperature before the
step of enzymatic digestion with 100 ng trypsin (Promega) for 18 h at 37
°C. Tryptic peptides were recovered by washing the gel pieces twice in
0.2 % trifluoroacetic acid - 50 % acetonitrile and once in 100 % aceto-
nitrile. The extracts were evaporated to dryness. The peptides were then
resuspended in 200 µL of loading buffer (0.1 % formic acid and 2 %
acetonitrile in H₂O), prior to LC-MS/MS injection.

2.6. Peptide LC-MS/MS analysis

Peptide separation and mass spectrometry were carried out on a
NanoElute2 LC system (Bruker) coupled to a timsTOF Pro mass spec-
trometer (Bruker) equipped with a CaptiveSpray source. Tryptic pep-
tides mixtures (1 µL) were first loaded at 50 bar onto a C18 precolumn
(Acclaim PepMap, 5 µm, 20 mm × 100 µm i.d.; Thermo Scientific) before
separation on a 25 cm × 75 µm analytical column (IonOpticks,
Australia) packed with 1.7 µm C18 particles and fitted with an inte-
grated emitter tip. The column was maintained at 50 °C, with peptides
eluted at a constant flow rate of 250 nL/min. The gradient was pro-
grammed as follows: from 2 % to 30 % buffer B (0.1 % formic acid in
acetonitrile) over 70 min, followed by a ramp to 85 % buffer B main-
tained for 12 min, and finally re-equilibration at 98 % buffer A (0.1 %
formic acid in water) for 2 min. Data were acquired on the timsTOF Pro
in dia-PASEF mode, using 24 isolation windows distributed across 12
dia-PASEF scans. The timsTOF Pro was operated with the following
parameters: mass range of 100–1700 m/z, ion mobility range of 1/K₀
= 0.60–1.60 V·s/cm², ramp time of 100 ms, duty cycle fixed at 100 %,
capillary voltage of 1600 V, dry gas flow of 3 L/min, and dry tempera-
ture set to 180 °C. For dia-PASEF acquisition, MS/MS spectra were
collected across a mass range of 352.7–953.7 Da, with an ion mobility
window of 1/K₀ = 0.70–1.10 (estimated cycle time: 1.38 s). Collision-
induced dissociation (CID) energies were ramped from 20 eV at 1/K₀
= 0.70 up to 59 eV at 1/K₀ = 1.60. Instrument calibration was performed
using three reference ions from the Agilent ESI LC/MS tuning mix (m/z
622.0289, 1/K₀ = 0.9848 V·s/cm²; m/z 922.0097, 1/K₀ = 1.1895 V·s/
cm²; and m/z 1221.9906, 1/K₀ = 1.3820 V·s/cm²).

2.7. Protein identification and data processing

Protein identification was carried out using DIA-NN (v1.8) in library-
free mode, employing deep learning-based spectral prediction. Peptides
were generated *in silico* with cleavage rules set for lysine (K\*) and
arginine (R\*), allowing a maximum of one missed cleavage, with peptide
lengths constrained between 7 and 30 amino acids. Fragment ion *m/z*
values were limited to 200–1800, while precursor ions were restricted to
300–1800 *m/z* and charge states from 1 + to 4 + .

Carbamidomethylation of cysteines was defined as a fixed modification, and N-terminal methionine excision was enabled. Protein identification was performed against a FASTA database including both an in-house contaminant collection and the *F. duncaniae* reference proteome (NCBI, GCA_002734145). DIA-NN generated a predicted spectral library and reanalyzed the data, prioritizing predicted spectra over experimental ones when higher confidence was achieved. Protein inference used the relaxed grouping strategy. Quantification outputs consisted of precursor-level matrices filtered at a 1 % false discovery rate (FDR), with mass accuracy thresholds set at 15 ppm for both MS1 and MS2 levels. Relative quantification was based on the report.tsv file produced by DIA-NN.

Peptides mapping to multiple entries in the protein.group column of the report.tsv file were excluded. Data quality control, normalization, filtering, and statistical processing were conducted with MCQR (Balliau et al., 2025). Peaks showing instability in retention time (standard deviation > 30 s) were discarded, and only peaks with retention times between 400 and 4800 s were retained. To account for global variation across LC-MS runs, normalization was applied using the diff.median. RT normalisation method. A repeatable peptide was defined as one detected in at least 11 out of 12 samples. Proteins were quantified if represented by at least two reproducible peptides. Missing intensity values were imputed by linear regression, using the values of other peptides belonging to the same protein. Protein-level intensities were then calculated as the sum of normalized signals from their specific reproducible peptides.

2.8. RNA extraction and purification

Strain A2-165 was exposed to oxidative stress by the addition of 440 µM H₂O₂, under the same conditions as for proteomic analysis. Samples were collected at T0 (before stress) and at 5, 10, 15, 30, and 60 min post-treatment, centrifuged at 3000 rpm for 3 min at 4 °C, and cell pellets were stored at -70 °C. For RNA extraction, pellets were resuspended in TRIS-EDTA buffer (25 mM TRIS, pH 7.5; 85 mM EDTA) and lysed with glass beads (<106 µm) in the presence of phenol:chloroform:isoamyl alcohol (25:24:1, equilibrated with 50 mM sodium acetate, pH 5) using a bead-beater (three cycles of 30 s at 6300 rpm, 30 s pause). Lysates were sequentially extracted with phenol:chloroform: isoamyl alcohol and chloroform, and RNA was precipitated overnight at -20 °C with isopropanol and sodium acetate, washed with 70 % ethanol, air-dried, and resuspended in DNase/RNase-free water. RNA concentration was measured at 260 nm using a NanoDrop™ spectrophotometer. Next, RNA samples were treated with DNase using the RNase-Free DNase Set (Qiagen) to eliminate any residual genomic DNA. Clean-up and concentration were performed using the RNA Clean-up and Concentration Micro Kit (Norgen Biotek).

2.9. Quantification of mRNA by qPCR

Complementary DNA (cDNA) was synthesized from 1 µg of total RNA using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems). The resulting cDNA was diluted 1:100 prior to qPCR analysis. Each qPCR reaction (total volume 20 µL) consisted of 12.5 µL of 2X SYBR Green Master Mix, 2.5 µL of DNase/RNase-free ultrapure water, 0.5 µL of each primer (forward and reverse), and 4 µL of diluted cDNA. qPCR was performed on a StepOne Plus thermocycler using the following program: an initial denaturation at 95 °C for 10 min, followed by 40 cycles of 95 °C for 15 s (denaturation) and 60 °C for 1 min (annealing/extension). Fluorescence was measured at the end of each cycle. A final melt curve analysis, ramping to 95 °C, was performed to determine the melting temperature of the PCR products and to verify the absence of non-specific amplification, such as primer-dimers. The cycle threshold (Ct) was defined as the point at which fluorescence exceeded the threshold level. Ct values were normalized to a reference gene encoding a translation elongation factor G, and relative gene expression was calculated
using the comparative ΔΔCt method.

## 2.10. Membrane fatty acids extraction and GC-MS analysis

Bacteria were harvested by centrifugation (3000 × g, 5 min), rapidly frozen in liquid nitrogen, and stored at -80 °C until further processing. Extraction and methylation of fatty acids (FA) were carried out on bacterial pellets as described (Touche et al., 2023). Whole-cell FA were first saponified and esterified using methanolic NaOH (1 mL of 3.75 M NaOH in 50 % (v/v) methanol and incubation for 30 min at 100 °C), followed by treatment with methanolic HCl (addition of 2 mL of 3.25 M HCl in 45 % (v/v) methanol solution and incubation for 10 min at 80 °C). Fatty acid methyl esters (FAME) were then extracted using a 1:1 (v/v)

diethyl ether/cyclohexane mixture. The organic phase was washed with a dilute base solution (0.3 M NaOH).

Analytical gas chromatography of FAME was carried out in a GC-MS Trace 1300 / ISQ 7000 system (Thermo Fisher Scientific) equipped with a BPX70 capillary column (25 m, 0.22-mm internal diameter; SGE, Victoria, Australia). The column temperature was initially maintained at 100 °C for 1 min, then gradually increased to 170 °C at a rate of 2 °C per minute. FA species were identified based on mass spectrometry (MS) databases (Replib, Mainlib, FAME2011). The relative abundance of each FA was calculated as a percentage of the total FAME peak area. Identified FA species were grouped into the following categories: saturated FA (SFA), unsaturated FA (UFA), saturated plasmalogens (SP), and unsaturated plasmalogens (UP).

<figure><img src="image_2.png" /><figcaption>**Fig. 1.** Growth and survival of *F. duncaniae* A2-165 in response to oxidative stress induced by H₂O₂. (A) Growth kinetics under various H₂O₂ concentrations. Data represent mean ± SD of three independent biological experiments. (B) Quantification of bacterial survival expressed as CFU/mL at three time points (T0, T1h, and T3h) after the addition of different H₂O₂ concentrations. Shown is the average of six independent biological experiments. (C) Changes in redox potential following the addition of H₂O₂ to the medium. (D) Representative transmission electron microscopy images of bacterial cells under control conditions and after exposure to oxidative stress induced by H₂O₂. Black scale bar, 5 µm. Yellow arrows indicate detachment of the cell wall or cytoplasmic efflux.</figcaption></figure>
## 2.11. Laurdan generalized polarization

The generalized polarization of the lipophilic dye Laurdan (6-dodecanoyl-2-dimethylaminonaphthalene) was used as a measure of the bacterial membrane fluidity (Wenzel et al., 2018). Bacteria were harvested by centrifugation (3000 × g, 5 min), washed once with PBS and incubated for 20 min in the dark with 10 µM of Laurdan (Sigma-Aldrich, France) from a 1 mM stock in dimethylformamide (DMF). The stained cells were washed four times with Laurdan 459 buffer (137 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄, 1.8 mM KH₂PO₄, 0.2 % glucose, 1 % DMF) and transferred to a dark clear-bottom 96-well plates (Corning, France). Laurdan fluorescence was measured between 400 nm and 540 nm under excitation at 350 nm using a Tecan Spark® at 30°C. Laurdan GP was calculated using the formula: $GP = (I_{450} - I_{500})/(I_{450} + I_{500})$, where I corresponds to the fluorescence intensity value at the recorded emission wavelength. An increase of Laurdan GP values was interpreted as a reduction of plasma membrane fluidity.

# 3. Results

## 3.1. Growth and survival of *F. duncaniae* A2–165 under H₂O₂ stress

In a first step, we evaluated the growth capacity of strain A2-165 in the presence of varying H₂O₂ concentrations (440 µM to 2640 µM, including an H₂O₂-free control). H₂O₂ was added at the onset of the exponential growth phase (10⁵ bacteria/mL), and its effect on growth kinetics was monitored over 24 h. At H₂O₂ concentrations of 440 µM and above, increasing H₂O₂ led to a prolonged lag phase and a lower maximum OD₆₀₀ compared to the unstressed condition (Fig. 1A). Remarkably, A2-165 was able to grow even at 2640 µM H₂O₂, although with a 14-hour lag phase, highlighting its capacity to survive and adapt under high oxidative stress.

We next quantified survival under H₂O₂ stress after 1- and 3-hours exposures across a broad range of concentrations by determining bacterial counts (CFU/mL). Following 1 h of exposure, CFU/mL decreased with increasing H₂O₂ concentrations, some bacteria still survived even at 4400 µM H₂O₂ (Fig. 1B). After 3 h, CFU/mL increased under all conditions, indicating that inhibitory effect of H₂O₂ was transient and highlighting the strain's capacity to withstand high oxidative stress. It should be noted that CFU counts reflect the ability to form colonies and may not capture cells in a Viable But Non-Culturable state, which has not been described for *Faecalibacterium*, so the term "survival" should be interpreted as maintenance of culturability under H₂O₂ stress rather than absolute cell viability. As shown in Fig. 1C, the addition of H₂O₂ to the medium caused a concentration-dependent increase in redox potential (up to -110 mV after 2 min at 1320 µM H₂O₂). In this study, redox potential measurements were used as an indicator of the extracellular oxidative environment experienced by *F. duncaniae* and were not intended to directly quantify intracellular redox homeostasis. After 1 h, the redox potential gradually returned to the baseline level of the control condition (-290 ± 20 mV) consistent with the transient stress observed during the first hour after H₂O₂ addition (Fig. 1B). The observed recovery of redox potential after H₂O₂ addition may reflect either bacterial detoxification or chemical decomposition of H₂O₂ in the medium.

In addition, we examined the subtle differences in cellular morphology induced by 1-hour exposure to H₂O₂ using electron microscopy (Fig. 1D). Cells exposed to 440 µM H₂O₂ displayed an intact surface morphology similar to that of the unstressed control. In contrast, at 880 µM H₂O₂, marked structural alterations were observed, including cell wall detachment and, in some cases, release of the cytoplasmic content, indicative of a loss of membrane integrity.

The characterization of these oxidative stress conditions with H₂O₂ was subsequently used to investigate the proteomic response of strain A2-165 to oxidative stress. **Proteomic response of *F. duncaniae* A2-165 to H₂O₂**

The oxidative stress response was investigated by whole-cell

proteomics using a concentration of 440 µM H₂O₂, which preserved cellular integrity (Fig. 1D), after 30 and 60 min of exposure (T30 and T60). These time points were selected to capture potential differences between early and later stages of the response, as previously observed in *Escherichia coli* (Rodríguez-Rojas et al., 2020). Following protein extraction, samples were analyzed at the PAPPSO proteomics platform (INRAE, Jouy-en-Josas). A total of 1887 proteins were initially identified across all samples. After peptide chromatographic filtering, normalization, and removal of non-reproducible or unreliable peptides, 1370 proteins were retained for statistical analyses. This represents approximately 50 % of the predicted proteome of *F. duncaniae* A2-165, which comprises 2815 predicted genes (Supplementary Table S1). To focus on proteins exhibiting substantial changes across the three experimental conditions (T0, T30, and T60), proteins with low variation were first filtered out. The selection criteria was a fold-change (maximum/minimum abundance across conditions) greater than 1.5. Subsequent ANOVA analysis identified 39 proteins whose abundance changed significantly between T30 (or T60) and T0. Multiple testing correction was applied uniformly across all comparisons using the Benjamini-Hochberg method to control the false discovery rate (FDR). Proteins with FDR-adjusted p-values < 0.05 were considered significantly altered. The protein abundance profiles are illustrated in Fig. 2A. Hierarchical clustering based on Euclidean distance revealed a clear segregation of the samples into three distinct clusters corresponding to the experimental time points (T0, T30, and T60), reflecting the temporal dynamics of the oxidative stress response.

Among the significantly more abundant proteins, several were directly associated with oxidative stress defense, including reverse rubrerythrin, rubrerythrin, and flavodoxin, which are known to contribute to O₂ or H₂O₂ detoxification in strict anaerobic bacteria (Fig. 2B) (Morvan et al., 2021). Changes were also detected in proteins involved in metal homeostasis, such as a P-type ATPase implicated in heavy metal transport, as well as the ferrous iron transport proteins A and B (Fig. 2B). We further observed the more abundance of proteins involved in DNA replication and repair (RecF, RecX, and exonucleases), which may be involved in the repair of DNA damaged due to oxidative stress. In addition, we also detected increased level of acetyl-CoA carboxylase, the enzyme catalyzing the first committed step of fatty acid biosynthesis. This finding points potential remodeling of plasma membrane fatty acid composition, aimed at maintaining the required membrane fluidity and thereby counteracting the deleterious effects of oxidative stress (Maslovska et al., 2023).

Remarkably, functional protein-protein interaction analysis with the set of 39 proteins revealed 23 proteins associated with the cobalamin biosynthesis pathway, 11 of which are directly involved in this metabolic process (Fig. 2C). This suggests a potential link between oxidative stress adaptation and cobalamin (also named vitamin B₁₂) metabolism. Finally, multiple transcriptional regulators were differentially abundant (Fig. 2B), indicating that global regulatory networks may be engaged to orchestrate the proteomic response. These observations raise the possibility that part of the observed proteomic remodeling reflects transcriptionally driven changes, with transcriptional regulators exerting influence over diverse functional pathways.

## 3.2. Transcriptional response of target genes to H₂O₂

We next examined whether variation in protein abundance correlated with transcriptional changes of the corresponding genes for six selected proteins of interest. The selected candidates included three genes encoding detoxification enzyme (ATP00803.1, ATP00391.1, ATO98703), two genes encoding proteins involved in metal ion transport (ATO99389.1, ATP00703.1), and one gene encoding an enzyme of the cobalamin biosynthesis pathway (ATP00528.1) (Table 1).

In this purpose, RNA was extracted from cultures exposed to the same oxidative stress conditions used in our proteomic analysis (440 µM H₂O₂). Samples were collected from unstressed cultures (T0) and after 5,
<figure><img src="image_3.png" /><figcaption>**Fig. 2.** Comparative protein abundance between H₂O₂ treated cells and the control condition in *F. duncaniae* A2–165. (A) Heatmap of the 39 differentially abundant proteins was generated in R (v4.3.1) from a row-standardized normalized matrix. Proteins were clustered using Euclidean distance and complete linkage, with a blue-to-red color scale indicating low to high abundance. The four biological replicates for each condition (T0, T30, and T60) are shown. (B) Fold-change in protein abundance between 30 min of oxidative stress (T30) and the control condition (T0). Proteins were classified in metabolic processes. (C) Functional enrichment</figcaption></figure>
analysis of protein-protein interaction networks with the set of 39 proteins based on the comparison of the three conditions (T0, T30, and T60) using the STRING tool. One cluster involves proteins related to the cobalamin biosynthesis pathway (shown in red circle).

<table><caption>**Table 1**

List of qPCR primers used in this study.</caption><thead><tr><th>Sequence</th><th>Target (locus_tag)</th></tr></thead><tbody><tr><td>ACACCCTGACCTTTGTGCTT</td><td>Forward ATP00803.1 Flavodiiron protein</td></tr><tr><td>CAGTCGGCGTTTTTCGTCAAA</td><td>Reverse ATP00803.1 Flavodiiron protein</td></tr><tr><td>CGCCAAGATGTGGTTCAAGG</td><td>Forward ATP00391.1 Rubrerythrin</td></tr><tr><td>GAAGCCCTCTTCCTTGGCTT</td><td>Reverse ATP00391.1 Rubrerythrin</td></tr><tr><td>GTCGTGACCGATAGCACCAA</td><td>Forward ATO98703 Reverse rubrerythrin</td></tr><tr><td>TTGCCATTTCGTGGACGGTA</td><td>Reverse ATO98703 Reverse rubrerythrin</td></tr><tr><td>CTATATGCCCGCACCTACCG</td><td>Forward ATP00910.1 Translation elongation Factor G</td></tr><tr><td>ACCGGAGTAGACACGGAAGT</td><td>Reverse ATP00910.1 Translation elongation Factor G</td></tr><tr><td>GCCGAGGGACATACAGTTGT</td><td>Forward ATO99389.1 Heavy metal translocating P-type ATPase</td></tr><tr><td>GGTCACCAGCTCTTCCAGAC</td><td>Reverse ATO99389.1 Heavy metal translocating P-type ATPase</td></tr><tr><td>CGACCTGAAGAAGCTGAGCA</td><td>Forward ATP00703.1 Ferrous iron transporter B</td></tr><tr><td>TCCACACTGCCGGTAAAGAC</td><td>Reverse ATP00703.1 Ferrous iron transporter B</td></tr><tr><td>GGGTCTACGGCATGGCAAG</td><td>Forward ATP00528.1 Precorrin-3B C(17)-methyltransferase</td></tr><tr><td>AGGGAGAGCACGCAGAAATC</td><td>Reverse ATP00528.1 Precorrin-3B C(17)-methyltransferase</td></tr></tbody></table>

15, 30, and 60 min of oxidative stress. Transcriptional levels of the six genes were quantified by RT-qPCR and normalized to a validated housekeeping gene, *tefG*, which encodes a translation elongation factor

and exhibits stable expression under oxidative stress conditions (Supplementary Fig. S2).

Exposure to H₂O₂ triggered robust transcriptional induction of all six genes (Fig. 3). For most genes, transcript levels peaked at 15 min, reflecting a rapid but transient transcriptional response. Notably, the reverse rubrerythrin and rubrerythrin encoding genes displayed maximal induction as early as 5 min, highlighting their particularly swift response. These results indicate that oxidative stress elicits a prompt transcriptional activation of genes involved in detoxification, metal ion transport, and cobalamin biosynthesis, supporting their functional roles in the cellular adaptation and stress defense mechanisms.

### 3.3. Direct protective effect of cobalamin against H₂O₂ stress

Following the observed induction and significant abundance of several enzymes involved in the cobalamin biosynthesis pathway, we hypothesized that *F. duncaniae* A2–165 might utilize this vitamin as a protective mechanism against oxidative stress. To test this hypothesis, bacterial growth was monitored under oxidative stress conditions, comparing cultures with and without cobalamin supplementation (Fig. 4A). The concentration of 0.01 g/L cobalamin was initially selected based on the rationale used for vitamin B8 in the optimized medium (Beney et al., 2022) to ensure a sufficiently high dose to potentially observe an effect. Cobalamin was introduced into the culture medium at

<figure><img src="image_4.png" /><figcaption>**Fig. 3.** Expression patterns of genes likely involved in response to oxidative stress in *F. duncaniae* A2–165. Gene expression was analyzed by qRT-PCR and normalized to the reference TefG encoding-gene. The results were expressed as a ratio after H₂O₂ exposure compared with the unstressed condition (time at 0 min). (A to F) Expression of the six genes of interest after 0, 5, 15, 30, or 60 min of exposure to 440 µM H₂O₂. The average and standard deviation of three independent biological experiments are shown. Statistically significant increases in expression compared with T0 are indicated with asterisks (Mann-Whitney; *, P, 0.05). Note that the ordinate scale is different for each panel.</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Fig. 4.** Impact of cobalamin supplementation on *F. duncaniae* A2-165 growth and resistance to oxidative stress. (A) Growth kinetics under various H₂O₂ concentrations in the presence or absence of 0.01 g/L cobalamin (N = 4). (B, C and D) Effect of cobalamin in 40 mL tubes on the maximal OD₉₀₀, cell counts (CFU/mL) and overall biomass (weight of the cellular pellet). Individual data points (n = 5 independent biological replicates) are shown together with mean ± SD. Statistically significant are indicated with asterisks (Mann-Whitney; *, P, 0.05). (E) Changes in redox potential following the addition of H₂O₂ to the medium supplemented with 0.01 g/L cobalamin. (F) Quantification of O₂⁻ produced by bacterial cells. O₂⁻ production was monitored by following the formation of XTT-formazan at 470 nm. At time zero (T0), cells were exposed to air. (G) Predicted homocysteine to S-adenosylmethionine biosynthetic pathway in strain A2-165 (H) Growth kinetics in the presence of various concentrations of methionine (N = 3 biological replicates).</figcaption></figure>
time zero (T0), before addition of H₂O₂. Cultures supplemented with cobalamin exhibited higher OD₆₀₀ values under both un-stressed conditions and in the presence of 440 µM H₂O₂ (Fig. 4A). Under oxidative stress with 880 µM or 1320 µM H₂O₂, addition of cobalamin had a pronounced effect on the growth rate during the exponential phase, with a minor effect on lag duration (Fig. 4A). This indicated that cobalamin primarily supports metabolic recovery and biomass accumulation after the initial oxidative stress, rather than acting as a primary first responder during the lag phase.

Since microplate cultures showed a higher OD₆₀₀ in the presence of cobalamin, we examined whether this increase truly reflected higher biomass production. Growth assays performed in larger volumes (40 mL culture tubes) confirmed that cobalamin supplementation increased the maximal OD₆₀₀, which correlated with higher viable cell counts (CFU/mL) and greater overall biomass (Fig. 4B–D). We next assessed whether the protective effect of cobalamin against H₂O₂ stress was dose-dependent by supplementing cultures with 0.01 or 1 g/L cobalamin. A 10-fold higher cobalamin concentration improved survival in the presence of 880 µM H₂O₂ (Supplementary Fig. S3). Thus, cobalamin clearly exerts a protective effect on *F. duncaniae* against oxidative stress. Subsequently, to elucidate the protective function of cobalamin, we examined its effect on redox potential following H₂O₂ addition. As shown in Fig. 4E, the presence of cobalamin did not significantly alter the H₂O₂-induced changes in redox potential. Then, we tested the antioxidant properties of cobalamin. *F. duncaniae* cells were briefly exposed to ambient air to induce superoxide anion production, which was measured using XTT (Fig. 4F). Cobalamin supplementation significantly reduced superoxide generation, demonstrating a direct antioxidant effect.

Metabolic reconstruction of *F. duncaniae* indicates that methionine synthase is the only enzyme requiring cobalamin as a cofactor (Fig. 4G) (Heinken et al., 2014). Since methionine and S-adenosylmethionine are central to protein synthesis and methylation-dependent regulation, we tested whether cobalamin's growth-promoting effect was mediated via the methionine pool by supplementing cultures with L-methionine at various concentrations. While methionine slightly enhanced growth, it did not replicate the effect of cobalamin (Fig. 4H), showing that cobalamin promotes growth through mechanisms independent of methionine availability. In addition, we tested the effect of 200 µM methionine supplementation in the growth medium on resistance to a range of H₂O₂ concentrations. This supplementation did not confer significant increased resistance, indicating that methionine does not provide a protective effect against H₂O₂ under the conditions tested (data not shown).

### 3.4. Temporal analysis of membrane fatty acid composition during bacterial growth

Given that proteomic analysis revealed an increased abundance of acetyl-CoA carboxylase in response to oxidative stress, we investigated the effect of H₂O₂ on membrane fatty acid (FA) composition. First, we performed a comprehensive characterization of the membrane FA profile across different growth phases in strain A2-165 (Fig. 5A and B). Gas chromatography coupled with GC-MS revealed a remarkably broad spectrum of membrane FA. These included conventional saturated and unsaturated FA, which contribute to membrane structural organization and fluidity, and specialized lipid components such as plasmalogen aldehydes (ALDE) and dimethyl acetals (DMA) (Supplementary Table S2). The presence of these compounds is particularly noteworthy, as they are often associated with unique membrane properties, including protection against reactive oxygen species (Dean and Lodhi, 2018). Genome analysis of strain A2-165 identified an operon comprising two genes encoding proteins homologous to PlsA and PlsR from *Clostridium perfringens*, which are involved in plasmalogen synthesis (Fig. 5C) (Jackson et al., 2021). The proposed reaction catalysed by plasmalogen synthase converts diacyl lipids into plasmalogens (Fig. 5D). Using the Schiff

reagent, which reacts with aldehydes, we visualised plasmalogen-producing bacterial colonies from strain A2-165 on agar plates (Fig. 5E).

Classification of conventional FA and plasmalogens into saturated and unsaturated categories revealed an increase in saturated species and a concomitant decrease in unsaturated species as the culture reached the stationary phase (Fig. 5F). Given that higher proportions of saturated FAs are expected to reduce membrane fluidity, we monitored fluidity throughout the growth phase using the Laurdan dye (Wenzel et al., 2018). Cells in the stationary phase displayed the lowest membrane fluidity (Fig. 5G), consistent with the decreased proportion of unsaturated FAs (Fig. 5F). These findings indicate that *F. duncaniae* membrane composition and fluidity are dynamically modulated during growth, emphasizing the importance of carefully considering growth phase when studying the effects of oxidative stress.

### 3.5. Effect of H₂O₂ exposure and cobalamin on membrane fatty acid profile

We further analyzed membrane FA composition after 1 h and 3 h of exposure to 440 or 880 µM H₂O₂, which was added during the early logarithmic phase. Addition of H₂O₂ at 440 or 880 µM resulted, after 1 h, in a significant increase in conventional saturated FAs, concomitant with a decrease in conventional unsaturated FAs (Fig. 6 A and B). Plasmalogen composition was not significantly affected (Supplementary Table S2). However, 3 h after H₂O₂ addition, no significant differences were observed compared with the unstressed control. This result indicates that the bacterial membrane of strain A2-165 tends to become more rigid under oxidative stress. This effect is transient during the first hour, consistent with the temporary effect on growth (Fig. 1B).

We then tested the effect of cobalamin on FA composition changes in response to oxidative stress. Cobalamin was added in the growth medium before addition of H₂O₂. In the presence of cobalamin, 1 h after oxidative stress, the levels of conventional saturated FAs remain significantly lower than in its absence, and conversely for unsaturated FAs (Fig. 6C and D). This finding highlights the protective role of cobalamin against oxidative stress, likely by stabilizing membrane structure and maintaining its functional integrity under challenging conditions.

## 4. Discussion

Although *Faecalibacterium* species are widely classified as extremely oxygen-sensitive, this study demonstrated that *F. duncaniae* A2-165 exhibited remarkable resilience to H₂O₂ stress, illustrating a distinction between atmospheric oxygen sensitivity and adaptive oxidative stress responses. We challenged the strain with H₂O₂ concentrations ranging from 440 to 4400 µM, thereby probing its ability to withstand high levels of oxidative stress under the experimental conditions. While increasing H₂O₂ concentrations prolonged the lag phase and reduced the maximum OD₆₀₀, strain A2-165 was still able to proliferate after exposure to 2640 µM H₂O₂, highlighting its adaptive potential. Survival says further demonstrated that the inhibitory effects of H₂O₂ were transient, with CFU/mL recovered after 3 h even at concentrations up to 4400 µM. In our conditions, strain A2-165 resisted higher H₂O₂ concentrations than recently reported (Zünd et al., 2025), which underscored the key role of culture medium composition in modulating resistance phenotypes. Although direct *in vivo* measurements of H₂O₂ in the colon remained challenging, studies such as Burguño et al. reported that epithelial cells produced H₂O₂ on the order of picomoles under chronic inflammation (Burguño et al., 2019). Taken together, these results underscored the robustness of A2-165 and its capacity to withstand physiologically relevant oxidative stress.

Proteomic analysis revealed a coordinated response to oxidative stress. Interestingly, *F. duncaniae* A2-165 showed the same induced proteins after 30 and 60 min of exposure, in contrast to *E. coli*, which
<figure><img src="image_6.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_7.png" /><figcaption>C</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>D</figcaption></figure>

<figure><img src="image_9.png" /><figcaption>E</figcaption></figure>

<figure><img src="image_10.png" /><figcaption>F</figcaption></figure>

<figure><img src="image_11.png" /><figcaption>G</figcaption></figure>

<figure><img src="image_12.png" /></figure>

(caption on next page)
<figure><img src="image_13.png" /><figcaption>**Fig. 5.** Membrane fatty acid profile of strain F. duncaniae A2-165 during various stages of growth. (A) FA species identified by gas chromatography are presented as the proportions of the total adjusted to 100%. Data represent mean ± SD of four independent biological experiments. (B) Schematic representation of the different growth phases tested. (C) Genetic architecture and domain organization of the putative plasmalogen synthase operon in strain A2-165 (top), and corresponding protein domain architecture (bottom). (D) Putative enzymatic reaction to convert diacyl lipids into plasmalogens. (E) Colonies of strain A2-165 grown on agar exposed to Schiff stain. Pink colonies correspond to plasmalogen production. (F) Relative abundance of unsaturated plasmalogens (UP), unsaturated fatty acids (UFA), saturated plasmalogens (SP), and saturated fatty acids (SFA) during the various stage of growth. Means ± SD are shown (G) Membrane fluidity evaluated using the Laurdan generalized polarization method (Laurdan GP). Laurdan GP = (I₄₄₀ - I₄₉₀)/(I₄₄₀ + I₄₉₀), where I₄₄₀ and I₄₉₀ are the emission intensities at 440 and 490 nm, respectively, when excited at 350 nm. Individual data points (n = 6 biological replicates) are shown together with mean ± SD. P values were determined by one-way ANOVA (***p < 0.0001; ** p < 0.0008).</figcaption></figure>

<figure><img src="image_14.png" /><figcaption>**Fig. 6.** Impact of oxidative stress and cobalamin on membrane fatty acid profile. FA species are presented as the proportions of the total adjusted to 100%. Data represent mean ± SD of four independent biological experiments. (A and B) Proportion of saturated and unsaturated fatty acids after 1 h and 3 h stress with H₂O₂. (C and D) Proportion of saturated and unsaturated fatty acids after H₂O₂ stress in the presence or absence of cobalamin on the growth medium. P values were determined by one-way ANOVA (***p < 0.0001; ** p < 0.0008; *p < 0.05).</figcaption></figure>

exhibited distinct early and late proteomic responses (Rodríguez-Rojas et al., 2020). This suggests that A2-165 promptly activated its entire oxidative stress resistance machinery upon encountering the stress. Such a rapid and sustained response likely reflects an evolutionary strategy to ensure immediate protection against ROS, minimizing cellular damage and maintaining redox homeostasis. Notably, proteins directly involved

in detoxification, including rubrerythrin, reverse rubrerythrin, and flavodoxin, were upregulated, supporting their role in H₂O₂ detoxification in F. duncaniae, as reported in other strict anaerobes (Morvan et al., 2021). We also observed that proteins linked to metal homeostasis, particularly iron transporters and P-type ATPases, were more abundant in A2-165 after H₂O₂ stress. Given the key role of iron metabolism in
oxidative stress, particularly through the generation of highly reactive
radicals via the Fenton reaction (Imlay et al., 1988; Luo et al., 1996),
these findings strongly suggest that modulation of metal ion transport
constitutes an essential adaptive mechanism in F. duncaniae
(Chandrangsu et al., 2017). Remarkably, it was recently demonstrated
that the effect of *Faecalibacterium* in reducing ovarian tumor growth was
associated with ferroptosis and metal ion homeostasis pathways (Ding
et al., 2026). The regulation of metal ion homeostasis by *Faecalibacter-
ium* and its potential effects on host health represent an important area
that deserves further research. Concurrently, DNA repair proteins, such
as RecF, RecX, and exonucleases, were upregulated, reflecting the need
to repair oxidative lesions and prevent mutagenesis (Carvajal-Garcia
et al., 2023; Foster et al., 2015). Indeed, oxidative stress was recognized
as one of the major sources of endogenous DNA damage in bacteria
(Foster et al., 2015). If not properly repaired, such damage could serve
as a significant driver of mutations, with broad implications for bacterial
evolution and the development of antibiotic resistance (Carvajal-Garcia
et al., 2023). Although our study focused on a single strain, previous
analyses have shown substantial variation in the distribution of putative
O₂/H₂O₂ detoxification systems among *Faecalibacterium* strains (Botin
et al., 2023). Future work examining additional strains will be crucial to
determine the extent to which the mechanisms described here are
conserved and to understand the variability in their functional contri-
butions to oxidative stress tolerance.

At the gene expression level, we identified two temporally distinct
induction profiles. The early peak in transcription for rubrerythrin and
reverse rubrerythrin-encoding genes within 5 min of oxidative stress
highlighted the swift activation of detoxification mechanisms, likely
essential for survival in fluctuating oxidative microenvironments, such
as those present during intestinal inflammation. The second profile
included four additional genes – encoding a flavodiiron protein, two
metal transporters, and a precorrin methyltransferrase – whose expres-
sion reached maximal levels after 15 min of H₂O₂ exposure. This in-
dicates that these two gene sets might be under the control of distinct
transcriptional regulators. Consistent with these findings, our previous
study in F. longum L2-6 (Botin et al., 2023) also reported temporally
distinct patterns of gene induction after oxidative stress, highlighting the
existence of a coordinated and hierarchical regulatory network.
Although *Faecalibacterium* species are adapted to a strictly anaerobic
environment, they appear to possess a sophisticated transcriptional
regulatory network capable of rapidly sensing and responding to
oxidative challenges. To date, the regulatory network controlling these
responses remain largely uncharacterized. Nevertheless, they merit
focused investigation (Auger et al., 2022b), as elucidating these net-
works is critical for understanding the molecular mechanisms that un-
derlie bacterial adaptation, survival, and interactions with the host in
the dynamic and oxygen-fluctuating environment of the colon. Oxida-
tive stress strategies have been previously reported in other strict an-
aerobes. For example, *Clostridium* species rely on superoxide reductases,
rubrerythrics, and peroxiredoxins, with regulation mediated by PerR,
OxyR, or σ<sup>B</sup>-like factors (Lotoux et al., 2025). *Bacteroides fragilis* induces
catalase, superoxyde dismutase, rubrerythrin and peroxydase enzymes
under oxidative stress, controlled by OxyR/BmoR regulators (Yekani
et al., 2021). Although the specific enzymes and regulators differ be-
tween taxa, the core adaptive strategy, rapid induction of ROS detoxi-
fication pathways, appears conserved among strict anaerobes, consistent
with our findings in F. duncaniae.

It is important to note that using H₂O₂ to simulate oxidative stress does not fully capture the complexity of the ROS environment in the intestine, which comprises multiple reactive oxygen and nitrogen species produced by host epithelial and immune cells, including hypochlorous acid and peroxynitrite. Moreover, single-strain cultures cannot reproduce interactions within the intestinal microbiota or host-related consequences of oxidative stress, such as disruptions of metabolic processes and iron homeostasis (Wang et al., 2023). Therefore, although our results provided useful insights into bacterial responses to oxidative

stress, further studies using *in vivo* models will be necessary to validate
these findings.

Interestingly, our analyses revealed a marked induction of enzymes
associated with cobalamin biosynthesis following H₂O₂ exposure,
despite the fact that the cobalamin biosynthetic pathway is incomplete
in *Faecalibacterium* spp. (Heinken et al., 2014). In gut and soil micro-
biomes, bacteria actively produce and exchange cobalamin and related
corrinoids, thereby promoting metabolic complementarity and com-
munity stability (Hallberg et al., 2024; Magnúsdóttir et al., 2015;
Sharma et al., 2019). Notably, *Bacteroides thetaiotaomicron* relies on the
surface protein BtuG for efficient cobalamin import and persistence in
the mammalian gut (Wexler et al., 2018). We did not identify any gene
encoding BtuG-like homologs in *Faecalibacterium* genomes, suggesting
that this genus may depend on alternative uptake systems to acquire
community-derived cobalamin precursors, or corrinoids needed to sus-
tain its metabolism.

In this work, we showed that cobalamin supplementation enhances
growth and improves survival under H₂O₂ stress in *F. duncaniae* A2–165.
Our data suggest that cobalamin does not act as a primary first responder
to acute oxidative stress, given its minimal effect on lag phase duration.
Instead, it appears to support metabolic recovery and biosynthesis
during exponential growth following stress, acting as a maintenance and
recovery factor. Unlike our findings, Bircher *et al.* reported no growth-
promoting effect of cobalamin under unstressed conditions (Bircher
et al., 2024). This discrepancy may reflect differences in culture media
and the use of a cobalamin concentration nearly 100-fold lower than in
our study. Notably, the protective effect of cobalamin against H₂O₂ stress
was not reproduced by methionine supplementation, suggesting that its
action extends beyond its role as a methionine synthase cofactor.
Although cobalamin is known to function as an antioxidant in various
cell and animal models (Birch et al., 2009; Chan et al., 2018; Li et al.,
2020), there is limited evidence in bacteria, except for a reported pro-
tective antioxidant role in the aerobic environmental bacterium *Lep-
tospirillum* (Ferrer et al., 2016). Further work will be required in
*Faecalibacterium* to determine whether this effect reflects a direct anti-
oxidant activity of cobalamin or results from downstream modulation of
redox-related metabolic pathways. In addition, the development of ge-
netic tools for *Faecalibacterium* or the use of heterologous systems could
in the future allow direct testing of the causal role of cobalamin in
oxidative stress resistance.

We performed a comprehensive analysis of the membrane fatty acid
composition of strain A2–165 across different growth phases. This
revealed the presence of plasmalogen-type fatty acids, which appear to
be commonly found in strict anaerobic bacteria (Ryan et al., 2023).
Plasmalogens are likely to contribute to membrane stability and fluidity,
providing an additional defense mechanism that helps bacteria cope
with the damaging effects of reactive oxygen species (Braverman and
Moser, 2012; Dean and Lodhi, 2018). Within the gut microbiota, they
may also play a role in microbiota-host interactions at the molecular
level (Ryan et al., 2023). This suggests that plasmalogens produced by
*F. duncaniae* could influence its communication with host intestinal
cells. This role warrants further investigation to elucidate the functional
significance of plasmalogens in both bacterial physiology and
host-microbe interactions. We also showed that exposure to H₂O₂
induced subtle but significant remodeling of FAs, suggesting that dy-
namic membrane adaptation constitutes an additional strategy to
maintain cellular homeostasis under stress. The observed increase in
saturated FA in response to H₂O₂ could result either from passive
oxidative damage or from an active adaptive response. Future studies
analyzing both the oxidation of membrane FA (e.g., lipid peroxidation)
and the expression of FA biosynthesis genes will be necessary to clarify
the underlying mechanism in *F. duncaniae*. Finally, cobalamin mitigates
H₂O₂ stress by stabilizing membrane FA composition in *F. duncaniae*.

Taken together, our findings highlight the role of cobalamin as a
microbial defense molecule, and underscore its potential importance in
enabling strict anaerobes to tolerate transient oxidative stress. Our
results have industrial applications: (i) since cobalamin availability positively influences biomass production in *F. duncaniae*, it could be used to optimize large-scale cultivation in bioreactors; and (ii) due to its antioxidant properties, cobalamin may enhance the viability and functional stability of encapsulated *Faecalibacterium*, as demonstrated for riboflavin-conjugated sodium alginate and glycol chitosan (Khan et al., 2014; Qiu et al., 2024). Beyond industrial relevance, these findings also carry applications for human health. Previous work has shown that cobalamin and related cofactors (corrinoids) can shape the composition, metabolic interactions, and competitive dynamics of human gut microbial communities (Degnan et al., 2014). By mitigating oxidative stress, cobalamin may help maintain the abundance and activity of health-promoting taxa, such as *Faecalibacterium*. This is particularly relevant in conditions characterized by elevated mucosal oxidative stress, including inflammatory bowel disease, metabolic disorders, and post-antibiotic dysbiosis. Our results therefore suggest that increased dietary cobalamin intake, or targeted delivery of cobalamin within microbiome-directed interventions (e.g., next-generation probiotics, synbiotics, or encapsulation matrices), may support the resilience and functional contribution of beneficial gut microbiota under fluctuating oxidative conditions. This opens promising perspectives for leveraging cobalamin as a modulator of gut microbial homeostasis in precision nutrition and microbiome-based therapies.

## 5. Conclusion

This study provides a comprehensive characterization of the mechanisms that enable *F. duncaniae* A2–165 to withstand high levels of H₂O₂, and identifies cobalamin as a key factor enhancing its resilience. We show that this strictly anaerobic bacterium deploys a rapid, coordinated, and multilayered defense against H₂O₂, involving detoxification enzymes, metal homeostasis systems, DNA repair pathways, and dynamic remodeling of membrane FA. Our findings demonstrate that exogenous cobalamin enhances growth, sustains membrane integrity, and significantly improves survival under stress. Beyond advancing our understanding of *Faecalibacterium* physiology, these results reveal cobalamin as a key microbial defense molecule that may help strict anaerobes endure transient oxidative stress within the gut environment. The implications extend to industrial cultivation strategies, where cobalamin supplementation could optimize biomass production and strain stability, as well as to microbiome-targeted interventions, aiming to preserve beneficial taxa in oxidative inflammatory conditions.

## Funding

The research leading to these results has received funding under grant (FAVEPRO) from Carnot Qualiment© (DOI: 10.17180/h5gd-gk88) supported by Agence Nationale de la Recherche.

## CRediT authorship contribution statement

**Christine Péchoux:** Methodology. **Jasmina Vidic:** Writing – review & editing, Methodology. **Alban Amoros:** Methodology. **Florence Dubois-Brissonnet:** Methodology, Formal analysis. **Beney Laurent:** Funding acquisition, Conceptualization. **Lydie Oliveira Correia:** Methodology. **Sébastien Dupont:** Writing – review & editing, Methodology. **Simon Ménard:** Methodology, Formal analysis, Data curation. **Rime Chebbo:** Methodology. **Bonastre Oliete:** Project administration, Funding acquisition, Conceptualization. **Sandrine Auger:** Writing – review & editing, Writing – original draft, Validation, Supervision, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Data curation, Conceptualization. **Jean-Marc Chatel:** Writing – review & editing, Funding acquisition. **Maria Alejandra de Angel Fontalvo:** Methodology, Formal analysis.

## Declaration of Competing Interest

The authors declare no conflict of interest.

## Acknowledgments

We are grateful to Melisanda Caillé for technical help, and Elena Bidnenko, PhD and Alain Guillot for scientific discussion. This work has benefited from the facilities and expertise of the Plateforme Analyse Protéomique de Paris Sud-Ouest (PAPPSO) which is supported by INRAE, the Ile-de-France regional council, IBiSA, and CNRS, and the Plateforme de Microscopie et Imagerie (MIMA2).

## Appendix A. Supporting information

Supplementary data associated with this article can be found in the online version at doi:10.1016/j.micres.2026.128455.

## Data availability

Data will be made available on request.

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