<figure><img src="image_1.png" /></figure>

# Improved organic matter biodegradation through pulsed H₂ injections during *in situ* biomethanation

M. Mahieux<sup>a,b</sup>, Q. Aemig<sup>b</sup>, C. Richard<sup>b</sup>, J-P. Delgenès<sup>a</sup>, M. Juge<sup>b</sup>, E. Trably<sup>a</sup>, R. Escudié<sup>a,*</sup>

<figure><img src="image_2.png" /></figure>

## HIGHLIGHTS

* Improvement of the organic substrate biodegradation was associated to H₂ injections.

* Non-dominant hydrogenotrophic methanogens became predominant with H₂ injections.

* CH₄ production was similar regardless of initial microbial activity levels.

* CH₄ production from organic matter increased of at least 626 NmL with H₂ injections.

## GRAPHICAL ABSTRACT

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

## ARTICLE INFO

### Keywords:

Biogas upgrading

Acidogenesis

Anaerobic Digestion

Methanogens

## ABSTRACT

During *in situ* biomethanation, microbial communities can convert complex Organic Matter (OM) and H₂ into CH₄. OM biodegradation was compared between Anaerobic Digestion (AD) and *in situ* biomethanation, in semi-continuous processes, using two inocula from the digester (D) and the post-digester (PoD) of an AD plant. The impact of H₂ on OM degradation was assessed using a fractionation method. Operational parameters included 20 days of hydraulic retention time and 1.5 gVs.L⁻¹.d⁻¹ of organic loading rate. During *in situ* biomethanation, 485 NmL of H₂ were injected for each feeding (3 times a week). Maximum organic COD removal was 0.6 gCOD in AD control and at least 1.6 gCOD for *in situ* biomethanation. Therefore, COD removal was 2.5 times higher with H₂ injections. These results bring out the potential of H₂ injections during AD, not only for CO₂ consumption but also for better OM degradation.

## 1. Introduction

The biomethane production industry, relying on the treatment of organic waste by Anaerobic Digestion (AD), plays a pivotal role in the global effort to decarbonize the energetic mix (REPowerEU, 2022). This current interest is evident, with biomethane production expected to expand worldwide from 3.5 to over 11 billion cubic meters between

2018 and 2022. Several project developments are driving this industry, mostly located in Europe and North America (currently representing altogether more than 90 % of the global biomethane production), with a significant anticipated contribution expected from Brazil and India between 2022 and 2026 (IEA, 2023).

Subsequently, the deployment and optimization of AD are key factors in unlocking the potential of organic matter as an energy source. AD

<footnote><sup>a</sup> INRAE, Univ. Montpellier, LBE, 102 Avenue des étangs, F-11100 Narbonne, France</footnote>

<footnote><sup>b</sup> ENGIE, Lab CRIGEN, 4 Rue Joséphine Baker, 93240 Stains, France</footnote>

<footnote>* Corresponding author.
E-mail address: renaud.escudie@inrae.fr (R. Escudié).</footnote>
is considered a mature industrial process. Conventional optimization strategies are based on both process management itself and applied treatment to substrates. In addition, the potential for using H₂ in *in situ* biomethanation has been explored, allowing the purification of biogas through the activity of hydrogenotrophic archaea, either in pure or mixed culture, within a dedicated reactor (Angelidaki et al., 2018). However, as hydrogenotrophic archaea are also present in the microbial community of the AD process, an alternative approach involves direct injection of H₂ into an AD process, leading to what is called *in situ* biomethanation (Rafrafi et al., 2021). For instance, the previous work of Luo et al. (2012) demonstrated that H₂ supplementation in AD reactors could enhance CH₄ production because H₂ and CO₂ are utilized by the microbial community to produce CH₄. Current H₂ production is mostly based on H₂ steam reforming, derived from the oil industry (Dincer and Acar, 2015). However, several technologies are considered to make H₂ production sustainable, such as water electrolysis coupled with intermittent renewable electricity production (Power-to-Gas processes) (Thema et al., 2019) and biological processes for H₂ production (Dincer and Acar, 2015).

H₂ is a well-known intermediate molecule for CH₄ production during the methanogenesis step of AD (Zhu et al., 2020). The introduction of exogenous H₂ during such a process was shown to negatively impact methanogenesis. Indeed, the accumulation of Volatile Fatty Acids (VFAs) due to both homoacetogenic activity stimulation and acetoclastic methanogenesis inhibition in the presence of high H₂ partial pressure has already been reported in the literature (Agneessens et al., 2018; Luo and Angelidaki, 2013). Conversely, no consensus is currently reached in the scientific community regarding the impact of exogenous H₂ addition on the organic matter degradation steps of AD (e.g., hydrolysis, acido-genesis, acetogenesis). Indeed, it was previously demonstrated by Cazier et al., (2019) that H₂, at a partial pressure of 1 bar, could inhibit hy-drolysis. However, Bassani et al., (2015) and Treu et al., (2018) both suggested that the development of syntrophic interactions between bacteria and archaea was promoted by H₂ injection during AD. These two studies suggested that the organic matter degradation process and, therefore, bacterial activities could be positively impacted by H₂ addi-tion during *in situ* biomethanation. The development of this positive effect during *in situ* biomethanation was not exposed in the literature. In addition, the possible impact of H₂ on bacterial activity could be vari-able, depending on the initial activity of the community. Indeed, several authors reported that the microbial composition of one inocula was impacting its adaptation capacity to H₂ injections. (Agneessens et al., 2018; Braga Nan et al., 2020).

For that purpose, it appeared to be necessary to evaluate the impact of H₂ on communities with close composition but high and low levels of hydrolytic activity. As a result, two inocula from the main and post-digester of the same AD unit were selected for this experiment. Post-digesters are conceived to receive digestate from the main digester, to collect the remaining methane from that digestate, before its storage in dedicated tanks (Boe et al., 2009). Therefore, in conventional AD units, main digesters are expected to exhibit high organic matter degradation activity, as they received complex organic matter as feeding (e.g., waste from agricultural and food industries) while post-digesters are expected to expose less active microbial consortium. Consequently, this study aimed to evaluate the effect of H₂ on hydrolytic, acidogenic, and ace-togenic activities, but also the response of microbial communities to complex organic feedstock input under high H₂ concentrations in the gas phase.

## 2. Materials and methods

### 2.1. Inocula and feedstock preparations

Two inocula corresponding to the main and post-digester of the same AD units were used at a concentration of 15 gV<sub>S</sub>.L<sup>-1</sup> during that exper-iment. They were sampled on the AD unit less than 5 days before that

experiment. These inocula were selected due to their rapid response toward H₂ injection, which was observed in previous work (Mahieux et al., 2024). Both were sieved through a 20-mesh sieve to remove a part of their residual organic matter and decrease their endogenous methane production.

The organic feeding consisted of a mixture of 2 % liquid pig manure, 29 % freeze-dried solid cow manure, 28 % rye silage (which was mortar ground), and 42 % food waste soup, all expressed in terms of Volatile Solid (VS). The food waste soup was prepared following a protocol already described elsewhere (Noguer et al., 2022). All feedstocks were used less than 7 days after their collection. The food waste soup was prepared the day of the mix preparation. The mixture was prepared and then diluted with distilled water to obtain a 30 gV<sub>S</sub>.L<sup>-1</sup> feeding solution and divided into aliquots before being stored at -20 °C.

The Total Solids (TS) and Volatile Solids (VS) contents of sieved inocula and feedstocks were measured as described in APHA standard methods (APHA, 2017). The correction of the TS and VS content of rye silage was performed following the method described in the work of Van Vlierberghe et al. (2022) and using the Porter and Murray equation. The biodegradable Chemical Oxygen Demand (COD) of the substrate was 0.024 gCOD.g<sup>-1</sup>. This COD content was evaluated using near-infrared spectroscopy, as already described elsewhere (Zennaro et al., 2022).

### 2.2. Operational setup

The culture medium was composed of 20 mL of phosphate buffer (0.5 M, pH 7.12 adjusted using NaOH), 2 mL of an oligo-element solu-tion, inoculum at a concentration of 15 gV<sub>S</sub>.L<sup>-1</sup>, 0.7 gV<sub>S</sub> of feedstock mixture. The medium was filled to 200 mL with distilled water. The composition of buffer and oligo-element solutions was similar to those used by Braga Nan et al. (2020).

For the experiment, several modified 1 L SchottFlask® with a working volume of 200 mL were used. These bottles were flushed for 15 min with nitrogen at the beginning of the experiment to ensure anaer-obic conditions. Feeding was operated for 48 days in semi-continuous conditions by setting a Hydraulic Retention Time (HRT) of 20 days and an organic loading rate of 1.5 gV<sub>S</sub>.L<sup>-1</sup>.d<sup>-1</sup> for the feedstock mix feeding.

Two conditions were assessed in this study: the Anaerobic Digestion condition (AD control condition), which solely received feedstock mixture, and the *in situ* biomethanation condition (*in situ* condition) (see supplementary material). In the latter, after one start-up week under similar feeding conditions as AD control, H₂ was additionally introduced rapidly from 1 to 1.45 bar. This gas injection corresponded to an addi-tion of 485 NmL of H₂ after each organic feeding. Both gas and organic feeding were operated three times per week. Organic feeding was con-ducted by adding 23.3 g of feedstock mix (equivalent to 0.7 gV<sub>S</sub>) and removing the same amount of culture media. The AD and *in situ* con-ditions were evaluated in triplicate and quadruplicate, respectively. Quadruplicates were chosen for the *in situ* condition to ensure a robust evaluation of this condition, as H₂ injection could trigger more vari-ability for CH₄ and VFA production. Both conditions were incubated on a mechanical shaking table to ensure continuous stirring in a mesophilic chamber at 35 °C during the entire experiment.

### 2.3. Analytical methods

Physical-chemical analyses were performed on the Bio2E platform (Bio2E, 2018). Gas composition was analyzed three times per week before and after feeding by gas chromatography using GC Perkin Elmer model Clarus 580 equipped with a Thermal Conductivity Detector (TCD detector) as previously described by Mahieux et al. (2024). Before gas analysis, SchottFlask® inner gas pressure was measured using a manometer Keller LEO 2 (KELLER AG, Winterthur, Switzerland).

Once a week, a liquid medium was sampled to determine the con-centrations of VFA and ammonium. For these analyses, liquid samples
underwent centrifugation for 15 min at 12,100 g, and the resulting supernatant was filtered using a 0.22 µm nylon filter (Fisherbrand®). Both the centrifugation pellet and the filtered supernatant were stored at -20 °C before their analysis. VFA concentration measurements were performed by gas chromatography using a Clarus 580 gas chromatograph (GC, Perkin Elmer), as reported by Noguer et al. (2022). The gas carrier was N₂ at 6 mL.min⁻¹. The GC was equipped with an Alltech-FFAP EC™1000 column coupled to a Flame Ionization Detector (FID) working at 280 °C. An Elite-FFAP cross-bond® carbowax® capillary column (15 m length, 0.53 mm diameter) was installed on the GC. Ammonium concentration was measured by automated spectrophotometric titration using Gallery™ instrument (Thermofischer Scientific) according to the manufacturer's instructions.

## 2.4. Fractionation method of organic matter

The fractionation method was conducted following the procedures outlined in Jimenez et al. (2015). The feedstock mixture used to supply the reactors and the digestate at the end of the experiment for the four conditions were analyzed. Initially, samples were centrifuged for 20 min at 18,600 g. The retrieved supernatant was assigned as the Dissolved Organic Matter fraction (DOM). Following centrifugation, TS and VS contents of sample pellets were determined before and after freeze-drying and grinding (1 mm). Then, 1 g of the resulting sample pellet was first subjected to automated sequential extraction using Dionex™ ASE™ 350. Different fractions of the sample were extracted sequentially using a 10 mM CaCl₂ solution, a 10 mM NaOH solution, and a 100 mM NaOH solution to determine the extractable Soluble from Particulate Organic Matter (SPOM), the Readily Extractable Organic Matter (REOM), and the Slowly Extractable Organic Matter (SEOM), respectively. Finally, a 72 % (v/v) H₂SO₄ solution was used as an extractant to manually determine the Poorly Extractable Organic Matter (PEOM). The remaining organic matter corresponded to the Non-Extractable Organic Matter (NEOM).

Chemical Oxygen Demand (COD) was measured using 2 mL of each fraction solution using Aqualytic® Vario COD kits (0–1500 mg O₂·L⁻¹) based on ultraviolet (UV) absorbance, which was read on a MultiDirect spectrophotometer from Aqualytic®. Total COD (mg O₂·gTS⁻¹) was estimated in duplicate. For that purpose, 0.25 g of freeze-dried and ground (1 mm) raw samples were dissolved in 10 mL of a 99 % (v/v) H₂SO₄ solution for 24 h. Those solutions were diluted in demineralized water before COD measurement. Fractions of COD (mgO₂·L⁻¹) were assessed with 2 mL of each extracted solution. The difference in total COD observed between the raw substrate and all assessed conditions after two HRTs corresponded to the Degraded Organic Matter (DEGOM). This fraction allowed the evaluation of the portion of organic matter efficiently degraded through AD in all conditions.

## 2.5. Microbial community analysis

### 2.5.1. Sequencing strategy

Reactors were sampled for microbial analysis on the inocula (Week 0), at the beginning of H₂ injection in the in situ condition (Week 1), after one HRT (Week 3), and after two HRT (Week 6) during the semi-continuous process for both conditions.

The DNA extraction and quality control, as well as the two different PCRs that were conducted to analyze separately bacteria and archaea that were present in samples, were operated on following the previous work of Braga Nan et al. (2020). The sequencing reaction was performed with an Illumina sequencer (2 × 300 pb paired-end run) at the GenoToul Platform (Toulouse, France). Sequencing results were analyzed with bioinformatic tools. Mothur version 1.48.0 was used for sequence cleaning, assembly and control. Sequence matching and taxonomic assignment were performed on SILVA version 132. The Operational Taxonomic Units (OTUs) were created considering a 3 % dissimilarity level for both archaeal communities and bacterial communities. Results

were registered in the Sequence Read Archive (https://www.ncbi.nlm.nih.gov/sra) under the BioProject accession number PRJNA956344, with SRA accession numbers SAMN34423353 to SAMN34423370 for bacteria and SAMN34423253 to SAMN34423270 for archaea.

### 2.5.2. Microbial quantification

Bacteria and Archaea quantification was assessed using quantitative Polymerase Chain Reaction (qPCR). A BioRad CFX96 Real-Time Systems C1000 Touch Thermal Cycler (Bio-Rad Laboratories, USA) was used to perform qPCR programs. Primers used for bacterial and archaeal quantification, as well as the qPCR protocols, were previously described by Mahieux et al. (2024).

### 2.5.3. Data treatment for visualization of microbial communities

Graphical visualizations were obtained using R software version 4.1.3. Phyloseq, microbiome, and Fantaxtic R packages were used to obtain visual representations of the microbial communities (McMurdie and Holmes, 2013; Teunisse, 2022).

## 2.6. Estimation of the methane production rate associated with organic matter degradation

All gas volumes were calculated based on normalized data (NmL), considering atmospheric pressure and a temperature of 0 °C. The mean total CH₄ production rate (CH₄ total, expressed in NmL CH₄·L⁻¹·d⁻¹) was calculated for each week of the experiment, using Eq (1), where CH₄(i) represents the total CH₄ production in the assessed condition at the time i (ti) of the experiment.

$$CH_4\text{total} = [CH_4(i + 1) - CH_4(i)]/[t_{i+1} - t_i] \quad (1)$$

Since no VFA accumulation was observed during the experiment, it was assumed that all the injected hydrogen was converted into CH₄. Subsequently, the mean CH₄ production rate associated with H₂ consumption (CH₄ biomethanation, expressed in NmL CH₄·L⁻¹·d⁻¹) was calculated as described in Eq. (2), where H₂(i) represents the total H₂ consumption observed at tᵢ, taking into account the stoichiometric reaction between CO₂ and H₂ to form CH₄.

$$CH_4\text{biomethanation} = [H_2(i + 1)/4 - H_2(i)/4]/[t_{i+1} - t_i] \quad (2)$$

Finally, the CH₄ production associated with organic matter degradation (CH₄ biodegradation expressed in NmL CH₄·L⁻¹·d⁻¹) in the in situ reactors was determined by retrieving from the total CH₄ production the portion of the CH₄ produced through biomethanation at each time of the experiment, as shown in Eq (3).

$$CH_4\text{biodegradation}(i) = CH_4\text{total}(i) - CH_4\text{biomethanation}(i) \quad (3)$$

A COD mass balance was performed for each week of the experiment and allowed to confirm the robustness of the obtained results (see supplementary materials).

# 3. Results and discussions

## 3.1. Assessment of process overall performances for both inocula

To assess the effect of adding H₂ on the degradation of organic matter, experiments were conducted in semi-continuous feeding mode in in situ biomethanation conditions and in AD control conditions (without H₂ injections). Furthermore, to characterize the impact of H₂ on bacterial activity, two different inocula were used: one inoculum, expected to exhibit high bacterial activity, was obtained from the main digester (referred to as D), and the other, expected to expose lower bacterial activity, was sampled from the post-digester (referred to as PoD) of the same AD unit. The performances of in situ biomethanation and AD reactors were evaluated through the production rates of CH₄ and
CO₂, as well as the consumption rates of H₂. These rates were averaged over each week of the experiment, considering that reactor feeding occurred three times per week. Moreover, three experimental steps were considered: week 0, without H₂ injection, so-called the start-up of the experiment; the first HRT (weeks 1 to 3) as an adaptation phase for *in situ* biomethanation; and the second HRT (weeks 4 to 6) as a steady state.

The ratio between the injected H₂ and the produced CO₂ is considered an important parameter for the stability of *in situ* biomethanation processes (Angelidaki et al., 2018). For the *in situ* biomethanation reactors using inoculum D, the average apparent H₂:CO₂ ratio observed after each H₂ injection was 3.1 ± 0.6:1 H₂:CO₂ mol:mol while the real H₂:CO₂ ratio, considering the total H₂ consumed and the total CO₂ production, was 2.7 ± 0.1:1 H₂:CO₂ mol:mol. Similar ratios were observed in the *in situ* reactor using the inoculum sampled from the post-digester (inoculum PoD). In comparison with the literature, this ratio was lower than the stoichiometric 4:1 H₂:CO₂ molar ratio of the methanation reaction, which was considered optimal for *in situ* biomethanation (Wahid et al., 2019). These low ratios were chosen to avoid CO₂ depletion during *in situ* biomethanation. This setup limited the strong impact of H₂ injection on pH, which could have led to VFA accumulation and process inhibition. Indeed, acetate accumulation is more likely to occur during *in situ* biomethanation in response to CO₂ depletion in a medium, as previously shown by Agneessens et al. (2018) work. Specifically, these authors showed that CO₂ depletion from the gas phase, and the resulting increase in pH could lead to methanogenesis inhibition and homoacetogenesis stimulation, leading to an increase of acetate concentrations in the media.

The total CH₄ production performances were described, (Fig. 1) while separating the part of the CH₄ coming from H₂ or OM consumption. The inocula obtained from the main digester (inoculum D) and the post-digester (inoculum PoD) of the same AD unit exhibited different initial behaviors. For the Week 0, during which neither the AD control nor *in situ* condition received H₂, inoculum D showed a high CH₄ production rate, while for the inoculum PoD, the production was very limited (606 ± 41 vs 137 ± 4 NmL CH₄·L⁻¹·d⁻¹ for inocula D and PoD, respectively). A part of the observed production could be associated with the remaining organic matter degradation using inoculum D, as reactors using that inoculum displayed a CH₄ production rate almost five times higher than those using the inoculum PoD. In addition, the CH₄ production rate for inoculum D was higher than the theoretical CH₄ production rate that could have been obtained with the fed substrate (413 NmL CH₄·L⁻¹·d⁻¹ expected from the substrate, considering its COD content), reinforcing the association of the additional CH₄ production with endogenous organic matter conversion. These results also suggest higher initial microbial activity in inoculum D than in inoculum PoD, considering their respective initial CH₄ production performances during

week 0. In addition, this first week of organic matter feeding (without H₂ addition in the *in situ* reactor) was associated with large VFA production regarding inoculum PoD, reaching 2.5 ± 0.2 g·L⁻¹ of acetate and 0.6 ± 0.3 g·L⁻¹ of propionate in all reactors inoculated with PoD at the beginning of week 1. This rapid accumulation of VFA suggests that the organic matter feeding induced an important environmental stress on the microbial community of the inoculum at the beginning of the experiment. Conversely, no significant VFA production was observed for reactors corresponding to inoculum D at the beginning of week 1.

Considering first the reactors inoculated with PoD, an improvement in the total CH₄ production rate was observed during the first part of the experiment under AD control conditions (from 116 ± 14 during Week 0 to 442 ± 109 NmL CH₄·L⁻¹·d⁻¹ during Week 2) as well as in the *in situ* condition (from 137 ± 4 during Week 0 to 658 ± 49 NmL CH₄·L⁻¹·d⁻¹ during Week 2). These results seemed to confirm an enhancement in the microbial consortia activity associated with that inoculum, both with and without H₂ addition. Conversely, using inoculum D, a stable total CH₄ production rate was observed in both AD control and *in situ* conditions from Week 1 to Week 3, which corresponds to the first HRT with H₂ addition in the *in situ* reactor (721 ± 13 and 379 ± 26 NmL CH₄·L⁻¹·d⁻¹ in average from Week 1 to 3, for the *in situ* condition and the AD control, respectively). The rapid stabilization of biomethanation was attributed to the low H₂:CO₂ ratio and the use of easily degradable substrates, such as food waste and rye silage (42 % and 28 % of the VS concentration in the feeding solution, respectively).

During the second HRT phase of the experiment, from Week 4 to Week 6, the CH₄ production rate remained stable in both AD control and *in situ* conditions (Fig. 1). This stability allowed considering this period as the steady state of the semi-continuous process. Limited VFA production was observed (Fig. 2) for that period. During this period, the CH₄ and VFA production patterns of inoculum D and PoD were almost identical. This similarity indicates that, after an initial gain of activity for inoculum PoD during the first HRT, both inocula exhibited comparable performance in AD and *in situ* biomethanation. Similar performances in the AD control were expected for these two inocula, as they were sampled on the same AD plant (close initial similar microbial communities) and fed similarly. Such similarity was not expected in the *in situ* performances of both inocula. Considering that H₂ injection was previously reported as triggering VFA production for non-adapted inocula (Agneessens et al., 2017), it could have been expected that H₂ injection had negatively impacted CH₄ production performance of both inocula and especially of inoculum PoD, which initially exhibited lower efficiency in CH₄ production compared to inoculum D.

Regarding the end of the process, propionic acid production, reaching a final concentration of 1 g·L⁻¹, was observed in all conditions, Fig. 2. This accumulation of VFA coincided with a decrease in

<figure><img src="image_4.png" /><figcaption>**Fig. 1.** Total methane production rate during the experiment (NmL CH₄·d⁻¹). All conditions were assessed in triplicates. D: AD Digestate of Digester as inoculum in anaerobic digestion process; D *in situ*: Digestate of Digester as inoculum in *in situ* biomethanation process; PoD AD: Digestate of Post-digester as inoculum in anaerobic digestion process; PoD *in situ*: Digestate of Post-digester as inoculum in *in situ* biomethanation process.</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Fig. 2.** VFA concentrations during the experiment (g.L⁻¹). All conditions were assessed in triplicates. D AD: Digestate of Digester as inoculum in anaerobic digestion process; D *in situ*: Digestate of Digester as inoculum in *in situ* biomethanation process; PoD AD: Digestate of Post-digester as inoculum in anaerobic digestion process; PoD *in situ*: Digestate of Post-digester as inoculum in *in situ* biomethanation process.</figcaption></figure>

ammoniacal nitrogen concentration and a decline in pH in all conditions (Table 1). Initially, the inocula exhibited high N-NH₄⁺ concentrations (i. e., 928 ± 19 and 1,270 ± 36 mg N-NH₄⁺·L⁻¹ for inoculum D and PoD, respectively). Subsequently, these inocula were considered adapted to relatively high free ammonia concentrations. Therefore, the decrease in N-NH₄⁺ concentration could have altered the media buffering capacity, resulting in a higher impact of VFA concentration on pH in this experiment (Capson-Tojo et al., 2020).

During the present experiment, no residual H₂ was detected in the gas phase, supporting that all added H₂ during one feeding time was consumed before the next feeding. Moreover, for both inocula, the VFA production observed after the beginning of H₂ injections in the *in situ*

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

Monitoring of pH and N-NH₃⁺ (mg.L⁻¹). All conditions were assessed in triplicates. D AD: Digestate of Digester as inoculum in anaerobic digestion process; D *in situ*: Digestate of Digester as inoculum in *in situ* biomethanation process; PoD AD: Digestate of Post-digester as inoculum in anaerobic digestion process; PoD *in situ*: Digestate of Post-digester as inoculum in *in situ* biomethanation process.</caption><thead><tr><th rowspan="2"></th><th colspan="2">D AD</th><th colspan="2">D in situ</th><th colspan="2">PoD AD</th><th colspan="2">PoD in situ</th></tr><tr><th>pH</th><th>N-NH₄⁺ (mg. L⁻¹)</th><th>pH</th><th>N-NH₄⁺ (mg. L⁻¹)</th><th>pH</th><th>N-NH₄⁺ (mg. L⁻¹)</th><th>pH</th><th>N-NH₄⁺ (mg. L⁻¹)</th></tr></thead><tbody><tr><td>T0</td><td>7.8 ± 0.1</td><td>928 ± 19</td><td>7.8 ± 0.0</td><td>928 ± 19</td><td>7.8 ± 0.1</td><td>1270 ± 36</td><td>7.8 ± 0.1</td><td>1270 ± 36</td></tr><tr><td>Week 0</td><td>7.5 ± 0.3</td><td>1078 ± 114</td><td>7.5 ± 0.3</td><td>1011 ± 18</td><td>7.6 ± 0.3</td><td>1185 ± 84</td><td>7.5 ± 0.3</td><td>1113 ± 126</td></tr><tr><td>Week 1</td><td>7.3 ± 0.2</td><td>516 ± 16</td><td>7.4 ± 0.1</td><td>518 ± 45</td><td>7.1 ± 0.2</td><td>574 ± 26</td><td>7.2 ± 0.1</td><td>580 ± 20</td></tr><tr><td>Week 2</td><td>7.1 ± 0.2</td><td>572 ± 9</td><td>7.4 ± 0.1</td><td>568 ± 18</td><td>7.0 ± 0.1</td><td>653 ± 40</td><td>7.3 ± 0.2</td><td>652 ± 41</td></tr><tr><td>Week 3</td><td>7.0 ± 0.1</td><td>379 ± 7</td><td>7.4 ± 0.1</td><td>347 ± 23</td><td>7.0 ± 0.1</td><td>477 ± 19</td><td>7.5 ± 0.2</td><td>428 ± 28</td></tr><tr><td>Week 4</td><td>6.9 ± 0.1</td><td>341 ± 16</td><td>7.4 ± 0.1</td><td>321 ± 14</td><td>6.9 ± 0.1</td><td>354 ± 26</td><td>7.3 ± 0.2</td><td>367 ± 27</td></tr><tr><td>Week 5</td><td>6.8 ± 0.1</td><td>314 ± 17</td><td>7.2 ± 0.2</td><td>254 ± 17</td><td>6.7 ± 0.2</td><td>325 ± 13</td><td>7.1 ± 0.2</td><td>293 ± 30</td></tr><tr><td>Week 6</td><td>6.6 ± 0.1</td><td>264 ± 11</td><td>7.0 ± 0.1</td><td>213 ± 20</td><td>6.5 ± 0.2</td><td>286 ± 7</td><td>6.9 ± 0.1</td><td>225 ± 53</td></tr></tbody></table>

conditions was similar to the one observed in AD controls (Fig. 2). Therefore, the VFA production was not associated with H₂ injections, and it was considered that all the consumed H₂ was converted into CH₄. As a result, the remaining CH₄ production was associated with organic matter conversion.

## 3.2. Influence of hydrogen on the organic matter conversion in methane

As detailed in Section 3.6, a mass balance was performed, taking into account that CH₄ was produced from both the injected H₂ (referred to as CH₄ biomethanation) and the degradation of organic matter (referred to as CH₄ biodegradation). Both CH₄ biomethanation and CH₄ biodegradation were depicted in Fig. 1 for all conditions.

During the first weeks of operation, the results suggested that the inocula was adapting to the conditions of *in situ* biomethanation and anaerobic digestion. For the PoD inoculum during Week 1, the CH₄ production rate associated with organic matter degradation was higher in the AD condition compared to the *in situ* condition (285 ± 32 vs 144 ± 28 NmL CH₄·L⁻¹·d⁻¹ for AD control and *in situ* conditions, respectively). However, this difference rapidly disappeared, as observed during Week 3 (442 ± 109 vs 437 ± 50 NmL CH₄·L⁻¹·d⁻¹, respectively, for AD control and *in situ* conditions, using the inoculum PoD). The capacity of both conditions to achieve similar methane production performances suggests that pulsed H₂ addition did not have a significant negative impact on organic matter degradation activity during this transitional phase. A similar pattern was also noted for reactors utilizing inoculum D. These results highlighted that, irrespective of the initial biodegradation activity of the consortia, pulsed H₂ injection did not inhibit organic matter degradation and even enabled a gain in biodegradation activity with the PoD inoculum. H₂ is a known inhibitor of various metabolic reaction chains that occur during the acidogenesis step (Bundhoo and Mohee, 2016) and has also been identified as an inhibitor of hydrolysis in specific conditions (Cazier et al., 2019) under high H₂ partial pressure (>0.5 bar). However, the present experiment showed that microbial consortia initially not adapted to H₂ injection could perform these activities without a significant negative impact of H₂. These results are consistent with the VFA consumption of the PoD inoculum in Fig. 2, where the VFAs produced during Week 0 (acclimation week, without H₂ addition) were rapidly consumed in the subsequent weeks, even with pulsed injections of H₂.

Remarkably, the degradation of organic matter was enhanced under *in situ* conditions when the inocula were acclimated to the operational conditions. For instance, at the beginning of the second HRT (Week 3),
with H₂ injection for the *in situ* condition, it was observed that CH₄ resulting from organic matter degradation was higher than in the AD control, regardless of the inocula used (403 ± 13 vs 522 ± 19 NmL CH₄·L⁻¹·d⁻¹ in AD control vs *in situ* condition, considering inoculum D and 380 ± 57 vs 489 ± 22 NmL CH₄·L⁻¹·d⁻¹ in AD control vs *in situ* condition, considering inoculum PoD). This improvement persisted until the end of the second HRT (average production from Week 3 to 6 of 349 ± 75 vs 523 ± 65 NmL CH₄·L⁻¹·d⁻¹ in AD control vs *in situ* condition, considering inoculum D, and 333 ± 64 vs 490 ± 47 NmL CH₄·L⁻¹·d⁻¹ in AD control vs *in situ* condition, considering inoculum PoD), suggesting that H₂, instead of inhibiting the first steps of AD, positively impacted the organic matter degradation during *in situ* biomethanation.

The improvement of CH₄ production rate associated with organic matter degradation could be linked to the increase in pH due to H₂ addition, as indicated in Table 1 (7.0 ± 0.1 vs 7.4 ± 0.1 in AD control vs *in situ* condition for inoculum D, and 7.0 ± 0.1 vs 7.5 ± 0.2 in AD control vs *in situ* condition for inoculum PoD during Week 3, which corresponds to the end of HRT 1). Indeed, the used substrate contained food waste and energy cover crops, potentially explaining the decrease in pH in AD control reactors during the experiment (from 7.0 ± 0.1 to 6.6 ± 0.1 and from 7.0 ± 0.1 to 6.5 ± 0.2, from Week 3 to Week 6, for AD control reactors using inoculum D and PoD during the second HRT, respectively). In contrast, the pH remained approximately 0.5 points higher in *in situ* conditions all throughout the experiments. As VFA concentrations were similar in all conditions, the pH difference between the AD and *in situ* reactors was associated with the difference in CO₂ content in the biogas for these two conditions (45 % vs 85 % of CO₂ retrieved in the gas phase on average for both inocula in the AD control vs *in situ* conditions, respectively, at the end of the experiment; see *supplementary materials*). Indeed, due to the injection of H₂, additional consumption of dissolved CO₂ occurred through the methanation reaction. As a result, the equilibrium between dissolved HCO₃⁻ and aqueous CO₂ led to the consumption of H⁺ proton, promoting an increase in pH in the *in situ* reactors compared to the AD control. As a result, higher pH conditions could have increased microbial biodegradation activity, leading to an improvement in associated CH₄ production performances (Díaz et al., 2020; Luo and Angelidaki, 2013).

### 3.3. Enhancement of organic matter degradation and accessibility by hydrogen injection

Methane production rates clearly showed an improvement in the degradation of organic matter during the *in situ* biomethanation process, regardless of the inoculum used. To reinforce this observation and further explore the biodegradation of the organic matter in the *in situ* and AD control reactors, the fractionation method proposed by Jimenez et al. (2015) was employed. This method was selected as it had previously been used to efficiently characterize AD hydrolysis (Jimenez et al., 2020) and simulate AD digestate bioaccessibility (Fernández-Domínguez et al., 2021) through the six fractions characterized by different accessibility of the Organic Matter (OM) (Dissolved OM (DOM), Soluble Particular OM (SPOM), Readily Extractible OM (REOM), Slowly Extractible OM (SEOM), Poorly Extractible OM (PEOM), Non Extractible OM (NEOM)), ordered from the most to the less accessible fraction. The difference in the total COD observed between the raw substrate and all assessed conditions after two HRT corresponded to the Degraded Organic Matter (DEGOM), as detailed in Section 3.4. This fraction allowed the evaluation of the portion of organic matter efficiently degraded through AD in all conditions. Fig. 3 presents the results of the fractionation analyses for the initial substrate and the digestate at the end of the experiments for all conditions (AD control and *in situ* biomethanation reactors inoculated with D and PoD).

Considering the substrate, the fractionation analysis revealed that 48 % of it was accessible, considering DOM, SPOM, SEOM, REOM, and PEOM as accessible fractions. This composition is consistent with the substrate used in this study, which was composed of solid and liquid

<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Bioaccessibility and conversion of the organic matter between the beginning and the end of the experiment. Fractions were ordered from down to top part of the barplot by increasing bioaccessibility level. DEGOM Degraded organic matter; DOM Dissolved organic matter; SPOM soluble particular organic matter; REOM readily extractable organic matter; SEOM slowly extractable organic matter; PEOM poorly extractable organic matter; NEOM non extractable organic matter. D AD:Digestate of Digester as inoculum in anaerobic digestion process; D *in situ*: Digestate of Digester as inoculum in *in situ* biomethanation process; PoD AD: Digestate of Post-digester as inoculum in anaerobic digestion process; PoD *in situ*: Digestate of Post-digester as inoculum in *in situ* biomethanation process.</figcaption></figure>

manure, energy covered crop silage, and food waste soup. Consistently, Fernández-Domínguez et al. (2021) reported an accessible portion of 46 % for a mixture of pig slurry and biowaste, which is close to the used substrate mixture.

Considering the degradation of the organic matter in all assessed conditions, Fig. 3 clearly shows that, regardless of the initial inoculum, the addition of H₂ significantly increases the DEGOM fraction. This result indicates that more organic matter was degraded under *in situ* biomethanation conditions compared to the AD control without external H₂. The distribution of organic matter fractions was consequently influenced by the addition of H₂, suggesting an increase in the hydrolysis of organic matter and in DEGOM in the case of *in situ* biomethanation.

Moreover, this influence on the degradation of organic matter was also observed with the other fractions. Indeed, while the SEOM and REOM fractions remained similar in the raw substrate and the digested media at the end of the experiment for all conditions, differences were observed for the DOM, SEOM, PEOM, and NEOM fractions.

The DOM fraction is considered to contain the part of the biomass that is already dissolved in water. This fraction was reduced by 10 % in all conditions in comparison with the raw substrate, except for the AD control condition using PoD as inoculum. For the latter, the DOM fraction was close to that of the substrate (29 % and 31 % for the raw substrate and the AD control, respectively). The DOM fraction has been reported to be deeply impacted by hydrolysis and organic matter solubilization phenomena during anaerobic digestion (Jimenez et al., 2020). As a result, the DOM could have been impacted by different microbial activities during the studied processes (Fernández-Domínguez et al., 2023). Indeed, solubilization of organic matter primarily extracted in other fractions (such as PEOM and SEOM) or microbial growth and its associated metabolite production could both have a strong impact on the
DOM fraction. Therefore, no clear links between the evolution of the DOM fraction and the improvement of organic matter degradation through H₂ could be made regarding the difference of 10% between the DOM fraction of the AD control condition and the *in situ* condition using inoculum PoD. Similarly, the increase in the SEOM fraction observed for all assessed conditions could not lead to a stated conclusion. Indeed, the increase in this fraction could result from plural organic matter conversion processes that might have differed in AD control and *in situ* biomethanation conditions.

Clearer results could be obtained through the evolution of the NEOM fraction. Indeed, the NEOM fraction is primarily associated with lignin-like molecules (Jimenez et al., 2015). This fraction exhibited a reduction in all conditions compared to the raw substrate. Moreover, the NEOM conversion was at least doubled in the *in situ* condition compared to the AD control condition (e.g., 28 % vs 10 % of the total COD of the sample corresponding to NEOM in the AD control vs *in situ* condition, using the inoculum D). The decrease in the NEOM portion could be attributed to the biodegradation activity of the microbial community. Indeed, through hydrolysis, the organic matter of the NEOM could have become more accessible and be extracted into the PEOM fraction, as the sum of the PEOM and NEOM fractions in all final digestates was close to the NEOM fraction of the substrate (Fig. 3). Following this transformation scheme, the organic matter contained in the PEOM fraction could have become more accessible for biodegradation, thus being associated with more accessible fractions (i.e., REOM, SEOM, SPOM, or DOM), facilitating, in this case, their conversion for CH₄ production. Subsequently, these results reinforce the statement of an improvement in organic matter conversion in the *in situ* biomethanation condition in comparison with the AD process.

To ensure the robustness of these results, which were surprising considering the inhibitory effect of H₂ on AD metabolism, the DEGOM fraction analyzed at the end of the experiment was compared to the methane produced during the experiment (from week 0 to week 6). In the case of inoculum D, it was estimated 11 % and 34 % of the 30 g·L⁻¹ of total COD in the substrate were consumed under AD control and *in situ* biomethanation conditions, respectively, representing an increase of 1.4 gCOD removal between the two conditions, for a 200 mL working volume. As the experiment lasted 2 HRT, 400 NmL of substrate were used, doubling the observed difference in COD removal (2.8 gCOD). The CH₄ production associated to OM degradation corresponded to 10.7 and 14.1

gCOD under AD control and *in situ* conditions, respectively, representing an increase of 3.4 gCOD. Similar results were obtained for reactors using the PoD inoculum, with a 2.4 gCOD removal increase (considering 400 mL of influent) and a 2.1 gCOD increase in CH₄ production between the control and *in situ* biomethanation conditions. Therefore, the combined analysis of the evolution of the DEGOM fraction and methane production clearly supports an enhancement of the organic matter biodegradation activity performed by the microbial community within H₂ addition in the reactor. As no other study using such a method to investigate the impact of H₂ on *in situ* biomethanation was retrieved, it was not possible to compare the present analysis with relevant literature.

### 3.4. Hydrogen-induced changes in microbial communities

The relative abundance of bacterial communities was monitored throughout the experiments (Fig. 4), by analyzing the initial inocula and the digested media sampled on week 1 (i.e., end of the start-up phase), week 3 (after one HRT), and week 6 (after two HRT). However, only minor changes were observed regarding the structure of the bacterial community, whatever the conditions. For both inocula, the dominant phyla were *Bacteroidetes* and *Firmicutes*, which are common phyla of bacteria found in AD process communities (Carballa et al., 2015). The proportions of these two phyla remained stable from the beginning to the end of the experiment in AD conditions. In contrast, for the *in situ* condition, a slight increase in the proportions of *Bacteroidetes* and a concomitant decrease in *Firmicutes* were observed.

Regarding the relative abundance at family level, the decrease in proportion of a family from the MBA03 order and the growth of bacteria from the *Enterococcaceae* family were mainly attributed to the operational conditions, as these changes in the bacterial communities were observed for all conditions. Conversely, the *Rikenellaceae* family appeared to be more predominant in the *in situ* condition compared to the AD control. The retrieved members of the *Rikenellaceae* family were mostly attributed to the genus *vadinBC27* (wastewater sludge group; see supplementary material). Members of this genus were previously described as amino acid degraders in anaerobic processes (Guo et al., 2014). In addition, it was previously suggested that *vadinBC27* (wastewater sludge group) could degrade amino acids through syntrophic interaction with hydrogenotrophic methanogens (Li et al., 2015).

<figure><img src="image_7.png" /><figcaption>**Fig. 4. Evolution of bacterial communities in assessed conditions.** Relative abundance of bacterial communities at Family level, classified by Phylum at the beginning of the experiment, at the start of H₂ injections (Week 1) after one HRT (Week 3) and after two HRT (Week 6). The 6 dominant Families retrieved per Phylum are displayed while other families are summed up in “Other” categories for each Order; Displayed Phyla represented more than 5 % of the overall bacterial community; D AD: Digestate of Digester as inoculum in anaerobic digestion conditions; D *in situ*: Digestate of Digester as inoculum in *in situ* biomethanation conditions; PoD AD: Digestate of Post-digester as inoculum in anaerobic digestion conditions; PoD *in situ*: Digestate of Post-digester as inoculum in *in situ* biomethanation conditions.</figcaption></figure>
Microorganisms from the *Ruminococcaceae* family were more abundant in AD control conditions than with H₂ injections. The effect of H₂ on that family remains uncertain, as these microorganisms are considered cellulolytic bacteria, promoting VFA and H₂ production through lignocellulosic biomass such as energy cover crops (Wojcieszak et al., 2017). Overall, no clear impact of H₂ on the bacterial community structure of the inocula could be stated from these results.

The lack of change in the main structure of the bacterial communities, despite the improved degradation of organic matter, may suggest a rapid adaptation of bacterial activity in the presence of H₂, as bacteria from *Bacteroidetes* and *Firmicutes* phyla are known for their diverse metabolic activities during AD (Campanaro et al., 2016; Treu et al., 2018; Westerholm et al., 2019). Besides, H₂ might impact only a minor part of the community, which may not be observable through the analysis of the overall bacterial community. For instance, syntrophic bacteria responsible for VFA degradation and H₂ production constitute a small portion of the AD community (Stams et al., 2012). These bacteria may be directly stimulated by the addition of H₂, either through the dissolved H₂ concentration or indirectly through the impact of H₂ on archaea structure and activity.

This hypothesis is supported by the high selective pressure exerted by H₂ on the archaeal communities in the *in situ* conditions compared to the AD control (Fig. 5). As already reported in the literature (Agneessens et al., 2017; Thapa et al., 2022), the injection of H₂ over several HRT led to substantial structural changes in the archaeal communities. In the present experiments, whatever the inoculum, the community composition was comprised of over 85 % archaea from the *Methanocorpusculum* genus at the end of the *in situ* experiment (i.e., after two HRT). Members of the *Methanocorpusculum* genus are known hydrogenotrophic methanogens (HM), exhibiting relatively rapid doubling times ranging from 8 to 14 h for identified species using H₂ and CO₂ as substrates (Zellner et al., 1989a; Zhao et al., 1989). Despite the low relative abundance of the *Methanocorpusculum* genus of less than 1 % in the inocula, it effectively outcompeted members of the *Methanobacterium* genus (also identified as HM) for H₂ consumption, with the latter representing over 40 % of the initial archaeal community in both inocula. The literature indicates that members of *Methanobacterium*, growing at mesophilic temperatures, have doubling times ranging from 5 h to 48 h when using H₂ and CO₂ as substrates (Garcia et al., 2000). In the present experiment, the strong development of the *Methanocorpusculum* genus suggests that

its identified members in the community exhibited faster growth using exogenous H₂ than the *Methanobacterium* genus that were initially dominant in the inoculum. In addition, members of *Methanobacterium* genus reported optimal growth temperature of 38.3 ± 3.5 °C, considering 19 species; (see supplementary material) (Garcia et al., 2000). In comparison, species from *Methanocorpusculum* genus reported optimal growth temperature of 35.2 ± 2.7 °C, considering 5 species, (see supplementary material) (Zellner et al., 1989b), which may have favored them during the experiment.

The growth of the *Methanocorpusculum* genus members increased the proportion of archaea in the community. Quantifying bacteria and archaea using qPCR revealed that the archaea/bacteria ratio (A/B ratio) (16S rDNA gene copy / 16S rDNA gene copy) increased from 2 % to 10 % and from 2 % to 9 % during the semi-continuous *in situ* biomethanation process using inoculum D and PoD, respectively. This increase in the archaeal population was not observed in AD control, where the A/B ratio remained stable at 2 % throughout the experiment for both inocula. The steadiness of the archaeal proportion in the AD control communities aligned with the balanced archaeal communities obtained in this process, indicating its stability under the control condition. Specifically, AD control communities were composed of three main genera of archaea, each representing approximately 30 % of the community at the end of the experiment: members of the *Methanobacterium* genus (HM), members of the *Methanosarcina* genus, and a genus of members of the *Batharchaeotha* phylum. While the metabolic activity of the *Bathyarchaeota* phylum members remains uncertain, members of *Methanosarcina* are known for their versatile methanogenic activity, with the capacity to produce CH₄ either from acetate or H₂ and CO₂ (De Vrieze et al., 2012; Evans et al., 2015; Maus et al., 2018). This composition of the archaeal communities in AD control was consistent with the archaeal communities of AD consortia observed in other studies (Braga Nan et al., 2020), suggesting stable methanogenesis under these conditions, despite changes in the community between the beginning and the end of the experiment.

However, the H₂ pulsed injection strategy used in the present experiment may have mitigated the known inhibitory effect of H₂ on the AD reaction chain, allowing an enrichment in hydrogenotrophic methanogens. As these species became dominant in the population, the overall H₂ uptake capacity of the community could have increased, reaching values close to the H₂ gas/liquid mass transfer rate of the experimental system. Subsequently, H₂ dissolved concentration could have been drastically reduced, limiting its effect on H₂ sensitive metabolisms.

At the same time, in the presence of an H₂-adapted archaeal population and with repeated H₂ additions, new syntrophic interactions might have been promoted, as previously reported by Basile et al. (2020). Additionally, the development of syntrophic interaction during *in situ* biomethanation of similar substrates was suggested in previous research (Treu et al., 2019). This synergy among microbial communities could have improved the AD reaction chain. Indeed, the H₂ uptake rate of the microbial community has been shown to impact metabolisms such as syntrophic acetate oxidation and VFA consumption pathways (Capson-Tojo et al., 2021). In the present experiment, the increased H₂ uptake rate, associated with the growth of hydrogenotrophic methanogens, could have indirectly improved acetogenic metabolisms, leading to improved OM biodegradation.

## 4. Conclusions

This study highlights that OM biodegradation was 2.5 times higher due to H₂ injections. This result was corroborated with the increase of least 626 NmL in CH₄ production associated to OM biodegradation with H₂ injections. Similar improvements in biodegradation activity were observed regardless of the initial microbial activity of the inocula used. Further investigations are required to elucidate the impact of H₂ on the AD reaction chain. These findings offer new perspectives for the

<figure><img src="image_8.png" /><figcaption>**Fig. 5.** Evolution of archaeal communities in assessed conditions. Relative abundance of archaeal communities at genus level at the beginning of the experiment, at the start of H₂ injections (Week 1) after one HRT (Week 3) and after two HRT (Week 6); D AD: Digestate of Digester as inoculum in anaerobic digestion conditions; D *in situ*: Digestate of Digester as inoculum in *in situ* biomethanation conditions; PoD AD: Digestate of Post-digester as inoculum in anaerobic digestion conditions; PoD *in situ*: Digestate of Post-digester as inoculum in *in situ* biomethanation conditions.</figcaption></figure>
development of *in situ* biomethanation, as it could enhance CH4 pro-
duction through both improved organic matter conversion and the CO2
transformation into CH4.

CRediT authorship contribution statement

**M. Mahieux:** Writing – review & editing, Writing – original draft, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. **Q. Aemig:** Writing – review & editing, Validation, Supervision, Methodology, Conceptualization. **C. Richard:** Writing – review & editing, Validation, Supervision, Investigation, Conceptualization. **J-P. Delgenès:** Writing – review & editing, Validation, Supervision. **M. Juge:** Project administration, Funding acquisition. **E. Trably:** Writing – review & editing, Validation, Supervision, Conceptualization. **R. Escudie:** Writing – review & editing, Validation, Supervision, Project administration, Methodology, Funding acquisition, Conceptualization.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Data availability

Data will be made available on request.

Acknowledgments

The National Association for Research and Technology (ANRT) is gratefully acknowledged for PhD fellowship allocated to Margot Mahieux under CIFRE convention N°2021/1463. The authors are grateful for Gaëlle Santa-Catalina's work concerning biomolecular tasks and 16S sequencing. They also thank Julie Jimenez for her advices concerning the used fractionation method and its analysis.

**Appendix A. Supplementary data**

Supplementary data to this article can be found online at https://doi.
org/10.1016/j.biortech.2024.131101.

References

Agneessens, L.M., Ottosen, L.D.M., Voigt, N.V., Nielsen, J.L., de Jonge, N., Fischer, C.H., Kofoed, M.V.W., 2017. In-situ biogas upgrading with pulse H₂ additions: The relevance of methanogen adaption and inorganic carbon level. Bioresour. Technol. 233, 256–263. https://doi.org/10.1016/j.biortech.2017.02.016.

Agnnessens, L.M., Ottosen, L.D.M., Andersen, M., Berg Olesen, C., Feilberg, A., Kofoed, M.V.W., 2018. Parameters affecting acetate concentrations during in-situ biological hydrogen methanation. Bioresour. Technol. 258, 33–40. https://doi.org/10.1016/j.biortech.2018.02.102.

Angelidaki, I., Treu, L., Tsapekos, P., Luo, G., Campanaro, S., Wenzel, H., Kougias, P.G., 2018. Biogas upgrading and utilization: Current status and perspectives. Biotechnol. Adv. 36, 452–466. https://doi.org/10.1016/j.biotechadv.2018.01.011.

APHA, 2017. Standard methods for the examination of water and wastewater analysis. (20th ed).

Basile, A., Campanaro, S., Kovalovszki, A., Zampieri, G., Rossi, A., Angelidaki, I., Valle, G., Treu, L., 2020. Revealing metabolic mechanisms of interaction in the anaerobic digestion microbiome by flux balance analysis. Metab. Eng. 62, 138–149. https://doi.org/10.1016/j.ymben.2020.08.013.

Bassani, I., Kougias, P.G., Treu, L., Angelidaki, I., 2015. Biogas upgrading via hydrogenotrophic methanogenesis in two-stage continuous stirred tank reactors at mesophilic and thermophilic conditions. Environ. Sci. Technol. 49, 12585–12593. https://doi.org/10.1021/acs.est.5b03451.

Bio2E, 2018. Environmental Biotechnology and Biorefinery Facility. https://doi.org/
10.15454/1.55723403446854E12.

Boe, K., Karakashev, D., Trably, E., Angelidaki, I., 2009. Effect of post-digestion temperature on serial CSTR biogas reactor performance. Water Res. 43, 669–676. https://doi.org/10.1016/j.watres.2008.11.037.

Braga Nan, L., Trably, E., Santa-Catalina, G., Bernet, N., Delgenès, J.-P., Escudie, R., 2020. Biomethanation processes: New insights on the effect of a high H₂ partial pressure on microbial communities. Biotechnol. Biofuels 13. https://doi.org/
10.1186/s13068-020-01776-y.

Bundhoo, M.A.Z., Mohee, R., 2016. Inhibition of dark fermentative bio-hydrogen production: A review. Int. J. Hydrogen Energy 41, 6713–6733. https://doi.org/
10.1016/j.ijhydene.2016.03.057.

Campanaro, S., Treu, L., Kougias, P.G., De Francisci, D., Valle, G., Angelidaki, I., 2016. Metagenomic analysis and functional characterization of the biogas microbiome using high throughput shotgun sequencing and a novel binning strategy. Biotechnol. Biofuels 9, 26. https://doi.org/10.1186/s13068-016-0441-1.

Capson-Tojo, G., Moscoviz, R., Astals, S., Robles, À., Steyer, J.-P., 2020. Unraveling the literature chaos around free ammonia inhibition in anaerobic digestion. Renew. Sustain. Energy Rev. 117, 109487 https://doi.org/10.1016/j.rser.2019.109487.

Capson-Tojo, G., Astals, S., Robles, À., 2021. Considering syntrophic acetate oxidation and ionic strength improves the performance of models for food waste anaerobic digestion. Bioresour. Technol. 341, 125802 https://doi.org/10.1016/j.biortech.2021.125802.

Carballa, M., Regueiro, L., Lema, J.M., 2015. Microbial management of anaerobic digestion: exploiting the microbiome-functionality nexus. Curr. Opinion Biotechnol. Environ. Biotechnol. Energy Biotechnol. 33, 103–111. https://doi.org/10.1016/j.copbio.2015.01.008.

Cazier, E.A., Trably, E., Steyer, J.-P., Escudie, R., 2019. Reversibility of hydrolysis inhibition at high hydrogen partial pressure in dry anaerobic digestion processes fed with wheat straw and inoculated with anaerobic granular sludge. Waste Manag. 85, 498–505. https://doi.org/10.1016/j.wasman.2019.01.019.

De Vrieze, J., Hennebel, T., Boon, N., Verstraete, W., 2012. Methanosarcina: The rediscovered methanogen for heavy duty biomethanation. Bioresour. Technol. 112, 1–9. https://doi.org/10.1016/j.biortech.2012.02.079.

Díaz, I., Fdz-Polanco, F., Mutsvene, B., Fdz-Polanco, M., 2020. Effect of operating pressure on direct biomethane production from carbon dioxide and exogenous hydrogen in the anaerobic digestion of sewage sludge. Appl. Energy 280, 115915. https://doi.org/10.1016/j.apenergy.2020.115915.

Dincer, I., Acar, C., 2015. Review and evaluation of hydrogen production methods for better sustainability. Int. J. Hydrogen Energy 40, 11094–11111. https://doi.org/
10.1016/j.ijhydene.2014.12.035.

Evans, P.N., Parks, D.H., Chadwick, G.L., Robbins, S.J., Orphan, V.J., Golding, S.D., Tyson, G.W., 2015. Methane metabolism in the archaeal phylum Batharchaeota revealed by genome-centric metagenomics. Science 350, 434–438. https://doi.org/
10.1126/science.aac7745.

Fernández-Domínguez, D., Patureau, D., Houot, S., Sertillanges, N., Zennaro, B., Jimenez, J., 2021. Prediction of organic matter accessibility and complexity in anaerobic digestates. Waste Manag. 136, 132–142. https://doi.org/10.1016/j.wasman.2021.10.004.

Fernández-Domínguez, D., Yekta, S.S., Hedenström, M., Patureau, D., Jimenez, J., 2023. Deciphering the contribution of microbial biomass to the properties of dissolved and particulate organic matter in anaerobic digestates. Sci. Total Environ. 877 https://doi.org/10.1016/j.scitotenv.2023.162882.

Garcia, J.-L., Patel, B.K.C., Ollivier, B., 2000. Taxonomic, phylogenetic, and ecological diversity of methanogenic archaea. Anaerobe 6, 205–226. https://doi.org/10.1006/anae.2000.0345.

Guo, X., Wang, C., Sun, F., Zhu, W., Wu, W., 2014. A comparison of microbial characteristics between the thermophilic and mesophilic anaerobic digesters exposed to elevated food waste loadings. Bioresour. Technol. 152, 420–428. https://doi.org/10.1016/j.biortech.2013.11.012.

IEA, 2023. Medium-Term Gas Report 2023 (No. Including the Gas Market Report, Q4-2023). Paris.

Jimenez, J., Aemig, Q., Doussiet, N., Steyer, J.-P., Houot, S., Patureau, D., 2015. A new organic matter fractionation methodology for organic wastes: Bioaccessibility and complexity characterization for treatment optimization. Bioresour. Technol. 194, 344–353. https://doi.org/10.1016/j.biortech.2015.07.037.

Jimenez, J., Charnier, C., Kouas, M., Latrille, E., Torrijos, M., Harmand, J., Patureau, D., Spérandio, M., Morgenroth, E., Béline, F., Ekama, G., Vanrolleghem, P.A., Robles, A., Seco, A., Batstone, D.J., Steyer, J.-P., 2020. Modelling hydrolysis: Simultaneous versus sequential biodegradation of the hydrolysable fractions. Waste Manag. 101, 150–160. https://doi.org/10.1016/j.wasman.2019.10.004.

Li, L., He, Q., Ma, Y., Wang, X., Peng, X., 2015. Dynamics of microbial community in a mesophilic anaerobic digester treating food waste: Relationship between community structure and process stability. Bioresour. Technol. 189, 113–120. https://doi.org/
10.1016/j.biortech.2015.04.015.

Luo, G., Angelidaki, I., 2013. Co-digestion of manure and whey for in situ biogas upgrading by the addition of H₂: Process performance and microbial insights. Appl. Microbiol. Biotechnol. 97, 1373–1381. https://doi.org/10.1007/s00253-012-4547-5.

Luo, G., Johansson, S., Boe, K., Xie, L., Zhou, Q., Angelidaki, I., 2012. Simultaneous hydrogen utilization and in situ biogas upgrading in an anaerobic reactor. Biotechnol. Bioeng. 109, 1088–1094. https://doi.org/10.1002/bit.24360.

Mahieux, M., Richard, C., Aemig, Q., Delgenès, J.-P., Juge, M., Trably, E., Escudie, R., 2024. Archaeal community composition as key driver of H₂ consumption rates at the start-up of the biomethanation process. Sci. Total Environ., 172922 https://doi.org/
10.1016/j.scitotenv.2024.172922.

Maus, I., Rumming, M., Bergmann, I., Heeg, K., Pohl, M., Nettmann, E., Jaenicke, S., Blom, J., Pühler, A., Schlüter, A., Sczyrba, A., Klocke, M., 2018. Characterization of Batharchaeota genomes assembled from metagenomes of biofilms residing in mesophilic and thermophilic biogas reactors. Biotechnol. Biofuels 11, 167. https://doi.org/10.1186/s13068-018-1162-4.

McMurdie, P.J., Holmes, S., 2013. phyloseq: An R package for reproducible interactive analysis and graphics of microbiome census data. PLoS One 8, e61217.
Noguer, M.C., Magdalena, J.A., Bernet, N., Escudié, R., Trably, E., 2022. Enhanced fermentative hydrogen production from food waste in continuous reactor after butyric acid treatment. Energies 15, 4048. https://doi.org/10.3390/en15114048.

Rafrafi, Y., Laguillaumie, L., Dumas, C., 2021. Biological methanation of H₂ and CO₂ with mixed cultures: Current advances, hurdles and challenges. Waste Biomass Valoriz. 12, 5259–5282. https://doi.org/10.1007/s12649-020-01283-z.

REPowerEU, 2022.

Stams, A.J.M., Sousa, D.Z., Kleerebezem, R., Plugge, C.M., 2012. Role of syntrophic microbial communities in high-rate methanogenic bioreactors. Water Sci. Technol. 66, 352–362. https://doi.org/10.2166/wst.2012.192.

Teunisse, G.M., 2022. Fantaxtic - Nested Bar Plots for Phyloseq Data (Version 2.0.1) [Computer software].

Thapa, A., Park, J.-G., Jun, H.-B., 2022. Enhanced ex-situ biomethanation of hydrogen and carbon dioxide in a trickling filter bed reactor. Biochem. Eng. J. 179, 108311 https://doi.org/10.1016/j.bej.2021.108311.

Thema, M., Bauer, F., Sterner, M., 2019. Power-to-Gas: Electrolysis and methanation status review. Renew. Sustain. Energy Rev. 112, 775–787. https://doi.org/10.1016/
j.rser.2019.06.030.

Treu, L., Campanaro, S., Kougias, P.G., Sartori, C., Bassani, I., Angelidaki, I., 2018.
Hydrogen-fueled microbial pathways in biogas upgrading systems revealed by
genome-centric metagenomics. Front. Microbiol. 9.

Treu, L., Tsapekos, P., Peprah, M., Campanaro, S., Giacomini, A., Corich, V., Kougias, P. G., Angelidaki, I., 2019. Microbial profiling during anaerobic digestion of cheese whey in reactors operated at different conditions. Bioresour. Technol. 275, 375–385. https://doi.org/10.1016/j.biortech.2018.12.084.

Van Vlierberghe, C., Carrere, H., Bernet, N., Santa-Catalina, G., Frederic, S., Escudie, R., 2022. Co-ensiling and field wilting investigated as preparation methods for the

ensiling of a wet harvested catch crop for biomethane production. Renew. Energy 195, 1230–1237. https://doi.org/10.1016/j.renenue.2022.06.078.

Wahid, R., Mulat, D.G., Gaby, J.C., Horn, S.J., 2019. Effects of H₂:CO₂ ratio and H₂ supply fluctuation on methane content and microbial community composition during in-situ biological biogas upgrading. Biotechnol Biofuels 12, 104. https://doi.org/10.1186/s13068-019-1443-6.

Westerholm, M., Schnürer, A., Westerholm, M., Schnürer, A. 2019. Microbial Responses to Different Operating Practices for Biogas Production Systems, in: Anaerobic Digestion. IntechOpen. https://doi.org/10.5772/intechopen.82815.

Wojcieszak, M., Pyzik, A., Poszytek, K., Krawczyk, P.S., Sobczak, A., Lipinski, L., Roubinek, O., Palige, J., Sklodowska, A., Drewniak, L., 2017. Adaptation of methanogenic inocula to anaerobic digestion of maize silage. Front. Microbiol. 8.

Zellner, G., Stackebrandt, E., Messner, P., Tindall, B.J., Conway de Macario, E., Kneifel, H., Sleytr, U.B., Winter, J., 1989. Methanocorpusculaceae fam. nov., represented by Methanocorpusculum parvum, Methanocorpusculum sinense spec. nov. and Methanocorpusculum bavaricum spec. nov. Arch. Microbiol. 151, 381–390. https://doi.org/10.1007/BF00416595.

Zennaro, B., Marchand, P., Latrille, E., Thoisy, J.-C., Houot, S., Girardin, C., Steyer, J.-P., Béline, F., Charnier, C., Richard, C., Accarion, G., Jimenez, J., 2022. Agronomic characterization of anaerobic digestates with near-infrared spectroscopy. J. Environ. Manage. 317, 115393 https://doi.org/10.1016/j.jenvman.2022.115393.

Zhao, Y., Boone, D.R., Mah, R.A., Boone, J.E., Xun, L., 1989. Isolation and characterization of Methanocorpusculum labreanum sp. nov. from the LaBrea Tar Pits. Int. J. Syst. Evol. Microbiol. 39, 10–13. https://doi.org/10.1099/00207713-39-1-10.

Zhu, X., Zhou, P., Chen, Y., Liu, X., Li, D., 2020. The role of endogenous and exogenous hydrogen in the microbiology of biogas production systems. World J. Microbiol. Biotechnol. 36, 79. https://doi.org/10.1007/s11274-020-02856-9.