# Phenolic compounds fingerprints show distinct ligninolytic bacteria behaviours during technical lignins conversion

Corinne Ivaldi, Brigitte Chabbert, Roland Molinié, David Crônier, David Mathiron, Jean Xavier Fontaine, François Mesnard, Frédéric Velard, Harivony Rakotoarivonina

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## To cite this version:

Corinne Ivaldi, Brigitte Chabbert, Roland Molinié, David Crônier, David Mathiron, et al. "Phenolic compounds fingerprints show distinct ligninolytic bacteria behaviours during technical lignins conversion." New Biotechnology, 2025, 89, pp.60-72. 10.1016/j.nbt.2025.06.004. hal-05144951

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Submitted on 24 Jul 2025

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# Phenolic compounds fingerprints show distinct ligninolytic bacteria behaviours during technical lignins conversion

Corinne Ivaldi<sup>a</sup>, Brigitte Chabbert<sup>a</sup>, Roland Molinié<sup>b</sup>, David Crônier<sup>a</sup>, David Mathiron<sup>c</sup>, Jean Xavier Fontaine<sup>b</sup>, François Mesnard<sup>b</sup>, Frédéric Velard<sup>d</sup>, Harivony Rakotoarivonina<sup>a,∗</sup> ◆

## ARTICLE INFO

### Keywords:

Phenol-oxidase activities
phenolic oligomers
lignins
metabolomics
UHPLC-ESI-HRMS

## ABSTRACT

Lignins, one of the main components of plant cell wall, and by-products of certain industries (paper and wood industries,...) are a renewable source of aromatic molecules. They can be degraded and transformed by microbial and enzymatic processes known to be respectful of the environment. Biological valorization of lignins remains challenging as biocatalysts are not sufficiently effective and efficient. Moreover, the chemical complexity and heterogeneity of lignins are a barrier to their use. Understanding the microbial behaviour on lignins by fingerprinting their efficient transformation could lead to the development of effective biological routes to valorise these aromatic polymers. Ligninolytic bacteria present some interesting features in term of ligninolytic enzymes productions, utilization of aromatic compounds *via* various intracellular pathways and the productions of molecules of interest from aromatic molecules. In this work, multiple approaches (growth studies, ligninolytic activities production, lignin modifications and phenolic compounds fingerprints) were used to understand the behaviour of two ligninolytic bacteria *Pandoraea norimbergensis* and *Comamonas composti*, in presence of lignins with different structures and origins. Results showed dissimilar growths profiles, lignin modifications, consumption and production of phenolic monomers and oligomers according to the bacteria and the lignin used. To achieve efficient transformation of lignins, suitable combination of biocatalysts and lignins is required and the microorganisms used must be selected on the basis of their metabolic capacity, and the structure and composition of lignin.

## 1. Introduction

Lignin, a three-dimensional polymer is tightly associated with cellulose and hemicellulose in plant cell wall. Lignin plays a role in structural strength, impermeability and water transport in plants [1]. It is composed of three phenolic units: *p*-hydroxyphenyl (H unit), guaiacyl (G unit) and syringyl (S unit) derived respectively from *p*-coumaryl alcohol, coniferyl alcohol and sinapyl alcohol which are linked by various carbon-carbon and ether bonds. Lignin content and monomer

composition vary according to botanical origin, tissue and cell wall strata [2]. Softwood contain mainly G units, hardwood G and S units while phenolic acids (i.e. ferulic and *p*-coumaric acids) are found in grass lignin in addition to the H, G and S units. Lignins are available at commercial or pilot scales as a coproduct of various processes (Kraft lignin for pulp and paper industry, lignocellulosic biorefineries...) and are commonly considered as technical lignins. Kraft lignin from wood represents 85 % of technical lignins production (around 650 kt/year) whereas soda/alkali lignins from non-woody biomasses represents

<footnote><sup>a</sup> Université de Reims Champagne Ardenne, INRAE, FARE, UMR A 614, Chaire AFERE, Reims, France</footnote>

<footnote><sup>b</sup> Université de Picardie Jules Verne, INRAE, BIOPI, UMRt BioEcoAgro, Amiens 1158, France</footnote>

<footnote><sup>c</sup> Université de Picardie Jules Verne, Plateforme Analytique, Amiens, France</footnote>

<footnote><sup>d</sup> Université de Reims Champagne-Ardenne, BIOS, Reims, France</footnote>

<footnote>*Abbreviations:* AS, acetosyringone; AV, acetovanillone; BM, basal medium; EC, extracellular; FA, ferulic acid; G, generation time; GC-MS, gas chromatography-mass spectrometry; HDMS, hexamethyldisilazane; IC, intracellular; LC-MS, Liquid chromatography-mass spectrometry; NMR, Nuclear magnetic resonance; OD, optical density; *p*-CA, *p*-coumaric acid; PCA, principal component analyses; POT, total phenol-oxidase; S, syringaldehyde; SA, syringic acid; SEM, scanning electron microscopy; UHPLC-EST-HRMS, ultra-high performance liquid chromatography – electrospray ionisation – high-resolution mass spectrometry; V, vanillin; VA, vanillic acid.</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: harivony.rakotoarivonina@univ-reims.fr (H. Rakotoarivonina).</footnote>
around 6.5 kt/year. Lignins from pretreatments during lignocellulosic
biorefinery (for bioethanol production) are estimated at ~140 kt/year,
mainly from pilot and demonstration scale. Lignin production will in-
crease in the coming years [3,4]. Technical lignins are highly modified
and heterogeneous, in terms of structure and chemical compositions
according to the extraction process and their botanical origin [5]. Kraft
lignin and alkali lignins have some common characteristics. They are
very condensed and rich in C-C linked substructures and contained
traces of sugars (<3 %) [6]. Because of their complexity lignins are
mainly used as energy source and burnt. In the context of climatic
change, to reduce the use of fossil fuels, materials and molecules alter-
native resources are sought [7]. Lignins can be used as a renewable
source of aromatic compounds, lignin-based materials, resins and ad-
hesives thereby upgrading their valorisation [5].

In the nature, lignins can be used by various microorganisms (fungi and bacteria) often organized in consortium [8]. Microorganisms first depolymerize lignin by producing extracellular oxidoreductases (mainly peroxidases and laccases) [9]. Fungi are the most studied for their lignin depolymerization actions [10]. Aromatic or lignin derived compounds released after depolymerization are used as carbon source (after they have been internalized inside of the cells via specific transporters) enabling microbial growth with or without the production of some molecules of interest [11,12].

The abilities of several bacteria belonging to various phyla *Burkholderiaceae*, *Sphingobacteriaceae*, *Pseudomonadaceae*, *Actinobacteria*, *Bacillaceae*, to use aromatic or lignin derived compounds is very well documented [9,13]. Numerous research studies deal with the ligninolytic potential of bacteria 1) by studying their ability to grow on lignin polymers, or on low molecular weight aromatic compounds derived from lignin (monomer, dimer, synthetic lignin); 2) by identifying genes encoding lignin modifying enzymes (laccases and peroxidases) or genes implicated in various pathways for aromatic compounds degradation and their expression. Thus, Gram-positive bacteria (*Rhodococci* and *Bacilli*) and many Gram-negative and aerobic bacteria like *Pandoraea* and *Comamonas* species are known for their ligninolytic properties by growing on technical lignins as Kraft lignin, and by expressing oxidoreductases genes or producing ligninolytic activities (laccase and peroxidase). They also can release phenolic compounds in the culture media [12,14–16]. During growth, laccase and peroxidase enzymes are secreted [11,17–20].

To elucidate the action of microorganisms during lignin trans-
formations, several approaches analyzed the changes in lignin structures
[21]. Vibrational spectroscopy such as infrared analysis allows the
identification of global structure changes within lignin through the
modification of functional groups. Whereas nuclear magnetic resonance
(NMR) can be used to point out specific modifications of lignin subunits,
inter-unit linkages or degree of lignin condensation...[21].

Metabolomic approaches have shown their relevance for the iden-
tification of lignin degradation products or metabolites produced after
growth on lignins [11,20,22]. Metabolomics profiles or fingerprints
could be achieved by either mass spectrometry (MS) associated with Gas
(GC) or Liquid (LC) chromatography or by nuclear magnetic resonance
(NMR) spectroscopy [23]. GC-MS have been implemented to identify the
phenolic monomers produced after the growth of ligninolytic microor-
ganisms on lignins or aromatic monomers [19,20].

Bacterial metabolomic profiles after growth have been studied by LC-MS/MS and allow the identification of main degradation products [21, 23]. Recently, lignin oligomers from Kraft lignin have been identified by a non-targeted UHPLC/HRMS approach [23]. Indeed, this approach has been successfully developed to characterize the metabolism of phenolic compounds in plants [24–26] allowing to identify or annotate not only phenolic monomers, but also phenolic oligomers.

Despite all these data, considering the complexity of lignins, these
studies do not necessarily reflect the ligninolytic capacity of the bacte-
ria. It is critical to understand the bacterial bioconversion of lignins
notably phenolic compounds, monomers but also some phenolic

oligomers that can represent an intermediary step during lignin
bioconversion and thus can reflect and/or reveal an efficient conversion
or transformation of lignins. To address these issues, in this study,
technical and commercial lignins with different compositions and
structures (Kraft lignin from conifers or soda lignin from grass (Proto-
bind P1000) were used. The lignins have been exposed against two
ligninolytic species *Pandoraea norimbergensis* and *Comamonas composti*
belonging to the Gram-negative bacterial groups (*Burkholderiaceae* and
*Comamonadaceae*). They are able to use and grow on lignins [19,20].
Kinetic growths studies of the bacteria in presence of the lignins as sole
carbon source were undertaken. Microscopic observations were carried
out to visualise the culture media (bacteria and lignins) before and after
growth. The phenol-oxidase activity produced by the bacteria and the
aromatic molecules released were quantified. A non-targeted finger-
printing analysis similar to metabolomics analyses by
ultra-high-performance liquid chromatography - electrospray ionisation
- high-resolution mass spectrometry (UHPLC-ESI-HRMS) focusing on
phenolic molecules was implemented to highlight possible variation in
the ability of the bacteria to use the different lignins. This approach
focused on the features of the aromatic compounds (monomers and
oligomers) present in the culture media at different time points during
growth. Finally, the structure of the lignins after growth was studied by
infrared spectroscopy.

## 2. Materials and methods

### 2.1. *Chemicals*

All the chemicals used come from Sigma-Aldrich (France) except for
glutaraldehyde (Electron Microscopy Sciences, England).

### 2.2. *Strain and culture media*

Kraft lignin was supplied by Sigma-Aldrich (France). Protobind P1000 lignin purchased from GreenValue SA (Orbe, Switzerland). The Kraft lignin is constituted by 85 % of lignin, traces of carbohydrates, and 3.6 % sulphur. It contained exclusively G-unit [16]. Protobind P1000 contained around 90 %, 2.4 % and 1.1 % DM of lignin, carbohydrates and sulphur respectively with a ratio of S/G/H units of 50/39/11 [6]. *Pandoraea norimbergensis* (DMSZ 24563) and *Comamonas composti* (DSMZ 21721) strains were cultivated on basal medium (BM) containing (g/L) 6.78 g of Na₂HPO₄; 3 g of KH₂PO₄; 0.5 g of NaCl; 0.13 g of (NH₄)₂SO₄; and 2 g yeast extract and supplemented with CaCl₂ 100 µM; MgSO₄ 2 mM. Glucose or lignins at 5 g/l (w/v) were added in the BM as main carbon source. For lignins containing media, BM and lignins were mixed before autoclaving at 120°C during 20 min. For culture media with glucose, heat sterilized glucose was added to the autoclaved BM.

### 2.3. *Bacterial growth*

Bacterial pre-cultures in 20 ml of BM supplemented by glucose were incubated at 30°C under constant agitation at 120 rpm until reaching an optical density at 600 nm (OD600 nm) of around 1. Bacteria were then inoculated at 1 % (v/v) in 20 ml or 200 ml volume of BM containing lignins as main carbon source. Controls (media without bacteria) and inoculated cultures were incubated at 30°C and 120 rpm for 8 days. Cultures and controls were performed in triplicate for each bacterium and each lignin. Bacterial growth was monitored by measuring OD600 nm in a spectrophotometer (Jenway 7300 spectrophotometer) using 1 ml samples taken after 1, 2, 3, 6 and 8 days of cultivations. Before measurement, to separate the lignin particles from the bacteria and the medium, the samples were centrifuged at 50 × g for 1 min. Control media treated as bacterial cultures were used as blanks for reading absorbances. Generation time (g) was calculated during exponential growth phase for the 200 ml volume cultures.
## 2.4. Microscopy analyses

Drops (20 µl) of fresh and homogeneous cultures or control media after 8 days of incubation, were deposited on a glass slide and covered with a coverslip, then directly observed under a light microscope (Nikon Eclipse Ci, Pays-bas) at × 1000 magnification with immersion oil objective. For scanning electron microscopy (SEM), the samples were prepared as described previously [27]. Briefly, drops were dried on glass coverslip, fixed in 2.5 % (w/v) glutaraldehyde in phosphate buffer saline (50 mM, pH 7), washed at room temperature in graded ethanol solutions (50 %, 70 %, 90 % and 100 %) and finally dehydrated with hexamethyldisilazane (HDMS). After air-drying at room temperature until complete evaporation of HDMS, samples were sputtered with a thin gold-palladium film using a JEOL ion sputter JFC 1100 instrument. Samples were then observed using a Schottky Field Emission Scanning Electron Microscope (JEOL JSM-7900 F, JEOL, France) with a primary beam energy of 2 kV. Observations were performed with the triplicates of cultures.

## 2.5. Extracts and residues preparation

From the 200 ml lignin cultures (for bacterial growth studies), 30 ml of culture samples were collected after 3- and 6-days incubation and prepared for the various analyses. Lignin residues when available (Protobind lignin) was separated to the culture by centrifugation at 50 × g for 1 min and were washed 3 times with distilled water, frozen at -20°C prior freeze drying. Culture media (20 ml), containing bacteria, were then centrifuged for 20 min at 5000 × g. The supernatant constituted the extracellular (EC) fractions. The EC fractions were aliquoted in the appropriate volumes and used for phenol-oxidase activities measurement and protein quantification or kept at -20°C for phenolic compounds and metabolite analyses. The bacterial cells pellets were resuspended in 1 ml of 50 mM Tris-HCl buffer (pH 7), and mechanically disrupted (Fastprep 24 5 G, MP Biomedicals, USA) prior to centrifugation for 5 min at 10000 × g. The recovered supernatant constituted the intracellular (IC) fractions. The IC fractions were aliquoted and immediately used for phenol-oxidase activities measurement and protein quantification.

## 2.6. Protein quantification

50 µl IC or EC fractions for each experimental triplicate were freeze-dried and subjected to total Nitrogen (N) quantifications by using a chromatographic column (CHN separation column 2 m, 6x5mm, SS, HSPQ, 2 mm unions, Eurovector, Italy) coupled to a TCD (Thermal Conductivity Detector) before being injected into an isotope mass spectrometer (Euro-EA, Eurovector, Italy). Proteins quantification refers to the nitrogen concentration (mg/ml) detected in the sample multiplied by the factor 6.25 usually used [28].

## 2.7. Phenol-oxidase activity measurement

Total phenol-oxidase (POT) activity (laccases and peroxidases) in EC and IC fractions was estimated after 3- and 6-days incubation using a pyrogallol as substrate as described previously [16]. The release of the oxidized product (purpurogallin) was monitored at 420 nm every 30 s for 5 min using the kinetic mode of a spectrophotometer (SPECORD PLUS®, Analytik Jena AG, Saint-Aubin, France) thermregulated at 30°C. The reaction medium (total volume of 1 ml) consisted of 50 mM Tris-HCl buffer pH 7, 8 mM pyrogallol substrate, 1 mM H₂O₂ and 100 µl of IC or EC fractions diluted or not. For laccase activity measurements, the reaction was performed in absence of H₂O₂. For EC fractions, activities in the non-inoculated media controls were determined and subtracted to the activities determined in the EC culture samples. Auto-oxidation of the substrate (8 mM pyrogallol) was also carried out in presence of 50 mM Tris-HCl pH 7 buffer at 30°C. Interference

oxidation activities of other non-enzymatic compounds from the cultures were quantified after denaturing the EC and IC fractions 10 min at 100°C. The values of the auto- and interference oxidations have been subtracted from the values of the assays and controls activities. Activities were expressed in IU·ml⁻¹ with one international unit (IU) corresponding to the apparition of 1 µmole of purpurogallin (oxidized pyrogallol) per minute at 30°C and pH 7. The molar extinction of purpurogallin ε420 nm = 1360 M⁻¹·cm⁻¹ used was determined earlier under the same conditions [16]. To determine peroxidase activity, lacase activity was subtracted to the total phenol-oxidases activity. All the experiment was performed in triplicate.

## 2.8. Phenolic compounds analysis in the culture media

Phenolic compounds were analysed using 5 ml of EC fractions. After acidification to pH 1 with 6 M HCl and extraction with diethyl ether, the organic fraction was dried and solubilized in 1.5 ml of a H₂O/methanol solution (50:50 v/v) then analysed by high-performance liquid chromatography (HPLC) on a Spherisorb S5-0DS2, RP18, 4.6 × 250 mm column equipped with a photodiode array detector (Waters, USA) as previously described [29]. The aromatic monomers were detected and quantified at 302 nm and 280 nm using 3,4,5-trimethoxycinnamic acid and 3-ethoxy-4-hydroxybenzaldehyde respectively as internal standards and reference phenolic monomers. Analyses were performed on the EC fractions obtained from triplicates controls and inoculated cultures after 3- and 6-days incubation.

## 2.9. Fingerprinting of lignin degradation products

Non-targeted analysis was performed using 5 ml of EC fractions taken at day 3 and day 6. Samples were prepared as for the analysis of phenolic molecules, except that there is no internal standard and the soluble fraction after evaporation were solubilised in 10 ml of a mixture of methanol/water (50/50 v/v). UHPLC-ESI-HRMS profiling analysis was carried out according to the previous analytical method [30]. Briefly, the analysis was performed on an ACQUITY UPLC I-class chain coupled to the Vion IMS Quadrupole time-of-flight high-resolution mass spectrometer (HRMS), equipped with an electrospray ionization (ESI) source (Z-spray) and an additional spray for the reference compound (Waters, Manchester, UK). A double detection was performed using a photodiode array (PDA) detector (UV detection between 190 and 500 nm) and by ESI-HRMS (range 50-2000 Da). The ionization was performed in positive and negative modes. Elution was performed with a mobile phase flow rate of 0.5 ml/min consisting of a mixture of water (A) and methanol (B) containing 0.1 % of formic acid. The gradient starts at t = 0, ratio (A/B) 80:20; at t = 0.5 min, 80:20; at t = 5 min, 40:60; at t = 6 min, 10:90; at t = 7 min, 10:90; at t = 7.5 min, 80:20; and at t = 10 min, 80:20. One µL of the sample was injected into a Kinetex Biphenyl column (2.1 x 100 mm; 1.7 µm) (Phenomenex, Torrance, California, USA) heated to 55 °C. Data were acquired in high-definition MS<sup>E</sup> (HDMS<sup>E</sup>) mode including a low-energy function (CE 6 eV) as well as a second high-energy function (CE 25-50 eV) for both positive and negative mode ionizations to obtain information on fragmentation. Data (negative mode) were acquired and processed with the UNIFI software (version 1.9.4, Waters), allowing to generate the data matrix with default parameters comprising, for each detected ion corresponding at a variable, the retention time, mass-to-charge ratio (m/z) value and chromatogram peak area. The non-targeted metabolomic data matrix was cleaned by removing the variables present in the blank, the variables with a significant variance (>35 %) in the quality control (QC), the variables with a retention time value below 0.75 min and above 5.75 min (after 6 min is the cleaning step) and the variables with a peak area value above 500. For discriminant lignin degradation products, three levels of identification or annotation were performed. The first level is a comparison with commercial standards based on similarity of accurate masses, retention times and fragmentation patterns (HDMS<sup>E</sup>
spectra) and allowed the identification of three products (variable 3, 5 and 7). The second level is a comparison with literature data that described lignin degradation products based on similarity of accurate masses, retention times and fragmentation patterns (highlighted in bold in SM2) and allowed annotation for five products (variable 12, 14, 17, 20, 22). The last level is a primary annotation based on molecular formula to define the number of lignin unit (mono- (M), di-(D) and trimers (T)) and was performed for all discriminant lignin degradation products i.e., all 29 variables. This annotation was correlated according to their elution order (retention times). All experiments were performed in triplicate.

## 2.10. Residual lignin analysis

Fourier Transform Infrared (FTIR) spectral analysis was performed on 2 mg freeze-dried residues corresponding to each experimental triplicate of bacterial cultures and lignin controls, was mixed with KBr (120 mg) using a Nicolet iS50 (Thermo Fisher Scientist, USA) spectrophotometer. Spectra were obtained from 16 scans recorded between 400 and 4000 $cm^{-1}$ with a spectral resolution of 4 $cm^{-1}$. All spectra were baseline corrected using OMNIC software (version 9.12, Thermo Fisher Scientist, USA) and normalized by fixing the area under spectrum between 1900 and 700 $cm^{-1}$ [16]. Infrared vibration bands were assigned according to literature data (Supplementary material 1).

## 2.11. Data analysis

One-way analysis of variance (ANOVA) was performed to assess the variation in protein concentrations, enzymatic activities and in phenolic monomers between culture conditions on each lignin using SigmaPlot® software (version 12.0, Systat, USA). For FTIR, means intensities of bands of interest between control and bacterial cultures were compared by the same ANOVA test. The Student Newman Keuls method was then used to perform pairwise multiple comparison. Results were considered significant at *p*-value < 0.05 unless specified in the text. For phenolic compounds fingerprinting, principal component analysis (PCA) based on a pareto scaling method using FactoMiner (version 2.4) and heatmap based on a natural logarithm scaling method using the heatmap package (version 1.0.12) were performed with the R software (version 3.6.3, company Foundation for Statistical Computing, Vienna, Austria).

# 3. Results

## 3.1. Bacterial growth on lignins and protein concentrations

Preliminary studies of the bacterial growths in 20 ml culture volumes, show the abilities of the two bacteria to use and to grow on Kraft and Protobind lignins as carbon source. These studies allowed to identify the different growth phases and to determine samplings at the beginning and the end of stationary phases (Supplementary material 2). Fig. 1 shows the growth of both bacteria on Kraft and Protobind lignins during 8 days at 30°C in large volume cultures. On Kraft lignin (Fig. 1a), the growth profiles of the two bacteria were roughly similar. *P. norimbergensis* started growing immediately after inoculation with a regular accelerating phase during the first 24 h whereas *C. composti* growth began after a 24 h lag phase. The exponential phase lasted approximately 24 h in both cultures and similar OD600nm (around 0.7) were obtained. The generation time (g) was 1.5-fold higher for *P. norimbergensis* compared to *C. composti* (*p*-value < 0.001) and represented 9.41 ± 1.97 h and 5.92 ± 0.53 h respectively. The stationary phase begun after 3 days of cultivations for both bacteria with an OD600nm of 0.78 for *P. norimbergensis* and 0.66 for *C. composti*. The protein concentrations in the different fractions (EC and IC) were quantified at the beginning and late stationary phases of cultivation. IC protein contents were similar at day 3 in the two bacteria and significantly increased at day 6 for *P. norimbergensis* (Table 1). Concerning EC proteins, the concentration increased significantly between day 3 and 6 for both bacteria and was higher in *C. composti* than in *P. norimbergensis*.

On Protobind lignin, growth profiles of the two bacteria were very different (Fig. 1b). The growth of *P. norimbergensis* consisted in two distinct stages. After a lag phase of 24 h, the bacterium showed a very short but quick exponential growth (g = 2,30 ± 0,01 h) to reach an OD600nm around 0.2, after which a 48 h stationary phase was observed. This first stage was followed by a second growth (g = 5,79 ± 0,23 h) lasting approximately 8 h until a maximal OD600nm of 0.65 ± 0.01 was reached at day 3. The growth declined sharply at the end of cultivation, after day 6. Protein quantifications showed a significant increase of EC proteins between day 3 and day 6 while IC proteins remained stable (Table 1).

*C. composti* started to grow after 2 days of cultivation with Protobind lignin. The growth was rather slow but regular until reaching a maximal OD600nm of around 0.44 without showing clear distinction between exponential and stationary phases between day 2 and day 6. The

<figure><img src="image_4.png" /><figcaption>**Fig. 1.** Bacterial growth for *P. norimbergensis* and *C. composti* on Kraft lignin (a) and Protobind lignin (b). Cultures were performed in triplicate and incubated at 30°C and 120 rpm during 8 days.</figcaption></figure>
<caption>**Table 1**</caption>

growth estimated with OD600nm and protein concentrations in extracellular (EC) and intracellular (IC) compartments for *P. norimbergensis* and *C. Composti* cultivated on Kraft and Protobind lignins.

<table><thead><tr><th rowspan="2"></th><th rowspan="2"></th><th colspan="2">Kraft lignin</th><th colspan="2">Protobind lignin</th></tr><tr><th>3 days</th><th>6 days</th><th>3 days</th><th>6 days</th></tr></thead><tbody><tr><td rowspan="2">OD 600 nm</td><td><i>P. norimbergensis</i></td><td>0.78 ± 0.02</td><td>0.76 ± 0.02</td><td>0.65 ± 0.01</td><td>0.59 ± 0.12</td></tr><tr><td><i>C. composti</i></td><td>0.66 ± 0.02</td><td>0.70 ± 0.01</td><td>0.27 ± 0.01</td><td>0.44 ± 0.2</td></tr><tr><td rowspan="2">EC protein</td><td><i>P. norimbergensis</i></td><td>0.69 ± 0.05</td><td>0.79 ± 0.01</td><td>1.19 ± 0.03</td><td>1.33 ± 0.08</td></tr><tr><td><i>C. composti</i></td><td>0.80 ± 0.08</td><td>0.90 ± 0.04</td><td>1.44 ± 0.04</td><td>1.55 ± 0.02</td></tr><tr><td rowspan="2">IC protein</td><td><i>P. norimbergensis</i></td><td>4.70 ± 0.07</td><td>4.94 ± 0.16</td><td>4.36 ± 0.02</td><td>4.41 ± 0.06</td></tr><tr><td><i>C. composti</i></td><td>4.60 ± 0.04</td><td>4.73 ± 0.02</td><td>4.15 ± 0.02</td><td>4.23 ± 0.02</td></tr></tbody></table>

Data represented mean ± standard deviation of OD 600 nm and protein concentration determined in triplicate cultures. The protein concentrations were calculated as nitrogen concentration*6,25 and expressed as mg protein/ml EC or IC.

estimated generation time was around 16.37 ± 5.96 h. The growth was also followed by a very pronounced decline phase after day 6 confirming the similar phenomenon observed in low volume culture (Supplementary material 2). Protein concentrations in IC and EC compartments significantly increased between day 3 and day 6. When comparing the two bacteria whatever the day considered, higher levels of EC proteins were obtained for *C. composti* compared to *P. norimbergensis* and the opposite trends were observed for IC proteins. Finally, EC protein concentrations were higher in Protobind cultures compared to Kraft lignin cultures.

## 3.2. Microscopic observation

Microscopic analyses were performed at the end of the cultivation (day 8). In contrast to the control media, light microscopic analyses of fresh bacterial culture allow the observation of rod shaped and motile cells (Figs. 2 -3).

Light microscopy observation of Kraft lignin in control media (without bacteria) show some globular structures (Fig. 2a). After bacterial growth, these few globular structures observed were not more visible and bacteria are isolated, in pairs, or grouped together in case of

*C. composti* (Fig. 2d, h). In the aim to get more details on the lignin particles and their surface, the bacterial cultures and the control media were observed by SEM. Fig. 2b, c showed the electron microscopy pictures of Kraft lignin control and indicate the presence of regular and smooth particles as well as some globular structures. After bacterial growth, very few particles were observed in the culture media (Fig. 2e, i), and they appeared rougher and irregular (Fig. 2 f, j). Moreover, at 5000 × magnification, isolated and some packed bacterial cells were clearly identified (Fig. 2 g, k see arrows). The bacterial size estimated by SEM is around 1 µm long.

Concerning Protobind lignin, after 8 days of incubation, light microscopy observation indicated the presence of numerous light-refractive granules in the control media (Fig. 3a). After bacterial growth, (Fig. 3d and g), these granules were still visible, they are more agglomerated forming black masses surrounded by bacteria. SEM confirmed the granular structure of Protobind lignin with various size (Fig. 3b, c). Fig. 3e, f, h, and i show bacteria free in the medium or coating the lignin granules (see arrows); the lignin granules seemed to be smaller and more porous.

<figure><img src="image_5.png" /><figcaption>**Fig. 2.** Microscopic observations of Kraft lignin after 8 days of incubation of each triplicate. Control media without bacteria (a,b,c). Lignin in presence of *P. norimbergensis* (d,e,f,g) and *C. composti* (h,i,j,k). Light microscope observations (a,d,h), scale bar = 20 µm. SEM observations at 1000 × (b,e,i, scale bar = 10 µm) and 5000 × magnification (c,f,g,j,k, scale bar = 1 µm). Black arrows indicate the position of bacteria.</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Microscopic observations of Protobind lignin after 8 days of incubation of each triplicat. Control media without bacteria (a,b,c). Lignin in presence of *P. norimbergensis* (d,e,f,) and *C. composti* (g,h,i). Light microscope observations (a,d,g), scale bar = 20 µm. SEM observations at 1000 × (b,e,h, scale bar = 10 µm) and 5000 × magnification (c,f,i, scale bar = 1 µm). Black arrows indicate the position of bacteria.</figcaption></figure>

### 3.3. Peroxidase and laccase activities

Total phenol-oxidase (POT) activities, laccase and peroxidase activities were quantified in the extracellular and intracellular fractions after day 3 and day 6 of growth on Kraft and Protobind lignins (Fig. 4). Overall, activities were detected in all compartments. IC POT (Fig. 4b, d) were higher than EC POT (Fig. 4a, c) whatever the considered bacteria or lignins.

On Kraft lignin cultures (Fig. 4a, b), *P. norimbergensis* displayed an EC POT activity of 42.36 ± 18.85 mIU/mg of proteins at day 3 constituted by a similar amount of laccase and peroxidase. Similar trends were observed at day 6. IC POT activity represented 27.20 ± 6.27 mIU/mg of proteins at 3 days and did not significantly increase after 6 days of incubation. In the case of *C. composti*, EC POT activity represented 64.78 ± 2.78 mIU/mg of proteins at day 3. Although the laccase proportion appeared higher (49.45 ± 15.63 mIU/mg of proteins), the difference with peroxidase activity (15.33 ± 13.30 mIU/mg of proteins), is not statistically significant. At day 6 a sharp rise in peroxidase activity (9-fold higher compared to day 3) led to a significant 3-fold increase in EC POT activity. The peroxidase activity was also 3.4-fold higher than laccase activity. *C. composti* produced around 581.08 ± 17.25 mIU/mg of proteins POT intracellularly at day 3, as mainly peroxidase activity. Both laccase and peroxidase activities increased between 3 and 6 days resulting in a statistically significant higher IC POT at day 6. Globally, except for EC POT at day 3, EC and IC POT activities determined in Kraft cultures were significantly higher for *C. composti* cultures compared to *P. norimbergensis*.

On Protobind lignin (Fig. 4c, d), after 3 days of cultivation, EC POT produced by *P. norimbergensis* was 40.39 ± 4.99 mIU/mg of proteins and did not vary significantly after day 6. IC POT activity reached a level of 20.52 ± 3.30 mIU/mg of proteins at day 3 and was ~ 3-fold higher at

day 6 with a significant increase of peroxidase. Concerning *C. composti*, EC POT activity represented 66.60 ± 7.20 mIU/mg of proteins at day 3. The laccase and peroxidase activities were similar during cultivation. IC POT activities produced by *C. composti* was higher than EC POT and represented 126.47 ± 11.43 mIU/mg of proteins. This level did not change during cultivation. IC peroxidase activity (92.59 ± 10.22 and 99.89 ± 5.23 mIU/mg of proteins at 3 and 6 days respectively) remained significantly higher than laccase.

### 3.4. Characterization of phenolic monomers

HPLC analysis of the phenolic monomers was performed at days 3 and 6 in both lignin cultures and control media (Fig. 5). In the control medium with Kraft lignin, the monomers identified were only vanillic acid (VA), vanillin (V), and acetovanillone (AV). In Protobind control medium, different phenolic monomers were released. In addition to vanillic acid (VA) and vanillin (V), syringic acid (SA), syringaldéhyde (S), p-coumaric acid (pCA), acetosyringone (AS) and ferulic acids (FA) were also detected. During Kraft and Protobind control media incubations, except for a decrease in vanillin content, the levels of the other phenolic monomers showed low variations.

Fig. 5a show the quantification of phenolic monomers in Kraft lignin cultures. With both bacteria, very few (only VA, AV and FA) and low amounts of phenolic monomers were quantified. For *P. norimbergensis*, VA and V completely disappeared at day 3. Similar trends were observed for *C. composti* with a complete disappearance of V and a 4.5-fold decrease of VA content compared with the control media (0.89 ± 0.02 mg/g of lignin in control media vs 0.19 ± 0.33 mg/g lignin. A small amount of FA was also detected in the *C. composti* Kraft lignin cultures. There was no apparent significant change in the AV quantified, whatever the conditions. Except AV, at day 6 no phenolic monomer was
<figure><img src="image_7.png" /><figcaption>Fig. 4. Phenol-oxidases (POT) activities production by *P. norimbergensis* and *C. composti* on Kraft (a, b) and Protobind (c, d) lignins after 3 and 6 days of cultivation at 30°C and pH 7. EC (a, c) and IC (b, d) activities were represented. Peroxidases activities were obtained by subtracting the laccases activities from the global phenol-oxidases activities. Experiments were performed in triplicate. \*p < 0.05, \*\*p < 0.01, \*\*\*p < 0.001.</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>Fig. 5. Phenolic monomers in the EC fractions after growth of *P. norimbergensis* and *C. composti* on Kraft (a) and Protobind lignins (b). (VA) vanillic acid, (SA) syringic acid, (V) vanillin, (AV) acetovanillone, (S) syringaldehyde, (pCA) p-coumaric acid, (AS) acetosyringone, and (FA) ferulic acid. Control corresponds to lignin culture medium without bacteria. Data represented mean ± standard deviation of phenolic content determined in triplicate cultures and controls. \*p < 0.05, \*\*p < 0.01, \*\*\*p < 0.001 compared to the control.</figcaption></figure>

quantified in the bacterial culture media.

On Protobind lignin, most of phenols present in the control media (VA, SA, V, S, p-CA and FA) were not detected after 3 days of *P. norimbergensis* cultivation (Fig. 5b). AS decreased in the control media (4.39 ± 1.02 mg/g of lignin) as compared to Protobind cultures (2.81 ± 0.20 mg/g of lignin). In *C. composti* cultures with Protobind lignin, the evolution of phenolic monomers was variable. V was not detected during the growth whereas higher amounts of VA and SA were obtained after cultivations. Compared to control media, VA contents significantly increased by 2.9-fold and 2 while SA contents increased 1.7-fold and 2.9-fold for day 3 and day 6 respectively. S which was not detected in the control media represented 2.03 ± 0.01 mg/g of lignin after 3 days of cultivation

and detected at trace levels after 6 days. The amounts of AS and FA were identical to those in the control media and did not change between the third and sixth days. Finally, a decrease of 1.4-fold for p-CA at day 3 was observed until complete disappearance at day 6.

### 3.5. Characterization of lignin degradation products

The lignin degradation products present in EC fractions of control media and bacterial cultures at day 3 and day 6 were analysed by a non-targeted metabolomics approach using UHPLC-ESI-HRMS in negative ionization mode. Approximately 300 and 900 ions were detected in the presence of Kraft and Protobind lignins, respectively. Analyses of lignin
degradation products by principal component analyses (PCA) were carried out by lignin type (Fig. 6). In both PCA, PC1 and PC2 allows the partition of samples corresponding to control, *P. norimbergensis* and *C. composti* (Fig. 6a, c). The score plot shows three clusters corresponding to the control and each bacterial culture. Regarding PC1 axis, the separation between *P. norimbergensis* and *C. composti* seems greater on Kraft lignin (PC1 = 53 %) than on Protobind lignin (PC1 = 36 %). The loading plot allowed the identifications of 29 discriminant ions (summarized in Supplementary material 3) which showed a difference in their projection on PC1 and PC2 according to the lignin types and the bacterial growths (Fig. 6b, d). These ions, detected as aromatic compounds were putatively annotated and classified as lignin monomers (M), dimers (D) and trimers (T) according to their molecular formula. Among the 29 discriminant variables, 7 products seem to be monomers of lignin, but only 3 were clearly identified (4-hydroxybenzaldehyde (M3), syringic acid M5 and ferulic acid (M7)) with the commercial standards. The 4 remaining monomers (M1, M2, M4 and M6) were detected in non-targeted UHPLC-ESI-HRMS approach but not quantified in the targeted HPLC-UV method (3.4. Characterization of phenolic monomers in EC fractions) due to their low response. These 4

compounds seem to be derivatives of the previous monomers with the presence of more oxygens in their elemental compositions which could be attributed to hydroxylation, carboxylation reactions. These modifications could explain the better response observed in negative ESI ionization mode compared to the other 3 monomers.

Unlike the HPLC-UV quantification method targeting the 9 phenolic monomers, only 6 monomers 4-hydroxybenzaldehyde (M3), SA (M5), FA (M7), VA, AS and V were detected by the non-targeted metabolomic approach. Their identity was clearly confirmed by comparison with the commercial standards. Among the 6 monomers described above, 3 monomers (M3, M5, M7) belong to the group of 29 discriminant ions and the other 3 (VA, AS and V) are present in the data matrix but not discriminant. The last 3 monomers (S, AV, *p*-CA, among the 9 phenolic monomers quantified by HPLC-UV) were not kept in the data matrix because they were under the intensity threshold. It should be noted that the MS response (negative ionization mode) of these phenolic monomers seems to be less efficient than in HPLC-UV detection.

To better identify evolution of the various aromatic metabolites hierarchical clustering analyses were performed using the fold change between metabolites levels in presence of bacteria and controls (Fig. 7).

<figure><img src="image_9.png" /><figcaption>**Fig. 6.** PCA analysis of metabolites present in EC fractions after growth of *P. norimbergensis* and *C. composti* on Kraft or Protobind lignins. (a) score plot Kraft lignin, (b) loading plot Kraft lignin, (c) score plot Protobind lignin, (d) loading plot Protobind lignin. Numbers in b and d correspond to the ions numbers given in **Supplementary material 3**.</figcaption></figure>
<figure><img src="image_10.png" /><figcaption>**Fig. 7.** Heatmap of discriminating metabolites (Supplementary material 3) in EC fractions after growth of *P. norimbergensis* and *C. composti* on Kraft (a) and Protobind (b) lignins. Natural logarithm of the peak area means (area/ml) for each condition and for each metabolite are expressed per one microliter injected of the solution. Colors in the heatmap are related to the sequential logarithmic scale presented in the right panel. Metabolites detected in EC fractions of both lignins are bold and marked by an asterisk (*).</figcaption></figure>

Three and four clusters for Kraft lignin heatmap and Protobind lignin heatmap respectively are characteristic (HCA inertia gain > 1) represented by a spacing in the Fig. 7. Four aromatic dimers (D10, D8, D9, D17) were detected on both lignins EC fractions. Hierarchical clustering and fold change (bacterial treatments/controls) over time showed a global decrease of components D9 and D17 whatever the bacterial treatments and the lignins considered, unlike compounds D8 and D10 which show a slight increase. The detection of compounds D8 and D10 were affected by the bacterial treatments, the incubation time and the type of lignin. On Kraft lignin, D10 (in the presence of *C. composti* only) increased from 3 days of growth whereas the detection level of D8 remained similar to the controls whatever the bacteria considered.

Ten other metabolites (mainly dimers) were present only in Kraft lignin condition, (Fig. 7a). The dimers D12 (annotated as G-8-O-4-G), D13, D15, D19, D20 globally increased in comparison with controls and particularly in the presence of *C. composti*. An accumulation of M2 monomer was observed in *P. norimbergensis* from 3 days of growth. The increase of dimers (D12, D20, D13, D19, D15) also varied with incubation time and depending on the bacteria. Thus, for example D12 level increased at day 3 in presence of *C. composti* then after decreased at day 6. The detection level of the dimers D25, D14, D22 and monomer M4 decreased, with a maximum drop at day 6 for both bacteria.

On Protobind EC fractions (Fig. 7b), 15 others aromatic compounds were detected and represented by monomers (including M3 4-hydroxybenzaldehyde, M5 syringic acid, M7 ferulic acid) and dimers. Moreover, several trimers (T26, T27, T28, T29) were found specifically on this lignin. Except syringic acid (M5) which increased drastically in presence of *C. composti*, all the other monomers decreased in presence of both

bacteria. The M3 monomer completely disappeared with both bacteria. Monomers M1, M5, M6 and M7 disappeared completely in presence of *P. norimbergensis*. The dimers D10, D11, D21, D23 and 24, were accumulated over time in the presence of both bacteria. A very high accumulation at day 6 was noticed for D18. The D16 dimer tends to accumulate in presence of *P. norimbergensis* and decrease in presence of *C. composti*. Finally, trimers T26, T27, T28, T29 are generally accumulated and particularly T27 and T29 on day 6 with *C. composti*.

### 3.6. Characterization of lignin after bacterial growth

At the end of cultivation (8 days), Kraft lignins in EC fractions of control media and bacterial cultures, were recovered, freeze-dried and were subjected to FTIR analyses. Thus, vibrations bands in the FTIR spectra of Kraft lignin cannot be attributed solely to the lignin, but also to components of the culture medium and to phenol monomers or dimers (Fig. 8a). However, bands characteristics of Kraft lignin (1600 cm⁻¹, 1505 cm⁻¹, 1462 cm⁻¹, 1452 cm⁻¹ and 1425 cm⁻¹, 1325 cm⁻¹) could be identified by comparing the spectra of culture media without lignin (Supplementary material 4). The intensity of the bands at 1600 cm⁻¹ (C=O stretching in conjugated p-substituted aryl ketone), 1505 cm⁻¹ (C=C aromatic skeletal vibration), and 1452 cm⁻¹ (C-H bending form methoxyl) decreased similarly and significantly after cultivation of both bacteria as compared to control.

On Protobind lignin, both residues and EC fractions in control media and bacterial cultures were analysed. As for Kraft EC, the spectra of the Protobind control media could be partly attributed to culture media. However, some vibration bands appeared characteristic of lignin
<figure><img src="image_11.png" /><figcaption>**Fig. 8.** FTIR spectra after 8 days incubation of Kraft lignin (a), Protobind EC fraction (b) and Protobind residue (c) with or without *P. norimbergensis* and *C. composti*.</figcaption></figure>

(1600 cm⁻¹, 1515 cm⁻¹, 1462 cm⁻¹, 1452 cm⁻¹ and 1425 cm⁻¹). Intensity of the bands at 1600 cm⁻¹, 1505 cm⁻¹ and 1425 cm⁻¹ (aromatic skeletal vibrations combined with methoxyl plane deformation) decreased similarly in EC fractions with both bacteria (Fig. 8b). Infrared analysis of the Protobind residue recovered after bacterial cultures (Fig. 8c) showed a few changes after growth of *P. norimbergensis* compared with control residues. Intensity of the bands around 1650 cm⁻¹ (C=O stretching in conjugated p-substituted aryl ketone), 1600 cm⁻¹ and 1325 cm⁻¹ (aromatic ring breathing with C-O stretching) was significantly lower as compared to the control (p-value < 0.010). For *C. composti*, the infrared spectrum of Protobind residue was clearly different from the spectra of the control without bacteria. The intensity at 1650 cm⁻¹ increased while intensity decreased significantly in the spectral region between 1460 cm⁻¹ and 1100 cm⁻¹, such as 1460 cm⁻¹ and 1427 cm⁻¹ (aromatic skeletal vibrations combined with methoxyl plane deformation), 1370 cm⁻¹ (deformation of the C-H of an aliphatic or phenolic alcohol), 1325 cm⁻¹ (phenolic OH), 1267 cm⁻¹ (aromatic ring with C-O stretch), 1215 cm⁻¹ (C–C plus C–O plus C=O stretch) and 1115 cm⁻¹ (asymmetric in-phase ring stretching, C–C and C–O stretching).

## 4. Discussion

Lignins are renewable sources of molecules and materials, but their complex structures make them recalcitrant to the action of biocatalysts (microorganisms and enzymes). Developing an efficient biological conversion route is one way of upgrading these polymers. Bacterial conversion of lignins occurred through two steps: the extracellular depolymerization of the polymeric lignins into low molecular weight aromatic compounds and the intracellular degradation (mineralization) of the lignin derivative molecules via various funnelling pathways [31]. Several strains of *Pandorea* and *Comamonas* have the ability to utilize and degrade lignins, and their functioning in the presence of lignin-derived compounds has been demonstrated. These studies, like most studies on bacterial conversion of lignin in the literature, are performed with Kraft lignin [11,18–20,32–35].

Because the structure of lignins change with the botanical origin of plant and with the extraction process used, it can induce different bacterial behaviours during lignin bioconversion [20]. Here, the behaviours of *Pandorea* *norimbergensis* and *Comamonas* *composti* on different technical lignins, Kraft and Protobind lignins were studied. Kraft lignin,
derived from softwood is rich in G units [31] while Protobind lignin, extracted from a mixture of grasses and extracted using an alkaline process contains a mixture of G, S and H units and some phenolic acids [6]. The multiple approaches used show the abilities of the two bacteria to grow on the two lignins by producing phenol-oxidase (peroxidase and laccase) activities intracellularly and extracellularly. Extracellular enzymes production and cell growth are tightly linked in lignin utilizing bacteria. These traits (specifically peroxidase activities production) are correlated to lignin degradation and this correlation is bacterial genus specific [36]. In our study, enzymes productions and growth profiles are different between the bacteria and according to the lignins. In Kraft lignin, the growth profiles follow comparable profiles in both bacteria and are similar to bacterial growths described earlier in the literatures [18,19,37,38]. The bacteria are not inhibited by the phenolic molecules (VA, AV, V) derived from Kraft lignin present in the culture medium or by the polymeric substrate itself confirming their ability to use Kraft lignin. The levels of extracellular peroxidase and laccase activities during growth are hardly comparable with literature data because of various experimental conditions used that induced different activity levels. However, an increased level of peroxidase during growth is accompanied by modifications in the aspect and structure of Kraft lignin (C=C aromatic skeletal, C-H bending form methoxyl vibrations bands...). Extracellular peroxidases are known as the main enzymes used by bacteria for lignin degradation. Several bacterial peroxidases are capable of various C-O bonds cleavage within lignins and bacterial secretome rich in peroxidase allow an extensive rate of lignin degradation [39-42]. The results confirm the abilities of the bacteria to use and to degrade kraft lignin.

To our knowledge, except studies involving the production of specific metabolites by engineered bacterial strains or bacterial enzymes [43,44], it is the first investigation dealing with conversion of Protobind lignin by bacteria. This study gives new insights into the abilities of bacteria to use lignin other than Kraft. Protobind lignin induces a different growth profile with a marked adaptation phase for both bacteria (presence of a lag phase for *C. composti* and diauxic growth for *P. norimbergensis*). Unlike Kraft lignin media, numerous phenolic monomers (VA, SA, AS, *p*-CA and FA) derived from G, H and S units were found in the Protobind culture medium. The presence of so many molecules can have two consequences: 1) either a rapid beginning of the growth as ligninolytic bacteria are capable of using these monomers directly [45] or 2) the complexity of these monomers and their amounts can prevent growth from starting. Indeed, ligninolytic bacteria use monomers derived from lignins depending on their intracellular metabolism [46]. The syringyl lignin-based derivatives, containing two methoxy would be more difficult to be degraded than guaiacyl and hydroxy phenyl derivatives [47,48]. In addition, some phenolic compounds and the polymeric lignin itself can have an inhibitory effect on the bacterial growth [49]. In all cases, the two bacteria therefore took some time to alleviate the inhibition and to activate efficient utilization of the Protobind lignin. After the adaptation phase, extracellular phenol-oxidase activities reached its maximal level and may be responsible of lignin utilization and the growth observed. The growth and the phenol-oxidase production observed are accompanied by the change in lignin morphology, clearly visible in SEM, and by the structural change of the Protobind lignin that show the abilities of the two bacteria to use and degrade this lignin.

During the bacterial growths on both lignins, levels of the different categories of enzymes activities produced change according to the lignins and to the bacteria. These behaviours have been observed in lignocellulolytic bacteria where the nature of the substrate has major impact on the lignocellulolytic enzymes production, and the enzymatic cocktails produced are well adapted to the biomass to process [16,50]. Furthermore, the two bacteria have higher intracellular phenol-oxidase activities than extracellular activities as observed in other species of *Comamonas* and *Pandoraea* and in other ligninolytic bacteria [19,20]. It is well known that after extracellular depolymerisation of lignins,

aromatic compounds are released and absorbed by the cells. This activates numerous intracellular oxidoreductases that are responsible for the intracellular catabolism of aromatic compounds derived from lignin [31].

The identification of aromatic /phenolic compounds derived from lignins in the extracellular medium can show the ability of ligninolytic microorganisms to effectively depolymerize these polymers. Thus, the literature is very rich in GC-MS analyses-based approaches which focused on the features of low molecular lignin aromatic compounds but mainly monomers [19,20,51]. Non-targeted analyses by using UHPLC-ESI-HRMS can give a fingerprint of phenolic compounds whose oligomers are difficult to identify using other approaches. The fingerprint analyses allowed the identification for 29 discriminant molecules (monomers and also oligomers) that can reflect the abilities of the two bacteria to use lignins and their distinct behaviour depending on the type of lignin and the bacteria. The targeted HPLC-UV identification of some phenolic monomers has completed the fingerprints obtained.

On Kraft lignin cultures with both bacteria, the fingerprint of phenolic molecules is more complex and shows the presence of numerous monomers and oligomers. Among the monomers annotated (including the phenolic monomers identified by HPLC), several have been previously identified by GC-MS during the bioconversion of Kraft lignin [18-20]. Several monomers disappeared confirming the abilities of these bacteria to use phenolic monomers as extensively demonstrated [32,44,52]. This can support the rapid growth observed on Kraft lignin.

This study showed for the first time an overview of phenolic oligomers and their evolution during Kraft lignin bioconversion. Numerous dimers are detected from Kraft cultures. Among them, D12, D14, D17, D20 and D22 have been annotated in several studies on Kraft lignin [23, 53,54]. D12 corresponded to G(8-O-4)G identified after Kraft lignin chemical treatment [53]. Several ones (D20, D13, D10, D15) have accumulated over time; while other decreased (D12 and D22 in *C. composti* or D 19, D17, D25 in *P. norimbergensis*), and some of them have almost completely disappeared from the extracellular medium (D9, D14, D22 in both bacteria). These molecules have been probably consumed /used by both bacteria. The mechanisms by which these molecules are used remain to be determined. It is possible that the dimers are cleaved into monomers by the action of phenol-oxidase activities produced by bacteria. Thus, the ability of bacterial peroxidases and laccases to degrade dimers (VGE and GGE) has been already demonstrated [55,56]. The monomers are then internalised by well-identified aromatic monomer transporters [31]. It is also possible that the dimers are internalised directly. Indeed, specific dimer (e.g. 5.5' dehydrodivanillate) transporters have been characterised in ligninolytic bacteria such as *Sphingobium* [57].

The growth of both bacteria, the variation in extracellular phenolic fingerprints (monomers and dimers) and the FTIR spectra are quite similar in the presence of Kraft lignin, suggesting that the same molecular mechanisms have been implemented, resulting in similar bacterial behaviours. Different ligninolytic bacteria were shown to activate the same pathways (β-oxidation pathway) to transform G-derived units into vanillic acid [31,47,58]. The G-derived oligomers and monomers released on Kraft lignin could induce the similar bacterial behaviours observed.

Concerning Protobind lignin, numerous phenolic monomers (VA, SA, AS, *p*-CA and FA) derived from G, H and S units were identified in the culture medium by HPLC and confirmed by the non-targeted fingerprint of phenolic monomers. Few studies concern bacterial degradation of lignins others than Kraft. After bacterial conversion of milled wood lignin extracted from corncob (grass as Protobind) a total of 20 monomers with G-S- H derived structures were identified in the culture media [20]. A group constituted solely of monomers identified by HPLC / UHPLC (including p- hydroxybenzaldehyde M3, Syringic acid M5, Ferulic acid M7 and V, VA, pCA) and dimers identified in the oligomeric fingerprints (e.g. D9 and D17, the probably derived G-units already found in the extracellular medium on Kraft cultures), disappeared or
decreased over time. It confirms the abilities of the two bacteria to use G-units as on Kraft lignin cultures. Moreover, the two bacteria are capable to activate simultaneously S and H units funneling pathways allowing them to utilize these phenolic compounds produced specifically during Protobind processing. This phenomenon was particularly marked in *P. norimbergensis*. Finally, this study shows for the first time the specific accumulation in the culture media of dimer (mainly D18) and trimers during Protobind lignin utilization whatever the bacteria. These molecules are likely the result of phenol-oxidase activity produced by the bacteria, which are capable of cleaving specific bonds within Protobind lignin. *C. composti* showed a greater accumulation of trimers T27 and T29, with a very marked change over time highlighting once again a difference in behaviour between the two bacteria. Further identification of these oligomers would provide a better understanding of the mechanisms of lignin degradation by the two bacteria, and make it possible the identification of potential phenolic compounds as platform molecules for green chemistry.

## 5. Conclusion

The aim of this study was to investigate the behaviours of two ligninolytic bacteria on different technical lignins. The two bacteria used exploit quickly Kraft lignin, and this latter seems to induce more common traits and behaviours. The utilization of Protobind lignin seems more complex. After adaptation, the bacteria are able to feed on this lignin for a limited time before declining, probably due to their inability to completely transform it. The two bacteria apparently use specific strategies, which might be due to variation in their ability to use mixed G and S units (Protobind lignin) in contrast to only G unit (Kraft lignin). Degradation of lignins was accompanied by phenol-oxidase activities production which are likely responsible of the structural modification of the lignins observed and of the liberation of phenolic monomers and oligomers. Fingerprinting of extracellular phenolic components mainly oligomers can be extended to other studies on ligninolytic bacteria. Identification of these oligomers can provide a better understanding of the behaviour of bacteria according to lignins.

## Funding

This work was carried out within the framework of the EVADELIM project supported by the SFR Condorcet and the University of Reims Champagne Ardenne. We thank the Grand Reims, for the financial support of the Chaire AFERE.

## CRediT authorship contribution statement

**David Mathiron:** Writing – review & editing, Methodology, Investigation, Data curation.
**Fontaine Jean-Xavier:** Writing – review & editing, Writing – original draft, Methodology, Investigation, Formal analysis, Data curation.
**Molinie Roland:** Writing – review & editing, Writing – original draft, Methodology, Investigation, Formal analysis, Data curation.
**David Crônier:** Methodology, Investigation.
**Corinne Ivaldi:** Writing – review & editing, Writing – original draft, Project administration, Investigation, Funding acquisition, Formal analysis, Data curation, Conceptualization.
**Brigitte Chabbert:** Writing – review & editing, Writing – original draft, Formal analysis, Data curation, Conceptualization.
**Frédéric Velard:** Writing – review & editing, Writing – original draft, Methodology, Investigation.
**Harivony Rako-toarivonina:** Writing – review & editing, Writing – original draft, Supervision, Project administration, Formal analysis, Data curation, Conceptualization.
**François Mesnard:** Writing – review & editing, Funding acquisition.

## Declaration of Competing Interest

The authors declare no competing interests.

## Acknowledgements

The authors thank Olivier Delfosse for the nitrogen quantification, François Gaudard for FTIR analysis, Serge Pilard for his involvement in the analysis of metabolites by UHPLC-ESI-HRMS.

## Appendix A. Supporting information

Supplementary data associated with this article can be found in the online version at doi:10.1016/j.nbt.2025.06.004.

## Data availability

Data will be made available on request.

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