# Tracking ectopic lignification in flax stems following scarification.

Anne-Sophie Blervacq, Dmitry Galinousky, Clemence Simon, Myriam Moreau, Anne Duputié, F. Baldacci-Cresp, Cedric Lion, Christophe Biot, Simon Hawkins, Godfrey Neutelings

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## To cite this version:

Anne-Sophie Blervacq, Dmitry Galinousky, Clemence Simon, Myriam Moreau, Anne Duputié, et al. Tracking ectopic lignification in flax stems following scarification. Plant Physiology and Biochemistry, 2025, Plant Physiol Biochem, 223, pp.109806. 10.1016/j.plaphy.2025.109806. hal-05043507

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Submitted on 23 Apr 2025

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# Tracking ectopic lignification in flax stems following scarification

Anne-Sophie Blervacq<sup>a,* ID</sup>, Dmitry Galinousky<sup>a,1</sup>, Clémence Simon<sup>a ID</sup>, Myriam Moreau<sup>b ID</sup>,
Anne Duputié<sup>c</sup>, Fabien Baldacci-Cresp<sup>d ID</sup>, Cedric Lion<sup>a ID</sup>, Christophe Biot<sup>a</sup>, Simon Hawkins<sup>a</sup>,
Godfrey Neutelings<sup>a</sup>

## ARTICLE INFO

Handling Editor: Shivendra Sahi

### Keywords:

Flax

Scarification

Ectopic lignin

Chemical reporters

Raman spectroscopy

Gene expression

## ABSTRACT

When flax (*Linum usitatissimum* L.) stems are scarified, major changes occur in the organization of cell walls within the tissues that border the wound. We sought to characterize the plant's response using a variety of approaches, with a particular focus on lignin deposition within the peripheral fiber cell walls of the stem. Raman spectroscopy and imaging first showed that changes occurred in the polysaccharide matrix of the parenchyma and fiber cell walls. These changes were accompanied by rapid deposition of lignin which initially diffuses centripetally and then, once the vascular cambium was reached, propagates in a periclinal manner until 150 µm from the edges of the wounded zone. Lignin biosynthesis appears to be the result of a *de novo* activity, as demonstrated by the concomitant accumulation of transcripts corresponding to lignin biosynthesis genes. In addition, using bioorthogonal chemistry approaches, we showed that wounding had enhanced the capacity of fiber cell walls to incorporate modified lignin precursors, in parallel with an increase in transcripts corresponding to peroxidases in the cortical tissues. This incorporation potential was identical for the 3 different types of reporters tested. Our findings demonstrated that mechanical stress can trigger lignification, in a polarized manner within the bast fibers, providing insights into the plasticity of cell wall composition and the potential for modulating fiber properties in flax.

## 1. Introduction

In flax, but also in some other plant species, bast fibers, located in the outer part of the stems, contain a thick gelatinous layer (G-layer), similar to the one found in the fibers of tension wood (Mellerowicz and Gorshkova, 2012). It is attached to the external S1 & S2 layers and faces the lumen of the cell. The most remarkable features in the composition of this layer are the presence of high amounts of cellulose, mostly present in a crystalline form, and very low amounts of xylan and lignin, making them different from the secondary cell wall of xylem cells. In fact, the amount of lignin in bast fibers has a major impact on their physical and chemical properties. This is determinant in the choice of the species when it comes to using their fibers for commercial textiles or bio-based composite materials. When considering different plant species, it is possible to obtain bast fibers with various lignin amounts

ranging from 1 % in ramie (*Boehmeria nivea*) up to 13 % in jute (*Corchorus capsularis* (Río et al., 2009)), and in kenaf (*Hibiscus cannabi-nus*, (Jin et al., 2012)). These very low amounts contribute to a better control of the subsequent biological separation of fibers from the bundles during the retting process (Wróbel-Kwiatkowska et al., 2007) during which the stalks are left in the field (dew retting) or in water tanks after plant harvesting and the cell matrix polysaccharides are partially digested by microorganisms. From a more scientific point of view, flax bast fibers can be considered as models for lignin regulation since their thick secondary cell walls contain less than those in xylem cell walls. For example, the currently cultivated form of flax, *Linum usitatissimum*, contains only between 1.5 % and 4.2 % lignin in the fiber cell walls while quantities in xylem tissues can be as high as 30 % (Day et al., 2005). This so-called hypolignification process correlates with the low transcript abundance of genes involved in the production of monolignols

<footnote><sup>a</sup> Univ. Lille, CNRS, UMR 8576 - UGSF - Unité de Glycobiologie Structurale et Fonctionnelle, F-59000, Lille, France</footnote>

<footnote><sup>b</sup> Univ. Lille, CNRS, UMR 8516 - LASIRE - Laboratoire de Spectroscopie pour les Interactions, la Réactivité et l'Environnement, F-59000, Lille, France</footnote>

<footnote><sup>c</sup> Univ. Lille, CNRS, UMR 8198 - EEP - Evolution Ecologie et Paléontologie, F-59000, Lille, France</footnote>

<footnote><sup>d</sup> Manager Analytical Sciences, Exothera, Allée Centrale - Zoning de Jumet 52, B-6040, Jumet, Belgium</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: Anne-Sophie.Blervacq@univ-lille.fr (A.-S. Blervacq).</footnote>

<footnote n="1">¹ Present address: Univ. Lille, INSERM UMR 1011 - Integrated Molecular Analysis of Liver Diseases, F-59045 Lille, France.</footnote>
and also in their oxidation before polymerization (R. Huis et al., 2012). In contrast, the production of a series of flax EMS-induced *btf* (*lignified bast fibers*) mutants containing fibers with high amounts of lignin, raised questions about the mechanisms responsible for the maintenance of a low lignin status in the fiber cells of wild type plants (Chantreau et al., 2014). EMS treatments led to an increase of the amounts of bast fiber lignin up to 17 % in the mutants without any changes in the xylem. Very clearly, EMS has made it possible to remove a lock on lignification in fibers but it still remains unclear if this inhibition occurs in the biosynthesis pathway in the wild type. Whole-genome transcriptomic results indicate that ectopic lignification of flax bast fibers was linked to an increase of transcripts controlling the production of lignin precursors (Chantreau et al., 2014). The increase of transcript accumulation corresponding to genes involved in monolignol oxidation suggests that they may be responsible for lignin production within the fibers of these mutants, and that transcription factors, or in general a defense pathway, may have been targeted by the mutagen. Modification of the lignin metabolic pathway by environmental parameters is therefore an interesting area of study, as it could lead to obtain variability in the composition and therefore the properties of the bast fibers.

Plants react to environmental stressors by exhibiting phenotypic plasticity, which can encompass both anatomical and metabolic alterations (Matsunaga et al., 2021). These changes are often dependent on modifications of the plant cell wall composition. Indeed, although fundamental molecular mechanisms of cell wall development are programmed in developmental patterning, they can be modified or re-activated in cells submitted to different environmental factors (Abdulrazzak et al., 2006; Nakamura et al., 2020; Mielke and Gasperini, 2019). For example, the amounts of lignin can rise consistently in secondary cell walls, or can even appear in cells only delimited by primary cell walls. This unexpected lignification, known as ectopic lignification (Moura et al., 2010; Caño-Delgado et al., 2003) is controlled at the genomic level as it can be induced by genetic mutations (Emonet and Hay, 2022). In other cases, lignin can also be deposited in secondary walls that are almost devoid of it. This was shown in bast (sclerenchyma) fibers in flax plants after the stem was damaged by scarification (Paul-Victor et al., 2017). In this case, lignin was deposited at the site of wounding and spread to the neighboring bast fibers. The effect of incisions on the plant stem was characterized at different levels such as healing efficiency, growth traits and stem biomechanics and their possible impact on textile quality and fiber yield. However, the molecular mechanisms underlying the modifications of the stem peripheral tissues and more specifically the fiber cell walls are totally unknown. In the *btf1* EMS-mutants, the fiber cell walls are enriched in pectin, hemi-cellulose, and glycoproteins when compared with the wild type. Modifications of the cell wall polysaccharide content have already been described during ectopic stress-induced lignification (Caño-Delgado et al., 2003). In a more general context, when induced by gene mutations, the proportion or content of amorphous or crystalline cellulose and also non-cellulosic polysaccharides can be modified and the polysaccharidic matrix can be substantially altered (Abdulrazzak et al., 2006; Chai et al., 2015; Chantreau et al., 2014).

The main goal of this work was to understand the biochemical mechanisms underlying the ectopic lignification induced by wound stress. To do so, we investigated the influence of a wounding stimulus on the composition of cell walls in the scarified zone, with a particular focus on the bast fibers. We proceeded to a deep analysis of *de novo* biosynthesis of lignin by histological, spectroscopic and bioorthogonal chemistry approaches shortly after wounding.

# 2. Material and methods

## 2.1. Plant material and experimental design

Flax plants (*Linum usitatissimum* L.) were grown from seeds sown in compost (Neuhaus) and cultivated in growth chambers (Angelantoni

Life Sciences, Italy) at 22 °C with a photoperiod of 16 h/8-h day/night. Eight weeks-old plant stems were wounded at 10 cm above cotyledon insertions with a razor blade by making a 2 cm incision in the stem parallel to the axis. The fiber-rich outer tissues, as previously defined (Day et al., 2005), were then collected at 1, 2, 7 and 14 days after wounding. In parallel, the corresponding tissues were also collected in unwounded control plants.

## 2.2. Histochemistry

Wounded and control stem fragments were collected on three plants at each time point and freehand cross sections were performed with a razor blade. The presence of lignin was determined by staining with the Wiesner reaction (Clifford, 1974) consisting in a first incubation in phloroglucinol (1 % w/v in ethanol) followed by HCl 6 N to reveal lignin in red.

A second histochemical analysis was performed by dual staining of cellulose and lignin. Sections were put on a glass slide and firstly stained in a drop of Alcian Blue (0.1 % w/v in water) for 2 min, then rapidly moved with a toothpick to a drop of Safranin O (0.1 % w/v in ethanol) for 15–20 s and finally rinsed in water. The cellulosic tissues appear in blue and lignin in red. The sections were analyzed with a photonic brightfield Olympus microscope BH-2 and pictures captured with a camera coupled to a computer (Bell photonics software).

## 2.3. Raman spectra acquisition, chemical imaging and data analysis

### 2.3.1. Sample preparation

Wounded and control stem fragments were collected and rapidly embedded in polyethylene glycol 1500 (VWR, France) (Gierlinger, 2014). Stem cross sections (10 µm) were obtained with a microtome (Leica RM65) equipped with an S65 metal blade. PEG was removed by multiple water baths and sections were mounted in water. Standard glass slides and glass coverslips were used.

### 2.3.2. Raman settings

Analyses were performed on a LabRam HR-Evolution (Horiba scientific, France). For excitation, 515 nm radiation from a Cobolt laser diode with a laser power of 10 mW incident on the sample was used. The spectrophotometer has an entrance slit of 100 µm, and is equipped with a 600 lines/mm⁻¹ grating that permits to achieve a spectral resolution of 1.8 cm⁻¹ per pixel. The Raman scattered light was detected with a CCD camera operating at 200K. An Olympus x100/0.9 objective focused the laser light on the samples with a laser spot of 0.8 µm. For spatially resolved measurements, an x-y motorized stage (Merzhäuser) with a minimum possible step size of 0.1 µm was used. The maps were recorded with spatial resolution of 1 µm in both directions (x and y). For wavelength calibration, a silicium plate was used as a reference for subsequent data pre-processing. Spectra were acquired at 515 nm, with 5 min bleaching, 100 accumulation/sec, from 300 cm⁻¹ to 1800 cm⁻¹ corresponding to the plant cell wall fingerprint region (Gorzsás et al., 2011; Gierlinger, 2014).

### 2.3.3. Raman spectra treatments

Raman spectra acquisition was performed on three controls or wounded individuals. A total of n = 31 single spectra were acquired to obtain average spectra (minimum of 10 single spectra x 3 individuals). For spectra illustration, baseline correction of spectra was applied (Horiba ®, LabSpec 6). Normalization of spectra was done by equalizing the area under each spectrum. To investigate how aromatics (related to free monolignols or polymerized lignin) moved within the cellulosic secondary cell wall of one single bast fiber present in the wounded zone, individual spectra were acquired all around a fiber, layer by layer.

### 2.3.4. Raman chemical imaging

Raman chemical imaging was performed on external tissues using an
"a priori" approach, consisting in the use of published data to identify the cell wall components according to their peak wavenumber (Supplementary Table 1). To enable comparison between different chemical images, a ratio was used based on the normalization on the cellulose peak at 379 cm⁻¹.

### 2.3.5. Raman chemometrics analysis

For chemometrics, we preprocessed spectra as follows: (i) peaks <300 cm⁻¹ and >1800 cm⁻¹ were removed for all spectra, (ii) we corrected the baseline using polynomial fitting (degree 6), (iii) we normalized spectra by equalizing the area under each spectrum [TotInt procedure]. We used both supervised and unsupervised methods to analyze variability among spectra from seven categories (31 spectra for each category: day 1, day 2 and the early phase control, day 7, day 14 and their respective controls).

To visualize differences in spectra, we used Principal Component Analysis (PCA), an unsupervised method that reduces dataset dimensionality and helps detect differences or similarities (Asri et al., 2022). In this analysis, scaling was done by the square root of the standard deviation (Pareto scaling), a method that reduces the importance of large values. We also performed a sparse Partial Least Square Discriminant Analysis (sPLS-DA), a supervised method that enables to select the most discriminating features (peaks) in the dataset (Li et al., 2020; Granato et al., 2018). We evaluated its performance by a 5-fold cross-validation with 100 repeats. All these analyses were carried both with the full dataset (847 values for 210 spectra) and with a reduced dataset (40 peaks corresponding to those defined in Supplementary Table 1, for the 210 spectra), but since results were highly similar, we here present the results for the full dataset. All analyses were performed in R 4.3.1 (R Development Core Team, 2023), using R libraries ChemoSpec (Hanson, 2022) and mixOmics (Rohart et al., 2017).

## 2.4. Chemical reporter imaging

### 2.4.1. Chemical reporter strategy

Wounded and control stem fragments were collected on three individuals at each time point. Freehand cross-sections (approximately 150–250 µm thick) were made at 10 cm above the cotyledon level using a razor blade and incubated in 300 µL sterile half strength Murashige & Skoog medium (½ MS) (Murashige and Skoog, 1962) containing 10 µM alkyne-tagged coniferyl alcohol (G<sub>ALK</sub>), 10 µM azide-tagged *p*-coumaryl alcohol (H<sub>AZ</sub>) and 10 µM methylcyclopropene tagged sinapyl alcohol (S<sub>CP</sub>) for 20 h in the light at 20 °C. After incorporation, samples were washed 4 times in ½ MS and submitted to bioorthogonal triple labeling.

### 2.4.2. Bioorthogonal triple labeling

Diels-Alder reaction with inverse electronic demand (DAR<sub>inv</sub>), strain-promoted azide-alkyne cycloaddition (SPAAC) and copper-catalyzed alkyne-azide cycloaddition (CuAAC) reactions were used consecutively to label the 3 tagged reporters S<sub>CP</sub>, H<sub>AZ</sub> and G<sub>ALK</sub> respectively. Cross-sections were incubated with 1) 300 µL DAR<sub>inv</sub> labeling solution containing 5 µM tetrazine-Cy5 (λ<sub>exc</sub> 649/λ<sub>em</sub> 670 nm) in ½ MS for 1 h in the dark and then washed 4 times in ½ MS. Then the cross-sections were incubated with 2) 300 µL SPAAC labeling solution containing 5 µM Dibenzylcyclooctyne-PEG<sub>4</sub>-5/6-Carboxyrhodamine 110 (DBCO-PEG<sub>4</sub>-Rhodamine Green, λ<sub>exc</sub> 501/λ<sub>em</sub> 526 nm) in ½ MS for 1 h in the dark and then washed 4 times in ½ MS. Lastly, sections were incubated with 3) 300 µL CuAAC labeling solution containing 2.5 mM sodium ascorbate, 0.5 mM CuSO<sub>4</sub> and 5 µM 5-carboxytetramethylrhodamine-azide (azide-fluor 545, λ<sub>exc</sub> 546/λ<sub>em</sub> 565 nm) in ½ MS for 1 h in the dark. Finally, the triple-labeled cross-sections were washed twice in ½ MS, 5 min, once in MeOH 70 %, 60 min, and 4 times in ½ MS, 5 min to fully remove any excess unreacted fluorophores and minimize fluorescence background. Then sections were mounted on slides in Fluoromount-G® (Thermo-Fisher Scientific, France) mounting medium.

### 2.4.3. Imaging

Sections were observed by confocal microscopy (Nikon A1R) using x20 (NA 0.8) and x60 (NA 1.4, oil immersion) objectives with the following excitation wavelengths and adapted bandpass filters: i) DBCO-PEG<sub>4</sub>-Rhodamine Green: exc: 488 nm, em: 525/50, ii) azide fluor 545: exc: 561 nm, em: 595/50, iii) tetrazine-Cy5: exc: 561 nm, em: 700/75, iv) lignin autofluorescence exc: 405 nm, em: 450/50. To ensure specificity, spectral acquisition and unmixing was applied to separate azide fluor 545 and tetrazine-Cy5 fluorescence. Microscope settings (gain, laser intensity, pinhole size) were kept constant throughout a given set of experiments and grey levels were used in Fiji (Schindelin et al., 2012) to make the data comparable.

## 2.5. RNA extraction and RT-qPCR

### 2.5.1. Sample preparation

At each time point, wound and control samples were collected from three biological repetitions of 4–5 individual plants. Wounded samples included the 2 mm borders of the scarified outer tissues peeled from the stems. The control samples included the outer tissues of stems collected at the same position from unwounded plants. Samples were stored at –80 °C before manipulation.

### 2.5.2. RNA extraction and cDNA synthesis

RNA extraction was carried out by TRI® reagent (Molecular Research Center, USA) following the manufacturer recommendations. Teflon pestles fixed on an electric homogenizer were used for sample grinding in the TRI reagent. The RNeasy Plant Mini Kit (QIAGEN, USA) was used for RNA samples purification and samples were treated with DNA-free™ Kit (Ambion, USA) to remove the genomic DNA. Spectrophotometer DS-11 (DeNovix) allowed the determination of concentration and quality of RNA solutions. cDNA was synthesized using the iScript™ Reverse Transcription Supermix for RT-qPCR (Bio-Rad, France). Each cDNA probe was accompanied by NRT (no reverse transcriptase) controls. The cDNA samples were diluted 5 times before using it in the qPCR.

### 2.5.3. RT-qPCR

iQ SYBR Green Supermix (Bio-Rad, France) was used for qPCR. The final volume of PCR mix was 20 µl, concentration of forward and reverse primers was 0.4 µM. The NRT and NTC (no template control) accompanied RT-qPCR, each reaction was performed in two technical repetitions. The reaction conditions and the primer design were optimized for a two-step PCR program performed on the AriaMx Real-Time PCR System (Agilent). For each reaction, high specificity of amplification was corroborated by the presence of a single peak on the melting curve. All PCR reactions were performed under the following conditions: 95 °C for 3 min, 40 cycles of 10 s at 95 °C and 10 s at 60 °C. For each primer pair, a melt curve with a 0.5 °C increment was generated in order to confirm the specificity of the amplification. Primers used in the current work are given in the Supplementary Table 2. We used the Primer-BLAST online service for the primer design (https://www.ncbi.nlm.nih.gov/tools/pri mer-blast/index.cgi). The PCR efficiency was obtained in the experi ment with the serially diluted samples (Rasmussen et al., 2000) and varies for used primers from 92 to 100 %. We used a relative quantifi cation method to analyze the gene expression (Livak and Schmittgen, 2001). To calculate the normalization factor, we used the geometric mean of the Cq-value of three reference genes – *LusGAPDH*, *LusETIF1*, and *LusETIF5A* (Rudy Huis, Hawkins, and Neutelings, 2010) whose expression stability was tested by the geNorm algorithm (Vandesompele et al., 2002) implemented in [ctrlGene] R-package. Non wounded plants were used as control samples to calculate the ddCq values reflecting relative changes in the gene expression. One-way ANOVA test with Tukey multiple comparisons of means was applied to the normalized and Log2-transformed data of the gene expression at different time points.
# 3. Results

## 3.1. Cell wall remodeling in the cortical parenchyma of the wounded zone

Flax stems were incised with a razor blade through the external tissues of 8 weeks-old plants and the samples collected at 5 different time points (Fig. 1). The effect of scarification was first evaluated by monitoring cell wall modifications in cortical parenchyma cells. The visual aspect of this tissue remains largely unchanged, with the exception of the presence of small clusters of hypertrophic cells within the wounded zone (WZ) close to the border (Supplementary Fig. 1).

The cell wall remodeling of the parenchyma cells was monitored by Raman spectroscopy in the area perpendicular to the wound over a distance of up to 250 µm (Fig. 2). Due to Raman spectral resolution (1 × 1 µm), it was possible to pick and to collect spectra according to specific spatial localization. During plant reaction after injury, collapsed cells progressively appeared as a mass in which no cell wall type could be clearly identified. This is probably due to the thinness and flexibility of primary cell walls. Therefore, it was only possible to perform the analyses during the first 48h after wounding (Fig. 2). In the parenchyma, cells were delimited by a typical primary cell wall. Wounding induced modification of amorphous cellulose (379-435-620-629-686-900-916-1337 cm⁻¹) and crystalline cellulose (1376 cm⁻¹) bands that all decreased in intensity during the early stages (day 1, day 2) compared to the control. Other bands corresponding to NCP (Non-Cellulose Polysaccharides) such as xylans and xyloglucan (455-517-1117-1729 cm⁻¹) also decreased. In contrast, relative intensities were higher during the first 48h after wounding for an additional band corresponding to cellulose (1050 cm⁻¹) but also cellulose/xyloglucan (1091 cm⁻¹) and for pectin bands (807-833-1452 cm⁻¹).

Wound-induced cell wall changes were not limited to polysaccharides. Several bands corresponding to the cutin polymer such as 710 and 1200 cm⁻¹ showed small differences compared to the control while greater variation was observed for the bands at 1300 and 1720 cm⁻¹ and reached a maximum value at day 2, 877 cm⁻¹. The Raman spectra also highlighted the presence of lignin within the parenchyma cell walls. Indeed, peaks appear after 24h at 1270 and 1424 cm⁻¹. A third peak present at 1452 cm⁻¹ also shows a strong increase but since it was also present in the control, it may be linked to the presence of

phenolic molecules other than lignin. Finally, the wide band [1557-1700 cm⁻¹] including aromatics, lignin and suberin is clearly amplified between day 0 and day 2 (Fig. 2).

## 3.2. Cell wall remodeling in the bast fibers of the wounded zone

As for the parenchyma cells, the impact of the wounding was assessed on the composition of the whole bast fiber cell walls located near to the WZ borders (Fig. 3). This very thick structure is stable over time so it was possible to perform analyzes during 14 days. When the whole bast fibers-cell wall was considered, Raman spectroscopy showed quick modifications appearing 24h after wounding. At this stage, amorphous cellulose bands (379-405 cm⁻¹), and pectin bands (833-852 cm⁻¹) showed higher intensities compared to the day 2 stage. After 48h, higher intensities for XG/cellulose (1091-1459 cm⁻¹), cellulose (1150 cm⁻¹), crystalline cellulose (1376 cm⁻¹) and xylan (1117 cm⁻¹) bands were measured.

During these early stages, a large band (1557-1700 cm⁻¹) corresponding to (lignin + aromatics) appeared at day 1, but it was not well-defined and showed a very low intensity (Fig. 3). At day 2, a band indicating the presence of aryl stretching vibrations (a4, 1599 cm⁻¹) was clearly recognizable. It was accompanied by small shoulder peaks at 1620-1652 cm⁻¹ (a5-a6) assigned to coniferyl alcohol and coniferaldehyde. Characteristic peaks corresponding to guaiacyl (G-unit) and syringyl (S-unit) monolignols were also identified at 1270 cm⁻¹ (a2) and 1332 cm⁻¹ (a3), respectively. For the late stage, at day 7, lignin/XG related peaks (1452-59 cm⁻¹ due to possible shift between samples) still remain at the day 2 levels (Fig. 3). When considering the [Lignin/aromatics] a2 – a6 bands, their intensities were maximal at day 14 (for example, the intensity of a4 is nearly x10 compared to the peak at day 2). They still persisted but were lower in ratio in the whole cell wall. Only a1, located at 1206 cm⁻¹, had very low values during the whole kinetics. It has been previously proposed that this peak could correspond to ρ-hydroxyphenyl (lignin H-unit). The peak a3 (1332-37 cm⁻¹ due to possible shift between samples) corresponds to both lignin S-unit but also to cellulose, due to deformation vibrations δ(HCC) and δ(HCO). In this case, variation of the peak intensity may be related to cellulose microfibril orientation. The minimum intensity observed at day 1 could be due to this cellulose microfibril orientation as the outer tissue is rich

<figure><img src="image_3.png" /><figcaption>**Figure 1.** Schematic representation of the wounding procedure in flax stems. A: position of the scarification in the plant stem. B: transverse section of the stem showing the depth of the wound extending from the epidermis to the vascular cambium. C: time points of the external tissue sampling.</figcaption></figure>
<figure><img src="image_4.png" /></figure>

**Figure [2]**. Cortical parenchyma cell wall remodeling in the wounded zone. Spectra were acquired for day 1 (orange) and day 2 (green), compared to control day 0 (blue) to detect remodeling events in the primary cell walls. Cellulose (c), xylan/Xyloglucan/hemicelluloses (xyl/XG/h), pectins (p), lignin + aromatics (a). Cutin (cu) related chemical bonds should be considered as precursors for suberin. See Supplementary Table 1 for peak assignments. Spectra were normalized by equalizing the area under the spectra.

<figure><img src="image_5.png" /></figure>

**Figure [3]**. Cell wall reorganization in bast fibers located close to the wound zone. Spectra were acquired for day 1 (orange) and day 2 (green), day 7 (brown) and day 14 (purple) to detect cell wall remodeling events in the primary cell walls. Cellulose (c), xylan/Xyloglucan/hemicelluloses (xyl/XG/h), pectins (p), lignin + aromatics (a). Cutin (cu) related chemical bonds should be considered as precursors for suberin. See Supplementary Table 1 for peak assignment. Spectra were normalized by equalizing the area under the spectra.

in bast fibers. But additional peak intensities increasing from day 2 to day 14 could also be related to ectopic lignification of this outer tissue.

During this late stage, the 1091 cm⁻¹ peak corresponding to both XG and cellulose decreased in comparison with the early stage signal. For cellulose, this is related to C-O-C stretching and is linked to cellulose-microfibril orientation-sensitive band. But in our studies, no modification of the laser angle was experimented to detect such a modification of orientation.

The high resolution of Raman spectroscopy enabled us to search for possible modifications in the various layers of the thick fiber cell walls. Our results were then presented according to the 3 layers as previously defined by (Gorshkova et al., 2022) i.e. the compound middle lamella (CML: up to 1 µm) including the middle lamella and the primary cell wall, the (S1+S2) layers belonging to the secondary cell wall (1–4 µm) and the gelatinous layer (Gn) contiguous (1–2 µm) to the cell lumen (Fig. 4A). During the early stage after wounding, the values in the bands

corresponding to cellulose in the CML (405–435 and 488 cm⁻¹) and (S1+S2) layer (900–969 cm⁻¹) increased, as did the cellulose 1376 cm⁻¹ band in the CML as soon as day 1 (Fig. 4B). In (S1+S2) after 48h, the 1150 cm⁻¹ cellulose band decreased in the same way as the 1376 cm⁻¹ crystalline cellulose band.

As for [lignin + aromatics], intensity started to increase rapidly within these 3 layers since day 1, then intensively covered the whole cell wall.

The wide band at 1557-1700 cm⁻¹ is particularly characteristic during this response. The intensity ratio 1599/379 cm⁻¹ (a4-coniferaldehyde/amorphous cellulose) could be used as an indicator of the progressive lignification in bast fibers, and gives us some spatial information (Fig. 4C). This ratio slightly increases during the early stage between day 1 and day 2, in a similar proportion whatever the cell wall layers tested. Then this ratio greatly increases (around 4x, as the late stage occurs) between day 2 and day 7, and at day 14, it returns to
<figure><img src="image_6.png" /><figcaption>**Figure [4]**. Spatial and temporal lignification of bast fiber cell wall after wounding. Bast fibers present in the WZ (0–100 µm) were considered. **A**: Cell wall distinct layers in bast fibers (CML: compound of the middle lamella, S1+S2, or Gn), and global bast fiber cell wall. **B**: Averaged spectra were measured within the 3 cell wall layers (CML, S1+S2, and Gn). Each stage spectrum was compared to their own controls (day 0, day 7-cont, day 14-cont). **C**: Evolution of the intensity ratio [1599/379 cm⁻¹] corresponding to [a4-coniferaldehyde/amorphous cellulose] in the different layers. Ratios were calculated on the average spectra at each time point. Spectra were normalized by equalizing the area under the spectrum.</figcaption></figure>
similar ratios observed at day 2.

### 3.3. Multivariate data analyses of bast fiber remodeling

#### 3.3.1. PCA analysis

In Raman analyses, a specific chemical bond is expressed through a peak, and a peak covers several wavenumbers. As a consequence, many intensities are correlated. Principal Component Analysis (PCA) allowed to reduce the dimensionality of the spectral profiles, whilst maintaining their variability. When analyzing the data related to the bast fibers, the eigenvalues indicate that the first two components, PC1 and PC2 explained respectively, 61 % and 20 % of the complete wounding kinetics (Fig. 5A). All three controls (day 0 for the early stage and, for the late stage, day 7-control and day 14-control) were located within a unique pool, with a few exceptions for control-day 14. In this experimental context, we can conclude that plant growth during the 2 weeks of experiments did not significantly affect bast fiber spectra profiles.

PC1 allowed discrimination of controls versus injured plants (day 1, day 2, day 7, day 14), notably early stage/controls (PC1 score <0) versus late stage (positive PC1 values). PC2 allowed partial discrimination between injured plants at day 1 and day 2 (positive values of PC2) versus their control confined with [PC2, PC1] scores close to 0. PC loadings (Fig. 5B) clearly established that PC1 is supported by peaks related to [lignin + aromatics], red asterisks and helped to explain the late stage. In contrast, PC2 is mostly supported by rearrangements in amorphous and crystalline cellulose (blue asterisks), and XG/xylan (green asterisks) and better explained the early stage.

When focusing on the early stage only (Fig. 6A–B), three clear clusters appear, with PC1 axis (54 %) explaining variability observed in day 1, and PC2 (18 %) explaining variability in day 2. At day 1, positive

contributions are due to cellulose (347-405-435-488-900-999 cm⁻¹) and lignin/aromatics + hemicellulose (1570-1652-1750 cm⁻¹) bands. On the other hand, negative contributions are observed for different cellulose bands (379-1091-1150-1452 cm⁻¹), crystalline cellulose (1376 cm⁻¹) and xylan/xyloglucan (457-517-1091-1117 cm⁻¹) bands. At day 2, global contributions are similar from those observed at day 1, except for the lignin 1452 cm⁻¹ and the G-unit bands (1270 cm⁻¹) that contribute to positive contributions.

For the late stage, there was a lot of variability within each group (Fig. 6). At day 7, PC1 (83 %) explained the variability between spectra of wounded and control fibers, with G-units (1270 cm⁻¹) and lignin/aromatics + hemicellulose (1570-1750 cm⁻¹) giving major contributions (Fig. 6C–D). The same holds for day 14 (Fig. 6E–F) where PC1 (68 %) explained the differences between wounded and control bast fibers. For the late stage, ectopic lignification was characterized by significant contributions of lignin/aromatics [1270-1332-1599-1620-1652 cm⁻¹]. The cell wall remodeling was also testified by modifications of other chemical bonds related to hemicellulose [455-517-1091-1117-1452-1729 cm⁻¹], cellulose [347-379-435-488-999-1091-1150-1376-1459 cm⁻¹] and β-glucan (969 cm⁻¹).

#### 3.3.2. sPLS-DA analysis

In order to facilitate processing of the high dimension of Raman datasets and also to select relevant variables with the Least Absolute Shrinkage and Selection Operator (LASSO) approach, it is possible to use sparse Partial Least Square (sPLS) regression which also takes in account multicollinearity. sPLS for Discrimination Analysis (sPLS-DA) will further help to achieve good predictive performance (i.e. good classification of spectra) and variable selection, with a preliminary test that should conclude that the error rate is very low and combines with a reduced number of components. sPLS-DA enables the selection of the most discriminative features in the data to classify samples. This analysis could not accurately discriminate all seven groups of plants (day 1, day 2, day 7, day 14 and their respective controls). This is likely because (i) all three controls have relatively similar spectra, (ii) spectra at day 7 and day 14 are not very different (Fig. 5A). Thus, we applied the sPLS-DA to two subdatasets: i) early stage [day 1, day 2, versus day 0], and ii) late stage [pooled wounded day 7+day 14, versus pooled day 7-control and day 14-control].

For the early-stage dataset, discrimination of all three groups was almost perfect with only two components (Fig. 7A–B). The first component showed that the amorphous cellulose peak at 405 cm⁻¹ was mainly responsible for discriminating the groups, while component 2 showed contributions of cellulose/pectin [600-700 cm⁻¹], and a mix of pectin/hemicelluloses covering a wide band at 1700-1791 cm⁻¹.

For the late stage, a good discrimination of the two groups (wounded vs control) required up to six components. Using only the first two axes results in a misclassification rate of 13 %, as shown by the overlap of the ellipses on Fig. 7C. The first component was considered as relevant and was mainly driven positively by the aromatics (1599-1620 cm⁻¹), and negatively by cellulose (379-488 cm⁻¹).

To sum up, during the first two days after wounding, rearrangements of amorphous cellulose, pectins and hemicelluloses occur within the bast fiber cell and simultaneously, [monolignol/lignin polymer] production seems to be induced. During the late stage (day 7 and day 14 after wounding), lignin and aromatics are overrepresented, while [amorphous cellulose + hemicellulose] continue their rearrangements. The Raman spectral profiles led to highlight the production of aromatics compounds, and probably, the ectopic production of lignin polymers. We checked this both with histochemistry, and with "in situ" chemical imaging in the WZ.

### 3.4. Dynamics of lignification determined by histological staining

As shown by the Raman approach, one of the major events occurring after scarification is the deposition of lignin within the wounding zone

<figure><img src="image_7.png" /><figcaption>**Figure 5.** PCA on total injury kinetics. Only bast fibers spectra were considered. A: Scatter plot of PC1 vs PC2. Three controls (day 0, day 7-cont and day 14-cont, with orange, dark grey and brown dots respectively). B: PC loadings based on the reference spectrum day 0. Positive and negative contributions were identified and assigned to cellulose (blue asterisks), hemicellulose (green asterisks) and [lignin/aromatics] (red asterisks).</figcaption></figure>
<figure><img src="image_8.png" /></figure>

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

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

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

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

<figure><img src="image_13.png" /></figure>

**Figure [6]**. PCA plots and PC loadings on early and late stages of wounding kinetic. Only bast fibers spectra were considered. A, B: early stage: PCA plot for day 1, day 2 and day 0. C, D: late stage day 7 and its own control day 7-control. E, F: late stage day 14 and its own control day 14-control. Reference spectra: B (day 0), D (day 7-control), F (day 14-control). Main positive and negative contributions were identified and assigned to cellulose (blue asterisks), hemicellulose (green asterisks) and [lignin/aromatics] (red asterisks).

and in the close bast fibers. Histochemistry was performed on wounded stem hand sections (Fig. 8) with Safranin (lignin)/Alcian Blue (cellulose) staining and Wiesner reaction with phloroglucinol-HCl, which specifically reveals the cinnamaldehyde functions of S and G unit derivatives in lignin by producing a red chromogen whose intensity depends on the quantities. The unwounded control was characterized by the almost total absence of staining with both approaches including around the fiber bundles and single fibers. In contrast, secondary xylem (wood) produced by the vascular cambium exhibits red staining, proving the progressive presence of lignin in their cell walls.

During the 24 h after wounding, an ectopic lignin deposition appeared following a periclinal deposition from 0 to 100 µm around the wounded borders, and within the external tissues covering both cortical parenchyma and fibers bundles (Fig. 8). Then, a centripetal diffusion from the external tissues to the vascular cambium occurred in both borders of the wounded zone during the next 24 h. No major increase in deposition was observed between two and seven days except a slight increase in the diameter of the zone of deposition of lignin (around 40–50 µm at day 2, to 70–100 µm at day 7) that could be only noticed in the two lips bordering the wounded zone cone. No main differences in terms of periclinal diffusion were noticed between day 7 and day 14.

### 3.5. Raman chemical imaging of the ectopic fiber lignification

Raman chemical imaging was firstly done according to the intensity of each major aromatic peak (G-unit: 1270; S-unit 1332 and lignin/aromatics: 1599 cm⁻¹) (Fig. 9). The exploration of other chemical bonds related to the other cell wall polymers is presented in Supplementary Fig. 2.

Deposition of ectopic lignin covers progressively all the wounded zone borders starting by dispersed dots (day 1) that enlarged rapidly (day 2) and then joined all together (day 7). These dots consisted of concentric depositions of [aromatics + lignin] chemical bands, and higher concentrations were localized in the center of such dots. These were firstly positioned as an anticlinal thin zone all along the borders, notably for chemical bonds related to G-unit. Periclinal diffusion occurred after 48h through the cortical parenchyma (0–100 µm from the borders). Lignin related chemical bands first surrounded the fibers at day 2 then moved within their secondary cell wall at day 7 reaching the maximum intensity (Fig. 9, day 7, red pixel corresponding to 80 arbitrary units). During the second week, an increase or an accumulation of deposition is noticed in a thicker area surrounding bast fibers (note the difference in scale). The alignment of the color scales between day 7 and day 14 is not possible due to the higher levels observed in day 14.

Bast fibers were not clearly labeled and only several spots appeared at day 7. Nevertheless, when crossed vectors were plotted through the
<figure><img src="image_14.png" /></figure>

**Figure [7]**. sPLS-DA analysis of Raman spectra of the cell walls of early (A–B) and late (C–D) datasets. Only bast fibers spectra were considered. A, B: early stage (day 1, day 2 and the control day 0). Plot (A) and PC loadings for the two first components (B). C, D: late stage (in this case day 7 and day 14 were pooled, as well as their own controls day 7-cont and day 14-cont). Plot (C) and X-Comp loadings for the two first components (D). a: aromatics; c: cellulose; h: hemicellulose; p: pectin.

<figure><img src="image_15.png" /></figure>

**Figure [8]**. Evolution of ectopic lignification after scarification. Lignification induced by scarification was imaged during 14 days in fresh hand sections stained with Safranin O-Alcian Blue (SAB, A to E) and phloroglucinol-HCl (F to J). The presence of ectopic lignin is underlined by a yellow dotted line. Black double arrowhead shows outer tissues. Simple yellow arrowhead exhibits the polar diffusion. Cortical parenchyma: cp; epidermis: ep; fiber: f; vascular cambium: vc; secondary xylem: xyl. Bar = 50 µm.

bast fiber cell wall, it was possible to detect [aromatics] from the wide window [1557-1700 cm⁻¹] (Fig. 10). The chemical environment is really different in outer tissues where the ectopic lignin appeared. The cell wall remodeling occurring during this process affected two main cell types: cortical parenchyma with large cells exhibiting only a thin primary cell wall, and bast fibers, firmly grouped into bundles, stuck all together with pectin, and exhibiting a thick secondary cell wall, with sheets of cellulose.

Periclinal deposition of this ectopic lignin is observed during the

early step (day 0-day 2). During the late stage (day 7-day 14), lignification occurred in depth, i.e. centripetally between day 2 and day 7. At day 7, some bast fibers close to the wounded zone pole also show [aromatics + lignin] bonds. During the second week (day 14) the amounts of these chemical bonds linked to lignins increase in a periclinal way, but also in depth. This spatio-temporal dynamics of ectopic lignin deposition, observed through Raman chemical imaging, is in accordance with imaging obtained with histochemistry.
<figure><img src="image_16.png" /></figure>

Figure [9]. Dynamics of ectopic lignin deposition within the wounded zone (0–100 µm). For Raman chemical imaging, spectra were first normalized on the 379 cm⁻¹ cellulose peak. Then the main aromatics peaks (1270-1332-1599 cm⁻¹) related to lignin were imaged using the integration of the surface under these peaks. Each map has its own color scale of intensity. Fiber lumens are identified by asterisks. The wounded border is indicated by white dotted lines on the right side of the panel. Respective chemical mapping sizes: control: 25 µm × 40 µm, day 1: 150 µm × 150 µm, day 2: 50 µm × 125 µm; day 7: 50 µm × 125 µm; day 14: 80 µm × 80 µm.

3.6. *Polarized and gradual lignification of bast fibers*

Ectopic lignification started from the wounded borders and first
diffused in a periclinal way. Here, we investigate the reception of the
signal only at the bast fiber-cell level. We wanted to state if the signal
was: i) simultaneously received by all cell types present in the wounded
borders, at the same time, and at the same level, whatever the cell wall-
faces considered, or ii) received progressively, in a polar manner, from
the wounded borders to a distance to be determined within the WZ.

This signal, and therefore, the ectopic lignification observed as a
response, reached the first bast fibers close to these wounded borders,
whereas such a cell type is usually considered as hypolignified (Day
et al., 2005). This unexpected lignification is able to progressively
invade the cellulosic secondary-cell wall of this particular cell. Aliphatic

compounds, phenolics and aromatics could be present in the wide band
[1557-1700 cm⁻¹] (Fig. 10) (for peak assignments, see Supplementary
Table 1).

The progressive deposition of [lignin + aromatics] all around fibers
in the WZ, lets their ratios to be much higher than within fibers, in which
their ratios are lower. It is difficult to access this information with
classical histochemistry due to this high difference in ratios. It is the
reason why fibers showed no red pixels (false color) within their cell
walls whereas surrounded cells and materials showed many. The three
confocal images (Fig. 10) therefore didn't seem informative at a first
view. Nevertheless, they are the base of the Raman polar vector. Indeed,
only Raman vector analysis helps us to appreciate the relative amounts
of [lignin + aromatics] despite this difference.

After 24h, Raman profiling led us to observe [lignin + aromatics]
<figure><img src="image_17.png" /><figcaption>**Figure [10]**. Centripetal and polarized lignification of the first bast fiber reached by wounded signals applied during 7 days. Raman chemical imaging of the wide band [1557-1700 cm⁻¹]. Epidermis is localized close to the yellow line. The first bast fibers analyzed are limited by a white dotted line. A vector (white arrow) was placed from wounded border-WB (left side of the panel) to the cortex (right side). Raman profiles were obtained from this polar vector A→B (wounded border → opposite pole). A spectrum was acquired every 0,5-1 µm along this vector. The presence of [aromatics + lignin] was evaluated as the band [1557-1700 cm⁻¹] was integrated. At day 1, first signals were perceived and produced a response through a peak of chemical bond related to [lignin + aromatics] (numbers 1 & 2). Then at day 2 and day 7, other peaks still appear (numbers 3 to 6). For imaging, spectra were normalized on the 379 cm⁻¹ cellulose peak. Lu: lumen, WB: wounded border.</figcaption></figure>

chemical bonds at low intensity in the first 2.5 µm zone from the wounded borders that could correspond to the CML layer of this bast fibers-cell wall at the wounded pole. At day 2, such chemical bonds are still present mainly in the first 4 µm (number 4), from 8 to 11 µm (lu), and from 12 to 16 µm (numbers 1 & 2). Such peaks correspond to the 16 µm first zone including S1+S2 and Gn facing the WZ but also Gn then S1+S2 from the wounded borders opposite face. The major one (lu, 2400 cpm) observed at day 2 is not due to aromatics present in the cell lumen, but to contributions from surrounded lignified cell walls (from the cell layer that is beyond the fiber tested). From either side of lumen (lu, the Gn facing the opposite wounded borders (portion 10–12 µm) exhibited higher intensity than the Gn facing wounded borders (portion 6–8 µm).

After 7 days, six peaks were detected covering the first 17.5 µm bast fiber surface. Intensities of all these peaks were very high, notably for peaks number 2 (14,500 cpm) and number 5 (15,000 cpm). Globally, at the tissue level, the wounding signal firstly diffuses periclinally through the outer tissues (epidermis, cortical parenchyma and fiber bundles). At the cell level, when it reaches the first bast fibers present in the WZ, it is diffused centripetally through this bast fibers-cell wall and crosses it. Such a signal therefore diffuses in a polar way, from the wounded face to the opposite face of this cell type. The limit of Raman spectroscopy is

that it does not allow us to discriminate previous lignin from the newly produced one. We only consider flax outer tissues in this study. Due to the hypolignified status of flax bast fibers (Day et al., 2005), we can state that spatio-temporal detection of chemical bonds related to [lignin + aromatics] is induced by such an injury.

### 3.7. *De novo* ectopic lignification monitoring with chemical reporters

Spatiotemporal dynamics of lignin biosynthesis can be traced by using a triple bioorthogonal labeling strategy following protocols previously developed in our laboratory (Simon et al., 2018; Morel et al., 2022) (Supplementary Fig. 3). This approach makes it possible to monitor the capacity of cell walls to incorporate chemical reporters H<sub>AZ</sub>, G<sub>ALK</sub> and S<sub>CP</sub> which are monolignol analogs that can be oxidized by the cell wall enzymatic machinery. Unlike histochemistry detection revealing the presence of pre-existing lignin, the triple bioorthogonal labeling strategy exclusively targets lignin produced *de novo*.

The chemical reporters can be included in the growing polymer in a similar way as natural monolignols. In a second step, bioorthogonal reactions can be sequentially used to covalently link each incorporated reporter to its appropriate fluorophore through their orthogonally
reactive chemical tags. We therefore used these tools to follow the
lignification process during *de novo* lignin synthesis in wounded flax
stem cross sections collected at different time points (Fig. 11). The
specificity of the labeling was validated by experiments using natural
non-tagged *p*-hydroxycoumaryl, coniferyl and sinapyl alcohols on
wounded stems (Supplementary Fig. 4). A second control was performed
with tagged reporters on unwounded stems (Supplementary Fig. 5). As
expected, the great majority of monolignol reporter incorporation
occurred in the differentiating xylem, while no bast fibers were labeled
in the absence of injury. In wounded stems, the incorporation pattern of
the three monolignol analogs were perfectly in line with the progressive
ectopic lignification kinetics previously described with Raman and his-
tochemistry (Fig. 11). Bast fibers directly adjacent to the wounded
border started incorporating the monolignol surrogates at day 2, mainly
at the cell junctions. Then the signals progressed through the entire fiber
bundle at day 7 and day 14, even in fibers not directly adjacent to the
incision. At day 7, all the cell junctions in the bast fiber bundles showed
a positive specific fluorescent signal. The cell walls of cortical paren-
chyma near to the wounded zone also progressively integrated the three
reporters from day 2 to day 14. Because lignin exhibits autofluorescence,
it is possible to co-localize the presence of this polymer with the
chemical reporters. Lignin deposition was then visible in a discreet
manner within the bast fibers adjacent to the injury at day 1 and day 2
after wounding. At day 7 and day 14, autofluorescence made individual
fiber cell outlines clearly visible in the entire bundle. In agreement with

the chemical reporter incorporation profiles, the cell walls of the sur-
rounding parenchyma cells, known to be delimited by primary cell
walls, also exhibited an autofluorescent signal indicating that lignin was
biosynthesized at this site. Autofluorescence was also detected in the
epidermis of the wounded samples (Fig. 11) as well as in the controls
(Supplementary Figs. 4 and 5) suggesting the permanent presence of
lignin but also suberin and cutin within this tissue, as described previ-
ously (Donaldson, 2020).

To get a more detailed examination of the spatial incorporation of
each monolignol into the fiber cell walls, we performed fluorescence
intensity profiles. A vector scan was applied through the wounded zone
from the first bast fiber until the basis of the outer tissue (Fig. 12A). This
approach is relevant to perform relative quantifications of fluorescence
intensities within a single channel and comparisons of profiles between
channels, but like in most fluorescence microscopy experiments, direct
comparisons of fluorescence intensity between channels was not
possible. Within the fiber bundles, the autofluorescent lignin was first
visible in the cells adjacent to the incision at day 2 and then progressed
towards the inside of the external tissue over a distance of 100 µm at day
7 and day 14 (Fig. 12B). The intensity of autofluorescence also increased
with time, confirming the temporal accumulation of lignin after
wounding. There was a slight discrepancy between the results obtained
by autofluorescence and Raman, which suggested significant lignifica-
tion at day 2, and histochemistry which detected the process at day 1.
This was potentially due to a difference in sensitivity between both

<figure><img src="image_18.png" /></figure>

**Figure 11.** Bioorthogonal lignin triple labeling in flax wounded zone. Monolignol surrogates were incorporated in freshly cut transversal cross-sections of flax stems at different times (day 1, day 2, day 7 or day 14) after stem wounding or without wounding for the control. False colors obtained by imaging are the following: lignin autofluorescence in blue, H<sub>AZ</sub> (SPAAC/DBCO-PEG4-RhodamineGreen) in green, G<sub>ALK</sub> (CuAAC/Azide-fluor 545) in red and S<sub>CP</sub> (DAR<sub>inv</sub>/tetrazine-Cy5) in magenta. Xyl: xylem, ep: epidermis, w: wounded pole, p: parenchyma. Bast fiber bundles are surrounded by white lines. Scale bar = 50 µm.
<figure><img src="image_19.png" /><figcaption>**Figure 12.** Bioorthogonal lignin labeling, intensity profiles of reporter incorporation in wounding zone. Incorporation of monolignol reporters in freshly cut transversal cross-sections of flax stems at different times (day 1, day 2, day 7 or day 14) after stem wounding. **A:** Brightfield and merged G<sub>ALK</sub> (red), H<sub>AZ</sub> (green) and S<sub>CP</sub> (magenta) channels of the wounding zone. **B:** For each time point, intensity profiles of lignin autofluorescence and chemical reporters are measured from the vector scan (yellow arrow) perpendicular from the wounding zone (wz) to the inner part (in) of outer tissues. Scale bar = 50 µm.</figcaption></figure>
experimental approaches.

Our results showed that the spatial incorporation profiles of HAZ, GALK and S<sub>CP</sub> reporters were very similar when compared to each other. Only very low signals were detected in the first 15 µm from the border at day 1. These signals then spread along the proximal 50 µm region at day 2 and even further over a distance of 100 µm for S<sub>CP</sub> and 140 µm for HAZ and GALK at day 7. At day 14, the incorporation of the three reporters was restricted to the 100 µm zone showing that the distal zone lost its activity. In parallel, the variation of relative intensity between the three reporters showed that in the first fibers of the bundle at day 1 and day 2, the peaks corresponding to HAZ and GALK both have quite similar intensity whereas the amplitude of S<sub>CP</sub> peaks is greater during this period. At day 7, signal spread is accompanied by an increase of fluorescence intensity for both HAZ and S<sub>CP</sub>, which present similar levels. In contrast, the intensity of the GALK peaks were lower than HAZ and S<sub>CP</sub>. At day 14, the signal did not extend further compared to day 7, however GALK intensity signals caught up with that of HAZ and S<sub>CP</sub>. This overall trend of decreasing then increasing GALK intensity over time compared to HAZ and SCP was also found in other samples analyzed (Fig. 12B and Supplementary Fig. 6), but it is necessary to consider the biological variability of each injured sample. These observations suggest a preferential incorporation of monolignols mimics over time during lignification in response to injury.

Remarkably, there was a high degree of synchronicity between the graph obtained by merging the data from the three chemical reporters and the lignin autofluorescence. This indicates that the increased incorporation capacity of the fiber cell walls is responsible for the *de novo* biosynthesis of lignin in response to injury. Furthermore, the end of the lignification process correlates with a decrease in these activities.

### 3.8. Gene expression related to lignin pathway

We monitored the expression level of 17 genes during 14 days of the experiment in the outer stem tissues of wounded flax plants. We

evaluated a gene expression level on five time points: 5 h, 1 day, 2, 7 and 14 days after wounding of the flax stems. The genes selected for the experiment are involved in directly or indirectly synthesizing cell wall polymers. Six of 17 selected genes (*LusPAL1*, *Lus4CL3*, *LusCAD1*, *LusF5H2*, *LusC3'H1* and *LusCCR1*) are linked to monolignol biosynthesis (*Le Roy* et al., 2017), three other genes (*LusPRX52*, *LusPRX53* and *LusPRX71*) encoding peroxidases were chosen because of their over-expression in flax mutants with enormous lignification of bast fibers (*Chantreau* et al., 2014). We also chose a UDP glycosyltransferase gene (the *LusUGT72N1*, a homologous to *UGT72E3* in *Arabidopsis*) because of its possible involvement in the monolignol homeostasis (*Baldacci-Cresp* et al., 2020). Finally, we added three *LusIRX* genes involved in xylan biosynthesis and four cellulose synthase genes namely *LusCESA1-A*, *LusCESA3-B*, *LusCESA4* and *LusCESA8-A*. The first two cellulose synthases refer to the biosynthesis of the primary and the last two - secondary cell wall (*Chantreau* et al., 2015). Our Raman results showed that scarification can also impact cell wall polysaccharides and we included *LusIRX* and *LusCESA* genes in our study.

The ddCq approach for the transcript quantification and a two step-normalization was used in our study (Livak and Schmittgen, 2001). The first one normalized the expression level (dCq values) against three reference genes previously validated as stable housekeeping genes (Rudy Huis, Hawkins, and Neutelings, 2010), and the second against control plants that were not wounded (Supplementary Fig. 7). The ANOVA test with Tukey multiple comparisons of means evidenced that 8 genes within the 17 studied showed statistically significant (confidence interval 95 %) changes between at least 2 different time points: *LusPAL1*, *LusCCR1*, *LusF5H2*, *LusPRX52*, *LusPRX53*, *LusUGT72N1*, *LusIRX9* and *LusCESA4*.

We tried to distinguish different groups amongst these 8 genes based on their expression profiles (Fig. 13) based on two criteria: similar profile changes and the same time points for maximum and minimum of their expression level. The first group included 4 genes: *LusCCR1*, *LusF5H2*, *LusPRX52* and *LusPRX53* (Fig. 13A). The expression profile of

<figure><img src="image_20.png" /><figcaption>**Figure 13.** Profiles of 8 differentially expressed genes (ddCq-values, Log2-scale) within the wound zone. The expression profiles of 4 (A), 2 (B) and single (C and D) genes were shown. Time points are 5 h (0.2 day), day 1, day 2, day 7, and day 14 after scarification. Points on the chart are mean values of the corresponding genes at each time point.</figcaption></figure>
these genes had 2 peaks: at day 1 and day 7; the *LusPRX52* gene at day 1 showed a 45.8-times up-regulation compared to the control plants, which was a maximum value among studied genes. Interestingly, both peroxidase genes *LusPRX52* and *LusPRX53* had a very close expression pattern. *LusIRX9* and *LusCESA4* formed a second group, showing the expression minimum at 5h after wounding (approximately 10 times lower compared to the control plants) and the following expression increased until day 1. Then gene expression decreased again on day 2 and reached the values of control plants level on day 7 (Fig. 13B). Two genes *LusPAL1* and *LusUGT72N1* differed from other genes by their expression profiles. *LusPAL1* had an increased expression in the early stage between 5h and day 2 before returning to the initial level after 7 days (Fig. 13C). *LusUGT72N1* showed a strong 27.5-fold expression decrease in the early stages until day 2 followed by a maximum expression peak at day 7 (Fig. 13D).

Expression profiles for these 8 differentially expressed genes on 5 time points were processed for principal component analysis. The generated PCA-plot showed 3 groups of time-associated samples: the first of them included samples collected at 5 h and day 14 after wounding, the second one - day 1 and day 7, and the last one - day 2 (Fig. 14). In the temporal interpretation, this could mean that samples of 5 h, day 1, and day 2 differ in their transcriptional profile, and during this timeline, the expression profiles of tissues are changing. Then a trajectory of tissue development reverses and goes back on the same track: on day 7, the profiles of the wounded outer stem tissues return to a state closer to that on day 1. On day 14, it again becomes similar at 5 h after wounding.

# 4. Discussion

## 4.1. Impact of wounding on cell morphology

Plants are permanently exposed to the risk of injury due to environmental mechanical constraints or the presence of herbivores. They have integrated during their evolution process a number of constitutive and induced defense mechanisms. The presence of lesions within the

tissues not only leads to the alteration of surrounding cells but also to a breakdown in protection against pathogens. Plants are then able to identify this deterioration as an altered self and induce specific protective responses (Savatin et al., 2014). The scientific literature gives a fairly clear idea of the sequence of events occurring in the wounded tissues, but these may vary between species or organs. For example, incisions were reported to induce proliferation of parenchyma cells in *Eucalyptus* bark (Hawkins and Boudet, 1996), in potato tuber (Lulai and Neubauer, 2014), and in medullary parenchyma (pith) of *Arabidopsis* mutants (Matsuoka et al., 2021; Zhong et al., 2000). After wounding, the cell growth regulation was modified by the increased expression of cytokinin biosynthetic (*StlPT*) and catabolic (*StCKX*) genes, but also a significant increase of free IAA content as demonstrated in the potato periderm (Lulai and Neubauer, 2014). Other hormones, such as Jasmonic Acid, Abscisic Acid and Ethylene, are also *de novo* synthetized and are considered to coordinate defense response (Bouquin et al., 1997). In our experimental conditions using flax stems, we observed that scarification induced the formation of bulged and expanded cells in the stem cortical parenchyma (0–100 µm from WZ borders, Supplementary Fig. 1).

## 4.2. Modifications of the cell surfaces

When the plant undergoes environmental modifications linked to biotic or abiotic stresses, impairment of the cell wall integrity appears to be a major event occurring within the concerned tissues (Gigli-Bisceglia et al., 2020). In our work, we first used spectroscopic and imaging approaches to characterize the modifications that took place within the walls of cells neighboring the scarified zone of the stem, with a special focus on the production of phenylpropanoid-derived polymers. The expression of genes involved in the production of cell wall constituents was then assessed in an attempt to shed light on the sequence of events and possible control points that occur during the synthesis of molecules involved in protecting cell interfaces.

Cutin and suberin both have close functional and chemical similarities but in addition to the fatty acid-containing structure characterizing cutin, suberin contains aromatic moieties synthesized via the general phenylpropanoid pathway (Nawrath, 2002). They play essential roles in sealing off tissues against loss of water and solutes, but also in reinforcing the cell wall (Beisson et al., 2012). Suberin is deposited on the inner side of primary cell walls as in cortical parenchyma and covalent bonds anchor the aromatic domain to cell wall polysaccharides (Yan and Stark, 2000). In this context, suberin was often reported as playing a protective role in wounded tissues (Wei et al., 2021; Yaqoob et al., 2022). This was confirmed in our experimental model by a Raman approach. In the parenchyma, the variation of the 877 cm⁻¹ band clearly reflects an increase in the amount of cutin over the first 2 days. This variation is accompanied by smaller variations in the specific bands at 710 and 1300 cm⁻¹.

Production of both suberin and lignin are linked by the biosynthesis of precursors derived from the general phenylpropanoid pathway (Xin and Herburger, 2021) leading mainly to the production of hydroxycinnamic acids for the first and hydroxycinnamic alcohols for the second. Within the wounded tissues in flax, spatial distribution was clearly demonstrated with the production of the two polymers in the parenchyma, and the exclusive presence of lignin in the bast fibers. Previous reports established co-production of both polymers in incised *Eucalyptus* (Hawkins and Boudet, 1996) and in lacerated *Phaseolus vulgaris* hypocotyls by vibrational spectroscopy (Busch et al., 2010). Temporality of the appearances of lignin/suberin were also investigated in *Populus* hybrids submitted to mechanical damage (Rains et al., 2022). In this species, polyphenolics accumulated 3 days post wounding, while aliphatic suberin deposition occurs 5 days post wounding reflecting a possible temporal regulation in the distribution of phenylpropanoids. The phenylpropanoid pathway seems therefore to be a key factor in the response to abiotic environmental stresses.

<figure><img src="image_21.png" /><figcaption>**Figure 14.** A PCA plot of the flax samples collected at different time points. ddCq-values of genes having statistically significant expression changes were used for plotting. Colors indicate different time points, and brown circles mark closer samples. day-s: day after scarification.</figcaption></figure>
### 4.3. Cell wall polysaccharides rearrangements

Lignification is a process regulated at different levels including precursor production, transport and *in situ* polymerization (Wang et al., 2013) and its deposition is coordinated with the formation of other cell wall polymers. The spectral analyses and associated PCA described in our work showed that crystalline cellulose related peaks decreased in intensity, and bonds related to xylan and xyloglucan appear as significant contributors to the component 1 in sPLS-DA in the late stage. In parallel, bands corresponding to lignin were increased. We can therefore suggest that the modification of crystalline cellulose levels plays a decisive role in the transport of monolignols and subsequently lignin polymerization in and around the bast fibers. Cellulose crystallinity had also been suggested as an important factor for lignification on the basis of transcriptomic analyses carried out in the lignified bast fiber (lbf) EMS mutants containing hyperlignified fibers (Chantreau et al., 2014). A link between cellulose and lignin was already clearly demonstrated since mutations in cellulose synthase genes and the use of the cellulose biosynthesis inhibitor isoxaben were shown to activate lignin synthesis and defense responses (Caño-Delgado et al., 2003). However, in our experimental model and unlike mutants, cellulose network was already established before the wounding process so it was possibly modified by internal rearrangements together with the production of xylan and additional cellulose in the secondary cell wall as shown by the variation of the expression of *IRX9* and *CESA4* genes.

It has been shown that the spatio-temporal dynamics of the cell wall in plants, and more particularly in *Arabidopsis*, are finely regulated at the level of gene expression (Brown et al., 2005; Mitsuda et al., 2007). Master regulators include NAC domain and Homeobox HD-ZIP Class III (HD-ZIPIII) transcription factors initiating cell specification and secondary cell wall synthesis in a combinatorial process. These are upstream regulators of MYB46 and MYB83 factors which have been shown to regulate the expression of genes controlling the biosynthesis of cellulose, lignin and xylan (Ko et al., 2014). As a result, their overexpression led to ectopic deposition of secondary walls (Zhong et al., 2007). Although the flax genes encoding the major secondary cell wall components have been previously identified (Le Roy et al., 2017; Chantreau et al., 2015), the transcription factors that could potentially regulate them are not yet known. A dichotomous regulatory system identical to that of *Arabidopsis* probably exists in flax and would be consistent with the expression profiles obtained by RT-qPCR. It is thus likely that wounding increases the activity of transcription factors controlling several genes linked to cell wall metabolism.

As wounding events progressed, cell wall rearrangements concerned amorphous cellulose and pectins. Indeed, relative signal intensities increased during the first 48h in pectin-maximum intensity peaks (807-833-1452 cm⁻¹) (Figs. 2 and 3). This was also confirmed by sPLS-DA first component (Fig. 7) with a main positive contribution of chemical bonds related to pectins/cellulose [600-700 cm⁻¹]. In the global plant wounding sensitivity (e.g. not only scarification), pectins and their methylation status, are linked to cell wall alteration simultaneously with defense gene expression (immune responses) (De Lorenzo et al., 2011; Lionetti et al., 2007; Taurino et al., 2014). The combination of pectin and oligogalacturonides (OGs) could be therefore considered as a danger signal (Boller and Felix, 2009; De Lorenzo et al., 2011). Such OGs are released upon the plant cell wall partial degradation. In the context of wounding by scarification in flax, cell walls from the cells present in the WZ were destroyed, and such spectral modifications could be associated with a kind of immune reaction or plant proprioception in their environment (Moulia et al., 2021).

### 4.4. Lignin diffusion within the WZ

Spatial and temporal regulation of lignin deposition was addressed in this work by using histochemical and spectroscopic approaches. Within 24 h post-wounding, lignin deposition first appeared periclinally around

the wound borders and then, over the next 24 h, it spread in a centripetal and radial manner towards the vascular cambium. Between day 2 and day 7, deposition slightly increased, and then no significant difference was observed between day 7 and day 14. This suggested that the signal transduction pathway is polarized according to the source of the stress. Such a polarized diffusion of lignification signal towards the xylem was also previously described in *Eucalyptus* bark after incision (Hawkins and Boudet, 1996) and more recently (Matsuoka et al., 2021), also demonstrated that abiotic stress led to a diffusion of a "signal" that could reach the xylem, far from the incision zone.

Within the fiber cell wall itself, lignin deposition is also polarized. It first appeared in the middle lamella and then progressively towards the Gn layer. Interestingly, this deposition profile occurring after stress induction is similar to that already described during the cell autonomous or non-autonomous lignification process in the xylem (Voxeur et al., 2015), the difference being that, as mentioned before, this lignification process takes place within a well-structured secondary cell wall including predeposited lignin. Due to the fact that the incision was made on the oldest part of the stem and that the volume of the lumen in the fiber cells is greatly reduced at this point, it is likely that most of the fibers have lost their metabolic capacity, suggesting that lignification is rather non-autonomous in this case. This hypothesis is supported by the capacity of the neighboring parenchyma cells to produce lignin as shown by Raman chemical imaging.

### 4.5. Monolignol production and disponibility

In order to confirm the possibility of a *de novo* biosynthesis of monolignols within the wounded cortical tissues, we established the expression profile of a number of candidate genes responsible for the biosynthesis of monolignols. Phenylalanine ammonia lyase (PAL) is a key enzyme of the phenylpropanoid pathway that catalyzes the deamination of phenylalanine to trans-cinnamic acid, a precursor of many different phenylpropanoid-related components. Among the 4 paralogs of the PAL genes in flax, *LusPAL1* was previously shown to be linked to the production of lignin (Le Roy et al., 2017). After wounding, transcripts corresponding to *LusPAL1* were accumulated with a peak at day 2 before returning to the same level as the control. The early activation of this gene is consistent with the rapid synthesis of lignin during this period. Both *CINNAMOYL COENZYME A REDUCTASE* (*LusCCR1*) and *FERULATE 5-HYDROXYLASE* (*LusF5H2*) genes are involved in the specific production of monolignols in flax (Le Roy et al., 2017), the second being associated with the formation of sinapyl alcohol, which will be integrated as a S unit in the polymer. Transcript accumulation for these two genes is more progressive compared to *LusPAL1* and is maximum at day 7. The presence of the specific S-unit 1332 cm⁻¹ peak in the Raman analysis during the late phases of wound healing can therefore be linked to the accumulation of the *LusF5H2* transcripts. Our results therefore suggest that lignification of wounded areas is indeed associated with an increase in the expression of some lignin biosynthesis genes.

Among the different candidate genes tested in this work, *LusUGT72N1* (*Lus10041715* according to the Phytozome nomenclature) showed a significant expression variation at day 2. This gene is closely related to the *Arabidopsis UGT72E* family which plays a role in the glycosylation of monolignol subunits (*Lim et al., 2005*) and, for *UGT72E3*, can act on the temporal regulation of lignification (*Baldacci-Cresp et al., 2020*). Glycosylation of lignin precursors is thought to prevent their transport across the plasma membrane and allow their storage in the vacuole (Le Roy et al., 2016). The corresponding enzymes would therefore play a role in their homeostasis, thereby contributing to the regulation of lignification. The reduction in *LusUGT72N1* expression on day 2 is consistent with this hypothesis and corresponds to the moment when phenylpropanoid production increases. Metabolism at this time tends to favor the flow of precursors towards the cell wall.
## 4.6. Lignification regulation by oxidase activities

Control of lignification can not only occur at the monolignol production level but also during their activation by oxidasic activities in *muro* (Wang et al., 2013). Wounding response was previously shown to release apoplastic peroxidases in the cell wall (Minibayeva et al., 2009). In flax stems, bast fibers and parenchyma cells exhibited ectopic lignin formation, so we can hypothesize that a modification in the pool of oxidases may occur at the cell wall level. The expression profiles of 3 PRX genes, *LusPRX52*, *LusPRX53* and *LusPRX71*, which were overexpressed in the *lbf1* mutants (Chantreau et al., 2014), were followed until day 14. The expression of *LusPRX52* showed an early peak at day 1 and *LusPRX52* at day 7 suggesting that these 2 genes may play a role in ectopic lignin deposition. However, identifying genes specifically responsible for monolignol activation is technically difficult because there is considerable redundancy between those encoding these activities. A typical example is the abolition of lignification in the Casparian strip, with a quintuple *prx* mutant (Rojas-Murcia et al., 2020). For this reason, we chose to investigate the global ability of the cell walls near the wound zone (WZ) to incorporate monolignol surrogates by using bioorthogonal chemistry. These monolignol analogs can be incorporated into the lignin polymer by the cell wall machinery in the same way as their natural counterparts (Simon et al., 2018; Morel et al., 2022). This allows for the identification of *de novo* lignin production zones and highlights activities specific to each monolignol. The absence of signals within the fibers of unwounded tissues showed that they are unable to incorporate monolignols, even when supplied in high amounts. Conversely, the imaging patterns obtained by bioorthogonal chemistry in the wounded stems are in line with the lignification patterns shown by Raman spectroscopy and lignin autofluorescence, both in the parenchyma and the fibers. Interestingly, the use of a vector scan, which allows comparison of the incorporation patterns, shows no difference in spatial integration profiles between the three monolignol surrogates. Although it was not possible to compare the signal intensities between each reporter, we can deduce that the enzymatic machinery is coordinated in such a way as to be able to incorporate the three monolignols simultaneously and that the composition of the lignin at this stage is perhaps not essential, the cell favoring instead a coarse synthesis from any monolignol.

## 5. Conclusions

In conclusion, we have successfully described the major events occurring during the post-wounding period in the flax stem. Using Raman profiling, Raman chemical imaging, targeted gene expression, and chemical reporter approaches, we characterized the modifications at both the tissue and cell levels. The most notable reaction is the deposition of lignin in the cell walls adjacent to the lesion. We detailed the events related to the production of this polymer, especially in bast fibers, which are known to be hypolignified. The plant proprioception process resulted in two main stages after wounding: an initial and very rapid reaction (days 0–2), followed by a modulation of the signal until days 7 and/or 14. While ectopic lignification and cell wall remodeling in bast fibers and cortical parenchyma were expected, the appearance of large, bulging cell clusters in the wound zone suggests a possible hormonal control mechanism.

## 6. Author's contributions

A.-S.B., M.M, acquired Raman spectral datasets and prepared illustrations. A.-S.B., M.M, F. B.-C. were in charge of Raman spectroscopy results description and its discussion parts. A.D. achieved multivariate analyses of these spectral datasets. D.G., G.N. developed the molecular approach and performed lignin gene expression analysis. C.S. was involved in bioorthogonal chemistry experiments, illustrations and analysis. C.S., C.B. and C.L were involved in chemical reporters technical

design, production of tagged monolignols and in vivo chemistry. A.-S.B., S.H., G.N. conceptualized the whole project and supervised the work. S. H. was involved in funds searching for this project. All co-authors contributed to writing the whole manuscript.

## Declaration of generative AI and AI-assisted technologies in the writing process

Authors have not used generative AI and AI-assisted technologies in the writing process of this manuscript.

## 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.

## Acknowledgements

We thank the Plateformes Lilloises en Biologie et Santé (PLBS)/Tis-Bio facilities (Dr. Corentin Spriet, U-Lille, France; https://tisbio.wixsite.com/tisbio) and the FRABio research federation for providing the technical environment to achieve this work. This study was supported by grants from the Ministère de l'Enseignement Supérieur, de la Recherche et de l'Innovation (France) and by the Human Frontier Science Program (LT000552/2020-L) to C.S. The study was also financed by the French Region Hauts de France and the EU (CPER/FEDER project ALIBIOTECH) to F. B.-C. DG was granted by College de France PAUSE program (https://www.college-de-france.fr/programme-pause) and the University of Lille. Acquisition and baseline correction of Raman spectra were carried out on the vibrational spectroscopy facility of the Advanced Characterization Platform of the Chevreul Institute (FR2638).

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.plaphy.2025.109806.

## Data availability

Data will be made available on request.

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