Resolving the metabolism of monolignols and other lignin-related aromatic compounds in Xanthomonas citri

Martim, Damaris B.; Brilhante, Anna J. V. C.; Lima, Augusto R.; Paixão, Douglas A. A.; Martins-Junior, Joaquim; Kashiwagi, Fernanda M.; Wolf, Lucia D.; Costa, Mariany S.; Menezes, Fabrícia F.; Prata, Rafaela; Gazolla, Matheus C.; Aricetti, Juliana A.; Persinoti, Gabriela F.; Rocha, George J. M.; Giuseppe, Priscila O.

doi:10.1038/s41467-024-52367-6

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Published: 12 September 2024

Resolving the metabolism of monolignols and other lignin-related aromatic compounds in Xanthomonas citri

Nature Communications volume 15, Article number: 7994 (2024) Cite this article

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Abstract

Lignin, a major plant cell wall component, has an important role in plant-defense mechanisms against pathogens and is a promising renewable carbon source to produce bio-based chemicals. However, our understanding of microbial metabolism is incomplete regarding certain lignin-related compounds like p-coumaryl and sinapyl alcohols. Here, we reveal peripheral pathways for the catabolism of the three main lignin precursors (p-coumaryl, coniferyl, and sinapyl alcohols) in the plant pathogen Xanthomonas citri. Our study demonstrates all the necessary enzymatic steps for funneling these monolignols into the tricarboxylic acid cycle, concurrently uncovering aryl aldehyde reductases that likely protect the pathogen from aldehydes toxicity. It also shows that lignin-related aromatic compounds activate transcriptional responses related to chemotaxis and flagellar-dependent motility, which might play an important role during plant infection. Together our findings provide foundational knowledge to support biotechnological advances for both plant diseases treatments and conversion of lignin-derived compounds into bio-based chemicals.

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Introduction

The bioconversion of aromatic compounds has a central role in carbon cycling¹, plant-pathogen interactions^2,3,4 and detoxification of organic pollutants⁵. This process typically starts in upper pathways (also named peripheral pathways), which converge the structural diversity of aromatic compounds to fewer intermediate metabolites, which are further funneled to central carbon metabolites through a narrower range of lower pathways^6,7. During this convergent process, industrially relevant molecules are formed. This offers opportunities for engineering microbial chassis to produce chemicals from complex mixtures of aromatic compounds derived from abundant wastes such as lignin and mixed-plastics^8,9.

Hundreds of microorganisms across diverse phyla are known to have the potential to metabolize lignin-related monomers, according to the eLignin database⁷. However, the metabolic pathways of only a few species have been characterized so far^7,10. Lignin-related monomers include the three main lignin precursors (p-coumaryl, coniferyl and sinapyl alcohols) and their respective p-hydroxyphenyl (H), guaiacyl (G) and sinapyl (S) derivatives. Although pathways for the catabolism of several lignin-related monomers have been described in the literature, information on upper pathways for some of them, such as p-coumaryl alcohol, sinapyl alcohol and sinapaldehyde, is still lacking^11,12,13.

The molecular mechanisms related to the bioconversion of aromatic compounds have been mainly studied in some model organisms, such as Pseudomonas putida KT2440^14,15 and Sphingobium sp. SYK-6^16,17, besides other species from the Rhodococcus¹⁸ and Burkholderia^19,20 genera. These models have been isolated either from soil or industrial wastewater, so our understanding of how microorganisms from other ecological niches metabolize lignin-related compounds remains elusive. Moreover, how these molecules impact microbial behavior and physiology is still partially understood^21,22.

Some plant pathogens, such as Xanthomonas species, have a vast arsenal of enzymes to degrade components of the plant cell wall such as xyloglucan²³ and xylan²⁴, using the released carbohydrates as a source of carbon, energy, and stimuli²⁵. However, little is known about their capacity and molecular strategies to metabolize other compounds available in the plant cell wall, especially the phenolics related to lignin, a major plant cell wall component^26,27,28,29. One of the plant defense mechanisms against Xanthomonas infection is to increase the lignification of the plant cell wall, which implies an increased secretion of monolignols in the infection site³⁰. Therefore, monolignol degradation could be an effective way for the pathogen to inhibit lignification. However, it is still unknown whether Xanthomonas species are biochemically capable of adopting such a strategy.

In this work, by combining RNA sequencing (RNA-seq) analysis, biochemical characterization, and gene knockout studies, we investigated the metabolism of lignin-related aromatics in the model phytopathogen Xanthomonas citri subsp. citri 306 (X. citri 306). Our data revealed missing steps and complete pathways for the catabolism of the three main lignin precursors, as well as reductive metabolic pathways and efflux approaches to cope with aryl aldehyde toxicity. This study also showed that lignin-related compounds activate transcriptional responses related to chemotaxis and flagellar-dependent motility in the phytopathogen, which might have an important role during the infection of the plant host. In summary, this work provides insights into the molecular mechanisms involved in plant-pathogen interactions and adds missing pieces to the known spectrum of molecular strategies for the bioconversion of lignin-related compounds.

Results

The model plant pathogen metabolizes a diverse range of lignin-related aromatics

To investigate if X. citri 306 can grow using lignin-related aromatic compounds as the main carbon source, we performed growth assays in minimal medium supplemented with 21 aromatic monomers, representative of H, G and S units (Fig. 1). Additionally, we analyzed two complex samples of lignin-derived compounds (LDC-I and LDC-II) produced from sugarcane bagasse (Supplementary Fig. 1 and Supplementary Table 1). The main aromatic monomers detected in the LDC-I sample were p-coumarate, ferulate, 4-hydroxybenzaldehyde, vanillin, along with organic acids (acetate, formate, lactate) and sugars (arabinose, glucose) (Supplementary Table 1). In LDC-II, the most abundant molecules detected were acetate and the aromatic compounds catechol, 3-methoxy-catechol, phenol, guaiacol and pyrogallol (Supplementary Table 1).

**Fig. 1: Summary of growth conditions and analyses of aromatics toxicity and depletion.**

Under the tested conditions, only 4-hydroxybenzoate (4HBA), LDC-I and LDC-II supported the growth of X. citri 306 (Fig. 1, Fig. 2a and Supplementary Fig. 2). After 30 h of X. citri 306 growth in the LDC-I condition, we observed the total depletion of 4-hydroxybenzaldehyde, glucose, acetate, and formate (Supplementary Fig. 3). Lactate, succinate and arabinose were partially depleted (40–55%), with negligible depletion of hydroxycinnamic acids (Supplementary Fig. 3). In the LDC-II condition added by glucose (5 mmol L⁻¹), we observed the total depletion of hydroquinone and acetate, and partial depletion of 3-methoxycatechol (59%), 4-methylcatechol (40%), and catechol (11%) (Supplementary Fig. 3).

**Fig. 2: *X. citri* 306 grows using 4-hydroxybenzoate and lignin-derived compounds as carbon sources and metabolizes the three main monolignols.**

To evaluate whether the lack of growth in some conditions was due to toxicity or to insufficient carbon and energy supply, we repeated the assay supplementing the medium with glucose. Glucose supplementation allowed the growth of X. citri 306 in the presence of the model aromatic compounds, except for those displaying severe toxicity at 5 mmol L⁻¹, such as some aryl aldehydes (Fig. 1 and Supplementary Fig. 2). Overall, the presence of 5 mmol L⁻¹ of the individual aromatic compounds decreased the growth rate when compared to the medium containing only glucose (XVM2m(G)), indicating toxicity. The aryl aldehydes displayed a more toxic effect than the correspondent aryl alcohols and aryl acids (Fig. 1 and Supplementary Fig. 2).

Next, we investigated if the aromatic compounds are depleted from the XVM2m(G) medium by X. citri 306, indicating they are either modified into other metabolites or funneled to the central carbon metabolism. For this purpose, we analyzed by HPLC the medium supernatant before and after bacterial growth using two input concentrations (millimolar and/or micromolar) of aromatic compounds (Fig. 1 and Supplementary Table 2). In micromolar aryl alcohol cultivations, only the three monolignols (p-coumaryl, coniferyl, and sinapyl alcohol) were effectively depleted from the medium by X. citri 306, generating oxidized metabolites detected in the medium (Fig. 1, Fig. 2b, and Supplementary Table 2). For all the tested aldehydes, a depletion higher than 70% was observed at the millimolar condition (except for 4-hydroxybenzaldehyde, <30%) (Fig. 1 and Supplementary Table 2). Among the aryl acids tested, X. citri 306 substantially depleted only 4-hydroxybenzoate. Together, these results indicate that X. citri 306 has pathways to metabolize aromatic compounds as complex as the monolignols and effectively metabolizes a more diverse range of aryl aldehydes compared to aryl alcohols and aryl acids provided in the culture medium (Fig. 1 and Supplementary Table 2).

Lignin-related compounds induce chemotaxis and flagellar-dependent motility

To investigate how X. citri 306 responds to lignin-related compounds and identify the metabolic pathways involved in their metabolism, we performed RNA-seq studies. Six model compounds (coniferyl alcohol, 4-hydroxybenzoate, 4-hydroxybenzaldehyde, vanillin, syringaldehyde, and benzaldehyde) were selected for the RNA-seq analyses as representatives of both different molecular structures (H, G and S-type units) and different entry points in the metabolic pathways. We also included in the tested conditions three complex mixtures of aromatic compounds (LDC-I, LDC-II, and aldehydes mix) (Supplementary Table 3).

A total of 278 to 1285 differentially expressed genes (DEGs) in the aromatic-containing conditions compared to the control XVM2m(G) were identified (Supplementary Data 1), evidencing the importance of these compounds in modulating various physiological processes. X. citri 306 discerned subtle structural variations among aromatic compounds, as shown by the incomplete overlap of upregulated genes in each condition (Fig. 3). For example, 4-hydroxybenzaldehyde activated the expression of around 400 genes that were not activated by 4-hydroxybenzoate, although they share a subset of about 200 genes activated by both (Fig. 3a). This discrepancy suggests that, although very similar in terms of molecular structure, 4-hydroxybenzaldehyde and 4-hydroxybenzoate can elicit substantially distinct transcriptional responses in X. citri 306, which might be related to the high toxicity of the aldehyde. Partially overlapping responses were also observed when comparing the same type of phenolic compound with distinct degrees of methoxylation (Fig. 3b) and complex mixtures with different compositions (Fig. 3c, Supplementary Table 1).

**Fig. 3: Transcriptional responses triggered by lignin-related aromatic compounds.**

Although X. citri 306 recognizes subtle changes in the structure of lignin-related aromatic compounds, Gene Ontology (GO) enrichment analysis revealed that biological processes such as signal transduction, bacterial-type flagellum assembly, bacterial-type flagellum-dependent cell motility, and chemotaxis were enriched and upregulated in all conditions featuring lignin-related aromatics (Fig. 3d, Supplementary Fig. 4 and Data 2), except in the LDC-I and LDC-II conditions. This might be due to the low individual concentration of aromatic inducers or to the interference of non-aromatic molecules present in these mixtures (Supplementary Fig. 3).

In the conditions containing individual aromatic compounds, the upregulation of chemotaxis and flagellar genes, including cheAZY (XAC1930-32), motAB (XAC3693-94), fliC (XAC1975), flgL (XAC1976), flgG (XAC1981), and flgE (XAC1983) suggests that the sensing of lignin-related aromatics stimulates a motile state in X. citri 306 (Supplementary Data 1). This observation is consistent with the results of the co-expression analysis, where a co-expressed gene module (M1), mainly composed of genes involved in flagellar assembly and chemotaxis, was identified (Fig. 3e). Higher activity within this module became especially pronounced in the presence of 4-hydroxybenzaldehyde (4HBZ) and syringaldehyde (SYALD) while showing reduced activity under conditions involving complex lignin-derived samples (LDC-I and LDC-II), which can be due to the comparatively lower concentration of individual aromatic compounds within these samples or to signal interference from other molecules present in these mixtures (Fig. 3e and Supplementary Fig. 3). Between the downregulated processes, translation was consistently enriched in all the tested conditions, except for 4HBA (Supplementary Fig. 4 and Data 2).

The first steps of monolignols catabolism are performed by aryl alcohol and aryl aldehyde dehydrogenases

The first step of coniferyl alcohol catabolism can be catalyzed by an NAD⁺ dependent aryl alcohol dehydrogenase (ADH), generating coniferaldehyde, which is then converted to ferulate by a NAD⁺-dependent aryl aldehyde dehydrogenase (ALDH)^32,33. For p-coumaryl and sinapyl alcohols, this information is still missing, but considering their chemical similarity to coniferyl alcohol, we hypothesized that their catabolism might follow a similar pathway. Thus, to uncover the genes responsible for monolignols catabolism, we searched for ADH and ALDH genes upregulated in the presence of lignin-related compounds in X. citri 306. Based on their higher upregulation levels, genomic context, and the presence of common catalytic domains reported for dehydrogenases active on aromatics, we selected eight ADH genes and three ALDH genes for cloning, heterologous expression, and biochemical activity screening (Fig. 4).

**Fig. 4: Transcriptomic analysis and activity screening reveal novel ADH and ALDH enzymes active on aromatic compounds.**

The activity screening revealed several alcohol and aldehyde dehydrogenases active on aromatic compounds, with variable preferences in terms of substrate, co-substrate (NAD(H)/NADP(H)) and reaction direction (oxidation and/or reduction), which will be detailed in later sections (Fig. 4 and Supplementary Tables 4, 5 and 6). Reaction products were confirmed by HPLC analyses, showing the aryl alcohol dehydrogenase activity of XAC0353, the aryl aldehyde dehydrogenase activity of XAC0129, XAC0354, and XAC0882 and the aryl aldehyde reductase activity of XAC1484 and XAC3477 (Supplementary Figs. 5–7 and Supplementary Table 6).

Between the screened genes, XAC0353 and XAC0354 called our attention because they are clustered in the genome and encode enzymatic activities compatible with the first steps of monolignols catabolism (Fig. 5a). XAC0353 gene product displayed a NAD⁺-dependent alcohol dehydrogenase activity over the three main monolignols, besides cinnamyl and 4-hydroxybenzyl alcohols, whereas XAC0354 gene product showed a NAD⁺-dependent aldehyde dehydrogenase activity on H-, G- and S-type hydroxycinnamic aldehydes, suggesting a role in the second step of monolignols catabolism in X. citri 306 (Fig. 4).

**Fig. 5: XAC0353 and XAC0354 play a role in the first steps of monolignols catabolism.**

To confirm these results, we measured the specific activity of XAC0353 and XAC0354 in several aromatic substrates (Fig. 5b, c). As expected, XAC0353 converted aryl alcohols in their respective aldehydes using NAD⁺ as co-substrate, showing higher specific activity over the monolignols compared to benzyl alcohol derivatives (Fig. 5b). Accordingly, the aldehyde dehydrogenase encoded by XAC0354 transformed aryl aldehydes in their respective acids in a NAD⁺-dependent manner, showing the highest specific activity on coniferaldehyde (100%), followed by p-coumaraldehyde (51%) and sinapaldehyde (34%), compared to the other tested aromatics (Fig. 5c). Both enzymes were assayed in similar conditions and displayed similar specific activity rates over their best substrates (monolignols and the respective aldehydes), which might be advantageous to avoid the accumulation of the intermediate aldehyde metabolites during the serial action of these enzymes.

To validate the in vivo involvement of these enzymes in monolignol catabolism, we characterized individual knockout strains of XAC0353 and XAC0354 genes. Growth assays in XVM2m(G) containing 50 μmol L⁻¹ monolignols revealed that XAC0353 deletion impaired monolignols consumption and decreased the excretion of hydroxycinnamic acids, which are the second intermediate metabolites of monolignols catabolism (Fig. 5d). The excretion of the first intermediates, the aryl aldehydes, was not observed, indicating they are rapidly converted to the respective acids. Regarding sinapyl alcohol, although we did not detect its presence in the growth assays, likely due to instability issues³⁴, we noticed a lower amount of sinapate being excreted by the KO53 strain compared to the WT, which is consistent with the results observed for the other two monolignols (Fig. 5d). Of note, the KO53 strain still retained a partial capacity to convert monolignols up to hydroxycinnamic acids, indicating the existence of at least another gene encoding for an enzyme with aryl alcohol dehydrogenase activity. Together, these results show that the deletion of XAC0353 gene partially compromises the catabolism of the monolignols, leading to a lower production of hydroxycinnamic acids, whose conversion to the next metabolite might be a limiting factor in X. citri 306.

The growth assays using 50 μmol L⁻¹ of hydroxycinnamic aldehydes showed no detectable excretion of hydroxycinnamic acids by the KO54 strain (ΔXAC0354), supporting a crucial role for this gene in the conversion of these aldehydes into acids (Fig. 5e). On the other hand, a higher amount of excreted monolignols (p-coumaryl and coniferyl alcohol) was observed for the KO54 strain compared to the WT, indicating that the deletion of the XAC0354 gene triggered a metabolic shift toward reactions that reduce the aldehydes into their respective monolignols. This is consistent with the complete depletion of the hydroxycinnamic aldehydes by the WT and the KO54 strain, although by preferentially following opposite metabolic directions.

Syringaldehyde induces the catabolism of hydroxycinnamic acids

Although X. citri 306 did not utilize hydroxycinnamic acids (HCAs) when available in the growth medium (Supplementary Table 2), HPLC analysis indicated that they are produced as intermediate metabolites of monolignols catabolism, along with smaller intermediates resultant from their degradation such as 4-hydroxybenzoate and syringate (Fig. 2b). Thus, these findings suggest that X. citri 306 possesses the necessary enzymatic systems for metabolizing HCAs produced intracellularly.

Genome mining revealed a gene cluster (XAC0881-84) homologous to the hca gene cluster responsible for the degradation of p-coumarate, ferulate, and sinapate in Xanthomonas campestris pv. campestris²⁹ (Supplementary Fig. 8a). In our RNA-seq data, this cluster was upregulated in the presence of syringaldehyde, an intermediate of the sinapate catabolism (Supplementary Fig. 8b and Data 1). E. coli cells co-expressing XAC0881 and XAC0883 transformed p-coumarate, ferulate and sinapate respectively into 4-hydroxybenzaldehyde, vanillin and syringaldehyde, experimentally demonstrating the predicted activities of these genes (Fig. 6).

**Fig. 6: Hydroxycinnamic acids catabolism is encoded by the *hca* gene cluster.**

The hca gene cluster also harbors XAC0882, which encodes for an aldehyde dehydrogenase active on 4-hydroxybenzaldehyde, vanillin, and syringaldehyde, according to our activity screening results and HPLC data (Fig. 4, Fig. 6b–d and Supplementary Table 6). This activity is compatible with the third step of HCAs catabolism, corroborating the involvement of the XAC0881-84 cluster in the metabolism of HCAs in X. citri 306. Besides XAC0882, another gene (XAC0129) also encodes for an aldehyde dehydrogenase active on aldehydes derived from HCAs catabolism (Fig. 4 and Supplementary Table 6), implying that there is a functional redundancy for this metabolic step in X. citri 306.

Novel reductive pathways for aryl aldehydes detoxification

Among the putative ADHs screened in the previous section, XAC1484 and XAC3477 displayed a NADPH-dependent reductase activity, catalyzing the reduction of aryl aldehydes in their respective alcohols, being active on derivatives of both cinnamyl and benzyl aldehydes (Fig. 4, Supplementary Table 4 and Supplementary Fig. 6 and 7). Curiously, XAC0353 also displayed a similar activity, but in a NAD(P)H-dependent manner, indicating that it might also contribute to the reduction of aryl aldehydes depending on the intracellular balance of NAD⁺/NAD(P)H and aryl alcohol/aryl aldehyde substrates. Indeed, X. citri 306 growing on a mixture of aldehydes at millimolar level showed to be capable of converting most of them into aryl alcohols, which were then exported to the extracellular medium (Fig. 7a). In such condition, the genes XAC1484 and XAC3477 were upregulated (Supplementary Data 1). This transcriptional evidence, along with biochemical activity data (Fig. 4, Supplementary Table 4 and Supplementary Fig. 6 and 7), support that XAC1484 and XAC3477 play a role in converting aryl aldehydes into their respective aryl alcohols in vivo. Moreover, XAC1484 is clustered with the genes XAC1482-83-85, which encodes for a putative Resistance-Nodulation-Division (RND) multidrug efflux transporter, which might be involved in the excretion of the aryl alcohols produced by XAC1484.

**Fig. 7: Reductive approaches provide an alternative route for aryl aldehydes detoxification.**

Since two metabolic directions, reductive or oxidative, are possible to the bioconversion of aryl aldehydes in X. citri 306, we argued how the concentration of these compounds affects their metabolic fate. To address this issue, we grew X. citri 306 in minimal medium supplemented with increasing concentrations of coniferaldehyde, used as a representative compound, and quantified its depletion as well as the production of intermediate metabolites excreted after 15 h of growth (Fig. 7b). In the lowest coniferaldehyde concentration tested, only ferulate excretion was detected. Increasing the concentration of coniferaldehyde, coniferyl alcohol became the predominant excreted metabolite (Fig. 7b). Thus, these results indicate that increasing concentrations of aryl aldehydes activate reductive pathways, providing an additional mechanism to cope with toxicity, which was generally higher for aryl aldehydes than to their alcohol or acid counterparts (Fig. 1).

H and G-type compounds are metabolic funneled via protocatechuate ortho-cleavage

The final step in the funneling pathways of H- and G-type monomers forms protocatechuate (PCA) (Fig. 8a)^7,10. In the case of H-type compounds, it is known that 4-hydroxybenzoate (4HBA) is converted to PCA by the enzyme p-hydroxybenzoate hydroxylase (EC 1.14.13.2)³⁵. In X. citri 306 genome, we found a gene (XAC0356) encoding for a protein homologous to the p-hydroxybenzoate hydroxylase (PobA) from Pseudomonas aeruginosa³⁶ and from X. campestris²⁸. XAC0356 was upregulated in the 4HBA condition (Fig. 8b). Its deletion abolished X. citri 306 growth in a medium containing 4HBA as the primary carbon source. It also disrupted 4HBA consumption when the ΔpobA strain grew in a medium containing glucose and 4HBA (Fig. 8c, d). These results demonstrate that XAC0356 encodes a functional PobA and is essential for the metabolism of 4HBA in X. citri 306.

**Fig. 8: H and G-type monomers are funneled to the protocatechuate ortho-cleavage pathway.**

In the last step of the funneling pathway for G-type monomers, vanillate needs to be demethylated to form PCA^7,12. Three main types of O-demethylase systems have been reported in the literature so far: tetrahydrofolate (THF)-dependent enzymes, Rieske-type oxygenases and cytochromes P450 oxygenases^37,38. In the X. citri 306 genome, we found genes homologous to Rieske-type oxygenases (XAC0311 and XAC0363) clustered with genes encoding for putative reductases (XAC0310 and XAC0362), as well as a predicted P450 gene (XAC3170, <24% sequence identity with GcoA)³⁹. In the vanillin condition, only XAC0362-63 were upregulated (Fig. 8e). The deletion of this gene pair (but not of XAC0310-11) resulted in vanillate accumulation after 40 h of bacterial cultivation in a medium containing vanillin (Fig. 8f). Enzymatic assays showed that XAC0362-63 display vanillate-O-demethylase activity, further supporting a role for these proteins in converting vanillate into PCA (Supplementary Fig. 9).

Once formed, PCA can be cleaved in three different positions: 2,3-cleavage, 4,5-(meta-cleavage), or 3,4-(ortho-cleavage)⁷. Conditions containing H and G-type aromatic compounds induced the expression of a gene cluster (XAC0364-71) homologous to the pcaIJFHGBDC cluster previously characterized in X. campestris²⁸, suggesting that X. citri 306 conserves the same PCA 3,4-(ortho-cleavage) pathway (Fig. 8g and Supplementary Data 1).

Benzaldehyde metabolism generates dead-end products

In the RNA-seq data, we observed that benzaldehyde upregulates the gene pobA and the pcaIJFHGBDC cluster (Fig. 8g and Supplementary Data 1) and this intriguing result prompted us to investigate if its metabolism involves these genes. To test this hypothesis, we performed activity assays and gene knockout studies targeting PobA (hydroxylase) and PcaHG (the first enzyme of the PCA ortho-cleavage pathway). PobA showed no detectable activity towards benzoate (Supplementary Fig. 10a). The ΔpobA strain was similar to the WT strain in the conversion of benzaldehyde and excretion of its respective intermediate metabolites (Supplementary Fig. 10b). The ΔpcaHG strain accumulated the intermediate metabolite PCA in positive controls containing 4-hydroxybenzoate or vanillin, but the same was not observed in presence of benzaldehyde (Supplementary Fig. 10c–e). Together, these results indicate that benzaldehyde metabolism has no connection with PobA and the PCA ortho-cleavage pathway, despite its role as an inducer of pobA and pca genes expression. Since benzaldehyde was depleted from the medium by X. citri 306 (Fig. 1), and it is a substrate of aryl aldehyde reductases (XAC1484 and XAC3477) (Supplementary Table 4) and dehydrogenase (XAC0129) (Supplementary Table 6), we then hypothesized that benzaldehyde metabolism generates benzyl alcohol and benzoate as dead-end products, which was confirmed by quantitative analysis using HPLC (Supplementary Fig. 10f).

Transcriptome response to syringaldehyde reveals the presence of a complete pathway for S-type lignin monomers catabolism

In nature, the catabolism of S-type monomers can follow at least three different pathways, categorized according to the generated intermediates: gallate (I), 2-pyrone-4,6-dicarboxylate (PDC) (II), or 4-carboxy-2-hydroxy-6-methoxy-6-oxohexa-2,4-dienoate (III). In X. citri 306, we found genes (XAC0882, XAC0878, XAC4155, XAC4156, and XAC4157) that were upregulated in the presence of syringaldehyde (Fig. 9a) and are homologous to desV, desB, ligK, ligU and ligJ involved in the catabolism of syringaldehyde via gallate in Sphingobium sp. SYK-6^12,40 (Fig. 9b and Supplementary Table 8).

**Fig. 9: Identification of enzymes involved in the metabolism of S-type aromatic compounds.**

The formation of gallate as an intermediate relies on a two-step O-demethylation process, catalyzed by tetrahydrofolate-dependent O-demethylases in Sphingobium sp. SYK-6⁴¹. However, no homologous to tetrahydrofolate-dependent O-demethylases was detected within the genome of X. citri 306. Conversely, the two gene pairs XAC0362-63 and XAC0310-11, homologous to Rieske-type O-demethylases, were upregulated in presence of syringaldehyde (Fig. 9a), indicating a possible role in the O-demethylation of syringate (SYR) and 3-O-methylgallate (3OMG).

Growth assays with the knockout strains of genes XAC0310−11 (KO310-11) and XAC0362-63 (KO362-63) in a minimal medium containing syringaldehyde revealed a higher accumulation of syringate by the KO362-63 strain, but not by KO310-11 (Fig. 9c). This observation suggests a role for XAC0362-63 in syringate demethylation. Whole-cell activity tests corroborated this finding, showing the production of 3OMG from syringate by E. coli cells co-expressing XAC0362-63. They also revealed that XAC0362-63 converts 3OMG into gallate (Fig. 9d, e).

The remaining pathway leading gallate up to the TCA cycle was inferred based on genome mining analysis, homology inference with previously characterized genes and syringaldehyde-specific transcriptional activation (Fig. 9a, b). It encompasses the putative gallate dioxygenase encoded by the genes XAC0878 (ligB, β-chain) and XAC0879 (ligA, α-chain), which are homologs to the respective N-terminal and C-terminal domains of gallate dioxygenases from P. putida KT2440 (GalA)⁴² and Sphingobium SYK-6 (DesB)⁴³ (Fig. 9b and Supplementary Fig. 11). Next steps are probably encoded by the XAC4155-56-57 genes, which are homologous to the ligK-ligU-ligJ genes previously characterized in Sphingobium sp. SYK-6¹² (Fig. 9a, b).

Discussion

This study uncovers complete pathways for the catabolism of the three main lignin precursors in the model plant pathogen X. citri 306 (Fig. 10). So far, only the microbial catabolism of coniferyl alcohol has been reported⁷, but in the context of a pathway having eugenol as the first substrate⁴⁴. Our study demonstrates the existence of catabolic pathways starting from coniferyl alcohol as well as p-coumaryl and sinapyl alcohols, adding new pieces in the puzzle of microbial metabolic pathways for lignin-related aromatic compounds.

Fig. 10: Schematic representation of the monolignols bioconversion pathways in *X. citri* 306, including proposed transporters for their assimilation and an efflux system for the secretion of aromatic compounds.

According to our data, the catabolism of monolignols in X. citri 306 starts with their uptake, probably facilitated by the putative outer membrane transporter MolK (XAC0352). MolK belongs to family COG4313, which has been implicated in the uptake of hydrophobic molecules⁴⁵. Next, the monolignols are dehydrogenated to aldehydes and subsequently to hydroxycinnamic acids (HCAs) mainly by two NAD⁺-dependent enzymes: the aryl alcohol dehydrogenase MolA (XAC0353) followed by the aryl aldehyde dehydrogenase MolB (XAC0354) (Fig. 10).

MolA is closely related to the CalA enzyme (61% sequence identity, 99% query cover) from Pseudomonas sp. HR199⁴⁶. CalA has been proposed to compose a pathway dedicated to the catabolism of eugenol in Pseudomonas sp. HR199, along with CalB, a coniferaldehyde dehydrogenase distantly related to MolB (30% sequence identity, 70% query cover)³². MolA and MolB co-occur in several plant-pathogenic, plant-symbiotic and environmental bacteria from the Xanthomonadales, Pseudomonadales, Burkholderiales and Rhizobiales orders, indicating that their biological roles go beyond the plant-pathogen context (Supplementary Fig. 12). Homologs closer to CalB were found exclusively in some Pseudomonas species and the triple co-occurrence of molA/calA, molB and calB genes was observed in some Pseudomonadales genomes, remaining to be determined its biological meaning.

Back to the pathways found in X. citri, after the action of MolA and MolB enzymes, the HCAs produced may be either excreted or converted into their respective hydroxybenzyl aldehyde derivatives via the CoA-dependent non-β-oxidation pathway encoded by the gene cluster (XAC0881-83) (Fig. 10). In contrast to X. campestris, which uptakes and metabolizes HCAs²⁹, X. citri 306 apparently metabolizes only HCAs produced intracellularly, likely due to the lack of transporters for their uptake. This adaptation correlates with studies showing that HCAs are commonly found in citrus fruits in their conjugated forms (ester- or glycoside-bond), with minimal concentrations in their free form^47,48. Part of the HCAs produced intracellularly is excreted by the cell, implying that HCAs deacetylation is probably a metabolic bottleneck to the flux towards the central carbon metabolism in X. citri 306.

Next, the H-G-S hydroxybenzyl aldehydes are converted to hydroxybenzoic acids by aryl aldehyde dehydrogenases, including XAC0882 and XAC0129. Then, 4HBA (H-type subunit) undergoes hydroxylation by the PobA enzyme (XAC0356) while vanillate (G-subunit) is O-demethylated by a Riske-type-oxygenase-reductase system VanAB (XAC0363-62), both steps resulting in the formation of PCA. Then, the PCA ring is ortho-cleaved and converted into β-ketoadipate by PcaHGBCD enzymes (XAC0367-68-69-71-70). Finally, PcaIJF enzymes (XAC0364-65-66) complete the conversions steps towards the tricarboxylic acid cycle (Fig. 10).

On another branch, syringate (S-type subunit) is O-demethylated to 3OMG and then to gallate by VanAB (XAC0363-XAC0362). Gallate is likely converted to 4-oxalomesaconate (OMA) by the action of LigAB (XAC0878-79) and follows a pathway up to pyruvate and oxaloacetate by enzymes encoded by the XAC4155-56-57 genes, which are homologous to ligK-ligU-ligJ from Sphingobium sp. SYK-6¹² (Figs. 9b and 10). This pathway was predicted based on transcriptional data and homology inference so future studies will be required to confirm the pathway and the metabolite intermediates leading gallate up to the TCA cycle in Xanthomonas.

Besides monolignols, X. citri 306 also uptakes their hydroxycinnamic and hydroxybenzoic aldehyde derivatives, driving them to two possible metabolic fates: oxidation to acids or reduction to alcohols. Two NADPH-dependent enzymes (XAC1484 and XAC3477) contribute to converting aryl aldehydes into aryl alcohols and the putative XAC1482-83-85 efflux system probably facilitates the aryl alcohol excretion. XAC1484 belongs to the Short-chain Dehydrogenases/Reductases (SDR) family whereas XAC3477 belongs to the Aldo/Keto Reductase (AKR) family, contrasting to the Medium-chain Dehydrogenase/Reductase (MDR) family members reported to play a role in aryl aldehyde reduction in other bacterial species^49,50,51,52. XAC1484 and XAC3477 share less than 41% sequence identity to functionally characterized enzymes, according to BLAST searches at the Swiss-Prot database⁵³. Thus, they represent novel reductases with potential biotechnological applications in aryl aldehydes detoxification⁵⁴ and production of value-added aryl alcohols^55,56,57.

The presence of the phenolic group seems to be mandatory for the complete catabolism of lignin-related compounds, since benzaldehyde, a non-phenolic aromatic, was converted to dead-end products (benzyl alcohol and benzoate) (Fig. 10). Other adaptative responses induced by aromatics include the upregulation of genes associated with chemotaxis and flagellar assembly, which might benefits X. citri fitness during host colonization. In X. campestris, the catabolism of plant-derived phenolic compounds has been shown to be important for virulence^27,28 and the induction of monolignols biosynthesis and plant cell wall lignification has been proved to be important host defense mechanisms^30,58,59. Thus, the capacity of X. citri 306 to sequester and metabolize key precursors of lignin biosynthesis may locally disrupt plant defense mechanisms in the benefit of the bacterium. This feature seems to be shared with other plant pathogens, as suggested by the conservation of the molRKAB operon in other Xanthomonas phytopathogenic species and of the molA and molB genes in phytopathogens from other genera (Supplementary Fig. 13).

In short, the metabolism of aromatic compounds in X. citri 306 converges to the central carbon metabolism but displays escape routes (efflux transporters) and reductases that likely function as complementary strategies to rapidly detoxify aryl aldehydes. Excretion of aryl acids or aryl alcohols by X. citri might serve as both a detoxification strategy and a mechanism to further hamper lignin biosynthesis, as these excreted compounds possibly compete with lignin precursors and prevent their oxidation by the plant laccases. These molecular strategies might inspire metabolic engineering approaches for the redesign of lignin biosynthesis in plants, aiming to facilitate biomass saccharification without compromising plant health^60,61. They also may serve as hotspots for the development of new treatments against plant diseases. From the industrial point of view, the knowledge provided here may support the development of microbial strains more tolerant to aryl aldehydes or proficient on converting lignin-derived compounds from agro-industrial side-streams into valuable bioproducts.

Methods

Bacterial strains and culture conditions

X. citri 306 was grown at 30 °C, 200 rpm, in LBON medium (10 g L⁻¹ bacto peptone and 5 g L⁻¹ yeast extract), minimal medium XVM2m (20 mmol L⁻¹ NaCl, 10 mmol L⁻¹ (NH₄)₂SO₄, 1 mmol L⁻¹ CaCl₂, 0.01 mmol L⁻¹ FeSO₄.7 H₂O, 5 mmol L⁻¹ MgSO₄, 0.16 mmol L⁻¹ KH₂PO₄, 0.32 mmol L⁻¹ K₂HPO₄, 0.03% m V⁻¹ casamino acids, pH 6.7) supplemented with different carbon sources, as better described in the next sections. Escherichia coli DH5α™ was used for DNA cloning, and E. coli BL21(DE3)-ΔslyD-pRARE2 or E. coli SHuffle® T7 Express lysY (New England Biolabs) were used for heterologous protein expression. E. coli strains were grown at 37 °C (or 20 °C), 200 rpm in LB medium (10 g L⁻¹ bacto peptone, 5 g L⁻¹ yeast extract, 10 g L⁻¹ NaCl), 2xYT (16 g L⁻¹ peptone, 10 g L⁻¹ yeast extract and 5 g L⁻¹ NaCl), or in M9 minimal medium (6.78 g L⁻¹ Na₂HPO₄, 3 g L⁻¹ KH₂PO₄, 0.5 g L⁻¹ NaCl, 1 g L⁻¹ NH₄Cl, 2 mmol L⁻¹ MgSO₄, 100 µmol L⁻¹ CaCl₂, pH 7.0). Bacterial growth was determined by measuring optical density at 600 nm (OD₆₀₀).

Preparation of lignin-derived compounds samples

Sugarcane bagasse was kindly provided by Isabel S/A, a Sugarcane ethanol company located in Novo Horizonte, São Paulo – Brazil. Fractions containing lignin-derived compounds were produced as previously described^62,63 (Supplementary Fig. 1). Briefly, sugarcane bagasse was subjected to an alkaline process (130 °C, 30 min, 1.5% NaOH, and 1:10 bagasse/alkaline solution ratio) in a 7.5 L reactor (Series 4580 HT, Parr) with temperature and stirring control. Next, the alkaline liquor was acidified to pH 2 with sulfuric acid (72%) to precipitate the insoluble lignin fraction⁶². After filtration, the liquid fraction containing aromatic compounds, here referred to as LDC-I, was obtained. The solid stream (precipitated lignin) was subjected to a hydrothermal depolymerization reaction at 350 °C, 165 bar, 90 min, and a lignin/water ratio of 1:50 in an inert atmosphere (N₂), and an agitation of 500 rpm. This reaction was taken in a high-pressure autoclave reactor (500 mL, model 4575 A, Parr), in batch mode, up to 344 bar, 500 °C, with control over temperature, pressure, and agitation (4848 reactor controller, Parr). At the end of the reaction, the depolymerized liquid stream was subjected to a liquid-liquid extraction step with ethyl acetate (Vetec PA ACS) (1:1, 3x), and this solvent was evaporated to obtain the LDC-II fraction (bio-oil). LDC-I and LDC-II samples were characterized by UV spectroscopy, GC-MS and HPLC, as described below and in the supplementary method.

Quantification of phenolic compounds by UV spectroscopy

Concentration of total aromatic compounds in the LDC-I sample was determined by UV spectroscopy at 280 nm, on 2 M NaOH solution, pH 12, using the following equation (1)⁶⁴.

$${{\rm{Aromatics}}}\,({{\rm{g}}}\,{{\rm{L}}}^{-1})\,=\,4.187 \times 10^{-2}*({{\rm{Abs}}}_{280 {{\rm{nm}}}}) - (3.279 \times 10^{-4})*{{\rm{dilution}}}$$

(1)

Next, it was diluted in XVM2m, filter-sterilized, and the aromatics final concentration in the medium was adjusted to 1 g L⁻¹ for X. citri 306 cultivations. LDC-II was solubilized on 1 mL 0.5 M NaOH solution, then diluted in minimal medium XVM2m for a final theoretical concentration of 1 g L⁻¹. After adjusting the pH to 6.7, the medium was filter-sterilized, and the concentration of aromatic compounds in the medium was estimated by 280 nm absorbance, at pH 12, according to the equation (1)⁶⁴. Then, this medium was diluted in XVM2m to adjust the aromatics concentration to 0.3 g L⁻¹.

X. citri 306 cultivations

For growth curves analysis, X. citri 306 was cultured in LBON medium, 100 μg mL⁻¹ ampicillin, overnight at 30 °C and 200 rpm. Then, the harvested cells were washed once with XVM2m and inoculated for an initial OD₆₀₀ of 0.05 in XVM2m supplemented with chemical standards of aromatic compounds, complex mixtures of lignin-derived compounds, or glucose (Supplementary Table 9). Except for LDC-I, the other conditions were also carried out in the presence of 5 mmol L⁻¹ glucose as an additional carbon source. The growth was monitored in 96 well plates incubated in a SpectraMax® M3 multi-mode microplate reader (Molecular Devices) using the SoftMax Pro software or in an Infinite 200 PRO plate reader (Tecan) using the i-control software, for 48 h, at 30 °C using n ≥ 3 biological replicates. Specific growth rates (μ) were obtained using the package growthrates as described by Petzoldt⁶⁵.

For HPLC analysis of conditions containing model compounds, X. citri 306 was cultivated in 125 mL flasks containing 15 mL of medium as detailed in Supplementary Table 10. The culture medium was sampled by removing 2 mL before and after specific post-inoculation time points (Supplementary Table 10), centrifuged at 4960 x g for 5 min to pellet the cells, and the supernatants were stored at −20 °C until the HPLC analysis described in the supplementary method.

For HPLC and GC-MS analysis of cultivations performed in minimal medium supplemented with LDC-I and LDC-II samples, X. citri 306 was grown in 10 mL of LBON and prepared as described above. The OD₆₀₀ was adjusted to 0.1 in 15 mL of XVM2m supplemented with 1 g L⁻¹ LDC-I or in XVM2m(G) supplemented with 0.3 g L⁻¹ LDC-II. The cells were incubated at 30 °C, 200 rpm. The culture medium was sampled by removing 2 mL in 0 h and 30 h post-inoculation, centrifuged at 4960 x g for 5 min, and the supernatants were utilized for further analyses (LDC-I (0 h), LDC-I (30 h), LDC-II (0 h), LDC-II (30 h). The experiment was conducted with n = 3 biological replicates. These samples were analyzed by GC-MS and HPLC for compounds identification and quantification as described in the supplementary method. Data were analyzed using OriginPro (2021).

X. citri 306 genome mining

For the initial prediction of the metabolic pathways related to the catabolism of aromatic compounds in X. citri 306, we used the BLASTp tool⁶⁶ to search in the genome of this bacterium for proteins homologous to those deposited in the manually curated eLignin database⁷. As a complementary tool, we also use the metabolic maps available in the KEGG database⁶⁷ for this strain as a reference.

RNA extraction and sequencing

X. citri 306 cells grown at 30 °C, 200 rpm on 40 mL XVM2m minimal medium supplemented with different carbon sources (Supplementary Table 3) were collected at the middle exponential phase (OD₆₀₀ = 0.05–0.1) from n = 4 biological replicates. Total RNA was extracted using the TRIzol/chloroform method⁶⁸. Samples were treated with RNase-free DNaseI (Invitrogen) and RNaseOUT (Invitrogen) for 30 min, at 37 °C, and purified with the RNeasy Mini Kit (Qiagen), following manufacturer’s recommendations. RNA samples concentration was determined using Nanodrop 1000 (Thermo Scientific), and their integrity was evaluated in an Agilent 2100 Bioanalyzer using the Agilent 2100 Expert Software (Agilent Technologies). The rRNA was depleted with the Ribo-Zero Plus rRNA depletion kit (Illumina Inc.). Subsequently, these samples were used to synthesize cDNA libraries with TruSeq Stranded Total RNA kit (Illumina Inc.), according to the manufacturer’s recommendations. The final RNA-seq libraries were quantified via qPCR using the QIAseq Library Quant assay kit (Qiagen), and library quality was verified using an Agilent 2100 Bioanalyzer (Agilent Technologies). Samples were pooled, and the RNA-seq was performed on an Illumina HiSeq 2500 platform equipped with HCS 2.2.68 software (SEQ facility LNBR-CNPEM, Campinas, Brazil).

RNA-seq data processing and analysis

The raw reads generated in the RNA-seq were filtered to remove adapters, primers and low-quality sequences using the fastp tool⁶⁹. Contaminant rRNA read sequences were removed using sortmeRNA⁷⁰. High-quality reads were mapped in the X. citri 306 genome (GI: 21240769) using the Bowtie2 tool⁷¹, allowing only two mismatches and unique alignments. Next, the Samtools program⁷² was used to process the alignment files, which were inspected using the Integrative Genome viewer program⁷³. Reads mapped to the X. citri 306 genome were subjected to the featureCounts tool⁷⁴ to estimate the number of reads mapped to each transcript. The processed data were summarized and plotted using the MultiQC package⁷⁵. Low count transcripts were removed, keeping only those that showed CPM (Counts Per Million) above 0.9, equivalent to a count of 10 to 15 reads per transcript. Differential expression analysis was carried out using edgeR package⁷⁶ by pairwise comparisons between X. citri 306 grown in XVM2m supplemented with lignin-related aromatics (Supplementary Table 3) and XVM2m(G) containing 5 mmol L⁻¹ glucose (reference medium). Differentially expressed genes were defined using log₂ Fold Change ≥ 1 (upregulated genes) or ≤ -1 (downregulated genes), and a p-adjusted ≤ 0.05 as thresholds. Variance analyses were conducted utilizing PCA (Principal Component Analysis) to assess data integrity and comparability. According to this analysis, one non-concordant replica from the coniferyl alcohol and one from 4-hydroxybenzaldehyde condition were excluded, and the differential expression analysis for these conditions was done with only n = 3 biological replicates. Functional and pathway enrichment analyses were performed separately to predict the functions of differentially expressed genes. Modular gene co-expression analysis was performed using the CEMiTool package³¹ and pathway enrichment analysis using the enrichKEGG function of the clusterProfiler R package⁷⁷. One thousand simulations were performed for each condition, and the groups were considered significantly enriched when they presented p ≤ 0.05. The Gene ontology (GO) enrichment analysis was performed using the clusterProfiler 3.14.3 R/Bioconductor package and the categories were considered enriched based on hypergeometric test, implemented in the enrich function of the package⁷⁸. Venn diagram plots were generated using the InteractiVenn web-based tool⁷⁹ (http://www.interactivenn.net/). de novo reconstruction of molRKAB operon from RNA-seq data was performed using the Trinity software package⁸⁰. Comparative analysis with the reference genome (AE008923.1) was performed using pyGenomeViz package (https://moshi4.github.io/pyGenomeViz/), which was also employed for the analysis of molRKAB operon conservation.

Gene cloning, protein expression and purification

The open reading frames selected for biochemical assays based on the RNA-seq analysis were amplified by PCR using specific primers and cloned in the expression vector pET28a(+) or pET21b(+) using the In-Fusion® HD kit (Takara Bio) or by using restriction enzymes and DNA ligase following manufacturer’s instructions (Supplementary Table 11). The constructs were confirmed by DNA sequencing and used to transform the E. coli strain BL21(DE3)-ΔSlyD-pRARE2 or BL21(DE3)-SHuffle®-lysY (for XAC0353 construct). The transformed cells were cultured in LB medium, 50 µg mL⁻¹ kanamycin or 100 µg mL⁻¹ ampicillin at 37 °C, 200 rpm until OD₆₀₀ ~ 0.8. Then, the protein expression was induced by adding 0.5 mmol L⁻¹ isopropyl-β-d-thiogalactopyranoside (IPTG) to the medium and incubating it at 20 °C for 16 h. Cells were harvested by centrifugation at 7,500 x g, at 4 °C for 30 min, resuspended on buffer as detailed in the Supplementary Table 12, and disrupted by sonication (pulses of 15 s with intervals of 30 s during 15 min, 30% amplitude). The cell extract was centrifuged at 35250 x g, at 4 °C for 30 min. The target proteins present in the supernatant were purified as described in the supplementary method. Protein concentration was estimated based on the absorbance of protein samples at 280 nm using the extinction coefficient calculated from their amino acid sequences using the ProtParam tool⁸¹.

Enzyme activity screening

The activity of putative ADHs was assessed by measuring the reduction of NAD(P)⁺ in the presence of aryl alcohols or the oxidation of NAD(P)H in the presence of aryl aldehydes (Supplementary Table 13). Putative ALDHs were evaluated for their activity over aryl aldehydes in presence of NAD(P)⁺ (Supplementary Table 13). Substrates and cofactors were purchased as detailed in Supplementary Table 14. The enzyme activity assays were performed in 100 µL of 50 mmol L⁻¹ HEPES buffer, pH 7.5, containing 0.25 mmol L⁻¹ NAD(P)⁺ or NAD(P)H and specific substrates at a final concentration of 0.25 mmol L⁻¹ (Supplementary Table 14). NAD(P)H production or consumption was monitored by absorbance at 340 nm and enzyme activity was defined as described in the supplementary method. The PobA activity was assessed in 100 µL of 50 mM Tris/sulfate buffer, pH 7.5, containing 0.5 mmol L⁻¹ NADPH, 60 µmol L⁻¹ FAD, and 0.5 mmol L⁻¹ of either 4-hydroxybenzoate or benzoate. The reaction was initiated by adding 1 µmol L⁻¹ of the PobA enzyme. A control reaction was conducted to monitor the absorption decrease at 340 nm in the absence of aryl substrates. Data were analyzed using OriginPro (2021).

Enzyme specific activity assay

The specific activity of XAC0353 and XAC0354 was accessed in 1 mL of 36 mmol L^-1 HEPES buffer, pH 7.5, containing 0.25 mmol L⁻¹ substrate, and 2 mmol L⁻¹ NAD⁺, at 30 °C for 10 min. In reactions involving coniferyl alcohol, cinnamyl alcohol, p-coumaryl alcohol, and sinapyl alcohol, XAC0353 was used at a concentration of 0.1 μmol L⁻¹. For the remaining aryl alcohols listed in Supplementary Table 14, a concentration of 1.2 μmol L⁻¹ of XAC0353 was utilized. XAC0354 at 0.08 μmol L⁻¹ was used in reactions containing aryl aldehydes (see Supplementary Table 14). Reactions were stopped by heat at 100 °C, 2 min, 650 rpm, and centrifuged (20,817 x g, 4 °C, 10 min). All assays were done using n = 3 biological replicates. The supernatant was used to measure the NADH production by HPLC as described by Sporty et al.⁸², with modifications (see details in the supplementary method). Data were analyzed using OriginPro (2021).

Whole-cell activity assays using aryl aldehydes or HCAs substrates

Whole-cell assays were performed as described by García-Hidalgo et al.⁴⁹. In short, E. coli BL21(DE3)-ΔSlyD-pRARE2 transformed with expression vectors containing the gene XAC0882, XAC0129 or co-expressing the genes XAC0881/XAC0883 (or with the empty vector, used as a negative control) were cultured in 5 mL of LB medium, overnight, at 30 °C and 200 rpm. Harvested cells were washed with M9 minimal medium and inoculated to an initial OD₆₀₀ of 0.8 in M9 medium containing 50 µg mL⁻¹ kanamycin or 100 µg mL⁻¹ ampicillin, 0.5 mmol L⁻¹ IPTG, 56 mmol L⁻¹ glucose, and 5 mmol L⁻¹ aryl aldehydes for putative dehydrogenases (XAC0129 and XAC0882) (Supplementary Table 14) or 5 mmol L⁻¹ p-coumarate, ferulate or sinapate for XAC0881/883. The cells were incubated at 30 °C and 200 rpm for 20 h. Samples were taken at the final time, centrifuged at 4960 x g for 5 min to pellet the cells, and the supernatants were analyzed as described in the HPLC section. All assays were done using n = 2 biological replicates.

Whole-cell activity assays for O-demethylases

O-demethylase whole-cell assays were performed as described by Lanfranchi et al.⁸³ with modifications. Briefly, the pairs XAC0310-XAC0311 or XAC0362-XAC0363 were co-expressed in E. coli BL21(DE3)-ΔslyD-pRARE2 in 50 mL of 2xYT medium supplemented with 1 mmol L⁻¹ L-cysteine, 0.1 mg mL⁻¹ FeCl₃, and 0.1 mg mL⁻¹ FeSO₄ during induction with 0.5 mmol L⁻¹ IPTG. After expression, the cells were centrifuged (2975 g, 4 °C, 10 min), washed with 20 mL of 50 mmol L⁻¹ Tris-HCl buffer, pH 7.5, and resuspended in 100 mmol L⁻¹ Tris-HCl, pH 7.5. The amount of buffer used for resuspension was adjusted to ensure that the OD₆₀₀ ( ~ 8.8) for the control cells (empty vector) and for the cells expressing the enzymes were the same, providing a theoretically equal number of cells across all samples. Reactions were prepared in 100 mmol L⁻¹ Tris-HCl, pH 7.5, containing 0.1 mmol L⁻¹ syringate, vanillate or 3OMG (final DMSO concentration 1%), 5 mmol L⁻¹ DTT, 0.1 mmol L⁻¹ FeSO₄, and 20% (v/v) of the cell stock in 2 mL Eppendorf tubes. The reactions were incubated at 30 °C, 150 rpm, for 21 h. After incubation, the reactions were stopped by heating at 90 °C for 10 min, then centrifuged (20,000 x g, 23 °C, 10 min). The supernatants were frozen at −20 °C before being analyzed by HPLC as described in the supplementary method. Stock solutions of substrates were prepared in 10 mmol L^-1 DMSO and stored at 4 °C. All assays were done using n = 2 biological replicates.

Gene deletion

Deletion mutants were generated using established methods⁸⁴ with some modifications. In brief, DNA fragments upstream and downstream of the target gene (~ 1 Kbp) were amplified by PCR from X. citri 306 genomic DNA using the primers listed in Supplementary Tables 15 and 16. The PCR fragments were cloned into the pJET1.2/blunt vector using The CloneJET PCR Cloning Kit (Thermo Scientific). Next, they were digested with specific restriction enzymes for sequential cloning into the suicide vector (pNPTS138) using T4 DNA ligase (Thermo Scientific). Alternatively, the PCR fragments were directly cloned into the suicide vector pNPTS138 using the commercial In-Fusion® HD kit (Takara Bio) or NEBuilder® HiFi DNA Assembly Master Mix kit (New England Biolabs). The final constructions (~ 2 Kbp) were confirmed by DNA sequencing. The recombinant plasmids were introduced into X. citri 306 by electroporation. The selection of knockout strains was performed as described in the supplementary method.

Characterization of gene knockout strains

The growth curve assay of the ΔpobA strain was conducted in a minimal medium XVM2m or XVM2m(G) supplemented with 4-hydroxybenzoate or benzaldehyde. The cultures were performed in sealed 96-well plates and incubated in a SPARK® multimode microplate reader (Tecan) at 30 °C with agitation. Data were collected with the SparkControl v.3 software. Each condition was replicated in at least three wells. For HPLC analysis, all mutant strains generated in this work were cultivated in 125 mL flasks containing 15 mL of medium as detailed in Supplementary Table 17. The culture medium was sampled by removing 2 mL before and after specific post-inoculation time points (Supplementary Table 17), centrifuged at 4960 x g for 5 min to pellet the cells, and the supernatants were stored at −20 °C until the analysis.

Phylogenetic analysis

To analyze the distribution of molA/molB and calA/calB genes in other bacterial genomes, a phylogenetic tree was constructed. The genome sequence of X. citri 306 along with 136 genomes were used for the analysis. The genomes were selected based on Sequence Similarity Network (SSN) analyses. Initially, individual references protein sequences of MolA (XAC0353), MolB (XAC0354), CalA (CAB69495.1) and CalB (CAA06926.1) were submitted to EFI-EST webtool⁸⁵ as seeds for SSN analyses to recover iso-functional clusters comprising the UniProt seed-related proteins sequences. The reference genome sequences of bacterial species identified on the iso-functional gene cluster harboring MolA and CalA were downloaded from the NCBI RefSeq database. This set of genome sequences was submitted to UBCG tool⁸⁶, and a phylogenetic tree was inferred based on 92 conserved single copy marker genes using maximum likelihood method implemented in RAxML with 100 bootstraps. The obtained tree was visualized in iTol (itol.embl.de). MolA/MolB and CalA/CalB related genes are highlighted in the tree according to protein sequence identity >50% and coverage >70% with references sequences as thresholds.

Statistics and reproducibility

Differentially expressed genes were evaluated according to negative binomial distribution implemented by edgeR⁷⁶ and defined using absolute log₂ Fold Change above 1 and adjusted p-values < 0.05 as thresholds. Exact adjusted p-values were provided in the Supplementary Data 1 file. Variance analyses were conducted utilizing PCA to assess data integrity and comparability. Outliers samples in PCA were excluded. The precise n number of replicate experiments are indicated in the figure legends. The investigators were not blinded to allocation during experiments and outcome assessment, since all analyses were objective in nature.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

RNA-seq data were deposited in the Gene Expression Omnibus database under accession number GSE252662. Other data generated in this study are presented in the supplementary material, supplementary data files and the source data file. Homologous sequences used for comparative purposes are found in Genbank under the accession codes CAB69495.1 [https://www.ncbi.nlm.nih.gov/protein/CAB69495.1], BAK68041.1 [https://www.ncbi.nlm.nih.gov/protein/BAK68041.1], BAH79100.1 [https://www.ncbi.nlm.nih.gov/protein/BAH79100.1], AAA75634.2 [https://www.ncbi.nlm.nih.gov/protein/AAA75634.2], BAK67507.1 [https://www.ncbi.nlm.nih.gov/protein/ BAK67507.1]. X. citri subsp. citri protein sequences, such as XAC0353, were retrieved from the KEGG database [https://www.genome.jp/entry/xac:XAC0353]. Databases used in this study: Uniprot: (https://www.uniprot.org/) eLignin (http://www.elignindatabase.com/)⁷ UniProtKB/Swiss-Prot (https://www.expasy.org/resources/uniprotkb-swiss-prot) KEGG RefSeq (https://www.ncbi.nlm.nih.gov/refseq/) Gene Expression Omnibus database (https://www.ncbi.nlm.nih.gov/geo/) Source data are provided with this paper.

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Acknowledgements

We are grateful to Renan H. S. Fernandes, for the preparation of LDC-I samples, to Adriano Freitas Lima, for their contribution on HPLC data collection, to Angelica Luana Carrilo Barra for her contribution on XAC0129 purification and to Mario Tyago Murakami for reviewing the manuscript and offering valuable feedback. This research used facilities of the Brazilian Biorenewables National Laboratory (LNBR), part of the Brazilian Center for Research in Energy and Materials (CNPEM), a private non-profit organization under the supervision of the Brazilian Ministry for Science, Technology, and Innovations (MCTI). The High-Performance Sequencing (SEQ) and Biophysics of Macromolecules (BFM) staff is acknowledged for their assistance during the experiments (Proposal 27931 and 20230475). We also thank the Molecular Chemistry Laboratory from LNBR for all support on HPLC and GC–MS analyses. This research was supported by the São Paulo Research Foundation - FAPESP (grants 2019/06921-7 to P.O.G. and scholarship 2019/08590-8 to D.B.M., 2022/01070-1 to A.J.V.C.B. and 2021/07139-0 to A.R.L.), Coordination for the Improvement of Higher Education Personnel – CAPES and by the Brazilian Scientific and Technological Development Council – CNPq.

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Graduate Program in Genetics and Molecular Biology, Institute of Biology, University of Campinas (UNICAMP), Campinas, São Paulo, Brazil
Damaris B. Martim, Anna J. V. C. Brilhante, Augusto R. Lima & Priscila O. Giuseppe
Brazilian Biorenewables National Laboratory (LNBR), Brazilian Center for Research in Energy and Materials (CNPEM), Campinas, São Paulo, Brazil
Damaris B. Martim, Anna J. V. C. Brilhante, Augusto R. Lima, Douglas A. A. Paixão, Joaquim Martins-Junior, Fernanda M. Kashiwagi, Lucia D. Wolf, Mariany S. Costa, Fabrícia F. Menezes, Rafaela Prata, Matheus C. Gazolla, Juliana A. Aricetti, Gabriela F. Persinoti, George J. M. Rocha & Priscila O. Giuseppe

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Damaris B. Martim
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Anna J. V. C. Brilhante
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Augusto R. Lima
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Douglas A. A. Paixão
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Joaquim Martins-Junior
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Fernanda M. Kashiwagi
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Lucia D. Wolf
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Mariany S. Costa
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Fabrícia F. Menezes
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Matheus C. Gazolla
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Juliana A. Aricetti
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Gabriela F. Persinoti
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George J. M. Rocha
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Priscila O. Giuseppe
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Contributions

D.B.M. performed X. citri 306 genome mining, X. citri 306 cultivations, RNA extraction, molA and molB gene knockout, ΔmolA, ΔmolB, ΔpobA and ΔpcaHG strains characterization, XAC0881, XAC0883 and dehydrogenases cloning, activity screening, and whole cell assays. A.J.V.C.B. performed XAC3477 purification, MolA and MolB cloning, expression, purification, and enzyme assays. A.R.L. performed XAC0129 cloning and ROs-type O-demethylases cloning, whole cell assays, gene knockout and strains characterization. L.D.W., M.S.C. and M.C.G. designed and performed HPLC assays. D.B.M., D.A.A.P., J.M.J., G.F.P. and P.O.G. designed and performed RNA-seq assays and analyses. F.F.M., J.A.A. and R.P. designed and performed GC-MS assays. J.M.J. and G.F.P. performed XAC0351-54 operon conservation analysis and phylogenetic analysis. F.M.K. performed pobA and pcaHG gene knockout. F.F.M., and G.J.M.R. designed and performed lignin depolymerization reactions. P.O.G. coordinated the work. D.B.M. and P.O.G. wrote the manuscript. All authors contributed to data analysis and manuscript revision.

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Correspondence to Priscila O. Giuseppe.

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Martim, D.B., Brilhante, A.J.V.C., Lima, A.R. et al. Resolving the metabolism of monolignols and other lignin-related aromatic compounds in Xanthomonas citri. Nat Commun 15, 7994 (2024). https://doi.org/10.1038/s41467-024-52367-6

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Received: 08 January 2024
Accepted: 03 September 2024
Published: 12 September 2024
DOI: https://doi.org/10.1038/s41467-024-52367-6

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