Low lignin solubility in aqueous solution is one of the major bottlenecks for lignin biodegradation and bioconversion. Alkaline solution contributes to improving lignin solubility, whereas most microbes can not survive in alkaline conditions. Herein, lignin dissolution behaviors in different pH solutions were systematically investigated, which indicated that solution pH above 10.5 contributed to high solubility of alkali lignin. To match with alkaline lignin aqueous system, several alkali-tolerant ligninolytic bacteria were isolated, most of which are distinct to previously reported ones. Then, the ligninolytic capabilities of these isolates were assessed in different pH conditions by determining their assimilation on alkali lignin, lignin-derived monomers and dimers, their decolorization capabilities, and their lignin peroxidase activities. Thereafter, the underlying ligninolytic and alkali-tolerant mechanisms of Sutcliffiella sp. NC1, an alkalophilic bacterium, was analyzed on the basis of its genome information. The results not only provide valuable information for lignin biodegradation and lignin valorization, but also expand knowledge on alkali-tolerant bacteria.
Declaration of Competing Interest
The authors declare no conflict of interest. Supplementary materials
Supplementary material associated with this article can be found, in the online version, at doi:10.1016/j.jobab.2023.09.004. 1 These authors contributed equally to this work.
Lignocellulosic biomass is regarded as the most likely alternative for fossil resources owing to its reserves for both energy and carbon. Lignin, cellulose and hemicellulose are three main components of lignocellulosic biomass, nonetheless the progress of lignin valorization is far behind the other two polysaccharides. One widely accepted refinery route for (hemi)cellulose is that lignocellulosic biomass is first pretreated to destroy its rigid structure. Then, exposed (hemi)cellulose are enzymatically hydrolyzed to monosaccharides, which are converted to target bio-chemicals by functional microbes or enzymes. Hitherto, multiple bio-chemicals have been produced via this refinery route, and some production lines have been or are being evaluated in commercial scale (Patel and Shah, 2021; Lynd et al., 2022). By contrast, lignin is designed to be burned for energy supply in most biorefinery scenarios. Considering the tremendous volume of lignin produced worldwide, there is growing interest in lignin valorization (Werner and Eltis, 2023).
Despite diverse solid linkages in lignin, some microbes still enable its decomposition. Inspired by the bioconversion routes for (hemi)cellulose, similar valorization routes are also proposed for lignin that it is first depolymerized to low-molecular-weight fragments, which are then converted to target compounds by wild or engineered microbes. Especially, consolidated processing for lignin depolymerization and bioconversion is feasible by a single powerful microbe or by a microbe consortium (Li et al., 2022; Liu et al., 2022; Wang et al., 2022). Compared with conventional lignin valorization routes, the advantage of the aforementioned lignin bioconversion route is that some microbes enable to funnel multiple lignin-derived compounds to target ones, not only improving product yields but also avoiding excessive purification operations (Linger et al., 2014). Recently, several versatile microbes have been exploited for lignin bioconversion and some valuable chemicals have been prepared from lignin by the aforementioned biological routes, such as microbial lipids, polyhydroxyalkanoate, organic acids, aromatics, etc. (Cai et al., 2021; Elmore et al., 2021; Suzuki et al., 2021; Zhao et al., 2021; Luo et al., 2022; Xu et al., 2022; Kumar et al., 2023; Nawaz et al., 2023; Saini et al., 2023; Weiland et al., 2023). One of the major obstacles for bioconversion of real lignin is that most types of lignin have low solubility in conventional aqueous medium due to their high molecular weights and hydrophobic groups. It is generally recognized that sufficient interaction of substrate with microbes/enzymes is a prerequisite for efficient bioconversion processes (Huang et al., 2022). On the contrary, low solubility of substrates in aqueous solution results in weak biochemical reaction. If lignin is completely dissolved in aqueous solution, ligninolytic microbes/enzymes will contact and function on lignin more efficiently.
Alkali pretreatment is a widely adopted pretreatment method for lignocellulosic biomass that alkaline reagents contribute to exposed cellulose and increased cellulose content through saponifying the ester bonds between hemicellulose and lignin, along with breaking chemical bonds of lignin. Meanwhile, some lignin components are dissolved in the alkaline pretreatment liquid, a good testimony to the high lignin solubility in alkaline solution. A recent study demonstrated that lignin solubility in alkaline solution varies greatly with the polarity of used solvents, the solution viscosity, and the cationic radius of used alkaline reagents (Melro et al., 2020). It is assumed that if alkaline solution was deployed for lignin bioconversion, high lignin solubility and homogeneous reaction system may promote the conversion processes. Nonetheless, most microbes are not compatible with alkaline solutions, impeding the implementation of this idea. Some special microbes have been dedicated to converting dissolved lignin components in alkaline pretreatment liquid or papermaking black liquor, nevertheless, the pH values of the applied solution were adjusted to about 7.0 during the bioconversion processes (Suzuki et al., 2021; Luo et al., 2022). From our point of view, if alkali-tolerant microbes were subjected in alkaline lignin solution, high solubility of lignin and high cell viability for lignin bioconversion might be addressed simultaneously.
Although most microbes are preferred for near-neutral pH environments, some are still able to tolerate acid or alkaline conditions, even exclusively live in extreme pH environments. Besides extreme pH environments, microbes are also discovered in other extreme habitats, such as those with high or low temperatures, high salinity, high pressure, high organic solvent contents, as well as heavy metals. These microbes are collectively called extremophiles. Special metabolic features and physiological properties enable these microbes as potential biocatalysts under harsh industrial conditions. In fact, extremophiles or extremozymes have brought immense benefits to the current bio-industry. For instance, thermostable amylases and Taq polymerases from hyperthermophiles play indispensable roles in starch-based fermentation industry and genetic engineering industry, respectively; some solvent tolerant microbes boost the biodegradation of spilled oils or other organic reagents; acid/alkali tolerant microbes contribute to easy fermentation processes by avoiding excessive neutralizing reagents addition (Nadaroglu et al., 2022; Zhu et al., 2022). In contrast to extensive applications of acid-tolerant microbes, alkali-tolerant microbes are rarely mentioned and the most acknowledged one is alkaliphiles adopted for producing alkaline detergent enzymes. Just recently, ligninolytic microbes in extreme environments have attracted increasing attention. For instance, ligninolytic microbes and enzymes in thermophilic environments were systematically characterized with stable isotope probing genome-resolved metagenomics and enzyme characterization, providing possibilities for efficient lignin bioconversion at elevated temperatures (Levy-Booth et al., 2022; An et al., 2023; Fall et al., 2023; Yang et al., 2023). By contrast, rare alkali-tolerant microbes have been systematically investigated. Few alkali-tolerant Bacillus species have been demonstrated possessing ligninolytic capability, whereas most of their ligninolytic profiles were investigated in near-neutral pH environments, missing the opportunity for unraveling their ligninolytic profiles in alkaline conditions.
With the assumption that the combination of alkaline solution and alkali-tolerant ligninolytic microbes enables efficient lignin biodegradation, multiple alkali-tolerant ligninolytic microbes were isolated in this study. Afterward, the ligninolytic capabilities of these isolates were detailed assessed by determining their assimilation on alkali lignin, lignin-derived dimers and lignin-derived monomers, their decolorization capabilities, as well as their lignin peroxidase activities. Thereafter, the underlying ligninolytic and alkali-tolerant mechanisms of Sutcliffiella sp. NC1, an alkalophilic ligninolytic bacterium, were analyzed on the basis of the genome information. The obtained results not only provide a potential bioconversion route for lignin, but also expand current knowledge on both alkali-tolerant bacteria and lignin biodegradation.
2.
Materials and methods
2.1
Soil samples, medium and reagents
Samples used for screening alkali-tolerant ligninolytic microbes were collected from several sites of China, including Xinle City in Hebei Province, Kaifeng City in Henan Province, Baotou City in the Inner Mongolia Autonomous Region, Lishui City and Shaoxing City in Zhejiang Province, Suzhou in City Anhui Province, Huai'an in Jiangsu Province, Wuhan in Hubei Province, Neijiang City and Yibin City in Sichuan Province. These samples were collected from lignocellulose-rich soils and the pH values of these samples were not considered at the sample collection stage.
Modified M9 minimal medium, consisting of 13.56 g/L Na2HPO4·12H2O, 6 g/L KH2PO4, 1 g/L NaCl, 2 g/L NH4Cl, 0.492 g/L MgSO4·7H2O, 0.111 g/L CaCl2, and 10 mL Hoagland trace element solution, was used for incubating lignocellulose-rich samples and investigating the effects of pH on cell growth of the isolates. The Luria-Bertani (LB) medium was used for culturing microbe seeds, investigating the effects of pH on the cell growth of the isolates, and investigating the decolorization capability of the isolates.
The alkali lignin (commodity code of 370, 959) and lignosulfonic acid sodium salt (commodity code of 471, 038) used in the present study are purchased from Sigma-Aldrich. The dealkaline lignin (CAS code of 9005–53–2) used is purchased from Aladdin Biochemical Technology Co., Ltd. (Shanghai, China). The lignin-derived dimers and monomers, glucose and other chemicals used are analytically or higher pure grade.
2.2
Investigating effects of pH on lignin solubility and particle size distribution
A series of 10 mL aqueous solutions with pH values of 7.0, 8.0, 9.0, 10.0, 10.5 and 11.0, prepared by adding different doses of NaOH, were applied to investigate the effects of pH on lignin solubility and the particle size distributions of dissolved lignin. In detail, 0.3 g alkali lignin was added into 10 mL different pH solutions individually, and then the obtained 30 g/L lignin solutions were incubated at 250 r/min for 12 h to make lignin sufficiently dissolve in aqueous solution. Thereafter, the well mixed lignin solutions were centrifuged at 5 000 r/min for 10 min to separate the undissolved lignin fractions from the aqueous solution. The solid fractions were washed thrice with deionized water and dried at 60 ℃ to constant weight. The lignin solubility (%, w) was calculated with the formula: lignin solubility = (0.3 – wsolid fraction)/0.3 × 100%. The supernatants were applied to characterize lignin particle size distribution in different pH solutions by dynamic light scattering. The dynamic light scattering was done with a Malvern Zetasizer (Nano ZS 90, Malvern Ltd., Malvern, UK). Because the dissolved lignin varied with solution pH, it was impractical to analyze the absolute value of signal intensities of dynamic light scattering and relative intensity (%) was adopted to analyze lignin particle size distribution. Moreover, spectral scan from 200 to 400 nm was implemented for dissolved lignin by a microplate reader (SpectraMax M3, Molecular Devices, CA, USA) to determine the alterations of chemical bonds in lignin.
2.3
Isolation and identification of alkali-tolerant ligninolytic microbes
Lignocellulose-rich soils were collected from several places in China by the members in our laboratory as stated in Section 2.1. Then, these samples were added into 50 mL sterile lignin-rich M9 medium (supplemented with 10 g/L lignin, pH of 11.0) at the dose of 5.0 g/L, aerobically cultured at 30 ℃ and 250 r/min for 48 h. Thereafter, 1 mL cultured broth was transferred into another 50 mL same medium and cultured in the same condition. After five transfers, the obtained broth was diluted for 104–106 folds, and plated on the LB solid medium. After cultivation at 30 ℃ for 72 h, the growing microbes were distinguished by colonial morphologies and the dominant species were streaked on fresh LB solid medium for isolation.
According to colonial morphologies, all isolates were bacteria, and thus 16 s ribosomal DNA (16 s rDNA) was applied to identify these isolates with the classical primers of 27F (AGAGTTTGATCCTGGCTCAG) and 1492R (GGTTACCTTGTTACGACTT). On the basis of sequenced 16 s rDNA, the classification of these isolates was implemented by sequence similarity analysis and evolutionary trees were drawn by Mega 11.0.
2.4
Determining assimilative capacities of isolated ligninolytic microbes on alkali lignin, lignin-derived monomers and lignin-derived dimers
To assess the ligninolytic capability of the isolates, their assimilative capacities on alkali lignin, lignin-derived monomers and lignin-derived dimers were determined, separately. In detail, p-coumaric acid, p-hydroxybenzoic acid, p-hydroxy benzaldehyde, and p-hydroxybenzylalcohol were selected as the representatives of H-lignin monomers (with no methoxy groups in the aromatic nucleus); ferulic acid and vanillic acid were selected as the representatives of G-lignin monomers (with one methoxy group in the aromatic nucleus); syringaldehyde, syringic acid, sinapic acid and 4-hydroxy-3, 5-dimethoxybenzyl alcohol were selected as the representatives of S-lignin monomers (with two methoxy groups in the aromatic nucleus); guaiacol, veratryl alcohol, protocatechuic acid, gallic acid and catechol were selected as other common lignin-derived monomers; biphenyl, benzyl phenyl ether, and 4, 4′-oxydiphenol were selected as lignin-derived dimers. Alkali lignin and glucose were also selected to determine the isolated strains' assimilative capacities.
First, 1 mL, –80 ℃ stored isolates were inoculated into 50 mL LB medium and then cultured at 30 ℃ and 250 r/min for 24 h. The cultured cells were washed thrice with sterile saline solution, and re-suspended in 4 mL sterile saline solution as seeds for the following culture. The prepared seeds were inoculated into 5 mL M9 medium (pH 11.0, supplemented with 5 mmol/L lignin-derived monomers and dimers, or 1.0 g/L alkali lignin) with the initial OD600 value of 0.2. After incubation at 30 ℃ and 250 r/min for 108 h, cell pellets were collected by centrifugation at 4 000 r/min for 10 min. The obtained cell pellets were washed trice by sterile saline solution and re-suspended in 5 mL sterile saline solution. The absorbance values of the suspensions were determined at 600 nm to indicate assimilative capacities of the isolates on lignin and lignin-derived compounds.
2.5
Investigating effects of pH on cell growths of isolated ligninolytic microbes
On the basis of growth performances on lignin and its derivatives, four isolates were selected to further test their viabilities in different pH of 6.0, 8.0, 10.0, and 11.0. Cell growth of the objective microbes was monitored when they were cultured in LB medium and M9 medium (supplemented with 10.0 g/L lignin) with different initial pH. Alkali lignin, lignosulfonic acid sodium salt, and dealkaline lignin were used to investigate the effects of pH on cell growths of the isolates. In detail, cell seeds were prepared with the above-stated procedure, and then the prepared seeds were inoculated into LB medium (initial pH of 6.0, 8.0, 10.0, or 11.0 before inoculation) or M9 medium (supplemented with 10.0 g/L lignin, initial pH of 6.0, 8.0, 10.0, and 11.0 before inoculation) at OD600 value of 0.2. During cultivation at 30 ℃ and 250 r/min, 1 mL sample was withdrawn at intervals to determine the cell growth. Particularly, due to the low solubility of alkali lignin in pH 6.0, 8.0, and 10.0 conditions, the pH values of withdrawn samples were adjusted to about 11.0 to dissolve the solid lignin fraction. Cells were obtained by centrifugation and washed thrice with sterile saline solution (pH 11.0). The washed cells were re-suspended in 1 mL sterile saline solution, and the turbidity (OD600 values) of the obtained bacterial suspension were determined by a spectrophotometer (TU-1810, Purkinje Ltd., Beijing, China). The two-dimensional heteronuclear single quantum coherence nuclear magnetic resonance (2D-HSQC NMR) was performed for alkali lignin incubated in neutral pH solution, incubated in solution of pH 11.0, and incubated in solution of pH 11.0 + Sutcliffiella sp. NC1 strain with the method described in a previous study (Cai et al., 2021).
2.6
Investigating decolorization capabilities of isolated ligninolytic microbes
As reported, most ligninolytic enzymatic systems are capable of functioning on synthetic industrial dyes and contributing to dye decolorization (Guo et al., 2021; Ali et al., 2022). Therefore, the determination of the isolates' decolorization capabilities was carried out to demonstrate the effectiveness of their ligninolytic enzymatic systems. The decolorization experiments were performed in LB medium (supplemented with 50 mg/L different types of dyes), with the initial medium pH of 7.0 or 10.0. Ten typical industrial dyes (Fig. S1) were used in this study, which were Coomassie Brilliant Blue G250 (CBB-G250), Reactive Blue 19 (RB-19), Methylene Blue (MB), Bromophenol Blue (BB), Toluidine Blue (TB), Chlorantine Fast Red 5B (CFR-5B), Congo Red (CR), Crystal Violet (CV), Amido Black 10B (AB-10B), and Malachite Green (PG). Prepared cell seeds were inoculated into LB medium (supplemented with different dyes) at the OD600 value of 0.2. The flasks were incubated at 30 ℃ and 250 r/min for 3 d, and then samples were withdrawn to determine the dye decolorization as follows. The withdrawn samples were centrifuged at 5 000 r/min for 10 min, and then the optical density of supernatants was detected by a spectrophotometer at the λmax values of tested dyes. The λmax values of applied dyes were as follows: 595 nm for CBB-G250; 595 nm for RB-19; 665 nm for MB; 590 nm for BB; 635 nm for TB; 510 nm for CFR-5B; 497 nm for CR; 590 nm for CV; 618 nm for AB-10B; 470 nm for PG. Percent decolorization was calculated by the formula: Decolorization = [(A0 – A)/A0] × 100%, where A0 is the initial absorbance of the objective dye solution, and A is the absorbance of dye solution treated with isolates for 3 d. The media contained equivalent dye and no isolates were applied as the control group.
2.7
Determining lignin peroxidase activities of isolated ligninolytic microbes
Lignin peroxidase activities of the isolated ligninolytic microbes were determined at pH values of 3.0, 7.0, and 10.0. In detail, target ligninolytic microbes were culture in LB medium at 30 ℃ and 250 r/min with an initial OD600 value of 0.2. To induce the generation of ligninolytic enzymes, 0.5 g/L alkali lignin was supplemented to the medium. Culture samples were withdrawn at 1st, 2nd, 4th, and 7th days to determine the lignin peroxidase activities. The reaction system consisted of 3.2 mL buffer solution (tartaric acid/sodium tartrate for pH 3.0, disodium hydrogen phosphate/sodium dihydrogen phosphate for pH 7.0, sodium carbonate/sodium bicarbonate for pH 10.0), 0.1 mL veratryl alcohol solution (40 mmol/L), 0.6 mL cell-free culture broth, and 0.1 mL H2O2 (4 mmol/L). Reactions were initiated by adding H2O2 and then the reaction systems were incubated at 30 ℃ for 3 min. The lignin peroxidase activity was quantified by monitoring the oxidation of veratryl alcohol to veratraldehyde at 310 nm (ε310 nm = 9 300 L/(mol·cm)). One unit of enzyme activity was defined as the amount of enzyme required to produce 1 μmol veratraldehyde from veratryl alcohol per minute.
2.8
Genome sequence and analysis for isolated ligninolytic microbes
To unravel the underlying mechanisms for the ligninolytic capability and alkali-tolerant capability of the isolated microbes, their genomes were sequenced with a combination of the third-generation single molecule sequencing of PacBio RSII single molecule real-time (SMRT) platform and the second-generation sequencing of Illumina Hiseq PE150 platform, serviced by Majorbio Company (Shanghai, China). The PacBio sequencing data were assembled by using the Unicycler (v0.4.8) hybrid assembly pipeline, and error correction of the PacBio assembly results was done using the Illumina reads by using Pilon (v1.22).
The predication of open reading frames (ORF) from assembled genome was performed by Glimmer 3.02. Genome annotation was carried out by aligning the coding protein by predicated ORF with Non-Redundant Protein Sequence Database (NR), Swiss-prot, String, and Gene Ontology (GO) databases using the BLAST 2.2.28+. Gene functions were predicated by Clusters of Orthologous Groups of proteins (COG), Kyoto Encyclopedia of Genes and Genomes (KEGG) and GO. Ribosomal RNA (rRNA) and transfer RNA (tRNA) were predicated by Barrnap 0.4.2 and tRNAscan-SE v1.3.1, respectively.
3.
Results and discussion
3.1
Effects of pH on lignin solubility and particle size distribution
Sufficient interaction of substrate with microbe and enzyme is considered a prerequisite for biodegradation, bioconversion and biocatalysis processes (Huang et al., 2022). Owing to high-molecular-weight and hydrophobic groups, most types of lignin have low solubility in aqueous solution, which reduces the interaction of lignin with ligninolytic cells and enzymes, leading to low lignin biodegradation and bioconversion efficiency. Improving lignin solubility is considered beneficial to the interaction of lignin with microbial cells and ligninolytic enzymes, and thus facilitating lignin bioconversion processes. Alkaline solution is a commonly used system to depolymerize and dissolve lignin, and several lignin valorization methods have been developed on the basis of the good solubility for lignin in alkaline solution (Liu et al., 2019). In this study, the solubility profiles of 30 g/L alkali lignin in different pH solution were determined first. As presented in Fig. 1a and 1b, when the initial pH was 7.0, only 16.6% (equivalent to 4.98 g/L) alkali lignin was dissolved in the solution. The solubility of alkali lignin significantly increased with the increased pH values of the aqueous solution. Especially, lignin solubility increased from 59.3% to 91.4% when the solution pH increased from 10.0 to 10.5. When pH 11.0 solution was used as the solvent for alkali lignin, nearly all the 30 g/L lignin was dissolved in the aqueous solution.
Fig.
1.
Effects of pH on lignin solubility and particle size distribution. (a) Solubilities of 30 g/L alkali lignin in different pH solutions; (b) Images of 30 g/L alkali lignin in different pH solutions; (c–h) Particle size distributions of the dissolved lignin in different pH solution (c, pH =7.0; d, pH =8.0; e, pH =9.0; f, pH =10.0; g, pH =10.5; h, pH =11.0); (i) Ultraviolet absorption profiles of lignin solution at initial pH of 7.0, 8.0, 9.0, 10.0, 10.5, and 11.0; (j) pH values of solution immediately added with 30 g/L and pH values after incubation at 250 r/min for 12 h; (k) pH changes during lignin solution at initial pH of 7.0, 8.0, 9.0, 10.0, 10.5, and 11.0.
Remarkably, the aqueous solution color became darker with increased lignin dissolved in the aqueous solution (Fig. 1b). The particle size distributions of the dissolved lignin in different pH solutions were also determined. The profiles of lignin particle size distributions were divided into two phases: when the solution pH increased from 7.0 to 10.0, the average diameter of dissolved lignin also significant increased. Particularly, the peak of the size distribution increased from 191.7 to 300.7 nm (Figs. 1c–1f, Table 1). The increased particle size of dissolved lignin is probably due to the fact that more high-molecular-weight lignin fractions were dissolved. Nonetheless, when the solution pH increased from 10.0 to 11.0, the particle size of dissolved lignin obviously decreased. Meanwhile, some lignin fractions with smaller particle size of 10–30 nm were generated at these conditions (Figs. 1f–1h). It was reported that high concentrations of OH– contributed to the cleavage of some lignin bonds (Xu et al., 2020), which was demonstrated by the ultraviolet wavelength scanning results that the absorption values of lignin solution at 220–240 nm and 270–290 nm were significantly decreased when the solution pH was increased to 10.5 and 11.0 (Fig. 1i).
Table
1.
Size distributions of dissolved lignin in aqueous system with different pH values.
To further demonstrate the function of OH– (introduced by NaOH) on alkali lignin, the pH values of solution immediately added with 30 g/L alkali lignin and the pH values after incubation at 250 r/min for 12 h were determined. As shown in Fig. 1j, after incubation for 12 h, the pH values of lignin-rich alkali solution were all significantly decreased, no matter the initial pH was 7.0 or 11.0. This phenomenon is similar to those observed by Zhao et al. (2021), and they attributed this phenomenon to that acidic groups in lignin (such as –COOH and Ar–OH) reacted with OH– in solution. Meanwhile, the cleavage of phenolic ether in high pH condition also provided phenolic hydroxyl groups reacting with OH− in solution. Another remarkable phenomenon is that the ΔpH values (the initial pH value minus the final pH value) increased when the pH values of solution used for lignin solubility increased from 7.0 to 10.5 (Fig. 1k), which was in line with the increased dissolved lignin. Whereas, when the pH values of solution used for lignin solubility increased from 10.5 to 11.0, the ΔpH value decreased, which was probably because that few additional lignin was dissolved in the aqueous solution in this process (there was already 91.4% lignin dissolved in aqueous solution at pH 10.5).
The above information highlighted that high pH not only contributed to high lignin solubility, but also reduced lignin particle size. To realize lignin bioconversion in alkaline solution, alkali-tolerant microbes are required. To screen microbes with traits of both ligninolytic and alkali-tolerant, lignocellulose-rich samples were inoculated into modified M9 medium with pH of 11.0 and alkali lignin as the sole carbon source. After five transfers, alkali-tolerant ligninolytic microbes were enriched, and then purified by streak plate method. Based on the morphology of emerging colonies, there were few, even only one, kinds of predominant microbes after enrichment culture. Similar with previous studies, major growing microbes belong to bacteria, which is probably ascribed to that the used mineral basal medium is beneficial to bacteria growth (Wu et al., 2022). The 16 s rDNA sequencing results suggested that some of these isolates exhibited a certain similarity in taxonomy, although the soil samples used for screening target microbes were collected from different areas. Among the thirteen purified alkali-tolerant lignin biodegradation bacteria, four strains were classified as Citricoccus sp.; three strains were classified as Alkalihalobacillus alcalophilus; three strains were classified as Sutcliffiella sp.; two strains were classified as Nesterenkonia sp.; one strain was classified as Enteractinococcus sp. (Fig. S2).
Hitherto, several ligninolytic bacteria have been isolated from diverse sites, such as leaf litter soils, sea sediment, papermaking black liquid, guts of wood-boring insects, etc., the majority of which belong to phyla Actinobacteria, Firmicutes, and Proteobacteria. Among these bacteria, the ligninolytic system of some outstanding strains has been deeply deciphered and further applied to produce value-added compounds from lignin-rich feedstock (Li et al., 2022; Liu et al., 2022). The obtained isolates in this study are significantly distinct to the previous reported ligninolytic bacteria that this is the first time to report Citricoccus sp., Nesterenkonia sp., Enteractinococcus sp., A. alcalophilus and Sutcliffiella sp. species on lignin assimilation (Fig. 2). In fact, in addition to reports on new strains identification, there have been rare reports for these five species, let alone detailed genetic information and industrial application. The discovery of these alkali-tolerant bacteria not only enlarges the species of ligninolytic microbes, but may also provide some new ligninolytic pathways for lignin bioconversion, as well as some new biological refinery routes for lignin. For instance, Halomonas sp., which can tolerate high salt concentration, has been applied as chassis for seawater-based unsterile open fermentation (Ye and Chen, 2021). With similar traits, the alkali-tolerant bacteria also have high potential as the chassis for the future unsterile open fermentation.
Fig.
2.
Maximum-likelihood tree showing evolutionary relationship between alkali-tolerant ligninolytic bacteria isolated in this study and the reported ligninolytic bacteria in previous studies.
3.3
Assimilative capabilities of isolates for lignin derivatives
Lignin biodegradation is an important constitution of global carbon cycle, without which the earth would be piled up with lignin due to the tremendous volume of lignin generated globally. Natural lignin biodegradation is generally divided into three steps: high-molecular-weight lignin is depolymerized to low-molecular-weight components (such as aromatic tripolymers, dimers and monomers); multiple low-molecular-weight linin components are funneled to few key aromatic intermediates, e.g., protocatechuic acid and catechol; aromatic nucleuses of the key intermediates are cleaved and the generated linear compounds are assimilated for cell growth (Kamimura et al., 2019; Bugg et al., 2020; Zhu et al., 2022). To test the ligninolytic capabilities of these isolates, they were individually cultured with alkali lignin, lignin-derived dimers and lignin-derived monomers as the sole carbon source. As presented in Fig. 3, all isolates utilized at least three lignin derivatives as the carbon source for cell growth. Generally, their assimilative capabilities decreased for H-lignin monomers, G-lignin monomers, and S-lignin monomers, which is in line with previous reports that methoxyl groups are among the most recalcitrant groups in lignin-derived monomers (Erickson et al., 2022; Harlington et al., 2022). For instance, when two S-lignin monomers (syringic acid and sinapic acid) were applied as substrates, there were almost no obvious cell growth for the 13 isolates in this study. In contrast, when H-lignin monomers (p-hydroxybenzonic acid and p-coumaric acid) were applied as the substrates, most tested isolates survived well on these aromatic compounds. The structural difference between "p-hydroxybenzonic acid and p-coumaric acid" and "syringic acid and sinapic acid" is that "syringic acid and sinapic acid" have two methoxy groups. In fact, mining new enzymes (Fetherolf et al., 2020; Sonoki et al., 2000) or engineering existing enzymes (Ellis et al., 2021; Harlington et al., 2022) for demethylation reactions are attracting increasing attention. Nonetheless, there are still some strains, especially the four Citricoccus sp. strains, skilling at degrading and assimilating S-lignin monomers. For instance, three of the four isolated Citricoccus sp. strains thrived on syringaldehyde and 4-hydroxy-3, 5-dimethoxybenzyl alcohol. Further genetic analysis on these isolates may provide new biocatalytic elements for O-demethylation reactions.
Fig.
3.
Cell growth of isolated ligninolytic bacteria on alkali lignin, lignin-derived dimers, lignin-derived monomers and glucose.
Interestingly, four of the 13 isolates exhibited poor or even no cell growth on glucose. As well acknowledged, glucose is the most natural available carbon source, and thus the majority of organisms have acquired glucose assimilation capability during their evolution. These isolates (such as Nesterenkonia sp. NC3) lacking of glucose assimilation capability must have undergone special events during their evolutionary processes. Recent studies revealed the underlying metabolic mechanisms for Sphingobium sp. SYK-6, an outstanding ligninolytic bacteria preferring lignin derivatives than glucose. Results highlighted that lignin-derived aromatics provide methyl groups for the biosynthesis of some essential amino acids (e.g., serine, histidine, and methionine) via tetrahydrofolate-dependent C1 metabolism in this strain (Varman et al., 2016). Nonetheless, which factors induced this evolution is still ambiguous. The glucose assimilation deficiency bacteria discovered in this study may be beneficial to uncover the key evolution events for gluose assimilation deficiency. Moreover, these glucose assimilation deficiency bacteria exhibit special potential in bioindustry. For instance, during biological detoxification processes and biopretreatment processes, engineers intend to biodegrade toxic compounds and destroy rigorous lignocellulosic structure without sacrificing glucose, and in these scenarios, microbes possessing metabolism for toxic compounds but not for glucose will show their talents (Zou et al., 2021). Taken assimilative capacities for lignin and its derivatives, Sutcliffiella sp. NC1, Citricoccus sp. NC2, Citricoccus sp. NR2, and Citricoccus sp. NT2 were selected for further investigation of their metabolic characteristics.
3.4
Cell growth of isolated ligninolytic bacteria in neutral and alkaline conditions
On the basis of their viabilities in extreme and common environments, extremophiles are divided into two categories: some can survive in both extreme and normal conditions, and others exclusively survive in harsh conditions (Nadaroglu, 2022). Here, to determine effects of pH on the isolated alkali-tolerant ligninolytic, Sutcliffiella sp. NC1 and three isolated Citricoccus sp. strains were cultured in LB medium and M9 medium (supplemented with 10 g/L alkali lignin) with initial pH of 6.0, 8.0, 10.0 and 11.0. As presented in Fig. 4a-4d, when cultured in LB medium, all the four tested bacteria survived well with the initial pH of 8.0–11.0. No cell growth was observed at pH 6.0 for Sutcliffiella sp. NC1, in contrast to the other three isolates exhibited normal cell growth under this pH condition. This suggested that Sutcliffiella sp. NC1 is an alkalophilic bacterium, instead of just an alkali-tolerant one. Moreover, the highest OD600 value was about 3.3 for Sutcliffiella sp. NC1 (cultured at an initial pH of 8.0 for 36 h), by contrast, the highest OD600 values for the three Citricoccus sp. strains were all higher than 10.0. The above phenomenon suggested that the growth of Sutcliffiella sp. NC1 is not vigorous as common microbes, even in the nutrient-rich LB medium. In fact, the cell growth of microbes and their inherent metabolisms are versatile, just like that some isolates obtained in this study are not capable of assimilating glucose well. These microbes were likely to provide distinct metabolic characteristics and genetic elements for bio-industry. The final broth pH increased or decreased to about 9.2–9.4 for all the tested four bacteria after 108 h cultivation when they were cultured in LB medium, independently of the various initial medium pH (Fig. 4i). It suggested that the four bacteria secreted some acid or alkaline compounds during their lifecycle to maintain the pH of the culture broth at about 9.2–9.4, which may be the optimum pH for their survival.
Fig.
4.
Growth profiles of Sutcliffiella sp. NC1 (a, e), Citricoccus sp. NC2 (b, f), Citricoccus sp. NR2 (c, g), and Citricoccus sp. NT2 (d, h) in LuriaBertani (LB) medium and modified M9 minimal medium with different initial pH. The pH values of culture broth at when Sutcliffiella sp. NC1, Citricoccus sp. NC2, Citricoccus sp. NR2 and Citricoccus sp. NT2 were cultured in LB medium for 108 h and cultured in modified M9 medium for 156 h (i).
When the tested four isolates were cultured in M9 medium (supplemented with 10 g/L alkali lignin), their growth profiles and final pH values were some differences to those observed in LB medium. No obvious growth was observed for all four tested bacteria at an initial pH of 6.0 (Fig. 4e–4h). The inability of these bacteria in M9 medium at pH 6.0 is correlate rather well with the nearly unchanged broth pH after 156 h incubation (Fig. 4i). These observations suggested that response of microbes to pH varies with used medium, and medium composition is an essential factor to be considered in future lignin biological conversion processes. Significant cell growth was detected for all four tested bacteria with alkali lignin as the main carbon source at initial pH of 10.0 and 11.0, further demonstrating their lignin assimilative capabilities. Particle size distributions determination of the dissolved lignin suggested that some low-molecular-weight lignin fractions disappeared after cultivation (Fig. 5, Table 2), which is consistent with the reports that the main role of ligninolytic bacteria is assimilating low-molecular-weight lignin components (Kamimura et al., 2019; Bugg et al., 2020). In addition, some lignin components with large particle size were getting reduced, which is probably because that these parts were depolymerized by the ligninolytic enzymes secreted by alkalophilic bacteria (Bugg et al., 2011; Zhang et al., 2022). The 2D-HSQC NMR results also suggested that compared with neutral pH solution, alkaline solution (pH 11.0) homogenized lignin by cleaving several linkages (Fig. 6a–6d). Incubating Sutcliffiella sp. NC1 in pH 11.0 solution further homogenized lignin components, especially some aromatic rings were cleaved because the aromatic areas in 2D-HSQC NMR are reduced (Fig. 6d, 6f).
Fig.
5.
Particle size distribution changes of alkaline lignin after incubation with Sutcliffiella sp. NC1 (a), Citricoccus sp. NC2 (b), Citricoccus sp. NR2 (c), and Citricoccus sp. NT2 (d) at initial pH value of 11.0.
Table
2.
The main peaks and their intensities of particle size distribution of alkaline lignin after incubation with the alkali-tolerant ligninolytic bacteria.
Fig.
6.
The two-dimensional heteronuclear single quantum coherence nuclear magnetic resonance (2D-HSQC NMR) spectra of alkali lignin incubated in neutral pH solution (a, b), incubated in solution of pH 11.0 (c, d), incubated in solution of pH 11.0 + Sutcliffiella sp. NC1 strain (e, f); (g) Structures identified in lignin.
Further, other two lignin sources of dealkaline lignin and lignosulfoic acid sodium salt were used as the sole carbon source to cultivate Sutcliffiella sp. NC1 and Citricoccus sp. NC2 with the initial medium pH values of 6.0, 8.0, 10.0, and 11.0, respectively. The cell growth profiles and the medium pH after 156 h cultivation were determined (Fig. S3). Similar to the results with alkali lignin as the sole carbon source, Sutcliffiella sp. NC1 and Citricoccus sp. NC2 showed better cell growth at the initial pH of 8.0–11.0, compared with the initial pH of 6.0, except that Sutcliffiella sp. NC1 did not show obvious cell growth with dealkaline lignin as the sole carbon source.
3.5
Decolorization capabilities and ligninolytic enzyme activities of isolates
A large number of ligninolytic enzymes are capable of functioning on industrial dyes due to the similar functional groups and chemical bonds between lignin and dyes. Therefore, some dyes are usually harnessed as indicators for screening ligninolytic microbes and decolorization capability are harnessed to evaluate the ligninolytic capability of relevant microbes. Herein, the decolorization capabilities on ten common industrial dyes of the isolates were assessed at pH 7.0 and 10.0. As presented in Figs. 7a and 7b, all the four isolates are capable of decolorizing the tested dyes. For instance, CBB-G250, MB, and CFR-5B are very stable in LB medium (pH 7.0) that nearly no spontaneous decolorization was observed after incubation for 3 d. Nonetheless, all the four isolates contributed to remarkable decolorization for these three stable dyes. When the initial medium pH was adjusted to 10.0, the decolorization results were different from those observed at pH 7.0. One obvious phenomenon is that spontaneous decolorization rates of some dyes changed due to the alteration of medium pH. For instance, the spontaneous decolorization of CR was 72.39% at pH 7.0, and it decreased to 6.61% when the medium pH increased to 10.0. By contrast, the spontaneous decolorization of BB was 9.20% with pH 7.0, and it increased to 42.00% when the medium pH increased to 10.0. The influence of pH on dye spontaneous decolorization is mainly due to the altered ionization degree (Donadelli et al., 2018). Regardless of the varied spontaneous decolorization, all the tested four bacteria significantly promote the decolorization of the tested dyes at an initial pH of 10.0 and some better decolorization results were achieved compared with the initial pH of 7.0 conditions. For instance, Sutcliffiella sp. NC1 contributed to 35.36% decolorization of CBB-G250 at initial pH of 7.0, and this value increased to 78.62% when the initial medium pH was 10.0. Among the four tested bacteria, Sutcliffiella sp. NC1 exhibited the most excellent decolorization capability at pH 10.0 that it contributed to the highest decolorization ratios to five of the ten applied dyes (Fig. 7b), although its cell growth was the weakest among these four strains. With the above information, all the four isolates are suitable for treating colored effluents, especially those with high pH, because most reported microbes with decolorization capabilities can not survive in alkaline pH condition (Xu et al., 2018).
Fig.
7.
Dye decolorization capabilities and lignin peroxidase activities of isolates. Decolorization capabilities of Sutcliffiella sp. NC1, Citricoccus sp. NC2, Citricoccus sp. NR2 and Citricoccus sp. NT2 with initial medium pH of 7.0 (a) and initial medium pH of 10.0 (b); Lignin peroxidase activities of Sutcliffiella sp. NC1 (c), Citricoccus sp. NC2 (d), Citricoccus sp. NR2 (e) and Citricoccus sp. NT2 (f) determined in the system pH of 3.0, 7.0 and 10.0.
To further demonstrate the ligninolytic capability of these isolates, their lignin peroxidase activity was determined at pH of 7.0 and 10.0. Lignin peroxidases (EC 1.11.1.14) catalyze the oxidation of various phenolic substrates with H2O2 as electron acceptor and are commonly served as the indicator enzyme for lignin bio-decomposition. As referenced, the optimum pH for common lignin peroxidase activity is among 2.0–5.0, and the pH of commonly used for lignin peroxidase activity determination is about 3.0 (Pham et al., 2021; Singh et al., 2021). Here, lignin peroxidase activities of the four isolates were detected at pH 3.0, 7.0 and 10.0 (Figs. 7c–7f). Particularly, the optimum pH of lignin peroxidases of Sutcliffiella sp. NC1 is 10.0, which is significantly different to those of other three Citricoccus sp. strains, whose highest lignin peroxidase activities were detected at pH 3.0. The Sutcliffiella sp. NC1 is an alkalophilic bacterium that it even can not survive in pH 6.0. Therefore, enzymes generated by Sutcliffiella sp. NC1 must adapt to alkaline conditions, especially extracellular enzymes. In addition to lignin peroxidases, it is predicted that many other enzymes in Sutcliffiella sp. NC1 also function well in alkaline conditions, making this bacterium a potential treasure for mining alkaline specific enzymes.
3.6
Unraveling ligninolytic and alkaline-tolerant mechanisms by genome analysis
On the basis of considerations of the cell growth profiles in different pH conditions, the assimilative capacities for lignin and its derivatives, the decolorization capabilities, and lignin peroxidase activities, Sutcliffiella sp. NC1 was considered the most outstanding strains among isolated alkali-tolerant ligninolytic microbes. As referenced, in 2020, the evolutionary relationships of Bacillus species were systematically clarified via phylogenomic and comparative analyses on more than 300 genomes, and 17 distinct Bacillus species clades were reclassified as novel Bacillaceae genera. By this time, the genus Sutcliffiella was proposed based on the name of the British microbiologist Prof. Iain Sutcliffe for his excellent works on microbial envelopes and lipoproteins, as well as his contributions to microbial taxonomy (Gupta et al., 2020). Although the thermophilic trait has been confirmed for some species in genus Sutcliffiella, this is the first report for the alkali-tolerant and ligninolytic traits for this genus. To better apply this alkalophilic bacterium in lignin bioconversion processes, a deep understanding of its biochemical and genetic basis is essential. Hence, its genome was sequenced and analyzed to unravel the underlying ligninolytic and alkali-tolerant mechanisms. The detailed genome information for Sutcliffiella sp. NC1 has been deposited in US National Center for Biotechnology Information (NCBI) with the accession number of CP115153. The general genome information for Sutcliffiella sp. NC1 is presented in Fig. 8 and Table 3. Here, we focused on the genetic basis for their ligninolytic and alkaline-tolerant capabilities.
Fig.
8.
The circle map of Sutcliffiella sp. NC1 genome. The six circles from outer to inner refer to genome size, protein-coding genes on the positive strand, CDs on negative chains, non-coding RNAs (rRNA and tRNA), GC contents, and GC skew value.
According to the functional annotation of the predicted genes, genes implicated in depolymerizing high-molecular-weight lignin, funneling diverse aromatic compounds to target aromatic intermediate, and cleaving aromatic ring for further metabolisms were all discovered in Sutcliffiella sp. NC1 (Table 4). In detail, the genes encoding typical lignin-degrading enzymes, including laccase, manganese peroxidase, dye-decolorizing peroxidase, and some versatile peroxidases, were all discovered in the genome of Sutcliffiella sp. NC1. Multiple enzymes implicated in funneling diverse aromatic compounds to target aromatic intermediate were also annotated from Sutcliffiella sp. NC1, such as alcohol dehydrogenase and aldehyde dehydrogenase responsible for the interconversion of aromatic acids, aromatic aldehydes, and aromatic alcohols; Decarboxylases, monooxygenase, and, responsible for the decarboxylation, hydroxylation and O-demethylation of lignin-derived aromatic monomers (Erickson et al., 2022); Ring-cleaving dioxygenases responsible for converting lignin-derived aromatics to linear compounds, and enzymes for further assimilating the generated linear compounds, such as enzymes in β-ketoadipate pathway. Moreover, biodegradation pathways for 4-hydroxyphenylpyruvate, 4-hydroxyphenylacetate, and phenylacetate were also annotated from Sutcliffiella sp. NC1, making this alkali-tolerant bacterium not only a competitive candidate for lignin valorization, but also a promising candidate for bioremediation.
Table
4.
Possible ligninolytic enzymes in Sutcliffiella sp. NC1.
Proteins
Non-redundant (NR) hits
Hit-description
Possible functions
Possible enzymes involved in lignin depolymerization
Laccase
WP_066412019.1 WP_066417352.1
Multicopper oxidase domain-containing protein
The most popular enzyme involved in lignin decomposition. It degrades both β−1 and β-O-4 dimers via Cα-Cβ cleavage, Cα oxidation and alkyl-aryl cleavage. It also performs aromatic ring cleavage and oxidize non-phenolic compounds when primary mediators are co-present.
One popular lignin-modifying peroxidase. It oxidizes Mn2+ into highly reactive Mn3+, which is stabilized by fungal chelators. Chelated Mn3+ in turn acts as diffusible redox-mediator that attacks phenolic lignin structures.
O-methyltransferase
WBL13923.1
O-methyltransferase
O-methyltransferase can eliminate the inhibiting effect of free-hydroxyl groups present at both ends of the lignin molecule and resulted in release of products from lignin degradation.
Other peroxidases
WP_066416151.1
Catalase/peroxidase
Some bacterial catalases/peroxidases catalyze the degradation of lignin-related compounds, such as some aromatic dimers.
WP_091696999.1
Superoxide dismutase
It catalyzes dismutation of the superoxide (O2–) radical into either O2 or H2O2, which are crucial for lignin depolymerization. Some superoxide dismutases have potentials to breakdown C–C, aryl-C bonds and remove methyl groups.
WP_010145496.1
Thioredoxin-dependent thiol peroxidase
They play important roles in protecting microbes from oxidative damages by detoxifying reactive intermediates formed during metabolism of aromatic compounds.
WP_191, 808, 760.1
Thiol peroxidase
WP_066421946.1
Dye-decolorizing peroxidase
It can decolorize multiple industrial dyes, catalyze the oxidation of various aromatic substrates, as well as depolymerize lignin, lignin-derived β-O-4 dimers and β-aryl ether dimers.
Possible enzymes involved in funneling diverse aromatic compounds to target aromatic intermediate
4-hydroxyphenylacetate 3-monooxygenase
WP_066421294.1
4-hydroxyphenylacetate 3-monooxygenase
It catalyzes 4-hydroxyphenylacetate to 3, 4-dihydroxyphenylacetate
4-hydroxyphenylpyruvate dioxygenase
WP_066419374.1
4-hydroxyphenylpyruvate dioxygenase
It catalyzes 4-hydroxyphenylpyruvate to homogentisate
2-polyprenyl-6-methoxyphenol hydroxylase
AST94230.1
2-polyprenyl-6-methoxyphenol hydroxylase
It catalyzes 2-polyprenyl-6-methoxyphenol to 2-methoxy-6-all-trans-polyprenyl-1, 4-benzoquinol
Phenylacetate-coa oxygenase subunits paac and paaj; 1, 2-phenylacetyl-coa epoxidase subunits A and B; phenylacetate degradation operon negative regulatory protein paax
It catalyzes biodegradation of phenylacetate.
FAD-dependent monooxygenase
WP_084380641.1
p-hydroxybenzoate hydroxylase
It catalyzes hydroxylation of Benzene ring (the most likely substrate is p-hydroxybenzoate, and it converts p-hydroxybenzoate to protocatechuate)
FAD-dependent monooxygenase
WP_066411735
p-hydroxybenzoate hydroxylase
Prephenate dehydratase
WP_066410993.1
Prephenate dehydratase
It catalyzes prephenate to phenylpyruvate.
Prephenate dehydrogenase
WP_066416897.1
Prephenate dehydrogenase
It catalyzes prephenate to 4-hydroxyphenylpyruvate.
Pyridoxal-dependent decarboxylase
WBL17492.1
Pyridoxal-dependent decarboxylase
It may catalyze decarboxylation reaction of aromatic acids.
NAD(P)-dependent alcohol dehydrogenase
WBL16690.1 WBL16773.1
NAD(P)-dependent alcohol dehydrogenase
It may catalyze interconversion of aromatic acids, aromatic aldehydes, and aromatic alcohols.
Iron-containing alcohol dehydrogenase
WBL14293.1
Iron-containing alcohol dehydrogenase
Aldehyde dehydrogenase
WBL14464.1
Aldehyde dehydrogenase
O-demethylase
WBL15586.1 WBL13758.1
Cytochrome P450
It may catalyze O-demethylation reactions of G- and S- type lignin-derived monomers.
Possible enzymes involved in cleaving aromatic rings and further metabolism
In addition to ligninolytic genes, genes responsible for the alkali-tolerant characteristic were also searched throughout the genome of Sutcliffiella sp. NC1. Based on multi-omics, physiological and genetic studies on alkali-tolerant or alkaliphilic microbes, four main mechanisms for alkaline pH homeostasis have been proposed, including (a) increasing acidic metabolites production via amino acid deaminases and/or sugar fermentation; (b) increasing adenosine triphosphate (ATP) synthase responsible for H+ entry to ATP generation; (c) changing cell surface properties; (d) increasing monovalent cation/proton antiporter activities, among which, monovalent cation/proton antiporters play the dominant role in most alkali-tolerant bacteria (Padan et al., 2005; Somayaji et al., 2022). For Sutcliffiella sp. NC1, abundant cation/proton antiporters are annotated from its genome, including eleven Na+/H+ antiporters, two K+/H+ antiporters, one Ca2+/H+ antiporter, and one cation/proton antiporter with unconfirmed cation types (Table 5). These cation/proton antiporters are proposed to diminish the establishment of pH gradients or to induce dissipation of the gradients by proton ATPases. Moreover, some other potential proteins for alkali-tolerate characteristics were also found in Sutcliffiella sp. NC1, such as several deaminases for acid metabolites generation, cardiolipin synthase for alkali-tolerate cell surface feature, H+-coupled flagellar motor for diminishing the establishment of a pH gradient, etc. Due to the few studies on Sutcliffiella species, let alone their alkalophilic characteristic, detailed function confirmation for the above-mentioned potential proteins will expand our understanding on the alkali-tolerate mechanisms of this strain, as well as provide excellent genetic elements for designing robust microbial cell factories.
Table
5.
Possible cation/proton antiporters responsible for alkali-tolerant in Sutcliffiella sp. NC1.
Proteins
NR hits
Hit-description
Possible functions
Na+/H+ antiporter
WBL15915.1
Na+/H+ antiporter NhaC
Na+/H+ antiporters are membrane proteins that play a major role in pH and Na+ homeostasis of cells throughout biological kingdom, from bacteria to humans and higher plants.
WBL16672.1
WBL16852.1
WBL14706.1
WBL14352.1
WBL14286.1
Na+/H+ antiporter subunit A
WBL14287.1
Na+/H+ antiporter subunit B
WBL14288.1
Na+/H+ antiporter subunit C
WBL14289.1
Na+/H+ antiporter subunit D
WBL14290.1
Na+/H+ antiporter subunit F1
WBL14292.1
Monovalent cation/H(+) antiporter subunit G
K+/H+ antiporter
WBL16037.1
K+/H+ antiporter
Function of K+/H+ antiporters is similar to that of Na+/H+ antiporters except that pH homeostasis depends on K+ efflux.
WBL16481.1
Ca2+/H+ antiporter
WBL15961.1
Ca2+/H+ antiporter
The Ca2+/H+ antiporters play an important role in maintaining cellular Ca2+ homeostasis in bacteria, yeast, and plants by promoting Ca2+ efflux across the cell membranes.
Other putative cation/proton antiporters
WBL16281.1
Putative cation/proton antiporter
Function is similar to above mentioned cation/proton antiporters, and it may use other cation effluxes in addition to Na+, K+, and Ca2+.
Different to many ligninolytic bacteria can funnel lignin components to polyhydroxyalkanoates, we did not detect the accumulation of this biopolymer in Sutcliffiella sp. NC1. This result is consistent with the genome in formation of Sutcliffiella sp. NC1 that no special gene clusters for polyhydroxyalkanoates biosynthesis are found in this strain. In this case, it was assumed that the consumed lignin is converted to the bacterial biomass, which is mainly composed by protein, lipids, polysaccharides, vitamins, etc. In other scenarios, if suitable metabolic engineering operations were applied for Sutcliffiella sp. NC1, the biodegradation pathways are blocked at some special nodes and thus target compounds will be accumulated. For instance, there are many ring-cleaving dioxygenases in Sutcliffiella sp. NC1 genome, and the deletion of these genes can result in the accumulation of special aromatic monomers from lignin (Cai et al., 2021; Liu et al., 2022).
4.
Conclusions
Towards the end, a new lignin bioconversion system was constructed by combining the advantages of high lignin solubility in alkaline solution and alkali-tolerant ligninolytic bacteria. In particular, several alkali-tolerant ligninolytic bacteria were isolated, and this is the first report on the ligninolytic capabilities of Citricoccus sp., Nesterenkonia sp., Enteractinococcus sp., A. alcalophilus and Sutcliffiella sp. species. Among these isolates, Sutcliffiella sp. NC1 was demonstrated as an alkalophilic bacterium, and its lignin peroxidases exhibited the highest activity in alkaline conditions. Moreover, the alkali-tolerant and ligninolytic mechanisms of Sutcliffiella sp. NC1 were preliminarily on the basis of its genome information.
Acknowledgements:
This work was supported by National Key R&D Program of China (No. 2021YFC2101301) and National Natural Science Foundation of China (No. 22278227).
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Table
2.
The main peaks and their intensities of particle size distribution of alkaline lignin after incubation with the alkali-tolerant ligninolytic bacteria.
Table
4.
Possible ligninolytic enzymes in Sutcliffiella sp. NC1.
Proteins
Non-redundant (NR) hits
Hit-description
Possible functions
Possible enzymes involved in lignin depolymerization
Laccase
WP_066412019.1 WP_066417352.1
Multicopper oxidase domain-containing protein
The most popular enzyme involved in lignin decomposition. It degrades both β−1 and β-O-4 dimers via Cα-Cβ cleavage, Cα oxidation and alkyl-aryl cleavage. It also performs aromatic ring cleavage and oxidize non-phenolic compounds when primary mediators are co-present.
One popular lignin-modifying peroxidase. It oxidizes Mn2+ into highly reactive Mn3+, which is stabilized by fungal chelators. Chelated Mn3+ in turn acts as diffusible redox-mediator that attacks phenolic lignin structures.
O-methyltransferase
WBL13923.1
O-methyltransferase
O-methyltransferase can eliminate the inhibiting effect of free-hydroxyl groups present at both ends of the lignin molecule and resulted in release of products from lignin degradation.
Other peroxidases
WP_066416151.1
Catalase/peroxidase
Some bacterial catalases/peroxidases catalyze the degradation of lignin-related compounds, such as some aromatic dimers.
WP_091696999.1
Superoxide dismutase
It catalyzes dismutation of the superoxide (O2–) radical into either O2 or H2O2, which are crucial for lignin depolymerization. Some superoxide dismutases have potentials to breakdown C–C, aryl-C bonds and remove methyl groups.
WP_010145496.1
Thioredoxin-dependent thiol peroxidase
They play important roles in protecting microbes from oxidative damages by detoxifying reactive intermediates formed during metabolism of aromatic compounds.
WP_191, 808, 760.1
Thiol peroxidase
WP_066421946.1
Dye-decolorizing peroxidase
It can decolorize multiple industrial dyes, catalyze the oxidation of various aromatic substrates, as well as depolymerize lignin, lignin-derived β-O-4 dimers and β-aryl ether dimers.
Possible enzymes involved in funneling diverse aromatic compounds to target aromatic intermediate
4-hydroxyphenylacetate 3-monooxygenase
WP_066421294.1
4-hydroxyphenylacetate 3-monooxygenase
It catalyzes 4-hydroxyphenylacetate to 3, 4-dihydroxyphenylacetate
4-hydroxyphenylpyruvate dioxygenase
WP_066419374.1
4-hydroxyphenylpyruvate dioxygenase
It catalyzes 4-hydroxyphenylpyruvate to homogentisate
2-polyprenyl-6-methoxyphenol hydroxylase
AST94230.1
2-polyprenyl-6-methoxyphenol hydroxylase
It catalyzes 2-polyprenyl-6-methoxyphenol to 2-methoxy-6-all-trans-polyprenyl-1, 4-benzoquinol
Phenylacetate-coa oxygenase subunits paac and paaj; 1, 2-phenylacetyl-coa epoxidase subunits A and B; phenylacetate degradation operon negative regulatory protein paax
It catalyzes biodegradation of phenylacetate.
FAD-dependent monooxygenase
WP_084380641.1
p-hydroxybenzoate hydroxylase
It catalyzes hydroxylation of Benzene ring (the most likely substrate is p-hydroxybenzoate, and it converts p-hydroxybenzoate to protocatechuate)
FAD-dependent monooxygenase
WP_066411735
p-hydroxybenzoate hydroxylase
Prephenate dehydratase
WP_066410993.1
Prephenate dehydratase
It catalyzes prephenate to phenylpyruvate.
Prephenate dehydrogenase
WP_066416897.1
Prephenate dehydrogenase
It catalyzes prephenate to 4-hydroxyphenylpyruvate.
Pyridoxal-dependent decarboxylase
WBL17492.1
Pyridoxal-dependent decarboxylase
It may catalyze decarboxylation reaction of aromatic acids.
NAD(P)-dependent alcohol dehydrogenase
WBL16690.1 WBL16773.1
NAD(P)-dependent alcohol dehydrogenase
It may catalyze interconversion of aromatic acids, aromatic aldehydes, and aromatic alcohols.
Iron-containing alcohol dehydrogenase
WBL14293.1
Iron-containing alcohol dehydrogenase
Aldehyde dehydrogenase
WBL14464.1
Aldehyde dehydrogenase
O-demethylase
WBL15586.1 WBL13758.1
Cytochrome P450
It may catalyze O-demethylation reactions of G- and S- type lignin-derived monomers.
Possible enzymes involved in cleaving aromatic rings and further metabolism
Table
5.
Possible cation/proton antiporters responsible for alkali-tolerant in Sutcliffiella sp. NC1.
Proteins
NR hits
Hit-description
Possible functions
Na+/H+ antiporter
WBL15915.1
Na+/H+ antiporter NhaC
Na+/H+ antiporters are membrane proteins that play a major role in pH and Na+ homeostasis of cells throughout biological kingdom, from bacteria to humans and higher plants.
WBL16672.1
WBL16852.1
WBL14706.1
WBL14352.1
WBL14286.1
Na+/H+ antiporter subunit A
WBL14287.1
Na+/H+ antiporter subunit B
WBL14288.1
Na+/H+ antiporter subunit C
WBL14289.1
Na+/H+ antiporter subunit D
WBL14290.1
Na+/H+ antiporter subunit F1
WBL14292.1
Monovalent cation/H(+) antiporter subunit G
K+/H+ antiporter
WBL16037.1
K+/H+ antiporter
Function of K+/H+ antiporters is similar to that of Na+/H+ antiporters except that pH homeostasis depends on K+ efflux.
WBL16481.1
Ca2+/H+ antiporter
WBL15961.1
Ca2+/H+ antiporter
The Ca2+/H+ antiporters play an important role in maintaining cellular Ca2+ homeostasis in bacteria, yeast, and plants by promoting Ca2+ efflux across the cell membranes.
Other putative cation/proton antiporters
WBL16281.1
Putative cation/proton antiporter
Function is similar to above mentioned cation/proton antiporters, and it may use other cation effluxes in addition to Na+, K+, and Ca2+.
Fig. 1. Effects of pH on lignin solubility and particle size distribution. (a) Solubilities of 30 g/L alkali lignin in different pH solutions; (b) Images of 30 g/L alkali lignin in different pH solutions; (c–h) Particle size distributions of the dissolved lignin in different pH solution (c, pH =7.0; d, pH =8.0; e, pH =9.0; f, pH =10.0; g, pH =10.5; h, pH =11.0); (i) Ultraviolet absorption profiles of lignin solution at initial pH of 7.0, 8.0, 9.0, 10.0, 10.5, and 11.0; (j) pH values of solution immediately added with 30 g/L and pH values after incubation at 250 r/min for 12 h; (k) pH changes during lignin solution at initial pH of 7.0, 8.0, 9.0, 10.0, 10.5, and 11.0.
Fig. 2. Maximum-likelihood tree showing evolutionary relationship between alkali-tolerant ligninolytic bacteria isolated in this study and the reported ligninolytic bacteria in previous studies.
Fig. 3. Cell growth of isolated ligninolytic bacteria on alkali lignin, lignin-derived dimers, lignin-derived monomers and glucose.
Fig. 4. Growth profiles of Sutcliffiella sp. NC1 (a, e), Citricoccus sp. NC2 (b, f), Citricoccus sp. NR2 (c, g), and Citricoccus sp. NT2 (d, h) in LuriaBertani (LB) medium and modified M9 minimal medium with different initial pH. The pH values of culture broth at when Sutcliffiella sp. NC1, Citricoccus sp. NC2, Citricoccus sp. NR2 and Citricoccus sp. NT2 were cultured in LB medium for 108 h and cultured in modified M9 medium for 156 h (i).
Fig. 5. Particle size distribution changes of alkaline lignin after incubation with Sutcliffiella sp. NC1 (a), Citricoccus sp. NC2 (b), Citricoccus sp. NR2 (c), and Citricoccus sp. NT2 (d) at initial pH value of 11.0.
Fig. 6. The two-dimensional heteronuclear single quantum coherence nuclear magnetic resonance (2D-HSQC NMR) spectra of alkali lignin incubated in neutral pH solution (a, b), incubated in solution of pH 11.0 (c, d), incubated in solution of pH 11.0 + Sutcliffiella sp. NC1 strain (e, f); (g) Structures identified in lignin.
Fig. 7. Dye decolorization capabilities and lignin peroxidase activities of isolates. Decolorization capabilities of Sutcliffiella sp. NC1, Citricoccus sp. NC2, Citricoccus sp. NR2 and Citricoccus sp. NT2 with initial medium pH of 7.0 (a) and initial medium pH of 10.0 (b); Lignin peroxidase activities of Sutcliffiella sp. NC1 (c), Citricoccus sp. NC2 (d), Citricoccus sp. NR2 (e) and Citricoccus sp. NT2 (f) determined in the system pH of 3.0, 7.0 and 10.0.
Fig. 8. The circle map of Sutcliffiella sp. NC1 genome. The six circles from outer to inner refer to genome size, protein-coding genes on the positive strand, CDs on negative chains, non-coding RNAs (rRNA and tRNA), GC contents, and GC skew value.
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