- Journal Details
- Format
- Journal
- eISSN
- 2544-4646
- First Published
- 04 Mar 1952
- Publication timeframe
- 4 times per year
- Languages
- English
Lignin is widely distributed in various plants and is the second most abundant natural organic polymer on the Earth (Zakzeski et al. 2010; Ragauskas et al. 2014). It is an amorphous and complex aromatic compound with a substantial molecular weight, which is mainly composed of three main lignin precursors (p-hydroxyphenyl, guaiacyl, and syringyl units) linked via C-C or C-O bonds formed by radical coupling reactions. It contains a variety of oxygen-containing functional groups, such as methoxy, hydroxyl, carboxyl, and other active structures (Zhu et al. 2017; Nishimura et al. 2018). In the plant cells, lignin can be converted via phenylalanine and tyrosine by transamination (Hatfield et al. 2017; Kang et al. 2019). Then, they are joined by chemical bonds such as ester bonds to form highly polymerized macromolecules. The long chains of cellulose are twisted into externally hydrophobic microfibrils, and then the lignin is combined with the allosteric hemicellulose by an electrostatic action. Hemicellulose bridges the hydrophobic regions of cellulose microfibrils, forming complex lignocellulosic composites (Kang et al. 2019). Hence, natural lignin could not be easily degraded in papermaking wastewater, agricultural straw returning, and clean biomass energy development. Because of their complicated structure, a significant obstacle obstructs the development and utilization of natural biomass energy such as straw, forage, and woody feeds.
A growing number of bacteria with lignin degradability have been discovered in recent years. Scientists realized that bacteria play an important role in the lignin industrial utilization process (Bugg et al. 2011a). Ligninolytic bacteria have extensive adaptability in industry and agriculture. Simultaneously, lignin waste can be converted to various value-added products by bacteria (Xu et al. 2018). Besides, both lignin peroxidase and manganese oxide enzyme are lignin-degrading proteases containing ferrous-ions, the former degrading the hydroxyl-containing aromatic ring inside the lignin, and the latter removing the methoxy group on the ring, then, make it easy to enter the next step of degradation (Bugg et al. 2011b; Bharagava et al. 2018). Many aerobic bacteria could degrade lignin, which belongs mainly to
The purpose of this study was to characterize and identify bacteria, which degrade lignin in nature, and to compare their degradation capabilities and analyze biodegradation products. Moreover, it could enjoy a broad array of uses in the comprehensive utilization of lignin resources in industry and agriculture.
10 g of each sample was placed in a sterile Erlenmeyer flask, and 50 ml of 0.9% physiological saline was added, then stirred violently by a vortex mixer. The mixed liquids were diluted by five levels (1 × 10–1, 1 × 10–2, 1 × 10–3, 1 × 10–4, 1 × 10–5), and 100 μl was inoculated into the ligninolytic selection medium. The medium contained 2.0 g (NH4)2SO4, 0.5 g MgSO4, 1.0 g K2HPO4, 0.5 g NaCl, 5.0 g alkaline lignin, 20.0 g agar powder, 1.0 l H2O. All the chemicals were purchased from Solarbio Biotechnology Co., Ltd. In the ultra-clean bench, it was coated with a triangular glass rod, and finally, the plate was inverted and cultured at 30°C for 48 h. Finally, single colonies were inoculated into LB tube medium.
The decolorization of aniline blue proved that bacteria could produce ligninolytic enzymes. Also, the decolorization of brilliant blue was considered as the laccase production in the plate. Therefore, 1% aniline blue or brilliant blue was added into the alkaline lignin medium by a sterile filter (0.45 μm), respectively. Afterward, the bacteria, for which had the hydrolysis circle was observed, were selected and purified. Finally, they were stored at –20°C with liquid paraffin.
The total DNA of the strains was extracted using the Omega Bacterial Genomic DNA Extraction Kit, and then the 16S rDNA sequence of the bacterial PCR was amplified by the 27F and 1492R primers as follows: 27F: (5’-AGAGTTTGATCCTGGCTCAG-3’), 1492R: (5’-TACGGCTACCTTGTTACGACTT-3’). The 16S rRNA amplicons were sequenced by the Nanchang Kechang Biotechnology Company.
The 16S rDNA sequences of the identified strains were imported into MEGA 7.0 for phylogenetic analysis, and the phylogenetic tree was constructed by the Neighbor-joining method.
The shape of bacteria was observed via scanning electron microscopy. 2 ml of the fermentation broth was added to the centrifuge tube, centrifuged at 8,000 g for 3 min at 4°C, the supernatant liquid was discarded, the precipitate was washed by adding a phosphate buffer solution (pH 7.2), and then centrifuged again. After repeating three times, the supernatant was discarded. 2.5% glutaraldehyde was added and fixed in a refrigerator at 4°C for 24 h. The ethanol was used for the gradient (30%, 50%, 70%, 80%, 90%) dehydration treatment. After gradient dehydration centrifugation, it was eluted twice with absolute ethanol, and the supernatant was discarded by centrifugation, and the bacteria were resuspended in absolute ethanol. The coverslips were immersed in 1 M HCl solution for 12 h, and the coverslips were washed with absolute ethanol, sonicated for 30 min and dried. 5–10 μl of the resuspended bacterial liquid was pipetted and added to cover glass. After drying, the sample was observed by scanning electron microscopy.
The Lip (lignin peroxidase) and Mnp (manganese peroxidase) of the bacteria were determined at 24 h, 48 h, and 72 h, respectively.
Lip activity was detected with the lignin peroxidase kit (Beijing Solabao Technology Co., Ltd.). Lip oxidized resveratrol to produce veratraldehyde with a specific absorption peak at 310 nm (Yang et al. 2017; Zhou et al. 2017). The bacterial suspension was centrifuged at 10,000 g for 10 min, and the supernatant was placed in a 2 ml centrifuge tube on ice for testing. The reaction system contained 1 mM resveratrol, 50 mM phosphate buffer (PBS), pH 7.2, and 0.1 mM hydrogen peroxide, and the supernatant was added in a volume of 100 μl. Ultrapure water was used as a control to measure the 10 S and 310 S absorbance at 310 nm which was recorded as A1 and A2, and ΔA = A2 – A1. The one enzyme activity unit was defined as the amount of enzyme required to oxidize 1 nmol of resveratrol per liter of culture suspension, and the molar extinction coefficient of veratraldehyde, ε1 was equal to 9,300 l · mol–1 · cm–1.
Mnp is also an oxidase that contains heme, which is oxidized with guaiacol to tetra-o-methoxyphenol in the presence of Mn2+ and has a characteristic absorption peak at 465 nm (Hwang et al. 2008). Mnp activity was detected by the manganese peroxidase kit (Beijing Suo Laibao Technology Co., Ltd.). The culture was centrifuged at 10,000 g for 10 min. The supernatant was placed on ice to be tested as a crude enzyme solution. A 100 μl sample and 900 μl of the substrate were thoroughly mixed in a 1 ml glass cuvette as a reaction system. After incubation at 37°C for 10 min, the absorbance at 465 nm was measured to calculate the difference ΔA, and ultrapure water was used as a control. An enzyme activity unit was defined as the amount of enzyme required to oxidize 1 nmol of guaiacol per minute per liter of the culture medium. The guaiacol extinction coefficient ε2 was equal to 12,100 l · mol–1 · cm–1, and the calculation formula was as follows:
d – cuvette light path, VA – total reaction volume, VS – sample volume in the reaction, T – reaction time.
The pH meter was calibrated, and the pH of the supernatant was determined and recorded. The supernatant’s pH was adjusted to about 2.0 with a concentrated hydrochloric acid (38%). After adjusting the pH, each broth was extracted using accelerated solvent extraction (ASE300). Using diatomaceous earth solidified one ml of the liquid, and solids were added. In the extraction vessel, ethyl acetate was used as an extractant, and after extraction, ethyl acetate was blown off using a nitrogen purifier to obtain an extracted product. After nitrogen drying, the product was dissolved in 100 μl of cyclohexane, dioxane, and ethyl acetate. At the same time, 50 μl of a trimethylsilyl trifluoroacetamide (BSTFA (N,O-bis(trimethylsilyl)) was added to facilitate measurement.
The dissolved sample was analyzed by GC-MS. The liquid was injected using Thermo Trace 1300 ISQ, the injection volume was 1 μl on the OM-5MS capillary column, the carrier gas was He gas, and the flow rate was controlled at 1 ml · min–1. The inlet temperature was set to 200°C, the column temperature was kept at 50°C for 4 min, then raised to 220°C for 25 min, the solvent delay time was 3 min, the transmission line and ion source temperature were set to 230°C, and 250°C, respectively (Barros et al. 2013). Electron ionization mass spectra were recorded in a Full Scan mode. The degradation product’s chemical structure was presumed based on the material retention time, electron mass spectrometry, and the NIST database.
Screening and identification of bacterial isolates based on different media.
| Name | Species | Accession number | Alkaline lignin | Aniline blue | Brilliant blue |
|---|---|---|---|---|---|
| YB5 | MT745877 | ∘ | + | – | |
| CL32 | MT745880 | ∘ | + | – | |
| LN2 | MT745878 | ∘ | ++ | – | |
| LN4 | MT745879 | ∘ | ++ | – |
The phylogenetic tree constructed by the neighbor-joining method is shown in Fig. 1A. As it is visible in Fig. 1B, the bacteria of YB5 strains were long rodshaped under SEM and with a length of 2.5 μm, and seemed to be splitting. The cells of both LN2 and LN4 strains were long rod-shaped, with a length of 1–1.5 μm. However, CL32 was a globular bacterium with a sphere diameter of fewer than 1 μm.
Fig. 1.
(A) Phylogenetic tree of four strains. Strains in this study were marked with a red triangle (
Fig. 2.
(A) Bubble chart about colony size on alkaline lignin medium and OD600 in the alkaline lignin liquid medium of four strains. The bubble center points indicate the OD600 values, and the bubble size indicates the colony size, but it is not an isometric diagram. (B) The degradation rate of alkaline lignin. After the bacteria were cultured in an alkaline lignin medium for three days, the alkaline lignin degradation rates of strains were calculated (the OD value determined the degradation rate).
Degradation of alkaline lignin is a biological process involving a reduction in carbon atoms and a decrease in molecular weight. It includes both incomplete degradation of macromolecules into small molecules, and the complete degradation of macromolecules into carbon dioxide and water. In this study, alkaline lignin was decomposed into small molecules or fully utilized to reduce the content of alkaline lignin in the environment. The histogram of the degradation rate of alkaline lignin was shown in Fig. 2B. The degradation rate of strains was determined after 72 h incubation in the alkaline lignin liquid medium. The results showed that the degradation rate of the YB5 strain reached 53.5%, it was slightly lower for LN4 than YB5 strain, and the degradation rate of CL32 was less than 40%, but the alkaline lignin degradation of LN2 strains was as low as 38.8%.
Both Lip and Mnp were essential oxidases in the lignin degradation process. The strains in this study failed to decolorize brilliant blue in the alkaline lignin selective medium. So, the enzyme activity of Lac was not determined. The dynamics of Lip and Mnp enzyme activity during three days were shown in Fig. 3.
Fig. 3.
Lip (A) and Mnp (B) activity dynamic of four strains. The color band around the line indicates the standard deviation. The width indicates the level of the standard deviation value. “a, b, c” indicates that the enzyme activity in the different periods was significantly different, and the same letter indicates any significant difference. “ND” – not detected. All activity assays were obtained from triplicate experiments.
The Lip activity of four strains was the highest on the third day (Fig. 3A). The YB5 strains enzyme activity continued to rise, and the difference between the third day and the second day was not significant (
The Mnp activity dynamic trend of YB5 and LN4 strains continued to increase and were the highest on the second and third day, respectively. This activity was significantly higher than on the first day (
Ethyl acetate, hexane and 1,4-dioxane are common chromatographic solvents. Hexane is a non-polar solvent, but dioxane is a polar solvent. Finally, the ethyl acetate polarity is normal. Due to different chemical polarity, these solvents are often selected for the detection of products of different-polarity. Characteristics of the products measured in different polar solvents are shown in Table II. Four strains were cultured for 72 h in the alkaline lignin medium and then treated to determine degradation products.
The compounds identified in three solvents extracts from the alkali lignin degraded by the bacterial strains and the control sample.
| RT (min) | Compounds | CK | YB5 | LN4 | LN2 | CL32 |
|---|---|---|---|---|---|---|
| Hexane as solvent | ||||||
| 19.24 | Pentonic acid lactone* | – | + | + | + | – |
| 22.65 | Hexadecanoate* | – | + | + | + | + |
| 24.46 | Octadecanoate* | – | + | + | + | + |
| 28.20 | Propyl hexadecanoate* | – | + | + | + | + |
| Dioxane as solvent | ||||||
| 8.45 | 2-Ethoxyethanol | + | + | + | + | + |
| 11.67 | 2,4-Hexadienal | + | + | + | + | + |
| 12.83 | Di (ethylene glycol) vinyl ether | + | + | + | + | + |
| 21.92 | Dimethylbiphenyl | + | – | – | – | – |
| 24.07 | 1,2,3-trimethyl-4-prop-1-enylnaphthalene | + | – | – | – | – |
| 26.87 | Diisobutyl phthalate | + | – | – | – | – |
| 29.32 | Dibutyl phthalate | + | – | – | – | – |
| Ethyl acetate as solvent | ||||||
| 21.54 | Acetosyringone | + | + | + | + | + |
| 22.93 | Diisooctyl phthalate | + | – | – | – | – |
| 23.6 | Palmitoleic acid | + | – | – | – | – |
| 23.79 | Dibutyl phthalate | + | – | – | – | – |
| 25.51 | Cis-Vaccenic acid | + | – | – | – | – |
Clearly, 2-ethoxyethanol, 2,4-hexadienal, di(ethylene glycol) vinyl ether, and acetosyringone were not degraded by the strains and were detected in both the CK group (only medium, no strain, as a control group), and in the treatment group. Hexadecanoate, octadecanoate, and propyl hexadecanoate were not detected in the CK group but were detected in the YB5, LN4, LN2 and CL32 strains. Pentonic acid lactone was detected in the YB5, LN4 and LN2 strains, but not in the control group and in CL32 strain.
Dimethylbiphenyl, 1,2,3-trimethyl-4-prop-1-enyl-naphthalene, diisobutyl phthalate, dibutyl phthalate, palmitoleic acid, cis-vaccenic acid was detected in the CK group. It is worth noting that in YB5, LN4, LN2 and CL32 strains the compounds that appeared in CK group were not detected. The chromatographs of compounds extracted with ethyl acetate, hexane, and 1,4-dioxane are showed in Supplementary Fig. 1–3.
The objective of this study was to screen the functional bacteria, which could degrade lignin in soil. There are many plant residues on the black soil in the forest, rich in humus and they potentially contain many lignin-degrading microorganisms. The screened bacteria were Bacillus, Acinetobacter, and Actinomycetes, whose ability to decolorize aniline blue and degrade lignin was reported. So, they could synthesize lignin peroxidase and manganese peroxidase, although none of them can synthesize laccase. So, they could synthesize lignin peroxidase and manganese peroxidase, although none of them can synthesize laccase.
It had been reported that many
Lignin peroxidase is an important lignin-degrading enzyme. Interestingly, although the enzyme from the YB5 strain has a higher degradation rate, the Lip enzyme activity was lowest on the first day. Bharagava and coworkers studied
In contrast, lignin-degrading bacteria could also be found in black liquor.
Sonoki and coworkers discovered that
Why are the degradation characteristics of these four strains so similar? The degradation ability of microbial communities after incubation in alkali lignin, which tends to be consistent, is also called a functional convergence. Carlos and coworkers mentioned it in their research, due to an enrichment of genes involved in benzoate degradation and catechol ortho cleavage pathways (Carlos et al. 2018). Go and coworkers isolated
In this study, four strains of lignin-degrading bacteria from soil, straw compost, and silage were screened.
Fig. 1.
Fig. 2.
Fig. 3.
The compounds identified in three solvents extracts from the alkali lignin degraded by the bacterial strains and the control sample.
| RT (min) | Compounds | CK | YB5 | LN4 | LN2 | CL32 |
|---|---|---|---|---|---|---|
| Hexane as solvent | ||||||
| 19.24 | Pentonic acid lactone* | – | + | + | + | – |
| 22.65 | Hexadecanoate* | – | + | + | + | + |
| 24.46 | Octadecanoate* | – | + | + | + | + |
| 28.20 | Propyl hexadecanoate* | – | + | + | + | + |
| Dioxane as solvent | ||||||
| 8.45 | 2-Ethoxyethanol | + | + | + | + | + |
| 11.67 | 2,4-Hexadienal | + | + | + | + | + |
| 12.83 | Di (ethylene glycol) vinyl ether | + | + | + | + | + |
| 21.92 | Dimethylbiphenyl | + | – | – | – | – |
| 24.07 | 1,2,3-trimethyl-4-prop-1-enylnaphthalene | + | – | – | – | – |
| 26.87 | Diisobutyl phthalate | + | – | – | – | – |
| 29.32 | Dibutyl phthalate | + | – | – | – | – |
| Ethyl acetate as solvent | ||||||
| 21.54 | Acetosyringone | + | + | + | + | + |
| 22.93 | Diisooctyl phthalate | + | – | – | – | – |
| 23.6 | Palmitoleic acid | + | – | – | – | – |
| 23.79 | Dibutyl phthalate | + | – | – | – | – |
| 25.51 | Cis-Vaccenic acid | + | – | – | – | – |
Screening and identification of bacterial isolates based on different media.
| Name | Species | Accession number | Alkaline lignin | Aniline blue | Brilliant blue |
|---|---|---|---|---|---|
| YB5 | MT745877 | ∘ | + | – | |
| CL32 | MT745880 | ∘ | + | – | |
| LN2 | MT745878 | ∘ | ++ | – | |
| LN4 | MT745879 | ∘ | ++ | – |