Isolation and characterization marine bacteria capable of degrading lignin-derived compounds

Peng Lu; Weinan Wang; Guangxi Zhang; Wen Li; Anjie Jiang; Mengjiao Cao; Xiaoyan Zhang; Ke Xing; Xue Peng; Bo Yuan; Zhaozhong Feng

doi:10.1371/journal.pone.0240187

Abstract

Lignin, a characteristic component of terrestrial plants. Rivers transport large amounts of vascular plant organic matter into the oceans where lignin can degrade over time; however, microorganisms involved in this degradation have not been identified. In this study, several bacterial strains were isolated from marine samples using the lignin-derived compound vanillic acid (4-hydroxy-3-methoxybenzoic acid) as the sole carbon and energy source. The optimum growth temperature for all isolates ranged from 30 to 35°C. All isolates grew well in a wide NaCl concentration range of 0 to over 50 g/L, with an optimum concentration of 22.8 g/L, which is the same as natural seawater. Phylogenetic analysis indicates that these strains are the members of Halomonas, Arthrobacter, Pseudoalteromonas, Marinomonas, and Thalassospira. These isolates are also able to use other lignin-derived compounds, such as 4-hydroxybenzoic acid, ferulic acid, syringic acid, and benzoic acid. Vanillic acid was detected in all culture media when isolates were grown on ferulic acid as the sole carbon source; however, no 4-hydroxy-3-methoxystyrene was detected, indicating that ferulic acid metabolism by these strains occurs via the elimination of two side chain carbons. Furthermore, the isolates exhibit 3,4-dioxygenase or 4,5-dioxygenase activity for protocatechuic acid ring-cleavage, which is consistent with the genetic sequences of related genera. This study was conducted to isolate and characterize marine bacteria of degrading lignin-derived compounds, thereby revealing the degradation of aromatic compounds in the marine environment and opening up new avenues for the development and utilization of marine biological resources.

Citation: Lu P, Wang W, Zhang G, Li W, Jiang A, Cao M, et al. (2020) Isolation and characterization marine bacteria capable of degrading lignin-derived compounds. PLoS ONE 15(10): e0240187. https://doi.org/10.1371/journal.pone.0240187

Editor: Pankaj Kumar Arora, Babasaheb Bhimrao Ambedkar University, INDIA

Received: December 25, 2019; Accepted: September 22, 2020; Published: October 7, 2020

Copyright: © 2020 Lu et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: All relevant data are within the paper and its Supporting Information files.

Funding: This study was supported by National Natural Science Foundation of China (31800327, 31570028), the project funded by Grant from Anhui National Natural Science Foundation (1708085QC67), the Priority Academic Program Development of Jiangsu Higher Education Institutions (PAPD), Postgraduate Research and Practice Innovation Program of Jiangsu Province, China (KYCX18_2138, KYCX17_1618). The authors declare that they have no conflicts of interest with the contents of this article.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Lignin, a major component of vascular plant tissue, is the second most abundant natural substance in nature after cellulose [1]. It plays a protective role against pathogenic and saprophytic organisms by guarding cellulose polymers from hydrolytic attack. Furthermore, it also allows for terrestrial plant growth by providing rigidity in the stems and supports the circulation of sap by waterproofing the vascular tissue [2].

The complex aromatic lignin heteropolymer is formed when crosslinked lignin monomers, such as guaiacyl, syringyl, and p-hydroxyphenyl, are further polymerized via β-aryl ether linkages, biphenyl bonds and heterocyclic linkages, by oxidase radicals. Although this molecule is highly recalcitrant, several enzymes with the ability to depolymerize lignin have been identified, such as lignin peroxidases (LiPs), manganese peroxidases (MnPs) and laccases [1]. These extracellular fungal enzymes attack lignin substructures in decaying plant cell walls, thereby releasing different lignin-derived breakdown products [2].

Vanillic acid, ferulic acid, p-hydroxybenzoic acid, syringic acid, benzoic acid and biphenyl are known lignin-derived compounds [3]. Detailed studies have been performed on the degradation of these low molecular compounds by soil bacteria [4, 5]. Research suggests that protocatechuic acid (PCA) is a key intermediate in the metabolism of lignin-derived compounds, resulting in aromatic ring cleavage. Three PCA metabolic pathways have been reported: 2,3-cleavage [6], 3,4-cleavage and 4,5-cleavage [7, 8]. The ring cleavage products from each pathway eventually enter the tricarboxylic acid cycle.

Rivers transport large amounts of vascular plant organic matter into the oceans, resulting in a carbon flux that is estimated to make up approximately 0.7% of terrestrial primary production [9]. The fact that lignin is only produced by terrestrial plants, and because it is a highly stable compound, makes it an excellent biomarker for the presence of terrestrial organic matter in lacustrine and marine environments [10]. Furthermore, when buried in sediments, this biomarker can aid in assessing the impact of land composition on marine environments [11, 12]. Marine microbes from the sea that are capable of degrading lignin potentially contribute to organic matter cycling [13]. Additional information can be inferred by determining the ratios of lignin-derived compounds. For instance, the synringyl/ vanillyl ratio, and the cinnamyl/ vanillyl phenols ratio can indicate the vegetative source; whereas the vanillic acid/ vanillin ratio and the syringic acid/ syringaldehyde ratio indicates the degradation state of lignin materials [14]. Research suggests that the average cycling time of dissolved lignin in marine environments is approximately 90 years [15], thereby highlighting the importance of understanding the degree and mechanisms of lignin degradation in the oceans, which, in turn allows, for a better understanding of carbon cycling within the marine environment.Some studies have examined lignin degradation by mixed microorganisms in marine environments [16, 17]; however, the microorganisms responsible for the degradation were not identified.

In this study, we isolated several bacterial strains from marine samples using the lignin-derived compound vanillic acid as the sole carbon and energy source. We then investigated the physiological characteristics of the isolates to determine their role in lignin mineralization in the marine environment.

Materials and methods

All experiments in the materials and methods were conducted according to protocols approved by Jiangsu Normal University and Anhui Normal University and completed by researchers of the manuscript. Field sampling is funded by the National Natural Science Foundation of China, and belongs to the research content of the project. The study area is not a special marine protected area and does not involve endangered or protected species.

Study site and sample collection

Seawater samples were collected from Lianyungang and sediments samples were collected from the East China Sea in Zhoushan, Zhejiang (30°58′20.71″N, 122° 95 41′22.10″E), People's Republic of China. The depth of the sediment sample was 10 m below sea level, the water temperature was 20°C. Further, the sediments samples were placed in sealed plastic boxes surrounded with ice and brought to the laboratory within 24h.

Chemicals and material

4-hydroxybenzoic acid (99.9%purity), benzoic acid (99.9% purity) and syringic acid (99.8% purity) were purchased from Sangon Biotech (Shanghai, China) at the highest purity available. Methanol (99.9%purity), trifluoroacetic acid (99.5%purity), and acetonitrile (99.9%purity) were purchased from Sigma-Aldrich (St. Louis, MO). Vanillic acid (99.9%purity), ferulic acid (99.9%purity) and protocatechuic acid (99.9%purity) were purchased from TCI Fine Chemicals (Tokyo, Japan). Marine Broth 2216 (DSMZ medium 604) was used as the nutrient medium. ONR7a medium (DSMZ medium #950, DSMZ, Germany), which contains only inorganic ions, was used as the defined seawater medium [18].

Isolation and characterization of vanillic acid degraders

Both the direct plating method and the enrichment technique were used to isolate vanillic acid-utilizing bacteria. The autoclaved medium was allowed to cool at room temperature and the filter sterilized vanillic acid was added into the medium. Briefly, direct plating was carried out on ONR7a agar plates amended with 0.2% vanillic acid. Cultures were incubated at 25°C until colonies appeared. The individual colonies that utilized vanillic acid as the sole carbon source for growth were selected and purified by re-streaking 3 times. The enrichment technique was performed in liquid ONR7a media amended with 0.2% vanillic acid. Cultures were incubated with shaking at 25°C until turbid. They were then plated on ONR7a agar after repeating this enrichment technique 5 times to obtain pure colonies [19].

To isolate the total DNA, each isolate was grown to exponential phase in 10 ml Marine Broth at 25°C. The resulting cells were washed twice with TES buffer (pH 8.0) containing 0.1 M NaCl, 10 mM Tris-HCl, and 1 mM EDTA and then suspended in 2.5 ml of the same buffer amended with 5 mg of lysozyme. After being kept at 37°C for 1h, 256 μl of 10% (wt/vol) sodium dodecyl sulfate (SDS) and 0.5 ml of TES containing 0.5 mg of protease K were added, and the mixture was incubated for 1 h at 65°C. Then, 1.8 ml of 10% SDS was added, and the mixture was incubated for another 1 h at 65°C. Total DNA was precipitated by adding 2.5 volumes of ethanol at room temperature. The DNA obtained by centrifugation at 6000 g for 10 min at room temperature was washed with 70% ethanol and dissolved in 1 ml TES for use.

The 16S rRNA gene-containing fragment of approximately 1.5 kb was amplified with the 27F (AGAGTTTGATCMTGGCTCAG) and 1492R (TACGGYTACCTTGTTACGACTT) primers using 2 μl DNA as the template, and the PCR products were sequenced [20]. PCR was done with a total volume of 50 μl, which contained 27.8 μl Sterilized H₂O, 5 μl 10×PCR Buffer, 5 μl dNTP Mix, 4μl MgCl₂, 3 μl Primer1, 3 μl Primer2, 0.2 μl Taq DNA Polymerase and 2 μl Template DNA. Amplification of the DNA was done for 35 cycles under the following conditions: denaturation, 95°C for 30 s; primer annealing, 56°C for 30 s; and primer ex tension, 72°C for 90 s. Miseq platform was used and the concentration of each sample was about 300ng μl^-1. The almost-complete 16S rDNA gene sequences of these isolates were obtained and used for initial BLAST searches in GenBank and for phylogenetic analyses. The sequences were aligned to their nearest phylogenetic neighbors using Genetyx software (GENETYX Corporation, Tokyo, Japan), and calculation of pairwise 16S rDNA gene sequence similarities was achieved using the EMBL sever (http://www.ebi.ac.uk/Tools/psa/). Phylogenetic trees were generated using same software. The robustness of the inferred topologies was tested by using 1,000 bootstrap re-samplings.

Optimum growth conditions for temperature and NaCl concentration

Each bacterium was grown to the stationary phase in Marine Broth 2216 at 25°C. The resulting cells were collected by centrifugation (5000×g, 5 min) and resuspended in 100 ml of the ONR7a medium to an optical density at 600 nm (OD₆₀₀) of 0.1. The cultures were incubated at different temperatures (20, 30, 35, 40, and 45°C) and NaCl concentration (0 g L^-1, 15 g L^-1, 22.8 g L^-1, 40 g L^-1, and 50 g L^-1). A portion of the culture was collected and measured at OD₆₀₀ after being incubated with shaking. Each experiment was done in parallel three times.

Degradation of different lignin-derived compounds by isolates

Each bacterium was grown to the stationary phase in Marine Broth 2216 at 25°C. The resulting cells were washed twice with ONR7a medium and then suspended in 5 ml of the same medium to give a turbidity of 0.05–0.1 at 600 nm. Each of the lignin-derived compounds (4-hydroxybenzoic acid, benzoic acid, ferulic acid and syringic acid) was added to a final concentration of 0.1%, and the mixture was shaken (150 rpm) at 30°C. Each experiment was done in one time.

Degradation products of ferulic acid by isolates

Isolates were inoculated into ONR7a medium supplemented with ferulic acid and incubated for 24 h under conditions optimized. The concentrations of degradation products were determined with HPLC.

Analysis of metabolism of lignin-derived compounds by HPLC and GC-MS

0.5 ml of the culture was collected and mixed with equal amounts of methanol. The mixture was filtered through a Millipore Filter Membrane (0.22 μm/50 mm) and then subjected to high-performance liquid chromatography (HPLC) analysis. An Alliance HPLC instrument (Waters, Milford, Mass.) with an octadecyl silica reverse-phase column (30 cm in length and 4.6 mm in diameter; Waters) was used at 40°C. A mobile phase consisting of solvent A (0.1% trifluoroacetic acid in water) and solvent B (acetonitrile) was used at a flow rate of 1 ml/min. After injection of 10 μl of a sample, 90% solvent A was run through the column for 4 min, after which the percentage of solvent B was gradually increased from 10% to 100% over 15 minutes. Finally, 100% of solvent B was run for 2 min. The absorbance spectrum of the eluate was monitored from 200 to 500 nm with a photodiode array detector (Waters 2998). The retention times of vanillic acid and ferulic acid were 6.237 and 7.332 min, respectively. All vanillic acid and ferulic acid we have used are standard. The test samples were extracted by ethyl acetate and dried over anhydrous Na₂SO₄ and the solvent was allowed to evaporate at 40°C. The residual was dissolved in an equal volume of methanol to original liquid sample and analyzed by gas chromatography-mass spectrometry (GC-MS) (Shimadzu QP5050A instrument) with a DB-5 column (length, 30 m; diameter, 0.25 mm) (J&W Scientific). The column temperature was initially kept at 50°C for 5 min before being increased to 300°C at a rate of 2.5°C per min. The injector and detector temperatures were both 300°C. Determination of vanillic acid, vanillin, vanillate, protocatechuate, and fumarate through SIM (selective reaction monitoring) with referenced ion fragments information as showed as following: 168.15m/z, 152.15m/z, 151.03m/z, 154.12m/z and 115.05m/z, respectively.

Enzyme assays for PCA dioxygenase

Isolates were grown in 600 ml of Marine Broth for 12 h at 25°C with shaking and then washed twice with ONR7a. We centrifuged the pre-cultured bacterial solution (5000× g, 5 min), removed the supernatant, then washed twice with seawater medium, removed the supernatant, and blotted dry with a pipette tip. Finally, the bacterial solution was dissolved in a seawater medium of 5 ml. The resulting cells were resuspended in 200 ml of ONR7a containing 0.2% vanillic acid, and the mixture was incubated with shaking at 25°C. The cells were harvested when approximately half of the vanillic acid was degraded, washed twice with 50 mM Tris-HCl (pH 8.0), and disrupted by ultrasonication. The amount of vanillic acid was determined by spectrophotometer. The cell-free extracts were obtained by centrifugation. The reaction mixture for assaying PCA dioxygenase activity consisted of 1900 μl of 50 mM Tris-HCl (pH 8.0), 40 μl of 1 mM PCA, and 60 μl of cell-free extract (100 μg ml^-1). The reaction mixture was incubated at 25°C and monitored spectrometrically from 200 to 500 nm with an EVOLUTION 260 BIO UV-Visible spectrophotometer (ThermoFisher, USA).

Results

Isolation of vanillic acid degraders

In this study, vanillic acid was used as lignin-derived low molecular weight compound to isolate vanillic acid-utilizing bacteria from marine and sediment samples collected from the coast of Lianyungang and the China East Sea, respectively. The predominant strains, which have adapted to their marine environment, could be isolated using the direct plating method, which resulted in isolation of bacteria with the highest growth rate in laboratory conditions. Three strains with the ability to use vanillic acid as the sole carbon and energy source were successfully isolated by direct plating, namely, strains ZXY12VA01, ZXY12VA14, and ZXY12VA15 (Table 1). Three additional strains with the ability to use vanillic acid as the sole carbon and energy source were isolated using the enrichment technique; namely, ZXY12VA05, ZXY12VA16, and ZXY12VA17.

Download:

Table 1. Strains isolated in this study.

https://doi.org/10.1371/journal.pone.0240187.t001

The colony morphology of the isolates when plated with vanillic acid are as follows: ZXY12VA01, white and small; ZXY12VA05, milk yellow, uplifted with a smooth edge, large; ZXY12VA14, milk yellow, smooth edge, uplifted; ZXY12VA15, white and small, uplifted, rapid growth; ZXY12VA16, small, dark brown, slow growth, secreted brown pigment; ZXY12VA17, primrose yellow and turned to yellow. All strains are Gram-negative, except strain ZXY12VA05, which is Gram-positive (Fig 1).

Download:

Fig 1.

Gram stain of isolates ZXY12VA01 (A), ZXY12VA05 (B), ZXY12VA14 (C), ZXY12VA15 (D), ZXY12VA16 (E), ZXY12VA17 (F).

https://doi.org/10.1371/journal.pone.0240187.g001

Electrophoresis results of PCR products (1500 bp) of isolated bacteria are shown in S1 Fig. Phylogenetic analysis based on the neighbor-joining algorithm revealed that strain ZXY12VA01 clusters with the Halomonas genus (Fig 2). The most closely related type strain is Halomonas titanicae BH1^T (99.87% 16S rDNA sequence similarity). Other related type strains are Halomonas variabilis DSM 3051^T (99.0%), Halomonas olivaria C17^T (99.0%), and Halomonas neptunia Eplume1^T (99.0%). Based on these data, strain ZXY12VA01 likely belongs to Halomonas, which belongs to the family Halomonadaceae. This heterogeneous family, originally proposed by Frenzmann et al. (1988), to date includes more than 60 species [21]. They have been isolated from many water and soil environments within saline, hypersaline or alkaline habitats. Halomonas titanicae BH1^T was isolated from rusticle samples collected at the site of the wreck using Bacto marine agar 2216 medium (Difco). It grows optimally at 30–37°C in the presence of 2–8% (w/v) NaCl.

Download:

Fig 2. Phylogenetic tree of thermophilic isolates based on 16S rDNA sequences.

https://doi.org/10.1371/journal.pone.0240187.g002

The most closely related type strain to ZXY12VA05 is Arthrobacter nicotinanae DSM 20123T (99.86%). Other related type strains include Arthrobacter mysorens LMG 16219^T, Arthrobacter arilaitensis Re117^T, and Arthrobacter uratoxydans DSM 20647^T. Based on these data, strain ZXY12VA05 belongs to the genus Arthrobacter. Members of the Arthrobacter genus have a high G+C content, are actinomycete-coryneform bacteria, and are all Gram-positive. This genus contains more than 40 species of bacteria isolated from diverse environments, including soil and clinical specimens. Most Arthrobacter strains are mesophiles; however, some members have been isolated from Antarctic lakes and deep-sea sediment [22].

Both ZXY12VA14 and ZXY12VA17 strains cluster with the genus Pseudoalteromonas (Fig 2). The most closely related type strain for ZXY12VA14 and ZXY12VA17 are Pseudoalteromonas mariniglutinosa KMM 3635^T (95%) and Pseudoalteromonas shioyasakiensis SE3^T (98%), respectively. Other related type strains are Pseudoalteromonas prydzensis MB8-11^T for ZXY12VA14; and Pseudoalteromonas arabiensis k53^T, Pseudoalteromonas lipolytica LMEB 39^T, and Pseudoalteromonas byunsanensis FR1199^T for ZXY12VA17. Members of Pseudoalteromonas are characteristically Gram-negative, rod-shaped, motile bacteria with one or two polar flagella. Most Pseudoalteromonas strains are mesophiles; however, some halotolerant psychrophilic members have been isolated from Antarctic sea ice [23]. Strain KMM 3635^T was isolated from a Chaetoceros lauderi diatom, which was collected from seawater in the Marseille Gulf [24]. Strain SE3T was isolated from Pacific Ocean sediment [25].

Phylogenetic analysis revealed that the most closely related type strain to ZXY12VA15 (Fig 2) is Marinomonas communis LMG 2864^T (99.15% 16S rDNA sequence similarity). Other related type strains include Marinomonas vaga ATCC 27119^T, Marinomonas aquimarina CECT 5080^T, and Marinomonas fungiae AN44^T. Based on these data, strain ZXY12VA15 belongs to the genus Marinomonas, which was created in 1983 and comprises 15 species. The members are Gram-negative, aerobic bacterium and have mainly been isolated from geographically diverse marine environments [26].

Strain ZXY12VA16 clusters with the Thalassospira genus (Fig 2). The most closely related type strain is Thalassospira povalilytica Zumi 95^T (99.62% 16S rDNA sequence similarity). Other related type strains include Thalassospira profundimaris WP0211^T, Thalassospira tepidiphila 1-1BT, and Thalassospira permensis NBRC 106175^T. Members of the Thalassospira genus are vibrioid to spiral cells; they are strict aerobes that are unable to grow under anaerobic conditions by fermentation, respiration or photoheterotrophy [27]. Thalassospira povalilytica Zumi 95^T was isolated from plastic rope litter found in the Tokyo Bay, Japan, and exhibits the ability to utilize polyvinyl-alcohol as a carbon source [28].

The sequence data reported in this study appear in the EMBL, GenBank and DDBI sequence databases under the accession numbers. The GenBank accession number of strains ZXY12VA01, ZXY12VA05, ZXY12VA14, ZXY12VA15, ZXY12VA16 and ZXY12VA17 are KT274805, KT274806, KT717632, KT274807, KT274808 and KT274809, respectively.

Optimum growth conditions for temperature and NaCl concentration

Marine and terrestrial environments are highly variable; however, microorganisms have the ability to rapidly evolve and adapt to changing environments. The optimum growth conditions of terrestrial bacteria in terms of temperature and NaCl concentration are generally 30°C and 0.5%, respectively. In contrast, marine microorganisms prefer to grow at temperatures under 30°C and at a high NaCl concentration. Thus, by investigating the optimal temperature and salt concentrations of the isolates, it may be possible to infer if these organisms originated from the terrestrial or the marine environment.

The optimum growth temperature of the isolates rang from 30 to 40°C (Fig 3). The optimum growth temperature for strains ZXY12VA01, ZXY12VA15, ZXY12VA16, and ZXY12VA17 is 35°C; whereas strains ZXY12VA05 and ZXY12VA14 grow optimally at 30°C. These isolates do not grow above 45°C or below 20°C.

Download:

Fig 3.

Optimum growth temperatures of isolates ZXY12VA01 (A), ZXY12VA05 (B), ZXY12VA14 (C), ZXY12VA15 (D), ZXY12VA16 (E), ZXY12VA17 (F).

https://doi.org/10.1371/journal.pone.0240187.g003

All isolates grow well in a wide NaCl concentration range from 0 to 50 g L^-1, with the optimum concentration being 22.8 g L^-1, which is the same as natural seawater (Fig 4). These 6 strains appear to have adapted to the NaCl concentration of seawater; therefore, they are not terrestrial microorganisms that were recently deposited into the marine environment.

Download:

Fig 4.

Optimum growth in different NaCl concentrations of isolates ZXY12VA01 (A), ZXY12VA05 (B), ZXY12VA14 (C), ZXY12VA15 (D), ZXY12VA16 (E), and ZXY12VA17 (F).

https://doi.org/10.1371/journal.pone.0240187.g004

Ability to utilize lignin-derived compounds

In addition to vanillic acid, it is possible for some organisms to utilize other lignin-derived low molecular weight compounds, such as benzoic acid, 4-hydroxybenzoic acid, ferulic acid, and syringic acid. Thus, the isolates were cultured for 10 days at 30°C in the selected lignin-derived compounds to test their ability to use these compounds as their sole carbon and energy source. 4-Hydroxybenzoic acid was the best substrate for all isolates because it resulted in cell densities of approximately 1.0 of OD₆₀₀ (Table 2). The cell densities grown on syringic acid and ferulic acid were much lower than those on 4-hydroxybenzoic acid; however, strains ZXY12VA05, ZXY12VA14, ZXY12VA16 grew as well on benzoic acid as on 4-hydroxybenzoic acid, while ZXY12VA01 and ZXY12VA15 did not. These results demonstrate that the strains isolated in this study have the ability to grow on several lignin-derived compounds and likely play an important role in lignin mineralization in marine environments.

Download:

Table 2. Vanillate-degrading isolates grown on different lignin-derived carbon sources.

https://doi.org/10.1371/journal.pone.0240187.t002

Detection of ferulic acid intermediates

Ferulic acid, an easily available component of lignin, plays an important role in the cross-linking of cell walls in various plants. The first step in ferulic acid degradation is the removal of its side chain. Two types of side chain removal have been reported for a number of microorganisms [29]. One type, which results in a 4-hydroxy-3-methoxystyrene intermediate, is catalyzed by a non-oxidative decarboxylase that eliminates one carbon from the ferulic acid side chain. The 4-hydroxy-3-methoxystyrene is subsequently transformed to vanillin and then to vanillic acid. The other type, which results in the direct formation of vanillin, involves the elimination of two carbons from the ferulic acid side chain. Vanillin is subsequently oxidized to vanillic acid by vanillin dehydrogenase.

To determine which type of metabolism is carried out by these strains, metabolites from cultures grown with ferulic acid as the sole carbon source were investigated using HPLC. Vanillic acid was detected in all culture media when isolates were grown on ferulic acid as the sole carbon source; however, no 4-hydroxy-3-methoxystyrene was detected (Figs 5 and 6), indicating that ferulic acid metabolism by these strains occurs via the elimination of two side chain carbons. Through this study, we found that ZXY12VA16 produced the most amount of vanillic acid. The ZXY12VA15 strain is a dominant strain of aromatic degrading bacteria. Normally, feruloyl-CoA synthetase and enoyl-CoA hydratase/aldolase are found in most FA-degrading strains. These strains convert ferulic acid to vanillin by a non-β-oxidative pathway [30]. It demonstrates that vanillic acid is an important intermediate in the metabolism of lignin and plays a leading role in the metabolism of aromatic compounds.

Download:

Fig 5. The formation of the intermediates in the biodegradation of ferulic acid in ZXY12VA01 shown in the HPLC analysis.

The major metabolites were detected on the 0h (A), 12h (B) and 24h (C), respectively.

https://doi.org/10.1371/journal.pone.0240187.g005

Download:

Fig 6.

Metabolism of ferulic acid to vanillic acid by isolates ZXY12VA01 (A), ZXY12VA05 (B), ZXY12VA14 (C), ZXY12VA15 (D), ZXY12VA16 (E), ZXY12VA17 (F).

https://doi.org/10.1371/journal.pone.0240187.g006

When strain ZXY12VA01, ZXY12VA05 and ZXY12VA14 was grown on 0.1% vanillic acid as the sole carbon, the formation of vanillin and protocatechuic acid was detected in the culture medium (Fig 7). Vanillic acid can be hydroformylated to form vanillin or demethylated to form protocatechuic acid. Protocatechuic acid is considered to be the target compound for ring fission. Protocatechuate has a vic-diol structure, which can be subjected to ring cleavage by dioxygenase. The formation of fumaric acid proves that the metabolite is finally decomposed into the tricarboxylic acid cycle (Fig 8).

Download:

Fig 7.

Growth of ZXY12VA01 (A), ZXY12VA05 (B) and ZXY12VA14 (C) on vanillic acid. The four major metabolites were detected on the 12h and 24h, respectively.

https://doi.org/10.1371/journal.pone.0240187.g007

Download:

Fig 8. Speculated biodegradation pathways of ferulic acid and vanillic acid by marine bacteria.

https://doi.org/10.1371/journal.pone.0240187.g008

Enzyme assay for aromatic ring-fission reaction

Vanillic acid metabolism involves removal of the methoxy group by a demethylase, resulting in the formation of a protocatechuic acid (PCA) intermediate. Ring cleavage of PCA occurs by three known dioxygenases: PCA 2,3-dioxygenase, PCA 3,4-dioxygenase, and PCA 4,5-dioxygenase and results in the ring-cleavage products 5-carboxy-2-hydroxymuconate-6-semialdehyde (350 nm), 5-carboxymuconate (290 nm), and 4-carboxy-2-hydroxymuconate-6-semialdehyde (410 nm), respectively. Each ring-fission product has a specific absorption peak, making it easy to determine which ring-cleavage occurred.

To induce vanillic acid-metabolizing enzymes, ZXY12VA15 was grown on vanillic acid and harvested when approximately half of the carbon source was degraded. The cell-free extracts were applied to the enzyme assay using PCA as the substrate. Spectrophotometric assays provided evidence for the formation of 4-carboxy-2-hydroxymuconate-6-semialdehyde (λ_max, 410 nm) at pH 8.0 (Fig 9). It was determined that strain ZXY12VA15 were capable of 4,5-cleavage.

Download:

Fig 9. Spectrophotometric analysis of ZXY12VA15 enzyme activity.

Absorbance increase of 4-carboxy-2-hydroxymuconate-6-semialdehyde (λ_max, 410 nm).

https://doi.org/10.1371/journal.pone.0240187.g009

Discussion

Genome-wide surveys were performed on the isolate representative species of related genera to identify genes associated with the PCA ring-cleavage pathway and the genomic information of strains were searched on KEGG.

Halomonas elongate DSM 2581 is a halotolerant bacteria isolated from a solar salt facility. It synthesizes the compatible solute Ectoine, thus allowing it to withstand NaCl concentrations well above 10% [31]. Contained in the H. elongata genome is a gene cluster (HELO_3955–3963) encoding PCA 3,4-dioxygenase and additional enzymes involved in the metabolism of ring-cleavage products.

Marinomonas mediterranea MMB-1 [32], Marinomonas sp. MWYL1 [33, 34], and Marinomonas posidonica IVIA-Po-181 [35] were isolated from marine samples. Their genomes have the same gene cluster, which encodes for PCA 4,5-dioxygenase and enzymes involved in the ring-cleavage product metabolism.

The members of Arthrobacter are among the most frequently isolated, indigenous, aerobic genera found in soils. Arthrobacter phenanthrenivorans Sph3 was isolated from a creosote-polluted site, where a wood-preserving factory had operated for over 30 years. It is able to metabolize phenanthrene as the sole carbon and energy source [36]. Arthrobacter aurescens TC1 was originally isolated from soil at an atrazine spill site [37]. Arthrobacter chlorophenolicus A6, a Gram-positive Actinobacterium, was enriched from a soil slurry using increasing concentrations of 4-chlorophenol and can degrade exceptionally high concentrations of 4-chlorophenol (up to 2.7 mM), along with other p-substituted phenols, including 4-nitrophenol and 4-bromophenol [38]. Arthrobacter arilaitensis Re117 was isolated from the surface of cheeses and possesses a salt-tolerance system [39]. Arthrobacter sp. FB24 was isolated from a soil sample that was obtained at a site contained by mixed waste that included both petroleum hydrocarbons and extremely high metal levels. The genomes of these Arthrobacter members have the same gene cluster encoding PCA 3,4-dioxygenase and enzymes involved in the ring-cleavage product metabolism.

Thalassospira xiamenensis M-5 (DSM17429) was isolated from an oil-waste pool at an oil storage dock. Strain M-5 has a gene cluster encoding PCA 3,4-dioxygenase and enzymes involved in the ring-cleavage product metabolism [40].

Pseudoalteromonas atlantica T6c is a biofilm-forming, Gram-negative, motile marine bacterium. It is chemoorganotrophic and secrets extracellular enzymes that are able to hydrolyze agar, alginate, and carrageenan [41]. Strain T6c has a gene cluster encoding PCA 4,5-dioxygenase and enzymes involved in the ring-cleavage product metabolism.

This study demonstrates that members of the genera Halomonas, Arthrobacter, Pseudoalteromonas, Marinomonas, and Thalassospira are involved in the degradation of lignin-derived compounds in marine environments. In addition, many environmental pollutants are aromatic compounds, and marine bacteria capable of degrading lignin can also degrade most environmental pollutants. The isolated lignin-degrading bacteria can be applied to environmental pollution treatment in a marine environment or a high-salt environment. Although we have isolated marine bacteria that can degrade vanillic acid, our further research will focus on applying to actual production.

Conclusion

In this study, 6 bacterial strains were isolated from marine samples using the lignin-derived compound vanillic acid as the sole carbon and energy source. It is concluded that these 6 strains have adapted to the NaCl concentration of seawater and are marine dominant bacteria widely present in the marine environment, rather than the soil microorganisms of the recent land. They are Phylogenetic analysis indicates that ZXY12VA01, ZXY12VA05, ZXY12VA14, ZXY12VA15, ZXY12VA16, and ZXY12VA17 are members of Halomonas, Arthrobacter, Pseudoalteromonas, Marinomonas, and Thalassospira, respectively. They are also able to use other lignin-derived compounds, such as 4-hydroxybenzoic acid, ferulic acid, syringic acid, and benzoic acid. The isolates showed 3,4-dioxygenase or 4,5-dioxygenase activity for protocatechuic acid ring-cleavage, which is consistent with information on related genera genome sequences.

Supporting information

S1 Fig. Electrophoresis image of 16S rDNA PCR products.

https://doi.org/10.1371/journal.pone.0240187.s001

(DOCX)

References

1. Pollegioni L, Tonin F, Rosini E. Lignin-degrading enzymes. Febs Journal, 2015, 282(7):1190–1213. pmid:25649492
- View Article
- PubMed/NCBI
- Google Scholar
2. Ruiz-Dueñas FJ, Martínez AT. Microbial degradation of lignin: how a bulky recalcitrant polymer is efficiently recycled in nature and how we can take advantage of this. Microb Biotechnol, 2009, 2:164–177. pmid:21261911
- View Article
- PubMed/NCBI
- Google Scholar
3. Kuzyk ZZA, Goñi MA, Stern GA, Macdonald RW. Sources, pathways and sinks of particulate organic matter in Hudson Bay: Evidence from lignin distributions. Marine Chemistry, 2008, 11:215–229.
- View Article
- Google Scholar
4. Masai E, Katayama Y, Fukuda M. Genetic and biochemical investigations on bacterial catabolic pathways for lignin-derived aromatic compounds. Biosci Biotechnol Biochem, 2007, 71:1–15. pmid:17213657
- View Article
- PubMed/NCBI
- Google Scholar
5. Peng X, Masai E, Kasai D, Miyauchi K, Katayama Y, Fukuda M. A second 5-carboxyvanillate decarboxylase gene, ligW2, is important for lignin-related biphenyl catabolism in Sphingomonas paucimobilis SYK-6. Appl Environ Microbiol, 2005, 71:5014–5021. pmid:16151081
- View Article
- PubMed/NCBI
- Google Scholar
6. Kasai D, Fujinami T, Abe T, Mase K, Katayama Y, Fukuda M, et al. Uncovering the protocatechuate 2,3-cleavage pathway genes. J Bacteriol, 2009, 191:6758–6768. pmid:19717587
- View Article
- PubMed/NCBI
- Google Scholar
7. Zhao KX, Huang Y, Chen X, Wang NX, Liu SJ. PcaO positively regulates pcaHG of the β-ketoadipate pathway in Corynebacterium glutamicum. J Bacteriol, 2010, 192:1565–1572. pmid:20081038
- View Article
- PubMed/NCBI
- Google Scholar
8. Wróbel AM, Gregoraszczuk EŁ. Differential effect of methyl-, butyl- and propylparaben and 17β-estradiol on selected cell cycle and apoptosis gene and protein expression in MCF-7 breast cancer cells and MCF-10A non-malignant cells. J Appl Toxicol, 2014, 34:1041–1050. pmid:24481588
- View Article
- PubMed/NCBI
- Google Scholar
9. Ittekkot V. Global trends in the nature of organic matter in river suspensions. Nature, 1988, 332:436–438.
- View Article
- Google Scholar
10. Labeeuw L, Martone P T, Yan B, Case R J. Ancient origin of the biosynthesis of lignin precursors. Biology Direct, 2015, 10(1). pmid:27392031
- View Article
- PubMed/NCBI
- Google Scholar
11. Hedges JI, Mann DC. The characterization of plant tissues by their lignin oxidation products. Geochim Cosmochim Acta, 1979, 43:1803–1807.
- View Article
- Google Scholar
12. Li X, Zhang T, Sun S, Lan H, Yu T. Lignin in marine environment and its analysis-A review. J Ocean Univ China, 2012, 11:501–506.
- View Article
- Google Scholar
13. Woo H L, Hazen T C. Enrichment of bacteria from eastern mediterranean sea involved in lignin degradation via the phenylacetyl-coa pathway. Frontiers in Microbiology, 2018, 9:922. pmid:29867833
- View Article
- PubMed/NCBI
- Google Scholar
14. Loh PS, Miller AE, Reeves AD, Harvey SM, Overnell J. Optimised recovery of lignin-derived phenols in a Scottish fjord by the CuO oxidation method. J Environ Monit, 2008, 10:1187–1194. pmid:19244642
- View Article
- PubMed/NCBI
- Google Scholar
15. Hernes PJ, Benner R. Transport and diagenesis of dissolved and particulate terrigenous organic matter in the North Pacific Ocean. Deep-Sea Res, 2002, 49:2119–2132.
- View Article
- Google Scholar
16. Hernes PJ, Benner R. Photochemical and microbial degradation of dissolved lignin phenols: Implications for the fate of terrigenous dissolved organic matter in marine environments. J Geophys Res, 2003, 108:3291–3299.
- View Article
- Google Scholar
17. Dittmar T, Lara RJ. Molecular evidence for lignin degradation in sulfate-reducing mangrove sediments (Amazonia, Rrasil). Geochim Cosmochim Acta, 2011, 65:1417–1428.
- View Article
- Google Scholar
18. Tian J, Zhu L, Wang W, Zhang L, Li Z, Zhao Q, et al. Genomic analysis of microbulbifer sp. strain A4B-17 and the characterization of its metabolic pathways for 4-hydroxybenzoic acid synthesis. Frontiers in Microbiology, 2018, 9:3115. pmid:30619190
- View Article
- PubMed/NCBI
- Google Scholar
19. Zhan H, Wang H, Liao L, Feng Y, Fan X, Zhang L, et al. Kinetics and novel degradation pathway of permethrin in Acinetobacter baumannii ZH-14. Frontiers in Microbiology, 2018, 9:98. pmid:29456525
- View Article
- PubMed/NCBI
- Google Scholar
20. Maejima Y, Kushimoto K, Muraguchi Y, Fukuda K, Miura T, Yamazoe A, et al. Proteobacteria and Bacteroidetes are major phyla of filterable bacteria passing through 0.22 μm pore size membrane filter, in Lake Sanaru, Hamamatsu, Japan[J]. Bioscience, Biotechnology, and Biochemistry, 2018, 82(7):1260–1263. pmid:29598452
- View Article
- PubMed/NCBI
- Google Scholar
21. Sánchez-Porro C, Kaur B, Mann H, Ventosa A. Halomonas titanicae sp. nov., a halophilic bacterium isolated from the RMS Titanic. Int J Syst Evol Microbiol, 2010, 60:2768–7274. pmid:20061494
- View Article
- PubMed/NCBI
- Google Scholar
22. Chen M, Xiao X, Wang P, Zeng X, Wang F. Arthrobacter ardleyensis sp. nov., isolated from Antarctic lake sediment and deep-sea sediment. Arch Microbiol, 2005, 183:301–305. pmid:15834596
- View Article
- PubMed/NCBI
- Google Scholar
23. Bowman JP. Pseudolteromonas prydzensis sp. nov., a psychrotrophic, halotolerant bacterium from Antarctic sea ice. Int J Syst Bacteriol, 1998, 48:1037–1041. pmid:9734061
- View Article
- PubMed/NCBI
- Google Scholar
24. Romanenko LA, Zhukova NV, Lysenko AM, Mikhailov VV, Stackebrandt E. Assignment of 'Alteromonas marinoglutinosa' NCIMB 1770 to Pseudoalteromonas mariniglutinosa sp. nov., nom. rev., comb. nov. Int J Syst Evol Microbiol, 2003, 53:1105–1109. pmid:12892134
- View Article
- PubMed/NCBI
- Google Scholar
25. Matsuyama H, Sawazaki K, Minami H, Kasahara H, Horikawa K, Yumoto I. Pseudoalteromonas shioyasakiensis sp. nov., a marine polysaccharide-producing bacterium. Int J Syst Evol Microbiol, 2014, 64:101–106. pmid:24021728
- View Article
- PubMed/NCBI
- Google Scholar
26. Chimetto LA, Cleenwerck I, Brocchi M, Willems A, De Vos P, Thompson FL. Marinomonas brasilensis sp. nov., isolated from the coral Mussismilia hispida, and reclassification of Marinomonas basaltis as a later heterotypic synonym of Marinomonas communis. Int J Syst Evol Microbiol, 2011, 61:1170–1175. pmid:20562247
- View Article
- PubMed/NCBI
- Google Scholar
27. López-López A, Pujalte MJ, Benlloch S, Mata-Roig M, Rosselló-Mora R, Garay E, et al. Thalassospira lucentensis gen. nov., sp. nov., a new marine member of the alpha-Proteobacteria. Int J Syst Evol Microbiol, 2002, 52:1277–1283. pmid:12148640
- View Article
- PubMed/NCBI
- Google Scholar
28. Nogi Y, Yoshizumi M, Miyazaki M. Thalassospira povalilytica sp. nov., a polyvinyl-alcohol-degrading marine bacterium. Int J Syst Evol Microbiol, 2014, 64:1149–1153. pmid:24408523
- View Article
- PubMed/NCBI
- Google Scholar
29. Peng X, Misawa N, Harayama S. Isolation and characterization of thermophilic bacilli degrading cinnamic, 4-coumaric, and ferulic acids. Appl Environ Microbiol, 2003, 69:1417–1427. pmid:12620824
- View Article
- PubMed/NCBI
- Google Scholar
30. Ni J, Gao Y Y, Tao F, Liu H Y, Xu P. Temperature-directed biocatalysis for the sustainable production of aromatic aldehydes or alcohols. Angewandte Chemie International Edition, 2018, 57(5):1214–1217. pmid:29178412
- View Article
- PubMed/NCBI
- Google Scholar
31. Schwibbert K, Marin-Sanguino A, Bagyan I, Heidrich G, Lentzen G, Seitz H, et al. A blueprint of ectoine metabolism from the genome of the industrial producer Halomonas elongata DSM 2581 T. Environ Microbiol, 2011, 13:1973–1994. pmid:20849449
- View Article
- PubMed/NCBI
- Google Scholar
32. Lucas-Elío P, Goodwin L, Woyke T, Pitluck S, Nolan M, Kyrpides NC, et al. Complete genome sequence of the melanogenic marine bacterium Marinomonas mediterranea type strain (MMB-1(T)). Stand Genomic Sci, 2012, 6:63–73. pmid:22675599
- View Article
- PubMed/NCBI
- Google Scholar
33. Leveau JH, Gerards S. Discovery of a bacterial gene cluster for catabolism of the plant hormone indole 3-acetic acid. FEMS Microbiol Ecol, 2008, 65:238–250. pmid:18205812
- View Article
- PubMed/NCBI
- Google Scholar
34. Alcolombri U, Laurino P, Lara-Astiaso P, Vardi A, Tawfik DS. DddD is a CoA-transferase/lyase producing dimethyl sulfide in the marine environment. Biochemistry, 2014, 53(34):5473–5475. pmid:25140443
- View Article
- PubMed/NCBI
- Google Scholar
35. Lucas-Elío P, Goodwin L, Woyke T, Pitluck S, Nolan M, Kyrpides NC, et al. Complete genome sequence of Marinomonas posidonica type strain (IVIA-Po-181(T)). Stand Genomic Sci, 2012, 7:31–43. pmid:23458837
- View Article
- PubMed/NCBI
- Google Scholar
36. Kallimanis A, Labutti KM, Lapidus A, Clum A, Lykidis A, Mavromatis K, et al. Complete genome sequence of Arthrobacter phenanthrenivorans type strain (Sphe3). Stand Genomic Sci, 2011, 4:123–130. pmid:21677849
- View Article
- PubMed/NCBI
- Google Scholar
37. Mongodin EF, Shapir N, Daugherty SC, DeBoy RT, Emerson JB, Shvartzbeyn A, et al. Secrets of soil survival revealed by the genome sequence of Arthrobacter aurescens TC1. PLoS Genet, 2006, 2:e214. pmid:17194220
- View Article
- PubMed/NCBI
- Google Scholar
38. Sahoo NK, Pakshirajan K, Ghosh PK. Evaluation of 4-bromophenol biodegradation in mixed pollutants system by Arthrobacter chlorophenolicus A6 in an upflow packed bed reactor. Biodegradation, 2014, 25:705–718. pmid:24934870
- View Article
- PubMed/NCBI
- Google Scholar
39. Monnet C, Loux V, Gibrat JF, Spinnler E, Barbe V, Vacherie B, et al. (2010) The Arthrobacter arilaitensis Re117 genome sequence reveals its genetic adaptation to the surface of cheese. PLoS One 5:e15489. pmid:21124797
- View Article
- PubMed/NCBI
- Google Scholar
40. Lai Q, Shao Z. Genome sequence of Thalassospira xiamenensis type strain M-5. Journal of Bacteriology, 2012, 194:6957. pmid:23209216
- View Article
- PubMed/NCBI
- Google Scholar
41. Goble AM, Fan H, Sali A, Raushel FM. Discovery of a cytokinin deaminase. Chem Biol, 2011, 6:1036–1040.
- View Article
- Google Scholar

[ref1] 1. Pollegioni L, Tonin F, Rosini E. Lignin-degrading enzymes. Febs Journal, 2015, 282(7):1190–1213. pmid:25649492
View Article
PubMed/NCBI
Google Scholar

[2] View Article

[3] PubMed/NCBI

[4] Google Scholar

[ref2] 2. Ruiz-Dueñas FJ, Martínez AT. Microbial degradation of lignin: how a bulky recalcitrant polymer is efficiently recycled in nature and how we can take advantage of this. Microb Biotechnol, 2009, 2:164–177. pmid:21261911
View Article
PubMed/NCBI
Google Scholar

[6] View Article

[7] PubMed/NCBI

[8] Google Scholar

[ref3] 3. Kuzyk ZZA, Goñi MA, Stern GA, Macdonald RW. Sources, pathways and sinks of particulate organic matter in Hudson Bay: Evidence from lignin distributions. Marine Chemistry, 2008, 11:215–229.
View Article
Google Scholar

[10] View Article

[11] Google Scholar

[ref4] 4. Masai E, Katayama Y, Fukuda M. Genetic and biochemical investigations on bacterial catabolic pathways for lignin-derived aromatic compounds. Biosci Biotechnol Biochem, 2007, 71:1–15. pmid:17213657
View Article
PubMed/NCBI
Google Scholar

[13] View Article

[14] PubMed/NCBI

[15] Google Scholar

[ref5] 5. Peng X, Masai E, Kasai D, Miyauchi K, Katayama Y, Fukuda M. A second 5-carboxyvanillate decarboxylase gene, ligW2, is important for lignin-related biphenyl catabolism in Sphingomonas paucimobilis SYK-6. Appl Environ Microbiol, 2005, 71:5014–5021. pmid:16151081
View Article
PubMed/NCBI
Google Scholar

[17] View Article

[18] PubMed/NCBI

[19] Google Scholar

[ref6] 6. Kasai D, Fujinami T, Abe T, Mase K, Katayama Y, Fukuda M, et al. Uncovering the protocatechuate 2,3-cleavage pathway genes. J Bacteriol, 2009, 191:6758–6768. pmid:19717587
View Article
PubMed/NCBI
Google Scholar

[21] View Article

[22] PubMed/NCBI

[23] Google Scholar

[ref7] 7. Zhao KX, Huang Y, Chen X, Wang NX, Liu SJ. PcaO positively regulates pcaHG of the β-ketoadipate pathway in Corynebacterium glutamicum. J Bacteriol, 2010, 192:1565–1572. pmid:20081038
View Article
PubMed/NCBI
Google Scholar

[25] View Article

[26] PubMed/NCBI

[27] Google Scholar

[ref8] 8. Wróbel AM, Gregoraszczuk EŁ. Differential effect of methyl-, butyl- and propylparaben and 17β-estradiol on selected cell cycle and apoptosis gene and protein expression in MCF-7 breast cancer cells and MCF-10A non-malignant cells. J Appl Toxicol, 2014, 34:1041–1050. pmid:24481588
View Article
PubMed/NCBI
Google Scholar

[29] View Article

[30] PubMed/NCBI

[31] Google Scholar

[ref9] 9. Ittekkot V. Global trends in the nature of organic matter in river suspensions. Nature, 1988, 332:436–438.
View Article
Google Scholar

[33] View Article

[34] Google Scholar

[ref10] 10. Labeeuw L, Martone P T, Yan B, Case R J. Ancient origin of the biosynthesis of lignin precursors. Biology Direct, 2015, 10(1). pmid:27392031
View Article
PubMed/NCBI
Google Scholar

[36] View Article

[37] PubMed/NCBI

[38] Google Scholar

[ref11] 11. Hedges JI, Mann DC. The characterization of plant tissues by their lignin oxidation products. Geochim Cosmochim Acta, 1979, 43:1803–1807.
View Article
Google Scholar

[40] View Article

[41] Google Scholar

[ref12] 12. Li X, Zhang T, Sun S, Lan H, Yu T. Lignin in marine environment and its analysis-A review. J Ocean Univ China, 2012, 11:501–506.
View Article
Google Scholar

[43] View Article

[44] Google Scholar

[ref13] 13. Woo H L, Hazen T C. Enrichment of bacteria from eastern mediterranean sea involved in lignin degradation via the phenylacetyl-coa pathway. Frontiers in Microbiology, 2018, 9:922. pmid:29867833
View Article
PubMed/NCBI
Google Scholar

[46] View Article

[47] PubMed/NCBI

[48] Google Scholar

[ref14] 14. Loh PS, Miller AE, Reeves AD, Harvey SM, Overnell J. Optimised recovery of lignin-derived phenols in a Scottish fjord by the CuO oxidation method. J Environ Monit, 2008, 10:1187–1194. pmid:19244642
View Article
PubMed/NCBI
Google Scholar

[50] View Article

[51] PubMed/NCBI

[52] Google Scholar

[ref15] 15. Hernes PJ, Benner R. Transport and diagenesis of dissolved and particulate terrigenous organic matter in the North Pacific Ocean. Deep-Sea Res, 2002, 49:2119–2132.
View Article
Google Scholar

[54] View Article

[55] Google Scholar

[ref16] 16. Hernes PJ, Benner R. Photochemical and microbial degradation of dissolved lignin phenols: Implications for the fate of terrigenous dissolved organic matter in marine environments. J Geophys Res, 2003, 108:3291–3299.
View Article
Google Scholar

[57] View Article

[58] Google Scholar

[ref17] 17. Dittmar T, Lara RJ. Molecular evidence for lignin degradation in sulfate-reducing mangrove sediments (Amazonia, Rrasil). Geochim Cosmochim Acta, 2011, 65:1417–1428.
View Article
Google Scholar

[60] View Article

[61] Google Scholar

[ref18] 18. Tian J, Zhu L, Wang W, Zhang L, Li Z, Zhao Q, et al. Genomic analysis of microbulbifer sp. strain A4B-17 and the characterization of its metabolic pathways for 4-hydroxybenzoic acid synthesis. Frontiers in Microbiology, 2018, 9:3115. pmid:30619190
View Article
PubMed/NCBI
Google Scholar

[63] View Article

[64] PubMed/NCBI

[65] Google Scholar

[ref19] 19. Zhan H, Wang H, Liao L, Feng Y, Fan X, Zhang L, et al. Kinetics and novel degradation pathway of permethrin in Acinetobacter baumannii ZH-14. Frontiers in Microbiology, 2018, 9:98. pmid:29456525
View Article
PubMed/NCBI
Google Scholar

[67] View Article

[68] PubMed/NCBI

[69] Google Scholar

[ref20] 20. Maejima Y, Kushimoto K, Muraguchi Y, Fukuda K, Miura T, Yamazoe A, et al. Proteobacteria and Bacteroidetes are major phyla of filterable bacteria passing through 0.22 μm pore size membrane filter, in Lake Sanaru, Hamamatsu, Japan[J]. Bioscience, Biotechnology, and Biochemistry, 2018, 82(7):1260–1263. pmid:29598452
View Article
PubMed/NCBI
Google Scholar

[71] View Article

[72] PubMed/NCBI

[73] Google Scholar

[ref21] 21. Sánchez-Porro C, Kaur B, Mann H, Ventosa A. Halomonas titanicae sp. nov., a halophilic bacterium isolated from the RMS Titanic. Int J Syst Evol Microbiol, 2010, 60:2768–7274. pmid:20061494
View Article
PubMed/NCBI
Google Scholar

[75] View Article

[76] PubMed/NCBI

[77] Google Scholar

[ref22] 22. Chen M, Xiao X, Wang P, Zeng X, Wang F. Arthrobacter ardleyensis sp. nov., isolated from Antarctic lake sediment and deep-sea sediment. Arch Microbiol, 2005, 183:301–305. pmid:15834596
View Article
PubMed/NCBI
Google Scholar

[79] View Article

[80] PubMed/NCBI

[81] Google Scholar

[ref23] 23. Bowman JP. Pseudolteromonas prydzensis sp. nov., a psychrotrophic, halotolerant bacterium from Antarctic sea ice. Int J Syst Bacteriol, 1998, 48:1037–1041. pmid:9734061
View Article
PubMed/NCBI
Google Scholar

[83] View Article

[84] PubMed/NCBI

[85] Google Scholar

[ref24] 24. Romanenko LA, Zhukova NV, Lysenko AM, Mikhailov VV, Stackebrandt E. Assignment of 'Alteromonas marinoglutinosa' NCIMB 1770 to Pseudoalteromonas mariniglutinosa sp. nov., nom. rev., comb. nov. Int J Syst Evol Microbiol, 2003, 53:1105–1109. pmid:12892134
View Article
PubMed/NCBI
Google Scholar

[87] View Article

[88] PubMed/NCBI

[89] Google Scholar

[ref25] 25. Matsuyama H, Sawazaki K, Minami H, Kasahara H, Horikawa K, Yumoto I. Pseudoalteromonas shioyasakiensis sp. nov., a marine polysaccharide-producing bacterium. Int J Syst Evol Microbiol, 2014, 64:101–106. pmid:24021728
View Article
PubMed/NCBI
Google Scholar

[91] View Article

[92] PubMed/NCBI

[93] Google Scholar

[ref26] 26. Chimetto LA, Cleenwerck I, Brocchi M, Willems A, De Vos P, Thompson FL. Marinomonas brasilensis sp. nov., isolated from the coral Mussismilia hispida, and reclassification of Marinomonas basaltis as a later heterotypic synonym of Marinomonas communis. Int J Syst Evol Microbiol, 2011, 61:1170–1175. pmid:20562247
View Article
PubMed/NCBI
Google Scholar

[95] View Article

[96] PubMed/NCBI

[97] Google Scholar

[ref27] 27. López-López A, Pujalte MJ, Benlloch S, Mata-Roig M, Rosselló-Mora R, Garay E, et al. Thalassospira lucentensis gen. nov., sp. nov., a new marine member of the alpha-Proteobacteria. Int J Syst Evol Microbiol, 2002, 52:1277–1283. pmid:12148640
View Article
PubMed/NCBI
Google Scholar

[99] View Article

[100] PubMed/NCBI

[101] Google Scholar

[ref28] 28. Nogi Y, Yoshizumi M, Miyazaki M. Thalassospira povalilytica sp. nov., a polyvinyl-alcohol-degrading marine bacterium. Int J Syst Evol Microbiol, 2014, 64:1149–1153. pmid:24408523
View Article
PubMed/NCBI
Google Scholar

[103] View Article

[104] PubMed/NCBI

[105] Google Scholar

[ref29] 29. Peng X, Misawa N, Harayama S. Isolation and characterization of thermophilic bacilli degrading cinnamic, 4-coumaric, and ferulic acids. Appl Environ Microbiol, 2003, 69:1417–1427. pmid:12620824
View Article
PubMed/NCBI
Google Scholar

[107] View Article

[108] PubMed/NCBI

[109] Google Scholar

[ref30] 30. Ni J, Gao Y Y, Tao F, Liu H Y, Xu P. Temperature-directed biocatalysis for the sustainable production of aromatic aldehydes or alcohols. Angewandte Chemie International Edition, 2018, 57(5):1214–1217. pmid:29178412
View Article
PubMed/NCBI
Google Scholar

[111] View Article

[112] PubMed/NCBI

[113] Google Scholar

[ref31] 31. Schwibbert K, Marin-Sanguino A, Bagyan I, Heidrich G, Lentzen G, Seitz H, et al. A blueprint of ectoine metabolism from the genome of the industrial producer Halomonas elongata DSM 2581 T. Environ Microbiol, 2011, 13:1973–1994. pmid:20849449
View Article
PubMed/NCBI
Google Scholar

[115] View Article

[116] PubMed/NCBI

[117] Google Scholar

[ref32] 32. Lucas-Elío P, Goodwin L, Woyke T, Pitluck S, Nolan M, Kyrpides NC, et al. Complete genome sequence of the melanogenic marine bacterium Marinomonas mediterranea type strain (MMB-1(T)). Stand Genomic Sci, 2012, 6:63–73. pmid:22675599
View Article
PubMed/NCBI
Google Scholar

[119] View Article

[120] PubMed/NCBI

[121] Google Scholar

[ref33] 33. Leveau JH, Gerards S. Discovery of a bacterial gene cluster for catabolism of the plant hormone indole 3-acetic acid. FEMS Microbiol Ecol, 2008, 65:238–250. pmid:18205812
View Article
PubMed/NCBI
Google Scholar

[123] View Article

[124] PubMed/NCBI

[125] Google Scholar

[ref34] 34. Alcolombri U, Laurino P, Lara-Astiaso P, Vardi A, Tawfik DS. DddD is a CoA-transferase/lyase producing dimethyl sulfide in the marine environment. Biochemistry, 2014, 53(34):5473–5475. pmid:25140443
View Article
PubMed/NCBI
Google Scholar

[127] View Article

[128] PubMed/NCBI

[129] Google Scholar

[ref35] 35. Lucas-Elío P, Goodwin L, Woyke T, Pitluck S, Nolan M, Kyrpides NC, et al. Complete genome sequence of Marinomonas posidonica type strain (IVIA-Po-181(T)). Stand Genomic Sci, 2012, 7:31–43. pmid:23458837
View Article
PubMed/NCBI
Google Scholar

[131] View Article

[132] PubMed/NCBI

[133] Google Scholar

[ref36] 36. Kallimanis A, Labutti KM, Lapidus A, Clum A, Lykidis A, Mavromatis K, et al. Complete genome sequence of Arthrobacter phenanthrenivorans type strain (Sphe3). Stand Genomic Sci, 2011, 4:123–130. pmid:21677849
View Article
PubMed/NCBI
Google Scholar

[135] View Article

[136] PubMed/NCBI

[137] Google Scholar

[ref37] 37. Mongodin EF, Shapir N, Daugherty SC, DeBoy RT, Emerson JB, Shvartzbeyn A, et al. Secrets of soil survival revealed by the genome sequence of Arthrobacter aurescens TC1. PLoS Genet, 2006, 2:e214. pmid:17194220
View Article
PubMed/NCBI
Google Scholar

[139] View Article

[140] PubMed/NCBI

[141] Google Scholar

[ref38] 38. Sahoo NK, Pakshirajan K, Ghosh PK. Evaluation of 4-bromophenol biodegradation in mixed pollutants system by Arthrobacter chlorophenolicus A6 in an upflow packed bed reactor. Biodegradation, 2014, 25:705–718. pmid:24934870
View Article
PubMed/NCBI
Google Scholar

[143] View Article

[144] PubMed/NCBI

[145] Google Scholar

[ref39] 39. Monnet C, Loux V, Gibrat JF, Spinnler E, Barbe V, Vacherie B, et al. (2010) The Arthrobacter arilaitensis Re117 genome sequence reveals its genetic adaptation to the surface of cheese. PLoS One 5:e15489. pmid:21124797
View Article
PubMed/NCBI
Google Scholar

[147] View Article

[148] PubMed/NCBI

[149] Google Scholar

[ref40] 40. Lai Q, Shao Z. Genome sequence of Thalassospira xiamenensis type strain M-5. Journal of Bacteriology, 2012, 194:6957. pmid:23209216
View Article
PubMed/NCBI
Google Scholar

[151] View Article

[152] PubMed/NCBI

[153] Google Scholar

[ref41] 41. Goble AM, Fan H, Sali A, Raushel FM. Discovery of a cytokinin deaminase. Chem Biol, 2011, 6:1036–1040.
View Article
Google Scholar

[155] View Article

[156] Google Scholar

Isolation and characterization marine bacteria capable of degrading lignin-derived compounds

Isolation and characterization marine bacteria capable of degrading lignin-derived compounds

Correction

Figures

Abstract

Introduction

Materials and methods

Study site and sample collection

Chemicals and material

Isolation and characterization of vanillic acid degraders

Optimum growth conditions for temperature and NaCl concentration

Degradation of different lignin-derived compounds by isolates

Degradation products of ferulic acid by isolates

Analysis of metabolism of lignin-derived compounds by HPLC and GC-MS

Enzyme assays for PCA dioxygenase

Results

Isolation of vanillic acid degraders

Optimum growth conditions for temperature and NaCl concentration

Ability to utilize lignin-derived compounds

Detection of ferulic acid intermediates

Enzyme assay for aromatic ring-fission reaction

Discussion

Conclusion

Supporting information

S1 Fig. Electrophoresis image of 16S rDNA PCR products.

References