Environmental Sustainability

, Volume 2, Issue 4, pp 381–389 | Cite as

Potential of Bacillus subtilis from marine environment to degrade aromatic hydrocarbons

  • Daisy VelupillaimaniEmail author
  • Arunachalam Muthaiyan
Original Article


Microbial degradation of aromatic pollutants has been a promising method for bioremediation and restoring environmental damage. Protocatechuate (PCA) is a common intermediate in the microbial degradation of several aromatic compounds. The present study reports the identification of protocatechuate 3,4-dioxygenase (3,4-PCD)—a key enzyme of the β-ketoadipate pathway, in Bacillus subtilis isolated from coastal water containing waste discharge from paper mills, textile industries, and timber processing factories. The strain efficiently degraded PCA up to 20 mM concentration. Enzyme assay indicated the production of 3,4-PCD and cleavage of the benzene ring at ortho position and the formation of keto compounds of the beta- ketoadipate pathway. The utilization of PCA by plasmid cured cells indicates PCA metabolism is encoded by chromosomal genes. The metabolic potential of the B. subtilis isolate makes it a promising tool for bioremediation of aromatic pollutants present in marine environments.


Bacillus subtilis Protocatechuate Intradiol cleavage Protocatechuate 3,4-dioxygenase Plasmid Bioremediation 


Biodegradation by the natural population of microorganisms is cost-effective and environmentally sustainable treatment strategy (Bjerketorp et al. 2018). Although microorganisms capable of degradation of aromatic compounds have been investigated extensively, information about microbial degradation in marine environments is still very limited (Vieira et al. 2018).

Lignin, is the most abundant renewable aromatic material on earth and the largest source of aromatic building blocks (Ponnusamy et al. 2019; Sun et al. 2018). It is a complex, chemically stable, aromatic heteropolymer present in the cell walls of vascular plants (Abdelaziz et al. 2016). Despite its recalcitrance to breakdown, studies indicate the degradation of lignin by soil microorganisms (Ayeronfe et al. 2018; Xu et al. 2018).

Lignin degradation by fungi has been extensively studied when compared to bacteria (DeAngelis et al. 2013; Fang et al. 2018). In bacterial lignin degradation, most of the current knowledge is still based primarily on studies of soil bacteria (Xu et al. 2019).

Lately, studies on bacterial lignolytic enzymes in the degradation of lignin and intermediate compounds have been intensified (Abdelaziz et al. 2016; de Gonzalo et al. 2016; Fisher and Fong 2014; Hogancamp and Raushel 2018). Microbial degradation of lignin employs different pathways to yield aromatic acid intermediates such as protocatechuate (PCA) and catechol, which further get metabolized via beta-ketoadipate pathway (BKP) (Wu et al. 2017). Ring-cleaving dioxygenases catalyze intradiol cleavage of aromatic compounds such as catechol and PCA to produce intermediates that enter the carbon cycle (Tian et al. 2017).

Protocatechuate 3,4-dioxygenase (3,4-PCD), a key enzyme in the BKP, plays an essential role in the breakdown of numerous aromatic and hydroaromatic substances (Zhu et al. 2018). Recent research has shown the potential use of immobilized 3,4-PCD in degradation of the toxic 3,4-dihydroxybenzoic acid (3,4-DHBA) from industrial food processing waste water effluents (Zhang et al. 2017). 3,4-PCD from Stenotrophomonas maltophilia (KB2) reportedly shows wide substrate specificity that makes it a useful tool in applications for bioremediation or bioaugmentation purposes (Garrido-Sanz et al. 2018; Guzik et al. 2014).

The dissimilation of ferulic acid, an important constituent of lignin, has been reported in Bacillus subtilis (Gurujeyalakshmi and Mahadevan 1987; Ravi et al. 2017). Clearly B. subtilis has metabolic potential to degrade aromatic substances that are part of the lignin polymer. Moreover, B. subtilis displays exceedingly low pathogenicity to humans and was granted GRAS (generally regarded as safe) status by the U.S. Food and Drug Administration (De Boer and Diderichsen 1991; Jeżewska-Frąckowiak et al. 2018). Therefore, the wide use of B. subtilis in the fermentation industry makes it a perfect choice for use in increased production of enzymes. In this study, the breakdown of PCA—one of the precursors of lignin degradation, by B. subtilis isolated from the marine environment—was investigated and the results are reported.

Materials and methods

Bacterial strain, growth conditions, and media

B. subtilis (MDB1), isolated from the coastal waters (Chennai, Tamilnadu, India) containing waste discharge from paper mills, textile industries, and timber processing factories, was obtained from Prof. A. Mahadevan, CAS in Botany, University of Madras, Chennai 600,025 and used in this study.

The culture was maintained in minimal medium (MM) containing (NH4)2HPO4—3.0 g, K2HPO4—1.2 g, NaCl—0.6 g in 1 l of water (pH 7.5) with glycerol (2.6%) as the sole carbon source. At 48 h, the cells were routinely subcultured to get uninduced cells. For short-term storage, the culture was grown in Luria–Bertani (LB) medium (Sambrook and Russell 2001) amended with PCA (5 mM). The media were adjusted to pH 7.5 and solidified with 2% agar when needed. For testing its ability to grow on glycerol, glucose, and PCA as sole carbon sources, 48-h culture of B. subtilis was used to inoculate MM broth amended with either PCA (5 mM), or glycerol (10%), or glucose (10 mM). The carbon sources were filter-sterilized through 0.45 µ Millipore filter and added to the autoclaved media before inoculation. The culture was prepared by aseptically transferring 0.1 ml of stock culture into 20 ml of the medium and incubated at 37 °C on a rotary incubator at 150 rpm. Growth was monitored at regular intervals by measuring the OD at 540 nm by spectrophotometer (Phillips, PU 8400).

Preparation of cell-free enzyme

PCD-grown B. subtilis cells were harvested by centrifugation at 12,000×g at 4 °C for 10 min. The cells were washed and suspended in 10 ml of phosphate buffer (0.1 M, pH 7.0) containing 17% sucrose (w/v), 0.1% ascorbic acid (w/v), and 0.1% cysteine–HCl. The cells were disrupted in an ultrasonic disintegrator (Braun, Model 2000) for 1 min and centrifuged at 15,000×g for 20 min at 4 °C to separate the cell debris and unbroken cells. The supernatant was further saturated with equal volume of ice-cold acetone and incubated at 0 °C for 24 h to maximize precipitation. The sample was dialyzed for another 48 h in Tris.HCl buffer (pH 7.2) to remove salt contamination, lyophilized, dissolved in 0.5 M Tris.HCl (pH 7.2), and assayed immediately for enzyme activity.

3,4-PCD enzyme assay

To screen for the presence of intradiol ring cleavage dioxygenase activity in crude cell extracts, Rothera’s test was performed. 3,4-PCD was assayed by the method of Stanier and Ingraham (1954). The reaction mixture contained PCA 0.1 mL (10 μg), 1 mL enzyme, and 1.9 mL 0.5 M phosphate buffer (pH 7.2). Cleavage of PCA by 3,4-PCD was assayed spectrophotometrically by following the disappearance of substrate at 290 nm and appearance of the product, beta- carboxy cis,cis–muconic acid at 260 nm. In the controls, the enzyme was replaced with glass-distilled water. The standard curve for PCA was determined by plotting the gradient concentration of PCA against their absorbance at 290 nm.

Antibiotic susceptibility test

The antibiogram of B. subtilis was determined by using antibiotic sensitivity discs (Hi Media; Mumbai). The antibiotics and concentration used are listed in Table 1. The antibiotic susceptibility spectrum was determined by Kirby-Bauer disc-diffusion method. LB-grown B. subtilis culture (100 μL) with OD 1.0 at 540 nm was used to inoculate Mueller–Hinton agar plates. The antibiotic discs were placed on the inoculated agar surface and the plates were incubated at 37 °C for 24 h. The zone of inhibition for different antibiotics was measured to obtain the susceptibility profile.
Table 1

Antibiotic sensitivity pattern of B. subtilis


Concentration (µg)

Susceptible (+)/Resistance (−)






























Nalidixic acid



Isolation and curing of plasmid DNA

Indigenous plasmid DNA from B. subtilis was isolated from cells grown for 16 h at 37 °C in LB medium amended with ampicillin (75 μg/ml). Extracted plasmid DNA was purified using the Qiagen-Qia prep mini column (Qiagen, Germany). Agarose gels (0.8% or 1% w/v) were used to resolve the DNA samples and stained with ethidium bromide (0.5% v/v).

Plasmid curing was carried out by growing B. subtilis cells in LB medium supplemented with mitomycin C (20 μg/mL). The treated culture was incubated at 37 °C in a rotary shaker at 150 rpm. To select for plasmid cured B. subtilis, log phase cells were serially diluted, spread on LB agar plates and incubated at 37 °C. Colonies were randomly picked and grown in 10 mL LB and screened for plasmids. The cured colonies were sub-cultured in growth medium containing streptomycin (100 µg/mL) and PCA to check for their degradative ability.


Growth of B. subtilis on single carbon-sources

The growth pattern of B. subtilis on glycerol, glucose, and PCA as sole carbon sources was determined (Fig. 1). On MM amended with glycerol (10%), the cells required a prolonged period of multiplication after a lag phase that extended to 8 h. The log phase was attained after 34 h of incubation. The stationary phase was reached at 36 h. When glucose (10 mM) was substituted for glycerol in the medium, the log phase was reached even before 6 h and continued until 32 h of incubation. Thus, glucose favored good growth of B. subtilis. The ability of B. subtilis to utilize PCA as the sole source of carbon and energy was tested by replacing the glycerol with 5 mM PCA. B. subtilis exhibited significant growth on media containing PCA as the sole carbon and energy source. PCA was readily utilized by B. subtilis cells. The cells had a lag phase of 8 h; the growth curve reached its maximum at 32 h of incubation and thereafter the cells entered the stationary phase. Growth increased with an increase in the concentration of PCA and reached maximum at 20 mM. However, increasing the concentration to 40 mM was toxic to the cells. PCA at 40 mM was inhibitory to the growth of B. subtilis (Fig. 2).
Fig. 1

Growth of B. subtilis on different carbon-sources. This replicated growth pattern (n = 3) of B. subtilis on MM amended with glycerol (10%), glucose (10 mM) and PCA (5 mM) shows good growth on all three carbon sources. The cells recorded an extended lag phase in glycerol and PCA while glucose favored good growth. Error bars represent the mean of data from triplicate assays

Fig. 2

Growth of B. subtilis on different concentrations of PCA. This replicated growth pattern (n = 3) of B. subtilis cells cultured on different concentrations of PCA shows an increase in growth corresponding to an increase in concentration. Maximum growth was recorded at 20 mM PCA. Error bars represent the mean of data from triplicate assays

Growth of B. subtilis on dual carbon-sources

Cells cultured on glycerol when transferred to MM medium containing both glycerol (10%) and PCA (5 mM) exhibited a prolonged lag phase. However, the growth rate of PCA-grown B. subtilis cells on glycerol + PCA was greatly enhanced. This growth pattern indicates the role of glycerol as a co-metabolite for the utilization of PCA as the sole source of carbon and energy in B. subtilis (Fig. 3). Additionally, the influence of glucose (10 mM) on the utilization of PCA by B. subtilis was investigated. Cells grown on PCA (5 mM) multiplied rapidly when transferred to MM containing PCA alone. However, glucose-grown cells when transferred to medium containing PCA alone, exhibited a lag phase of 8 h (Fig. 4). The growth patterns for PCA and glucose-grown cells on glucose were almost similar (Fig. 5). Cells exposed to PCA and transferred to medium containing both glucose and PCA continued to grow up to 32 h without any lag phase. The growth pattern of glucose-grown cells on glucose + PCA was the same up to 8 h, thereafter the cells entered the decline phase (Fig. 6). The initial growth pattern on dual carbon sources was almost identical to the growth on glucose alone. This indicated the preferential utilization of glucose during the log phase. Regardless of their prior induction on PCA, B. subtilis utilized PCA only after a lag phase of 4 h (Fig. 3). The presence of glucose (10 mM) along with PCA (5 mM) resulted in diauxic growth of B. subtilis.
Fig. 3

Growth of B. subtilis on glycerol + PCA. This replicated growth pattern (n = 3) of B. subtilis cells cultured on glycerol and transferred to MM amended with Glycerol (10%), and PCA (5 mM) shows prolonged lag phase. However, growth rate of PCA- grown cells was greatly enhanced on MM containing glycerol + PCA. Error bars represent the mean of data from triplicate assays

Fig. 4

Growth of B. subtilis on PCA + Glucose. This replicated growth pattern (n = 3) of B. subtilis cells cultured on PCA (5 mM) and transferred to MM with PCA alone multiplied rapidly. However, glucose-grown cells when transferred to medium containing PCA alone, exhibited a lag phase of 8 h. Error bars represent the mean of data from triplicate assays

Fig. 5

Growth of B. subtilis on PCA (alone) and glucose (alone). This replicated growth pattern (n = 3) of B. subtilis for PCA- and glucose-grown cells on glucose alone were almost similar without a lag phase. Error bars represent the mean of data from triplicate assays

Fig. 6

Growth of B. subtilis on glucose + PCA. This replicated growth pattern (n = 3) of glucose-grown B. subtilis cells on glucose + PCA showed similar growth rate up to 8 h but the growth rate decreased thereafter. Error bars represent the mean of data from triplicate assays

3,4-PCD enzyme assay

PCA up to 20 mM favored growth of B. subtilis cells. Clearly, PCA induced the enzymes associated with its oxidation. Hence, 3,4-PCD enzyme activity was assayed. The crude enzyme from B. subtilis cells grown on PCA did not develop deep yellow color, indicating the absence of meta cleavage of the aromatic ring. The contents turned deep purple in color indicating ortho cleavage of PCA and appearance of keto compounds of β-ketoadipate pathway.

The presence of beta- carboxy cis, cis–muconic acid was indicated by the increase in absorbance at 260 nm and confirmed the ortho cleavage of PCA by 3,4-PCD (Fig. 7).
Fig. 7

Intradiol cleaving enzyme assay. Enzyme was assayed in a spectrophotometer based on the disappearance of the substrate PCA at 290 nm (λ max for PCA) and appearance of the product, β-carboxy cis, cis–muconate at 260 nm (λ max for β-carboxy cis, cis–muconate). Increase in absorbance at 260 nm indicated appearance of β-carboxy cis, cis–muconate and confirms ortho cleavage of PCA by 3, 4-PCD. Error bars represent the mean of data from triplicate assays

Antibiotic susceptibility test

The antibiotic susceptibility spectrum of B. subtilis to antibiotics was evaluated. Depending upon the zone of inhibition and using the standard chart for antibiotic discs, it was evident that B. subtilis was resistant to streptomycin (25 μg/disc). The cells were sensitive to novobiocin, erythromycin, carbenicillin, tetracycline, chloramphenicol, kanamycin, neomycin, bacitracin, ampicillin, and nalidixic acid (Table 1).

Isolation and curing of plasmid DNA

B. subtilis MDB1 was screened for the presence of plasmids. The yield of plasmid DNA varied from 5 to 10 μg/mL culture. A single plasmid with low mobility was observed on the gel (Fig. 8). To check whether the PCA degradation trait of B. subtilis is encoded in plasmid DNA or genomic DNA, plasmid curing was performed (Fig. 9). Plasmid-cured cells were sub-cultured in medium containing streptomycin (100 µg/mL) and PCA to check for degradative ability. Growth of colonies indicates the Streptomycin resistant and PCA degrading genes are not carried on the plasmid but encoded in chromosomal DNA.
Fig. 8

DNA profile of B. subtilis. Lanes 1 & 2—arrow indicates the mega plasmid DNA above the chromosomal DNA; Lane 3—marker λ Hind III digest

Fig. 9

Curing of plasmid DNA in B. subtilis cells. Lane 1 arrow indicates plasmid DNA cured; Lane 2- marker λ Hind III digest


Lignin, present in agricultural and industrial waste is a complex, recalcitrant compound and poses a significant challenge to the environment (Ayeronfe et al. 2018). Marine sediments serve as a suitable sink for aromatic compounds derived from industrial wastes and sewage effluents in aquatic environments (Zhuang et al. 2019). Hence, there is a great need for increasing the knowledge about the microbial degradation of recalcitrant compounds present in the marine environment, especially mechanisms, enzymology and genetic basis of degradation. In present study, we have described the aromatic degradation ability of the marine isolate, B. subtilis MDB1.

Since this B. subtilis strain was isolated from polluted coastal water, it might have survived along with other easily metabolizable carbon compounds that serve as an initial energy source required for breakdown of complex recalcitrant aromatic compounds. PCA has been reported as an intermediate in the biodegradation of many aromatic substances (Arunachalam et al. 2003; Guevara et al. 2019; Wu et al. 2017). Therefore, we studied the effect of simple carbon sources on the utilization of PCA by B. subtilis. According to our results, glucose supported good growth of B. subtilis, while an extended lag phase was observed in glycerol and PCA. This is similar to reports of preferential utilization of glucose as a carbon source by Serratia marcescens, Klebsiella pneumonia and Citrobacter sp. (Chandra et al. 2011).

Microorganisms readily metabolize preferred carbon sources which results in accumulation of non-preferred, complex aromatic compounds that are not easily degraded (Buffing et al. 2018). Moreover, some bacteria utilized another carbon compound to derive energy required for lignin degradation (Zhu et al. 2017). However, in our study B. subtilis was able to utilize PCA (20 mM) as a sole carbon source without the support of easily metabolizable glucose. Our results are similar to reports of Stenotrophomonas sp. and B. subtilis isolated from a palm oil plantation that reduced alkali lignin without the requirement of additional carbon sources (Azman et al. 2019).

In our growth study on single carbon sources, PCA (20 mM) favored the growth of B. subtilis whereas a higher concentration (40 mM) had an inhibitory effect. This result is similar to reports of inhibitory effect of high concentration of PCA due to interference with substrate transport across the cell membrane in Pseudomonas putida and disruption of cytoplasmic membranes in Escherichia coli (Bernal-Mercado et al. 2018; Harwood et al. 1994).

Easily metabolizable sugars such as glucose when added to the culture media serve as cometabolites and enhance utilization of aromatic compounds by lignin degrading microbial population (Chandra et al. 2011; Kuang et al. 2018; Xu et al. 2018). Therefore, in this study we tested the utilization of PCA by glucose-grown cells of B. subtilis. In dual carbon source growth study, B. subtilis cells preferentially utilized glucose and upon its completion, a second phase of growth occurred with the utilization of PCA. This diauxic pattern of growth occurs in cells when induction to the less preferred carbon source was prevented in the presence of the preferred substrate (Wang et al. 2019). The preferential utilization of substrates causes carbon catabolite repression of complex compounds, increasing their recalcitrance in the environment (Choudhary et al. 2017; Martínez-Valenzuela et al. 2018). Hence, the utilization of glucose and PCA by B. subtilis in the present study suggests its potential application in degradation of aromatic pollutants present with other nutrients in the marine environment.

Dioxygenases are natural enzymes for lignin degradation (Wang et al. 2017). In our study, the enzyme assay results indicated the ortho-cleavage of PCA by 3,4-PCD. The present study  reports the presence of 3,4-PCD, a key enzyme of the PCA degradation in B. subtilis. The intradiol or ortho- ring cleavage pathway commonly known as the BKP has a significant role in the degradation of many aromatic substances in nature (Tian et al. 2017; Żur et al. 2018). 3,4-PCD, has great potential for environmental bioremediation (Zhang et al. 2017). The reported efficiency of immobilized 3,4-PCD paves a future application for water purification (Das et al. 2016).

Plasmids encode beneficial traits that include the ability to degrade different aromatic compounds. Genes for aromatic metabolic pathways are often plasmid-encoded (Suvorova and Gelfand 2019). Association between drug resistance and aromatic degradation pathways encoded by plasmids has been reported in bacteria (Jutkina et al. 2011; Yan and Wu 2017). Therefore, resistance of B. subtilis MDB1 to antibiotics was evaluated by antimicrobial sensitivity discs. B. subtilis MDB1 was resistant to streptomycin. However, the absence of plasmids did not affect streptomycin resistance and PCA oxidation. Plasmid cured B. subtilis cells utilized PCA as the sole carbon source. Clearly, the plasmid in B. subtilis is not associated with PCA degradation. These findings demonstrate that the PCA metabolic machinery of B. subtilis is encoded by chromosomal genes. This is similar to other reports of chromosomal catabolic pathways for the assimilation of aromatic compounds (Guevara et al. 2019; Morales et al. 2004). Chromosomal genes encoding metabolic pathways are more stable when compared to plasmid encoded metabolic machinery (Xin et al. 2019). This renders B. subtilis MDB1 a good candidate as an inexpensive and sustainable alternative for facilitating bioremediation of environmental pollutants.

In summary, besides the bioremediation aspect of lignin degradation, interest in lignin as an underexploited carbon source has significantly increased in the last two decades, as evidenced by published reports on lignin depolymerization and valorization (Brink et al. 2019; Chio et al. 2019; Henson et al. 2018; Hirose et al. 2018; Mei et al. 2019; Ponnusamy et al. 2019; Shanmugam et al. 2019; Xu et al. 2018; Zhang et al. 2019). Lignocellulose can serve as a sustainable bioenergy source to meet the growing demand for fuels and chemicals (Sun et al. 2018). Moreover, marine microbial enzymes are naturally halotolerant and better retain their conformation even under high ionic strength conditions (Woo and Hazen 2018). Findings of the present study contribute valuable information to this emerging new facet of biodegradation research that involves discovery of bacterial strains in the marine environment for production of useful products (fuels, chemicals and materials) from recalcitrant biomass.


This study has explored the aromatic degradation potential of a marine isolate B. subtilis MDB1. Information about microbial degradation in marine environments is still very limited. Considering the increasing interest in discovery of efficient lignin depolymerizing microbes, these findings provide new direction towards the underexplored marine environment as a potential reservoir for aromatic degrading bacteria. Although 3,4-PCD plays a pivotal role in the degradation of many aromatic substances in nature, the complete sequencing of the B. subtilis (168) genome makes no mention of PCD genes. Therefore, the preliminary findings from this study form the framework for further advanced work involving the characterization and sequencing of PCD genes in B. subtilis. This will be a valuable contribution to supplement the lack of information about protocatechuate-degrading gene sequence in B. subtilis. Microbial degradation of aromatic environmental pollutants involves complex processes in microbial communities. The ability to form spores, secrete halotolerant enzymes and its low pathogenicity to humans may confer a competitive advantage to B. subtilis in bioremediation of contaminated coastal waters. Although, lignin waste disposal can be achieved by various chemical and physical methods, microbial biodegradation is eco-friendly and the natural competency of B. subtilis will facilitate an enduring sustenance of the marine environment.



The bacterial culture and resources provided by Prof. A. Mahadevan, Center for Advanced Studies in Botany, University of Madras, Chennai, India to conduct this study is thankfully acknowledged. The authors thank Amanda Newsum, Editor, Grand Canyon Education, Phoenix, Arizona, USA, for helping with the editing needs of this manuscript.


  1. Abdelaziz OY, Brink DP, Prothmann J et al (2016) Biological valorization of low molecular weight lignin. Biotechnol Adv 34:1318–1346Google Scholar
  2. Arunachalam M, Mohanraj M, Mohan N, Mahadevan A (2003) Biodegradation of catechin. Proc Indian Nat Sci Acad B69(4):353–370Google Scholar
  3. Ayeronfe F, Kassim A, Ishak N et al (2018) A review on microbial degradation of lignin. Adv Sci Lett 24:4407–4413. Google Scholar
  4. Azman NF, Megat Mohd Noor MJ, MD Akhir FN et al (2019) Depolymerization of lignocellulose of oil palm empty fruit bunch by thermophilic microorganisms from tropical climate. Bioresour Technol. Google Scholar
  5. Bernal-Mercado AT, Vazquez-Armenta FJ, Tapia-Rodriguez MR et al (2018) Comparison of single and combined use of catechin, protocatechuic, and vanillic acids as antioxidant and antibacterial agents against uropathogenic Escherichia coli at planktonic and biofilm levels. Mol (Basel Switz) 23(11):2813. Google Scholar
  6. Bjerketorp J, Röling W, Feng XM, Garcia AH, Heipieper HJ, Håkansson S (2018) Formulation and stabilization of an Arthrobacter strain with good storage stability and 4- chlorophenol-degradation activity for bioremediation. App Microbiol Biotechnol 102(4):2031–2040. Google Scholar
  7. Brink DP, Ravi K, Lidén G, Gorwa-Grauslund MF (2019) Mapping the diversity of microbial lignin catabolism: experiences from the eLignin database. Appl Microbiol Biotechnol 103:3979–4002Google Scholar
  8. Buffing MF, Link H, Christodoulou D, Sauer U (2018) Capacity for instantaneous catabolism of preferred and non-preferred carbon sources in Escherichia coli and Bacillus subtilis. Sci Rep 8(1):11760. Google Scholar
  9. Chandra R, Abhishek A, Sankhwar M (2011) Bacterial decolorization and detoxification of black liquor from rayon grade pulp manufacturing paper industry and detection of their metabolic products. Bioresour Technol 102:6429–6436. Google Scholar
  10. Chio C, Sain M, Qin W (2019) Lignin utilization: a review of lignin depolymerization from various aspects. Renew Sustain Energy Rev 107:232–249. Google Scholar
  11. Choudhary A, Modak A, Apte SK, Phale PS (2017) Transcriptional modulation of transport- and metabolism-associated gene clusters leading to utilization of benzoate in preference to glucose in Pseudomonas putida CSV86. Appl Env Microbiol 83(19):e01280-17. Google Scholar
  12. Das R, Hamid SBA, Annuar MSM (2016) Highly efficient and stable novel nanobiohybrid catalyst to avert 3, 4-dihydroxybenzoic acid pollutant in water. Sci Rep 6:33572. Google Scholar
  13. De Boer AS, Diderichsen B (1991) On the safety of Bacillus subtilis and B. amyloliquefaciens: a review. Appl Microbiol Biotechnol 36(1):1–4Google Scholar
  14. de Gonzalo G, Colpa DI, Habib MHM, Fraaije MW (2016) Bacterial enzymes involved in lignin degradation. J Biotechnol 236:110–119. Google Scholar
  15. DeAngelis KM, Sharma D, Varney R, Simmons B, Isern NG et al (2013) Evidence supporting dissimilatory and assimilatory lignin degradation in Enterobacter lignolyticus SCF1. Front Microbiol 4:280. Google Scholar
  16. Fang X, Li Q, Lin Y et al (2018) Screening of a microbial consortium for selective degradation of lignin from tree trimmings. Bioresour Technol 254:247–255. Google Scholar
  17. Fisher AB, Fong SS (2014) Lignin biodegradation and industrial implications. AIMS Bioeng 1:92–112. Google Scholar
  18. Garrido-Sanz D, Manzano J, Martín M, Redondo-Nieto M, Rivilla R (2018) Metagenomic analysis of a biphenyl-degrading soil bacterial consortium reveals the metabolic roles of specific populations. Front Microbiol 9:232. Google Scholar
  19. Guevara G, Lopez MC, Alonso S, Perera J, Navarro-Llorens JM (2019) New insights into the genome of Rhodococcus ruber strain Chol-4. BMC Genomics 201920:332. Google Scholar
  20. Gurujeyalakshmi G, Mahadevan A (1987) Degradation of guaicol glyceryl ether (GGE) by B. subtilis. Appl Microbiol Biotechnol 26:289–293Google Scholar
  21. Guzik U, Hupert-Kocurek K, Sitnik M, Wojcieszyńska D (2014) Protocatechuate 3, 4- dioxygenase: a wide substrate specificity enzyme isolated from Stenotrophomonas maltophilia KB2 as a useful tool in aromatic acid biodegradation. J Mol Microbiol Biotechnol 24(3):150–160Google Scholar
  22. Harwood CS, Nicholas NN, Kim MK, Ditty JL, Parales RE (1994) Identification of the pcaRKF gene cluster from Pseudomonas putida: involvement in chemotaxis, biodegradation and transport of 4- hydroxybenzoate. J Bacteriol 176:6479–6488Google Scholar
  23. Henson WR, Campbell T, DeLorenzo DM et al (2018) Multi-omic elucidation of aromatic catabolism in adaptively evolved Rhodococcus opacus. Metab Eng 49:69–83. Google Scholar
  24. Hirose J, Tsuda N, Miyatake M et al (2018) Draft genome sequence of Pseudomonas sp. strain LLC-1 (NBRC 111237), capable of metabolizing lignin-derived low-molecular- weight compounds. Genome Announc 6:e00308-18. Google Scholar
  25. Hogancamp TN, Raushel FM (2018) Functional annotation of LigU as a 1,3-allylic isomerase during the degradation of lignin in the protocatechuate 4,5-cleavage pathway from the soil bacterium Sphingobium sp. SYK-6. Biochemistry 57:2837–2845. Google Scholar
  26. Jeżewska-Frąckowiak J, Seroczyńska K, Banaszczyk J, Jedrzejczak G, Żylicz-Stachula A, Skowron PM (2018) The promises and risks of probiotic Bacillus species. Acta Biochim Pol 65(4):509–519. Google Scholar
  27. Jutkina J, Heinaru E, Vedler E, Juhanson J, Heinaru A (2011) Occurrence of plasmids in the aromatic degrading bacterioplankton of the baltic sea. Genes 2(4):853–868. Google Scholar
  28. Kuang F, Li Y, He L et al (2018) Cometabolism degradation of lignin in sequencing batch biofilm reactors. Env Eng Res 23:294–300. Google Scholar
  29. Martínez-Valenzuela Marcela et al (2018) Expression of the sRNAs CrcZ and CrcY modulate the strength of carbon catabolite repression under diazotrophic or non-diazotrophic growing conditions in Azotobacter vinelandii. PloS One 13(12):e0208975. Google Scholar
  30. Mei Q, Shen X, Liu H, Han B (2019) Selectively transform lignin into value-added chemicals. Chinese Chem Lett 30:15–24. Google Scholar
  31. Morales G, Linares JF, Beloso A, Albar JP, Martínez JL, Rojo F (2004) The Pseudomonas putida Crc global regulator controls the expression of genes from several chromosomal catabolic pathways for aromatic compounds. J Bacteriol 186(5):1337–1344. Google Scholar
  32. Ponnusamy VK, Nguyen DD, Dharmaraja J et al (2019) A review on lignin structure, pretreatments, fermentation reactions and biorefinery potential. Bioresour Technol 271:462–472Google Scholar
  33. Ravi K, García-Hidalgo J, Gorwa-Grauslund MF, Lidén G (2017) Conversion of lignin model compounds by Pseudomonas putida KT2440 and isolates from compost. Appl Microbiol and Biotechnol 101(12):5059–5070. Google Scholar
  34. Sambrook J, Russell DW (2001) Molecular cloning: a laboratory manual, 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring HarborGoogle Scholar
  35. Shanmugam S, Sun C, Chen Z, Wu YR (2019) Enhanced bioconversion of hemicellulosic biomass by microbial consortium for biobutanol production with bioaugmentation strategy. Bioresour Technol 279:149–155. Google Scholar
  36. Stanier RY, Ingraham JL (1954) Protocatechuic acid oxidase. J Biol Chem 210:799–808Google Scholar
  37. Sun Z, Fridrich B, de Santi A, Elangovan S, Barta K (2018) Bright side of lignin depolymerization: toward new platform chemicals. Chem Rev 118(2):614–678. Google Scholar
  38. Suvorova IA, Gelfand MS (2019) Comparative genomic analysis of the regulation of aromatic metabolism in betaproteobacteria. Front Microbiol 10:642. Google Scholar
  39. Tian M, Du D, Zhou W, Zeng X, Cheng G (2017) Phenol degradation and genotypic analysis of dioxygenase genes in bacteria isolated from sediments. Braz J Microbiol 48(2):305–313. of the Brazilian Society for Microbiology) Google Scholar
  40. Vieira G, Magrini MJ, Bonugli-Santos RC, Rodrigues M, Sette LD (2018) Polycyclic aromatic hydrocarbons degradation by marine-derived basidiomycetes: optimization of the degradation process. Braz J Microbiol 49(4):749–756. of the Brazilian Society for Microbiology) Google Scholar
  41. Wang Y, Li J, Liu A (2017) Oxygen activation by mononuclear nonheme iron dioxygenases involved in the degradation of aromatics. J Biol Inorg Chem JBIC 22(2–3):395–405. Google Scholar
  42. Wang X, Xia K, Yang X, Tang C (2019) Growth strategy of microbes on mixed carbon sources. Nat Comm 10(1):1279. Google Scholar
  43. Woo HL, Hazen TC (2018) Enrichment of bacteria from eastern mediterranean sea involved in lignin degradation via the Phenylacetyl-CoA pathway. Front Microbiol 9:922. Google Scholar
  44. Wu W, Dutta T, Varman AM, Eudes A, Manalansan B, Loqué D, Singh S (2017) Lignin valorization: two hybrid biochemical routes for the conversion of polymeric lignin into value-added chemicals. Sci Rep 7(1):8420. Google Scholar
  45. Xin Y, Mu Y, Kong J, Guo T (2019) Targeted and repetitive chromosomal integration enables high-level heterologous gene expression in Lactobacillus casei. Appl Environ Microbiol 85(9):e00033-19. Google Scholar
  46. Xu R, Zhang K, Liu P et al (2018) Lignin depolymerization and utilization by bacteria. Bioresour Technol 269:557–566Google Scholar
  47. Xu Z, Lei P, Zhai R, Wen Z, Jin M (2019) Recent advances in lignin valorization with bacterial cultures: microorganisms, metabolic pathways, and bio-products. Biotechnol for biofuels. 12:32. Google Scholar
  48. Yan S, Wu G (2017) Reorganization of gene network for degradation of polycyclic aromatic hydrocarbons (PAHs) in Pseudomonas aeruginosa PAO1 under several conditions. J Appl Gen 58(4):545–563. Google Scholar
  49. Zhang LS, Fang Y, Zhou Y, Ye BC (2017) Improvement of the stabilization and activity of protocatechuate 3,4-dioxygenase isolated from Rhizobium sp. LMB-1 and immobilized on Fe3O4 nanoparticles. Appl Biochem Biotechnol 183(3):1035–1048. Google Scholar
  50. Zhang R, Li C, Wang J, Yan Y (2019) Microbial ligninolysis: toward a bottom-up approach for lignin upgrading. Biochem 58:1501–1510. Google Scholar
  51. Zhu D, Zhang P, Xie C, Zhang W, Sun J, Qian W, Yang B (2017) Biodegradation of alkaline lignin by Bacillus ligniniphilus L1. Biotechnol for Biofuels 10:44. Google Scholar
  52. Zhu D, Si H, Zhang P, Geng A, Zhang W et al (2018) Genomics and biochemistry investigation on the metabolic pathway of milled wood and alkali lignin-derived aromatic metabolites of Comamonas serinivorans SP-35. Biotechnol Biofuels 11:338. Google Scholar
  53. Zhuang L, Tang Z, Ma J, Yu Z, Wang Y, Tang J (2019) Enhanced anaerobic biodegradation of benzoate under sulfate-reducing conditions with conductive iron-oxides in sediment of pearl river estuary. Front Microbiol 10:374. Google Scholar
  54. Żur J, Piński A, Marchlewicz A, Hupert-Kocurek K, Wojcieszyńska D, Guzik U (2018) Organic micropollutants paracetamol and ibuprofen-toxicity, biodegradation, and genetic background of their utilization by bacteria. Env Sci Poll Res Int 25(22):21498–21524. Google Scholar

Copyright information

© Society for Environmental Sustainability 2019

Authors and Affiliations

  1. 1.Centre for Advanced Studies in BotanyUniversity of MadrasChennaiIndia
  2. 2.Division of Arts and SciencesUniversity of New MexicoGallupUSA
  3. 3.College of Science, Engineering and TechnologyGrand Canyon UniversityArizonaUSA

Personalised recommendations