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Research Article
17 September 2020

The Hydroxyquinol Degradation Pathway in Rhodococcus jostii RHA1 and Agrobacterium Species Is an Alternative Pathway for Degradation of Protocatechuic Acid and Lignin Fragments


Deletion of the pcaHG genes, encoding protocatechuate 3,4-dioxygenase in Rhodococcus jostii RHA1, gives a gene deletion strain still able to grow on protocatechuic acid as the sole carbon source, indicating a second degradation pathway for protocatechuic acid. Metabolite analysis of wild-type R. jostii RHA1 grown on medium containing vanillin or protocatechuic acid indicated the formation of hydroxyquinol (benzene-1,2,4-triol) as a downstream product. Gene cluster ro01857-ro01860 in Rhodococcus jostii RHA1 contains genes encoding hydroxyquinol 1,2-dioxygenase and maleylacetate reductase for degradation of hydroxyquinol but also putative mono-oxygenase (ro01860) and putative decarboxylase (ro01859) genes, and a similar gene cluster is found in the genome of lignin-degrading Agrobacterium species. Recombinant R. jostii mono-oxygenase and decarboxylase enzymes in combination were found to convert protocatechuic acid to hydroxyquinol. Hence, an alternative pathway for degradation of protocatechuic acid via oxidative decarboxylation to hydroxyquinol is proposed.
IMPORTANCE There is a well-established paradigm for degradation of protocatechuic acid via the β-ketoadipate pathway in a range of soil bacteria. In this study, we have found the existence of a second pathway for degradation of protocatechuic acid in Rhodococcus jostii RHA1, via hydroxyquinol (benzene-1,2,4-triol), which establishes a metabolic link between protocatechuic acid and hydroxyquinol. The presence of this pathway in a lignin-degrading Agrobacterium sp. strain suggests the involvement of the hydroxyquinol pathway in the metabolism of degraded lignin fragments.


Microbial metabolism of protocatechuic acid (3,4-dihydroxybenzoic acid) via the β-ketoadipate pathway is known to mediate the degradation of many substituted benzoic acids in a range of soil bacteria able to degrade aromatic compounds (1). The β-ketoadipate pathway has also recently emerged as an important central pathway for degradation of lignin fragments by lignin-degrading bacteria, such as Rhodococcus jostii RHA1 and Pseudomonas putida KT2440 (2), and has been exploited to generate bioproducts from lignin degradation (35).
In the course of metabolic engineering studies to generate pyridine-dicarboxylic acid products from lignin degradation via protocatechuic acid (5), a deletion of the pcaHG genes encoding protocatechuate 3,4-dioxygenase in Rhodococcus jostii RHA1 was made, which revealed, to our surprise, that this gene deletion mutant still was able to grow on protocatechuic acid as the sole carbon source; hence, there is a second pathway for degradation of protocatechuic acid.
Examination of the genome of Rhodococcus jostii RHA1 revealed that there is a gene cluster for conversion of hydroxyquinol (benzene-1,2,4-triol) via intradiol oxidative cleavage to maleylacetate and reduction to β-ketoadipate, encoded by genes ro01857-ro01860 (Fig. 1). The hydroxyquinol catabolic pathway has been observed in Burkholderia cepacia AC1100 (6), Sphingomonas wittichi RW1 (7), and Rhodococcus sp. strain PN1 (8). Kasai et al. have implicated R. jostii RHA1 genes ro01857-ro01861 in γ-resorcylate catabolism via decarboxylation of gamma-resorcylate (2,6-dihydroxybenzoic acid) and hydroxylation of resorcinol to form hydroxyquinol (9). Transcription of genes ro01857-ro01860 was shown to be regulated by gene regulator TsdR, to which gamma-resorcylate binds as an effector (9). The oxidative decarboxylation of vanillic acid to 2-methoxyhydroquinone has been reported in extracts of Sporotrichum pulverulentum (10); hence, the oxidative decarboxylation of protocatechuic acid to form hydroxyquinol has some biochemical precedent. Here, we present evidence that conversion of protocatechuic acid to hydroxyquinol occurs in R. jostii RHA1 and is mediated by flavin-dependent mono-oxygenase (ro01860) and decarboxylase (ro01859) enzymes.
FIG 1 Gene cluster (A) and hypothesis (B) for the hydroxyquinol pathway in Rhodococcus jostii RHA1.


Growth phenotypes of pcaHG gene knockout strain of Rhodococcus jostii RHA1.

The pcaHG genes encoding protocatechuate 3,4-dioxygenase, which catalyzes the first step of the β-ketoadipate pathway, were deleted in Rhodococcus jostii RHA1, using the pk18mobsacB plasmid (see Fig. S1 in the supplemental material), containing the sacB gene as a counterselectable marker (E. M. Spence, L. Calvo-Bado, P. Mines, T. D. H. Bugg, submitted for publication) (11). The growth phenotypes of these mutant strains were investigated, using a range of different carbon sources in M9 minimal medium, as shown in Table 1.
TABLE 1 Growth phenotypes of wild-type R. jostii RHA1 and ΔpcaHG gene deletion strainsa
R. jostii RHA1 constructM9/vanillic acidM9/vanillinM9/4-hydroxy-benzoic acidM9/ferulic acidM9/protocatechuic acid
Wild type++++++++++++++
Strains were grown with 0.1% carbon source in liquid M9 minimal medium. +++, strong growth (OD600, >0.6) after 48 h; ++, growth (OD600, 0.3 to 0.6) after 48 h; +, weak growth (OD600, 0.2 to 0.3) after 48 h; −, no growth. The OD600 was 0.15 to 0.2 at the start of culture.
As shown in Table 1, the ΔpcaHG strain was unable to grow on 0.1% vanillic acid or 4-hydroxybenzoic acid as the carbon source, consistent with their metabolism via the β-ketoadipate pathway, but it was able to grow on solid or liquid M9 medium containing 0.1 to 0.75% (wt/vol) protocatechuic acid as a carbon source (Fig. S2 and S3), implying that another pathway could be used by R. jostii RHA1 to metabolize protocatechuic acid. The lack of growth on 0.1% vanillic acid, which can be converted to protocatechuic acid, suggests that this alternative pathway is only induced at higher concentrations of protocatechuic acid. Growth on 0.1% protocatechuic acid was accompanied by a dark brown coloration, either on solid or liquid medium, which was not observed when the wild-type strain was grown on M9 with protocatechuic acid (Fig. S3).

Metabolite analysis in Rhodococcus jostii RHA1.

Wild-type R. jostii RHA1 was grown in either Luria-Bertani broth or M9 minimal medium containing 0.1% (wt/vol) vanillin, vanillic acid, or protocatechuic acid, and after acidification, metabolites formed were extracted into ethyl acetate. The metabolites were then analyzed by C18 reverse-phase high-performance liquid chromatography (HPLC) and by gas chromatography-mass spectrometry (GC-MS), and the observed peaks were compared with authentic standards.
The results are shown in Table 2. Consistent with the known vanillic acid degradation pathway (12), vanillin was converted into vanillic acid, and both vanillin and vanillic acid were converted into protocatechuic acid. Vanillin and protocatechuic acid were also converted into a new peak at a retention time of 11 min, which matched an authentic sample of hydroxyquinol, consistent with an oxidative decarboxylation of protocatechuic acid to hydroxyquinol (Fig. S4). Guaiacol was detected as a metabolite of vanillic acid by GC-MS, both in M9 minimal medium and by treatment of vanillic acid with cell extract of R. jostii RHA1 in the absence of cofactors, consistent with a decarboxylation of vanillic acid to guaiacol (Fig. S5). When cell extract was used in the presence of 0.1 mM NADH, catechol was also observed (Fig. S5), but catechol was not observed as a metabolite from protocatechuic acid. Unexpectedly, the formation of vanillic acid as a metabolite was also observed by growth of R. jostii on M9 minimal medium containing 0.1% protocatechuic acid (Fig. S6), implying that methylation of protocatechuic acid can occur.
TABLE 2 Metabolites detected by C18 reverse-phase HPLC or GC-MS after growth of R. jostii RHA1 or treatment with R. jostii RHA1 cell extracta
SubstrateMediumMetabolites observed
Vanillic acidProtocatechuic acidHydroxyquinolGuaiacolCatechol
Vanillic acidLB ++   
 M9   + (GC) 
 Extractb   + (GC) 
 Extract + NADHc ++  +
Protocatechuic acidM9+ +  
R. jostii RHA1 was grown in either Luria-Bertani broth (LB) or M9 minimal medium containing 0.1% (wt/vol) carbon source. +, metabolite observed; ++, strong formation of metabolite; GC, metabolite detected by GC-MS.
Cell extract contained 100 μg protein in 50 mM Tris buffer, pH 7.5 (5 ml).
With addition of 0.1 mM NADH. Retention times and gradients are described in Materials and Methods.
Growth of the ΔpcaHG Rhodococcus jostii gene deletion mutant on M9 minimal medium containing protocatechuic acid, as noted above, generated a dark brown coloration. Analysis of supernatant from this incubation by liquid chromatography-mass spectrometry (LC-MS) at extracted ion m/z 127, corresponding to benzene-1,2,4-triol, gave a peak that matched the retention time of a commercial sample of hydroxyquinol (Fig. 2), verifying the conversion of protocatechuic acid into hydroxyquinol.
FIG 2 LC-MS analysis for hydroxyquinol product (extracted ion m/z 127). (A) Analysis of incubation of ΔpcaHG R. jostii deletion mutant with protocatechuic acid. (B) Analysis of commercial sample of hydroxyquinol (benzene-1,2,4-triol).

Investigation of hydroxyquinol gene cluster in Rhodococcus jostii and Agrobacterium species.

A gene cluster closely related to the ro01857-ro01860 gene cluster of R. jostii RHA1 was found in the genome sequence of a lignin-degrading Agrobacterium sp. strain identified from municipal waste soil (13) (Fig. 3). The Agrobacterium sp. gene cluster (genes agro_00119-agro_00125) also contains a putative flavin reductase gene, agro_00119, adjacent to flavin mono-oxygenase agro_00120, and an aromatic acid transporter gene (agro_00125). Gene agro_00121 is annotated as a putative decarboxylase (42.7% identity to DBD23_ASPOR Aspergillus oryzae 2,3-dihydroxybenzoate decarboxylase) within the PF04909.11 amidohydrolase superfamily. The Agrobacterium sp. genome also contained a gene cluster encoding the β-ketoadipate pathway (genes agro_04310-agro_04320), a DyP-type peroxidase (gene agro_00063), and a putative β-etherase LigE (gene agro_01577), consistent with activity for lignin degradation (Table S1). Related hydroxyquinol pathway gene clusters are also found on the genomes of Agrobacterium tumefaciens F2 (genes Agau_C100398 to Agau_C100407), Agrobacterium sp. strain H13-3 (genes AgroH133_08581 to AgroH133_08584), and Rhizobium species such as Rhizobium sp. strain Kim5 (Kim5_PC00119 to Kim5_PC00122).
FIG 3 Comparison of gene clusters for hydroxyquinol utilization in Rhodococcus jostii RHA1 and Agrobacterium species.
The putative mono-oxygenase and decarboxylase genes from R. jostii RHA1 and Agrobacterium species were cloned into pET-S expression vector and expressed in E. coli BL21(DE3) as N-His6 fusion proteins. The recombinant mono-oxygenase enzymes Ro01860 and Ag00120 both expressed strongly and, after purification on a nickel-nitrilotriacetic acid column, gave the expected 60-kDa protein bands by SDS-PAGE (Fig. S7). The putative decarboxylase enzymes Ro01859 and Ag00121 both expressed weakly, giving weaker bands by SDS-PAGE corresponding to the expected 44.7- and 50.1-kDa proteins, respectively (Fig. S7). Western blots were carried out to confirm that the desired decarboxylase protein had been expressed, which confirmed that a protein of the correct size had been expressed at low levels (Fig. S8).
The purified recombinant enzymes were incubated with protocatechuic acid in the presence of 10 μM FAD and 200 μM NADPH to test for conversion to hydroxyquinol, and the reactions were monitored by C18 reverse-phase HPLC. Incubation of 1 mM protocatechuic acid with 100 μg purified Ro01860 and Ro01859 enzymes gave no new product peak after 1 h; however, addition of 100 μg purified mono-oxygenase Ro01860 and cell extract containing recombinant decarboxylase Ro01859 (100 μg protein) to 1 mM protocatechuic acid was found to generate a new peak at 10.7 min corresponding to hydroxyquinol (Fig. 4A), but this peak was not formed by cell extract lacking Ro01859. Using 100 μg recombinant mono-oxygenase Ro01860 or Ag00120 alone, some consumption of protocatechuic acid was observed (35% and 77%, respectively), but hydroxyquinol was not formed, although a new peak at a retention time of 12.8 min was generated that did not coelute with hydroxyquinol (Fig. S9). Using purified recombinant decarboxylase enzyme Ro01859 or Ag00121 alone, no reaction with protocatechuic acid was observed. We hypothesize that the purification of decarboxylase Ro01859 renders the enzyme inactive, perhaps due to loss of an essential cofactor, but that overexpressed decarboxylase Ro01859 and purified mono-oxygenase Ro01860 together are able to catalyze the conversion of protocatechuic acid to hydroxyquinol.
FIG 4 Reverse-phase HPLC analysis of incubations of protocatechuic acid (PCA) (A), catechol (B), hydroquinone (1,4-HQ) (C), and gentisic acid (D) with purified mono-oxygenase Ro01860 and overexpressed decarboxylase Ro01859. Biotransformations contained 100 μg of each protein in PBS buffer (1.0 ml), containing 1 mM substrate, 10 μM FAD, and 200 μM NADPH, and were incubated for 60 min at 25°C. Hydroxyquinol product is marked with a blue box, and other substrates are labeled. Gentisic acid substrate elutes at >15 min.
Several other potential substrates for the mono-oxygenase and decarboxylase enzymes were also tested. Formation of hydroxyquinol was also observed upon incubation of Ro01859/Ro01860 with catechol (Fig. 4B), 1,4-hydroquinone (Fig. 4C), or gentisic acid (2,5-dihydroxybenzoic acid) (Fig. 4D), indicating that hydroxylation/decarboxylation of several compounds can be carried out by this pair of enzymes in vitro, as discussed below. The formation of a small amount of hydroxyquinol was also observed when catechol was treated with mono-oxygenase Ro01860 alone; approximately 2% of the product formed with Ro01859/Ro01860. No reaction of Ro01860 alone was observed with 1,4-hydroquinone, 2,3-dihydroxybenzoic acid, 2,4-dihydroxybenzoic acid, or gentisic acid (2,5-dihydroxybenzoic acid).


The hydroxyquinol degradation pathway was previously reported in Burkholderia cepacia AC1100 (6), Sphingomonas wittichi RW1 (7), and Rhodococcus sp. strain PN1 (8) as an aromatic catabolic pathway, and a hydroxyquinol degradation gene cluster has been observed in Cupriavidus necator JMP134 (14). This pathway has been implicated in degradation of 4-nitrophenol in Arthrobacter chlorophenolicus A6 (15) and in γ-resorcylate catabolism in R. jostii RHA1 (9). Following our initial observation that a ΔpcaHG gene deletion mutant of R. jostii RHA1 was able to grow on minimal medium containing protocatechuic acid, we hypothesized that protocatechuic acid could be converted via flavin-dependent mono-oxygenase (ro01860) and a decarboxylase (ro01859) into hydroxyquinol. We have shown by metabolite analysis that protocatechuic acid can be converted into hydroxyquinol by wild-type R. jostii RHA1 cells, by the ΔpcaHG gene deletion strain, and in vitro using recombinant Ro01860/Ro01859 enzymes. Therefore, these observations establish a metabolic link between protocatechuic acid and hydroxyquinol in R. jostii, which can be used as an alternative pathway for metabolism of protocatechuic acid in this bacterium.
The catalytic mechanism of these two enzymes is likely to proceed as shown in Fig. 5. Phenol hydroxylation ortho or para to a phenolic hydroxyl group is well precedented in the flavin mono-oxygenase family, via a flavin hydroperoxide reaction intermediate (16). Several soil bacteria that can metabolize chlorinated phenols (1719) or 4-nitrophenols (20, 21) are known to express flavin-dependent mono-oxygenase enzymes that catalyze hydroxylation para to a phenolic hydroxyl group, followed by loss of either a 4-chloro or 4-nitro substituent. In this case, hydroxylation in the 1-position would be catalyzed by mono-oxygenase Ro01860, to give a diffusible intermediate, which is then decarboxylated by Ro01859, using the carbonyl group at C-4 as an electron sink to aid decarboxylation (Fig. 5). Such a mechanism would explain why a combination of Ro01860 and Ro01859 is needed to effect this oxidative decarboxylation and why no activity was observed using decarboxylase Ro01859 alone. Using mono-oxygenase Ro01860 alone, the intermediate would be formed, leading to consumption of protocatechuic acid, as observed, but the intermediate is likely to be chemically unstable and could not be characterized.
FIG 5 Proposed catalytic mechanism for mono-oxygenase Ro01860 and decarboxylase Ro01859.
Formation of hydroxyquinol in vitro was also observed from catechol, 1,4-hydroquinone, or 2,5-dihydroxybenzoic acid (gentisic acid). Only a small amount of hydroxylation of catechol by mono-oxygenase Ro01860 alone was observed, implying that the two enzymes are more active in combination, perhaps forming a complex. Kasai et al. have shown that the R. jostii ro01859 and ro01860 gene products are able to convert 2,6-dihydroxybenzoic acid to hydroxyquinol: Ro01859 decarboxylates γ-resorcylate to resorcinol, and Ro01860 converts resorcinol to hydroxyquinol (9). Hence, it appears that these enzymes are able to catalyze several hydroxylation/decarboxylation reactions. Kasai et al. reported that gene regulator TsdR negatively regulates the expression of these genes and binds to γ-resorcylic acid (2,6-dihydroxybenzoic acid). The observation that the ΔpcaHG deletion mutant can grow on M9-0.1% protocatechuic acid but not on 0.1% vanillic acid or 0.1% 4-hydroxybenzoic acid suggests that the conversion of protocatechuic acid to hydroxyquinol takes place only at higher concentrations of protocatechuic acid; hence, protocatechuic acid may bind more weakly to gene regulator TsdR. Kasai et al. reported activity for resorcinol hydroxylation by Ro01860 using NADH as a cofactor (9), whereas we observed activity for protocatechuic acid conversion by Ro01860 and Ro01859 using NADPH as a cofactor (NADH was not tested); therefore, it appears that Ro01860 can accept either NADH or NADPH for different transformations.
Our metabolite analysis also indicates two other transformations that can occur in R. jostii RHA1, as shown in Fig. 6. First, we have observed the conversion of vanillic acid to guaiacol via decarboxylation, followed by demethylation to catechol. Decarboxylation of vanillic acid to guaiacol was previously seen in Bacillus megaterium and Streptomyces (22); a gene cluster responsible for this transformation has been identified in Streptomyces sp. strain D7 (23), and a vanillate decarboxylase enzyme has been purified from Nocardia sp. strain NRRL 5646 (24). However, there is no vanillic acid decarboxylase gene annotated on the R. jostii RHA1 genome, so the gene responsible for this transformation is not known. The decarboxylation of vanillate to guaiacol might explain an earlier observation where an R. jostii vanillate mono-oxygenase (ro04165) gene deletion strain failed to accumulate vanillic acid when grown on medium containing wheat straw lignocellulose, whereas a Δvdh vanillate dehydrogenase gene deletion strain did accumulate vanillin (3). We did not observe any conversion of protocatechuic acid to catechol, and there is no protocatechuic acid decarboxylase gene annotated on the R. jostii RHA1 genome, although such a decarboxylase gene has been identified in Klebsiella pneumoniae (25). The formation of guaiacol is also consistent with the presence of a gene cluster for guaiacol metabolism in R. jostii RHA1 at ro08067/08068, containing a homologue for a P450-dependent guaiacol demethylase enzyme, which has been characterized from Amycolatopsis sp. strain ATCC 39116 (26).
FIG 6 Biochemical transformations of vanillic acid and protocatechuic acid in Rhodococcus jostii RHA1 observed in this work and the transformation reported by Kasai et al. (9).
Second, a surprising observation was the conversion of protocatechuic acid to vanillic acid via methylation of the 3-hydroxy group. There is a methyltransferase gene (ro04166) located in the gene cluster for vanillate mono-oxygenase (ro04165) that may be responsible for this methylation reaction. This reaction is surprising, as it would apparently set up a futile cycle between vanillic acid and protocatechuic acid; therefore, expression of the methyltransferase and demethylase genes would need to be tightly regulated.
We have elsewhere observed 2-methoxyhydroquinone, the methylated form of hydroxyquinol, as a metabolite of lignin oxidation by lignin-degrading enzyme manganese superoxide dismutase from Sphingobacterium sp. strain T2 (27). A pathway for degradation of (+)-pinoresinol, a component of lignin structure, via 2-methoxyhydroquinone has also recently been elucidated in Pseudomonas sp. strain SG-MS2 (28, 29). Hence, it seems likely that hydroxyquinol can be generated from other lignin oxidation reactions involving aryl-Cα oxidative cleavage of the aryl-C3 unit found in lignin. In a recent survey of the genomes of lignin-degrading bacteria (30), the hydroxyquinol gene cluster was only observed in Rhodococcus jostii RHA1 and Agrobacterium species, but lignin-degrading Paenibacillus species and Ochrobactrum species strains also contained mhq genes, thought to be involved in hydroquinone breakdown (31). The pathways branching from vanillic acid and protocatechuic acid shown in Fig. 6 will reduce metabolic flux through the β-ketoadipate pathway; hence, an understanding of these pathways will underpin efforts to engineer R. jostii RHA1 for efficient conversion of lignin to renewable chemicals (35).


Bacterial strains, plasmids, and chemicals.

Rhodococcus jostii RHA1 was used as the ancestral strain (32). The ΔpcaHG R. jostii strain contains a deletion in chromosomal pcaHG genes, as described here. The Agrobacterium species was isolated from municipal waste as described previously (13). For routine growth and maintenance, R. jostii or Agrobacterium sp. cells were cultured in liquid or solid Luria-Bertani broth (LB) medium, or, for R. jostii, on M9 minimal medium supplemented with 0.1% benzoic acid at 30°C and with shaking at 180 rpm, if required. Bacterial growth was measured by absorbance at 595 or 600 nm.
Expression vector pET-S is a derivative of pET151 topo (Invitrogen) in which a Sumo tag was amplified from pOPINS3C ( using primers Pfor, CCCCCCCATATGGCACACCATCACCACC (NdeI site underlined), and Prev, AAAAAAATCGATGGATTTAAATGGGCTAGCGGATCCACCGCTGCTGATCTGTTCG (ClaI, SwaI, NheI, and BamHI sites underlined, respectively), and cloned into NdeI/ClaI sites of pET151 topo. All chemicals were purchased form Sigma-Aldrich unless otherwise stated.

Construction of gene deletion mutants.

The ΔpcaHG gene deletion strain was constructed using the pk18mobsacB plasmid, which uses the sacB gene (confers sucrose sensitivity) as a counterselectable marker (11). PCR was used to amplify two 1-kb regions of chromosomal DNA on either side of the genes to be deleted. The amplified 1-kb upstream and downstream DNA sequences were ligated into pk18mobsac, and the resulting construct (see Fig. S1 in the supplemental material) was confirmed by DNA sequencing and restriction digestion. The recombinant plasmid was taken up into R. jostii by electroporation and recombinant colonies selected by kanamycin resistance. Isolation of the double crossover gene deletion was carried out using sucrose resistance counterselection (11), and the markerless gene deletion was confirmed by internal and external PCR analysis.

Metabolite analysis.

Cultures of R. jostii RHA1 were grown in either Luria-Bertani medium or M9 minimal medium containing 1 mM vanillic acid, vanillin, or protocatechuic acid at 30°C for 24 to 48 h. Aliquots (500 μl) of culture were removed and combined with 500 μl of HPLC-grade methanol-0.1% trifluoroacetic acid. Samples were vortexed and then centrifuged (microcentrifuge) for 15 min. Biotransformations using R. jostii cell extract contained 100 μg protein, 1 mM substrate, with or without addition of 0.1 mM NADH, in 50 mM Tris buffer, pH 7.5, and were incubated at 20°C for 1 to 2 h. The incubation was then acidified to pH 1 (1 M HCl) and the products were extracted into ethyl acetate, solvent was evaporated at reduced pressure, and the sample was redissolved in methanol. HPLC analysis of the supernatant was performed with a Hyperclone 5-μm C18 reverse-phase column (Phenomenex), using an Agilent 1200 series HPLC analyzer. The HPLC solvents were water-0.1% trifluoroacetic acid (solvent A) and methanol-0.1% trifluoroacetic acid (solvent B). The applied gradient was 10 to 30% B over 5 min, 30 to 40% B over 15 min, 40 to 70% B over 10 min, 70 to 100% B over 5 min, and 100 to 10% B over 10 min, at a flow rate of 0.5 ml min−1. UV detection was at 270 nm. Samples for LC-MS analysis were suspended in 1:1 methanol (MeOH)-water and were separated on a Phenomenex Luna C18 column (5 μm, 100 Å, 50 mm, 4.6 mm) on an Agilent 1200 analyzer and Bruker HCT Ultra mass spectrometer (flow rate, 0.5 ml/min) with monitoring at 270 nm. Solvent A was water-0.1% formic acid, and solvent B was MeOH-0.1% formic acid. The gradient was 15% solvent B for 5 min, 15 to 25% B from 5 to 15 min, 25 to 70% B from 15 to 23 min, and 70 to 100% from 23 to 30 min. GC-MS analysis (ion-trap analyzer) was performed using a Varian 4000 gas chromatograph-mass spectrometer with a Varian Factor Four column (length, 30 m; inner diameter, 0.25 mm; thickness, 0.25 μm). Electron impact mass spectra (EI-MS) were recorded at an ionization energy of 70 eV.

Expression and purification of R. jostii and Agrobacterium mono-oxygenase and decarboxylase enzymes.

R. jostii ro01859 (decarboxylase) and ro01860 (mono-oxygenase) genes and Agrobacterium agro_00120 (mono-oxygenase) and agro_00121 (decarboxylase) genes were amplified from genomic DNA using PCR, using the oligonucleotide primers given below, and ligated into the pET-S expression vector, using XbaI and ClaI restriction sites, resulting in the expression of an N-His6 fusion protein. Oligonucleotide primers for PCR were the following: ro01859, forward, AAAAAAGCTAGCCAGGGCAAGATCGCACTGG; reverse, AAAAAAATCGATTCACCGGTCGAGTTTGAACAAC; ro01860, forward, AAAAAAAGCTAGCATGTCTGCCTTCGCACAG; reverse, AAAAAAATCGATAGGTGTTTGTCGGTGGTG; ag00120, forward, AAAAAAAGCTAGCATGAACGATATGAGCCATGCG; reverse, AAAAAAATCGATTCAATATTGGCCCTTGGGTTC; ag00121, forward, AAAAAAAGCTAGCATGCAAGGCAAGGTCGCTC; reverse, AAAAAAATCGATTCAGTTCCCGTCGAGTTTGAAC.
Recombinant plasmids were expressed in Escherichia coli BL21(DE3). For protein purification, constructs were grown in Luria-Bertani broth (500 ml) in the presence of ampicillin (50 μg/ml), and protein expression was induced by addition of isopropyl-β-d-thiogalactopyranoside (IPTG) (250 μM) at an optical density at 600 nm (OD600) of 0.6. Expression of mono-oxygenases Ro01860 and Ag00120 was carried out in the presence of 1 mM riboflavin to achieve reconstitution of the flavin cofactor. Cultures were grown for 16 h at 16°C, and then cells were harvested by centrifugation (4,000 × g, 20 min). Cells were resuspended in 50 mM sodium phosphate buffer, pH 8.0, containing 300 mM NaCl and sonicated in a Constant Systems cell disruptor (20,000 lb/in2), and then cell debris was removed by centrifugation (20,000 × g, 20 min). Cell extract was applied to a His Gravitrap column (GE Healthcare) and washed with 50 mM sodium phosphate buffer, pH 8.0, containing 300 mM NaCl and 20 mM imidazole, and then protein was eluted with 50 mM sodium phosphate buffer, pH 8.0, containing 300 mM NaCl and 250 mM imidazole. Purified protein was then desalted using a PD-10 desalting column (GE Healthcare).
SDS-PAGE was carried out using a Thermo Fisher Scientific Invitrogen Mini Gell tank, using a 10% Bis-Tris protein gel, according to the manufacturer’s instructions. Thermo Fisher Scientific PageRuler prestained protein ladder was used for molecular weight markers. Western blotting was carried out by using a Thermo Fisher Scientific Invitrogen Novex iBlot 2 nitrocellulose transfer stack and a Thermo Fisher Scientific iBlot 2 gel transfer device, according to the manufacturer’s instructions, using Thermo Fisher Scientific 6×-His tag monoclonal antibody.

Biotransformation using mono-oxygenase Ro01860 and decarboxylase Ro01859.

Biotransformations using recombinant Ro01860 and Ro01859 were carried out using 100 μg of each protein in PBS buffer (1.0 ml) containing 1 mM substrate in the presence of 10 μM FAD and 200 μM NADPH and incubated for 60 min at 25°C. Aliquots (100 μl) were removed and combined with 100 μl of HPLC-grade methanol-0.1% trifluoroacetic acid. Samples were vortexed and then centrifuged (microcentrifuge) for 15 min and analyzed by HPLC using a Zorbax Eclipse plus (Agilent) C18 reverse-phase column, using the gradient and conditions described above. Control incubations lacking enzyme were also carried out, which showed no substrate conversion.

Data availability.

Accession numbers used in this study were ABG93670 (R. jostii ro01859 decarboxylase), ABG93671 (R. jostii ro01860 flavin-dependent mono-oxygenase), WP_149145897.1 (Agrobacterium sp. agro_00120 flavin-dependent mono-oxygenase), and WP_149145898.1 (Agrobacterium sp. agro_00121 decarboxylase).


This research was supported by research grants from the IB Catalyst CRD Industrial Research Program (reference TS/M008282/1), BBSRC (reference BB/P01738X/1), and FAPESP (research grant 2015/50590-4) and by the ERASMUS exchange program (to L.D.).

Supplemental Material

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Harwood CS, Parales RE. 1996. The β-ketoadipate pathway and the biology of self-identity. Annu Rev Microbiol 50:553–590.
Bugg TDH, Ahmad M, Hardiman EM, Rahmanpour R. 2011. Pathways for degradation of lignin in bacteria and fungi. Nat Prod Rep 28:1883–1896.
Sainsbury PD, Hardiman EM, Ahmad M, Otani H, Seghezzi N, Eltis LD, Bugg TDH. 2013. Breaking down lignin to high-value chemicals: the conversion of lignocellulose to vanillin in a gene deletion mutant of Rhodococcus jostii RHA1. ACS Chem Biol 8:2151–2156.
Vardon DR, Franden MA, Johnson CW, Karp EM, Guarnieri MT, Linger JG, Salm MJ, Strathmann TJ, Beckham GT. 2015. Adipic acid production from lignin. Energy Environ Sci 8:617–628.
Mycroft Z, Gomis M, Mines P, Law P, Bugg TDH. 2015. Biocatalytic conversion of lignin to aromatic dicarboxylic acids in Rhodococcus jostii RHA1 by re-routing aromatic degradation pathways. Green Chem 17:4974–4979.
Daubaras DL, Saido K, Chakrabarty AM. 1996. Purification of hydroxyquinol 1,2-dioxygenase and maleylacetate reductase: the lower pathway of 2,4,5-trichlorophenoxyacetic acid metabolism by Burkholderia cepacia AC1100. Appl Environ Microbiol 62:4276–4279.
Armengaud J, Timmis KN, Wittich RM. 1999. A functional 4-hydroxy-salicylate/hydroxyquinol degradative pathway gene cluster is linked to the initial dibenzo-p-dioxin pathway genes in Sphingomonas sp. strain RW1. J Bacteriol 181:3452–3461.
Yamamoto K, Nishimura M, Kato D, Takeo M, Negoro S. 2011. Identification and characterization of another 4-nitrophenol degradation gene cluster, nps, in Rhodococcus sp. strain PN1. J Biosci Bioeng 111:687–694.
Kasai D, Araki N, Motoi K, Yoshikawa S, Iino T, Imai S, Masai E, Fukuda M. 2015. γ-Resorcylate catabolic-pathway genes in the soil actinomycete Rhodococcus jostii RHA1. Appl Environ Microbiol 81:7656–7665.
Buswell JA, Ander P, Pettersson B, Eriksson K. 1979. Oxidative decarboxylation of vanillic acid by Sporotrichum pulverulentum. FEBS Lett 103:98–101.
van der Geize R, Hessels GI, van Gerwen R, van der Meijden P, Dijkhuizen L. 2001. Unmarked gene deletion mutagenesis of kstD, encoding 3-ketosteroid-D1-dehydrogenase, in Rhodococcus erythropolis SQ1 using sacB as a counter-selectable marker. FEMS Microbiol Lett 205:197–202.
Chen HP, Chow M, Liu CC, Lau A, Liu J, Eltis LD. 2012. Vanillin catabolism in Rhodococcus jostii RHA1. Appl Environ Microbiol 78:586–588.
Rashid GMM, Duran-Pena MJ, Rahmanpour R, Sapsford D, Bugg TDH. 2017. Delignification and enhanced gas release from soil containing lignocellulose by treatment with bacterial lignin degraders. J Appl Microbiol 123:159–171.
Pérez-Pantoja D, De la Iglesia R, Pieper DH, González B. 2008. Metabolic reconstruction of aromatic compound degradation from the genome of the amazing pollutant-degrading bacterium Cupriavidus necator JMP134. FEMS Microbiol Rev 32:736–794.
Nordin K, Unell M, Jansson JK. 2005. Novel 4-chlorophenol degradation gene cluster and degradation route via hydroxyquinol in Arthrobacter chlorophenolicus A6. Appl Environ Microbiol 71:6538–6544.
Massey V. 1994. Activation of molecular oxygen by flavins and flavoproteins. J Biol Chem 269:22459–22462.
Beadle CA, Smith ARW. 1982. The purification and properties of 2,4-dichlorophenol hydroxylase from a strain of Acinetobacter species. Eur J Biochem 123:323–332.
Xun L, Orser CS. 1991. Purification and properties of pentachlorophenol hydroxylase, a flavoprotein from Flavobacterium sp. strain ATCC 39723. J Bacteriol 173:4447–4453.
Wieser M, Wagner B, Eberspächer J, Lingens F. 1997. Purification and characterization of 2,4,6-trichlorophenol-4-monooxygenase, a dehalogenating enzyme from Azotobacter sp. strain GP1. J Bacteriol 179:202–208.
Leungsakul T, Johnson GR, Wood TK. 2006. Protein engineering of the 4-methyl-5-nitrocatechol monooxygenase from Burkholderia sp. strain DNT for enhanced degradation of nitroaromatics. Appl Environ Microbiol 72:3933–3939.
Min J, Lu Y, Hu X, Zhou N-Y. 2016. Biochemical characterization of 3-methyl-4-nitrophenol degradation in Burkholderia sp. strain SJ98. Front Microbiol 7:791.
Crawford RL, Olson PP. 1978. Microbial catabolism of vanillate: decarboxylation to guaiacol. Appl Environ Microbiol 36:539–543.
Chow KT, Pope MK, Davies J. 1999. Characterization of a vanillic acid non-oxidative decarboxylation gene cluster from Streptomyces sp. D7. Microbiology 145:2393–2403.
Dhar A, Lee K-S, Dhar K, Rosazza JPN. 2007. Nocardia sp. vanillic acid decarboxylase. Enzyme Microb Technol 41:271–277.
Sonoki T, Morooka M, Sakamoto K, Otsuka Y, Nakamura M, Jellison J, Goodell B. 2014. Enhancement of protocatechuate decarboxylase activity for the effective production of muconate from lignin-related aromatic compounds. J Biotechnol 192:71–77.
Mallinson SJB, Machinova MM, Silveira RL, Garcia-Borras M, Gallup N, Johnson CW, Allen MD, Skaf MS, Crowley MF, Neidle EL, Houk KN, Beckham GT, DuBois JL, McGeehan JE. 2018. A promiscuous cytochrome P450 aromatic O-demethylase for lignin bioconversion. Nat Commun 9:2487.
Rashid GMM, Taylor CR, Liu Y, Zhang X, Rea D, Fülöp V, Bugg TDH. 2015. Identification of manganese superoxide dismutase from Sphingobacterium sp. T2 as a novel bacterial enzyme for lignin oxidation. ACS Chem Biol 10:2286–2294.
Shettigar M, Balotra S, Cahill D, Warden AC, Lacey MJ, Kohler HPE, Rentsch D, Oakeshott JG, Pandey G. 2017. Isolation of the (+)-pinoresinol-mineralizing Pseudomonas sp. strain SG-MS2 and elucidation of its catabolic pathway. Appl Environ Microbiol 84:e02531-17.
Shettigar M, Balotra S, Kasprzak A, Pearce SL, Lacey MJ, Taylor MC, Liu JW, Cahill D, Oakeshott JG, Pandey G. 2020. Oxidative catabolism of (+)-pinoresinol is initiated by an unusual flavocytochrome encoded by translationally coupled genes within a cluster of (+)-pinoresinol-coinduced genes in Pseudomonas sp. strain SG-MS2. Appl Environ Microbiol 86:e00375-20.
Granja-Travez RS, Persinoti GF, Squina FM, Bugg TDH. 2020. Functional genomic analysis of bacterial lignin degraders: diversity in mechanisms of lignin oxidation and metabolism. Appl Microbiol Biotechnol 104:3305–3320.
Tago K, Sato J, Takesa H, Kawagishi H, Hayatsu M. 2005. Characterization of methylhydroquinone-metabolizing oxygenase genes encoded on plasmid in Burkholderia sp. NF100. J Biosci Bioeng 100:517–523.
Seto M, Kimbara K, Shimura M, Hatta T, Fukuda M, Yano K. 1995. A novel transformation of polychlorinated biphenyls by Rhodococcus sp. strain RHA1. Appl Environ Microbiol 61:3353–3358.

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Published In

cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 86Number 1917 September 2020
eLocator: e01561-20
Editor: Rebecca E. Parales, University of California, Davis
PubMed: 32737130


Received: 29 June 2020
Accepted: 27 July 2020
Published online: 17 September 2020


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  1. Agrobacterium sp.
  2. hydroxyquinol pathway
  3. lignin degradation
  4. Rhodococcus jostii



Edward M. Spence
Department of Chemistry, University of Warwick, Coventry, United Kingdom
Heather T. Scott
Department of Chemistry, University of Warwick, Coventry, United Kingdom
Louison Dumond
Department of Chemistry, University of Warwick, Coventry, United Kingdom
Leonides Calvo-Bado
Department of Chemistry, University of Warwick, Coventry, United Kingdom
Sabrina di Monaco
Department of Chemistry, University of Warwick, Coventry, United Kingdom
James J. Williamson
Department of Chemistry, University of Warwick, Coventry, United Kingdom
Gabriela F. Persinoti
Laboratório Nacional de Biorrenováveis (LNBR/CNPEM), Campinas, Brazil
Programa de Processos Tecnológicos e Ambientais, Universidade de Sorocaba, Sorocaba, Brazil
Department of Chemistry, University of Warwick, Coventry, United Kingdom


Rebecca E. Parales
University of California, Davis


Address correspondence to Timothy D. H. Bugg, [email protected].

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