A Multicomponent THF Hydroxylase Initiates Tetrahydrofuran Degradation in Cupriavidus metallidurans ZM02

Hao Ren; Haixia Wang; Yang Wang; Yiyang Chen; Zhenmei Lu

doi:10.1128/aem.01880-21

. 2022 Mar 22;88(6):e01880-21. doi: 10.1128/aem.01880-21

A Multicomponent THF Hydroxylase Initiates Tetrahydrofuran Degradation in Cupriavidus metallidurans ZM02

Hao Ren ^a, Haixia Wang ^a, Yang Wang ^b, Yiyang Chen ^a, Zhenmei Lu ^a,^✉

Editor: Jeremy D Semrau^c

PMCID: PMC8939321 PMID: 35108100

ABSTRACT

Tetrahydrofuran (THF) has been recognized as a water contaminant because of its human carcinogenicity, extensive use, and widespread distribution. Previously reported multicomponent monooxygenases (MOs) involved in THF degradation were highly conserved, and all of them were from Gram-positive bacteria. In this study, a novel THF-degrading gene cluster (dmpKLMNOP) encoding THF hydroxylase was identified on the chromosome of a newly isolated Gram-negative THF-degrading bacterium, Cupriavidus metallidurans ZM02, and functionally characterized. Transcriptome sequencing and RT-qPCR demonstrated that the expression of dmpKLMNOP was upregulated during the growth of strain ZM02 on THF or phenol. The deletion of oxygenase alpha or beta subunit or the reductase component disrupted the degradation of THF but did not affect the utilization of its hydroxylated product 2-hydroxytetrahydrofuran. Cupriavidus pinatubonensis JMP134 heterologously expressing dmpKLMNOP from strain ZM02 could grow on THF, indicating that the THF hydroxylase Dmp_ZM02KLMNOP is responsible for the initial degradation of THF. Furthermore, the THF and phenol oxidation activities of crude enzyme extracts were detected, and the highest THF and phenol catalytic activities were 1.38 ± 0.24 μmol min⁻¹mg⁻¹ and 1.77 ± 0.37 μmol min⁻¹mg⁻¹, respectively, with the addition of NADPH and Fe²⁺. The characterization of THF hydroxylase associated with THF degradation enriches our understanding of THF-degrading gene diversity and provides a novel potential enzyme for the bioremediation of THF-containing pollutants.

IMPORTANCE Multicomponent MOs catalyzing the initial hydroxylation of THF are vital rate-limiting enzymes in the THF degradation pathway. Previous studies of THF degradation gene clusters have focused on Gram-positive bacteria, and the molecular mechanism of THF degradation in Gram-negative bacteria has rarely been reported. In this study, a novel THF hydroxylase encoded by dmpKLMNOP in strain ZM02 was identified to be involved in both THF and phenol degradation. Our findings provide new insights into the THF-degrading gene cluster and enzymes in Gram-negative bacteria.

KEYWORDS: Cupriavidus metallidurans ZM02, THF hydroxylase, tetrahydrofuran, phenol, initial degradation

INTRODUCTION

The oxygen heterocyclic compound tetrahydrofuran (THF) is an important chemical feedstock for polymer synthesis (1), a popular promoter molecule (2), and a polar solvent for applying coatings to various materials (3). THF is widely distributed in industrial effluents produced by the pharmaceutical, pesticide, and chemical industries and has been found to be a contaminant in groundwater (4, 5). Many toxicological studies on rats exposed to THF have demonstrated that THF can enhance tumor formation (6, 7), cause DNA damage (8), and induce nervous system dysfunction (9). Biodegradation is considered a potential remediation strategy for the removal of THF contaminants. To date, more than 20 strains, mainly including Gram-positive bacteria (Rhodococcus sp. [10, 11], Pseudonocardia sp. [12], and Mycobacterium sp. [13]), Gram-negative bacteria (Pseudomonas sp. [14] and Xanthobacter sp. [15]), and fungi (Graphium sp. [16]) have been identified that can aerobically utilize THF as a sole carbon source and energy source.

In the proposed THF degradation pathway, THF is initially oxidized at the carbon adjacent to the oxygen atom to form 2-hydroxytetrahydrofuran (2-OH THF), which involves a monooxygenase-catalyzed reaction (17, 18). 2-OH THF is first oxidized to γ-butyrolactone, which is further oxidized to 4-hydroxybutyrate before being transformed to succinate (11, 16, 19). Finally, succinate is completely mineralized through the tricarboxylic acid (TCA) cycle. In addition, 2-OH THF can spontaneously form its tautomer 4-hydroxybutyraldehyde, which is then converted to 4-hydroxybutyrate (16). The initial key metabolite 2-OH THF was first detected during THF degradation in the filamentous fungus Pseudallescheria boydii ZM01 (18). The production of γ-butyrolactone was verified after incubation of THF with Graphium sp. (16) and Rhodococcus sp. strain DTB pregrown on diethyl ether (19). The metabolite 4-hydroxybutyrate was identified in intracellular extracts of Rhodococcus aetherivorans strain M8 after derivatization with phenol boronate (11). According to previous extensive investigations of metabolites, the THF degradation pathway in the reported THF-degrading bacteria is highly similar.

The initial oxidation of THF is a multicomponent monooxygenase (MO)-catalyzed reaction, and previous studies have demonstrated that two different soluble di-iron monooxygenases (SDIMOs) are involved in the hydroxylation of THF and 1,4-dioxane (14D), which shares similar chemical properties with THF. The THF-degrading gene cluster thmADBC encoding THF MO is widely distributed in several closely related Pseudonocardia strains (including ENV478, CB1190, and K1) and Rhodococcus sp. strain YYL and is responsible for the initial degradation of THF and 14D (20 –22). Decreased translation of the thmB gene encoding the THF MO beta subunit by antisense RNA resulted in the loss of its ability to metabolize THF and 14D in Pseudonocardia sp. ENV478 (21). A host R. jostii RHA1 heterologously expressing thmADBC from P. dioxanivorans CB1190 and P. tetrahydrofuranoxydans K1 was able to transform THF and 14D, which explicitly verified the THF and 14D oxidation activity of THF MO (20). All of the reported THF MO gene clusters in Pseudonocardia and Rhodococcus strains are highly conserved, with the same genetic arrangement and high amino acid sequence similarity between corresponding subunits (94 to 99%). Furthermore, an investigation of the THF-degrading consortium by metagenomic sequencing revealed a 9,387-bp THF degradation gene cluster, thmX, encoding a THF MO from an as-yet-uncultivated Pseudonocardia species (23). The amino acid sequence similarities of the four ORFs between thmX and other reported THF MO gene clusters from P. tetrahydrofuranoxydans K1 and Pseudonocardia sp. ENV478 are between 79 and 93%. In addition, a novel propane MO gene cluster, prmABCD, in Mycobacterium dioxanotrophicus PH-06 was upregulated when using THF and 14D as induced substrates based on RNA sequencing and reverse transcription-quantitative PCR (RT-qPCR) (24). The prmABCD gene cluster was cloned and heterologously expressed in M. smegmatis mc²-155, and the transformant clones were capable of degrading THF, 14D, and propane, which demonstrates the catalytic function of propane MO to initiate the oxidation of THF and 14D (25). Previous studies providing a comprehensive comparison of catalytic behaviors of propane MO and THF MO revealed that the half-saturation coefficient (K_m) for propane MO is nearly four times lower than that of THF MO, which suggests propane MO has a higher affinity to 14D (26). Notably, all the THF degradation gene clusters reported thus far are from Gram-positive bacteria, and knowledge of the molecular mechanisms of THF degradation in Gram-negative bacteria is lacking.

In the present study, a new Gram-negative THF-degrading bacterium from THF-contaminated activated sludge was isolated, and the intermediate metabolites during THF degradation were identified. A novel THF degradation gene cluster, dmpKLMNOP, encoding a multicomponent THF hydroxylase responsible for THF degradation by strain ZM02 was characterized, which is substantially different from previously reported THF degradation gene clusters in Gram-positive bacteria. This research advances our understanding of the variety of THF-degrading genes and substrate diversity of THF hydroxylase, providing a new approach for the design of novel probes to rapidly assess the THF biodegradation potential at various THF-contaminated sites.

RESULTS

Isolation of the THF-degrading bacterium and detection of metabolites during THF degradation.

A new THF-degrading bacterium, designated strain ZM02, was isolated from activated sludge contaminated by THF. Strain ZM02 was able to aerobically utilize THF as a sole carbon and energy source, and 20 mM THF was significantly degraded in 168 h without a lag phase (Fig. 1A). In addition, strain ZM02 was capable of utilizing phenol for growth (Fig. 1B). The colonies of strain ZM02 were smooth, flat, creamy in color, and with regular margins after cultivation on Luria-Bertani (LB) agar plates for 4 days at 30°C (see Fig. S1A in the supplemental material). Cells are Gram-negative, short rods and occur singly (see Fig. S1B to D), which is consistent with the typical microscopic features of Cupriavidus sp. (27 –29). A phylogenetic tree constructed by 16S rRNA sequences (OK103907) showed that strain ZM02 shared a high degree of identity (100%) with Cupriavidus metallidurans (AY860234) (see Fig. S2). Thus, we finally identified the isolated THF-degrading bacterium C. metallidurans ZM02 based on morphological characteristics and phylogenetic relationship analysis. C. metallidurans ZM02 has been deposited in the China Center for Type Culture Collection (CCTCC), Wuhan, China, under accession no. CCTCC AB 2019263.

FIG 1 — Growth of *C. metallidurans* ZM02 with 20 mM THF (A) or 2.5 mM phenol (B) as the sole carbon and energy source and the associated degradation. Negative controls (NCs) without inoculation were prepared to evaluate the effects of THF or phenol volatilization. The data are averages of triplicates, and error bars indicate the standard deviations.

Intermediate metabolites in extracellular supernatants and intracellular extracts were detected during THF degradation by strain ZM02. With the degradation of THF, no metabolites were detected in extracellular supernatants (Fig. 2A), while two peaks appeared at retention times of 5.123 and 6.816 min (products I and II, respectively) in intracellular extracts (Fig. 2B, Fig. S3). The retention times of products I and II are consistent with the retention times of the authentic compound acetic acid and γ-butyrolactone, respectively (Fig. 2C). The metabolites accumulated in the cells were analyzed using gas chromatography-mass spectrometry (GC-MS). The GC-MS spectrum of product I was highly matched with acetic acid based on the comparison and analysis with the NIST database, and product I was finally identified as acetic acid. The GC-MS spectrum of γ-butyrolactone and 4-hydroxybutyrate are similar. To identify product II, the intracellular extracts were derivatized using N-methyl-N-trimethylsilyl trifluoroacetamide (MSTFA), and MSTFA-derived 4-hydroxybutyrate was detected by GC-MS with fingerprint ions of m/z 147 and m/z 233 (Fig. 2E). In addition, the MSTFA derivative of succinate, which was a downstream product of THF degradation, was also detected and identified (Fig. 2F). The metabolite 2-OH THF was not detected during THF degradation by strain ZM02 (see Fig. S3), which may be attributed to these intermediate metabolites being transient and rapidly converted.

FIG 2 — Analysis of intermediate metabolites during THF degradation by *C. metallidurans* ZM02. (A and B) Detection of THF metabolites in culture supernatants (A) and cell extracts (B) of strain ZM02 by GC analysis using 20 mM THF as a substrate. (C) Retention times of different authentic compounds by GC analysis. (D) The metabolite acetic acid was identified based on GC-MS analysis. (E and F) GC-MS spectra of the derivatives of 4-hydroxybutyrate (E) and succinate (F) in the cell extracts of strain ZM02.

Characterization of the putative THF hydroxylase gene cluster in C. metallidurans ZM02.

Genome sequencing demonstrated that the genome of C. metallidurans ZM02 was 6.26 Mb with a 63.07% GC content, including two chromosomes and a circular plasmid. Genome annotation analysis revealed that only one SDIMO gene cluster, designated dmpKLMNOP and located on chromosome 2, was found in the whole genome of strain ZM02 (see Table S1). Considering the key role of SDIMOs in THF degradation that has been reported, the dmpKLMNOP gene cluster, which has six complete open reading frames (ORFs) with a total size of approximately 4.6 kb and a GC content of 64% (Fig. 3), was speculated to be involved in THF degradation by strain ZM02. The putative THF hydroxylase Dmp_ZM02KLMNOP consists of four components, including the oxygenase Dmp_ZM02LNO, the reductase DmpP_ZM02, the accessory protein DmpK_ZM02, and the assembly protein DmpM_ZM02. In addition, a dmpR gene encoding an NtrC family transcriptional activator and a catA gene encoding catechol 1,2-dioxygenase are located upstream and downstream of dmpKLMNOP, respectively.

FIG 3 — Characterization of the THF-degrading gene cluster in *C. metallidurans* ZM02 and the proposed THF degradation pathway in reported THF-degrading strains. (A) Putative THF hydroxylase gene cluster (*dmpKLMNOP*) in strain ZM02. The numbers on the left and right ends indicate locations in the chromosome. The nucleotide sequence length of each gene is listed below that gene. (B) The THF degradation pathway is proposed based on the pathway originally suggested by Skinner (16) but supplemented with genes involved in THF degradation. THF hydroxylase, aldehyde dehydrogenase, and succinate semialdehyde dehydrogenase were encoded by *dmpKLMNOP*, *aldH*, and *sad*, respectively. The red boxes with dotted lines represent metabolites identified during THF degradation by strain ZM02.

The genetic arrangement of the putative THF hydroxylase gene cluster dmpKLMNOP in C. metallidurans ZM02 is different from those of the representative THF MO gene cluster thmADBC in R. ruber YYL, the propane MO gene cluster prmABCD in M. dioxanotrophicus PH-06, and the toluene MO gene cluster tmoABCDEF in Azoarcus sp. DD4 (Fig. 4A). The amino acid sequence alignment of the oxygenase large subunit indicated that DmpN_ZM02 showed less than 30% sequence identity to ThmA (R. ruber YYL), PrmA (M. dioxanotrophicus PH-06), and TmoA (Azoarcus sp. DD4). In addition, the amino acid sequence identity between the putative THF hydroxylase gene cluster dmpKLMNOP in strain ZM02 and other reported THF-degrading gene clusters in Gram-positive bacteria were both lower than 41% (Fig. 4A), which indicates that dmpKLMNOP is a completely different THF-degrading gene cluster. A phylogenetic tree constructed using the large subunit of SDIMOs showed that Dmp_ZM02KLMNOP belongs to SDIMOs and has a close relationship with phenol MO (group 2) (Fig. 4B). The putative THF hydroxylase in strain ZM02 shares the same number and type of subunits as the reported phenol MOs in phenol-degrading bacteria. The proteins encoded by the operon of strain ZM02 are more similar to those of R. eutropha strain E2 than to those of A. calcoaceticus strain NCIB8250 (Fig. 4A). Based on the analysis of the phylogenetic relationship between putative THF hydroxylase and phenol MOs, the putative THF hydroxylase Dmp_ZM02KLMNOP in C. metallidurans ZM02 is most phylogenetically related to phenol MOs in Ralstonia sp., and DmpN_ZM02 has a high degree of sequence identity to PhyC_KN1 (74%) in Ralstonia sp. KN1 and PoxD_E2 (73%) in Ralstonia sp. E2 (see Fig. S4).

FIG 4 — Structure alignment and phylogenetic analysis of *dmpKLMNOP*. (A) Amino acid sequence identity analysis between the putative THF hydroxylase gene cluster *dmpKLMNOP* in *C. metallidurans* ZM02 and other reported THF/phenol-degrading gene clusters. The blue box represents the reported THF/14D-degrading gene clusters, and the purple box represents the reported phenol-degrading gene clusters. The arrows indicate the transcript size and direction of the transcription of each gene, and genes with the same filled color are isoenzymes. The numbers under the genes represent the amino acid sequence identity of each component of MOs with the corresponding component in strain ZM02. (B) Unrooted phylogenetic tree generated based on amino acid sequences of the large subunit of SDIMOs. SDIMOs divided into the same group are represented by the same color. The strains marked with a gray triangle can utilize THF as a sole carbon source. Group 1, toluene/benzene MOs; group 2, phenol MOs; group 3, methane/butane MOs; group 4, alkene MOs; group 5, THF MOs; group 6, propane MOs.

The expression of THF hydroxylase is upregulated during growth on THF and phenol.

To determine whether dmpKLMNOP expression in C. metallidurans ZM02 was inducible, the expression levels during growth on THF, phenol, and citric acid were determined using transcriptome sequencing and RT-qPCR. Transcriptome analysis revealed that a total of 37 genes had expression fold changes higher than 4.0 in THF-induced cells (see Table S2), and the most highly upregulated genes were those in the THF hydroxylase operon. In addition, dmpLMNOPR and catA expression was upregulated when utilizing phenol as a substrate (see Fig. S5). RT-qPCR results confirmed that relative to citric acid, THF upregulated the expression of dmpLNPR and catA, and the expression fold changes were 10.5, 10.6, 11.2, 3.7, and 11.1 for the five subunits, respectively (Fig. 5). In addition, the expression levels of the THF hydroxylase gene cluster were higher after incubation with THF than after incubation with phenol (Fig. 5; see also Fig. S5 in the supplemental material). Notably, the significant upregulation of dmpR and catA expression in strain ZM02 under THF incubation implies that these genes play a role in the THF degradation pathway.

FIG 5 — Expression of *dmpL*, *dmpN*, *dmpP*, *dmpR*, and *catA* genes during *C. metallidurans* ZM02 growth on THF (5 mM) or phenol (5 mM) relative to that on citric acid (5 mM). The expression of THF degradation-related genes was verified by RT-qPCR. The 16S rRNA gene was used as a reference gene. At least three independent RNA samples were collected for each substrate.

Deletion of the THF hydroxylase gene inactivates the degradation of both THF and phenol in strain ZM02.

To validate the contribution of THF hydroxylase to THF degradation, we deleted dmpL, dmpN, and dmpP from the genome of strain ZM02. All knockout mutants were successfully obtained by a homologous recombination strategy and then inoculated into ammonium mineral salt (AMS) medium to verify their utilization of THF, phenol, catechol, and other THF-degrading intermediate metabolites. The ΔdmpL, ΔdmpN, and ΔdmpP deletion mutants were not able to utilize THF as a sole carbon source (Table 1), indicating that an intact dmpKLMNOP gene cluster is essential for THF degradation activity in strain ZM02. However, the deletion of dmpL, dmpN, and dmpP did not affect the utilization of the intermediate metabolites 2-OH THF or γ-butyrolactone (Table 1), which indicates that THF hydroxylase encoded by dmpKLMNOP may catalyze the initial step in the THF degradation pathway. In addition, the THF hydroxylase gene deletion mutants cannot use phenol for growth but can utilize catechol (Table 1), which indicates that THF hydroxylase encoded by dmpKLMNOP in strain ZM02 can also catalyze the conversion of phenol to catechol. To further investigate the transcriptional regulatory function of the dmpR gene in THF and phenol utilization in strain ZM02, the ΔdmpR deletion mutant was obtained, and deletion of the dmpR gene completely disabled THF and phenol utilization (Table 1). These results demonstrated that dmpR is obligatory for the transcription of dmpKLMNOP in strain ZM02 during growth on THF or phenol.

TABLE 1.

Growth of the wild-type strains and gene knockout strains on different substrates^a

Substrate	Strain (mean growth [g dry weight] ± SD)
Substrate	ΔdmpL	ΔdmpN	ΔdmpP	ΔdmpR	ZM02	JMP134
Tetrahydrofuran	NG	NG	NG	NG	0.30 ± 0.02	NG
2-Hydroxytetrahydrofuran	0.39 ± 0.03	0.40 ± 0.00	0.37 ± 0.03	0.52 ± 0.00	0.42 ± 0.01	0.37 ± 0.03
γ-Butyrolactone	0.59 ± 0.01	0.66 ± 0.02	0.58 ± 0.04	0.63 ± 0.01	0.65 ± 0.03	0.18 ± 0.00
Phenol	NG	NG	NG	NG	0.64 ± 0.01	0.79 ± 0.03
Catechol	0.58 ± 0.11	0.79 ± 0.03	0.50 ± 0.09	0.72 ± 0.04	0.66 ± 0.05	0.32 ± 0.01

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All of the strains were cultured in AMS medium containing 5 mM of each substrate at an initial OD₆₀₀ of 0.05 for 5 days at 30°C with 200 rpm. NG, no growth.

The THF hydroxylase encoded by dmpKLMNOP can oxidize THF.

Different recombinant vectors harboring all or part of the THF hydroxylase gene cluster were constructed to investigate whether all six genes are required for THF hydroxylase activity or some are involved in auxiliary functions. C. pinatubonensis JMP134 was chosen as a host for heterologous expression of the THF hydroxylase gene cluster because (i) it belongs to the same genus as strain ZM02 and (ii) C. pinatubonensis JMP134 is unable to grow on THF but can degrade the key intermediate metabolite 2-OH THF (Table 1). C. pinatubonensis JMP134 containing recombinant plasmid pBBR-dmpKLMNOP, which expresses the whole THF hydroxylase, grew on THF. However, plasmids pBBR-dmpLMNOP and pBBR-dmpKLMN, which lack dmpK and dmpOP, respectively, cannot support the growth of C. pinatubonensis JMP134 on THF. Surprisingly, C. pinatubonensis JMP134 harboring the plasmid pBBR-dmpKLMNO but not dmpP exhibited THF degradation ability, demonstrating that five genes (dmpKLMNO) must be present to allow C. pinatubonensis JMP134 growth on THF. Notably, coexpression of dmpR and dmpKLMNOP in C. pinatubonensis JMP134 increased the THF degradation rate (Fig. 6), indicating that dmpR promotes the transcriptional activity of the dmp operon that encodes THF hydroxylase for the catabolism of THF.

FIG 6 — (A and B) THF consumption (A) and growth (B) by *C. pinatubonensis* JMP134 containing different expression plasmids. Solid lines represent the residual THF concentration, and dashed lines represent the growth of *C. pinatubonensis* JMP134. The empty plasmid pBBR1MCS2 was also transformed into *C. pinatubonensis* JMP134 as a control group. All points represent the mean values of triplicate trials with error bars denoting the standard deviations.

THF and phenol oxidation activities of THF hydroxylase were detected by crude enzyme assays.

Different expression plasmids were constructed and expressed in host C. pinatubonensis JMP134 or E. coli BL21(DE3). SDS-PAGE analysis revealed that the DmpL_ZM02, DmpN_ZM02, and DmpP_ZM02 proteins were primarily present as particulate material in C. pinatubonensis JMP134 containing plasmid pBBR-dmpKLMNOP, and only a small soluble amount was found in the supernatant of C. pinatubonensis JMP134 containing pHGE-dmpKLMNOP (see Fig. S6A). No THF- or phenol-stimulated NAD(P)H oxidizing activity was detected in the crude extracts of C. pinatubonensis JMP134 containing plasmid pBBR-dmpKLMNOP or pHGE-dmpKLMNOP (Tables 2 and 3), which may be due to the insoluble expression and low expression level of the target protein. To improve the soluble expression of Dmp_ZM02KLMNOP, E. coli BL21(DE3) capable of expressing chaperonin GroELS was used as an expression host (see Fig. S6B). The chaperonin GroELS assists in correct folding and increases the solubility of target proteins produced in E. coli BL21(DE3).

TABLE 2.

Crude enzyme activity of different THF hydroxylase expression strains with THF

Reaction conditions	Mean enzyme activity (μmol min⁻¹ mg⁻¹) ± SD^a
Reaction conditions	C. pinatubonensis JMP134(pBBR-dmpKLMNOP)	C. pinatubonensis JMP134(pHGE-dmpKLMNOP)	E. coli BL21(DE3)(pET28a-dmpKLMNOP)
Cell extract+NADH+THF	ND	ND	0.29 ± 0.11
Cell extract+NADH+Fe²⁺	ND	ND	ND
Cell extract+NADH+Fe²⁺+THF	ND	ND	0.53 ± 0.07
Cell extract+NADPH+THF	ND	ND	1.04 ± 0.20
Cell extract+NADPH+Fe²⁺	ND	ND	ND
Cell extract+NADPH+Fe²⁺+THF	ND	ND	1.38 ± 0.24

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ND, not detectable. The numbers indicate the THF-stimulated NAD(P)H oxidation rates.

TABLE 3.

Crude enzyme activity of different THF hydroxylase expression strains with phenol

Reaction conditions	Mean enzyme activity (μmol min⁻¹ mg⁻¹) ± SD^a
Reaction conditions	C. pinatubonensis JMP134(pBBR-dmpKLMNOP)	C. pinatubonensis JMP134(pHGE-dmpKLMNOP)	E. coli BL21(DE3)(pET28a-dmpKLMNOP)
Cell extract+NADH+phenol	ND	ND	0.82 ± 0.09
Cell extract+NADH+Fe²⁺	ND	ND	ND
Cell extract+NADH+Fe²⁺+phenol	ND	ND	1.19 ± 0.14
Cell extract+NADPH+phenol	ND	ND	1.54 ± 0.23
Cell extract+NADPH+Fe²⁺	ND	ND	ND
Cell extract+NADPH+Fe²⁺+phenol	ND	ND	1.78 ± 0.37

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ND, not detectable. The numbers indicate the phenol-stimulated NAD(P)H oxidation rates.

Since THF and 2-OH THF had no characteristic absorption peaks in the range from 200 to 800 nm (see Fig. S7), it is difficult to show the NAD(P)H-dependent consumption of the THF substrate or the NAD(P)H-dependent appearance of the catalytic product 2-OH THF spectrophotometrically. The THF and phenol catalytic activity of THF hydroxylase were evaluated based on the oxidation of the electron donor NADPH or NADH. An obvious decrease in the characteristic absorption peak of NADH or NADPH (λ_max, 340 nm) was observed in cell extracts of E. coli BL21(DE3) harboring plasmid pET28a-dmpKLMNOP (Fig. 7C and D). Compared with NADH as an electron donor, the THF and phenol catalytic activity of THF hydroxylase was higher when using NADPH as an electron donor. In addition, the addition of Fe²⁺ as a cofactor improved the THF or phenol catalytic activity (Tables 2 and 3). The highest THF and phenol catalytic activity improved to 1.38 ± 0.24 μmol min⁻¹mg⁻¹ and 1.77 ± 0.37 μmol min⁻¹mg⁻¹, respectively, with the addition of NADPH and Fe²⁺ (Tables 2 and 3).

FIG 7 — Detection of THF catalytic activity in cell extracts of strain *E. coli* BL21(DE3) containing plasmid pET28a-*dmpKLMNOP* or empty vector pET28a. The reaction was initiated by the addition of 0.2 mM THF, and the spectra were recorded every minute after the addition of THF for 10 min. The arrows indicate the directions of spectral changes. (A) The reaction mixtures of the control group contained cell extracts of *E. coli* BL21(DE3) containing plasmid pET28a-*dmpKLMNOP*, NADH, and Fe²⁺. (B) The reaction mixtures of the control group contained cell extracts of *E. coli* BL21(DE3) containing plasmid pET28a-*dmpKLMNOP*, NADPH, and Fe²⁺. (C) The reaction mixtures of the control group contained cell extracts of *E. coli* BL21(DE3) containing empty vector pET28a, NADPH, THF, and Fe²⁺. (D) The reaction mixtures contained cell extracts of *E. coli* BL21(DE3) containing plasmid pET28a-*dmpKLMNOP*, NADH, THF, and Fe²⁺. (E) The reaction mixtures contained cell extracts of *E. coli* BL21(DE3) containing plasmid pET28a-*dmpKLMNOP*, NADPH, THF, and Fe²⁺.

The representative phenol-degrading bacteria containing similar THF-degrading gene clusters cannot degrade THF alone.

The THF hydroxylase in strain ZM02 and phenol MO, which catalyzes the degradation of phenol in phenol-degrading bacteria, are highly similar, and the gene clusters of the two MOs are similar in genetic composition and arrangement. To evaluate whether phenol-degrading bacteria have the potential to degrade THF alone, three representative phenol-degrading bacteria containing dmp-like clusters were inoculated into AMS medium with THF. The amino acid sequence of the phenol MO gene cluster in C. pinatubonensis JMP134 is very similar to that of the THF hydroxylase gene cluster in strain ZM02, and the identities of homologous subunits are between 67 and 95% (Fig. 8A). The amino acid sequence identities of THF hydroxylase are 50, 50, 50, 71, 44, and 58% and 47, 43, 53, 67, 37, and 55% to the corresponding enzyme encoded by dmpK, dmpL, dmpM, dmpN, dmpO, and dmpP from Sphingomonas melonis TY and Alcaligenes faecalis JQ135, respectively (Fig. 8A). Representative phenol-degrading bacteria C. pinatubonensis JMP134, A. faecalis JQ135, and S. melonis TY could grow with phenol as the sole carbon source (data not shown) but could not grow with THF as the sole carbon source (Fig. 8B).

FIG 8 — Comparative analysis of the THF-degrading bacterium *C. metallidurans* ZM02 and other representative phenol-degrading bacteria. (A) Comparative analysis of the THF hydroxylase gene cluster *dmpKLMNOP* in *C. metallidurans* ZM02 and phenol MO gene clusters in representative phenol-degrading bacteria. The arrows indicate the size and direction of the transcription of each gene, and genes with the same fill color are isoenzymes. The numbers under the genes represent the amino acid sequence identity of each component of phenol MO with the corresponding component in strain ZM02. (B) The growth and degradation curves of different phenol-degrading bacteria utilizing 2 mM THF as a substrate. The solid lines indicate the THF concentrations, and the dotted lines indicate the OD₆₀₀ values.

DISCUSSION

SDIMOs are multicomponent nonheme bacterial enzymes capable of hydroxylating a broad variety of substrates, such as aromatic hydrocarbons, alkanes, alkenes, and chlorinated solvents (30 –33). SDIMOs are usually transcribed from a single operon encoding four to six polypeptides and are categorized into six subgroups (group 1 to group 6) based on catalytic substrate specificity, phylogenetic analyses of the MO large subunit, the number of subunits, and genetic arrangement (25, 34). Previous investigations on the prevalence of SDIMOs indicated that there is an incredibly wide distribution of SDIMOs in bacterial consortia from various contaminated sites and a high diversity of SDIMOs among bacteria obtained in pure culture (35 –38). SDIMOs play an indispensable role in the oxidation of THF, and two subgroups of SDIMOs (THF and propane MOs) derived from Gram-positive THF-degrading bacteria have been demonstrated to be associated with THF degradation (21, 25). In this study, a novel THF-degrading gene cluster, dmpKLMNOP, encoding THF hydroxylase in the Gram-negative bacterium C. metallidurans ZM02, was identified and characterized.

According to our investigation, the THF hydroxylase in strain ZM02 shares the same number and type of subunits and the same genetic arrangement as the phenol MO that is widespread in phenol-degrading bacteria (Fig. 4A), which implies that THF hydroxylase is highly homologous to phenol MO. The phenol MO gene cluster is generally found in Gram-negative bacteria (39), and more than 10 such gene clusters have been reported, including mphKLMNOP (40), dmpKLMNOP (41), mopKLMNOP (42), phcKLMNOP (43, 44), afpKLMNOP (45), and aphKLMNOP (46). Notably, the THF hydroxylase gene cluster dmpKLMNOP in C. metallidurans ZM02 was unlikely to have been recently horizontally transferred from other phenol-degrading bacteria for the following reasons: (i) the GC content of THF hydroxylase (64%) was close to that of the genome of C. metallidurans ZM02 (63%), and (ii) no transposase gene or insertion sequences were found near the THF hydroxylase gene cluster dmpKLMNOP on the chromosome of C. metallidurans ZM02, which is different from the case of the THF MO gene cluster thmADBC (47) and the phenol MO gene cluster afpKLMNOP (45). Interestingly, phenol-degrading bacteria containing similar THF hydroxylase gene clusters are abundant in the natural environment, but the ability of phenol-degrading bacteria to degrade THF alone has rarely been reported. The representative phenol-degrading bacteria C. pinatubonensis JMP134, A. faecalis JQ135 and S. melonis TY, which contain dmp clusters similar to those of C. metallidurans ZM02, can degrade phenol but not THF. The evolutionary trajectory of THF hydroxylase is likely to remain unclear, and certain factors may affect the degradation of THF by phenol-degrading bacteria containing similar THF-degrading gene clusters: (i) THF hydroxylase originates from phenol MO, which has the potential to catalyze the oxidation of THF and phenol, while the original phenol MO cannot degrade THF; and (ii) the expression of the THF hydroxylase gene cluster in strain ZM02 can be induced by THF alone, but the expression of the phenol MO gene cluster, which is closely evolutionarily related to the THF hydroxylase gene cluster, cannot be induced by THF in phenol-degrading bacteria.

Phenol MO catalyzes the hydroxylation of a series of aromatic compounds, such as phenol, chlorophenol, cresol isomers, dimethylphenol isomers, benzene, toluene, biphenyl, naphthalene, and alkylphenol (48, 49). In addition, the phenol MO from Pseudomonas stutzeri OX1 is a promiscuous hydroxylase that can catalyze the ortho-hydroxylation of several nonnatural substrates, including catechol, 4-hydroxybenzoic acid, and resorcinol (50). Disruption of the dmpL, dmpN, and dmpP genes abolished the growth of strain ZM02 in AMS medium containing THF or phenol, which indicates that the THF hydroxylase encoded by dmpKLMNOP is involved in both THF and phenol degradation. THF hydroxylase, which is probably evolved from phenol MO that can catalyze phenol hydroxylation at the ortho position to form catechol (51, 52), can also oxidize THF at the carbon position adjacent to the oxygen atom to form 2-OH THF. In the present study, the metabolites 4-hydroxybutyrate and succinate were detected and identified in the intracellular extracts, which suggests that THF is metabolized intracellularly by strain ZM02. Notably, the accumulation of acetic acid, a metabolic by-product, was observed during the degradation of 20 mM THF by strain ZM02, which may be due to the overflow metabolism phenomenon (53, 54).

The oxygenase component of phenol MO is usually constituted by three polypeptides (L, N, and O) and has a catalytic function as a dimeric (LNO)₂ subcomplex (50). A reductase component (DmpP) containing an [2Fe-2S] iron-sulfur cluster and a FAD as a cofactor is responsible for transferring electrons from NADH or NADPH electron donors to the oxygenase component (55). The assay above strongly suggests that dmpKLMNOP encodes the complete components of THF hydroxylase that together catalyze the oxidation of THF and phenol. Surprisingly, C. pinatubonensis JMP134, harboring a plasmid that expresses five polypeptides (DmpK_ZM02 to DmpO_ZM02) but not DmpP_ZM02, also exhibited the same THF degradation ability as complete THF hydroxylase. Why can C. pinatubonensis JMP134 heterologously expressing dmpKLMNO degrade THF? dmpP encodes the FAD- and [2Fe-2S]-containing reductase component of THF hydroxylase, which is responsible for transferring electrons into the diiron cluster in the active site. We speculate that genes encoding proteins with a similar function to DmpP_ZM02 in the genome of C. pinatubonensis JMP134 play an alternative role during the degradation of THF.

In summary, we identified a novel THF hydroxylase encoded by dmpKLMNOP in a newly isolated Gram-negative THF-degrading bacterium, Cupriavidus metallidurans ZM02 and demonstrated its catalytic function of initiating the oxidation of THF and phenol. Our study uncovered a THF hydroxylase involved in both phenol and THF degradation. The THF oxidation activity of THF hydroxylase was verified in C. pinatubonensis JMP134 heterologously expressing the dmpKLMNOP gene cluster. In addition, substantial evidence for the active expression of THF hydroxylase gene cluster in E. coli BL21(DE3) has been obtained. Our findings expand our understanding of the diversity of THF degradation gene clusters and broaden the substrate spectrum of THF hydroxylase.

MATERIALS AND METHODS

Bacterial strains, culture conditions, and chemicals.

The bacterial strains and plasmids used in this study are listed in Table 4, and all primers are listed in Table 5. The THF-degrading strain was isolated from activated sludge and cultivated in AMS medium at 30°C on a rotary shaker (200 rpm) with 20 mM THF as the sole carbon source. The AMS medium was prepared with the following components (per L): 0.66 g (NH₄)₂SO₄, 1 g MgSO₄·7H₂O, 0.017 g CaCl₂·2H₂O, 1 mL stock A (5 g Fe-Na EDTA and 2 g Na₂MoO₄·2H₂O per L, stored at 4°C), 1 mL trace element stock solution (0.5 g FeSO₄·7H₂O, 0.4 g ZnSO₄·7H₂O, 0.02 g MnSO₄·H₂O, 0.015 g H₃BO₃, 0.01 g NiCl₂·6H₂O, 0.25 g EDTA, 0.05 g CoCl₂·6H₂O and 0.005 g CuCl₂·2H₂O per L, stored at 4°C), and 20 mL 1 M phosphate buffer (113 g K₂HPO₄ and 47 g KH₂PO₄ per L).

TABLE 4.

Bacterial strains and plasmids used in this study

Strain or plasmid	Characteristics^a	Reference or source
Strains
C. metallidurans ZM02	Wild type, THF-degrading strain; G-, Tc^s	This study
C. pinatubonensis JMP134	Wild type, phenol-degrading strain, used for gene expression; G-, Km^s	60
S. melonis TY	Wild type, phenol-degrading strain	61
A. faecalis JQ135	Wild type, phenol-degrading strain	62
E. coli
DH5α	supE44 lacU169(80dlacZΔM15) hsdR17 recA1 endA1 gyrA96 Δthi relA1	Tsingke
BL21(DE3)	Chaperonin gene groELS integrated in chromosome	63
ΔdmpL	ZM02 mutant with dmpL gene replaced by Tc resistance gene	This study
ΔdmpN	ZM02 mutant with dmpN gene replaced by Tc resistance gene	This study
ΔdmpP	ZM02 mutant with dmpP gene replaced by Tc resistance gene	This study
ΔdmpR	ZM02 mutant with dmpR gene replaced by Tc resistance gene	This study
JMP134-pBBR	JMP134 transformed with pBBR1MCS2; Km^r	This study
JMP134-pBBR-dmpKLMNOPR	JMP134 transformed with pBBR-dmpKLMNOPR; Km^r	This study
JMP134-pBBR-dmpKLMNOP	JMP134 transformed with pBBR-dmpKLMNOP; Km^r	This study
JMP134-pBBR-dmpKLMNO	JMP134 transformed with pBBR-dmpKLMNO; Km^r	This study
JMP134-pBBR-dmpLMNOP	JMP134 transformed with pBBR-dmpLMNOP; Km^r	This study
JMP134-pBBR-dmpKLMN	JMP134 transformed with pBBR-dmpKLMN; Km^r	This study
JMP134-pHGE	JMP134 transformed with pHGE; Km^r	This study
JMP134-pHGE-dmpKLMNOP	JMP134 transformed with pHGE-dmpKLMNOP; Km^r	This study
BL21(DE3)-pET28a	BL21(DE3) transformed with pET28a; Km^r	This study
BL21(DE3)-pET28a-dmpKLMNOP	BL21(DE3) transformed with pET28a-dmpKLMNOP; Km^r	This study

Plasmids
pEX18Tc	Gene knockout vector, oriT⁺, sacB⁺; Tc^r	64
pEX18Tc-dmpL	dmpL gene knockout vector; Tc^r	This study
pEX18Tc-dmpN	dmpN gene knockout vector; Tc^r	This study
pEX18Tc-dmpP	dmpP gene knockout vector; Tc^r	This study
pEX18Tc-dmpR	dmpR gene knockout vector; Tc^r	This study
pBBR1MCS2	Gene expression vector; Km^r	64
pBBR-dmpKLMNOPR	Vector for expression of dmpKLMNOPR; Km^r	This study
pBBR-dmpKLMNOP	Vector for expression of dmpKLMNOP; Km^r	This study
pBBR-dmpKLMNO	Vector for expression of dmpKLMNO; Km^r	This study
pBBR-dmpKLMN	Vector for expression of dmpKLMN; Km^r	This study
pBBR-dmpLMNOP	Vector for expression of dmpLMNOP; Km^r	This study
pHGE	P_tac, IPTG-inducible expression vector; Km^r	65
pHGE-dmpKLMNOP	Vector for inducible expression of dmpKLMNOP; Km^r	This study
pET28a	Gene expression vector; Km^r	Novagen
pET28a-dmpKLMNOP	Vector for inducible expression of dmpKLMNOP; Km^r	This study

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Tc^r, tetracycline resistance; Km^r, kanamycin resistance; Km^s, kanamycin sensitivity; G-, gram-negative bacterium.

TABLE 5.

Primers used in this study

Primer	Sequence (5′–3′)^a	Purpose
dmpL-UF	CGGTACCCGGGGATCCCGTAGCCAAGCTGCACCCA	Construct dmpL gene knockout vector
dmpL-UR	ATAAACAAATAGGGGTTCCGCGGGTGGTTGTCTCCTCGGATGT
dmpL-DF	AGCCGGGCCACCTCGACCTGACGGAGACAACCATGAGCGAC
dmpL-DR	GGCCAGTGCCAAGCTTGAACGTCACCGTGGCCATGT
dmpN-UF	CGGTACCCGGGGATCCTCGCTCTTGCGCACGAAGAAAC	Construct dmpN gene knockout vector
dmpN-UR	ATAAACAAATAGGGGTTCCGCGACAAGGAGACCAGATCATGCCAG
dmpN-DF	AGCCGGGCCACCTCGACCTGAGGTTTGTCTCCTGTCTTGGGGTT
dmpN-DR	GGCCAGTGCCAAGCTTATTTCGTGGACAAGCGAGGGC
dmpP-UF	CGGTACCCGGGGATCCCTGGTCCTGCAACGCTAAAGC	Construct dmpP gene knockout vector
dmpP-UR	ATAAACAAATAGGGGTTCCGCGAGTATTTCCACCACAAAGGAGACA
dmpP-DF	AGCCGGGCCACCTCGACCTGAGGCAACCTCCTGCAGCAAGT
dmpP-DR	GGCCAGTGCCAAGCTTTTCGAGCAGAACGGTGGGGC
dmpR-UF	CGGTACCCGGGGATCCAGGCGGTAATCGACGCGTTG	Construct dmpR gene knockout vector
dmpR-UR	ATAAACAAATAGGGGTTCCGCGCCTTGTCACTCCAGATTCCGGC
dmpR-DF	AGCCGGGCCACCTCGACCTGAGGTTGGGACAGTTGGCCCAG
dmpR-DR	GGCCAGTGCCAAGCTTTGTGCGCGGACGGATTACCG
Tc-F	CGCGGAACCCCTATTTGTTTAT	Amplify Tc gene
Tc-R	TCAGGTCGAGGTGGCCCGGC	Amplify Tc gene
pEX18Tc-F	ACTCATTAGGCACCCCAGGC	Verify gene knockout vector
pEX18Tc-R	CAAGCTCGCCATTCGCCATT	Verify gene knockout vector
ΔdmpL-UF	TTGACCACTTCCAGGCCCAGC	Verify dmpL gene knockout mutant
ΔdmpL-UR	GGAAGCAGCCCAGTAGTAGGT
ΔdmpL-DF	ATATCGCCGACATCACCGATG
ΔdmpL-DR	CGCTCGCATTCGGTTTCCTTG
ΔdmpN-UF	TCTGCCGCCGACAGGAACTT	Verify dmpN gene knockout mutant
ΔdmpN-UR	AGTGCGGCGACGATAGTCATG
ΔdmpN-DF	CGTCCTGTGGATTCTCTACGCC
ΔdmpN-DR	AGACCTTCGCGCATATCGCG
ΔdmpP-UF	TGTTCCTGTCCCTGCGGCAC	Verify dmpP gene knockout mutant
ΔdmpP-UR	GCTGACTGGGTTGAAGGCTCT
ΔdmpP-DF	GACTGTTGGGCGCCATCTCC
ΔdmpP-DR	CTACAACGGGGACATGGCCAC
ΔdmpR-UF	CTGCCAGGCCATGTCTTCCAT	Verify dmpR gene knockout mutant
ΔdmpR-UR	AGGGCGTGCAAGATTCCGAA
ΔdmpR-DF	ATGCAGGAGTCGCATAAGGG
ΔdmpR-DR	CGACATGGGTCAGCACCAGAT
pBBR-dmpKR-F	CGGTATCGATAAGCTCTACTTCTTCAACCGATACGCCAGC	Design pBBR-dmpKLMNOPR expression vector
pBBR-dmpKR-R	CGGGCTGCAGGAATTTCAGAGTGAGCGGAACAGCG	Design pBBR-dmpKLMNOPR expression vector
pBBR-dmpKP-F	CGGTATCGATAAGCTATGAGATATCCCCCTCAGGCAC	Design pBBR-dmpKLMNOP expression vector
pBBR-dmpKP-R	CGGGCTGCAGGAATTTCAGAGTGAGCGGAACAGCG	Design pBBR-dmpKLMNOP expression vector
pBBR-dmpKO-F	CGGTATCGATAAGCTATGAGATATCCCCCTCAGGCAC	Design pBBR-dmpKLMNO expression vector
pBBR-dmpKO-R	CGGGCTGCAGGAATTTCAGTTGGCCGAGCCGGC	Design pBBR-dmpKLMNO expression vector
pBBR-dmpKN-F	CGGTATCGATAAGCTATGAGATATCCCCCTCAGGCAC	Design pBBR-dmpKLMN expression vector
pBBR-dmpKN-R	CGGGCTGCAGGAATTTCACGAACGGTTGTTGCCTTG	Design pBBR-dmpKLMN expression vector
pBBR-dmpLP-F	CGGTATCGATAAGCTATGCAAATCGATATCCGGACTG	Design pBBR-dmpLMNOP expression vector
pBBR-dmpLP-R	CGGGCTGCAGGAATTTCAGAGTGAGCGGAACAGCG	Design pBBR-dmpLMNOP expression vector
pBBR-F	AGCGCGCAATTAACCCTCACT	Verify gene expression vector
pBBR-R	ACTATAGGGCGAATTGGAGCTCC	Verify gene expression vector
pHGE-dmpKP-F	CACAGGAGAGAATTCATGACCACGCCCATCGTCCA	Design pHGE-dmpKLMNOP expression vector
pHGE-dmpKP-R	CAAAACAGCCAAGCTTTCAGAGCGAGCGGAACAGC	Design pHGE-dmpKLMNOP expression vector
pHGE-F	ATGAGCCGTGTTTTCTGGACGAT	Verify gene expression vector
pHGE-R	ATGCCTGGCAGTTCCCTACTCT	Verify gene expression vector
pET28-dmpKP-F	AGGAGATATACCATGAGATATCCCCCTCAGGCAC	Design pET28a-dmpKLMNOP expression vector
pET28-dmpKP-R	AGCCGGATCTCAGTGGTGGTGGTGGTGGTGTCAGAGTGAGCGGAACAGCG	Design pET28a-dmpKLMNOP expression vector
pET28a-F	AGCCCAGTAGTAGGTTGAGGCC	Verify gene expression vector
pET28a-R	AGCAAAAAACCCCTCAAGACCCG	Verify gene expression vector
dmpL-qPCR-F	GGAGAAGAACCTTGATTTC	RT-PCR
dmpL-qPCR-R	TCCATCGTGTGATACATC	RT-PCR
dmpN-qPCR-F	GAAGTGGTCAAGTTCATG	RT-PCR
dmpN-qPCR-R	TGTAGTCCATCATCATCG	RT-PCR
dmpP-qPCR-F	CCGATGACGAACTGTATT	RT-PCR
dmpP-qPCR-R	GAAGTCGTAGCCGAAATG	RT-PCR
dmpR-qPCR-F	CTTCTATGGCGAGTTTCT	RT-PCR
dmpR-qPCR-R	CATTCGGTTTCCTTGTAG	RT-PCR
catA-qPCR-F	GGCAACTACTCGTACTTC	RT-PCR
catA-qPCR-R	GAGACGAAGAAATGGATATG	RT-PCR
16S rRNA-qPCR-F	CCTACGGGAGGCAGCAG	RT-PCR, 16S rRNA
16S rRNA-qPCR-R	ATTACCGCGGCTGCTGG	RT-PCR, 16S rRNA

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Underlined nucleic acid sequences represent homologous sequences derived from plasmids.

E. coli DH5α and BL21(DE3) strains were used for the construction and expression of recombinant plasmids and were cultivated at 37°C in LB broth or on LB plates. C. pinatubonensis JMP134 was used as a heterologous expression host and grown aerobically in LB or AMS medium at 30°C. The phenol-degrading bacteria A. faecalis JQ135 and S. melonis TY were cultivated in AMS medium containing THF and phenol at 30°C. The suicide plasmid pEX18Tc was used for gene deletion. The broad-host-range expression vectors pBBR1MCS2 and pHGE were used to construct the coexpression vectors expressed in C. pinatubonensis JMP134, and vector pET28a was used for the expression of THF hydroxylase in E. coli BL21(DE3). The antibiotics used for selection in this study were at the following concentrations: kanamycin (Km), 20 μg mL⁻¹; and tetracycline (Tc), 50 μg mL⁻¹. THF (>99% purity), γ-butyrolactone (>99% purity), catechol (>99% purity), NADH, and NADPH were purchased from Aladdin Industrial Corporation (Shanghai, China). Phenol (>99% purity) was purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). 2-OH THF (>95% purity) was purchased from Macklin Biochemical Co., Ltd. (Shanghai, China). THF was stored in brown, narrow-mouthed bottles, and all liquid culture experiments containing THF were performed in 250 mL narrow-mouthed shake flasks. The bottle mouth was sealed with aluminum foil to reduce the volatilization of THF during the cultivation process. The sonicator with a 2 mm ultrasonic horn used in this research was purchased from Ningbo Scientz Biotechnology Co., Ltd. (JY92-N; Ningbo, China).

Isolation of THF-degrading bacteria and detection of THF metabolic intermediates.

THF-degrading bacteria were enriched and isolated from activated sludge collected from the Harbin Sewage Treatment Plant, Heilongjiang Province, China. After 28 days of continuous-passage acclimatization in AMS medium with THF concentrations from 20 to 50 mM at 30°C, a stable THF-degrading consortium was obtained. The enriched culture was serially diluted and plated on AMS agar plates containing 20 mM THF at 30°C for 4 days to isolate pure cultures. Strain ZM02 cultured to logarithmic phase in LB medium was collected by centrifugation at 4,600 × g for 5 min at 4°C, and the bacterial pellets were resuspended and washed twice with sterile water to remove the carried-over carbon source. The THF and phenol degradation abilities were evaluated in 100 mL of AMS with 20 mM THF and 2.5 mM phenol at an initial optical density at 600 nm (OD₆₀₀) of 0.05, respectively. A negative control without inoculation was prepared to evaluate the effect of THF volatilization. The residual substrate concentration and THF degradation-related metabolites were detected by GC-2014C gas chromatography equipped with a flame ionization detector and an AOC-20i autoinjector (Shimadzu, Shanghai, China). Samples were centrifuged at 10,000 × g for 2 min, and the supernatant was subsequently filtered through a 0.45-μm filter for pretreatment. The THF concentration was detected as described in a previous study (56), and the phenol concentration was measured by the same program with a slight modification as follows: the column temperature was set to 160°C and then held for 12 min. Standard curves of THF and phenol were appropriately prepared, and the residual substrate concentration was calculated. The limit of detection for THF and phenol was 0.01 mM using the methods described above.

To detect the intermediate metabolites during THF degradation by strain ZM02, 10 mL of cultivation liquid was collected and centrifuged at 15,000 × g for 2 min, and the supernatant was pretreated as described above before GC analyses. In addition, the cells were washed twice and resuspended in ice-cold 20 mM phosphate buffer (pH 7.4). Cell suspensions were then disrupted by sonication in an ice-water bath for 5 min (sonication for 3 s with 4-s intervals). Cell debris was removed by centrifugation at 15,000 × g for 10 min at 4°C, and cell extracts were collected for the detection of metabolites. For derivatization, 40 μL of MSTFA was added to the samples, followed by incubation at 70°C for 40 min, and then the samples were cooled to room temperature for analysis. The detection of intermediate metabolites by GC was conducted as described in our previous study (18).

Genome and transcriptome sequencing.

In this study, genomic DNA extraction, sequencing library construction, and sequencing were outsourced to Novogene Biotech Company (Beijing, China). Whole-genome sequencing of strain ZM02 was performed using a PacBio platform, and gene prediction and annotation were accomplished by Rapid Annotations using Subsystems Technology (RAST; http://rast.nmpdr.org/) (57). The Kyoto Encyclopedia of Genes and Genomes (KEGG), Clusters of Orthologous Groups (COG), Non-Redundant (NR) Protein Database, and Gene Ontology (GO) were used for general function annotation. For transcriptome sequencing, strain ZM02 was cultured and harvested after growth to mid-logarithmic phase on 5 mM THF, phenol, and citric acid. Total RNA was extracted with a spin column bacterial total RNA purification kit (Sangon Biotech, Shanghai, China). RNA sequencing was accomplished using the Illumina HiSeq4000 platform (giving 400 × coverage) by MAGIGENE Biotech Company (Shenzhen, China).

Reverse transcription-quantitative PCR.

Total RNA of strain ZM02 cultured to logarithmic phase was prepared as described above. RNA integrity was assessed by agarose gel electrophoresis, and RNA concentration was detected by a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA). cDNA was synthesized using high-capacity cDNA reverse transcription kits (Applied Biosystems, Foster City, CA) and then stored at −20°C. The RT-qPCR mixture contained 0.4 μL of diluted cDNA (25 ng μL⁻¹), 10 μL of 2×SYBR green PCR master mix (Applied Biosystems), 0.4 μM concentrations of each forward and reverse primer, and enough DNA-free water to reach a total volume of 20 μL. RT-qPCR was performed in a Rotor-Gene Q real-time PCR detection system (Qiagen, Germany) with the following conditions: 95°C for 2 min and 40 cycles of 95°C for 10 s, 60°C for 20 s, and 72°C for 15 s. Specific primers targeting each subunit of the hydroxylase gene cluster were designed by Beacon Designer 7 (Table 5). Differential gene expression was quantified by using the 2^−ΔΔ^CT method with 16S rRNA as the reference gene. Treatments with citric acid as the sole carbon source were used as controls to calculate the expression levels of target genes in the presence of THF or phenol.

Gene knockout experiments.

Deletion mutants were constructed by homologous recombination. Gene knockout vectors containing two DNA fragments homologous to the upstream and downstream regions (800 bp), Tc resistance gene, and suicide plasmid pEX18Tc digested by BamHΙ and HindΙΙΙ were constructed by using an In-Fusion HD cloning kit (TaKaRa Bio, USA). The upstream and downstream DNA fragments from C. metallidurans ZM02 were amplified by PCR using the corresponding primers listed in Table 5. The corresponding deletion plasmids were electroporated into strain ZM02 at 2.5 kV for 12 ms, and double-crossover mutants were screened on LB plates containing Tc (50 μg mL⁻¹). The mutants were verified by PCR and DNA sequencing.

Cloning and heterologous expression of the putative THF-degrading gene cluster.

Combinations of THF-degrading gene clusters were cloned from strain ZM02 genomic DNA using the primers listed in Table 5. The products were inserted into the EcoRI and HindIII sites of pBBR1MCS2 using the method described above. All the inserted fragments were sequenced to ensure that no mutations had been introduced in the heterologous expression vectors. The recombinant strains were grown on 100 mL of AMS medium containing 5 mM THF with an initial OD₆₀₀ of 0.05 at 30°C. C. pinatubonensis JMP134 containing only the empty plasmid pBBR1MCS2 was used as a control.

Preparation of cell extracts and crude enzyme assays.

The complete ORF of THF hydroxylase was amplified using the genomic DNA of strain ZM02 and inserted into the EcoRI and HindIII sites of plasmids pHGE and pET28a, resulting in plasmids pHGE-dmpKLMNOP and pET28a-dmpKLMNOP, respectively. Strain JMP134 containing plasmid pHGE-dmpKLMNOP was inoculated into 100 mL of AMS medium with 5 mM THF at the initial OD₆₀₀ of 0.05 and cultivated for 4 days with the addition of 0.1 mM IPTG (isopropyl-β-d-thiogalactopyranoside). E. coli BL21(DE3) carrying the recombinant plasmid pET28a-dmpKLMNOP or empty vector pET28a was grown in LB medium at 37°C to an OD₆₀₀ of 0.6 and then induced with 0.5 mM IPTG for 14 h at 30°C. The method used for the preparation of cell extracts described previously (58, 59) was modified as follows: 50 mL of cultivation liquid was centrifuged at 15,000 × g for 10 min, and cells were harvested and then resuspended in 10 mL of ice-cold 50 mM Tris-HCl buffer (pH 7.4). The suspension was disrupted by sonication in an ice-water bath for 30 min (sonication for 3 s with 4-s intervals), and cell debris was removed by ultracentrifugation at 100,000 × g for 1 h at 4°C. Protein concentrations were estimated by using a BCA kit (Beyotime, Shanghai, China). The reaction mixtures contained cell extract (5 mg mL⁻¹), 0.2 mM THF/phenol, 0.1 mM NADH/NADPH, and 0.1 mM Fe²⁺, and the final volume was adjusted to 1 mL with 50 mM Tris-HCl buffer (pH 7.4). The reaction mixtures using cell extracts of E. coli BL21(DE3) containing the empty vector or containing all of these reaction components except substrate were used as negative controls. The assay was initiated by the addition of THF or phenol. The molar extinction coefficients for NADH and NADPH were 4,530 M⁻¹ cm⁻¹ and 5,220 M⁻¹ cm⁻¹ at 340 nm, respectively. The activity of the crude enzyme was assayed by measuring the decrease in absorbance at 340 nm due to the consumption of NADH or NADPH in the presence of THF or phenol. One unit of enzyme activity was defined as the amount of enzyme required to reduce 1 μM electron donor per min at 30°C.

Data availability.

The 16S rRNA sequence and the genome sequence of C. metallidurans ZM02 have been deposited in GenBank under accession numbers OK103907 and CP083718 to CP083720, respectively. The transcriptomic data for strain ZM02 growing on different substrates have been deposited in the SRA database under accession numbers SRR17236270 to SRR17236278.

ACKNOWLEDGMENTS

This study was financially supported by grants from the National Natural Science Foundation of China (41630637, 41721001, and 32170107).

We are grateful to Luying Xun (Shandong University, China) for providing C. pinatubonensis JMP134 and Haichun Gao (Zhejiang University, China) for the gift of plasmid pHGE used in this study.

Footnotes

Supplemental material is available online only.

Supplemental file 1

Fig. S1 to S7 and Tables S1 to S2. Download aem.01880-21-s0001.pdf, PDF file, 0.8 MB^{(843.8KB, pdf)}

Contributor Information

Zhenmei Lu, Email: lzhenmei@zju.edu.cn.

Jeremy D. Semrau, University of Michigan-Ann Arbor

REFERENCES

1.Li Y, Yu BA. 2010. Glycosylation initiated cationic ring-opening polymerization of tetrahydrofuran to prepare neo-glycopolymers. Chem Commun (Camb) 46:6060–6062. 10.1039/c0cc00566e. [DOI] [PubMed] [Google Scholar]
2.Liu JX, Yan YJ, Yan YG, Zhang J. 2019. Tetrahydrofuran (THF)-mediated structure of THF• (H₂O)n_=1-10: a computational study on the formation of the THF hydrate. Crystals 9:73. 10.3390/cryst9020073. [DOI] [Google Scholar]
3.Rabek JF, Skowronski TA, Ranby B. 1980. Studies on photodegradation of poly(vinyl-chloride) accelerated by α-hydroperoxy-tetrahydrofuran. Polymer 21:226–229. 10.1016/0032-3861(80)90065-8. [DOI] [Google Scholar]
4.Isaacson C, Mohr TK, Field JA. 2006. Quantitative determination of 1,4-dioxane and tetrahydrofuran in groundwater by solid phase extraction GC/MS/MS. Environ Sci Technol 40:7305–7311. 10.1021/es0615270. [DOI] [PubMed] [Google Scholar]
5.Ikehata K, Wang-Staley L, Qu XY, Li Y. 2016. Treatment of groundwater contaminated with 1,4-dioxane, tetrahydrofuran, and chlorinated volatile organic compounds using advanced oxidation processes. Ozone Sci Eng 38:413–424. 10.1080/01919512.2016.1198686. [DOI] [Google Scholar]
6.Gamer AO, Jaeckh R, Leibold E, Kaufmann W, Gembardt C, Bahnemann R, van Ravenzwaay B. 2002. Investigations on cell proliferation and enzyme induction in male rat kidney and female mouse liver caused by tetrahydrofuran. Toxicol Sci 70:140–149. 10.1093/toxsci/70.1.140. [DOI] [PubMed] [Google Scholar]
7.Chhabra RS, Herbert RA, Roycroft JH, Chou B, Miller RA, Renne RA. 1998. Carcinogenesis studies of tetrahydrofuran vapors in rats and mice. Toxicol Sci 41:183–188. 10.1006/toxs.1997.2399. [DOI] [PubMed] [Google Scholar]
8.Hermida SAS, Possari EPM, Souza DB, Campos IPD, Gomes OF, Di Mascio P, Medeiros MHG, Loureiro APM. 2006. 2′-Deoxyguanosine, 2′-deoxycytidine, and 2′-deoxyadenosine adducts resulting from the reaction of tetrahydrofuran with DNA bases. Chem Res Toxicol 19:927–936. 10.1021/tx060033d. [DOI] [PubMed] [Google Scholar]
9.Malley LA, Christoph GR, Stadler JC, Hansen JF, Biesemeier JA, Jasti SL. 2001. Acute and subchronic neurotoxicological evaluation of tetrahydrofuran by inhalation in rats. Drug Chem Toxicol 24:201–219. 10.1081/dct-100103720. [DOI] [PubMed] [Google Scholar]
10.Yao YL, Lv ZM, Min H, Lv ZH, Jiao HP. 2009. Isolation, identification and characterization of a novel Rhodococcus sp. strain in biodegradation of tetrahydrofuran and its medium optimization using sequential statistics-based experimental designs. Bioresour Technol 100:2762–2769. 10.1016/j.biortech.2009.01.006. [DOI] [PubMed] [Google Scholar]
11.Tajima T, Hayashida N, Matsumura R, Omura A, Nakashimada Y, Kato J. 2012. Isolation and characterization of tetrahydrofuran-degrading Rhodococcus aetherivorans strain M8. Process Biochem 47:1665–1669. 10.1016/j.procbio.2011.08.009. [DOI] [Google Scholar]
12.Inoue D, Tsunoda T, Sawada K, Yamamoto N, Saito Y, Sei K, Ike M. 2016. 1,4-Dioxane degradation potential of members of the genera Pseudonocardia and Rhodococcus. Biodegradation 27:277–286. 10.1007/s10532-016-9772-7. [DOI] [PubMed] [Google Scholar]
13.Kim YM, Jeon JR, Murugesan K, Kim EJ, Chang YS. 2009. Biodegradation of 1,4-dioxane and transformation of related cyclic compounds by a newly isolated Mycobacterium sp. PH-06. Biodegradation 20:511–519. 10.1007/s10532-008-9240-0. [DOI] [PubMed] [Google Scholar]
14.Chen JM, Zhou YY, Chen DZ, Jin XJ. 2010. A newly isolated strain capable of effectively degrading tetrahydrofuran and its performance in a continuous flow system. Bioresour Technol 101:6461–6467. 10.1016/j.biortech.2010.03.064. [DOI] [PubMed] [Google Scholar]
15.Chen DZ, Jin XJ, Chen J, Ye JX, Jiang NX, Chen JM. 2016. Intermediates and substrate interaction of 1,4-dioxane degradation by the effective metabolizer Xanthobacter flavus DT8. Int Biodeterior Biodegradation 106:133–140. 10.1016/j.ibiod.2015.09.018. [DOI] [Google Scholar]
16.Skinner K, Cuiffetti L, Hyman M. 2009. Metabolism and cometabolism of cyclic ethers by a filamentous fungus, a Graphium sp. Appl Environ Microbiol 75:5514–5522. 10.1128/AEM.00078-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
17.Bernhardt D, Diekmann H. 1991. Degradation of dioxane, tetrahydrofuran and other cyclic ethers by an environmental Rhodococcus strain. Appl Microbiol Biotechnol 36:120–123. 10.1007/BF00164711. [DOI] [PubMed] [Google Scholar]
18.Ren H, Li H, Wang H, Huang H, Lu Z. 2020. Biodegradation of tetrahydrofuran by the newly isolated filamentous fungus Pseudallescheria boydii ZM01. Microorganisms 8:1190. 10.3390/microorganisms8081190. [DOI] [PMC free article] [PubMed] [Google Scholar]
19.Moreno-Horn M, Garbe LA, Tressl R, Gorisch H. 2005. Transient accumulation of (γ)-butyrolactone during degradation of bis(4-chloro-n-butyl) ether by diethylether-grown Rhodococcus sp. strain DTB. Appl Microbiol Biotechnol 69:335–340. 10.1007/s00253-005-1984-4. [DOI] [PubMed] [Google Scholar]
20.Sales CM, Grostern A, Parales JV, Parales RE, Alvarez-Cohen L. 2013. Oxidation of the cyclic ethers 1,4-dioxane and tetrahydrofuran by a monooxygenase in two Pseudonocardia species. Appl Environ Microbiol 79:7702–7708. 10.1128/AEM.02418-13. [DOI] [PMC free article] [PubMed] [Google Scholar]
21.Masuda H, McClay K, Steffan RJ, Zylstra GJ. 2012. Biodegradation of tetrahydrofuran and 1,4-dioxane by soluble diiron monooxygenase in Pseudonocardia sp. strain ENV478. J Mol Microbiol Biotechnol 22:312–316. 10.1159/000343817. [DOI] [PubMed] [Google Scholar]
22.Thiemer B, Andreesen JR, Schrader T. 2003. Cloning and characterization of a gene cluster involved in tetrahydrofuran degradation in Pseudonocardia sp. strain K1. Arch Microbiol 179:266–277. 10.1007/s00203-003-0526-7. [DOI] [PubMed] [Google Scholar]
23.Qi MB, Huang H, Zhang Y, Wang HX, Li HB, Lu ZM. 2019. Novel tetrahydrofuran (THF) degradation-associated genes and cooperation patterns of a THF-degrading microbial community as revealed by metagenomic. Chemosphere 231:173–183. 10.1016/j.chemosphere.2019.05.137. [DOI] [PubMed] [Google Scholar]
24.He Y, Mathieu J, Yang Y, Yu PF, da Silva MLB, Alvarez PJJ. 2017. 1,4-Dioxane biodegradation by Mycobacterium dioxanotrophicus PH-06 is associated with a Group-6 soluble di-iron monooxygenase. Environ Sci Technol Lett 4:494–499. 10.1021/acs.estlett.7b00456. [DOI] [Google Scholar]
25.Deng DY, Li F, Li MY. 2018. A novel propane monooxygenase initiating degradation of 1,4-dioxane by Mycobacterium dioxanotrophicus PH-06. Environ Sci Technol Lett 5:86–91. 10.1021/acs.estlett.7b00504. [DOI] [Google Scholar]
26.Li F, Deng DY, Li MY. 2020. Distinct catalytic behaviors between two 1,4-dioxane-degrading monooxygenases: kinetics, inhibition, and substrate range. Environ Sci Technol 54:1898–1908. 10.1021/acs.est.9b05671. [DOI] [PubMed] [Google Scholar]
27.Mazhar SH, Herzberg M, Ben Fekih I, Zhang CK, Bello SK, Li YP, Su JM, Xu JQ, Feng RW, Zhou SG, Rensing C. 2020. Comparative insights into the complete genome sequence of highly metal-resistant Cupriavidus metallidurans strain BS1 isolated from a gold-copper mine. Front Microbiol 11:47. 10.3389/fmicb.2020.00047. [DOI] [PMC free article] [PubMed] [Google Scholar]
28.Zhao XQ, Wang RC, Lu XC, Lu JJ, Li J, Hu H. 2012. Tolerance and biosorption of heavy metals by Cupriavidus metallidurans strain XXKD-1 isolated from a subsurface laneway in the Qixiashan Pb-Zn sulfide minery in eastern China. Geomicrobiol J 29:274–286. 10.1080/01490451.2011.619637. [DOI] [Google Scholar]
29.Estrada-de los Santos P, Martinez-Aguilar L, Lopez-Lara IM, Caballero-Mellado J. 2012. Cupriavidus alkaliphilus sp. nov., a new species associated with agricultural plants that grow in alkaline soils. Syst Appl Microbiol 35:310–314. 10.1016/j.syapm.2012.05.005. [DOI] [PubMed] [Google Scholar]
30.Notomista E, Lahm A, Di Donato A, Tramontano A. 2003. Evolution of bacterial and archaeal multicomponent monooxygenases. J Mol Evol 56:435–445. 10.1007/s00239-002-2414-1. [DOI] [PubMed] [Google Scholar]
31.van Beilen JB, Funhoff EG. 2007. Alkane hydroxylases involved in microbial alkane degradation. Appl Microbiol Biotechnol 74:13–21. 10.1007/s00253-006-0748-0. [DOI] [PubMed] [Google Scholar]
32.Zhou NY, Jenkins A, Chan Kwo Chion CK, Leak DJ. 1999. The alkene monooxygenase from Xanthobacter strain Py2 is closely related to aromatic monooxygenases and catalyzes aromatic monohydroxylation of benzene, toluene, and phenol. Appl Environ Microbiol 65:1589–1595. 10.1128/AEM.65.4.1589-1595.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]
33.Tinberg CE, Song WJ, Izzo V, Lippard SJ. 2011. Multiple roles of component proteins in bacterial multicomponent monooxygenases: phenol hydroxylase and toluene/o-xylene monooxygenase from Pseudomonas sp. OX1. Biochemistry-US 50:1788–1798. 10.1021/bi200028z. [DOI] [PMC free article] [PubMed] [Google Scholar]
34.Coleman NV, Bui NB, Holmes AJ. 2006. Soluble di-iron monooxygenase gene diversity in soils, sediments and ethene enrichments. Environ Microbiol 8:1228–1239. 10.1111/j.1462-2920.2006.01015.x. [DOI] [PubMed] [Google Scholar]
35.He Y, Mathieu J, da Silva MLB, Li MY, Alvarez PJJ. 2018. 1,4-Dioxane-degrading consortia can be enriched from uncontaminated soils: prevalence of Mycobacterium and soluble di-iron monooxygenase genes. Microb Biotechnol 11:189–198. 10.1111/1751-7915.12850. [DOI] [PMC free article] [PubMed] [Google Scholar]
36.Coleman NV, Yau S, Wilson NL, Nolan LM, Migocki MD, Ly MA, Crossett B, Holmes AJ. 2011. Untangling the multiple monooxygenases of Mycobacterium chubuense strain NBB4, a versatile hydrocarbon degrader. Environ Microbiol Rep 3:297–307. 10.1111/j.1758-2229.2010.00225.x. [DOI] [PubMed] [Google Scholar]
37.Sales CM, Mahendra S, Grostern A, Parales RE, Goodwin LA, Woyke T, Nolan M, Lapidus A, Chertkov O, Ovchinnikova G, Sczyrba A, Alvarez-Cohen L. 2011. Genome sequence of the 1,4-dioxane-degrading Pseudonocardia dioxanivorans strain CB1190. J Bacteriol 193:4549–4550. 10.1128/JB.00415-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
38.Deng D, Pham DN, Li F, Li M. 2020. Discovery of an inducible toluene monooxygenase that cooxidizes 1,4-dioxane and 1,1-dichloroethylene in propanotrophic Azoarcus sp. strain DD4. Appl Environ Microbiol 86:e01163-20. 10.1128/AEM.01163-20. [DOI] [PMC free article] [PubMed] [Google Scholar]
39.Harzallah B, Bousseboua H, Jouanneau Y. 2017. Diversity shift in bacterial phenol hydroxylases driven by alkyl-phenols in oil refinery wastewaters. Environ Sci Pollut Res Int 24:14376–14386. 10.1007/s11356-017-8950-4. [DOI] [PubMed] [Google Scholar]
40.Xu Y, Chen M, Zhang W, Lin M. 2003. Genetic organization of genes encoding phenol hydroxylase, benzoate 1,2-dioxygenase alpha subunit and its regulatory proteins in Acinetobacter calcoaceticus PHEA-2. Curr Microbiol 46:235–240. 10.1007/s00284-002-3840-4. [DOI] [PubMed] [Google Scholar]
41.Nordlund I, Powlowski J, Shingler V. 1990. Complete nucleotide sequence and polypeptide analysis of multicomponent phenol hydroxylase from Pseudomonas sp. strain CF600. J Bacteriol 172:6826–6833. 10.1128/jb.172.12.6826-6833.1990. [DOI] [PMC free article] [PubMed] [Google Scholar]
42.Schirmer F, Hillen W. 1998. The Acinetobacter calcoaceticus NCIB8250 mop operon mRNA is differentially degraded, resulting in a higher level of the 3′ CatA-encoding segment than of the 5′ phenolhydroxylase-encoding portion. Mol Gen Genet 257:330–337. 10.1007/pl00008621. [DOI] [PubMed] [Google Scholar]
43.Santos PM, Sa-Correia I. 2007. Characterization of the unique organization and coregulation of a gene cluster required for phenol and benzene catabolism in Pseudomonas sp. M1. J Biotechnol 131:371–378. 10.1016/j.jbiotec.2007.07.941. [DOI] [PubMed] [Google Scholar]
44.Teramoto M, Futamata H, Harayama S, Watanabe K. 1999. Characterization of a high-affinity phenol hydroxylase from Comamonas testosteroni R5 by gene cloning, and expression in Pseudomonas aeruginosa PAO1c. Mol Gen Genet 262:552–558. 10.1007/s004380051117. [DOI] [PubMed] [Google Scholar]
45.Zhu CG, Zhang LY, Zhao LP. 2008. Molecular cloning, genetic organization of gene cluster encoding phenol hydroxylase and catechol 2,3-dioxygenase in Alcaligenes faecalis IS-46. World J Microbiol Biotechnol 24:1687–1695. 10.1007/s11274-008-9660-3. [DOI] [Google Scholar]
46.Arai H, Akahira S, Ohishi T, Maeda M, Kudo T. 1998. Adaptation of Comamonas testosteroni TA441 to utilize phenol: organization and regulation of the genes involved in phenol degradation. Microbiology 144:2895–2903. 10.1099/00221287-144-10-2895. [DOI] [PubMed] [Google Scholar]
47.Kohlweyer U, Thiemer B, Schrader T, Andreesen JR. 2000. Tetrahydrofuran degradation by a newly isolated culture of Pseudonocardia sp. strain K1. FEMS Microbiol Lett 186:301–306. 10.1111/j.1574-6968.2000.tb09121.x. [DOI] [PubMed] [Google Scholar]
48.Ma F, Shi SN, Sun TH, Li A, Zhou JT, Qu YY. 2013. Biotransformation of benzene and toluene to catechols by phenol hydroxylase from Arthrobacter sp. W1. Appl Microbiol Biotechnol 97:5097–5103. 10.1007/s00253-012-4301-z. [DOI] [PubMed] [Google Scholar]
49.Nguyen NT, Hsieh HC, Lin YW, Huang SL. 2011. Analysis of bacterial degradation pathways for long-chain alkylphenols involving phenol hydroxylase, alkylphenol monooxygenase and catechol dioxygenase genes. Bioresour Technol 102:4232–4240. 10.1016/j.biortech.2010.12.067. [DOI] [PubMed] [Google Scholar]
50.Wang J, Shen X, Wang J, Yang Y, Yuan Q, Yan Y. 2018. Exploring the promiscuity of phenol hydroxylase from Pseudomonas stutzeri OX1 for the biosynthesis of phenolic compounds. ACS Synth Biol 7:1238–1243. 10.1021/acssynbio.8b00067. [DOI] [PubMed] [Google Scholar]
51.Powlowski J, Shingler V. 1994. Genetics and biochemistry of phenol degradation by Pseudomonas sp. CF600. Biodegradation 5:219–236. 10.1007/BF00696461. [DOI] [PubMed] [Google Scholar]
52.Merimaa M, Heinaru E, Liivak M, Vedler E, Heinaru A. 2006. Grouping of phenol hydroxylase and catechol 2,3-dioxygenase genes among phenol- and p-cresol-degrading Pseudomonas species and biotypes. Arch Microbiol 186:287–296. 10.1007/s00203-006-0143-3. [DOI] [PubMed] [Google Scholar]
53.De Mey M, Lequeux GJ, Beauprez JJ, Maertens J, Van Horen E, Soetaert WK, Vanrolleghem PA, Vandamme EJ. 2007. Comparison of different strategies to reduce acetate formation in Escherichia coli. Biotechnol Prog 23:1053–1063. 10.1021/bp070170g. [DOI] [PubMed] [Google Scholar]
54.Veit A, Polen T, Wendisch VF. 2007. Global gene expression analysis of glucose overflow metabolism in Escherichia coli and reduction of aerobic acetate formation. Appl Microbiol Biotechnol 74:406–421. 10.1007/s00253-006-0680-3. [DOI] [PubMed] [Google Scholar]
55.Cafaro V, Izzo V, Scognamiglio R, Notomista E, Capasso P, Casbarra A, Pucci P, Di Donato A. 2004. Phenol hydroxylase and toluene/o-xylene monooxygenase from Pseudomonas stutzeri OX1: interplay between two enzymes. Appl Environ Microbiol 70:2211–2219. 10.1128/AEM.70.4.2211-2219.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
56.Liu ZB, He ZX, Huang H, Ran XB, Oluwafunmilayo AO, Lu ZM. 2017. pH Stress-induced cooperation between Rhodococcus ruber YYL and Bacillus cereus MLY1 in biodegradation of tetrahydrofuran. Front Microbiol 8:2297. 10.3389/fmicb.2017.02297. [DOI] [PMC free article] [PubMed] [Google Scholar]
57.Overbeek R, Olson R, Pusch GD, Olsen GJ, Davis JJ, Disz T, Edwards RA, Gerdes S, Parrello B, Shukla M, Vonstein V, Wattam AR, Xia FF, Stevens R. 2014. The SEED and the Rapid Annotation of microbial genomes using Subsystems Technology (RAST). Nucleic Acids Res 42:D206–D214. 10.1093/nar/gkt1226. [DOI] [PMC free article] [PubMed] [Google Scholar]
58.Zhou NY, Al-Dulayymi J, Baird MS, Williams PA. 2002. Salicylate 5-hydroxylase from Ralstonia sp. strain U2: a monooxygenase with close relationships to and shared electron transport proteins with naphthalene dioxygenase. J Bacteriol 184:1547–1555. 10.1128/JB.184.6.1547-1555.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
59.Zhou NY, Fuenmayor SL, Williams PA. 2001. Nag genes of Ralstonia (formerly Pseudomonas) sp. strain U2 encoding enzymes for gentisate catabolism. J Bacteriol 183:700–708. 10.1128/JB.183.2.700-708.2001. [DOI] [PMC free article] [PubMed] [Google Scholar]
60.Gao R, Liu HL, Xun LY. 2017. Cytoplasmic localization of sulfide: quinone oxidoreductase and persulfide dioxygenase of Cupriavidus pinatubonensis JMP134. Appl Environ Microbiol 83:e01820-17. 10.1128/AEM.01820-17. [DOI] [PMC free article] [PubMed] [Google Scholar]
61.Wang HX, Xie CX, Zhu PP, Zhou NY, Lu ZM. 2016. Two novel sets of genes essential for nicotine degradation by Sphingomonas melonis TY. Front Microbiol 7:2060. [DOI] [PMC free article] [PubMed] [Google Scholar]
62.Qiu JG, Zhang JJ, Zhang YT, Wang YH, Tong L, Hong Q, He J. 2017. Biodegradation of picolinic acid by a newly isolated bacterium Alcaligenes faecalis strain JQ135. Curr Microbiol 74:508–514. 10.1007/s00284-017-1205-2. [DOI] [PubMed] [Google Scholar]
63.Guo Y, Li DF, Ji HN, Zheng JT, Zhou NY. 2020. Hexachlorobenzene monooxygenase substrate selectivity and catalysis: structural and biochemical insights. Appl Environ Microbiol 87:e01965-20. 10.1128/AEM.01965-20. [DOI] [PMC free article] [PubMed] [Google Scholar]
64.Wang HX, Wang XY, Ren H, Wang XJ, Lu ZM. 2020. 3-Hydroxypyridine dehydrogenase HpdA is encoded by a novel four-component gene cluster and catalyzes the first step of 3-hydroxypyridine catabolism in Ensifer adhaerens HP1. Appl Environ Microbiol 86:e01313-20. 10.1128/AEM.01313-20. [DOI] [PMC free article] [PubMed] [Google Scholar]
65.Guo KL, Wang W, Wang HX, Lu ZM, Gao HC. 2019. Complex oxidation of apocytochromes c during bacterial cytochrome c maturation. Appl Environ Microbiol 85:e01989-19. 10.1128/AEM.01989-19. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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Supplementary Materials

Supplemental file 1

Fig. S1 to S7 and Tables S1 to S2. Download aem.01880-21-s0001.pdf, PDF file, 0.8 MB^{(843.8KB, pdf)}

Data Availability Statement

[B1] 1.Li Y, Yu BA. 2010. Glycosylation initiated cationic ring-opening polymerization of tetrahydrofuran to prepare neo-glycopolymers. Chem Commun (Camb) 46:6060–6062. 10.1039/c0cc00566e. [DOI] [PubMed] [Google Scholar]

[B2] 2.Liu JX, Yan YJ, Yan YG, Zhang J. 2019. Tetrahydrofuran (THF)-mediated structure of THF• (H₂O)n_=1-10: a computational study on the formation of the THF hydrate. Crystals 9:73. 10.3390/cryst9020073. [DOI] [Google Scholar]

[B3] 3.Rabek JF, Skowronski TA, Ranby B. 1980. Studies on photodegradation of poly(vinyl-chloride) accelerated by α-hydroperoxy-tetrahydrofuran. Polymer 21:226–229. 10.1016/0032-3861(80)90065-8. [DOI] [Google Scholar]

[B4] 4.Isaacson C, Mohr TK, Field JA. 2006. Quantitative determination of 1,4-dioxane and tetrahydrofuran in groundwater by solid phase extraction GC/MS/MS. Environ Sci Technol 40:7305–7311. 10.1021/es0615270. [DOI] [PubMed] [Google Scholar]

[B5] 5.Ikehata K, Wang-Staley L, Qu XY, Li Y. 2016. Treatment of groundwater contaminated with 1,4-dioxane, tetrahydrofuran, and chlorinated volatile organic compounds using advanced oxidation processes. Ozone Sci Eng 38:413–424. 10.1080/01919512.2016.1198686. [DOI] [Google Scholar]

[B6] 6.Gamer AO, Jaeckh R, Leibold E, Kaufmann W, Gembardt C, Bahnemann R, van Ravenzwaay B. 2002. Investigations on cell proliferation and enzyme induction in male rat kidney and female mouse liver caused by tetrahydrofuran. Toxicol Sci 70:140–149. 10.1093/toxsci/70.1.140. [DOI] [PubMed] [Google Scholar]

[B7] 7.Chhabra RS, Herbert RA, Roycroft JH, Chou B, Miller RA, Renne RA. 1998. Carcinogenesis studies of tetrahydrofuran vapors in rats and mice. Toxicol Sci 41:183–188. 10.1006/toxs.1997.2399. [DOI] [PubMed] [Google Scholar]

[B8] 8.Hermida SAS, Possari EPM, Souza DB, Campos IPD, Gomes OF, Di Mascio P, Medeiros MHG, Loureiro APM. 2006. 2′-Deoxyguanosine, 2′-deoxycytidine, and 2′-deoxyadenosine adducts resulting from the reaction of tetrahydrofuran with DNA bases. Chem Res Toxicol 19:927–936. 10.1021/tx060033d. [DOI] [PubMed] [Google Scholar]

[B9] 9.Malley LA, Christoph GR, Stadler JC, Hansen JF, Biesemeier JA, Jasti SL. 2001. Acute and subchronic neurotoxicological evaluation of tetrahydrofuran by inhalation in rats. Drug Chem Toxicol 24:201–219. 10.1081/dct-100103720. [DOI] [PubMed] [Google Scholar]

[B10] 10.Yao YL, Lv ZM, Min H, Lv ZH, Jiao HP. 2009. Isolation, identification and characterization of a novel Rhodococcus sp. strain in biodegradation of tetrahydrofuran and its medium optimization using sequential statistics-based experimental designs. Bioresour Technol 100:2762–2769. 10.1016/j.biortech.2009.01.006. [DOI] [PubMed] [Google Scholar]

[B11] 11.Tajima T, Hayashida N, Matsumura R, Omura A, Nakashimada Y, Kato J. 2012. Isolation and characterization of tetrahydrofuran-degrading Rhodococcus aetherivorans strain M8. Process Biochem 47:1665–1669. 10.1016/j.procbio.2011.08.009. [DOI] [Google Scholar]

[B12] 12.Inoue D, Tsunoda T, Sawada K, Yamamoto N, Saito Y, Sei K, Ike M. 2016. 1,4-Dioxane degradation potential of members of the genera Pseudonocardia and Rhodococcus. Biodegradation 27:277–286. 10.1007/s10532-016-9772-7. [DOI] [PubMed] [Google Scholar]

[B13] 13.Kim YM, Jeon JR, Murugesan K, Kim EJ, Chang YS. 2009. Biodegradation of 1,4-dioxane and transformation of related cyclic compounds by a newly isolated Mycobacterium sp. PH-06. Biodegradation 20:511–519. 10.1007/s10532-008-9240-0. [DOI] [PubMed] [Google Scholar]

[B14] 14.Chen JM, Zhou YY, Chen DZ, Jin XJ. 2010. A newly isolated strain capable of effectively degrading tetrahydrofuran and its performance in a continuous flow system. Bioresour Technol 101:6461–6467. 10.1016/j.biortech.2010.03.064. [DOI] [PubMed] [Google Scholar]

[B15] 15.Chen DZ, Jin XJ, Chen J, Ye JX, Jiang NX, Chen JM. 2016. Intermediates and substrate interaction of 1,4-dioxane degradation by the effective metabolizer Xanthobacter flavus DT8. Int Biodeterior Biodegradation 106:133–140. 10.1016/j.ibiod.2015.09.018. [DOI] [Google Scholar]

[B16] 16.Skinner K, Cuiffetti L, Hyman M. 2009. Metabolism and cometabolism of cyclic ethers by a filamentous fungus, a Graphium sp. Appl Environ Microbiol 75:5514–5522. 10.1128/AEM.00078-09. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B17] 17.Bernhardt D, Diekmann H. 1991. Degradation of dioxane, tetrahydrofuran and other cyclic ethers by an environmental Rhodococcus strain. Appl Microbiol Biotechnol 36:120–123. 10.1007/BF00164711. [DOI] [PubMed] [Google Scholar]

[B18] 18.Ren H, Li H, Wang H, Huang H, Lu Z. 2020. Biodegradation of tetrahydrofuran by the newly isolated filamentous fungus Pseudallescheria boydii ZM01. Microorganisms 8:1190. 10.3390/microorganisms8081190. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B19] 19.Moreno-Horn M, Garbe LA, Tressl R, Gorisch H. 2005. Transient accumulation of (γ)-butyrolactone during degradation of bis(4-chloro-n-butyl) ether by diethylether-grown Rhodococcus sp. strain DTB. Appl Microbiol Biotechnol 69:335–340. 10.1007/s00253-005-1984-4. [DOI] [PubMed] [Google Scholar]

[B20] 20.Sales CM, Grostern A, Parales JV, Parales RE, Alvarez-Cohen L. 2013. Oxidation of the cyclic ethers 1,4-dioxane and tetrahydrofuran by a monooxygenase in two Pseudonocardia species. Appl Environ Microbiol 79:7702–7708. 10.1128/AEM.02418-13. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B21] 21.Masuda H, McClay K, Steffan RJ, Zylstra GJ. 2012. Biodegradation of tetrahydrofuran and 1,4-dioxane by soluble diiron monooxygenase in Pseudonocardia sp. strain ENV478. J Mol Microbiol Biotechnol 22:312–316. 10.1159/000343817. [DOI] [PubMed] [Google Scholar]

[B22] 22.Thiemer B, Andreesen JR, Schrader T. 2003. Cloning and characterization of a gene cluster involved in tetrahydrofuran degradation in Pseudonocardia sp. strain K1. Arch Microbiol 179:266–277. 10.1007/s00203-003-0526-7. [DOI] [PubMed] [Google Scholar]

[B23] 23.Qi MB, Huang H, Zhang Y, Wang HX, Li HB, Lu ZM. 2019. Novel tetrahydrofuran (THF) degradation-associated genes and cooperation patterns of a THF-degrading microbial community as revealed by metagenomic. Chemosphere 231:173–183. 10.1016/j.chemosphere.2019.05.137. [DOI] [PubMed] [Google Scholar]

[B24] 24.He Y, Mathieu J, Yang Y, Yu PF, da Silva MLB, Alvarez PJJ. 2017. 1,4-Dioxane biodegradation by Mycobacterium dioxanotrophicus PH-06 is associated with a Group-6 soluble di-iron monooxygenase. Environ Sci Technol Lett 4:494–499. 10.1021/acs.estlett.7b00456. [DOI] [Google Scholar]

[B25] 25.Deng DY, Li F, Li MY. 2018. A novel propane monooxygenase initiating degradation of 1,4-dioxane by Mycobacterium dioxanotrophicus PH-06. Environ Sci Technol Lett 5:86–91. 10.1021/acs.estlett.7b00504. [DOI] [Google Scholar]

[B26] 26.Li F, Deng DY, Li MY. 2020. Distinct catalytic behaviors between two 1,4-dioxane-degrading monooxygenases: kinetics, inhibition, and substrate range. Environ Sci Technol 54:1898–1908. 10.1021/acs.est.9b05671. [DOI] [PubMed] [Google Scholar]

[B27] 27.Mazhar SH, Herzberg M, Ben Fekih I, Zhang CK, Bello SK, Li YP, Su JM, Xu JQ, Feng RW, Zhou SG, Rensing C. 2020. Comparative insights into the complete genome sequence of highly metal-resistant Cupriavidus metallidurans strain BS1 isolated from a gold-copper mine. Front Microbiol 11:47. 10.3389/fmicb.2020.00047. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B28] 28.Zhao XQ, Wang RC, Lu XC, Lu JJ, Li J, Hu H. 2012. Tolerance and biosorption of heavy metals by Cupriavidus metallidurans strain XXKD-1 isolated from a subsurface laneway in the Qixiashan Pb-Zn sulfide minery in eastern China. Geomicrobiol J 29:274–286. 10.1080/01490451.2011.619637. [DOI] [Google Scholar]

[B29] 29.Estrada-de los Santos P, Martinez-Aguilar L, Lopez-Lara IM, Caballero-Mellado J. 2012. Cupriavidus alkaliphilus sp. nov., a new species associated with agricultural plants that grow in alkaline soils. Syst Appl Microbiol 35:310–314. 10.1016/j.syapm.2012.05.005. [DOI] [PubMed] [Google Scholar]

[B30] 30.Notomista E, Lahm A, Di Donato A, Tramontano A. 2003. Evolution of bacterial and archaeal multicomponent monooxygenases. J Mol Evol 56:435–445. 10.1007/s00239-002-2414-1. [DOI] [PubMed] [Google Scholar]

[B31] 31.van Beilen JB, Funhoff EG. 2007. Alkane hydroxylases involved in microbial alkane degradation. Appl Microbiol Biotechnol 74:13–21. 10.1007/s00253-006-0748-0. [DOI] [PubMed] [Google Scholar]

[B32] 32.Zhou NY, Jenkins A, Chan Kwo Chion CK, Leak DJ. 1999. The alkene monooxygenase from Xanthobacter strain Py2 is closely related to aromatic monooxygenases and catalyzes aromatic monohydroxylation of benzene, toluene, and phenol. Appl Environ Microbiol 65:1589–1595. 10.1128/AEM.65.4.1589-1595.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B33] 33.Tinberg CE, Song WJ, Izzo V, Lippard SJ. 2011. Multiple roles of component proteins in bacterial multicomponent monooxygenases: phenol hydroxylase and toluene/o-xylene monooxygenase from Pseudomonas sp. OX1. Biochemistry-US 50:1788–1798. 10.1021/bi200028z. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B34] 34.Coleman NV, Bui NB, Holmes AJ. 2006. Soluble di-iron monooxygenase gene diversity in soils, sediments and ethene enrichments. Environ Microbiol 8:1228–1239. 10.1111/j.1462-2920.2006.01015.x. [DOI] [PubMed] [Google Scholar]

[B35] 35.He Y, Mathieu J, da Silva MLB, Li MY, Alvarez PJJ. 2018. 1,4-Dioxane-degrading consortia can be enriched from uncontaminated soils: prevalence of Mycobacterium and soluble di-iron monooxygenase genes. Microb Biotechnol 11:189–198. 10.1111/1751-7915.12850. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B36] 36.Coleman NV, Yau S, Wilson NL, Nolan LM, Migocki MD, Ly MA, Crossett B, Holmes AJ. 2011. Untangling the multiple monooxygenases of Mycobacterium chubuense strain NBB4, a versatile hydrocarbon degrader. Environ Microbiol Rep 3:297–307. 10.1111/j.1758-2229.2010.00225.x. [DOI] [PubMed] [Google Scholar]

[B37] 37.Sales CM, Mahendra S, Grostern A, Parales RE, Goodwin LA, Woyke T, Nolan M, Lapidus A, Chertkov O, Ovchinnikova G, Sczyrba A, Alvarez-Cohen L. 2011. Genome sequence of the 1,4-dioxane-degrading Pseudonocardia dioxanivorans strain CB1190. J Bacteriol 193:4549–4550. 10.1128/JB.00415-11. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B38] 38.Deng D, Pham DN, Li F, Li M. 2020. Discovery of an inducible toluene monooxygenase that cooxidizes 1,4-dioxane and 1,1-dichloroethylene in propanotrophic Azoarcus sp. strain DD4. Appl Environ Microbiol 86:e01163-20. 10.1128/AEM.01163-20. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B39] 39.Harzallah B, Bousseboua H, Jouanneau Y. 2017. Diversity shift in bacterial phenol hydroxylases driven by alkyl-phenols in oil refinery wastewaters. Environ Sci Pollut Res Int 24:14376–14386. 10.1007/s11356-017-8950-4. [DOI] [PubMed] [Google Scholar]

[B40] 40.Xu Y, Chen M, Zhang W, Lin M. 2003. Genetic organization of genes encoding phenol hydroxylase, benzoate 1,2-dioxygenase alpha subunit and its regulatory proteins in Acinetobacter calcoaceticus PHEA-2. Curr Microbiol 46:235–240. 10.1007/s00284-002-3840-4. [DOI] [PubMed] [Google Scholar]

[B41] 41.Nordlund I, Powlowski J, Shingler V. 1990. Complete nucleotide sequence and polypeptide analysis of multicomponent phenol hydroxylase from Pseudomonas sp. strain CF600. J Bacteriol 172:6826–6833. 10.1128/jb.172.12.6826-6833.1990. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B42] 42.Schirmer F, Hillen W. 1998. The Acinetobacter calcoaceticus NCIB8250 mop operon mRNA is differentially degraded, resulting in a higher level of the 3′ CatA-encoding segment than of the 5′ phenolhydroxylase-encoding portion. Mol Gen Genet 257:330–337. 10.1007/pl00008621. [DOI] [PubMed] [Google Scholar]

[B43] 43.Santos PM, Sa-Correia I. 2007. Characterization of the unique organization and coregulation of a gene cluster required for phenol and benzene catabolism in Pseudomonas sp. M1. J Biotechnol 131:371–378. 10.1016/j.jbiotec.2007.07.941. [DOI] [PubMed] [Google Scholar]

[B44] 44.Teramoto M, Futamata H, Harayama S, Watanabe K. 1999. Characterization of a high-affinity phenol hydroxylase from Comamonas testosteroni R5 by gene cloning, and expression in Pseudomonas aeruginosa PAO1c. Mol Gen Genet 262:552–558. 10.1007/s004380051117. [DOI] [PubMed] [Google Scholar]

[B45] 45.Zhu CG, Zhang LY, Zhao LP. 2008. Molecular cloning, genetic organization of gene cluster encoding phenol hydroxylase and catechol 2,3-dioxygenase in Alcaligenes faecalis IS-46. World J Microbiol Biotechnol 24:1687–1695. 10.1007/s11274-008-9660-3. [DOI] [Google Scholar]

[B46] 46.Arai H, Akahira S, Ohishi T, Maeda M, Kudo T. 1998. Adaptation of Comamonas testosteroni TA441 to utilize phenol: organization and regulation of the genes involved in phenol degradation. Microbiology 144:2895–2903. 10.1099/00221287-144-10-2895. [DOI] [PubMed] [Google Scholar]

[B47] 47.Kohlweyer U, Thiemer B, Schrader T, Andreesen JR. 2000. Tetrahydrofuran degradation by a newly isolated culture of Pseudonocardia sp. strain K1. FEMS Microbiol Lett 186:301–306. 10.1111/j.1574-6968.2000.tb09121.x. [DOI] [PubMed] [Google Scholar]

[B48] 48.Ma F, Shi SN, Sun TH, Li A, Zhou JT, Qu YY. 2013. Biotransformation of benzene and toluene to catechols by phenol hydroxylase from Arthrobacter sp. W1. Appl Microbiol Biotechnol 97:5097–5103. 10.1007/s00253-012-4301-z. [DOI] [PubMed] [Google Scholar]

[B49] 49.Nguyen NT, Hsieh HC, Lin YW, Huang SL. 2011. Analysis of bacterial degradation pathways for long-chain alkylphenols involving phenol hydroxylase, alkylphenol monooxygenase and catechol dioxygenase genes. Bioresour Technol 102:4232–4240. 10.1016/j.biortech.2010.12.067. [DOI] [PubMed] [Google Scholar]

[B50] 50.Wang J, Shen X, Wang J, Yang Y, Yuan Q, Yan Y. 2018. Exploring the promiscuity of phenol hydroxylase from Pseudomonas stutzeri OX1 for the biosynthesis of phenolic compounds. ACS Synth Biol 7:1238–1243. 10.1021/acssynbio.8b00067. [DOI] [PubMed] [Google Scholar]

[B51] 51.Powlowski J, Shingler V. 1994. Genetics and biochemistry of phenol degradation by Pseudomonas sp. CF600. Biodegradation 5:219–236. 10.1007/BF00696461. [DOI] [PubMed] [Google Scholar]

[B52] 52.Merimaa M, Heinaru E, Liivak M, Vedler E, Heinaru A. 2006. Grouping of phenol hydroxylase and catechol 2,3-dioxygenase genes among phenol- and p-cresol-degrading Pseudomonas species and biotypes. Arch Microbiol 186:287–296. 10.1007/s00203-006-0143-3. [DOI] [PubMed] [Google Scholar]

[B53] 53.De Mey M, Lequeux GJ, Beauprez JJ, Maertens J, Van Horen E, Soetaert WK, Vanrolleghem PA, Vandamme EJ. 2007. Comparison of different strategies to reduce acetate formation in Escherichia coli. Biotechnol Prog 23:1053–1063. 10.1021/bp070170g. [DOI] [PubMed] [Google Scholar]

[B54] 54.Veit A, Polen T, Wendisch VF. 2007. Global gene expression analysis of glucose overflow metabolism in Escherichia coli and reduction of aerobic acetate formation. Appl Microbiol Biotechnol 74:406–421. 10.1007/s00253-006-0680-3. [DOI] [PubMed] [Google Scholar]

[B55] 55.Cafaro V, Izzo V, Scognamiglio R, Notomista E, Capasso P, Casbarra A, Pucci P, Di Donato A. 2004. Phenol hydroxylase and toluene/o-xylene monooxygenase from Pseudomonas stutzeri OX1: interplay between two enzymes. Appl Environ Microbiol 70:2211–2219. 10.1128/AEM.70.4.2211-2219.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B56] 56.Liu ZB, He ZX, Huang H, Ran XB, Oluwafunmilayo AO, Lu ZM. 2017. pH Stress-induced cooperation between Rhodococcus ruber YYL and Bacillus cereus MLY1 in biodegradation of tetrahydrofuran. Front Microbiol 8:2297. 10.3389/fmicb.2017.02297. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B57] 57.Overbeek R, Olson R, Pusch GD, Olsen GJ, Davis JJ, Disz T, Edwards RA, Gerdes S, Parrello B, Shukla M, Vonstein V, Wattam AR, Xia FF, Stevens R. 2014. The SEED and the Rapid Annotation of microbial genomes using Subsystems Technology (RAST). Nucleic Acids Res 42:D206–D214. 10.1093/nar/gkt1226. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B58] 58.Zhou NY, Al-Dulayymi J, Baird MS, Williams PA. 2002. Salicylate 5-hydroxylase from Ralstonia sp. strain U2: a monooxygenase with close relationships to and shared electron transport proteins with naphthalene dioxygenase. J Bacteriol 184:1547–1555. 10.1128/JB.184.6.1547-1555.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B59] 59.Zhou NY, Fuenmayor SL, Williams PA. 2001. Nag genes of Ralstonia (formerly Pseudomonas) sp. strain U2 encoding enzymes for gentisate catabolism. J Bacteriol 183:700–708. 10.1128/JB.183.2.700-708.2001. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B60] 60.Gao R, Liu HL, Xun LY. 2017. Cytoplasmic localization of sulfide: quinone oxidoreductase and persulfide dioxygenase of Cupriavidus pinatubonensis JMP134. Appl Environ Microbiol 83:e01820-17. 10.1128/AEM.01820-17. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B61] 61.Wang HX, Xie CX, Zhu PP, Zhou NY, Lu ZM. 2016. Two novel sets of genes essential for nicotine degradation by Sphingomonas melonis TY. Front Microbiol 7:2060. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B62] 62.Qiu JG, Zhang JJ, Zhang YT, Wang YH, Tong L, Hong Q, He J. 2017. Biodegradation of picolinic acid by a newly isolated bacterium Alcaligenes faecalis strain JQ135. Curr Microbiol 74:508–514. 10.1007/s00284-017-1205-2. [DOI] [PubMed] [Google Scholar]

[B63] 63.Guo Y, Li DF, Ji HN, Zheng JT, Zhou NY. 2020. Hexachlorobenzene monooxygenase substrate selectivity and catalysis: structural and biochemical insights. Appl Environ Microbiol 87:e01965-20. 10.1128/AEM.01965-20. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B64] 64.Wang HX, Wang XY, Ren H, Wang XJ, Lu ZM. 2020. 3-Hydroxypyridine dehydrogenase HpdA is encoded by a novel four-component gene cluster and catalyzes the first step of 3-hydroxypyridine catabolism in Ensifer adhaerens HP1. Appl Environ Microbiol 86:e01313-20. 10.1128/AEM.01313-20. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B65] 65.Guo KL, Wang W, Wang HX, Lu ZM, Gao HC. 2019. Complex oxidation of apocytochromes c during bacterial cytochrome c maturation. Appl Environ Microbiol 85:e01989-19. 10.1128/AEM.01989-19. [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

A Multicomponent THF Hydroxylase Initiates Tetrahydrofuran Degradation in Cupriavidus metallidurans ZM02

Hao Ren

Haixia Wang

Yang Wang

Yiyang Chen

Zhenmei Lu

Roles

ABSTRACT

INTRODUCTION

RESULTS

Isolation of the THF-degrading bacterium and detection of metabolites during THF degradation.

FIG 1.

FIG 2.

Characterization of the putative THF hydroxylase gene cluster in C. metallidurans ZM02.

FIG 3.

FIG 4.

The expression of THF hydroxylase is upregulated during growth on THF and phenol.

FIG 5.

Deletion of the THF hydroxylase gene inactivates the degradation of both THF and phenol in strain ZM02.

TABLE 1.

The THF hydroxylase encoded by dmpKLMNOP can oxidize THF.

FIG 6.

THF and phenol oxidation activities of THF hydroxylase were detected by crude enzyme assays.

TABLE 2.

TABLE 3.

FIG 7.

The representative phenol-degrading bacteria containing similar THF-degrading gene clusters cannot degrade THF alone.

FIG 8.

DISCUSSION

MATERIALS AND METHODS

Bacterial strains, culture conditions, and chemicals.

TABLE 4.

TABLE 5.

Isolation of THF-degrading bacteria and detection of THF metabolic intermediates.

Genome and transcriptome sequencing.

Reverse transcription-quantitative PCR.

Gene knockout experiments.

Cloning and heterologous expression of the putative THF-degrading gene cluster.

Preparation of cell extracts and crude enzyme assays.

Data availability.

ACKNOWLEDGMENTS

Footnotes

Contributor Information

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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