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. 2026 Aug 6;92(9):e01277-26. doi: 10.1128/aem.01277-26

The PucR-type activator IppR regulates 2-isopropylphenol hydroxylation in strain Rhodococcus sp. D-6

Qian Zhu 1, Kangning Wei 1, Yike Lyu 1, Qi Xie 1, Kaihua Pan 1, Qian Li 1, Weihao Zhu 1, Zhijian Ke 2,3, Jiguo Qiu 1, Mingliang Zhang 1, Junqiang Hu 1, Shang Dai 1,✉, Qing Hong 1,✉
Editor: Marina Lotti4
PMCID: PMC13599513  PMID: 42560057

ABSTRACT

Although genes involved in isoprocarb (IPC) degradation, including the IPC hydrolase gene ipcH and the 2-isopropylphenol (IPP) hydroxylation gene cluster ippA1A2, have been identified in Rhodococcus sp. D-6, the transcriptional regulatory mechanism remains unclear. In this study, we characterized a PucR-type transcriptional regulator, IppR, which controls the inducible expression of the ippA1A2 cluster. Gene disruption and complementation analyses revealed that ippR was essential for activation of ippA1A2 expression in response to IPP in strain D-6. Quantitative reverse transcription-PCR (qRT-PCR) and promoter activity assays demonstrated that IPP was the effector molecule of IppR. Electrophoretic mobility shift assay (EMSA) showed that IppR bound to a 24-bp motif in the promoter region, with the CC/CG box within this motif identified as the critical site for IppR binding. Moreover, IppR residues likely involved in IPP binding include Val188, Leu192, Val218, Trp257, Trp259, His297, and Ala300, along with Lys382 and Thr384, which are predicted to be essential for DNA recognition. This study elucidates the regulatory mechanism of IPC degradation in strain D-6, and the identification of IppR expands the diversity of regulatory models within the PucR-type regulator.

IMPORTANCE

Carbamate insecticides pose a potential threat to the environment and human health. Bacteria play an important role in the biodegradation of these compounds. Rhodococcus sp. D-6, which is capable of degrading isoprocarb (IPC), a representative carbamate insecticide, was isolated in our previous study. The expression of the IPC hydrolase gene ipcH was constitutive, while the expression of the ippA1A2 cluster responsible for hydroxylation of 2-isopropylphenol (IPP), the hydrolysis product of IPC, was inducible. The present study functionally characterized the PucR-type transcriptional regulator IppR, which activates the transcription of the ippA1A2 cluster to mediate IPP hydroxylation in strain D-6. This study advances our understanding of the regulatory mechanisms underlying IPC degradation.

KEYWORDS: Rhodococcus sp. D-6, isoprocarb degradation, two-component 2-isopropylphenol monooxygenase gene ippA1A2, transcriptional regulation, PucR-type regulator

INTRODUCTION

Isoprocarb [2-(1-methylethyl)-phenol methylcarbamate, IPC] is a representative carbamate insecticide widely used for pest control in agriculture and forestry (1, 2). Due to its extensive application and chemical stability, IPC residues have been detected in various environments (3–5). IPC exerts its insecticidal activity by inhibiting the activity of acetylcholinesterase, a key enzyme involved in neural signal transmission (6). Therefore, residual IPC in the environment poses potential hazards to humans and non-target organisms (7–10). IPC has been classified as a moderately toxic compound by the World Health Organization (WHO). Investigating the microbial degradation mechanism of IPC not only is of theoretical significance but also holds potential for bioremediation of IPC-contaminated environments.

To date, several IPC-degrading strains have been reported, including members of the genera Rhodococcus (11), Lactobacillus (12), Streptococcus (12), Acinetobacter (13), Sphingobium (14), and Bacillus (15). Among them, the IPC degradation mechanism of Rhodococcus sp. D-6, a strain isolated in our laboratory, has been extensively characterized (11). In strain D-6, IPC is first hydrolyzed by the hydrolase IpcH to produce 2-isopropylphenol (IPP). IPP is subsequently hydroxylated by the two-component flavin-dependent monooxygenase IppA1A2 to generate 2-isopropylhydroquinone (11, 16). The resulting 2-isopropylhydroquinone undergoes sequential reactions, including oxidative ring opening, dehydrogenation, and reduction, eventually entering the tricarboxylic acid cycle as a carbon and energy source for the growth of strain D-6 (16). Notably, strain D-6 can utilize IPP, the hydrolysis product of IPC, as its sole growth substrate (16). However, the regulatory mechanism of IPC degradation in strain D-6 remains unclear.

PucR family transcriptional regulators participate in diverse physiological pathways, including amino acid metabolism, fatty acid biosynthesis, purine metabolism, and antibiotic biosynthesis (17–26). They play essential roles in bacterial central metabolism and environmental adaptation. Unlike PucR from Bacillus subtilis 168, which functions as both an activator and repressor of the purine catabolic pathway, all other characterized PucR-type regulators act exclusively as transcriptional activators (17–26). PucR family members share two conserved structural domains: a GGDEF-like domain and a PucR-type C-terminal helix-turn-helix (HTH) domain. The GGDEF-like domain is widely present among functionally characterized PucR-type regulators, except for GabR, a regulator of 4-aminobutyrate utilization in Corynebacterium glutamicum ATCC 13032 (22, 23). This domain has been hypothesized to mediate effector binding by Megalizzi et al., but experimental validation is currently lacking (23). The C-HTH domain of the PucR family, which binds promoter DNA to modulate transcription of target genes, exists in all functionally characterized PucR family regulators (19, 22, 23).

Previous studies showed that the ipcH gene is constitutively expressed in strain D-6, while the ippA1A2 cluster is inducible (11, 16). In this study, the PucR-type regulator IppR was identified as the transcriptional activator of the ippA1A2 cluster in response to IPP in strain D-6. The regulatory mechanism was investigated, including the transcriptional units, DNA-binding sites, effector molecule, and key amino acids essential for effector binding and DNA recognition.

RESULTS

IppR positively regulates IPP degradation in strain D-6

Genome analysis of strain D-6 revealed a putative transcriptional regulator gene, ippR, upstream of the ippA1A2 genes (Fig. 1B and Fig. S1). BLASTP analysis conducted in the NCBI non-redundant database indicated that IppR shared the highest sequence similarity (80.53%) with an uncharacterized PucR family regulator from Rhodococcus aetherivorans (WP_387433404), with 99% coverage. Phylogenetic analysis further demonstrated that IppR clustered with SrmR, MabR, and SdaR within a subclade of the PucR family (Fig. S1). Among functionally characterized PucR-type regulators, IppR showed the highest amino acid sequence identity to SrmR (29.66%), a regulator of spiramycin biosynthesis in Streptomyces ambofaciens BES2281 (17, 18). IppR shared 27.54% sequence identity with MabR, which regulates mycolic acid biosynthesis in Mycobacterium tuberculosis H37Rv (19, 20), and 27.66% identity with SdaR, which regulates the metabolism of d-galactarate, d-glycerate, and d-glucarate in Escherichia coli K12 (21). Multiple sequence alignment revealed that IppR contains a GGDEF-like domain (Val197-Ala311) and a PucR-type C-HTH domain (Asp350-Ala419), both of which are characteristic features of PucR family transcriptional regulators (Fig. 2A). Collectively, these results identify IppR as a new member of the PucR-type regulator.

Fig 1.

Bar chart shows ippA1 and ippA2 expression peaking in D-6 and D-6 delta ippR/ippR with IPP, near zero with glucose or in D-6 delta ippR, confirmed by gel electrophoresis of cDNA fragments.

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(A) Hydroxylation of 2-isopropylphenol (IPP) to 2-isopropylhydroquinone by the two-component monooxygenase IppA1A2. (B) Schematic diagram of the ippA1A2 gene cluster in strain D-6. The amplification fragments for transcriptional unit evaluation are shown under the gene cluster as lines. (C) Transcriptional analysis of ippA1 and ippA2 in strain D-6, the ippR knockout strain D-6ΔippR, and the ippR-complementary strain D-6ΔippR/ippR in the presence of 1.04 mM glucose or 1.04 mM IPP. The transcriptional level of the 16S rRNA gene was used as an internal standard, and the data in each column were calculated by the 2−ΔΔCT threshold cycle (CT) method using three replicates. (D) PCR amplification of the ippA1A2 cluster using DNA (D), total RNA (R), and cDNA (cD) as the templates. The amplified products were detected by electrophoresis. (E) DNA elements in the promoter of IPP1 operon. The IppR-binding site in the promoter region of IPP1 operon is indicated by a gray box. Fragments FIPP1 and PIPP1 were underlined.

Fig 2.

Sequence alignment of IppR, SrmR, MabR, and SdaR showing conserved residues in the GGDEF-like and PucR C-terminal HTH domains, with the predicted IppR structure highlighting distinct N-terminal and C-terminal domains.

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(A) Alignment of IppR with the PucR-type transcriptional regulator. IppR was aligned with SrmR (CAA45051), MabR (P9WPH5), and SdaR (P37047) using ClustalX. The GGDEF-like domain and the PucR C-terminal helix-turn-helix (HTH) domain were underlined. The key amino acids of IppR for binding to IPP and the promoter region of the IPP1 operon were indicated by a black arrow and a blue arrow, respectively. (B) Structure of IppR predicted by ColabFold. The N-terminal domain was shown in wheat color, and the C-terminal domain was shown in light blue color.

To investigate the role of IppR in regulating the expression of the gene cluster, the ippR knockout strain D-6ΔippR and its complementary strain D-6ΔippR/ippR were constructed. Cells of strains D-6, D-6ΔippR, and D-6ΔippR/ippR were incubated in MSM supplemented with 1.04 mM IPP as the sole growth substrate. As shown in Fig. S2A, the wild-type strain D-6 exhibited a 6-h lag phase and completely degraded 1.04 mM IPP within 21 h, whereas strain D-6ΔippR lost the ability to degrade IPP. Additionally, the cell density of strain D-6 cultured with 1.04 mM IPP increased significantly from 3.0 × 106 CFU/mL to 7.1 × 107 CFU/mL (Fig. S2B), indicating that IPP served as the carbon and energy source for cell growth of strain D-6. In contrast, the cell density of strain D-6ΔippR cultured with IPP showed no significant change (Fig. S2B). Moreover, the degradation and growth characteristics of the complementary strain D-6ΔippR/ippR were comparable to those of the wild-type strain D-6 (Fig. S2).

The transcriptional levels of ippA1 and ippA2 were evaluated in strains D-6, D-6ΔippR, and D-6ΔippR/ippR grown on IPP or glucose using qRT-PCR. In strain D-6, the transcription levels of ippA1 and ippA2 in the presence of IPP were 25- and 27-fold higher, respectively, than those in the presence of glucose (Fig. 1C). However, no significant differences in transcriptional levels of these genes were observed in the ippR knockout strain D-6ΔippR between IPP- and glucose-grown cells (Fig. 1C). In the ippR complementary strain D-6ΔippR/ippR, the transcriptional levels of these genes were similar to those of wild-type strain D-6 (Fig. 1C). These results indicate that IppR functions as a transcriptional activator of the ippA1A2 cluster.

Organization and transcriptional analysis of the ippA1A2 cluster

To investigate the transcript of the ippA1A2 cluster, total DNA, RNA, and cDNA extracted from IPP-induced cells of strain D-6 were used as templates for PCR amplification. The PCR products were verified by gel electrophoresis. Among the four amplified fragments, only fragments F2 and F3 were amplified from cDNA (Fig. 1D). These results indicate that ippA1, orf2560, and ippA2 are transcribed as a single operon, designated the IPP1 operon.

To identify the promoter region of the IPP1 operon, its TSS was determined by 5′-RACE using total RNA extracted from IPP-induced cells of strain D-6. The TSS of the IPP1 operon was “A” at 97 bp upstream of the ippA2 start codon (Fig. S3). The promoter region of IPP1 operon, a 128-bp fragment located upstream of TSS, was designated as fragment PIPP1 (Fig. 1E). A detailed analysis of this promoter region is shown in Fig. 1E.

IPP as an effector that enhances IppR binding to the promoter region of the IPP1 operon

The ippR was cloned from strain D-6 and expressed in E. coli BL21(DE3). Recombinant IppR was purified as a C-terminal 6×His-tagged protein using an Ni-NTA agarose resin matrix. The monomeric molecular weight of IppR was determined to be approximately 45.0 kDa by SDS-PAGE, consistent with its expected molecular weight (Fig. S4A). As shown in Fig. S5, gel filtration chromatography revealed that native IppR eluted as a dimer (the calculation method is described in the legend of Fig. S5). Collectively, these results demonstrate that IppR exists as a homodimer in vitro.

EMSA was performed to determine whether IppR binds to the promoter region of the IPP1 operon (fragment FIPP1, the 225-bp fragment spanning from −128 to +97 of the TSS). The results showed that IppR bound to fragment FIPP1 along with increasing amounts (0–1.26 μM) of IppR, with a detectable FIPP1-IppR complex forming at 0.63 μM IppR (Fig. 3A). In contrast, no DNA-protein complex was observed with a nonspecific control DNA fragment derived from ippA1, confirming the binding specificity between IppR and fragment FIPP1 (Fig. 3A). Subsequently, the effect of IPP on the binding of IppR to fragment FIPP1 was evaluated by EMSA. When IPP was present in the incubation mixture of IppR and fragment FIPP1, the binding capacity of IppR to the promoter was enhanced. As shown in Fig. 3A, the formation of the FIPP1-IppR complex required only 0.42 μM IppR in the presence of IPP, while 0.63 μM IppR was required in its absence. Additionally, at identical IppR concentrations (0.63, 0.84, and 1.05 μM), IPP treatment lowered the amount of free fragment FIPP1 and promoted the formation of the FIPP1-IppR complex (Fig. 3A). These results indicate that IppR binds to the promoter region of the IPP1 operon, and IPP is an effector to enhance this interaction.

Fig 3.

EMSA gel and bar charts show IPP disrupts IppR binding to fragment FIPP1, with beta-galactosidase activity rising markedly at 3 and 6 hours in IPP-treated D-6 and D-6 delta ippA1 strains versus glucose controls.

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Determination of the effector of IppR. (A) Effects of IPP on the binding of IppR to the promoter region of IPP1 operon (fragment FIPP1). EMSAs with fragment FIPP1 (40 nM) and purified IppR (0–1.26 μM) without or with 0.21 mM IPP. The control DNA was a 125-bp fragment that was amplified from the ippA1 gene. (B) The β-galactosidase activity assay was performed with strain D-6 carrying the PIPP1-lacZ transcriptional fusion, grown in the presence of 1.04 mM glucose or IPP. (C) The β-galactosidase activity assay was performed with strain D-6ΔippA1 carrying the PIPP1-lacZ transcriptional fusion, grown in the presence of 1.04 mM glucose and IPP.

To further investigate the effect of IPP on the binding of IppR to the promoter region of the IPP1 operon, the promoter region was cloned into the β-galactosidase reporter plasmid pME6522 (27), generating the plasmid pME6522-PIPP1. The resulting promoter-lacZ fusion plasmid pME6522-PIPP1 was subsequently introduced into strain D-6, generating derivative strain D-6-PIPP1. β-Galactosidase activity in strain D-6-PIPP1 was induced using glucose and IPP. As illustrated in Fig. 3B, strain D-6-PIPP1 exhibited low β-galactosidase activity (45 Miller units) in the presence of glucose, whereas IPP significantly increased its activity to 414 Miller units. These results suggest that IPP acts as the effector molecule of IppR. To validate this, the plasmid pME6522-PIPP1 was introduced into the ippA1 knockout strain D-6ΔippA1, which is unable to degrade IPP (16). As shown in Fig. 3C, IPP still significantly induced β-galactosidase activity in strains D-6ΔippA1-PIPP1. Additionally, IPC, 2-isopropylhydroquinone, and its downstream metabolites exhibit no effector activity toward IppR (data shown in the supplemental text). These results confirm that IPP functions as the effector molecule of IppR and activates transcription of the ippA1A2 cluster.

Key amino acids of IppR involved in IPP binding

To identify the amino acids in IppR required for IPP binding, molecular docking analyses of IPP to IppR were performed. Five structural models of IppR were predicted using the ColaFold platform based on the AlphaFold2 algorithm (Fig. S6). The reliability of these models was evaluated primarily using the mean predicted local distance difference test (pLDDT) scores, together with predicted aligned error (PAE) profiles (Fig. S6). Among the five models, model IV showed the highest mean pLDDT score (88.0), compared with models I (87.1), II (85.5), III (87.6), and V (85.2) (Fig. S6). Therefore, model IV was selected for subsequent structural analysis. The predicted structure of the IppR monomer consists of two distinct regions: an N-terminal domain (N-IppR, Met1 to Glu195) and a C-terminal domain (C-IppR, Asp196 to Arg420) (Fig. 2), consistent with the domain organization of Mycobacterium tuberculosis MabR (PDB IDs: 9F81 and 9F80), the only structurally characterized PucR-type regulator reported to date (23). For the N-terminal domain, both IppR and MabR adopt an all-α structure. Specifically, the predicted N-IppR is composed of two 310-helices (η1 and η2) and six α-helices (α1 to α6), while N-MabR consists of six α-helices. For the C-terminal domain, both IppR and MabR possess a GGDEF-like domain and a PucR-type C-HTH domain. Specifically, the GGDEF-like domain of C-IppR (Val197-Ala311) comprises three α-helices (α7 to α9) and four β-strands (β1 to β4), while the corresponding domain in MabR contains three 310-helices, five α-helices, and five β-strands. The PucR-type C-HTH domain of IppR (Asp350-Ala419) contains four α-helices (α12 to α15), similar to MabR, which also adopts four α-helices. In addition, the two domains of IppR are connected by two 310-helices (η3 and η4), two α-helices (α10 and α11), and one β-strand (β5), while only two α-helices mediate the interdomain connection in MabR.

To predict the interaction between IPP and IppR, molecular docking of IPP was performed against the putative active sites of IppR using the AutoDock program. The results showed that the substrate binding pocket of IPP was located within the GGDEF-like domain and the adjacent region. The IPP binding pocket was comprised of five antiparallel β-strands (β1 to β5) surrounded by four α-helices (α6 to α9) (Fig. 4A). Visualization analysis revealed that the vicinity of IPP contained several hydrophobic residues, including Val188, Leu192, Val218, Trp257, Trp259, and Ala300, as well as the amphipathic residue His297 (Fig. 4B). Val188, Trp257, Ala300, and the amphipathic His297 engage in hydrophobic interactions with IPP. These residues encircle IPP, creating a hydrophobic microenvironment in the predicted effector-binding site (Fig. 4B). The parallel orientation of the IPP benzene ring relative to the aromatic ring of Trp257 enables strong π-π stacking interactions (Fig. 4B), which likely stabilize IPP within the IppR binding pocket and enhance its binding affinity.

Fig 4.

3D structure of IppR's IPP-binding domain with key residues Trp-257, Trp-259, His-297, and Val-188 shown, alongside EMSAs revealing mutants lose DNA-binding ability, restored by IPP addition.

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Identification of key amino acids for IPP binding in IppR. (A) The IPP-binding domain of IppR. (B) Residues in the IPP-binding pocket of IppR. Residues in yellow are hydrophobic residues, residues in orange are amphipathic residues, and the molecule in blue is IPP. π-π stacking interaction are shown in yellow dotted lines. (C) EMSAs using fragment FIPP1 (40 nM) with IppR and its mutant proteins (1.26 μM). (D) EMSAs using fragment FIPP1 (40 nM) with IppR and its mutant proteins (0.42 μM) in the presence of 0.21 mM IPP.

To assess the role of these residues in IPP binding, Val188, Leu192, Val218, Trp257, Trp259, His297, and Ala300 were individually substituted with glycine, generating mutants IppR-V188G, IppR-L192G, IppR-V218G, IppR-W257G, IppR-W259G, IppR-H297G, and IppR-A300G (Fg. S4B). These mutant proteins could still bind to the fragment FIPP1 when their concentration reached 1.26 μM (Fig. 4C). However, the fragment FIPP1 was not shifted when incubated with the mutant proteins (0.42 μM) in the presence of 0.21 mM IPP, while the FIPP1-IppR complex formed under the same conditions (Fig. 4D). These findings indicate that the mutant proteins maintained DNA-binding capability to fragment FIPP1 but lost responsiveness to IPP. Moreover, strains D-6ΔippR/ippR-V188G, D-6ΔippR/ippR-L192G, D-6ΔippR/ippR-V218G, D-6ΔippR/ippR-W257G, D-6ΔippR/ippR-W259G, D-6ΔippR/ippR-H297G, and D-6ΔippR/ippR-A300G were unable to degrade IPP (Table S1). These results are consistent with Val188, Leu192, Val218, Trp257, Trp259, His297, and Ala300 playing key roles in IPP binding.

Binding sites of IppR to the promoter region of the IPP1 operon

To determine the key binding sites of IppR, the fragment FIPP1 was divided into six sub-fragments FIPP1-1 to FIPP1-6 and analyzed by EMSA (Fig. 5A). As shown in Fig. 5B, fragment FIPP1 and sub-fragments FIPP1-2, FIPP1-3, FIPP1-5, and FIPP1-6 were shifted upon incubation with IppR, forming the DNA-IppR complex. These five fragments shared a 24-bp overlapping DNA motif 5′-ACCGGATGCGACCTGCGCATTGGT-3′.

Fig 5.

Diagram of ippA1 and ippR regions with sub-fragments and an EMSA gel showing DNA-IppR complex formation. Fragments containing the CC/CG box show shifted bands, indicating IppR binding.

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Identification of IppR-binding site within the promoter regions of fragments FIPP1. (A) Schematic diagram of fragment FIPP1 and its sub-fragments FIPP1-1 to FIPP1-6 used to determine the IppR-binding site. The locations of the fragments are shown below. The nucleotide sequence above the fragments shows the IppR-binding site within the promoter regions of fragment FIPP1, and the CC/CG box was highlighted in red. (B) EMSA of fragment FIPP1 and sub-fragments FIPP1-1 to FIPP1-6 in panel A with IppR. The lanes contain the following: the DNA fragment (40 nM) alone (−) and the DNA fragment (40 nM) with 1.26 μM IppR (+).

To further investigate essential binding sites within the 24-bp motif, molecular docking with the target motif to IppR dimer was performed using the HDOCK Server (28, 29). The predicted DNA-IppR complex structure indicated that the IppR dimer recognizes the 24-bp DNA motif through the C-HTH domain of each monomer. As shown in Fig. 6A, the α14 of IppR (the third α-helix within the C-HTH domain of each IppR monomer) acts as the recognition helix and inserts into the DNA major groove. In the complex, Lys382 forms hydrogen bonds with C-12, C-13, C-16, and G-17 (Fig. 6A). To investigate the role of the CC/CG box in IppR binding, various mutant DNA fragments of FIPP1 were analyzed using EMSAs. As shown in Fig. 6B, only the M1-1-FIPP1 fragment could bind to IppR to form DNA-IppR complex, while all other mutant fragments lost binding activity. These results further suggest the critical role of the CC/CG box in IppR sequence recognition.

Fig 6.

Structural model of IppR dimer bound to DNA, with Lys382 and Thr384 contact residues shown. EMSA gels reveal that CC and CG box mutations abolish DNA-IppR complex formation, confirmed by IppR-K382G and IppR-T384G mutants.

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(A) Structure model of the DNA-IppR complex. The IppR dimer is bound to the 24-bp DNA motif. The two monomers of IppR are represented in light blue and light green, respectively. The close-up view shows IppR bound to DNA, with the interacting bases in yellow and the predicted contact residues Lys382 and Thr384 as sticks. (B) Diagram of the wild-type sequence of fragment FIPP1 and its mutant sequences was shown at the top. The mutant nucleotides between the wild-type sequence and the mutant type sequences are shown in red. Lane 0, the wild-type promoter fragment FIPP1 (40 nM) alone, lanes WT-FIPP1 to MT-FIPP1, the mutant DNA fragment (40 nM) with purified IppR (1.26 μM). (C) EMSAs using fragment FIPP1 (40 nM) with purified IppR and its mutant proteins (1.26 μM).

Key amino acids of IppR involved in binding to the promoter DNA

In addition to Lys382, which forms hydrogen bonds with the CC/CG box, Thr384 also forms non-specific hydrogen bonds with the phosphate backbone of the 24-bp DNA motif (Fig. 6A). To determine the roles of Lys382 and Thr384 for DNA binding in IppR, two variants (IppR-K382G and IppR-T384G) were constructed (Fig. S4C). As shown in Fig. 6C, the DNA-binding abilities of both variants were significantly altered, resulting in a complete disruption of fragment FIPP1-binding capacity. Furthermore, strains D-6ΔippR/ippR-K382G and D-6ΔippR/ippR-T384G lost the ability to degrade IPP (Table S1). These results are consistent with Lys382 and Thr384 playing crucial roles in the DNA-binding function of IppR.

DISCUSSION

This study presents evidence that IppR is a PucR-type transcriptional activator that positively regulates the transcription of the ippA1A2 genes, which encode an IPP hydroxylase. This role is consistent with most characterized PucR-type regulators, which act as transcriptional activators (17–26). A notable exception is PucR itself, which acts as a dual regulator with both an activator and a repressor activity. Based on the presented data, we propose that IppR binds IPP as an effector and functions similarly as the PucR-type activators. However, further studies are required to elucidate the mechanism of action of PucR family transcriptional regulators.

The GGDEF-like domain shares evolutionary similarity with the canonical GGDEF domain, which plays a pivotal role in binding the substrate guanosine triphosphate in diguanylate cyclase (30–32). Structurally, the GGDEF domain of diguanylate cyclase adopts a typical fold consisting of five antiparallel β-strands surrounded by five peripheral α-helices. The structure of the GGDEF-like domain of IppR comprises four anti-parallel β-strands surrounded by three α-helices. Despite differences in secondary structural element composition, the two domains exhibit a conserved overall topology characterized by a central β-sheet wrapped by peripheral α-helices. Megalizzi et al. hypothesized that the GGDEF-like domain of PucR-type regulators could potentially function as the effector-binding site (23). However, this hypothesis has not been experimentally verified. In this study, homology modeling combined with molecular docking analysis identified that the GGDEF-like domain of IppR, along with its neighboring amino acid residues, collectively form the substrate-binding pocket for IPP (Fig. 4A). The IppR binding pocket forms a hydrophobic microenvironment consistent with the hydrophobic characteristics of IPP. Hydrophobic interactions with IPP and π-π stacking interaction mediated by Trp257 enhance the stabilization of IPP within the active site (Fig. 4B). Collectively, these results confirm that the GGDEF-like domain and its vicinity are the essential binding centers for the effector molecule IPP. This study offers direct evidence that the GGDEF-like domain of PucR family regulators is directly involved in effector binding and functional regulation, representing the first report of the GGDEF-like domain exerting regulatory effects through effector binding.

Based on previous reports, PucR-type regulatory proteins exhibit diverse target binding sites on promoter DNA. Beier et al. found that PucR, a regulatory protein controlling the purine catabolic pathway in Bacillus subtilis 168, recognizes a conserved binding motif on promoter DNA (24). In contrast, other characterized PucR-type members bind promoter DNA via recognition of inverted repeat sequences, including AdeR (25), PrcR (26), MabR (19), and GabR (22). AdeR binds the promoter of L-alanine dehydrogenase-encoding ald to regulate gene transcription (25). PrcR regulates proline utilization in strain 168 by binding to the promoter DNA of the putBCP operon (26). In Mycobacterium tuberculosis H37Rv, MabR controls mycolic acid biosynthesis (19). GabR in Corynebacterium glutamicum ATCC 13032 regulates 4-aminobutyric acid metabolism (22). Additionally, no target DNA-binding sites have been reported for SrmR and SdaR, two members of the PucR family. In this study, the key promoter region bound by IppR was identified as 5′- ACCGGATGCGACCTGCGCATTGGT-3′ (Fig. 5). Molecular docking and site-directed mutagenesis experiments validated that mutation of the CC/CG box within this region eliminated the retardation of the DNA fragment by IppR (Fig. 6A and B), indicating that the CC/CG box is essential for stable formation of the DNA-IppR complex. This binding site differs from previously reported PucR family binding patterns, providing a novel motif type for promoter recognition by PucR family transcriptional regulators.

The HTH domain is widespread among diverse regulators and mainly functions in recognizing and binding promoter DNA of target genes (33–40). Members of the PucR family harbor a C-terminal HTH domain that is critical for DNA recognition (23). Consistent with these observations, our structural characterization identifies that IppR contains a PucR-type HTH domain at its C-terminal. Structural analysis of the predicted IppR-DNA complex demonstrates that the third α-helix within this domain inserts into the DNA major groove, thereby mediating promoter DNA binding (Fig. 6B). In comparison, structural analysis of Mycobacterium tuberculosis MabR, the sole structurally characterized PucR-type regulator, has confirmed that its C-terminal HTH domain uses the second α-helix to insert into the DNA major groove for sequence-specific binding (23). Lys382 of IppR is predicted to form hydrogen bonds with the CC/CG box, while Thr384 forms non-specific hydrogen bonds with the phosphate backbone of the 24-bp core DNA-binding motif (Fig. 6A). These two residues are predicted to be essential for IppR recognition and binding to the promoter DNA of the IPP1 operon. Notably, although several PucR-family regulators have been functionally characterized, the precise PucR-type regulators’ DNA binding mode and the critical residues mediating their interaction have not been defined to date. Our findings not only provide a basis for clarifying the molecular mechanism of DNA-IppR binding but also confirm that the HTH domain is the key functional domain responsible for DNA recognition and binding in PucR family regulators.

MATERIALS AND METHODS

Chemicals and media

2-Isopropylphenol (IPP; 98.0% purity) was purchased from Macklin Biochemical Co., Ltd. (Shanghai, China). Luria-Bertani (LB, pH 7.0) medium consisted of the following components (in g liter−1): 10.0 tryptone, 5.0 yeast extract, and 10.0 NaCl. Mineral salt medium (MSM, pH 7.0) consisted of the following components (in g liter−1): 1.5 K2HPO4, 0.5 KH2PO4, 0.2 MgSO4·7H2O, 1.0 NH4NO3, and 1.0 NaCl. IPP (1.04 mM) or glucose (1.04 mM) was used as the sole growth substrate for cell growth of strain D-6 and its related mutant strains unless stated otherwise.

Bacterial strains, plasmids, and culture conditions

All bacterial strains and plasmids utilized in this study were listed in Table S2, and the primers were listed in Table S3. Escherichia coli strains were cultivated at 37°C in LB medium. Rhodococcus sp. D-6, an isoprocarb (IPC)-degrading strain utilizing IPC as the sole carbon and energy source for its growth, and strain D-6∆ippA1, an ippA1 knockout strain that lost its IPP degradation ability, were previously isolated and constructed by our lab (11, 16). Strain D-6 and its derivatives were grown at 30°C in LB medium. Antibiotics were supplemented according to Table S2. Streptomycin (Str), tetracycline (Tc), and kanamycin (Km) were supplemented to final concentrations of 25 μg mL−1, 15 μg mL−1, and 50 μg mL−1, respectively.

Genetic manipulations

The ippR gene was knocked out through a double-crossover event. Two DNA fragments, corresponding to the 1,300 bp upstream and downstream flanking regions of the ippR gene, were amplified using primer pairs ippR-upF/ippR-upR and ippR-downF/ippR-downR (listed in Table S3), respectively. These two PCR fragments were combined through overlap extension PCR and subsequently cloned into BamHI- and EcoRI-digested pK18mobSacB using the ClonExpress II one-step cloning kit (Vazyme, Nanjing, China), resulting in the generation of pK18-ippR (41). Subsequently, pK18-ippR was introduced into strain D-6 via electroporation. Single-crossover mutants were screened on LB plates supplemented with Km (50 μg mL−1) and Str (25 μg mL−1) and confirmed through PCR amplification. Double-crossover mutants were selected on LB plates containing Str (25 μg mL−1) and 20% sucrose and verified by PCR amplification. The double-crossover knockout strain was sequenced to ensure accurate gene deletion and designated as D-6ΔippR.

To obtain the ippR-complementary strain D-6ΔippR/ippR, the ippR gene was amplified with the primer pair ippR-hbF/ippR-hbR (listed in Table S3) from genomic DNA of strain D-6. The resulting PCR product was ligated into pRESQ, generating pRE-ippR, which was then electroporated into strain D-6∆ippR for complementation (42).

The construction of point mutant strains was similar to the procedure used for ippR complementation, with the differences described below. Primers carrying desired mutation sites were used for fragment amplification. The resulting plasmids were electroporated into strain D-6ΔippR to generate corresponding complementary strains.

Growth and degradation assay

Strain D-6 and its derivates were inoculated into LB medium, harvested by centrifugation, and washed three times with MSM. The washed cells were then resuspended in MSM containing 1.04 mM IPP, with an initial density of approximately 3.0 × 106 CFU/mL. The residual concentration of IPP was determined by HPLC analysis. Concurrently, the growth of these strains was quantified by enumerating viable colonies on LB agar plates supplemented with 25 mg/L Str. All experiments were conducted in triplicate.

Quantitative reverse transcription-PCR

Strains D-6, D-6ΔippR, and D-6ΔippR/ippR were cultured to an optical density at 600 nm (OD600) of 0.6 in LB medium containing Str (25 μg mL−1). Cells were then harvested by centrifugation (6,000 × g for 5 min), washed three times with fresh MSM, and resuspended in MSM to a final OD600 of 1.0 at 30°C in the presence of 1.04 mM IPP. MSM containing 1.04 mM glucose was used as the negative control. Upon reaching 40% to 60% of IPP degradation, total RNA was extracted using an RNA isolation kit (TaKaRa, Dalian, China). cDNA was synthesized using the PrimeScript RT reagent kit (Vazyme, Nanjing, China) and subsequently used as a template for quantitative PCR (qPCR). The 16S rRNA gene served as an internal standard control, and relative gene expression was calculated based on the 2−ΔΔCT threshold cycle (CT) method.

Determination of the transcription start sites

Total RNA was extracted from strain D-6 cultured in MSM supplemented with 1.04 mM IPP using an RNA isolation kit (TaKaRa, Dalian, China). The transcription start site (TSS) of the IPP1 operon, including genes ippA1, orf2560, and ippA2, was identified using 5′-rapid amplification of cDNA ends (5′-RACE) (Sangon, Shanghai, China). First-strand cDNA was reverse transcribed using primer TSS-A-1 (listed in Table S3), followed by cDNA tailing with terminal transferase and dGTP. The dG-tailed cDNA was then amplified using the abridged anchor primer (APP) and TSS-A-2 (listed in Table S3). The resulting amplification product underwent a nested PCR with AAP and primer TSS-A-3 (listed in Table S3) to generate the final DNA fragment. The final PCR product was cloned into the pMD19-T vector (TaKaRa, Japan) and sequenced using M13F-47 and M13R-48 primers. The promoter region of the IPP1 operon was predicted using BDGP (43) and BacPP (44).

Protein expression and purification

The ippR gene was amplified from strain D-6 using the primer pair ippR-F/ippR-R (listed in Table S3) and cloned into pET-29a(+) to yield pET-ippR (45, 46). The plasmid pET-ippR was transformed into E. coli BL21(DE3), and the cells were cultured at 37°C in LB medium with 50 µg mL−1 Km until OD600 reached approximately 0.6. Subsequently, cells were induced at 16°C for 10 h by adding 0.1 mM isopropyl-β-d-thiogalactopyranoside. After induction, cells were harvested, washed, and lysed by sonication. Subsequently, the C-terminal 6×His-tagged protein IppR was purified using an Ni-NTA agarose resin matrix. The concentration and molecular weight of IppR were determined by the Bradford method (47) and sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), respectively.

The multimeric state of native IppR was determined by gel filtration chromatography (48). IppR was loaded onto a Superdex 200 Increase 10/300 GL column (GE Healthcare) with 20 mM Tris-HCl buffer (pH 7.4, containing 0.1 M NaCl) as the mobile phase (0.4 mL min−1) at 25°C. Standard proteins, including β-galactosidase (116 kDa), phosphorylase b (97 kDa), bovine serum albumin (66 kDa), and ovalbumin (45 kDa), were used. The molecular weight (MW) of IppR was calculated from the linear regression of the standard curve, which was generated by plotting the logarithm of the molecular weights of the standards against their corresponding retention volume (V), following the equation Log10(Mw) = a × V + b, where a is the slope and b is the intercept of the calibration curve.

The construction, expression, and purification of mutant IppR proteins were performed similarly to those of wild-type IppR, with the differences described below. Mutants of the ippR gene were constructed using overlap PCR described by Zhu et al. (49).

Electrophoretic mobility shift assay

The promoter regions of the IPP1 operon were amplified from the genomic DNA of strain D-6 using the primer pair FIPP1-F/FIPP1-R presented in Table S3. A 125-bp region of ippA1 used as the negative control was amplified using the primer pair ippA1-F/ippA1-R (Table S3). To explore the core binding sites of IppR, subfragments FIPP1-1 to FIPP1-6 of the promoter region of the IPP1 operon were amplified using primers listed in Table S3. Point mutations in the promoter were generated via PCR, with the principle of mutating G and C to A and T, respectively. All mutated promoter DNA was amplified using primers listed in Table S3. The 40 nM promoter probe was incubated with purified IppR (0–1.26 µM) in EMSA binding buffer, which consisted of 20 mM Tris-HCl, 1 mM dithiothreitol, 0.5 mM EDTA, 50 mM KCl, and 5% (vol/vol) glycerol. To determine whether the effector of IppR was IPP, 0.21 mM IPP was added to the reaction system. The reaction mixtures were incubated at 25°C for 30 min and subsequently resolved on a 5% (vol/vol) native polyacrylamide gel by electrophoresis in 0.5 × Tris-glycine-EDTA buffer for 1 h. The gels were stained with SYBR green I for 15 min to visualize the DNA and DNA-protein complexes.

Reporter plasmid construction and β-galactosidase activity assay

The promoter region of the IPP1 operon was amplified using primer pair ippA2-6522-F and ippA2-6522-R listed in Table S3. The purified PCR product was digested with EcoRI and PstI and ligated with pME6522 (27) to construct plasmid pME6522-PIPP1. Subsequently, plasmid pME6522-PIPP1 was electroporated into strains D-6 and D-6ΔippA to yield strains D-6-PIPP1 and D-6ΔippA-PIPP1, respectively.

β-Galactosidase activity was assayed using strain D-6-PIPP1 grown in MSM supplemented with 1.04 mM IPP or 1.04 mM glucose, respectively. Cultures were incubated at 30°C with shaking at 180 rpm. β-Galactosidase activity assays utilized o-nitrophenyl-β-d-galactopyranoside (ONPG) as substrate, with 300 μL culture samples harvested for analysis. The observed activity was normalized to the optical density of the culture at 600 nm, the optical density of o-nitrophenol (ONP) at 420 nm, and expressed in Miller units (50). One Miller unit was defined as the amount of enzyme that catalyzed the conversion of ONPG into 1 µmol ONP per min.

Structure modeling

The structural model of IppR was predicted using ColabFold (AlphaFold2.ipynb-Colaboratory), and the 3D conformer of IPP was obtained from PubChem (https://pubchem.ncbi.nlm.nih.gov/) (51, 52). IPP and the 24-bp motif were docked to IppR using AutoDock Vina software (53, 54) and HDOCK server (http://hdock.phys.hust.edu.cn/) (28, 29), respectively. Figures were generated using PyMOL software.

Analytical techniques

To analyze IPP, 1 mL of degradation culture (MSM containing IPP inoculated with strain D-6 and its variant strains) was thoroughly mixed with 1 mL of methanol and centrifuged for 5 min at 12,000 × g. The supernatants were subsequently analyzed by HPLC (Thermo Fisher Scientific). The mobile phase consisted of methanol-water-glacial acetic acid (50:49:1, vol/vol/vol) at a flow rate of 1.0 mL min−1. Elution was monitored at 215 nm, with an injection volume of 20 μL and a column temperature of 40°C.

ACKNOWLEDGMENTS

This work was supported by the National Natural Science Foundation of China (32400084 and 32400085), the Natural Science Foundation of Jiangsu Province (BK20250193), and the National Key R&D Program of China (2022YFA0912500).

Contributor Information

Shang Dai, Email: daishang@njau.edu.cn.

Qing Hong, Email: hongqing@njau.edu.cn.

Marina Lotti, University of Milano-Bicocca, Milan, Italy.

DATA AVAILABILITY

The genome sequence of Rhodococcus sp. D-6 has been deposited in the GenBank database under the accession numbers CP132970 to CP132972 (11).

SUPPLEMENTAL MATERIAL

The following material is available online at https://doi.org/10.1128/aem.01277-26.

Supplemental material. aem.01277-26-s0001.pdf.

Tables S1 to S3, Fig. S1 to S6, and supplemental text.

aem.01277-26-s0001.pdf (846.4KB, pdf)
DOI: 10.1128/aem.01277-26.SuF1

ASM does not own the copyrights to Supplemental Material that may be linked to, or accessed through, an article. The authors have granted ASM a non-exclusive, world-wide license to publish the Supplemental Material files. Please contact the corresponding author directly for reuse.

REFERENCES

  • 1. Chapalamadugu S, Chaudhry GR. 1992. Microbiological and biotechnological aspects of metabolism of carbamates and organophosphates. Crit Rev Biotechnol 12:357–389. doi: 10.3109/07388559209114232 [DOI] [PubMed] [Google Scholar]
  • 2. Silberman J, Taylor A. 2023. Carbamate toxicity. In StatPearls. StatPearls Publishing, Treasure Island (FL). https://www.ncbi.nlm.nih.gov/books/NBK482183/. [PubMed] [Google Scholar]
  • 3. Xu L, Granger C, Dong H, Mao Y, Duan S, Li J, Qiang Z. 2020. Occurrences of 29 pesticides in the Huangpu River, China: highest ecological risk identified in Shanghai metropolitan area. Chemosphere 251:126411. doi: 10.1016/j.chemosphere.2020.126411 [DOI] [PubMed] [Google Scholar]
  • 4. Kurnia A, Ardiwinata AN, Harsanti ES, Sutriadi MT, Mulyono A, Makmur RF, Hindersah R. 2024. Chlorpyriphos and isoprocarb residues in the soil and food crop at agricultural area of Citarum watershed. International conference on organic and applied chemistry (ICOAC) 2022; Malang, Indonesia. Vol. 2957. doi: 10.1063/5.0184011 [DOI] [Google Scholar]
  • 5. Chen Y, Huang R, Guan Y, Zhuang T, Wang Y, Tan R, Wang J, Zhou R, Wang B, Xu J, Zhang X, Zhou K, Sun R, Chen M. 2021. The profiling of elements and pesticides in surface water in Nanjing, China with global comparisons. Sci Total Environ 774:145749. doi: 10.1016/j.scitotenv.2021.145749 [DOI] [PubMed] [Google Scholar]
  • 6. Wang C, Lu G, Cui J. 2012. Responses of AChE and GST activities to insecticide coexposure in Carassius auratus. Environ Toxicol 27:50–57. doi: 10.1002/tox.20612 [DOI] [PubMed] [Google Scholar]
  • 7. Wang S, Han X, Yu T, Liu Y, Zhang H, Mao H, Hu C, Xu X. 2022. Isoprocarb causes neurotoxicity of zebrafish embryos through oxidative stress-induced apoptosis. Ecotoxicol Environ Saf 242:113870. doi: 10.1016/j.ecoenv.2022.113870 [DOI] [PubMed] [Google Scholar]
  • 8. Ma J, Chen J, Wang P, Tong S. 2008. Comparative sensitivity of eight freshwater phytoplankton species to isoprocarb, propargite, flumetralin and propiconazol. Pol J Environ Stud 17:525–529. [Google Scholar]
  • 9. Park H, Song G, Lim W. 2021. Isoprocarb induces acute toxicity in developing zebrafish embryos through vascular malformation. J Anim Reprod Biotechnol 36:17–24. doi: 10.12750/JARB.36.1.17 [DOI] [Google Scholar]
  • 10. Rahman MF, Siddiqui MK, Mahboob M, Mustafa M. 1990. Haematological and hepatotoxic effects of isoprocarb in chicken. J Appl Toxicol 10:187–192. doi: 10.1002/jat.2550100308 [DOI] [PubMed] [Google Scholar]
  • 11. Zhu Q, Liu H, Pan K, Zhu W, Qiao Y, Li Q, Hu J, Zhang M, Qiu J, Yan X, Ge J, Hong Q. 2024. The novel hydrolase IpcH initiates the degradation of isoprocarb in a newly isolated strain Rhodococcus sp. D-6. J Hazard Mater 476:135045. doi: 10.1016/j.jhazmat.2024.135045 [DOI] [PubMed] [Google Scholar]
  • 12. Chen FF, Pang JN, Ma CM, Qi L, Li T. 2018. Biodegradation of carbamate pesticide by Lactic acid bacteria. J Chin Inst Food Sci Technol 18:102–108. doi: 10.16429/j.1009-7848.2018.10.014 [DOI] [Google Scholar]
  • 13. Wu Q, Li F, Zhu X, Ahn Y, Zhu Y. 2022. Isolation and characterization of cyromazine degrading Acinetobacter sp. ZX01 from a Chinese ginger cultivated soil. Environ Sci Pollut Res 29:67765–67775. doi: 10.1007/s11356-022-20538-x [DOI] [PubMed] [Google Scholar]
  • 14. Wu J, Gao A, wang B, Yang Y, Li N, Zhang H. 2025. Cloning and expression of the isoprocarb hydrolase gene cehA from the newly isolated isoprocarb-degrading strain Sphingobium sp. R-7. Curr Microbiol 82:sp. doi: 10.1007/s00284-025-04183-4 [DOI] [PubMed] [Google Scholar]
  • 15. Hu K, Wang X, Zhu J, Liu A, Ao X, He L, Chen S, Zhou K, Yang Y, Zou L, Liu S. 2020. Characterization of carbaryl-degrading strain Bacillus licheniformis B-1 and its hydrolase identification. Biodegradation 31:139–152. doi: 10.1007/s10532-020-09899-7 [DOI] [PubMed] [Google Scholar]
  • 16. Zhu Q, Wei K, Pan K, Hu G, Zhu W, Huang Y, Wang C, Li Q, Zhang M, Qiu J, Hong Q. 2025. Identification of a novel ipp gene cluster responsible for 2-isopropylphenol degradation in strain Rhodococcus sp. D-6. Appl Environ Microbiol 91:e0099525. doi: 10.1128/aem.00995-25 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17. Geistlich M, Losick R, Turner JR, Rao RN. 1992. Characterization of a novel regulatory gene governing the expression of a polyketide synthase gene in Streptomyces ambofaciens. Mol Microbiol 6:2019–2029. doi: 10.1111/j.1365-2958.1992.tb01374.x [DOI] [PubMed] [Google Scholar]
  • 18. Karray F, Darbon E, Nguyen HC, Gagnat J, Pernodet JL. 2010. Regulation of the biosynthesis of the macrolide antibiotic spiramycin in Streptomyces ambofaciens. J Bacteriol 192:5813–5821. doi: 10.1128/JB.00712-10 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19. Salzman V, Mondino S, Sala C, Cole ST, Gago G, Gramajo H. 2010. Transcriptional regulation of lipid homeostasis in mycobacteria. Mol Microbiol 78:64–77. doi: 10.1111/j.1365-2958.2010.07313.x [DOI] [PubMed] [Google Scholar]
  • 20. Tsai YT, Salzman V, Cabruja M, Gago G, Gramajo H. 2017. Role of long-chain acyl-CoAs in the regulation of mycolic acid biosynthesis in mycobacteria. Open Biol 7:170087. doi: 10.1098/rsob.170087 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21. Monterrubio R, Baldoma L, Obradors N, Aguilar J, Badia J. 2000. A common regulator for the operons encoding the enzymes involved in D-galactarate, D-glucarate, and D-glycerate utilization in Escherichia coli . J Bacteriol 182:2672–2674. doi: 10.1128/JB.182.9.2672-2674.2000 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22. Zhu L, Mack C, Wirtz A, Kranz A, Polen T, Baumgart M, Bott M. 2020. Regulation of γ-aminobutyrate (GABA) utilization in corynebacterium glutamicum by the PucR-Type transcriptional regulator GabR and by alternative nitrogen and carbon sources. Front Microbiol 11:544045. doi: 10.3389/fmicb.2020.544045 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23. Megalizzi V, Tanina A, Grosse C, Mirgaux M, Legrand P, Dias Mirandela G, Wohlkönig A, Bifani P, Wintjens R. 2024. Domain architecture of the Mycobacterium tuberculosis MabR (Rv2242), a member of the PucR transcription factor family. Heliyon 10:e40494. doi: 10.1016/j.heliyon.2024.e40494 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24. Beier L, Nygaard P, Jarmer H, Saxild HH. 2002. Transcription analysis of the Bacillus subtilis PucR regulon and identification of a cis-acting sequence required for PucR-regulated expression of genes involved in purine catabolism. J Bacteriol 184:3232–3241. doi: 10.1128/JB.184.12.3232-3241.2002 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25. Lin TH, Wei GT, Su CC, Shaw GC. 2012. AdeR, a PucR-type transcription factor, activates expression of l-alanine dehydrogenase and is required for sporulation of Bacillus subtilis . J Bacteriol 194:4995–5001. doi: 10.1128/JB.00778-12 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26. Huang SC, Lin TH, Shaw GC. 2011. PrcR, a PucR-type transcriptional activator, is essential for proline utilization and mediates proline-responsive expression of the proline utilization operon putBCP in Bacillus subtilis . Microbiology (Reading) 157:3370–3377. doi: 10.1099/mic.0.054197-0 [DOI] [PubMed] [Google Scholar]
  • 27. Wang L, Tang H, Yu H, Yao Y, Xu P. 2014. An unusual repressor controls the expression of a crucial nicotine-degrading gene cluster in Pseudomonas putida S16. Mol Microbiol 91:1252–1269. doi: 10.1111/mmi.12533 [DOI] [PubMed] [Google Scholar]
  • 28. Li H, Huang E, Zhang Y, Huang SY, Xiao Y. 2022. HDOCK update for modeling protein-RNA/DNA complex structures. Protein Sci 31:e4441. doi: 10.1002/pro.4441 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29. Yan Y, Zhang D, Zhou P, Li B, Huang SY. 2017. HDOCK: a web server for protein-protein and protein-DNA/RNA docking based on a hybrid strategy. Nucleic Acids Res 45:W365–W373. doi: 10.1093/nar/gkx407 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30. Pei J, Grishin NV. 2001. GGDEF domain is homologous to adenylyl cyclase. Proteins 42:210–216. doi: 10.1002/1097-0134(20010201)42:2<210::aid-prot80>3.0.co;2-8 [DOI] [PubMed] [Google Scholar]
  • 31. Paul R, Weiser S, Amiot NC, Chan C, Schirmer T, Giese B, Jenal U. 2004. Cell cycle-dependent dynamic localization of a bacterial response regulator with a novel di-guanylate cyclase output domain. Genes Dev 18:715–727. doi: 10.1101/gad.289504 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32. Ryjenkov DA, Tarutina M, Moskvin OV, Gomelsky M. 2005. Cyclic diguanylate is a ubiquitous signaling molecule in bacteria: insights into biochemistry of the GGDEF protein domain. J Bacteriol 187:1792–1798. doi: 10.1128/JB.187.5.1792-1798.2005 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33. Aravind L, Anantharaman V, Balaji S, Babu MM, Iyer LM. 2005. The many faces of the helix-turn-helix domain: transcription regulation and beyond. FEMS Microbiol Rev 29:231–262. doi: 10.1016/j.femsre.2004.12.008 [DOI] [PubMed] [Google Scholar]
  • 34. Li CX, Liao LS, Wan XD, Zhang FF, Zhang T, Luo XM, Zhao S, Feng JX. 2021. PoxCbh, a novel CENPB-type HTH domain protein, regulates cellulase and xylanase gene expression in Penicillium oxalicum. Mol Microbiol 116:140–153. doi: 10.1111/mmi.14696 [DOI] [PubMed] [Google Scholar]
  • 35. Wintjens R, Rooman M. 1996. Structural classification of HTH DNA-binding domains and protein-DNA interaction modes. J Mol Biol 262:294–313. doi: 10.1006/jmbi.1996.0514 [DOI] [PubMed] [Google Scholar]
  • 36. Seetharaman J, Kumaran D, Bonanno JB, Burley SK, Swaminathan S. 2006. Crystal structure of a putative HTH-type transcriptional regulator yxaF from Bacillus subtilis. Proteins 63:1087–1091. doi: 10.1002/prot.20924 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37. Rampioni G, Polticelli F, Bertani I, Righetti K, Venturi V, Zennaro E, Leoni L. 2007. The Pseudomonas quorum-sensing regulator RsaL belongs to the tetrahelical superclass of H-T-H proteins. J Bacteriol 189:1922–1930. doi: 10.1128/JB.01552-06 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38. Nikolskaya AN, Galperin MY. 2002. A novel type of conserved DNA-binding domain in the transcriptional regulators of the AlgR/AgrA/LytR family. Nucleic Acids Res 30:2453–2459. doi: 10.1093/nar/30.11.2453 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39. Rivera-Cancel G, Motta-Mena LB, Gardner KH. 2012. Identification of natural and artificial DNA substrates for light-activated LOV-HTH transcription factor EL222. Biochemistry 51:10024–10034. doi: 10.1021/bi301306t [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40. Kotoky R, Ogawa N, Pandey P. 2022. The structure-function relationship of bacterial transcriptional regulators as a target for enhanced biodegradation of aromatic hydrocarbons. Microbiol Res 262:127087. doi: 10.1016/j.micres.2022.127087 [DOI] [PubMed] [Google Scholar]
  • 41. Schäfer A, Tauch A, Jäger W, Kalinowski J, Thierbach G, Pühler A. 1994. Small mobilizable multi-purpose cloning vectors derived from the Escherichia coli plasmids pK18 and pK19: selection of defined deletions in the chromosome of Corynebacterium glutamicum. Gene 145:69–73. doi: 10.1016/0378-1119(94)90324-7 [DOI] [PubMed] [Google Scholar]
  • 42. van der Geize R, Hessels GI, van Gerwen R, van der Meijden P, Dijkhuizen L. 2002. Molecular and functional characterization of kshA and kshB, encoding two components of 3-ketosteroid 9alpha-hydroxylase, a class IA monooxygenase, in Rhodococcus erythropolis strain SQ1. Mol Microbiol 45:1007–1018. doi: 10.1046/j.1365-2958.2002.03069.x [DOI] [PubMed] [Google Scholar]
  • 43. Reese MG. 2001. Application of a time-delay neural network to promoter annotation in the Drosophila melanogaster genome. Comput Chem 26:51–56. doi: 10.1016/s0097-8485(01)00099-7 [DOI] [PubMed] [Google Scholar]
  • 44. de Avila e Silva S, Echeverrigaray S, Gerhardt GJL. 2011. BacPP: baction—aterial promoter prediction—A tool for accurate sigma-factor specific assignment in enterobacteria. J Theor Biol 287:92–99. doi: 10.1016/j.jtbi.2011.07.017 [DOI] [PubMed] [Google Scholar]
  • 45. Zhu Q, Wei K, Pan K, Li Q, Zhu W, Zhang M, Hong Q. 2025. Comamonas trifloxystrobinivorans sp. nov., encoding a novel esterase responsible for the degradation of trifloxystrobin. Int J Syst Evol Microbiol 75. doi: 10.1099/ijsem.0.006796 [DOI] [PubMed] [Google Scholar]
  • 46. Li N, Chen L, Chen E, Yuan C, Zhang H, He J. 2022. Cloning of a novel tetrahydrofolate-dependent dicamba demethylase gene from dicamba-degrading consortium and characterization of the gene product. Front Microbiol 13:978577. doi: 10.3389/fmicb.2022.978577 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47. Bradford MM. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72:248–254. doi: 10.1016/0003-2697(76)90527-3 [DOI] [PubMed] [Google Scholar]
  • 48. Cheng M, Pei D, He X, Liu Y, Zhu P, Yan X. 2021. The operon encoding hydrolytic dehalogenation of 4-chlorobenzoate is transcriptionally regulated by the TetR-type repressor FcbR and its ligand 4-chlorobenzoyl coenzyme A. Appl Environ Microbiol 87:e02652-20. doi: 10.1128/AEM.02652-20 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49. Zhu S, Qiu J, Wang H, Wang X, Jin W, Zhang Y, Zhang C, Hu G, He J, Hong Q. 2018. Cloning and expression of the carbaryl hydrolase gene mcbA and the identification of a key amino acid necessary for carbaryl hydrolysis. J Hazard Mater 344:1126–1135. doi: 10.1016/j.jhazmat.2017.12.006 [DOI] [PubMed] [Google Scholar]
  • 50. Sambrook J, Russell DW. 2001. Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. [Google Scholar]
  • 51. Kim G, Lee S, Levy Karin E, Kim H, Moriwaki Y, Ovchinnikov S, Steinegger M, Mirdita M. 2025. Easy and accurate protein structure prediction using ColabFold. Nat Protoc 20:620–642. doi: 10.1038/s41596-024-01060-5 [DOI] [PubMed] [Google Scholar]
  • 52. Zhao Y, Xu G, Xu Z, Guo B, Liu F. 2023. LexR positively regulates the LexABC efflux pump involved in self-resistance to the antimicrobial Di- N -oxide phenazine in Lysobacter antibioticus. Microbiol Spectr 11:e0487222. doi: 10.1128/spectrum.04872-22 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53. Trott O, Olson AJ. 2010. AutoDock Vina: improving the speed and accuracy of docking with a new scoring function, efficient optimization, and multithreading. J Comput Chem 31:455–461. doi: 10.1002/jcc.21334 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54. Seeliger D, de Groot BL. 2010. Ligand docking and binding site analysis with PyMOL and Autodock/Vina. J Comput Aided Mol Des 24:417–422. doi: 10.1007/s10822-010-9352-6 [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental material. aem.01277-26-s0001.pdf.

Tables S1 to S3, Fig. S1 to S6, and supplemental text.

aem.01277-26-s0001.pdf (846.4KB, pdf)
DOI: 10.1128/aem.01277-26.SuF1

Data Availability Statement

The genome sequence of Rhodococcus sp. D-6 has been deposited in the GenBank database under the accession numbers CP132970 to CP132972 (11).

Articles from Applied and Environmental Microbiology are provided here courtesy of American Society for Microbiology (ASM)

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