Transcriptional regulation of two redundant 3-bromo-4-hydroxybenzoate catabolic operons via two different regulatory modes in Pigmentiphaga kullae strain H8

TL;DR

This study reveals how two redundant gene clusters in a bacterium are regulated differently to help it adapt to complex environments.

Contribution

The paper identifies two distinct regulatory mechanisms for redundant catabolic operons in Pigmentiphaga kullae strain H8.

Findings

BhbR1 activates transcription by shortening its DNA binding region in response to 3-Br-4-HB.

BhbR2 activates transcription by binding a palindromic sequence without effector-induced changes.

Both regulators show reduced activation at high 3-Br-4-HB concentrations, emphasizing the need for redundancy.

Abstract

Gene redundancy endows bacteria with enhanced adaptability to complex and fluctuating environments but results in genetic costs. Transcriptional regulation is considered an effective strategy for harmonizing adaptive benefits with physiological burdens. In our previous study, two redundant gene clusters (phbh1pcaApcaBorf404bhbR1 and phbh2pcaB2pcaA2bhbR2) involved in 3-bromo-4-hydroxybenzoate (3-Br-4-HB) catabolism were identified in Pigmentiphaga kullae strain H8. The LysR-type transcription regulator BhbR1, encoded by the bhbR1 gene, activated phbh1pcaApcaBorf404 transcription. Through DNase I footprinting assays, the presence of the inducer 3-Br-4-HB was found to shorten the BhbR1-bound region in the promoter, uncovering the protected -35 box, thereby activating transcription. The MarR-family transcription factor (MFTF) BhbR2, encoded by the bhbR2 gene, was different from typical inhibitive MFTFs and activated phbh2pcaB2pcaA2 transcription. BhbR2 was found to bind a 17-bp imperfect palindromic sequence (TTGATT-N5-AATCAA) in the target promoter. Intriguingly, the presence of 3-Br-4-HB neither dissociated BhbR2 from the promoter nor modified its binding site, indicating a novel regulatory mode. Despite a coincident trend in activating their respective operons in response to different concentrations of 3-Br-4-HB, BhbR1 and BhbR2 both showed a significant attenuation of the activation effect at high concentrations (>480 μM), highlighting the requisite co-existence of redundant 3-Br-4-HB catabolic operons and their regulatory genes. This study presents two distinct transcriptional regulation mechanisms of these two redundant 3-Br-4-HB catabolic operons in strain H8, expanding our understanding of the diversity of transcriptional regulation for enhancing adaptation.

In bacteria, catabolic genes for pollutant degradation often possess functionally redundant duplicates, providing a genetic basis for rapid adaptation to complex polluted environments. Synergic regulation plays an important role in balancing the physiological burden of extra genetic material with the adaptive benefits conferred by genetic redundancy. Although the co-existence of two redundant 3-bromo-4-hydroxybenzoate (3-Br-4-HB)-catabolic operons has been proven to enhance the metabolic robustness and adaptability of the host strain Pigmentiphaga kullae H8, how these two inducible catabolic operons are regulated remains unclear. This study identified two regulators, the LysR-type transcription regulator BhbR1 and the MarR-family transcription factor BhbR2, which activated transcription of the two 3-Br-4-HB-catabolic operons using different modes, and also revealed interactions of these two regulators with their effectors and target promoters. These findings not only clarify two distinct transcriptional strategies employed by redundant catabolic operons but also enhance our understanding of the significance of regulatory diversity for bacterial adaptation to complex polluted environments.

Figures

Organization and transcriptional analysis of two 3-Br-4-HB-degrading gene clusters (bhb1 and bhb2) in Pigmentiphaga kullae strain H8. (A) Degradation pathway of 3-Br-4-HB. (B) Schematic diagram illustrating the genetic organization of the bhb1 and bhb2 clusters. Two promoters Pphbh1 and PpcaA2, corresponding to two operons phbh1pcaApcaBorf404 and phbh2pcaB2pcaA2, are indicated by bent arrows. The amplification fragments (RT1-RT8) for transcriptional unit assessment are shown as black lines under the bhb1 and bhb2 clusters. (C) Transcriptional unit analysis of the bhb1 and bhb2 clusters. Total RNA and cDNA were used as templates for PCR amplification of eight fragments (RT1–RT8, as shown in panel B), which were then detected by electrophoresis. (D) Organization of the intergenic region between orf408 and pbhb1. (E) Organization of the intergenic region between bhbR2 and pcaA2. Start codons (ATG) are indicated by arrows, and the putative -10 boxes and -35 boxes are underlined. Transcriptional start sites are indicated by bent arrows, with the direction of the arrows indicating the direction of gene transcription.

Roles of BhbR1 and BhbR2 in 3-Br-4-HB degradation and transcriptional regulation of degradation genes. (A) Degradation of 3-Br-4-HB by strain H8Δphbh2, the bhbR1-mutant H8Δphbh2ΔbhbR1, and the bhbR1-complemented strain H8Δphbh2ΔbhbR1 (pBBR-bhbR1). (B) Relative transcription analysis of phbh1, pcaB, and bhbR1 in 3-Br-4-HB-induced and -uninduced cells of strains H8Δphbh2, H8Δphbh2ΔbhbR1, and H8Δphbh2ΔbhbR1 (pBBR-bhbR1). (C) Degradation of 3-Br-4-HB by strain H8Δphbh1, the bhbR2-mutant H8Δphbh1ΔbhbR2, and the bhbR2-complemented strain H8Δphbh1ΔbhbR2 (pBBR-bhbR2). (D) Relative transcription analysis of phbh2, pcaB2, and bhbR2 in 3-Br-4-HB-induced and -uninduced cells of strains H8Δphbh1, H8Δphbh1ΔbhbR2, and H8Δphbh1ΔbhbR2 (pBBR-bhbR2). Error bars represent the standard deviations of three replicates.

Identification of inducers recognized by BhbR1 and BhbR2 and the effect of 3-Br-4-HB concentration on target gene transcription. (A) Transcription analysis of phbh1 and pcaB in strain H8Δphbh2, and phbh2 and pcaB2 in strain H8Δphbh1, was conducted in the presence or absence of 3-Br-4-HB, Br-PCA, 4-HB, or DBHB. (B) The β-galactosidase activity assay was conducted in strains H8Δphbh2 (pME-PbR1) and H8Δphbh1 (pME-PbR2) upon induction with 0.2 mM of 3-Br-4-HB, Br-PCA, DBHB, and 4-HB. (C) Transcription analysis of target genes pcaB and pcaB2 in strain H8Δphbh1Δphbh2 was performed in response to varying concentrations of 3-Br-4-HB (0, 30, 60, 120, 240, 480, 720, and 960 µM). Error bars represent the standard deviations of three replicates. Different letters indicate differences as determined by one-way ANOVA (P < 0.05).

Identification of the promoter binding sites of BhbR1 and BhbR2. (A) EMSA of BhbR1 binding with the promoter DNA probe PbR1 (A) and BhbR2 binding with the promoter DNA probe PbR2 (B). “−” indicates samples incubated without 3-Br-4-HB, while “+” indicates samples incubated with 0.1 mM 3-Br-4-HB. Each lane contained 35 nM of promoter DNA probe. A 152-bp DNA fragment amplified from the 16S rRNA gene of strain H8 was used as a control DNA probe (28 nM). Protein-DNA complexes are indicated by brackets. DNase I footprinting assays of BhbR1 binding with the probe PbR1 in the absence (C) or presence (E) of 50 µM 3-Br-4-HB, and BhbR2 binding with the probe PbR2 in the absence (D) or presence (F) of 50 µM 3-Br-4-HB. The 6-carboxyfluorescein (FAM)-labeled DNA probe (350 ng) was incubated with 5 µg of BhbR1 or BhbR2, indicated by blue lines, as well as incubated without BhbR1 or BhbR2, indicated by red lines. Regions protected by BhbR1 or BhbR2 from DNase I cleavage are indicated with black dotted boxes, and the sequences of protected regions are shown at the bottom. (G) DNA elements in the promoters Pphbh1 and PpcaA2. The promoter regions protected by BhbR1 and BhbR2 in the absence (−) or presence (+) of 3-Br-4-HB are marked with light gray stripes. The regulatory binding sites for BhbR1 (ATA-N9-TAT) and BhbR2 (TTGATT-N5-AATCAA) are framed with red dashed lines. Two putative low-affinity activation binding sites (ABS1 and ABS2) in the promoter Pphbh1 are framed with blue dashed lines. The putative -10 and -35 boxes are underlined by black dashed and solid lines, respectively. TSSs are indicated by orange bent arrows, with the direction of the arrows indicating the direction of gene transcription.

SPR assays for binding dynamics of the regulators with the inducer and target promoters. SPR assays analyzing interactions of the inducer 3-Br-4-HB with regulators BhbR1 (A) and BhbR2 (B). A serial dilution of 3-Br-4-HB (0.39–6.25 µM) was flowed over the CM7 sensor chip. SPR assays of BhbR1 binding with the fragments Fphbh1 in the absence (C) or presence (E) of 5 µM 3-Br-4-HB, and BhbR2 binding with the fragments FpcaA2 in the absence (D) or presence (F) of 5 µM 3-Br-4-HB. Biotinylated fragments Fphbh1 and FpcaA2 were captured on an SA sensor chip, while nonspecific biotinylated fragments F16S, derived from the 16S rRNA gene of strain H8, served as a control. Various concentrations of BhbR1 (68.75–1,100 nM) and BhbR2 (31.25–500 nM) were passed over the immobilized DNA fragments. The sensor surface was regenerated after each cycle using a short treatment with 0.5% (wt/vol) SDS. Derived dissociation constants (KD) are displayed above the equilibrium binding curves.

Tables

TABLE 1. Strains and plasmids used in this study

Strains and plasmids	Relevant characteristicsa	Source or reference
Pigmentiphaga kullae strains
H8	3-Br-4-HB-degrading strain, wild type; Sper, Strr	(21)
H8Δphbh1	Mutant of strain H8 with phbh1 knockout; Sper, Strr	(21)
H8Δphbh2	Mutant of strain H8 with phbh2 knockout; Sper, Strr	(21)
H8Δphbh2ΔbhbR1	Double mutant of strain H8 with phbh2 and bhbR1 knockout; Sper, Strr	This study
H8Δphbh1ΔbhbR2	Double mutant of strain H8 with phbh1 and bhbR2 knockout; Sper, Strr	This study
H8Δphbh2ΔbhbR1 (pBBR-bhbR1)	Mutant H8Δphbh2ΔbhbR1 complemented with bhbR1 gene by harboring the plasmid pBBR-bhbR1; Sper, Strr, Kmr	This study
H8Δphbh1ΔbhbR2 (pBBR-bhbR2)	Mutant H8Δphbh1ΔbhbR2 complemented with bhbR2 gene by harboring the plasmid pBBR-bhbR2; Sper, Strr, Kmr	This study
H8Δphbh1Δphbh2	Double mutant of strain H8 with phbh1 and phbh2 knockout; Sper, Strr	(21)
H8Δphbh2 (pME-PbR1)	Mutant H8Δphbh2 harboring the plasmid pME-PbR1; Sper, Strr, Tcr	This study
H8Δphbh1 (pME-PbR2)	Mutant H8Δphbh1 harboring the plasmid pME-PbR2; Sper, Strr, Tcr	This study
E. coli strains
DH5α	Host strain for cloning vectors	Lab stored
HB101 (pRK2013)	Conjugation helper strain; Kmr	Lab stored
BL21(DE3)	Host strain for expression vectors	Lab stored
Plasmids
pBBR1MCS-2	Broad host range vector; Kmr	(50)
pBBR-bhbR1	The bhbR1 gene cloned into pBBR1MCS-2; Kmr	This study
pBBR-bhbR2	The bhbR2 gene cloned into pBBR1MCS-2; Kmr	This study
pJQ200SK	Suicide vector; P15A, ori, sacB, RP4, (pBluescriptSK); Gmr	Lab stored
pJQ-ΔbhbR1	pJQ200SK containing the upstream and downstream fragments of bhbR1 gene for the gene targeting of bhbR1 gene; Gmr	This study
pJQ-ΔbhbR2	pJQ200SK containing the upstream and downstream fragments of bhbR2 gene for the gene targeting of bhbR2 gene; Gmr	This study
pET29a(+)	Expression vector; Kmr	Lab stored
pET-bhbR1	The bhbR1 gene cloned into pET29a(+); Kmr	This study
pET-bhbR2	The bhbR2 gene cloned into pET29a(+); Kmr	This study
pMD19-T	TA clone vector; Ampr	TaKaRa
pMD-PbR1	pMD19-T harboring the fragment of DNA probe PbR1; Ampr	This study
pMD-PbR2	pMD19-T harboring the fragment of DNA probe PbR2; Ampr	This study
pME6522	Vector for promoter-lacZ transcriptional fusions; Tcr	(35)
pME-PbR1	pME6522 harboring the DNA fragment PbR1 containing the promoter Pphbh1; Tcr	This study
pME-PbR2	pME6522 harboring the DNA fragment PbR2 containing the promoter PpcaA2; Tcr	This study

TABLE 2. Primers used in this study

Primers	Sequence (5′−3′)a	Purpose
Gene knockout, complementation, and overexpression
bhbR1-kn1 bhbR1-kn2	CAATCTAGACTGCTGGCCAACATGGTGGA (XbaI) GTCTGCAACAGGGGCGTGCTGCAGTTCGACCTTGGGGAAG	Amplification of the upstream homologous arm of bhbR1 for gene knockout
bhbR1-kn3 bhbR1-kn4	CTTCCCCAAGGTCGAACTGCAGCACGCCCCTGTTGCAGACCAACTCGAGTGTGGGTGCACAAGGACACG (XhoI)	Amplification of the downstream homologous arm of bhbR1 for gene knockout
bhbR1-f bhbR1-r	GTAGAATTCATCATGGCGGCTTCGACAA (EcoRI)GAAGGTACCAACCGCGGGCCCTTCTA (KpnI)	Amplification of bhbR1 for gene complementation
bhbR1-29f bhbR1-29r	GCACATATGACGCTGCAGCAACTGCGCG (NdeI)GCACTCGAGCCGGCCCATGCCGGCGAT (Xho I)	Amplification of bhbR1 gene for cloning into pET29a(+) vector
bhbR2-kn1 bhbR2-kn2	CAATCTAGAGTGCAGGAGCTGATACAG (XbaI)CCTTCCGAACTGGCCGAACGGGAAAGAATCCGGGAGCT	Amplification of the upstream homologous arm of bhbR2 for gene knockout
bhbR2-kn3 bhbR2-kn4	CGAGCTCCCGGATTCTTTCCCGTTCGGCCAGTTCGGAACAAGGGCCCGTCAGACCGCTCCCTTGT (ApaI)	Amplification of the downstream homologous arm of bhbR2 for gene knockout
bhbR2-f bhbR2-r	GCAGGTACCTTGGGTAGAGGGAAAA (KpnI)CAAGGATCCTACCAGACGCCCGCCAGGT (BamHI)	Amplification of bhbR2 for gene complementation
bhbR2-29f bhbR2-29r	GCACATATGTCCTTCGAGCAGCCCGCAA (NdeI)GCACTCGAGCCAGACGCCCGCCAGGTC (XhoI)	Amplification of bhbR2 gene for cloning into pET29a(+) vector
Transcriptional organization
RT-F1 RT-R1	TGGGCGTGGTGGCCCGCAAGG CTTCCCGAGGTGCCCACGCTC	Amplification of 724 bp of bhbR1-orf404-spanning region
RT-F2 RT-R2	CACCTGGTTGAAGGTGGCCAC TCGTCGGCACCGGCGGCA	Amplification of 685 bp of orf404-pcaB-spanning region
RT-F3 RT-R3	CAGCGGCACGACCTTGATGTT GGGCATAGACGGCACCTATGT	Amplification of 780 bp of pcaB-pcaA-spanning region
RT-F4 RT-R4	TAGAGCCCGGCTCCCACCGTC CCTGAACGAAGGCCGCATCGT	Amplification of 734 bp of pcaA-phbh1-spanning region
RT-F5 RT-R5	CGTTCGGCCAGTTCGGAA CAGCTTGAGCAGGAAGTA	Amplification of 592 bp of bhbR2-pcaA2-spanning region
RT-F6 RT-R6	ACGAAGAGGCCTACATGA CGTCGTAGACCAGCGATT	Amplification of 556 bp of pcaA2-pcaB2-spanning region
RT-F7 RT-R7	GCAAGATGGACCAGGACT CACTTCATGCTGGGGATA	Amplification of 553 bp of pcaB2-phbh2-spanning region
RT-F8 RT-R8	GGTGATGAAGACCTACGA CCAATTGCGCAAGGTGTT	Amplification of 757 bp of phbh2-orf3312-spanning region
RT-qPCR
Qphbh1-f Qphbh1-r	CATGGTCCACGAAGGCATA CAGGTCCTTGACGATTTCA	Amplification of a 121-bp fragment of phbh1 for RT-qPCR
QpcaB-f QpcaB-r	CATAGGCAGTTCGCACGTT GTACTTCGTTCTCCAGCCA	Amplification of a 125-bp fragment of pcaB for RT-qPCR
Qphbh2-f Qphbh2-r	CGGTGATGAAGACCTACGA GAAGCTCAGCATGGCGAAG	Amplification of a 125-bp fragment of phbh2 for RT-qPCR
QpcaB2-f QpcaB2-r	ACCTGGAATTCTCGCAGCA GTATGGTCGAAGCACAGGT	Amplification of a 124-bp fragment of pcaB2 for RT-qPCR
QbhbR1-f QbhbR1-r	GGTCATGGCGCTGGTCAAT AGGCAGATCTCGAAGCTGT	Amplification of a 132-bp fragment of bhbR1 for RT-qPCR
QbhbR2-f QbhbR2-r	GTTCAGCATGTCGTCGAGC CATCGAACTGACCGACAAG	Amplification of a 127-bp fragment of bhbR2 for RT-qPCR
Q16S-f Q16S-r	GTTCTTCGGAGCTTGGTAG ACATCATCCACCGCTTGTG	Amplification of a 115-bp fragment of 16S rRNA gene for RT-qPCR
5′-RACE
phbh1-GSP1 phbh1-GSP2 phbh1-GSP3	GTTTCGCCTTCCTGCTCGAAGC CGATTTCATGCTGGGGATAGA CGAAGCGTATGTCTATGCCTT	5′-RACE of the operon phbh1pcaApcaBorf404
pcaA2-GSP1 pcaA2-GSP2 pcaA2-GSP3	CGTGGTCGTTGTAGACCATGAT ATGGAGGGCACGTGAGAAGTA GTCAGACCGCTCCCTTGTCGT	5′-RACE of the operon pcaA2pcaB2phbh2
EMSA and DNase I footprinting assays
PbR1-f PbR1-r	GAACTCGAGGTGCTGTCGATTTGTGCCA (XhoI)GAAACTAGTCGTGGGTCGTAGTTCCGT (SpeI)	Amplification of DNA probe PbR1 (243 bp) containing the promoter Pphbh1 for EMSA
PbR2-f PbR2-r	GAACTCGAGTGATGCCGTAGCGGCCTT (XhoI)GAAACTAGTGGGCAATGCTCCTTGGGTA (SpeI)	Amplification of DNA probe PbR2 (258 bp) containing the promoter PpcaA2 for EMSA
Pb16S-f Pb16S-r	GCGGCGAGTGGCGAACGGGTGAG TAATCCGACATCGGCCGCTCCAA	Amplification of a nonspecific probe Pb16S (152 bp) from 16S rRNA gene of strain H8 as control for EMSA
FAM-M13F M13R	GTAAAACGACGGCCAGT (5′-FAM labeled)CAGGAAACAGCTATGAC	Amplification of DNA probes for DNase I footprinting assays
SPR assay
Fpb1-Btn-f Fpb1-r	GTCGTAGTTCCGTTGCCGGG (5′-biotin labeled)ACCAGCAAGGAACGTGCCCC	Amplification of biotinylated DNA fragments Fpbhb1 (144 bp) for SPR
Fpb2-f Fpb2-Btn-r	CGAAGGACATGGCTGCTCCTGGGCAATGCTCCTTGGGTAG (5′-biotin labeled)	Amplification of biotinylated DNA fragments FpcaA2 (153 bp) for SPR
F16S-Btn-f F16S-r	GCGGCGAGTGGCGAACGGGTGAG (5′-biotin labeled)TAATCCGACATCGGCCGCTCCAA	Amplification of nonspecific biotinylated DNA fragments F16S (152 bp) for SPR
β-galactosidase activity assay
pME-PbR1-F	ATCGGAATTCGTGCTGTCGATTTGTGCCA (EcoRI)	Amplification of the DNA fragment PbR1 (243 bp) containing the promoter Ppbhb1 for constructing pME-PbR1
pME-PbR1-R	ATCGCTGCAGCGTGGGTCGTAGTTCCGT (PstI)
pME-PbR2-F	ATCGGAATTCTGATGCCGTAGCGGCCTT (EcoRI)	Amplification of the DNA fragment PbR2 (258 bp) containing the promoter PpcaA2 for constructing pME-PbR2
pME-PbR2-R	ATCGCTGCAGGGGCAATGCTCCTTGGGTA (PstI)

Full text

INTRODUCTION

Bacteria play crucial roles in natural biogeochemical cycles, particularly in the degradation of environmental xenobiotics. They can metabolize various pollutants, including organohalogen compounds, polycyclic aromatic hydrocarbons, pesticides, plastics, and petroleum hydrocarbons, by expressing specific degrading enzymes (1–5). However, dynamic changes in the types and concentrations of pollutants in the environment necessitate effective strategies for regulating the expression of these genes. Transcriptional regulation serves as a fundamental mechanism to directly and effectively control gene expression (6, 7). It enables bacteria to quickly respond to specific compounds, control the expression of specialized catabolic genes, efficiently utilize these compounds as energy and carbon sources, and cope with nutritional requirements and toxicity pressures imposed by pollutants (8). Therefore, transcriptional regulation is critical for bacterial adaptation to complex and ever-changing polluted environments, broadening their ecological niches.

Genetic redundancy is a common occurrence across all three kingdoms of life (9). Intriguingly, species in highly disturbed habitats tend to have increased numbers of redundant genes compared to those in stable settings (10, 11). An increasing number of studies have indicated that genetic redundancy can confer hosts with adaptive benefits for coping with environmental disturbances, including buffering gene mutations, increasing protein dosage and function, and acquiring novel adaptive traits through neo- and sub-functionalization (9, 12–14). Despite its benefits, the expansion of genetic material imposes an additional physiological burden on the host. For instance, it consumes cellular resources and energy for gene replication, transcription, and translation, potentially compromising competitive fitness (15). Bacterial genomes often contain redundant genes for degrading pollutants, though their transcription varies in response to diverse environmental and physiological stimuli (16–19). Although experimental evidence is scarce, transcriptional regulation is intuitively believed to compensate for the physiological costs caused by genetic redundancy. Understanding the regulation mechanisms of these redundant genes that confer selective advantages and improve host adaptability to environmental disturbances has attracted widespread interest.

In our previous study, Pigmentiphaga kullae strain H8, a bacterium capable of efficiently degrading 3-bromo-4-hydroxybenzoate (3-Br-4-HB), was isolated from soil highly contaminated with halogenated hydroxybenzoate (20, 21). Two redundant gene clusters, designated as bhb1 (phbh1pcaApcaBorf404bhbR1) and bhb2 (phbh2pcaB2pcaA2bhbR2), were confirmed to be involved in 3-Br-4-HB catabolism at both physiological and genetic levels (Fig. 1A and B). Notably, the redundancy of bhb1 and bhb2 clusters not only enables the host to metabolize anthropogenic 3-Br-4HB but also helps it overcome the toxicity and metabolic disturbance caused by high concentrations of 3-Br-4-HB, enhancing its adaptation to habitats highly contaminated with 3-Br-4-HB (21). The transcription of genes for key degradation enzymes, such as phbh1 and phbh2, is inducible; however, their transcriptional regulation mechanisms remain unclear.

Organization and transcriptional analysis of two 3-Br-4-HB-degrading gene clusters (bhb1 and bhb2) in Pigmentiphaga kullae strain H8. (A) Degradation pathway of 3-Br-4-HB. (B) Schematic diagram illustrating the genetic organization of the bhb1 and bhb2 clusters. Two promoters Pphbh1 and PpcaA2, corresponding to two operons phbh1pcaApcaBorf404 and phbh2pcaB2pcaA2, are indicated by bent arrows. The amplification fragments (RT1-RT8) for transcriptional unit assessment are shown as black lines under the bhb1 and bhb2 clusters. (C) Transcriptional unit analysis of the bhb1 and bhb2 clusters. Total RNA and cDNA were used as templates for PCR amplification of eight fragments (RT1–RT8, as shown in panel B), which were then detected by electrophoresis. (D) Organization of the intergenic region between orf408 and pbhb1. (E) Organization of the intergenic region between bhbR2 and pcaA2. Start codons (ATG) are indicated by arrows, and the putative -10 boxes and -35 boxes are underlined. Transcriptional start sites are indicated by bent arrows, with the direction of the arrows indicating the direction of gene transcription.

Here, a LysR-type transcriptional regulator BhbR1, encoded by a regulatory gene bhbR1, and a MarR-type transcriptional regulator BhbR2, encoded by a regulatory gene bhbR2, were found to be involved in the catabolism of 3-Br-4-HB in strain H8. The regulatory mechanisms of BhbR1 and BhbR2 for activating the transcription of two 3-Br-4-HB catabolic operons were analyzed at both physiological and genetic levels. This study will expand our understanding of the diversity of transcriptional regulation of redundant genes.

RESULTS

Transcriptional analysis of two 3-Br-4-HB-degrading gene clusters

Strain H8 harbors two spatially separate but functionally redundant 3-Br-4-HB-degrading gene clusters, bhb1 and bhb2, located approximately 3 Mb apart on its genome (Fig. 1B). Reverse transcription PCR analysis of the intergenic regions between two adjacent open reading frames (orfs) was performed to delineate the transcriptional units of these clusters. Using the total RNA extracted from 3-Br-4-HB-induced cells of strain H8 and its reverse-transcribed cDNA as templates, three distinct segments, RT2, RT3, and RT4, were successfully amplified from the bhb1 cluster (Fig. 1C), indicating that the bhb1 cluster is organized into two operons: phbh1pcaApcaBorf404 and bhbR1. Amplification of two segments (RT6 and RT7, Fig. 1C) from the bhb2 cluster suggested that bhb2 comprised two operons: phbh2pcaB2pcaA2 and bhbR2.

To determine the promoter regions of 3-Br-4-HB-catabolic operons, phbh1pcaApcaBorf404 and phbh2pcaB2pcaA2, transcriptional start sites (TSSs) were identified by 5′ rapid amplification of cDNA ends (5′-RACE) using total RNA extracted from 3-Br-4-HB-induced cells of strain H8. The sequencing results of 5′-RACE are shown in the supplemental material (Fig. S1). As shown in Fig. 1D and E, the TSSs were pinpointed at a G residue 44 bp upstream of the phbh1 translational start codon and a C residue 79 bp upstream of the pcaA2 translational start codon, respectively. The software BPROM (http://www.softberry.com/berry.phtml?topic=bprom&group=programs&subgroup=gfindb) was utilized to predict the -10 and -35 boxes of promoters. The predicted -10 (CGTGATAC) and -35 (TTTTCG) boxes of the phbh1 promoter (designated P_phbh1_), and the predicted -10 (AGCTAGAGT) and -35 (TTGATT) boxes of the pcaA2 promoter (designated P_pcaA2_) are shown in Fig. 1D and E, respectively.

We applied the RhoThermPredict algorithm to identify Rho-dependent terminators within the bhb1 and bhb2 gene clusters (22). In the phbh1pcaApcaBorf404 operon*,* a Rho utilization site (RUT site) was predicted downstream of orf404 (155–233 nt from the stop codon), followed by a PAUSE-CONSENSUS sequence (CAACGGTCTGC, 68–79 nt downstream of the RUT site) and a palindromic sequence (GGCCGCCGGGGCGGCC, nt 30–46) (italics denote a palindromic sequence) further downstream (Fig. S2A). Similarly, in the phbh2pcaB2pcaA2 operon*,* a RUT site was predicted downstream of phbh2 (+2 to −73 nt from the stop codon), accompanied by a PAUSE-CONSENSUS sequence (CGTCCGAGCCGA) and a palindromic sequence (GCGGCAGGGCCGC, nt 27–40) further downstream (Fig. S2B).

BhbR1 and BhbR2 positively regulate the transcription of phbh1pcaApcaBorf404 and phbh2pcaB2pcaA2, respectively

Two transcriptional regulatory genes, bhbR1 and bhbR2, were found in the bhb1 and bhb2 clusters, respectively. BhbR1, encoded by the bhbR1 gene and composed of 302 amino acids, showed the highest identity with the LysR-type transcriptional regulator (LTTR) TsaR (34.2%) from Comamonas testosterone (23). Phylogenetic analysis of BhbR1 and its closely related members showed that BhbR1, TsaR, GlaR, and TdcA formed a subclade from other LTTRs (Fig. S3A), indicating that BhbR1 is a new member of the LTTR family. BhbR2 is encoded by the bhbR2 gene and consists of 179 amino acids; it was found to be most closely related to members of the MarR (multiple antibiotic resistance regulatory factor) family of transcriptional factors (MFTFs). Phylogenetic analysis revealed that BhbR2 is a new member of the MarR family, and the majority of MFTFs phylogenetically related to BhbR2 have been characterized as transcriptional repressors, including CbaR, EmrR, and HucR (Fig. S3B) (24–26).

Because both hydroxylases PHBH1 and PHBH2 catalyze the initial reaction of 3-Br-4-HB degradation in strain H8 (21), double-mutant strains, H8Δphbh2ΔbhbR1 and H8Δphbh1ΔbhbR2, and their corresponding complemented strains, H8Δphbh2ΔbhbR1 (pBBR-bhbR1) and H8Δphbh1ΔbhbR2 (pBBR-bhbR2), were constructed to determine the transcriptional regulatory roles of BhbR1 and BhbR2. The degradation assay showed that the bhbR1-mutant strain H8Δphbh2ΔbhbR1 completely lost the capacity to degrade 3-Br-4-HB, whereas complementation with bhbR1 in the mutant strain recovered this ability (Fig. 2A). Additionally, real-time quantitative PCR (RT-qPCR) analyses showed that in the presence of 3-Br-4-HB, transcription of target genes phbh1 and pcaB was not activated in the mutant H8Δphbh2ΔbhbR1; contrastingly, the transcription levels of phbh1 and pcaB significantly increased in strain H8Δphbh2 and the bhbR1-complemented strain H8Δphbh2ΔbhbR1 (pBBR-bhbR1) following 3-Br-4-HB induction (Fig. 2B). These results suggest that BhbR1 is involved in the catabolism of 3-Br-4-HB and activates the transcription of the phbh1pcaApcaBorf404 operon in strain H8. Unlike typical LTTRs, BhbR1 also activates its own gene transcription, although to a lesser extent compared to the target genes (Fig. 2B).

Roles of BhbR1 and BhbR2 in 3-Br-4-HB degradation and transcriptional regulation of degradation genes. (A) Degradation of 3-Br-4-HB by strain H8Δphbh2, the bhbR1-mutant H8Δphbh2ΔbhbR1, and the bhbR1-complemented strain H8Δphbh2ΔbhbR1 (pBBR-bhbR1). (B) Relative transcription analysis of phbh1, pcaB, and bhbR1 in 3-Br-4-HB-induced and -uninduced cells of strains H8Δphbh2, H8Δphbh2ΔbhbR1, and H8Δphbh2ΔbhbR1 (pBBR-bhbR1). (C) Degradation of 3-Br-4-HB by strain H8Δphbh1, the bhbR2-mutant H8Δphbh1ΔbhbR2, and the bhbR2-complemented strain H8Δphbh1ΔbhbR2 (pBBR-bhbR2). (D) Relative transcription analysis of phbh2, pcaB2, and bhbR2 in 3-Br-4-HB-induced and -uninduced cells of strains H8Δphbh1, H8Δphbh1ΔbhbR2, and H8Δphbh1ΔbhbR2 (pBBR-bhbR2). Error bars represent the standard deviations of three replicates.

Mutation of bhbR2 in the strain H8Δphbh1ΔbhbR2 completely abolished 3-Br-4-HB degradation. The degradation ability was restored in the bhbR2-complemented strain H8Δphbh1ΔbhbR2 (pBBR-bhbR2), indicating the involvement of BhbR2 in the catabolism of 3-Br-4-HB in strain H8 (Fig. 2C). Furthermore, the bhbR2-mutant strain H8ΔbhbR2Δphbh1 did not upregulate transcription of the target genes phbh2 and pcaB2 with 3-Br-4-HB induction. However, significant transcriptional upregulation of the target genes was observed in the bhbR2-harboring strain H8Δphbh1 and the bhbR2-complemented strain H8Δphbh1ΔbhbR2 (pBBR-bhbR2) (Fig. 2D). These findings suggest that BhbR2 is an essential activator for the transcription of the phbh2pcaA2pcaB2 operon in response to 3-Br-4-HB.

Characterization of inducible effectors recognized by BhbR1 and BhbR2

To identify the inducers that activate target gene transcription by BhbR1 and BhbR2, we analyzed the transcription levels of phbh1, pcaB, phbh2, and pcaB2 in the presence of 3-Br-4-HB, the intermediate metabolite 3-bromo-4,5-dihydroxybenzoate (Br-PCA), and the structural analogs 3,5-dibromo-4-hydroxybenzoate (DBHB) and 4-hydroxybenzoate (4-HB) by RT-qPCR. As shown in Fig. 3A, transcription levels of four target genes were significantly upregulated under 3-Br-4-HB induction, indicating that 3-Br-4-HB is an inducer for both BhbR1 and BhbR2. In addition, transcription of phbh1 and pcaB showed moderate upregulation relative to the control when induced with Br-PCA and DBHB, suggesting that BhbR1 has non-specific inducer recognition. The presence of 4-HB did not trigger any notable transcriptional response.

Identification of inducers recognized by BhbR1 and BhbR2 and the effect of 3-Br-4-HB concentration on target gene transcription. (A) Transcription analysis of phbh1 and pcaB in strain H8Δphbh2, and phbh2 and pcaB2 in strain H8Δphbh1, was conducted in the presence or absence of 3-Br-4-HB, Br-PCA, 4-HB, or DBHB. (B) The β-galactosidase activity assay was conducted in strains H8Δphbh2 (pME-PbR1) and H8Δphbh1 (pME-PbR2) upon induction with 0.2 mM of 3-Br-4-HB, Br-PCA, DBHB, and 4-HB. (C) Transcription analysis of target genes pcaB and pcaB2 in strain H8Δphbh1Δphbh2 was performed in response to varying concentrations of 3-Br-4-HB (0, 30, 60, 120, 240, 480, 720, and 960 µM). Error bars represent the standard deviations of three replicates. Different letters indicate differences as determined by one-way ANOVA (P < 0.05).

The β-galactosidase activity assay was conducted to further consolidate the recognition of inducers by BhbR1 and BhbR2. Strains H8Δphbh2 (pME-PbR1) and H8Δphbh1 (pME-PbR2) were induced with 0.2 mM concentrations of 3-Br-4-HB, Br-PCA, DBHB, and 4-HB, respectively. As shown in Fig. 3B, for strain H8Δphbh2 (pME-PbR1), β-galactosidase activity increased 51.7-fold with 3-Br-4-HB, 38.2-fold with Br-PCA, and 34.9-fold with DBHB, relative to non-induced controls. In contrast, β-galactosidase activity in H8Δphbh1 (pME-PbR2) was significantly increased only with 3-Br-4-HB as an inducer, with a 145.6-fold increase over non-induced cells. These results align with RT-qPCR data, further demonstrating that 3-Br-4-HB is a shared inducer for both BhbR1 and BhbR2, while BhbR1 also recognizes Br-PCA and DBHB.

To investigate the effect of inducer concentration on the activity of regulators, the transcriptional differences of target genes in the two operons induced by different concentrations of 3-Br-4-HB (0, 30, 60, 120, 240, 480, 720, and 960 µM) were further analyzed. Simultaneously, to avoid the fluctuation of inducer concentration caused by the degradation of 3-Br-4-HB, transcriptional analysis of target genes pcaB and pcaB2 was performed using the double mutant strain H8*∆phbh1∆phbh2* that had complete loss of 3-Br-4-HB-degrading ability. As shown in Fig. 3C, pcaB and pcaB2 exhibited the strongest transcriptional activity at inducer concentrations ranging from 60 to 240 µM and 30 to 480 µM, respectively. However, there was a significant decrease in transcriptional activity for both pcaB and pcaB2 as the inducer concentration increased; when the inducer concentration reached 960 µM, the transcription of pcaB and pcaB2 was only upregulated by 438- and 348-fold, respectively. These data suggest that high concentrations of 3-Br-4-HB induce physiological toxicity, affecting gene transcription.

Identification of the binding sites of promoter regions by BhbR1 and BhbR2

The C-terminal His-tagged BhbR1 and BhbR2 were over-expressed in Escherichia coli BL21 (DE3), purified using Ni-NTA affinity chromatography, and verified by SDS-PAGE (Fig. S4). Gel filtration analysis revealed that active BhbR1 exists as a tetramer in PBS (8.0 g NaCl, 0.2 g KCl, 1.5 g Na_2_HPO_4_, 0.24 g KH_2_PO_4_, and 1 L deionized water, pH 7.8) (Fig. S5B), whereas BhbR2 forms a dimer (Fig. S5D).

Electrophoretic mobility shift assays (EMSAs) were performed to determine the interactions between the regulators (BhbR1 and BhbR2) and their target promoter DNA probes: PbR1 (a 243-bp DNA fragment containing the promoter P_phbh1_) and PbR2 (a 253-bp DNA fragment containing the promoter P_pcaA2_). As shown in Fig. 4A and B, regardless of the presence of the inducer 3-Br-4-HB, both BhbR1 and BhbR2 bound to their corresponding target promoter probes and formed protein-DNA complexes, BhbR1-PbR1 and BhbR2-PbR2, delaying the migration of the two promoter probes. In contrast, nonspecific DNA probe Pb16S, a 152-bp 16S rRNA gene fragment from strain H8, showed no interaction with either regulator; this confirmed the specificity of BhbR1 and BhbR2 for their target regions of P_phbh1_ and P_pcaA2_, respectively.

Identification of the promoter binding sites of BhbR1 and BhbR2. (A) EMSA of BhbR1 binding with the promoter DNA probe PbR1 (A) and BhbR2 binding with the promoter DNA probe PbR2 (B). “−” indicates samples incubated without 3-Br-4-HB, while “+” indicates samples incubated with 0.1 mM 3-Br-4-HB. Each lane contained 35 nM of promoter DNA probe. A 152-bp DNA fragment amplified from the 16S rRNA gene of strain H8 was used as a control DNA probe (28 nM). Protein-DNA complexes are indicated by brackets. DNase I footprinting assays of BhbR1 binding with the probe PbR1 in the absence (C) or presence (E) of 50 µM 3-Br-4-HB, and BhbR2 binding with the probe PbR2 in the absence (D) or presence (F) of 50 µM 3-Br-4-HB. The 6-carboxyfluorescein (FAM)-labeled DNA probe (350 ng) was incubated with 5 µg of BhbR1 or BhbR2, indicated by blue lines, as well as incubated without BhbR1 or BhbR2, indicated by red lines. Regions protected by BhbR1 or BhbR2 from DNase I cleavage are indicated with black dotted boxes, and the sequences of protected regions are shown at the bottom. (G) DNA elements in the promoters Pphbh1 and PpcaA2. The promoter regions protected by BhbR1 and BhbR2 in the absence (−) or presence (+) of 3-Br-4-HB are marked with light gray stripes. The regulatory binding sites for BhbR1 (ATA-N9-TAT) and BhbR2 (TTGATT-N5-AATCAA) are framed with red dashed lines. Two putative low-affinity activation binding sites (ABS1 and ABS2) in the promoter Pphbh1 are framed with blue dashed lines. The putative -10 and -35 boxes are underlined by black dashed and solid lines, respectively. TSSs are indicated by orange bent arrows, with the direction of the arrows indicating the direction of gene transcription.

DNase I footprint assays were carried out to identify the detailed BhnR1- and BhbR2-binding sites in their target promoter regions. In the absence of the inducer, BhbR1 protected a 54-bp DNA sequence (5′-TGCTGATAGCAAACCGTTATCGAGATAAGGAAAAACTGGCTTTTCGCTGTGCGC-3′), located from -22 to -75 relative to the TSS of the promoter P_phbh1_ (Fig. 4C). This protected region contains a high-affinity regulatory binding site (RBS), a 15-bp imperfect palindromic sequence (ATA-N_9_-TAT) with a consensus T-N_11_-A motif of a typical LTTR RBS, usually centered near position -65 in LTTR promoter regions (8, 27), and a putative low-affinity activation binding site (ABS1) ACT-N_10_-TGT overlapping the -35 box (Fig. 4G). The presence of 3-Br-4-HB shortened the BhbR1-protected region, located from -33 to -75 relative to the TSS of P_phbh1_ (Fig. 4E). This suggests that the activation binding site of BhbR1 was shifted to a position upstream of ABS1, designated as ABS2 (Fig. 4G).

BhbR2 consistently shielded a 22-bp invariant sequence (5′-TCCTTGATTGTTCTAATCAACC-3′) spanning from -22 to -43 in relation to the TSS of promoter P_pcaA2_, regardless of the presence of 3-Br-4-HB (Fig. 4D and F), which was consistent with the results of EMSA. The protected region covered the -35 box of P_pcaA2_ and included an AT-rich non-continuous palindromic sequence (TTGATTGTTCTAATCAA), which was consistent with the characteristics of typical MFTFs.

Determination of regulator binding dynamics with the inducer and the target promoter

Surface plasmon resonance (SPR) assays were performed to evaluate the binding dynamics between the regulators BhbR1 and BhbR2 and the inducer 3-Br-4-HB, as well as the promoters P_phbh1_ and P_pcaA2_, respectively. As shown in Fig. 5A and B, the equilibrium dissociation constants (K_D_) for the binding of BhbR1 and BhbR2 with 3-Br-4-HB were 2.2 and 2.1 µM, respectively, indicating comparable affinity. BhbR1 displayed a strong affinity (K_D_ = 0.7 nM) toward the biotinylated promoter fragment F_pbhb1_, whereas BhbR2 exhibited a low affinity (K_D_ = 397.0 nM) for the biotinylated promoter fragment F_pcaA2_ (Fig. 5C and D). However, upon the addition of 3-Br-4-HB, the affinity of BhbR1 for F_phbh1_ decreased, as indicated by an increased K_D_ value of 1.20 nM, whereas the affinity of BhbR2 for F_pcaA2_ was enhanced, resulting in a reduced K_D_ value of 259.0 nM (Fig. 5E and F). These results indicate that the addition of 3-Br-4-HB subtly modulates the binding and dissociation thresholds of BhbR1 and BhbR2 with their respective target promoters.

SPR assays for binding dynamics of the regulators with the inducer and target promoters. SPR assays analyzing interactions of the inducer 3-Br-4-HB with regulators BhbR1 (A) and BhbR2 (B). A serial dilution of 3-Br-4-HB (0.39–6.25 µM) was flowed over the CM7 sensor chip. SPR assays of BhbR1 binding with the fragments Fphbh1 in the absence (C) or presence (E) of 5 µM 3-Br-4-HB, and BhbR2 binding with the fragments FpcaA2 in the absence (D) or presence (F) of 5 µM 3-Br-4-HB. Biotinylated fragments Fphbh1 and FpcaA2 were captured on an SA sensor chip, while nonspecific biotinylated fragments F16S, derived from the 16S rRNA gene of strain H8, served as a control. Various concentrations of BhbR1 (68.75–1,100 nM) and BhbR2 (31.25–500 nM) were passed over the immobilized DNA fragments. The sensor surface was regenerated after each cycle using a short treatment with 0.5% (wt/vol) SDS. Derived dissociation constants (KD) are displayed above the equilibrium binding curves.

DISCUSSION

3-Br-4-HB is both naturally occurring as well as a synthetic compound widely used as an intermediate in chemical synthesis (1, 28, 29). Although the microbial degradation of 3-Br-4-HB has been extensively studied at the genetic and biochemical levels (1, 30), the regulatory mechanisms remain unclear. In this study, we demonstrated that the LysR-type transcriptional regulator BhbR1 and the MarR-family transcription factor BhbR2 trigger the transcription of two redundant 3-Br-4-HB-degrading gene clusters in response to 3-Br-4-HB, thereby governing 3-Br-4-HB metabolism in strain H8. This research not only fills a gap in the regulatory mechanisms of 3-Br-4-HB microbial metabolism but also deepens our understanding of the transcriptional regulation of redundant genes in microorganisms.

LTTRs are the most abundant and extensively studied transcriptional regulators in bacteria, managing various physiological and metabolic processes by activating or repressing target gene transcription (31–33). In the regulation of aromatic catabolism, LTTRs typically activate the transcription of catabolic genes while repressing their own gene transcription (8). In contrast, BhbR1 not only strongly activates target operon transcription but also slightly activates its own gene transcription (Fig. 2B). Structurally, LTTRs contain a highly conserved winged-helix-turn-helix (wHTH) DNA-binding domain at the N-terminus and an effector-binding domain at the C-terminus (34). Physiologically active LTTRs typically form homotetramers composed of two dimers, each binding to a high-affinity RBS and a low-affinity ABS on the target promoter, respectively (35, 36). DNase I footprinting assays revealed that the addition of 3-Br-4-HB shortened the BhbR1-binding region, allowing a BhbR1 dimer to slide from ABS1 to ABS2 (Fig. 4E and G), thereby exposing the -35 box and initiating transcriptional activation. This interaction between BhbR1 and the target promoter resembles the proposed “sliding dimer” model of LTTRs: the conformational contraction induced by effectors affects the entire protein-DNA complex, leading to the relaxation of the DNA bending and exposure of the -35 box, which facilitates the recruitment of RNA polymerase to activate transcription (27, 33, 37, 38).

MFTFs are widely distributed in bacteria and archaea and play important roles in various biological processes, including antibiotic resistance, stress responses, virulence, and the catabolism of aromatic compounds (39–43). Typically, MFTFs consist of 148–196 amino acids and feature a wHTH DNA-binding motif and a ligand-binding site, enabling conformational changes in response to specific signals (8, 44). Most MFTFs are homodimers, with each wHTH binding to an inverted repeat sequence within a palindromic sequence (16–20 bp) in the promoter region (45). The overlap of the binding sites of MFTFs with the -10 and -35 boxes leads to transcriptional repression of target genes; upon binding effectors, MFTFs undergo a conformational change, causing dissociation from the promoter DNA and relieving transcriptional repression (45, 46). To date, most MFTFs have been identified as transcriptional repressors, with only a few acting as activators (41, 47, 48). In this study, transcriptional activation of the phbh2pcaA2pcaB2 operon was observed exclusively in the presence of bhbR2 (Fig. 2C and D), suggesting that BhbR2 functions as a transcriptional activator in the catabolism of 3-Br-4-HB. Notably, the MFTFs with the closest phylogenetic relationship to BhbR2 in the phylogenetic tree have been identified as transcriptional repressors (Fig. S3B), highlighting the unique role of BhbR2. Furthermore, EMSA and DNase I footprinting assays showed that the addition of the inducer 3-Br-4-HB did not dissociate BhbR2 from the promoter DNA or alter its promoter-binding site (Fig. 4F). Collectively, these findings suggest a novel regulatory mechanism by which BhbR2 positively regulates the transcription of the phbh2pcaA2pcaB2 operon, although further investigation is needed to fully understand this process.

Regulating the transcription of redundant genes in response to specific signals is an effective strategy for balancing selective advantages and physiological burdens. A classic example was found in Pseudomonas putida mt-2, where the transcription of the redundant catechol 1,2-dioxygenase gene catA2 was selectively triggered in response to intracellular excess catechol, functioning as a “metabolic safety valve” to alleviate the toxicity of excessive catechol (49). In strain H8, the redundancy of the two 3-Br-4-HB-catabolic operons has been confirmed to help the host cope with toxic stress and metabolic disturbances caused by high concentrations of 3-Br-4-HB (21). When exposed to varying concentrations of 3-Br-4-HB, BhbR1 and BhbR2 consistently activated transcription of their respective operons, with no selective transcription favoring either operon in response to high or low concentrations of the inducer (Fig. 3C). This observation was further supported by the similar K_D_ values of BhbR1 and BhbR2 with 3-Br-4-HB (Fig. 5A and B). Nevertheless, high concentrations of 3-Br-4-HB (0.72 and 0.96 mM) adversely affected strain H8 cells, resulting in a significant attenuation of the transcriptional activation mediated by BhbR1 and BhbR2 (Fig. 3C). Therefore, equipping redundant 3-Br-4-HB-catabolic operons with two distinct regulatory systems is an effective safeguard measure to ensure the production of sufficient degradation enzymes, enabling host cells to cope with the toxic effects of high concentrations of 3-Br-4-HB in the habitat.

MATERIALS AND METHODS

Chemicals, bacterial strains, culture medium, plasmids, and oligonucleotides

All chemical standards (purities ≥ 98%), 3-Br-4-HB, Br-PCA, DBHB, and 4-HB, were purchased from Bidepharm (Shanghai, China) or J&K Chemical Co., Ltd. (Shanghai, China). Molecular biology reagents, such as restriction enzymes, high-fidelity DNA polymerase, PrimeScript RT reagent Kit with gDNA Eraser, TB Green Premix Ex Taq II (Tli RNaseH Plus) Kit, and BCA Protein Assay Kit, were purchased from TakaRa Biotechnology Co., Ltd. (Dalian, China). ClonExpress II One Step cloning kit was purchased from Vazyme Biotech Co., Ltd. (Nanjing, China).

The strains and plasmids used in this study are summarized in Table 1, and oligonucleotide primers are listed in Table 2. The method and results of phylogenetic analysis of Pigmentiphaga kullae strain H8 are shown in Text S1. Lysogenic broth (LB) medium (10.0 g tryptone, 5.0 g yeast extract, 5.0 g NaCl, and 1 L deionized water) was used to cultivate E. coli strains and Pigmentiphaga kullae strains in culture tubes or Erlenmeyer flasks with shaking (180 rpm) at 37°C. Mineral salts medium (MSM; 1.0 g NH_4_Cl, 1.5 g K_2_HPO_4_, 0.5 g KH_2_PO_4_, 0.2 g MgSO_4_, 1.0 g NaCl, and 1 L deionized water, pH 7.0) supplemented with 0.2 mM 3-Br-4-HB was used for degradation assays. YMS medium (MSM supplemented with 0.2% [wt/vol] yeast extract) was used to prepare substrate-induced cells. The solid medium was prepared by adding 2.0% (wt/vol) agar to the liquid medium. Antibiotic concentrations were used as follows: kanamycin at 50 mg L^−1^, spectinomycin at 80 mg L^−1^, and ampicillin and gentamicin at 100 mg L^−1^.

Detection of TSSs of the phbh1pcaApcaBorf404 and phbh2pcaB2pcaA2 operons

The TSSs of the phbh1pcaApcaBorf404 and phbh2pcaB2pcaA2 operons were identified using a 5′/3′-RACE Kit (2nd Generation, Roche, Switzerland) following the manufacturer’s instructions. Briefly, total RNA was extracted from strain H8 cells induced by 0.2 mM 3-Br-4-HB, using TRIzol reagent (Invitrogen). First-strand cDNA was synthesized with a primer (phbh1-GSP1 or pcaA2-GSP1) by reverse transcription. The cDNA was incubated with terminal transferase and dATP for homopolymeric tailing. Tailed cDNA was amplified using an oligo dT-anchor primer from the kit and a specific primer (phbh1-GSP2 or pcaA2-GSP2). The obtained PCR product was used as a template for nested PCR with the oligo dT-anchor primer and a specific primer (phbh1-GSP3 or pcaA2-GSP3), and the resulting product was cloned into pMD19-T (TaKaRa) for sequencing.

Gene disruption and complementation

The upstream and downstream homologous arms of bhbR1 and bhbR2 were amplified with primer pairs, bhbR1-Kn1/Kn2 with bhbR1-Kn3/Kn4, and bhbR2-Kn1/Kn2 with bhbR2-Kn3/Kn4, respectively. The upstream and downstream fragments of both bhbR1 and bhbR2 were fused by overlapping PCR and inserted into pJQ200SK to generate bhbR1-disrupted plasmid pJQ-ΔbhbR1 and bhbR2-disrupted plasmid pJQ-ΔbhbR2. Then, pJQ-ΔbhbR1 and pJQ-ΔbhbR2 were transferred into strains H8Δphbh2 and H8Δphbh1, respectively, by triparental mating with the helper strain E. coli HB101 (pRK2013). The bhbR1-disrupted mutant H8Δphbh2ΔbhbR1 and bhbR2-disrupted mutant H8Δphbh1ΔbhbR2 were obtained using a method that has been previously described (21).

The PCR-amplified sequences of bhbR1 and bhbR2 were individually cloned into the broad-host-range vector pBBR1MCS-2 using a ClonExpress II one-step cloning kit (Vazyme, Nanjing, China). The resulting constructs (pBBR-bhbR1 and pBBR-bhbR2) were transformed into mutants H8Δphbh2ΔbhbR1 and H8Δphbh1ΔbhbR2, respectively, using triparental mating. The bhbR1-complemented strain H8Δphbh2ΔbhbR1 (pBBR-bhbR1) and the bhbR2-complemented strain H8Δphbh2ΔbhbR2 (pBBR-bhbR2) were then used for further experiments.

RT-qPCR

Cells of various Pigmentiphaga kullae strains were grown in LB broth overnight, harvested by centrifugation, and washed twice with fresh YMS medium. Cells were then inoculated into YMS medium at an OD_600_ value of 0.4 and incubated with 0.2 mM 3-Br-4-HB, Br-PCA, or DBHB for 1.5 h or 0.2 mM 4-HB for 0.5 h to prepare substrate-induced cells. Cells were grown in YMS medium without additives for 1.5 h to generate uninduced control cells. The extraction of total RNA from induced and uninduced cells, generation of cDNA, and RT-qPCR were then carried out according to the previously described methods (32). Relative transcription of target genes was quantified using the 2^-ΔΔCT^ threshold cycle (C_T_) method (51), and the 16S rRNA gene of strain H8 was used as a reference for normalization. The ΔC_T_ represents the difference between transcription of the tested gene and 16S rRNA gene in the same treatment group, and ΔΔC_T_ represents the difference between ΔC_T_ (induced group) and the ΔC_T_ (uninduced group). All samples were run in triplicate.

Construction of promoter-lacZ transcriptional fusions

Two promoter-lacZ (P_phbh1_ and P_pcaA2_) plasmids were constructed to identify inducible effectors by detecting β-galactosidase activity. The P_phbh1_-containing DNA fragment PbR1 and the P_pcaA2_-containing DNA fragment PbR2 were amplified from genomic DNA of strain H8 using the primer pairs pME-PbR1-F/pME-PbR1-R and pME-PbR2-F/pME-PbR2-R, respectively, as listed in Table 2. The PCR products were digested with EcoRI and PstI and then ligated into the EcoRI/PstI-digested pME6522 to generate reconstructed plasmids pME-PbR1 and pME-PbR2. Subsequently, these plasmids were transformed into strains H8Δphbh2 and H8Δphbh1 to produce H8Δphbh2 (pME-PbR1) and H8Δphbh1 (pME-PbR2), respectively.

The cells of strains H8Δphbh2 (pME*-PbR1*) and H8Δphbh1 (pME*-PbR2*) were inoculated into YMS medium at an OD_600_ of 0.3. After adding 0.2 mM 3-Br-4-HB, Br-PCA, DBHB, or 4-HB, the cells were incubated at 37°C and 180 rpm for 3 h. The cells were then harvested by centrifugation and used for β-galactosidase activity analysis. β-galactosidase activity was measured using SDS- and chloroform-permeabilized cells as described (35, 52). All experiments were conducted in triplicate.

Protein expression and purification

The bhbR1 and bhbR2 genes were amplified with primer pairs bhbR1-29f/r and bhbR2-29f/r, respectively, and then cloned into the pET29a(+) vector to generate expression plasmids pET-bhbR1 and pET-bhbR2. The expression and purification of proteins with a C-terminal His tag were performed following a previously described method (32). The purified BhbR1 and BhbR2 were analyzed by SDS-PAGE and dialyzed in 3 L of PBS overnight to remove imidazole. Concentrations of purified proteins were determined using a BCA Protein Assay Kit (TaKaRa, Dalian, China) according to the manufacturer’s instructions.

Gel filtration chromatography

Before gel filtration, purified BhbR1 and BhbR2 were concentrated using an ultrafiltration device (Amicon) with a 10-kDa cutoff membrane. To calibrate the molecular weights of BhbR1 and BhbR2, a panel of commercially available proteins, including myosin (200.0 kDa), phosphorylase B (97.2 kDa), serum albumin (66.4 kDa), egg albumin (44.3 kDa), papain (23.4 kDa), and lysozyme (14.7 kDa), was used as molecular weight markers. A solution containing BhbR1, BhbR2, or protein markers was charged onto a Superose 6 10/300 GL column (GE Healthcare) for gel filtration at 25°C using an ÄKTA Purifier chromatography system (GE Healthcare). The elution process was carried out with PBS at a flow rate of 0.4 mL min^−1^ for 1 h, with detection at a wavelength of 280 nm; elution volumes were systematically recorded.

Electrophoretic mobility shift assay

The EMSA was conducted following previously described methods (32, 53). The DNA probes PbR1 (a 243-bp DNA fragment containing P_phbh1_ sequence) and PbR2 (a 258-bp DNA fragment containing P_pcaA2_ sequence) were amplified with primer pairs PbR1-f/r and PbR2-f/r, respectively. A non-specific 152-bp DNA probe amplified from the 16S rRNA gene of strain H8 using primer pairs Pb16S-f/r served as control DNA. The DNA probes (35 nM) were then incubated with increasing amounts of purified BhbR1 (0–0.6 µM) and BhbR2 (0–1.92 µM) in EMSA buffers. The buffer for BhbR1 contained 100 mM Tris–HCl (pH 8.0), 50 mM KCl, 5% (vol/vol) glycerol, and 1 mM dithiothreitol (DTT), while the buffer for BhbR2 contained 50 mM Tris-HCl (pH 8.0), 30 mM KCl, 5% (vol/vol) glycerol, 1 mM EDTA, and 5 mM DTT. After incubation at 25°C for 30 min, the resultant mixtures were immediately loaded onto a 5% native polyacrylamide gel at 4°C and electrophoresed at 100 V for 1.5 h using a 0.5× TGE buffer (45 mM Tris-boric acid and 1 mM EDTA, pH 8.0). The gels were subsequently stained with ethidium bromide for 10 min at room temperature to visualize DNA-protein interactions.

DNase I footprinting assay

DNA probes PbR1 and PbR2 were cloned into pMD19-T (TaKaRa) to construct recombinant plasmids pMD-PbR1 and pMD-PbR2, respectively. Subsequently, fluorescent 6-carboxyfluorescein (FAM)-labeled probes were generated by PCR amplification with primers FAM-M13F and M13R, using pMD-PbR1 and pMD-PbR2 as templates. For each DNase I footprinting assay, the binding reaction and sample analysis were conducted following a protocol described previously (32), modified by using the GeneScan-LIZ600 size standard from Applied Biosystems for sample analysis.

SPR assay

SPR assays were performed to analyze intermolecular interactions using a Biomoleculer Interaction Analysis System (Biacore T200, Cytiva, Sweden) at room temperature. To determine the interaction between the regulator and the inducer, purified BhbR1 and BhbR2 were immobilized onto a CM7 sensor chip (Cytiva, Sweden) by amine coupling, achieving approximately 10,000 response units for each protein using the EDC/NHS method. A serial dilution of the inducer 3-Br-4-HB was prepared in PBST buffer (10 mM Na_2_HPO_4_, 2 mM KH_2_PO_4_, pH 7.4, 137 mM NaCl, 2.7 mM KCl, and 0.05% Tween 20) and was flowed across the chip at a rate of 30 µL min^−1^. The equilibrium dissociation constant (K_D_) was determined using the Biacore T200 Evaluation software with the steady-state affinity model.

To determine kinetic constants of regulator-promoter interactions, biotinylated DNA fragments F_pbhb1_ (144 bp) and F_pcaA2_ (153 bp) were generated by PCR amplification using primer pairs Fpb1-Btn-f/Fpb1-r and Fpb2-f/Fpb2-Btn-r, respectively. These biotinylated fragments were captured on a streptavidin sensor chip (Cytiva, Sweden) before introducing target proteins and monitoring interactions. As a control, a 152-bp nonspecific biotinylated DNA fragment derived from the 16S rRNA gene of strain H8 was amplified using primers F16S-Btn-f and F16S-r. Concentration gradients of BhbR1 and BhbR2 were prepared in HEPES-EP buffer (10 mM HEPES, 150 mM NaCl, 0.05% surfactant P20, and 3 mM EDTA, pH 7.4), with or without 5 µM 3-Br-4-HB; these were then applied to the chip. Kinetic analyses were conducted using the Biacore T200 Evaluation software with the kinetics affinity model.

Chemical analysis

Qualitative and quantitative determination of 3-Br-4HB were carried out using a Dionex UltiMate 3000SD HPLC system equipped with a diode array detector and a Dionex C_18_ reversed-phase column (4.6 × 250 mm, 5-mm particle size), following the previously described method (21).