The predicted σ54-dependent regulator EtpR is essential for expression of genes for anaerobic p-ethylphenol and p-hydroxyacetophenone degradation in “Aromatoleum aromaticum” EbN1
© Büsing et al. 2015
Received: 25 June 2015
Accepted: 15 October 2015
Published: 2 November 2015
The denitrifying betaproteobacterium "Aromatoleum aromaticum" EbN1 anaerobically utilizes a multitude of aromatic compounds via specific peripheral degradation routes. Compound-specific formation of these catabolic modules is assumed to be mediated by specific transcriptional activators. In case of the recently elucidated p-ethylphenol/p-hydroxyacetophenone pathway, the highly substrate-specific regulation was implicated to involve the predicted σ54-dependent, NtrC-type regulator EbA324. The latter was suggested to control the expression of the two neighboring gene clusters encoding the catabolic enzymes as well as a corresponding putative solvent efflux system. In the present study, a molecular genetic approach was used to study the predicted function of EbA324.
An unmarked in frame ΔebA324 (here renamed as ΔetpR; p-ethylphenol regulator) deletion mutation was generated. The ΔetpR mutant was unable to grow anaerobically with either p-ethylphenol or p-hydroxyacetophenone. Growth similar to the wild type was restored in the ΔetpR mutant background by in trans expression of plasmid-born etpR. Furthermore, expression of the "p-ethylphenol" gene clusters as well as corresponding protein formation was shown to depend on the presence of both, EtpR and either p-ethylphenol or p-hydroxyacetophenone. In the wild type, the etpR gene appears to be constitutively expressed and its expression level not to be modulated upon effector presence. Comparison with the regulatory domains of known phenol- and alkylbenzene-responsive NtrC-type regulators of Pseudomonas spp. and Thauera aromatica allowed identifying >60 amino acid residues in the regulatory domain (in particular positions 149 to 192 of EtpR) that may contribute to the effector specificity viz. presumptively restricted effector spectrum of EtpR.
This study provides experimental evidence for the genome predicted σ54-dependent regulator EtpR (formerly EbA324) of "A. aromaticum" EbN1 to be responsive to p-ethylphenol, as well as its degradation intermediate p-hydroxyacetophenone, and to control the expression of genes involved in the anaerobic degradation of these two aromatic growth substrates. Overall, the presented results advance our understanding on the regulation of anaerobic aromatic compound catabolism, foremost based on the sensory discrimination of structurally similar substrates.
The "Aromatoleum"/Thauera/Azoarcus cluster within Betaproteobacteria comprises most of the currently known denitrifiers capable of anaerobic degradation of aromatic compounds [1, 2]. "Aromatoleum aromaticum" EbN1 is a metabolically versatile and the first genome-sequenced representative of this cluster, completely oxidizing >20 different aromatic compounds under anoxic conditions. These include the alkylbenzenes toluene and ethylbenzene, phenol, as well as the alkylphenols p-cresol and p-ethylphenol [3–5]. The multiple sensory/regulatory proteins predicted from the genome were suggested to constitute a fine-tuned regulatory network . Subsequent experimental studies indeed implicated the latter in substrate-specific formation of catabolic modules [4, 5, 7, 8], as well as in the adaptation to substrate-limiting  and stress conditions . Synthesis of the hitherto accomplished physiological-proteomic insights qualifies "A. aromaticum" EbN1 as a promising systems biology model .
In the present study, an unmarked ∆etpR in frame deletion mutation was generated to verify the predicted regulatory function of EtpR. The ∆etpR mutant and the in trans etpR-complemented mutant were characterized by means of physiological experiments as well as on the molecular level.
Results and discussion
Generation of the ∆etpR and etpR-complemented mutants
The unmarked in frame ∆etpR deletion mutation was generated to test the predicted regulatory function of EtpR in mediating p-ethylphenol- and p-hydroxyacetophenone-specific expression of the two “p-ethylphenol” gene clusters. In the ΔetpR mutant, only the start and stop codons of etpR were preserved to maintain the reading frame (Fig. 1a). Accordingly, no PCR product could be obtained using etpR-specific primers and only a small 318 bp amplicon was observed applying primers targeting the up- and downstream intergenic regions of etpR (wild type amplicon 2,133 bp; Fig. 1b). In addition, correctness of the deletion site and its 5'- and 3'-flanking regions (i.e., ∆etpR genotype) was confirmed by nucleotide sequencing (Additional file 1: Figure S1). This newly generated ∆etpR mutant was in trans complemented via an etpR-bearing broad-host range plasmid, yielding constitutive expression of etpR (see below). The resultant etpR-complemented mutant had the genotype ΔetpR, pBBR1MCS-2 ΩetpR.
The ∆etpR mutant cannot grow with p-ethylphenol and p-hydroxyacetophenone
Selected growth parameters determined for wild type, ΔetpR mutant and etpR-complemented mutant (etpR-compl.) of "A. aromaticum" EbN1a
Consumption rate (mM h−1)
Bz1 + pHac2c
Bz1 + pEp2c
Growth with benzoate
Growth with p-ethylphenol and p-hydroxyacetophenone
The ΔetpR mutant could not grow with p-ethylphenol and p-hydroxyacetophenone (Fig. 2a, c). The minute initial increase in optical density can probably be attributed to consumption of residual benzoate, co-transferred during initial inoculation. The unexpected complete depletion of p-ethylphenol from ΔetpR mutant cultures after ~160 h of incubation is most likely due to unspecific membrane accumulation, as it is not coupled to denitrification and also observable in wild type cultures with de novo protein synthesis inhibited by kanamycin (see Additional file 1: Figure S2). Noteworthy, the etpR-complemented mutant started to grow after a markedly shorter lag-phase as compared to the wild type, even though maximum growth rates and substrate depletion profiles of the two strains were similar (Fig. 2a, c; Table 1). This shorter lag-phase may be due to the higher EtpR abundance in the complemented strain (see below).
Growth with a binary mixture of benzoate and p-hydroxyacetophenone
When the three strains were shifted from benzoate to a binary mixture of benzoate and p-hydroxyacetophenone, highest rates of growth and benzoate consumption were observed for the ΔetpR mutant (Table 1), while the concentration of p-hydroxyacetophenone remained unchanged in this strain (Fig. 2d). However, growth rate and ODmax were lower as compared to growth with benzoate, most likely due to the toxicity of p-hydroxyacetophenone. Rates for growth and benzoate consumption of the wild type with the binary substrate mixture were higher as compared to the etpR-complemented mutant, but the rate of concomitant p-hydroxyacetophenone depletion was lower (Table 1). Hence, the etpR-complemented mutant seems to more preferentially consume p-hydroxyacetophenone than benzoate as compared to the wild type (Fig. 2d). Similar results were obtained for growth experiments with a binary mixture of benzoate and p-ethylphenol (Additional file 1: Figure S3).
EtpR mediates substrate-specific expression of the "p-ethylphenol" gene clusters
The simultaneous utilization of benzoate and p-hydroxyacetophenone (see preceeding paragraph), i.e., absence of catabolite repression, is a prerequisite for subsequent analysis of transcript and proteomic profiles. In the ∆etpR mutant benzoate sustains growth while at the same time the hypothesized loss of response to the effector p-hydroxyacetophenone due to etpR deletion can be tested. p-Hydroxyacetophenone was used as substitute of p-ethylphenol due to (i) its higher water solubility (not requiring provision via a carrier phase), (ii) the apparent absence of a passive uptake as observed for p-ethylphenol (Fig. 2a, c) and (iii) the uniform induction of gene expression and protein formation by both substrates .
Non-detection of the EtpR protein in the wild type suggests protein abundance below the detection limit of the applied method. In contrast, EtpR was detected in all samples of the etpR-complemented mutant with high confidence indicating a markedly higher abundance of EtpR. The absence of "p-ethylphenol" gene cluster expression in benzoate-utilizing cells of the etpR-complemented mutant, despite the artificially high EtpR abundance in this strain, demonstrates the effector dependence of EtpR for transcriptional activation.
Hints on constitutive expression of etpR
Wild type EtpR abundance probably sufficient for full-level transcription
The etpR-complemented mutant strongly expressed etpR under all three tested substrate conditions (>28-fold vs. wild type; Fig. 4), agreeing with the medium copy number of the vector (around ten), the strong vector-inherent promoter, and the exclusive detection of the EtpR protein. Notably, the artificially high abundance of EtpR in the etpR-complemented mutant did not yield an increased abundance of proteins involved in p-ethylphenol degradation (at ½ODmax) as compared to the wild type (Fig. 3b). Hence, maximum transcription/translation levels are apparently achieved also with the lower EtpR abundance occurring in the wild type, which is also reflected by the similar μmax of both strains with p-ethylphenol or p-hydroxyacetophenone. The higher EtpR level in the etpR-complemented mutant may allow for reaching the maximum levels of catabolic proteins faster, which may explain the significantly shorter lag-phase as compared to the wild type (Fig. 2a, c; Table 1).
Sequence comparison of regulatory domains
Known phenol- or alkylbenzene-specific NtrC-type regulators
DmpR and PhlR are closely related regulators of aerobic phenol catabolism in Pseudomonas sp. CF600 [17, 22] and P. putida H , respectively. They are activated by direct binding of phenol or derivatives thereof, e.g., cresol and dimethylphenol isomers [24, 25] (Additional file 1: Figure S4). Notably, DmpR also recognizes o- and m-ethylphenol . Similarly, DmpR of Thauera aromatica K172  and PdeR of "A. aromaticum" EbN1 [4, 6] are suggested to regulate anaerobic phenol degradation; differential proteomics indicated PdeR to also respond to p-cresol . XylR of P. putida regulates aerobic toluene degradation, and is activated by this as well as other alkylbenzenes such as m- and p-xylene .
Alteration of regulatory domain residues in XylR and DmpR from Pseudomonas spp. yielded broadened or restricted effector spectra, or even completely inhibited the response to aromatic effectors [19, 20, 27–34]. Most of these residues are confined to a stretch of 75 amino acids, defined as effector specifying region (ESR) . 3D-models of XylR and DmpR predicted structural features for shaping an effector-binding pocket and interaction with the central domain [36, 37]. The current knowledge on the regulatory domain is compiled in Additional file 1: Figure S5.
Comparison of EtpR to known regulators
The phylogeny of the inspected regulatory domains reflects effector spectra as well as deployment for anaerobic vs. aerobic catabolism. EtpR branches off from the "anaerobic" phenol-specific regulators (PdeR of "A. aromaticum" EbN1 and DmpR of T. aromatica K172), while the three together separate from those of aerobic phenol degradation (DmpR and PhlR of Pseudomonas spp.). The alkylbenzene-specific XylR of P. putida is more distantly related to the other five regulators (Additional file 1: Figure S4).
Current proteomic data demonstrated specific formation of the "p-ethylphenol"-proteins only in cells anaerobically growing with p-ethylphenol or p-hydroxyacetophenone, but not with 20 other aromatic substrates [5, 11]. Therefore, activation of EtpR may be restricted to the former two. Such a narrow effector spectrum would distinct EtpR from the above described three regulators of Pseudomonas spp..
To identify primary sequence features, possibly linked to the effector spectrum of EtpR, aligned regulatory domain sequences were analyzed (Additional file 1: Figure S5), with the ESR emphasized in Fig. 5 (amino acid positions 130–205 in EtpR). Across these six complete regulatory domains, 43 residues (20.7 %) were strictly conserved, comprising most of the reported conserved residues of NtrC-type regulators . Sixty-two other residues (29.8 %; colored circles) are conserved to differing degrees and are mostly ESR-located; they may therefore be effector specifying for EtpR and are briefly described in the following.
Recognition of the phenolic moiety may involve 8 residues (4 in the ESR; brown circles) that are exclusively conserved in the five phenolic compound-sensing regulators (incl. EtpR). Further 6 residues (light green circles) could also contribute to phenol-specificity, as they are conserved in DmpR, PhlR and PdeR, but not in alkylbenzene-sensing XylR and alkylphenol-sensing EtpR.
Notably, a total of 28 residues (dark green circles) are conserved in all proteins except for EtpR, with 16 of them located in the ESR. The sensory relevance of these residues is reflected by four of them being located in the predicted binding pocket of DmpR and XylR  and two at locations that broaden the effector spectrum of DmpR [30, 31]. From an inversed perspective, these 28 residues are distinct in EtpR. They may therefore be involved in specific sensing of, e.g., the keto group in p-hydroxyacetophenone, since the latter is not known to be an effector of the other five compared regulators. Alternatively, these 28 residues may at least indirectly contribute to a structural shaping of the EtpR-specific sensory properties. These scenarios may also account for another group of 20 residues (blue circles), which are differently conserved in DmpR, XylR and PhlR of Pseudomonas spp. as compared to DmpR of T. aromatica and PdeR of "A. aromaticum" EbN1, but are again distinct in EtpR.
More than 17 residue changes (marked by a "+" sign) in the regulatory domains of Pseudomonas DmpR and XylR (13 in the ESR or close by) broaden their effector spectra (partly also towards p-ethylphenol) [19, 31, 34]. They may not be effector specifying for EtpR, as none of the changes yields EtpR-residues.
Notably, the effector specificity of EtpR may not only be attributed to single residues, since also the interaction of different regions contributes to defining effector spectra, i.e., loops (yellow areas) and interactions of regulatory and central domain (black arrows) .
Aerobic aromatic compound degradation pathways of Pseudomonas spp. may accommodate several, structurally related aromatic substrates, e.g., toluene, m- and p-xylene as well as 1,2,4-trimethylbenzene in case of P. putida . This substrate promiscuity of single pathways is also reflected in rather broad effector spectra of the involved transcriptional regulators [16, 25]. In contrast, the substrate ranges of the individual anaerobic pathways in "A. aromaticum" EbN1 appear to be more restricted and the substrate-specific expression of their genes is assumed to be individually controlled by corresponding one- or two-component regulatory systems . The observed highly p-ethylphenol/p-hydroxyacetophenone-specific induction of the anaerobic p-ethylphenol degradation pathway by EtpR can be attributed to a concurrence of unique sets of amino acids accumulating in the ESR (accounting for 36.8 %), in particular between position 149 and 192. Since they are most specific for EtpR as compared to the other regulators, they possibly specify the sensing of phenolic-, ketonic- and/or alkylbenzylic-moieties, and determine the observed narrow effector spectrum of EtpR. This further supports a general role of ESRs in mediating sensory recognition of aromatic compounds as previously reported for DmpR and XylR of Pseudomonas spp. . The apparent constitutive, low level expression of etpR in the wild type allows for a full level formation of catabolic proteins that may be only reached faster at higher regulator abundance as observed for the etpR-complemented mutant. Hence, the extent of "p-ethylphenol" gene cluster expression should depend on the presence of the substrates/effectors p-ethylphenol and p-hydroxyacetophenone rather than on the abundance of the regulator itself (EtpR). Overall, the present study demonstrates EtpR (EbA324) to serve as transcriptional regulator in the p-ethylphenol catabolism, and represents to our knowledge the first molecular genetic study on a σ54-dependent regulator in an anaerobic aromatic compound degrader.
Bacterial strains and cultivation
Strains and plasmids used in this study
Genotype and/or characteristics
“Aromatoleum aromaticum” EbN1
EbN1-RR001 (ΔetpR mutant)
EbN1-RR002 (etpR-complemented mutant)
ΔetpR, pBBR1MCS-2 ΩetpR
EbN1-RR003 (Wild type containing pBBR1MCS-2)
Wild type, pBBR1MCS-2
EbN1-RR004 (ΔetpR mutant containing pBBR1MCS-2)
Escherichia coli S17-1
Pro, thi, hsdR, Tra +, recA−, Trr, Smr, ΩRP4-TE::Mu-Kn::Tn7
KmR, sacB modified from B. subtilis, lacZα
KmR, sacB modified from B. subtilis, lacZα, acsA from "A. aromaticum" EbN1
KmR, sacB modified from B. subtilis, lacZα; acsA, ebA326 and part of ebA327 from "A. aromaticum" EbN1
KmR, mob, lacZα
KmR, mob, lacZα, etpR from "A. aromaticum" EbN1
Generation of the in frame ∆etpR deletion mutation
Oligonucleotide primers applied in this study
Nucleotide sequence (5' → 3')b
Product length (bp)
Gene specific primer pairs
Generation of Δ etpR deletion mutation
Identification of Δ etpR genotype
Generation of in trans complementation of etpR
For unmarked knockout of the etpR (ebA324) gene, a knockout vector based on the suicide vector pK19mobsacB  was constructed in an E. coli S17-1 background, containing 2.4 kbp of the 5'-region and 1.3 kbp of the 3'-region of etpR. Initially, the 5'-region containing parts of the acsA gene were cloned into pK19mobsacB using SphI and BamHI restriction sites as reported , yielding the plasmid pK19 ΩacsA. Subsequently, the 3'-region, containing ebA326 and parts of the ebA327 gene, was cloned into plasmid pK19 ΩacsA using the BamHI restriction site, resulting in the pK19 ΩacsAebA326/7 knockout vector (Table 2). In the final vector construct, the start and stop codons of etpR were maintained, separated by a 6 bp BamHI restriction site ("GTGGGATCCACT" blow up in Fig. 1a). Homologous regions were amplified by PCR from genomic DNA of "A. aromaticum" EbN1 using a high fidelity polymerase (Phusion®; ThermoFisher Scientific, Dreieich, Germany; Table 3). The correctness of nucleotide sequences was verified by sequence analysis as described before  and pK19 ΩacsAebA326/7 was transferred by conjugation from the E. coli S17-1 donor strain to "A. aromaticum" EbN1 according to the protocol described previously . Integration of the knockout vector (single-cross over) gave rise to kanamycin resistant colonies and was verified by PCR using a primer pair (∆etpR_F/R) targeting the up- and downstream intergenic regions of etpR, yielding two amplicons of 318 bp and 2.13 kbp, respectively (Table 3; Fig. 1). The second cross-over (i.e., removal of the plasmid) was induced by several transfers of the single cross-over mutant in liquid medium without kanamycin. Colonies capable of growing on sucrose-containing medium were screened using the same primer pair as described above to identify a ∆etpR genotype in the deletion mutant strain EbN1-RR001 (Fig. 1). The genotype was verified by sequencing.
Complementation of etpR in trans into the ∆etpR deletion mutant
A complementation vector for in trans expression of etpR was generated in an E. coli S17-1 background, based on the broad-host range vector pBBR1MCS-2 . A 2.4 kbp nucleotide sequence was amplified by PCR, containing etpR as well as 300 bp upstream of the etpR start codon to also include the ribosomal binding site (Table 3; Fig. 1). This amplicon was cloned into the pBBR1MCS-2 vector using KpnI and XhoI restriction sites and verified by sequencing. The vector was transferred via conjugation to the ∆etpR mutant yielding the etpR-complemented mutant strain EbN1-RR002 (genotype: ΔetpR, pBBR1MCS-2 ΩetpR) (Table 2; Fig. 1). Conjugation via agar-plate mating, identification for positive clones on selective media and PCR-based verification were carried out as previously described . For control experiments, the pBBR1MCS-2 vector without etpR was conjugationally transferred to the wild type strain and the ΔetpR mutant yielding strain EbN1-RR003 (genotype: wild type, pBBR1MCS-2) and strain EbN1-RR004 (genotype: ΔetpR, pBBR1MCS-2), respectively (Table 2).
Growth experiments with the wild type, the ∆etpR mutant and the etpR-complemented mutant were carried out to characterize the phenotype of the generated ∆etpR deletion mutation. All three strains were adapted to anaerobic growth with benzoate for at least five passages. Anaerobic cultivation was conducted under nitrate-limited conditions with 400 ml medium in 500 ml flat bottles, sealed with rubber stoppers. Pre-cultures were provided with 4 mM benzoate as growth substrate and cells transferred at half-maximal optical density (½ ODmax) to fresh medium supplemented with either of the following substrates: (i) p-ethylphenol (0.5 % (w/v) in 14 ml of the inert carrier phase 2,2,4,4,6,8,8-heptamethylnonane (HMN)), (ii) p-hydroxyacetophenone (2 mM), (iii) benzoate (4 mM) and (iv) a binary mixture of benzoate (4 mM) and either p-hydroxyacetophenone (2 mM) or p-ethylphenol (0.5 % (w/v) in 10 ml HMN). In case of cultures with benzoate or a binary substrate mixture, the medium contained 10 mM nitrate to achieve higher cell densities; while with p-ethylphenol and p-hydroxyacetophenone provided as single substrate only 7 mM nitrate were added to the medium.
Growth until depletion of the electron acceptor nitrate was monitored by measuring the optical density at 660 nm (UV–vis Spectrometer 1240; Shimadzu, Duisburg, Germany) and analysing the substrate concentrations in the culture supernatants with an UltiMate 3000 RSLC system (ThermoFisher Scientific, Germering, Germany). The supernatants were diluted and acidified (pH 2.0, 6 % acetonitrile) prior to analysis. Separation of analytes was achieved with a Dionex Acclaim 120 reversed-phase separation column (250 mm length, 2.1 mm inner diameter, 5 μm bead size; ThermoFisher Scientific). The column was temperature controlled at 25 °C and operated with a non-linear gradient of acetonitrile (5–90 % (v/v), pH 2.8) as eluent at a flow rate of 0.5 ml min−1: 2 min at 5 %, 5 to 14 % in 1 min, 14 to 39 % in 10.5 min, 39 to 90 % in 3 min, and 3 min at 90 %. Retention times (compound-specific wavelengths in parentheses) were the following: p-ethylphenol, 16.6 min (220 nm); p-hydroxyacetophenone, 9.4 min (270 nm); and benzoate, 11.8 min (236 nm).
For selected samples nitrate concentrations were determined by means of an ICS 1100 ion chromatography system (ThermoFisher Scientific). Nitrite and nitrate were separated using an IonPac™ AG9-HC separation column (250 mm length, 4 mm inner diameter, 9 μm bead size; ThermoFisher Scientific) and detected at a wavelength of 210 nm. Separation was achieved with sodium carbonate (9 mM) as the eluent administered at a flow rate of 1 ml min−1. The retention times (detection limits in parentheses) were as follows: nitrite, 9.1 min (1 μM); nitrate, 13.3 min (1 μM).
To assess the possibility of a passive depletion of p-ethylphenol from the medium, benzoate-adapted cells of the wild type were pre-grown with benzoate and transferred to anoxic medium with p-ethylphenol as described before. After 15 h, growth was inhibited by the addition of 50 μg ml−1 kanamycin. The concentration of p-ethylphenol in the cultures was analyzed by the RSLC system as given above.
Cultivation for transcript and proteomic analysis
For profiling of compound-specific transcripts and proteome signatures, substrate-adapted wild type, the ∆etpR mutant and the etpR-complemented mutant were grown anaerobically with either (i) p-hydroxyacetophenone, (ii) benzoate or (iii) a binary mixture of benzoate and p-hydroxyacetophenone as sole organic substrate(s). Cells were cultivated in 250 ml flat bottles with 200 ml medium and harvested during linear growth at ½ ODmax as described by Champion et al. . For each strain and growth condition, three replicate cultures were harvested for transcript and proteomic analyses, respectively (in total 6 replicate cultures each). Cell pellets for transcript analysis were treated with RNAprotect® Bacteria Reagent (Qiagen GmbH, Hilden, Germany) according to the manufacturer’s instructions.
Preparation of mRNA and reverse transcription (RT)-PCR
Preparation of total RNA was performed according to the protocol of Oelmüller et al. (22) using cells treated with RNAprotect® Bacteria Reagent. Complete removal of DNA from the RNA preparation was verified by PCR. The quality (i.e., integrity) of the isolated RNA was analysed with MOPS-gels according to standard protocols . cDNA was generated from two independent RNA preparations per strain and substrate condition, respectively, using the antisense primer of the target genes (Table 3). Reverse transcription was performed with 2.5 μg RNA applying the RevertAid H Minus Reverse Transcriptase (ThermoFisher Scientific) according to the manufacturer’s instructions. cDNA was amplified by PCR using the PCR MasterMix (Promega, Mannheim, Germany). Depending on the PCR efficiency of the gene-specific primer pairs, 1.0 or 2.0 μl of cDNA preparation were used as template per 20 μl PCR experiment comprising 20 or 39 PCR cycles.
To exclude polar effects resulting from the in frame ΔetpR deletion mutation and to qualitatively study gene expression of the two "p-ethylphenol" gene clusters located up and downstream of etpR, transcripts representative of both of them (Fig. 3) were analysed. Target genes were chosen such that both the first and last genes as well as the middle of each of the two gene clusters were covered, i.e., acsA, hped and pchF for the "catabolic" gene cluster, and ebA335, ebA327 and ebA326 for the "efflux" gene cluster.
Relative expression levels of etpR in the wild type and the eptR-complemented mutant were determined by real-time RT-PCR. BcrC (encoding the catalytic γ-subunit of benzoyl-CoA reductase) was selected as reference gene since benzoyl-CoA is a common intermediate in anaerobic degradation of benzoate as well as p-hydroxyacetophenone, and since bcrC expression is not regulated under the two investigated substrate conditions . cDNA was generated from three individual RNA preparations per strain and growth condition as described above. Real-time PCR was carried out as reported by Kühner et al.  using an iQ5 real-time PCR detection system (Bio-Rad, Munich, Germany) and a qPCR MasterMix Plus for SyBR green I with fluorescin (Eurogentec, Cologne, Germany). The correctness of obtained PCR products was verified by sequencing. Determination of PCR efficiencies was performed as described by Ramakers et al.  and relative expression levels were calculated according to Pfaffl et al. . At least three replicates of each individual cDNA preparation were analysed (in total 18 qPCR experiments). The wild type grown with p-hydroxyacetophenone served as reference state for calculation of relative gene expression levels (Fig. 4).
Whole cell shotgun analysis of substrate-adapted cells was performed as described recently . Essentially, tryptic peptides were separated by a nanoLC system (UltiMate3000 nanoRSLC; ThermoFisher Scientific) operated in a trap-column mode and equipped with a 25 cm separation column (C18, 2 μm bead size, 75 μm inner diameter; ThermoFisher Scientific), applying a 280 min linear acetonitrile gradient. The nanoLC eluent was continuously analyzed by an online-coupled ion-trap mass spectrometer (amaZon speed ETD; Bruker Daltonik GmbH, Bremen, Germany) using a captive spray ion source (Bruker Daltonik GmbH). Per full scan MS (mass range 400–1400 m/z), 20 MS/MS spectra of the most intense doubly (or more highly) charged ions were acquired applying subsequent precursor exclusion for 0.2 min. Protein identification was performed using the ProteinScape platform (version 3.1; Bruker Daltonik GmbH) on an in-house Mascot server (version 2.3; Matrix Science Ltd, London, UK) based on the genome sequence of "A. aromaticum" EbN1  and applying a target-decoy strategy as described . Search results of the three biological replicates per test state were compiled and only proteins identified by at least 2 peptides were considered.
We are grateful to D. Thies (Bremen) and C. Hinrichs (Oldenburg) for technical assistance. This study was supported by the FOL program of the Carl von Ossietzky University Oldenburg and the Deutsche Forschungsgemeinschaft (GRK 1885 "Molecular basis of sensory biology").
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- Widdel F, Rabus R. Anaerobic biodegradation of saturated and aromatic hydrocarbons. Curr Opin Biotechnol. 2001;12:259–76.View ArticlePubMedGoogle Scholar
- Widdel F, Musat F. Diversity and common principles in enzymatic activation of hydrocarbons. In: Timmis KN, editor. Handbook of Hydrocarbon and Lipid Microbiology. Berlin: Springer; 2010. p. 984–1009.Google Scholar
- Rabus R, Widdel F. Anaerobic degradation of ethylbenzene and other aromatic hydrocarbons by new denitrifying bacteria. Arch Microbiol. 1995;163:96–103.View ArticlePubMedGoogle Scholar
- Wöhlbrand L, Kallerhoff B, Lange D, Hufnagel P, Thiermann J, Reinhardt R, et al. Functional proteomic view of metabolic regulation in “Aromatoleum aromaticum” strain EbN1. Proteomics. 2007;7:2222–39.View ArticlePubMedGoogle Scholar
- Wöhlbrand L, Wilkes H, Halder T, Rabus R. Anaerobic degradation of p-ethylphenol by “Aromatoleum aromaticum” strain EbN1: pathway, regulation, and involved proteins. J Bacteriol. 2008;190:5699–709.PubMed CentralView ArticlePubMedGoogle Scholar
- Rabus R, Kube M, Heider J, Beck A, Heitmann K, Widdel F, et al. The genome sequence of an anaerobic aromatic-degrading denitrifying bacterium, strain EbN1. Arch Microbiol. 2005;183:27–36.View ArticlePubMedGoogle Scholar
- Kühner S, Wöhlbrand L, Fritz I, Wruck W, Hultschig C, Hufnagel P, et al. Substrate-dependent regulation of anaerobic degradation pathways for toluene and ethylbenzene in a denitrifying bacterium, strain EbN1. J Bacteriol. 2005;187:1493–503.PubMed CentralView ArticlePubMedGoogle Scholar
- Trautwein K, Wilkes H, Rabus R. Proteogenomic evidence for β-oxidation of plant-derived 3-phenylpropanoids in “Aromatoleum aromaticum” EbN1. Proteomics. 2012;12:1402–13.View ArticlePubMedGoogle Scholar
- Trautwein K, Lahme S, Wöhlbrand L, Feenders C, Mangelsdorf K, Harder J, et al. Physiological and proteomic adaptation of “Aromatoleum aromaticum” EbN1 to low growth rates in benzoate-limited, anoxic chemostats. J Bacteriol. 2012;194:2165–80.PubMed CentralView ArticlePubMedGoogle Scholar
- Trautwein K, Kühner S, Wöhlbrand L, Halder T, Kuchta K, Steinbüchel A, et al. Solvent stress response of the denitrifying bacterium “Aromatoleum aromaticum” strain EbN1. Appl Environ Microbiol. 2008;74:2267–74.PubMed CentralView ArticlePubMedGoogle Scholar
- Rabus R, Trautwein K, Wöhlbrand L. Towards habitat-oriented systems biology of “Aromatoleum aromaticum” EbN1 - Chemical sensing, catabolic network modulation and growth control in anaerobic aromatic compound degradation. Appl Microbiol Biotechnol. 2014;98:1–18.View ArticleGoogle Scholar
- Studholme DJ, Dixon R. Domain architectures of σ54-dependent transcriptional activators. J Bacteriol. 2003;185:1757–67.PubMed CentralView ArticlePubMedGoogle Scholar
- Schumacher J, Joly N, Rappas M, Zhang X, Buck M. Structures and organisation of AAA+ enhancer binding proteins in transcriptional activation. J Struct Biol. 2006;156:190–9.View ArticlePubMedGoogle Scholar
- Bush M, Dixon R. The role of bacterial enhancer binding proteins as specialized activators of σ54-dependent transcription. Microbiol Mol Biol Rev. 2012;76:497–529.PubMed CentralView ArticlePubMedGoogle Scholar
- Helmann JD, Chamberlin MJ. Structure and function of bacterial sigma factors. Annu Rev Biochem. 1988;57:839–72.View ArticlePubMedGoogle Scholar
- Abril MA, Michan C, Timmis KN, Ramos JL. Regulator and enzyme specificities of the TOL plasmid-encoded upper pathway for degradation of aromatic hydrocarbons and expansion of the substrate range of the pathway. J Bacteriol. 1989;171:6782–90.PubMed CentralPubMedGoogle Scholar
- Shingler V, Bartilson M, Moore T. Cloning and nucleotide sequence of the gene encoding the positive regulator (DmpR) of the phenol catabolic pathway encoded by pVI150 and identification of DmpR as a member of the NtrC family of transcriptional activators. J Bacteriol. 1993;175:1596–604.PubMed CentralPubMedGoogle Scholar
- Fernández S, de Lorenzo V, Pérez-Martín J. Activation of the transcriptional regulator XylR of Pseudomonas putida by release of repression between functional domains. Mol Microbiol. 1995;16:205–13.View ArticlePubMedGoogle Scholar
- Shingler V, Pavel H. Direct regulation of the ATPase activity of the transcriptional activator DmpR by aromatic compounds. Mol Microbiol. 1995;17:505–13.View ArticlePubMedGoogle Scholar
- Delgado A, Salto R, Marqués S, Ramos JL. Single amino acids changes in the signal receptor domain of XylR resulted in mutants that stimulate transcription in the absence of effectors. J Biol Chem. 1995;270:5144–50.View ArticlePubMedGoogle Scholar
- Ramos JL, Marqués S, Timmis KN. Transcriptional control of the Pseudomonas TOL plasmid catabolic operons is achieved through an interplay of host factors and plasmid-encoded regulators. Annu Rev Microbiol. 1997;51:341–73.View ArticlePubMedGoogle Scholar
- Powlowski J, Shingler V. Genetics and biochemistry of phenol degradation by Pseudomonas sp. CF600. Biodegradation. 1994;5:219–36.View ArticlePubMedGoogle Scholar
- Herrmann H, Müller C, Schmidt I, Mahnke J, Petruschka L, Hahnke K. Localization and organization of phenol degradation genes of Pseudomonas putida strain H. MGG Mol Gen Genet. 1995;247:240–6.View ArticlePubMedGoogle Scholar
- Herrmann H, Janke D. Involvement of the plasmid pPGH1 in the phenol degradation of Pseudomonas putida strain H. FEMS Microbiol Lett. 1987;43:133–7.View ArticleGoogle Scholar
- Shingler V, Moore T. Sensing of aromatic compounds by the DmpR transcriptional activator of phenol-catabolizing Pseudomonas sp. strain CF600. J Bacteriol. 1994;176:1555–60.PubMed CentralPubMedGoogle Scholar
- Breinig S, Schiltz E, Fuchs G. Genes involved in anaerobic metabolism of phenol in the bacterium Thauera aromatica. J Bacteriol. 2000;182:5849–63.PubMed CentralView ArticlePubMedGoogle Scholar
- Delgado A, Ramos JL. Genetic evidence for activation of the positive transcriptional regulator Xy1R, a member of the NtrC family of regulators, by effector binding. J Biol Chem. 1994;269:8059–62.PubMedGoogle Scholar
- Ng LC, O’Neill E, Shingler V. Genetic evidence for interdomain regulation of the phenol-responsive σ54-dependent activator DmpR. J Biol Chem. 1996;271:17281–6.View ArticlePubMedGoogle Scholar
- Salto R, Delgado A, Michán C, Marqués S, Ramos JL. Modulation of the function of the signal receptor domain of XylR, a member of a family of prokaryotic enhancer-like positive regulators. J Bacteriol. 1998;180:600–4.PubMed CentralPubMedGoogle Scholar
- Wise AA, Kuske CR. Generation of novel bacterial regulatory proteins that detect priority pollutant phenols. Appl Environ Microbiol. 2000;66:163–9.PubMed CentralView ArticlePubMedGoogle Scholar
- Sarand I, Skärfstad E, Forsman M, Romantschuk M, Shingler V. Role of the DmpR-mediated regulatory circuit in bacterial biodegradation properties in methylphenol-amended soils. Appl Environ Microbiol. 2001;67:162–71.PubMed CentralView ArticlePubMedGoogle Scholar
- Garmendia J, Devos D, Valencia A, de Lorenzo V. À la carte transcriptional regulators: unlocking responses of the prokaryotic enhancer-binding protein XyIR to non-natural effectors. Mol Microbiol. 2001;42:47–60.View ArticlePubMedGoogle Scholar
- De Las Heras A, de Lorenzo V. Cooperative amino acid changes shift the response of the σ54-dependent regulator XylR from natural m-xylene towards xenobiotic 2,4-dinitrotoluene. Mol Microbiol. 2011;79:1248–59.View ArticlePubMedGoogle Scholar
- Pavel H, Forsman M, Shingler V. An aromatic effector specificity mutant of the transcriptional regulator DmpR overcomes the growth constraints of Pseudomonas sp. strain CF600 on para-substituted methylphenols. J Bacteriol. 1994;176:7550–7.PubMed CentralPubMedGoogle Scholar
- Skärfstad E, O’Neill E, Garmendia J, Shingler V. Identification of an effector specificity subregion within the aromatic-responsive regulators DmpR and XylR by DNA shuffling. J Bacteriol. 2000;182:3008–16.PubMed CentralView ArticlePubMedGoogle Scholar
- Devos D, Garmendia J, de Lorenzo V, Valencia A. Deciphering the action of aromatic effectors on the prokaryotic enhancer-binding protein XylR: A structural model of its N-terminal domain. Environ Microbiol. 2002;4:29–41.View ArticlePubMedGoogle Scholar
- Suresh PS, Kumar R, Kumar A. Three dimensional model for N-terminal A domain of DmpR (2-Dimethylphenol) protein based on secondary structure prediction and fold recognition. In Silico Biol. 2010;10:223–33.PubMedGoogle Scholar
- Wöhlbrand L, Rabus R. Development of a genetic system for the denitrifying bacterium “Aromatoleum aromaticum” strain EbN1. J Mol Microbiol Biotechnol. 2009;17:41–52.View ArticlePubMedGoogle Scholar
- Green M, Sambrook J. Molecular Cloning: A Laboratory Manual, vol. 1. 4th ed. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press; 2012.Google Scholar
- Kovach ME, Elzer PH, Hill DS, Robertson GT, Farris MA, Roop RM, et al. Four new derivatives of the broad-host-range cloning vector pBBR1MCS, carrying different antibiotic-resistance cassettes. Gene. 1995;166:175–6.View ArticlePubMedGoogle Scholar
- Champion KM, Zengler K, Rabus R. Anaerobic degradation of ethylbenzene and toluene in denitrifying strain EbN1 proceeds via independent substrate-induced pathways. J Mol Microbiol Biotechnol. 1999;1:157–64.PubMedGoogle Scholar
- Ramakers C, Ruijter JM, Lekanne Deprez RH, Moorman AFM. Assumption-free analysis of quantitative real-time polymerase chain reaction (PCR) data. Neurosci Lett. 2003;339:62–6.View ArticlePubMedGoogle Scholar
- Pfaffl MW. A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res. 2001;29:2002–7.View ArticleGoogle Scholar
- Zech H, Hensler M, Koßmehl S, Drüppel K, Wöhlbrand L, Trautwein K, et al. Adaptation of Phaeobacter inhibens DSM 17395 to growth with complex nutrients. Proteomics. 2013;13:2851–68.PubMedGoogle Scholar
- Simon R, Priefer U, Pühler A. A broad host range mobilization system for in vivo genetic engineering: transposon mutagenesis in Gram negative bacteria. Bio/Technology. 1983;1:784–91.View ArticleGoogle Scholar
- Schäfer A, Tauch A, Jäger W, Kalinowski J, Thierbach G, Pühler A. 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. 1994;145:69–73.View ArticlePubMedGoogle Scholar
- Büsing I, Höffken W, Breuer M, Wöhlbrand L, Hauer B, Rabus R. Molecular genetic and crystal structure analysis of 1-(4-hydroxyphenyl)-ethanol dehydrogenase from “Aromatoleum aromaticum” EbN1. J Mol Microbiol Biotechnol. 2015;25:327–39.Google Scholar