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Environmental Microbiology (2015) 17(1), 119–133

doi:10.1111/1462-2920.12511

Transcriptional and translational control through the 5′-leader region of the dmpR master regulatory gene of phenol metabolism

Anjana Madhushani,1 Teresa del Peso-Santos,1 Renata Moreno,2 Fernando Rojo2 and Victoria Shingler1* 1 Department of Molecular Biology, Umeå University, Umeå, SE 90187, Sweden. 2 Departamento de Biotecnologia Microbiana, Centro Nacional de Biotecnologia, CSIC, Madrid, Spain. Summary Expression of pathways for dissimilation of toxic aromatic compounds such as (methyl)phenols interfaces both stress-response and carbon catabolite repression control cascades. In Pseudomonas putida, carbon catabolite repression is mediated by the protein Crc – a translational repressor that counteracts utilization of less-preferred carbon sources as growth substrates until they are needed. In this work we dissect the regulatory role of the 5′-leader region (5′-LR) of the dmpR gene that encodes the master regulator of (methyl)phenol catabolism. Using deletion and substitution mutants combined with artificial manipulations of Crc availability in P. putida, we present evidence that a DNA motif within the 5′-leader region is critical for inhibition of the output from the Pr promoter that drives transcription of dmpR, while the RNA chaperone Hfq facilitates Crc-mediated translation repression through the 5′-leader region of the dmpR mRNA. The results are discussed in the light of a model in which Hfq assists Crc to target a sequence within a loop formed by secondary structure of the 5′-LR mRNA. Our results support the idea that Crc functions as a global translational inhibitor to co-ordinate hierarchical carbon utilization in Pseudomonads. Introduction The hallmark ability of bacteria to rapidly adapt to shifting nutritional conditions entails selection of the most Received 19 March, 2014; accepted 11 May, 2014. *For correspondence. E-mail [email protected]; Tel. (+46) 90 785 2534; Fax (+46) 90 772 630.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd

energetically favourable carbon source on offer. Pseudomonas putida strains are metabolically versatile bacteria that can use a wide range of alternative organic compounds available in the diverse soil and water habitats they occupy (Nelson et al., 2002). Their innately broad metabolic capacity is frequently augmented by catabolic plasmids that encode the ability to utilize aromatic pollutants, such as phenolics, as sole carbon and energy sources. However, production of the multiple enzymes of pathways for the catabolism of aromatic compounds is energetically expensive and aromatic compounds are generally toxic even to bacteria that can utilize them as growth substrates. Hence, the regulatory circuits that control their expression are highly attuned to host physiology and interfaces both global stressresponse and carbon-catabolism control networks to result in the hierarchical utilization of preferred substrates over less energetically favourable (non-preferred) aromatic substrates (reviewed in Shingler, 2003; 2004; Rojo, 2010). The pVI150 plasmid-encoded dmp-system (dimethylphenol) confers the ability to grow on (methyl)phenols to its native P. putida CF600 host and to the genome sequenced P. putida KT2440 strain (Shingler et al., 1989). Within the dmp-system, DmpR is the obligatory transcription activator of the σ54-dependent Po promoter that drives transcription of the dmp-operon and thus expression of the suite of Dmp-enzymes required for growth at the expense of (methyl)phenols (see Fig. 1A for schematic overview; reviewed in Shingler, 2004). Because DmpR can only assemble into its active multimeric form upon binding pathway substrates, it is the direct aromatic sensor that ultimately controls the ability to utilize (methyl)phenols as carbon sources (Shingler and Moore, 1994; O’Neill et al., 2001; Wikström et al., 2001, and references therein). Transcription from the σ70-dependent Pr promoter, which drives expression of the dmpR master regulatory gene, and transcription from the σ54-dependent Po dmpoperon promoter are extensively integrated within the nutritional and stress network orchestrated by the nucleotide alarmone ppGpp and the regulatory protein DksA (Sze and Shingler, 1999; Laurie et al., 2003;

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Fig. 1. DNA encoding the 5′-LR of dmpR inhibits transcription from Pr. A. Schematic of the pVI150-encoded dmp-system composed of dmpR gene and dmp-operon (not to scale). The locations of the DNA binding sites of DmpR (upstream activation sites, UAS2 and UAS1; inverted arrows) and IHF (shaded box) are shown relative to the σ54-Po operon promoter and the σ70-Pr promoter of dmpR that drive divergent but non-overlapping transcription from a 266 bp intergenic region. Inactive DmpR dimers (grey ovals) require binding of an aromatic effector before ATP binding can trigger multimerization into its hexameric active form, which is strictly required for transcription from the Po promoter. The expanded region below shows the sequence of the Pr promoter. The -35 and -10 elements of σ70-Pr promoter (del Peso-Santos et al., 2012) are highlighted in bold with matches to the consensus elements of σ70-promoters underlined. The 123 bp region that encodes the 5′-LR between the +1 A start site and the ATG initiation codon of dmpR (Johansson et al., 2008) is shown with the +1 and ATG codon highlighted by bold italics. Fusion points to the luxAB genes in the transcriptional reporter plasmids used in panels B–D are indicated. B. The graph shows the growth (squares) and luciferase activity profiles (circles) of LB-cultured P. putida KT2440 harbouring either a Pr transcriptional reporter plasmid with the 5′-LR region (pVI938, closed symbols) or an equivalent reporter lacking the 5′-LR (pVI2310, open symbols). Data are the average +/− standard errors of duplicate determinations from each of two independent experiments normalized by setting stationary phase values of the +5′-LR plasmid as 1. C. As panel B, but showing the stationary phase values obtained using the series of Pr transcriptional reporters (illustrated in panel A) with progressively shorter 3′ ends of the 5′-LR DNA. Data are the average of duplicate determinations from each of two independent experiments +/− S.D. D. The graph shows the mRNA stabilities of luxAB transcripts produced from Pr reporter plasmids with (pVI938, closed symbols) or without (pVI2310, open symbols) the 5′-LR. Data are the average +/− standard errors of triplicate determinations in each of two independent experiments, and are expressed by setting the values at time point 0 as 100%. Differences in relative values at this point were 6.5 +/− 0.1 fold higher for pVI2310 (-5′-LR), than pVI938 (+5′-LR). Data are the average of triplicate determinations from each of two independent experiments +/− standard deviation (SD).

Bernardo et al., 2006; 2009; Szalewska-Palasz et al., 2007; Johansson et al., 2008). These two global regulatory molecules frequently act together by directly targeting RNA polymerase (RNAP) to alter its properties and are both involved in increasing the levels of all the proteins required for σ54-Po promoter activity. These include (i) the pool of σ54-RNAP holoenzyme, (ii) the DNAbending protein integration host factor (IHF), which facili-

tates efficient interaction between DmpR and σ54-RNAP, and (iii) DmpR itself, through directly stimulating the performance of σ70-RNAP at the Pr promoter of the dmpR gene (Johansson et al., 2008; del Peso-Santos et al., 2011). Furthermore, in the presence of (methyl)phenols and under nutrient-limiting conditions that elicit ppGpp production, activity at the DmpR-dependent Po promoter further stimulates transcription from the non-overlapping

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 119–133

Control through the 5′-leader region of the dmpR gene Pr promoter, creating a feed-forward loop to enhance Pr activity under energy-limiting conditions (Johansson et al., 2008). Integration within the ppGpp/DksA network provides a general mechanism for subverting expression of the specialized Dmp-enzymes for phenol catabolism when preferred alternative carbon sources are present. However, specific mechanisms of carbon repression control (CRC) also exist to coordinate utilization of preferred substrates over energetically less-favourable ones. Although specific CRC systems are common in bacteria, the molecular mechanisms underlying selective utilization of available carbon sources differs greatly between species (Gorke and Stulke, 2008; Rojo, 2010; Valderrama et al., 2014 and references therein). In Pseudomonads, the translational repressor protein Crc is the primary mediator of CRC (Moreno et al., 2009a; Sonnleitner et al., 2009). Messenger RNAs that are specifically targeted by Crc bear in common a catabolite activity (CA)-motif close to or overlapping the ribosome binding site (RBS) (Moreno et al., 2009b; Sonnleitner et al., 2009; 2012). The levels of Crc vary little under different growth regimes; however, when preferred carbon sources are depleted, Crc-mediated translational repression is antagonized by production of specific small RNAs (sRNAs) which contain multiple CA-motifs – CrcZ in Pseudomonas aeruginosa, CrcZ and its homologue CrcY in P. putida. These sRNAs are believed to mediate de-repression by sequestering Crc, leading to efficient translation of Crc-targeted mRNAs that encode regulators, uptake systems and enzymes required for utilization of non-preferred substrates (Moreno et al., 2010, Rojo, 2010; Sonnleitner and Haas, 2011, and references therein). The impact of Crc regulation on the dmp-system has not previously been investigated. In addition to stimulation of Pr activity by ppGpp and DksA under nutritional stress conditions, a 5′-leader region (5′-LR) of dmpR has also been implicated in governing DmpR levels (Johansson et al., 2008), suggesting additional control mechanisms. 5′-LRs of bacterial mRNAs frequently serve as regulatory hubs to control expression of downstream protein-coding sequences through forming riboswitches, or serving as the targets for RNA binding proteins or interacting sRNAs. Pairing of sRNAs with their target mRNA is usually facilitated by the abundant Hfq protein – an RNA chaperone that can simultaneously bind both the sRNA and mRNA to promote appropriate base pairing (see Garst et al., 2011, Storz et al., 2011 and De Lay et al., 2013 for recent reviews). In this work, we present evidence that the 5′-LR region of the dmpR gene is intimately involved in controlling the levels of DmpR by two distinct mechanisms. At the transcriptional level, reduced output from the Pr promoter was traced to a DNA motif (ATAAATA) present in the initially

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transcribed region, while translational repression was traced to a CA-motif of the 5′-LR mRNA. We found that, in vitro, Crc was unable to form a complex with the CA-motif of the RNA on its own, but could do so in the presence of the Hfq RNA chaperone that bound the CA-motif directly. This latter finding is in line with the recent discovery that Crc and Hfq act together to form complexes with other RNA targets containing CA-motifs (Moreno et al., 2014), which has implications for how Crc availability is controlled through sequestration by CrcZ and its homologues. Results The initially transcribed 5′-leader DNA inhibits transcription from Pr The σ54-Po and σ70-Pr promoters drive divergent transcription from within the 406 bp intergenic region between the initiation codons of dmpR and the dmp-operon (Fig. 1A). Previous primer extension and deletion analysis has demonstrated that the initially transcribed region of the dmpR gene encodes a 123 nt long 5′-LR of the dmpR mRNA (Fig. 1A; Johansson et al., 2008). The presence of the DNA encoding the 5’-LR has an ∼ 5-fold repressive effect on the output from Pr in P. putida KT2440 (Fig. 1B) and Escherichia coli (data not shown) cultured in Luria– Bertani (LB), as detected using luciferase (luxAB) transcriptional reporter plasmids that either possess or lack this region. The enhanced output seen at the exponentialto-stationary phase transition, which occurs irrespective of the presence or absence of the 5′-LR, is due to nutrition depletion and consequent ppGpp production that results in direct stimulation of the performance of σ70-RNAP at the Pr promoter of the dmpR gene (Johansson et al., 2008; del Peso-Santos et al., 2011). Additional deletion analysis, using otherwise identical luciferase transcriptional reporter plasmids that possess progressively shorter regions of the 5′-LR, demonstrated that possession of the promoter-distal portion of the 5′-LR DNA is critical for its inhibitory effect on Pr output (compare pVI1027 and pVI2307, Fig. 1C). 5′-LRs can be targeted to modulate gene expression by a variety of different mechanisms that involve upregulation or downregulation of transcription or, alternatively, can function at the level of translation efficiency and/or mRNA stability. In close agreement with luciferase activity assays, quantitative PCR revealed ∼ 6.5-fold higher levels of dmpR transcripts from P. putida KT2440 harbouring the luciferase transcriptional reporter plasmid lacking the 5′-LR DNA as compared with a counterpart that possessed the 5′-LR. Determination of mRNA stabilities of these transcripts showed indistinguishable halflives of ∼ 5 min (Fig. 1D). Because the luxAB reporter genes of the transcriptional reporter plasmids come

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 119–133

122 A. Madhushani et al. equipped with their own RBS, the combined data in Fig. 1 lends strong support to the idea that the inhibitory effect of the 5′-LR is mediated at the DNA (rather than mRNA stability) level, through a mechanism that alters the number of full-length transcripts from the Pr promoter.

Transcriptional inhibition by the 5′-leader DNA is orientation-independent and can exert its effects on diverse RNA polymerase holoenzymes To be able to further manipulate the 5′-LR DNA we employed a luciferase transcriptional reporter plasmidbearing DNA encompassing the -38 to +1 region of the Pr promoter (see plasmid 2, Fig. 2A). This plasmid has a BglII site located immediately downstream of the +1 transcriptional start that was used in conjunction with a downstream NotI site to reconstruct the 5′-LR in either its native or inverse orientation, and to insert an equivalent size region of unrelated ‘random’ DNA (plasmids 3 to 5, Fig. 2A). The activities of the resulting Pr-luxAB derivatives were then compared with native counterparts with or without the 5′-LR in P. putida KT2440 using a luciferase plate test assay. Comparison of the in vivo output from Pr using these constructs showed that the repressive effect of the 5′-LR is reproduced when reconstituted in either orientation (compare 3 and 4 with 1 and 2, Fig. 2B). Furthermore, the data also showed that repression by the 5′-LR DNA is specific since it was not mimicked by the presence of random DNA, which resulted in a derivative that exhibited the same elevated levels of Pr output as seen in the absence of the 5′-LR (compare 2 and 5, Fig. 2B). These results reinforce the notion that the repressive effect of the 5′-LR is mediated at the DNA rather than the RNA level.

¢ ′ ′ ′ ′

Given that the 5′-LR inhibits output from the weak σ70dependent Pr promoter, we were interested to determine if the 5′-LR DNA could also inhibit output from strong σ70-promoters and/or promoters dependent on other σ-factors. To address this question we generated analogous pairs of luciferase transcriptional reporter plasmids with known promoters, with or without the 5′-LR DNA upstream of the luxAB genes. The specific promoters chosen for this analysis represent a range of promoter strengths and contain the distinctive promoter DNA motifs for different forms of the RNAP holoenzyme (see Supporting Information Fig. S1). The promoters used include the strong σ70-λPL promoter and σ70-PrT-11A promoter – a hyperactive derivative of Pr (del Peso-Santos et al., 2011), the weak σ70-Paer1, σFliA-Paer2 and σE-PhtrA promoters, and the strong σ54-Po promoter. Comparison of the in vivo output from these reporters showed that the 5′-LR DNA can mediate a repressive effect on output from both powerful and weak σ70dependent promoters, and promoters dependent on the alternative σ-factors σFliA, σE and σ54 (Supporting Information Fig. S1). Thus, we conclude from this data that, in addition to being orientation-independent, the transcriptional inhibition mechanism can exert its effect on diverse RNA polymerase holoenzymes.

Transcriptional inhibition by the 5′-leader DNA maps to a promoter-distal ATAAATA DNA motif The data in Fig. 1C implicated the promoter-distal DNA of the 5′-LR as critical in mediating transcriptional inhibition. To ascertain if all or only the promoter-distal DNA of the 5′-LR is required to inhibit output from Pr, we used the same strategy as outlined in the preceding section to generate an additional series of transcriptional reporters

Fig. 2. A short Pr promoter-distal ATAAATA motif is responsible for transcriptional inhibition. A. Schematic illustration of the extent of the 5′-LR in Pr-luxAB transcriptional reporter plasmids. Open regions within the shaded box indicate the extent of DNA deletions within the 5′-LR DNA, while an oblique line indicates the presence of a non-native BglII site introduced to allow manipulation of the 5′-LR DNA. Plasmids are: 1 (pVI1017); 2 (pVI955); 3 (pVI2311); 4 (pVI2313); 5 (pVI2312), 6 (pVI2314 Δ1, pVI2315 Δ2, pVI2316 Δ3, and pVI2317 Δ4); 7 (pVI2318 Δ5); 8 (pVI2319 Δ6); 9 (pVI2320 Δ7); 10 (pVI2321 Δ8). Key DNA sequences of plasmids harbouring the Δ1–Δ8 deletions are also shown. B. Representative luciferase plate test assays employing P. putida KT2440 harbouring the indicated reporter plasmids.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 119–133

Control through the 5′-leader region of the dmpR gene that progressively lack larger portions of the 5′-LR as depicted in Fig. 2A. The first four deletion derivatives (Δ1– Δ4), which lacked up to 70% of the 5′-LR DNA, all mediated inhibition, while deletion of all but the 3′-distal end of the 5′-LR DNA (as in Δ5) did not (compare plasmids 6 and 7 in Fig. 2B). Examination of the DNA sequences of Δ4 and Δ5 indicated that repression was mediated via a 15–16 bp AT-rich region present in Δ4 but lacking in Δ5 (Fig. 2B). Further deletion analysis (Δ6–Δ8, Fig. 2A and B) traced this property to a 7 bp ATAAATA DNA motif, which is present in all derivatives that mediate inhibition of Pr output but absent from all those that do not. The symmetry of the ATAAATA motif likely underlies the ability of the 5′-LR DNA to exert its effects in both orientations. The 5′-LR mediates translational control of the dmpR mRNA As illustrated in Fig. 2B, the Δ8 derivative only possesses the promoter-distal 21 bp of the 5′-LR DNA. These 21 bp encompasses the ATAAATA DNA motif and the RBS for translation of dmpR. Because the Δ8 derivative still maintains wild-type output from Pr, this deletion was chosen to determine potential effects of the 5′-LR on translation of the dmpR mRNA. To this end, Δ8 was introduced into a Pr–dmpR expression plasmid (Fig. 3A). Activity of this derivative in comparison with an otherwise identical wildtype Pr-dmpR expression plasmid was monitored using PP980 – a P. putida KT2440 derivative carrying a Po-luxAB transcriptional fusion on its chromosome. Because the Po promoter is strictly dependent of DmpR for activity, this genetic system provides a read-out of DmpR levels in the cell. Comparison of the activities of the DmpR-dependent Po promoter revealed a major difference, with the Δ8 derivative resulting in vastly higher output (> 60-fold) than the wild-type counterpart during the exponential phase of growth (Fig. 3B), strongly suggesting elevated levels of DmpR during this growth phase. This was verified by Western analysis that showed readily detectable exponential phase levels of DmpR from the Δ8 derivative (Fig. 3C). In contrast, at the exposure shown, exponential phase DmpR levels were undetectable from the derivative possessing a wild-type 5′-LR. Because the Δ8 deletion derivative does not result in increased transcriptional output from the Pr promoter (Figs 2B and 3D), these data demonstrate that the 5′-LR region controls the efficiency of translation of dmpR from its mRNA. As illustrated in Fig. 3A, the Δ8 deletion removes half of a potential RNA CA-motif associated with translational repression mediated by Crc (consensus AAnAAnAA; Moreno et al., 2009b). Thus, Δ8 may constitute a

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CA-target-site-deficient mutant. Because this potential CA-motif overlaps the RBS for dmpR, we also constructed a Pr-dmpR derivative with a full-length 5′-LR but with the same half of the CA-motif substituted by four consecutive C residues (denoted 4C, Fig. 3A), which should leave the RBS intact. As shown by the data in Fig. 3, the 4C-substitution derivative exhibited very similar properties as the Δ8-deletion derivative, tracing the effect on translation of dmpR to this region of the 5′-LR. Taken together, the data in Fig. 3 implicates the identified AAAAAUAA RNA motif as a site for translational control of dmpR. Translational control through the 5′-LR requires Crc As outlined in the introduction, the available levels of Crc in the cell are thought to decrease in the stationary phase because of the action of two Crc-sequestering sRNAs (CrcZ and CrcY). Deletion of both CrcZ and CrcY is required to observe changes in the action of Crc at the translational level (Moreno et al., 2012). Therefore, to test the influence of Crc on translation of dmpR, we used both a Crc null and a double CrcZ/CrcY-null derivative of P. putida KT2440 for comparison with the wild-type strain. A Po-luxAB transcriptional reporter plasmid carrying Pr-dmpR in its native configuration completed the genetic system (Fig. 4A). As in the preceding section, the Po-luxAB luciferase reporter activity was used as the read-out of the DmpR levels in the cell. Because lack of the CA-motif for Crc increased DmpR protein levels during exponential growth and consequently elevated output from the DmpR-dependent Po promoter (Fig. 3), lack of Crc would be expected to produce the same phenotype. As anticipated, lack of Crc resulted in higher luciferase activities in the exponential phase, where Crc is normally available to inhibit translation of dmpR in LB-cultured cells (compare Fig. 4B and C). Conversely, lack of CrcZ and CrcY resulted in decreased activity in the stationary phase (compare Fig. 4B and D), which is consistent with lack of Crc sequestration by these stationary phase-produced sRNAs. It is notable from Fig. 4C that lack of Crc has an adverse effect on growth of P. putida in rich medium. Therefore, as a complementary approach, we also artificially decreased available intracellular levels of Crc using plasmids that independently overexpress CrcZ or CrcY from an IPTG inducible promoter. Induced expression of either of these two Crc-sequestering sRNA resulted in greatly increased output from the DmpR-dependent Po promoter during exponential growth and a reduced growth rate (Fig. 5A and B), reminiscent of that seen in the Crcnull P. putida strain (Fig. 4C). As expected, the reporter system used is immune to the effects of overexpression of

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Fig. 3. Translational control of the dmpR mRNA. A. Schematic illustration of the chromosomal DmpR-dependent Po-luxAB reporter strain fusion and the plasmid-encoded Pr-dmpR expression plasmids used to indirectly monitor the effect of manipulations of the 5′-LR on DmpR levels. The expanded region below shows the promoter-distal sequence of the 5′-LR DNA present in the indicated constructs aligned with the motif encoding a potential CA-motif of the mRNA (highlighted in bold, consensus Moreno et al., 2009a,b). The shaded box illustrates the likely extent of the RBS that overlaps the CA-motif. B. The graph shows the luciferase activity profiles of the Po-luxAB reporter strain PP980 carrying different Pr-dmpR expression plasmids grown in LB in the presence of 2 mM 2-methylphenol – the most potent effector of DmpR activity. Plasmids were the wild-type Pr-WT-dmpR fusion (pVI401; filled circles) or its Crc-motif-deficient counterparts Pr-Δ8-dmpR (pVI2342; open down triangles) or Pr-4C-dmpR (pVI2343; open triangles). The results are the average of duplicate determinates from two to four independent cultures. For clarity, only a single growth profile (filled squares) is shown. C. Western blot analysis of DmpR levels in cells as under panel B) harvested during the exponential and stationary phases of growth. Images are from 10 μg and 5 μg crude extracts separated by 10% SDS-PAGE and analysed using anti-DmpR antibodies. D. The graph shows the exponential phase output of comparable transcriptional Pr-luxAB transcriptional reporter plasmids in PP980 grown in LB in the absence of 2 mM 2-methylphenol. Plasmids were Pr-WT-luxAB fusion (pVI2338) or its CA-motif-deficient counterparts Pr-Δ8-luxAB (pVI2339) and Pr-4C-luxAB (pVI2340). Data are the average +/− standard errors of eight determinations from independent cultures normalized by setting values for the Pr-WT plasmid (pVI2338) as 1.

either CrcZ or CrcY in a Crc-null background (Fig. 5C and D), confirming that the phenotype derives from the ability of CrcZ and CrcY to modulate Crc activity. Taken together, the data in Figs 4 and 5 illustrate that Crc exerts a negative effect on translation of dmpR during the exponential phase, which is counteracted by the sRNAs CrcZ and CrcY in the stationary phase. These data corroborate the prevailing model for control of Crc function based on other Crc-targeted processes (Rojo, 2010 and references therein).

Both Hfq and Crc are required to form a stable complex with the AAAAAUAA RNA motif RNA binding activity observed with His-tagged Crc preparations has been attributed to contaminating E. coli Hfq, which co-purifies in minute amounts because of its naturally His-rich C-terminal extension (Milojevic et al., 2013). Therefore, we prepared both P. putida Crc and P. putida Hfq-His through alternative strategies to avoid such contamination. As detailed in Supporting Information, Crc was

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Fig. 4. Translational regulation of dmpR by Crc and the sRNAs CrcZ and CrcY. A. Schematic illustration of the Po-luxAB/Pr-dmpR pVI466 reporter used to indirectly monitor the effect on DmpR levels as described under Fig. 3. The graphs show growth (black squares) and luciferase activity profiles (cyan circles) of the DmpR-dependent Po promoter carried on pVI466 in: B. The wild-type P. putida KT2440 strain. C. A Crc-null counterpart (KT2440-crc::Gm). D. A double CrcZ/CrcY-null derivative (PP3386; KT2440-ΔcrcY::Gm/ΔcrcZ::Tc). E. A Hfq-null derivative (KT2440-Δhfq). F. A double Hfq/Crc-null derivative (KT2440-Δhfq crc::Gm). G. Western blot analysis of DmpR protein levels in cells grown as under panels B–F and harvested at different times during the exponential (Exp.; OD600 0.2–0.3) and stationary phase (Stat.; OD600 2–3) of growth. Images are from 5 μg and 10 μg crude extracts separated by 10% SDS-PAGE and analysed using anti-DmpR antibodies. Note that the prolonged growth time required to reach stationary phase for the Hfq-null and Hfq/Crc-null strains (∼ 20 h) resulted in detection of a cross-reactive band.

purified via a cleavable intein tag, while Hfq-His derivatives were purified from a Hfq-null E. coli expression strain. These protein preparations were used to assess the ability of Crc to bind the identified AAAAAUAA RNA motif within the 5′-LR of the dmpR mRNA using RNA electro-mobility shift assays (EMSAs). The assays employed 26 nt long 5′-FAM-labelled fluorescent RNA probes that encompassed the potential wild-type CA-motif

(Fig. 6) or a variant with the 4C substitution within the CA-motif (Supporting Information Fig. S2). Consistent with recent findings, P. putida Crc alone was unable to bind to the CA-motif (Fig. 6A and Supporting Information Fig. S2A). Hfq-His could form a complex with the RNA; however, only when present at high concentrations (≥ 12.5 μM, Fig. 6B and C). In marked contrast, a distinct complex – migrating higher than that with Hfq-His

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126 A. Madhushani et al. Fig. 5. Effects of CrcZ and CrcY overexpressions on expression of DmpR. Growth (squares) and luciferase activity profiles (circles) of the DmpR-dependent Po promoter carried on the dmpR-Pr-Po-luxAB plasmid pVI466 as under Fig. 4. Assays were performed with: P. putida KT2440 harbouring: A. CrcZ overexpression plasmid (p424-Z). B. CrcY overexpression plasmid (p424-Y); Crc-null counterpart of the P. putida KT2440 strain harbouring: C. Plasmid p424-Z. D. Plasmid p424-Y. Cultures were grown in the absence (closed symbols) or presence (open symbols) of 1 mM IPTG to induce expression of the Crc-sequestering sRNAs CrcZ (from p424-Z) or CrcY (from p424-Y).

– was formed in the co-presence of low concentrations of Crc and Hfq-His (e.g. 1 μM), i.e. at concentrations that do not elicit a complex when either protein was present alone (Fig. 6D and E). These results clearly demonstrate that both proteins contribute to the formation of this complex. As expanded on in the discussion, these results suggest that the RNA chaperone Hfq is required for interaction of Crc with RNA, or that Crc modifies the RNA binding of Hfq (or vice versa). Using the 4C mutant RNA probe (where the CA-motif is disrupted by substitution with 4C residues: AAAAAUAA to CCCCAUAA), we found that both the formation of the apparent low affinity complex with Hfq-His alone and that of the apparent high affinity complex in the co-presence of Crc and Hfq-His was dependent on the integrity of the CA-motif (Supporting Information Fig. S2B–E). Since all complex formation was abolished by the 4C substitution, these results pinpoint the 5′-proximal 4A nucleotides of the CA-motif as critical for the action of Hfq. The ringshaped hexamer of E. coli Hfq exhibits two distinct RNA binding interfaces – a proximal face that binds U-rich sequences and a distal face that binds A-rich ARN sequences (where R is a purine and N any ribonucleotide; reviewed in De Lay et al., 2013). The 102 residue Hfq from E. coli and the 86 residue Hfq from P. putida are highly homologous, exhibiting > 90% identity in their N-terminal 72 residues (Supporting Information Fig. S3). Therefore, to explore the role of Hfq further, we purified

two mutant versions of P. putida Hfq-His with single amino acid substitutions: Y25D or K56A, which based on mutagenesis of E. coli Hfq would independently abolish its binding to A-rich or U-rich RNA targets, respectively (Mikulecky et al., 2004; Link et al., 2009). Consistent with the requirement for the 5′-proximal 4As of the CA-motif, the Y25D substitution (A-rich binding deficient) rendered Hfq incapable of binding the CA-motif either in isolation or in the presence of Crc. Conversely, the K56A substitution (U-rich binding deficient) behaved essentially as the wildtype (Fig. 6C and F). Hence, from this data we conclude that the A-rich RNA binding properties of the distal face Hfq are essential for the co-action of Hfq and Crc in generating the high affinity stable complex with the CA-motif of the dmpR mRNA. Hfq is required for efficient Crc-mediated control in vivo The results in Fig. 6 and Supporting Information Fig. S2 identify Hfq as a co-actor in translational regulation mediated by Crc. As such, these results would predict that lack of Hfq, like lack of Crc, would alleviate translational inhibition of dmpR during the exponential phase of growth. To test this prediction we monitored intracellular levels of DmpR in a P. putida Hfq-null strain in comparison to those of the wild-type and Crc-null strains using a DmpRdependent Po-luxAB reporter and Western analysis as previously.

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Fig. 6. Hfq assists Crc association with RNA in vitro. EMSA analysis employing a 26 nt RNA probe (+89 to +115 relative to the transcriptional start from Pr) encompassing the potential Crc binding site (bold and underlined). A. Reactions in the absence and presence of increasing concentrations of Crc (0.15, 0.3, 0.6, 1.2, 2.4 and 4.8 μM). B. Reactions in the absence and presence of increasing concentrations of wild-type Hfq-His (0.8, 1.6, 3.125, 6.25, 12.5 and 25 μM). C. Titrations of wild-type (WT) and the indicated mutant variants of Hfq-His (0, 12.5, 25 or 50 μM). D. Reactions in the presence of 1 μM Hfq-His and increasing amounts of Crc (0, 0.09, 0.187, 0.375, 0.75, 1.5 and 3 μM). E. Reactions in the presence of 1 μM Crc and increasing amounts of Hfq-His (0, 0.187, 0.375, 0.75, 1.5, 3 and 6 μM). F. Reactions with 0.5 μM Crc in the absence or presence of 3 μM of the indicated Hfq-His protein. Open arrowheads indicate free RNA; filled arrowheads indicate riboprotein complexes.

Because Hfq is a global regulator that assists interactions between diverse small regulatory RNAs and their target mRNAs, lack of cellular Hfq will have pleiotropic effects. The Hfq-null derivative of P. putida KT2440 is very slow growing (Fig. 4E) making direct comparison with the wild-type and Crc strains somewhat questionable. Nevertheless, as in cells lacking Crc, transcription from the DmpR-dependent Po promoter was high during the exponential phase in the Hfq-null strain, which contrasts the lack of activity in this phase of growth in the wild-type strain (compare Fig. 4B, C and E). Consistently, enhanced DmpR levels were detected in extracts from cells of the Crc-null and Hfq-null strains cultured to the exponential phase, relative to the levels found in the wildtype strain (Fig. 4G). These data support the idea that Crc is dependent on Hfq to mediate its repressive effect. To

further investigate this idea, we used a double Crc/Hfqnull strain that exhibited the same poor growth as the Hfq-null strain (Fig. 4F). In contrast to the major exponential phase difference observed between the wild-type and Crc-null strains (> 15-fold), lack of Crc in the Hfq-null background only had a very minor effect (∼ 1.2-fold). These data reinforce the notion that Hfq is required to enable Crc to mediate a repressive effect on translation of dmpR and that this repressive effect is mediated through the CA-motif of the 5′-LR mRNA. Discussion The aromatic sensor–regulator DmpR controls transcription from the σ54-Po promoter and thus formation of the catabolic enzymes encoded by the dmp-operon. In this

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128 A. Madhushani et al. work, we demonstrate that the 5′-LR of the dmpR gene functions as a regulatory hub to control DmpR levels by two distinct mechanisms. At the level of transcription, inhibition of full-length transcripts was traced to an A-rich ATAAATA DNA motif located downstream of the Pr promoter (Figs 1 and 2). At the translational level, we found that Hfq aids Crc to promote formation of a stable ribonucleoprotein complex through a CA-motif of the 5′-LR RNA in vitro and facilitates Crc-dependent repression in intact cells (Figs 4–6). Many chromosomal and plasmid-encoded phenolic catabolic systems exist that have analogous genetic organizations as that found in the dmp-system, i.e. with a dmpR-like gene divergently transcribed from genes encoding catabolic enzymes (Shingler, 2004). Within these different systems, the sequence of the 5′-LR region is more highly conserved than other regulatory regions, and the coding regions of cognate genes (Supporting Information Fig. S4), and all maintain the integrity of the ATAAATA- (DNA) and CA(RNA) motifs. Thus, the notable conservation of the 5′-LR region suggests strong evolutionary pressure to maintain the regulatory processes that target these motifs. On the mechanisms of transcriptional repression through the ATAAATA DNA motif Inhibition of the number of full-length transcripts mediated by the entire 5′-LR DNA (Fig. 1) can be simply mimicked by the presence of the ATAAATA motif at different locations relative to the Pr promoter (Fig. 2). While the ATAAATA motif is naturally located distally from Pr (108 bp downstream), it can also exert its effect when relocated to 14 bp downstream from Pr (as in the largest deletion, Δ8). Because the ATAAATA motif can function in the absence of native upstream or downstream sequences (Figs 1 and 2), it appears unlikely that reduced transcript levels are the result of any kind of transcription attenuation mechanism, which all rely on changes in the secondary structure of upstream and/or downstream RNA and interaction of an RNA element of a stem loop structure with the transcriptional machinery (reviewed in Naville and Gautheret, 2009). We can envisage two plausible alternative mechanisms that could account for the repressive effect of this motif. The first of these involves transcriptional pausing because of its similarity to the -10 element of σ70-promoter (consensus TATAAT). Because σ70 can stay partially attached to RNAP during the first ∼200 nt of elongation of the RNA, its binding to promoter mimics within initially transcribed regions can impede the progression of RNAP to result in a lower net number of full-length transcripts (Ring et al., 1996; Ko et al., 1998; Leibman and Hochschild, 2007). However, this does not appear to be the explanation in the case of the ATAAATA motif, since it can mediate inhi-

bition from promoters dependent on diverse RNAP holoenzymes that recognize very different promoter DNA elements (Supporting Information Fig. S1). An alternative likely scenario is that the ATAAATA motif serves as a binding site for a repressor that would impede progression of RNA polymerase. Because repression at the level of the transcription was also seen in the heterologous host E. coli, such a putative repressor would have to be encoded within the genomes of both E. coli and P. putida KT2440. The ATAAATA motif bears similarity to the consensus sequence for E. coli HNS (TCGATAAATT). HNS-like proteins bind AT-rich DNA through a ‘Q/RGR’-motif of the protein (Gordon et al., 2011). The genome of P. putida KT2440 encodes five HNS paralogues (Renzi et al., 2010), namely PP0017, PP1366 (also known as TurA), PP2947, PP3693 and PP3765 (also known as TurB). By analogy to the repressive action of TurA on toluene and xylene metabolism (Rescalli et al., 2004), any of these proteins could potentially target the ATAAATA motif to bring about reduced levels of transcription. However, evaluation of this possibility by monitoring transcription from Pr with or without the 5′-LR in P. putida strains individually devoid of each of these HNS-like proteins, refuted this idea (T. del PesoSantos et al., unpublished). However, since the ‘Q/RGR’motif functions in otherwise structurally unrelated proteins (Gordon et al., 2011), additional candidates are likely. The identity of the potential ATAAATA binding repressor is the subject of ongoing research. On the mechanisms of translational repression through the CA-RNA motif Our dissection of the role of Crc and the CA-motif in mediating translational repression of dmpR corroborates and extends the prevailing model for the action and control of Crc activity to include the hexameric Hfq RNA chaperone. In vivo, lack of either of these proteins results in high DmpR levels under conditions where translation of dmpR is normally inhibited (Figs 3–5). In vitro, Crc is unable to form a riboprotein complex with the CA-motif directly (Fig. 6A); however, in the presence of Hfq, Crc triggers formation of a stable complex that is quite distinct from that produced by binding of Hfq alone (Fig. 6). We found that the ability of Crc to trigger formation of the stable riboprotein complex is dependent on both the ability of Hfq to bind to the CA-motif and on the integrity of the A-rich RNA binding properties of the distal face of Hfq (Fig. 6 and Supporting Information Fig. S2). Therefore, it seems reasonable to propose that binding of Hfq initiates the series of events that leads to formation of the stable complex. These findings are in agreement with recent results showing Crc and Hfq form co-riboprotein complexes with the CA-motifs of CrcZ (Moreno et al., 2014).

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 119–133

Control through the 5′-leader region of the dmpR gene Hfq is an adaptable protein – it can simultaneously bind sRNAs and mRNA, modify the structure of mRNAs and/or interact with proteins involved in transcription, translation and mRNA stability (reviewed in De Lay et al., 2013). By analogy to the action of Hfq in assisting other RNA-based regulatory processes, initial binding of Hfq may simply present the RNA in a configuration suitable for Crc binding, or may promote Hfq/Crc protein : protein interactions that modify the RNA binding properties of one or other of the two proteins. However, distinguishing between these mechanistic possibilities requires further experimentation. Irrespective of precise details of the complex, Hfq would predictably assist Crc targeting of CA-motifs of mRNAs, and its association with the multiple CA-motifs of CrcZ and CrcY that are thought to sequester Crc when preferred carbon sources are absent. The CA-motif of the dmpR mRNA is located on a loop within one of two alternative potential structures of the 5′-LR, but is masked in the alternative structure (Fig. 7), raising the possibility that initial binding of Hfq result in restructuring of the RNA. The CA-motif partially overlaps the region that is needed for binding of the ribosomes. The model that emerges from our findings is that a Crc/ Hfq riboprotein complex and ribosomes would compete for their overlapping binding sites on the 5′-LR RNA. Within this scenario, in the presence of a preferred carbon source (high available Crc levels) translation would be blocked, resulting in low levels of DmpR. As a consequence of low levels of DmpR, the Po promoter that drives expression of the specialized catabolic enzymes would be essentially inactive and so (methyl)phenols, as non-preferred carbon sources, would not be consumed. However, in the absence of a preferred carbon source (low Crc availability because of sequestering by CrcZ and CrcY), the RBS is free, resulting in high levels of DmpR, Po activity and thus expression of the enzymes for (methyl)phenol catabolism.

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Concluding remarks Under energy-limiting conditions, the stimulatory action of ppGpp and DksA on Pr promoter activity and de-repression of Crc/Hfq-mediated translational repression would combine robust transcription of dmpR with efficient translation of its mRNA, and thus result in high levels of the Dmp-enzymes for metabolism of the nonpreferred substrates – (methyl)phenols. Conversely, under high-energy growth conditions, low ppGpp levels and Crc/Hfq-mediated translational repression would work hand-in-hand to subvert expression of DmpR at both the transcriptional and translational levels. The synergism of inputs from these global regulatory networks makes seeming biological sense – effective muting of the dmpsystem when preferred carbon sources are present. However, the transcriptional inhibition mediated through the ATAAATA DNA motif remains an enigma – what is the regulatory logic of having this counteractive system operating simultaneously? One speculative explanation could be that signal-responsive control through this negative regulatory element may form a so-called ‘incoherent feedforward loop’, in which simultaneously acting opposing forces can result in pulse generators and/or repose accelerators (Alon, 2007). It is notable that the ATAAATA-DNA and the CA-RNA motifs are confined to the Pr promoter-distal region of the 5′-LR, yet the sequence of the entire 5′-LR is highly conserved in related systems. While this may simply reflect evolutionary selection for an appropriate secondary structure for interaction of Crc/Hfq with the RNA, examination of the sequence of the 5′-LR presents intriguing alternative possibilities because of two additional potential regulatory features. The first of these is a short upstream open reading frame (uORF, highlighted in grey in Fig. 7). Translation of this uORF could potentially mediate translational coupling to modulate translation of the downstream dmpR

Fig. 7. Predicted secondary structures and regulatory features of the dmpR 5′-LR mRNA. The schematic shows two alternative predicted secondary structures of the 5′-LR of dmpR with the CA-motif shown in orange and the RBS highlighted in pale orange. Other potential regulatory features within this structure are: (1) A short uORF (grey) with a potential Val (GUG) or Met (AUG) start codon, bounded by a UAG termination codon (highlighted in red); and (2) a potential target site for the small regulatory RNA PhrS (blue). The alignment to the right shows the sequences of the PhrS-creg region that interacts with its known target within the 5′-LR of pqsR mRNA compared with its potential target within the 5′-LR of the dmpR mRNA.

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130 A. Madhushani et al. gene, as has been found for uORFs of the regulatory fur gene, the magnesium transporter gene mgtA, and the quinolone gene pqsR (Vecerek et al., 2007; Park et al., 2010; Sonnleitner et al., 2011). The second is a sequence with high similarity to the target site of the sRNA PhrS that modulates translational coupling of the uORF and pqsR (Sonnleitner et al., 2011). Our current dissection of the role of Crc in controlling DmpR levels should greatly facilitate future work to unravel the significance of these additional potential regulatory features of the 5′-LR. Experimental procedures Bacterial strains and growth conditions Bacterial strains are listed in Supporting Information Table S1. All P. putida strains used are based on the genome sequenced KT2440 strain (Nelson et al., 2002). These include its Crc-null derivative in which the crc gene is interrupted by a gentamicin resistance cassette (KT2440crc::Gm; Aranda-Olmedo et al., 2005), a Hfq-null complete gene deletion derivative that lacks any resistance marker (KT2440-Δhfq; generously provided by V. de Lorenzo), and a double Hfq/Crc-null derivative (KT2440-Δhfq -crc::Gm) that was generated from the Hfq-null strain using the gentamicin insertion allele of crc carried on the pCRC10-Gm (Moreno et al., 2007). PP980 (KT2440-Po-luxAB) carries a mono-copy Po-luxAB reporter fusion inserted into its chromosome via mini-Tn5-Km2 (Pavel et al., 1994), while PP1423 (KT2440dmpR-Tel) expresses DmpR from its native promoter from the host chromosome by virtue of an insertion introduced via mini-Tn5-Tel (Sze et al., 2002). E. coli and P. putida strains were cultured at 37°C and 30°C, respectively, in LB/Lennox medium (AppliChem GmbH). Where needed, cultures were amended with carbenicillin (Cb, 100 μg ml−1 for E. coli or 1000 μg ml−1 for P. putida) to select for the resident plasmid.

Promoter-luxAB transcriptional reporter plasmids Plasmids (Supporting Information Table S2) were constructed by standard DNA techniques with the fidelity of linker- or PCRderived DNA fragments confirmed by DNA sequencing. All promoter-luxAB transcriptional reporter plasmids are based on the broad-host-range promoter probe vector pVI928 (Johansson et al., 2008) that carries the promoter-less luxAB genes of Vibrio harveyi, downstream of a poly-cloning site. Construction of the Pr-luxAB reporter plasmids with modified 5′-leader regions and other transcriptional reporter plasmids with a variety of promoters controlling transcription of the luxAB genes are detailed in Supporting Information Appendix S1.

Pr-dmpR expression plasmids (pVI2341 to pVI2343, Supporting Information Table S2): NotI sites flanking the -555 to +1842 Pr-dmpR fragments of plasmids harbouring modifications of the 5′-LR (pVI2334– pVI2336) were used for introduction of expression cassettes into the broad-host range plasmid pVI398 (Pavel et al., 1994). DNA fragments were inserted in the same orientation

as in pVI401, which carries the equivalent native Pr-dmpR fragment (Sze et al., 1996).

Generation of P. putida KT2440 mutant strains A double CrcZ/CrcY-null derivative of P. putida KT2240 was generated by double site recombination essentially as described in Moreno et al., 2012. First, the gentamicin gene replacement allele of crcY carried on the pKNG101-based plasmid pKNΔcrcY-Gm (Supporting Information Table S2) was introduced into P. putida KT2440 by conjugation from the replication-permissive E. coli strain S17-1 λpir. First-site recombinants were selected on rich media containing gentamycin. Second-site recombinants were then enriched for by growth in M9-minimal medium containing 10 mM benzoate as carbon source and 10% sucrose for counterselection of the pKNG101 encoded sacB gene (Kaniga et al., 1991). Fidelity of the recombination event of the resulting strain (denoted PP3362; KT2440-ΔcrcY::Gm) was verified by screening for loss of pKNG101 encoded streptomycin resistance phenotype, and by diagnostic PCR using primers that flank the site of insertion in crcY gene. The same strategy was then used to introduce the tetracycline gene replacement allele of crcZ carried on the pKNG101-based plasmid pKNΔcrcZ-Tc (Supporting Information Table S2) into KT2440ΔcrcY::Gm, resulting in the strain PP3386 (KT2440ΔcrcY::Gm/ΔcrcZ::Tc).

Luciferase activity assays Quantitative luciferase assays were performed on cultures grown and assayed at 30°C as previously described (Sze and Shingler, 1999). To ensure balanced growth, overnight cultures were diluted 1:50 and grown into exponential phase before a second dilution to an OD600 of 0.05–0.08 and initiation of the experiment. Light emission from 100 μl of whole cells using a 1:2000 dilution of decanal (luciferase substrate) was measured using an Infinite M200 (Tecan) luminometer. Specific activity is expressed as relative luciferase units per OD600 of 1.0. For simple screening of promoter activity, individual colonies of P. putida KT2440 harbouring different transcriptional reporter plasmids were streaked on solid medium. After overnight growth at 30°C, 100 μl of 1:1 diluted decanal was added to the lid of inverted plates and light emission documented using X-ray film.

Western analysis Samples from luciferase assay cultures were collected during the exponential (OD600 of ∼ 0.2–0.3) and stationary (OD600 of 2.0–3.0) phases of growth. Crude extract preparation, separation of proteins by SDS-PAGE, and subsequent electrotransfer to PVDF membrane (Amersham Biosciences) were as previously described (Shingler and Pavel, 1995). DmpR was detected using affinity purified polyclonal rabbit antibodies raised against the N-terminal 232 residues of DmpR (Bernardo et al., 2006) and revealed using ECL-Prime reagents (GE Healthcare) and X-ray film.

Quantitative PCR and mRNA stability assays P. putida KT2440 harbouring luxAB transcriptional reporter plasmids pVI938 (+5′-LR) or pVI2310 (-5′-LR) were grown

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 119–133

Control through the 5′-leader region of the dmpR gene as described for luciferase assays until early stationary phase (OD600 of ∼ 3.0). New transcription was then prevented by the addition of rifampicin to a final concentration of 200 μg ml−1. Cells from 20 ml rapidly chilled cultures were harvested at the indicated times. Total RNA from collected cell pellets was isolated using TRI Reagent® and RNAqueous® kit (Ambion) as recommended by the manufacturer. RNA preparations were treated with 20 units of RNase-free DNaseI (Ambion) at 37°C for 30 min and then clarified by phenol/chloroform extraction. Agarose gel electrophoresis was used to confirm the integrity of the final RNA preparations. Transcript levels were quantified by reverse transcriptase quantitative PCR (RT-qPCR) performed using the BioRad iScriptTM One-Step SYBR Green RT-PCR Kit and iCycler as recommended by the manufacturer. Triplicate 25 μl reactions contained 5 ng RNA and 7.5 pmols of primers (Supporting Information Table S3). Pr-derived transcripts from transcriptional reporter plasmids were quantified using primers that amplify a 200 bp luxABspecific fragment (Tm = 76.8°C) from the cDNA generated using random hexanucleotides. Internal standards for normalization were provided by equivalent triplicate reactions containing primers that amplify a 166 bp specific fragment (Tm = 76.8°C) from cDNA generated from transcripts of the rpoN (σ54) gene. Data are the average of triplicate determinations in each of two independent experiments.

RNA electro-mobility shift assays P. putida Hfq-His (wild-type and mutant derivatives) and wildtype Crc proteins were expressed and purified as detailed in Supporting Information Appendix S1. EMSA analysis of binding of these proteins to RNA probes was essentially as previously described (Moreno et al., 2009b). Reactions (20 μl in 10 mM Hepes-KOH pH 7.9, 35 mM KCl, 2 mM MgCl2) contained 1 μg yeast tRNA, 10 nM 5′-FAM-labelled RNA probe (DNA Technology, Denmark) and the indicated concentrations of purified proteins. Reactions were incubated for 40 min at 20°C prior to addition of 4 μl of 40% sucrose and analysis on a non-denaturing 6% polyacrylamide gel containing TBE buffer (45 mM Tris-HCl, pH 8.3, 43 mM boric acid, 2 mM MgCl2, 5% glycerol). Electrophoresis was performed in TBE buffer at 4°C and the results documented using LAS 4000 ImageQuant (GE Healthcare).

Acknowledgements We thank Eleonore Skärfstad and Master’s students (Ana R. Maceiras de Oliveira and Sarp Bamyaci) for their aid in some of the experimental work .We are indebted to V. de Lorenzo for the P. putida Hfq-null strain and B. E. Uhlin for the E. coli MG1655 Hfq-null strain. This work was supported by The Swedish Research Council (Grant Number 6212011-4791), Carl Trygger’s Foundation for Scientific Research (Grant Number CTS-11-420) to VS, the European Molecular Biology Organization through a Long-Term Research Fellowship (Grant Number 540-2009 to T. d. P.-S.), the J. C. Kempe Foundation (to AM) and the Spanish Ministry of Economy and Competitiveness (Grant BFU201232797 to FR).

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Control through the 5′-leader region of the dmpR gene Sze, C.C., Moore, T., and Shingler, V. (1996) Growth phasedependent transcription of the σ54-dependent Po promoter controlling the Pseudomonas-derived (methyl)phenol dmp operon of pVI150. J Bacteriol 178: 3727–3735. Sze, C.C., Bernardo, L.M., and Shingler, V. (2002) Integration of global regulation of two aromatic-responsive σ54dependent systems: a common phenotype by different mechanisms. J Bacteriol 184: 760–770. Valderrama, J.A., Shingler, V., Carmona, M., and Diaz, E. (2014) AccR is a master regulator involved in carbon catabolite repression of the anaerobic catabolism of aromatic compounds in Azoarcus sp. CIB. J Biol Chem 289: 1892–1904. doi:10.1074/jbc.M113.517714. Vecerek, B., Moll, I., and Bläsi, U. (2007) Control of Fur synthesis by the non-coding RNA RyhB and ironresponsive decoding. EMBO J 26: 965–975. Wikström, P., O’Neill, E., Ng, L.C., and Shingler, V. (2001) The regulatory N-terminal region of the aromatic-responsive transcriptional activator DmpR constrains nucleotidetriggered multimerisation. J Mol Biol 314: 971–984.

Supporting information Additional Supporting Information may be found in the online version of this article at the publisher’s web-site: Fig. S1. The 5′-LR DNA of dmpR inhibits transcription from promoters dependent on alternative σ-factors. A. Schematic illustration of transcriptional reporter plasmids with (+) and without (-) DNA encoding the 5′-LR of dmpR. The oblique line indicates the presence of a non-native BglII site upstream of the 5′-LR DNA. Sequences to the right show the promoter regions controlling transcription of the luxAB genes in the indicated reporters. Blue inverted arrow indicates the location of the UAS1 and UAS2 sites for DmpR binding. B. Representative images of luciferase plate test assays employing P. putida harbouring the indicated reporter plasmids as listed in panel A. P. putida KT2440 was used to assay activities of all but the σ54-dependent Po promoter, which was assayed using PP1423 (KT2440-dmpR-Tel) that supplies its obligatory activator (DmpR) from the chromosome. In the latter case, plates were additionally supple-

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mented with 2 mM of 2-methylphenol – the most potent effector of DmpR activity. Note that exposure time varied depending on the strength of the promoter under scrutiny, with short exposure time leading to non-detection of Pr activity as in the far right-hand autoradiogram. Fig. S2. RNA complex formation with Hfq-His and Crc/HfqHis requires an intact AAAAAUAA RNA motif. A–D. EMSA analyses as described under Fig. 6, except using a FAM-labelled 26 nt probe harbouring the 4C substitution. Note that in contrast to the data in Fig. 6 using the wild-type (WT) RNA probe, no complex is formed with this mutant RNA probe either with high levels of Hfq-His alone or in the co-presence of low levels of Crc and Hfq-His. E. EMSA analyses as described under Fig. 6, in which complex formation of the FAM-labelled 26 nt wild-type probe in the presence of 0.5 μM Crc and 3 μM Hfq is specifically competed by unlabelled wild-type probe (0, 25, 50 or 100 fold excess), but not by probes harbouring either the 4C substitution or a CUCU substitution. First lane, control for free FAM-labelled WT probe. Fig. S3. Alignment of Hfq. Sequences of P. putida KT2440 (PP), P. aeruginosa PA01 Hfq (PA) and E. coli MG1655 (EC) Hfq are shown aligned with differences highlighted in yellow. The conserved residues substituted in the P. putida Hfq-His mutant proteins Y25D and K56A are shown in red, while the four His residues of E. coli Hfq are shown in blue. Fig. S4. Marked conservation of DNA encoding the 5′-LR among (methyl)phenol degradative systems. The upper schematic illustrates the pVI150-encoded dmp-system of CF600 (as in Fig. 1) with the locations of four sub-regions used in blast searches highlighted. The table shows the percent identity found in highly related chromosomally encoded (P. putida BH, P. putida KCTC1452 and P. putida 35X) and plasmids-encoded (pPGH1 of P. putida H) (methyl)phenol degradative systems. Note that the 5′-LR DNA is more highly conserved than other intergenic DNA or DNA encoding cognate regulators of DmpK counterparts. Table S1. Bacterial strains used in this study. Table S2. Plasmids used in this study. Table S3. PCR primers used in this study. Table S4. Double-stranded DNA linkers used in this study. Appendix S1. Supporting material.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 119–133

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Transcriptional and translational control through the 5'-leader region of the dmpR master regulatory gene of phenol metabolism.

Expression of pathways for dissimilation of toxic aromatic compounds such as (methyl)phenols interfaces both stress-response and carbon catabolite rep...
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