A GntR family transcriptional regulator (PigT) controls gluconate-mediated repression and defines a new, independent pathway for regulation of the tripyrrole antibiotic, prodigiosin, in Serratia

Peter C. Fineran, Lee Everson, Holly Slater and George P. C. Salmond

Department of Biochemistry, University of Cambridge, Cambridge CB2 1QW, UK

Correspondence
George P. C. Salmond
gpcs{at}mole.bio.cam.ac.uk


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Biosynthesis of the red, tripyrrole antibiotic prodigiosin (Pig) by Serratia sp. ATCC 39006 (39006) is controlled by a complex regulatory network involving an N-acyl homoserine lactone (N-AHL) quorum-sensing system, at least two separate two-component signal transduction systems and a multitude of other regulators. In this study, a new transcriptional activator, PigT, and a physiological cue (gluconate), which are involved in an independent pathway controlling Pig biosynthesis, have been characterized. PigT, a GntR homologue, activates transcription of the pigA–O biosynthetic operon in the absence of gluconate. However, addition of gluconate to the growth medium of 39006 repressed transcription of pigA–O, via a PigT-dependent mechanism, resulting in a decrease in Pig production. Finally, expression of the pigT transcript was shown to be maximal in exponential phase, preceding the onset of Pig production. This work expands our understanding of both the physiological and genetic factors that impinge on the biosynthesis of the secondary metabolite Pig in 39006.


Abbreviations: 39006, Serratia sp. ATCC 39006; BHL, N-butanoyl-L-homoserine lactone; Car, carbapenem; CRP, cAMP receptor protein; HHL, N-hexanoyl-L-homoserine lactone; HTH, helix–turn–helix; N-AHL, N-acyl homoserine lactone; Pig, prodigiosin; QS, quorum sensing; RBS, ribosome-binding site; WT, wild-type

The GenBank/EMBL/DDBJ accession number for the pigT ORF sequence reported in this paper is AJ973142.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Serratia sp. ATCC 39006 (39006) is a taxonomically ill-defined member of the Enterobacteriaceae that produces a number of interesting secondary metabolites. Like many strains of Serratia marcescens, 39006 displays swimming motility and produces the red, tripyrrole antibiotic prodigiosin (Pig; 2-methyl-3-pentyl-6-methoxyprodigiosin), a molecule which also displays anticancer and immunosuppressant activities (Manderville, 2001; Perez-Tomas et al., 2003). The biosynthetic pathway of Pig has recently been elucidated in this laboratory (Harris et al., 2004; Williamson et al., 2005). In addition to Pig, 39006 also synthesizes a broad-spectrum {beta}-lactam antibiotic, carbapenem (Car; 1-carbapen-2-em-3-carboxylic acid), which is also synthesized by some strains of Erwinia carotovora ssp. carotovora and Photorhabdus luminescens ssp. laumondii TT01 (reviewed by Coulthurst et al., 2005). 39006 also shares with Erwinia species the ability to macerate plant cell walls via production of secreted pectate lyases and cellulases (Crow, 2001; Slater et al., 2003). Furthermore, recent studies have demonstrated that 39006 is virulent in a Caenorhabditis elegans infection model (Coulthurst et al., 2004).

Previous research in this laboratory has demonstrated numerous regulatory mechanisms, both genetic and environmental, involved in the control of secondary metabolite and (in some cases) exoenzyme production. Briefly, an N-acyl homoserine lactone (N-AHL) quorum-sensing (QS) system (SmaIR) functions, via a de-repression mechanism, to control Pig, Car and exoenzymes in response to increasing concentrations of N-butanoyl-L-homoserine lactone (BHL) and N-hexanoyl-L-homoserine lactone (HHL) synthesized by SmaI (Fineran et al., 2005; Slater et al., 2003; Thomson et al., 2000). QS controls the transcription of the Pig and Car biosynthetic genes by affecting expression of at least four other regulatory genes (carR, pigR, pigQ and rap) (Fineran et al., 2005; Slater et al., 2003).

Another key regulator is PigP, a novel putative DNA-binding protein, which controls expression of Pig and Car via modulation of at least seven other regulatory genes (carR, pigQ, pigR, pigS, pigV, pigX and rap) (Fineran et al., 2005). In addition, a GacAS family two-component system (PigQW) regulates Pig production via activation of the biosynthetic operon pigA–O (Fineran et al., 2005). We have also shown that Pig and Car levels are altered in response to phosphate concentration, sensed by the pstSCAB/phoBR system (Slater et al., 2003). Our current model of secondary metabolite production involves an intricate hierarchical network of transcriptional regulation that integrates numerous cues, including cell density, phosphate and unknown signal(s), presumably detected by the sensor kinase PigW (Fineran et al., 2005).

In the current study, we characterize a new physiological cue (gluconate) and a predicted sensor/effector (PigT) that regulate Pig production, in 39006, independently of the known regulators mentioned above. PigT is homologous to the Escherichia coli GntR protein, which is a repressor of the GntI group of genes required for gluconate utilization (Izu et al., 1997; Tong et al., 1996). We demonstrate that PigT activates transcription of the Pig biosynthetic operon (pigA–O), whereas addition of gluconate causes a reduction in transcription of pigA–O. Furthermore, a putative PigT binding site was identified in the promoter of pigA, based on sequence similarity to the gnt operator site (Porco et al., 1997). Therefore, PigT is predicted to activate pigA–O transcription directly. We demonstrate that PigT increases expression of a pigA promoter cloned in E. coli and that this activation is inhibited by gluconate. The transcription profile of a chromosomal pigT : : lacZ fusion and primer extension of pigT demonstrated that pigT expression was maximal in exponential phase and decreased in stationary phase. The gluconate/PigT system represents a new, independent pathway involved in regulation of Pig production.


   METHODS
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ABSTRACT
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METHODS
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Bacterial strains, plasmids, phage and culture conditions.
Bacterial strains and plasmids are listed in Table 1. Serratia sp. ATCC 39006 derivative strains were grown at 30 °C and E. coli strains were grown at 37 °C in Luria broth (LB; 5 g yeast extract l–1, 10 g Bacto tryptone l–1 and 5 g NaCl l–1) or minimal medium [0·1 %, w/v, (NH4)2SO4, 0·41 mM MgSO4, 0·2 %, w/v, glucose, 40 mM K2HPO4, 14·7 mM KH2PO4, pH 6·9–7·1] in shake flasks at 300 r.p.m., or on minimal or LB agar supplemented with 1·5 % (w/v) agar (LBA) (Miller, 1972). Bacterial growth (OD600) was measured in a Unicam He{lambda}ios spectrophotometer. When required, LB and minimal medium were supplemented with antibiotics at the following final concentrations: kanamycin (Km) 50 µg ml–1, spectinomycin (Sp) 50 µg ml–1, streptomycin (Sm) 50 µg ml–1, ampicillin (Ap) 100 µg ml–1 and tetracycline (Tc) 35 µg ml–1. The generalized transducing phage {pi}OT8 was used for transduction of chromosomal mutations, as described previously (Thomson et al., 2000).


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Table 1. Bacterial strains, bacteriophage and plasmids used in this study

 
DNA manipulations and sequencing of pigT.
All molecular biology techniques, unless stated otherwise, were performed by standard methods (Sambrook et al., 1989). Oligonucleotide primers were obtained from Sigma Genosys and are listed in Table 2. DNA sequencing was performed at the DNA Sequencing Facility, Department of Biochemistry, University of Cambridge on an Applied Biosystems 3730xl DNA Analyser. Nucleotide sequence data were analysed using GCG (Genetics Computer Group, University of Wisconsin) and compared with GenBank DNA or non-redundant protein sequence databases using BLAST (Altschul et al., 1997).


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Table 2. Oligonucleotide primers used in this study

 
Random transposon mutagenesis of Serratia sp. ATCC 39006 strain LacA was performed by conjugation with either E. coli S17-1 {lambda}pir harbouring plasmid pUTmini-Tn5lacZ1 or E. coli SM10 {lambda}pir harbouring plasmid pUTmini-Tn5Sm/Sp, as described previously (Thomson et al., 2000). Mutations were moved by {pi}OT8-mediated transduction into ‘clean’ LacA backgrounds to ensure that the phenotypes were associated with single transposon insertions. Genomic DNA from the pigT mutant (HSPIG36) was prepared and subjected to Southern blot and hybridization analysis using the pig cluster as a probe [pTON245 labelled with DIG using the Random Primed DNA Labelling Kit (Roche)]. The precise site of the transposon insertion in strain HSPIG36 was determined by ‘single primer site PCR’, followed by DNA sequence analysis across the transposon/chromosomal junction, as described previously (Fineran et al., 2005). Additionally, random primed PCR was used to complete the approximately 2 kb sequence containing pigT in a method similar to that described previously (Jacobs et al., 2003). Briefly, PCR was performed using a random primer mix (PF106, PF107 and PF108) and the pigT-specific primers HS3 (5' sequence) or gntR seq R (3' sequence) on LacA chromosomal DNA. A second PCR was performed using an aliquot of DNA from the first PCR with primers PF109 and either PF110 (5' sequence) or PF113 (3' sequence). The resulting mix of PCR fragments was purified and sequenced with PF110 or PF113, as appropriate. Sequencing was completed on the opposite strands using primers PF115 and PF116.

Construction of a plasmid (pTA38) that expresses native PigT.
A construct that enabled expression of native, untagged PigT was created as outlined below. The pigT gene was amplified by PCR, using primers PF103 and PF69, which contain EcoRI and PstI restriction sites, respectively. Additionally, primer PF103 contains a consensus ribosome-binding site (RBS, AGGAGGA). The PCR fragment of pigT was cloned into pQE-80L, previously digested with the enzymes EcoRI and PstI. The resulting plasmid, pTA38, was confirmed by DNA sequencing. Expression of plasmid pTA38 in both E. coli and 39006 hosts was induced with 1 mM IPTG.

Generation of a pigT : : mini-Tn5Sm/Sp mutant by transposon exchange mutagenesis.
Previously, we reported an efficient in vivo method based on phage transduction that allows exchange of a chromosomal transposon insertion for an alternative transposon (Fineran et al., 2005). Using this exchange method, strain HSPIG36S (pigT : : mini-Tn5Sm/Sp) was constructed from the parental strain HSPIG36 (pigT : : mini-Tn5lacZ1). The location of the new transposon was determined by PCR amplification across the insertion site using primers facing out of the transposon (LER1 and SP2 for mini-Tn5Sm/Sp) and the pigT-specific primers HS3 and HS4. PCR products in the expected size range were analysed further by sequencing and the point of insertion was determined. The pigT : : mini-Tn5Sm/Sp mutation was transduced into a ‘clean’ LacA background using {pi}OT8, and the nature of the transductants was confirmed by phenotype and PCR analysis.

Marker exchange mutagenesis of pigW.
Strain PIG62L (pigW : : lacZKm) was generated by a marker exchange strategy. Firstly, primers PF95 and PF96, containing XbaI and SalI restriction sites, respectively, were used to PCR-amplify a fragment of the pigW gene. This fragment was ligated into pBluescript II KS+digested with XbaI and SalI, and the resulting construct (pTA35) was confirmed by DNA sequencing. Secondly, a promoterless lacZ gene and a Km resistance cassette were amplified from plasmid pUTmini-Tn5lacZ1 using primers PF97 and PF98, which contain NcoI and NheI restriction sites, respectively. The lacZKm fragment was ligated to specific NcoI and NheI sites internal to the pigW gene in pTA35, generating plasmid pTA36. Next, the entire pigW : : lacZKm fragment was excised from plasmid pTA36 on the unique XbaI and SalI restriction sites and ligated into the marker exchange vector pKNG101 digested with XbaI and SalI, generating plasmid pTA37. Marker exchange with plasmid pTA37 was performed using a sucrose selection protocol similar to that described elsewhere (Kaniga et al., 1991). The pigW mutant was confirmed by PCR, sequencing and phenotypic analysis.

Construction of pigA promoter : : lacZ fusions and assay conditions.
The pigA promoter was delineated into four different size fragments and cloned into the promoterless lacZ plasmid pRW50 (Lodge et al., 1992). The plasmids, named pTA15, pTA30, pTA31 and pTA32 in order of decreasing size, were created by cloning BamHI/HindIII-digested PCR products [generated with forward primers HS78, PF70, PF71 and PF72, respectively, and the reverse primer HS79 (Table 2, Fig. 3)] into BamHI/HindIII-digested pRW50. The nature of all plasmids was confirmed by DNA sequencing.



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Fig. 3. In E. coli, the pigA promoter is activated by PigT and contains a predicted palindromic PigT binding site. (a) Partial sequence of the pigA promoter that was cloned into contructs pTA15, pTA30, pTA31 and pTA32 (indicated). Features of the promoter include the ROP1 (repressor of pigment) region (bold italics), the transcriptional start site (+1), the predicted –10 and –35 elements (boxed) (Slater et al., 2003), and the proposed PigT binding site (bold boxed, centred at –78·5) based on the GntR binding site (Porco et al., 1997). A putative cAMP receptor protein (CRP) binding site centred at –64·5 relative to the +1 overlaps the PigT box and is shown underlined with the two conserved half sites in italics. Primer binding sites are shown underlined and the translational initiation codon of pigA is shown in bold (ATG). (b) Schematic representation of the orfY–pigA intergenic region and the pRW50-derived pigA promoter fusion constructs (pTA15, pTA30, pTA31 and pTA32). The scale of the promoter region is indicated, but genes are not drawn to scale. {beta}-Galactosidase activity of the pigA promoter fusions in E. coli YU563 in the absence (pQE-80L, –) or presence (pTA38, +) of PigT is indicated. Data shown are the mean±SD of at least three independent experiments.

 
Promoter activity assays were performed in an E. coli gntR mutant (YU563) to avoid interference by native GntR in E. coli. YU563 was transformed with the pRW50-derived construct (pTA15, 30, 31 or 32) and either the vector control (pQE-80L) or the PigT expression plasmid (pTA38). Additionally, E. coli strains containing pRW50 had no detectable {beta}-galactosidase activity. Overnight cultures of strains tested were grown with Ap and Tc selection for approximately 16 h and subcultured into 25 ml LB containing Ap, Tc and IPTG in 250 ml conical flasks to a starting OD600 of 0·04. The bacterial cultures were incubated at 300 r.p.m. at 37 °C for approximately 3 h, or until the OD600 reached between 0·4 and 0·6. Aliquots (1 ml) of culture were removed, pelleted by centrifugation and subjected to {beta}-galactosidase assays, as outlined below.

{beta}-Galactosidase assays.
{beta}-Galactosidase activity in bacterial cultures grown in liquid media was determined using ONPG as substrate, as described elsewhere (Miller, 1972). Enzyme activity was expressed as the initial reaction rate per ml of sample per OD600 of the bacterial culture tested ({Delta}A420 min–1 ml–1 OD600–1). Results presented are the mean±standard deviation (SD) of three independent experiments, unless stated otherwise.

Bioassays of prodigiosin, carbapenem, N-AHLs, exoenzymes and motility.
The assays for Pig and Car were performed as described previously (Slater et al., 2003). Pig production was plotted as (A534 ml–1 OD600–1)x50. Detection of N-AHLs was performed using the Serratia LIS bioassay described in Thomson et al. (2000). Activity of pectate lyase and cellulase was analysed on agar plates containing the corresponding substrates, as described elsewhere (Andro et al., 1984). Motility was assessed on tryptone swarm agar (TSA) plates (10 g Bacto tryptone l–1, 5 g NaCl l–1 and 3 g agar l–1). Overnight bacterial cultures were adjusted to an OD600 of 0·2, 3 µl was spotted onto the plates, and halo size was examined after growth for 16 h at 30 °C.

Primer extension and RNA studies.
A hot-acidic phenol method (Aiba et al., 1981) was used to extract total RNA from 39006. Primer extension analysis for the pigA transcript was performed as described previously (Slater et al., 2003) using primer HS34. All primer extension reactions were performed with 25 µg total RNA and 0·2 pmol of the appropriate 32P-labelled primer. Primer extension analysis of pigT was performed using primer gntR SS(2), which anneals 122 bp downstream of the predicted pigT translational start site. mRNA transcripts were quantified using ImageJ, a densitometry program available at http://rsb.info.nih.gov/nih-image/Default.html.


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
A GntR homologue, PigT, regulates Pig production and swimming motility
From a mutagenesis programme aimed at identifying regulators of secondary metabolite production, one mutant (HSPIG36) was analysed in more detail because it showed a strong defect in Pig production (compare Fig. 1a with Fig. 1b). The impact of the mutation in HSPIG36 on other phenotypes (swimming motility, Pel, Cel, Car and N-AHL production) was assessed in plate assays from stationary-phase overnight cultures. Mutant HSPIG36 showed reduced swimming motility compared to the ‘wild-type’ (WT refers to the parental precursor strain LacA throughout this paper) (compare Fig. 1c with Fig. 1d). There was no detectable difference between the WT and mutant HSPIG36 in the exoenzyme assays or in N-AHL production (data not shown).



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Fig. 1. Mutation of pigT or addition of gluconate to 39006 culture media affects Pig production in 39006. Pig production by strains (a) LacA and (b) HSPIG36 (pigT) grown on LB agar supplemented with 1·5 % (w/v) agar (LBA). Swimming motility of strains (c) LacA and (d) HSPIG36 (pigT) grown on tryptone swarm agar (TSA). Complementation of Pig production in (e) HSPIG36 (pigT) carrying plasmid pTA38 (PigT) compared to (f) HSPIG36 (pigT) with the vector control (pQE-80L). Pig production by strain LacA on minimal medium with either 0·2 % (w/v) glucose (g) or 0·2 % (w/v) gluconate(h) as the sole carbon source. Pigproduction phenotype of strain LacA grown on 0·2 % glucose minimal medium (i) or on 0·2 % glucose minimal medium supplemented with 0·02 % gluconate (j). Production of Pig by strain LacA on (k) LBA or (l) LBA containing 0·02 % gluconate.

 
The site of the mini-Tn5lacZ1 insertion in HSPIG36 was determined as described in Methods. The transposon interrupted a 999 bp ORF with homology to gntR from E. coli. We designated this gene in 39006 as pigT. The predicted PigT protein is similar (77 % identity/81 % similarity) to GntR from E. coli K-12 (Tong et al., 1996), a member of the GalR/LacI family of transcriptional regulators (Fukami-Kobayashi et al., 2003; Peekhaus & Conway, 1998). In E. coli, in the absence of gluconate, GntR binds DNA via an N-terminal helix–turn–helix (HTH) domain and represses the GntI group of genes involved in gluconate utilization (Peekhaus & Conway, 1998). The location and sequence of the gene 5' of pigT is similar (67 % identity/73 % similarity of 45 amino acids aligned) to YhhW from E. coli K-12. Furthermore, an ORF 3' of pigT is similar (57 % identity/67 % similarity) to GntK from E. coli K-12. These results indicate that pigT shares a similar, although not identical, genomic context to gntR of E. coli K-12.

To demonstrate that the transposon disruption in pigT was responsible for the Pig phenotype, complementation was performed. Plasmid pTA38, which expresses native PigT, was able to complement the Pig phenotype in HSPIG36, whereas a pQE-80L control could not (compare Fig. 1e with Fig. 1f). Therefore, PigT affects both swimming motility and Pig production in 39006 and shares sequence similarity with GntR from E. coli.

PigT is a transcriptional regulator of Pig
To examine the impact of PigT on Pig production in more detail, Pig was assessed throughout growth in LB in the pigT mutant and compared to that of the WT (Fig. 2a). Pig production was approximately 10 % of the levels observed in the WT. To enable construction of double chromosomal mutants, strain HSPIG36S (pigT : : mini-Tn5Sm/Sp) was constructed (see Methods). HSPIG36S was shown to have the same swimming motility and Pig production defects as HSPIG36. Expression of a chromosomal pigA : : lacZ fusion was studied throughout growth in the WT and pigT mutant backgrounds (Fig. 2b), and was found to be reduced to approximately 30 % of WT levels. In addition, primer extension studies demonstrated that the pigA–O transcript in the pigT mutant was similarly reduced compared to that of the WT (Fig. 2c). The pigT mutant was unaffected for Car and BHL/HHL production when examined throughout growth in LB (data not shown). These results were consistent with a model in which PigT regulates Pig production by controlling the transcription of the biosynthetic genes (pigA–O).



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Fig. 2. PigT regulates Pig production by controlling transcription of the Pig biosynthetic operon pigA–O. (a) Pig levels in WT (squares) and HSPIG36 (pigT) (triangles) throughout growth in LB. (b) {beta}-Galactosidase activity was measured from a chromosomal pigA : : lacZ fusion in an otherwise WT background (squares) or in a strain containing a mini-Tn5Sm/Sp chromosomal insertion in pigT (triangles). (c) Primer extension analysis of the pigA–O transcript from WT and HSPIG36 (pigT) throughout growth in LB. The pigA transcript was quantified and the pigT/WT ratios are given below the lanes. In (a)and (b), solid symbols and lines represent Pig and {beta}-galactosidase activity, respectively, whereas the open symbols and dashed lines represent the growth curves of the corresponding strains. Data shown are the mean±standard deviation (SD) of at least three independent experiments.

 
PigT does not operate via known regulators
In recent reports, we elucidated a complex circuit controlling Pig production that involved numerous transcriptional regulators, including the SmaIR QS system (Fineran et al., 2005; Slater et al., 2003; Thomson et al., 1997, 2000). Given this plethora of known regulatory inputs, it was possible that PigT was controlling pigA expression indirectly via one or more of these other known secondary metabolite regulators. Therefore, the effect of a chromosomal mini-Tn5Sm/Sp mutation in pigT on the expression of lacZ transcriptional fusions in pigP, pigQ, pigR, pigS, pigV, pigW, pigX, rap and smaI was examined in early-exponential, mid-exponential, late-exponential/transition and stationary phase. However, pigT did not affect {beta}-galactosidase activity from any of these fusions (data not shown). Furthermore, chromosomal mini-Tn5Sm/Sp mutations in smaI, pigP, pigQ, pigW, pigX, pstS and rap did not alter expression of the pigT : : mini-Tn5lacZ1 fusion in early-exponential, mid-exponential, late-exponential/transition and stationary phase (data not shown). Therefore, PigT defines a new prodigiosin regulator operating via a novel route, independent of the members of the regulatory network examined in this study. PigT could be activating pigA–O transcription directly, or could be repressing expression of an, as yet, unidentified repressor of Pig.

PigT is predicted to act directly at the pigA promoter
The transcriptional hierarchy data described above indicate that pigT does not control the transcription of the other regulators examined, consistent with the notion that PigT directly activates pigA–O transcription. Examination of the sequence 5' of pigA–O revealed a potential PigT binding site (ATGTTACTGGTAACTG) centred at –78·5 relative to the pigA transcriptional start site (Fig. 3a) (Slater et al., 2003). The bold type represents the two conserved half sites of the predicted PigT/GntR binding sites. This predicted binding site is based on the E. coli GntR palindromic consensus binding sequence (ATGTTA-(N4, GC-rich)-TAACAT) (Porco et al., 1997) present in the promoter regions of GntR-regulated genes. The position of this binding site suggests that PigT may be binding directly to the pigA–O promoter region and functioning as a Class I activator to regulate transcription (Busby & Ebright, 1999).

To test if PigT directly activates transcription of the Pig biosynthetic operon, the pigA promoter was cloned into the low-copy, promoterless lacZ expression vector pRW50 (Lodge et al., 1992). Promoter delineation studies were performed on the pigA promoter to determine regions required for PigT activation and promoter activity (see Methods). The results showed that, in an E. coli gntR mutant (YU563), the presence of PigT caused an increase in transcription from the pigA promoter (Fig. 3b). Furthermore, the shortest construct (pTA32), which still possessed the –35 and –10 elements, but not the predicted PigT binding site, was not activated by PigT. These results suggest that PigT directly activates transcription of pigA–O and this activation requires a 150 bp region 5' of the pigA promoter. Within this region there is an inverted repeat sequence similar to the E. coli GntR box, which we predict is a PigT box.

We had reported previously that transposon insertions within the ROP1 (repressor of pigment) region of the pigA promoter in 39006 resulted in a hyper-pigmented phenotype and an increase in pigA : : lacZ expression (Slater et al., 2003). It was suggested that the ROP1 region might (1) be a repressor binding site, (2) be important for DNA conformation or (3) encode a small RNA that acts negatively on Pig production. Deletion of the ROP1 region had little effect on promoter activity in E. coli (Fig. 3b). Furthermore, PigT was still able to activate pigA promoter constructs that lacked the ROP1 region, indicating that ROP1 is not required for the PigT-mediated activation of pigA. Thus, the DNA region between –193 and –44 inclusive is required for PigT-mediated activation of pigA transcription.

Expression of pigT is maximal in exponential phase
To assess the expression of pigT, {beta}-galactosidase activity was measured from a chromosomal pigT : : lacZ fusion throughout growth (Fig. 4a). In addition, primer extension analysis of the pigT transcript was performed (Fig. 4b). These experiments showed that levels of the pigT transcript peak in exponential phase (6 h) and then decline as the cultures reach stationary phase (12 h).



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Fig. 4. Expression of pigT is maximal in the exponential growth phase and decreases in stationary phase. (a) {beta}-Galactosidase activity was measured from pigT : : lacZ (HSPIG36) throughout growth in LB. (b) Primer extension analysis of the pigT transcript from a WT culture throughout growth in LB. In (a), bars represent {beta}-galactosidase activity, whereas the open symbols and dashed lines represent the growth curves of the corresponding strain. Data shown are the mean±SD of at least three independent experiments.

 
Gluconate regulates Pig production transcriptionally
The identification of a putative GntR homologue (PigT) that controls Pig encouraged us to examine whether gluconate affects pigmentation. 39006 grown on minimal medium with 0·2 % (w/v) gluconate as sole carbon source was non-pigmented when compared with the 0·2 % (w/v) glucose-grown controls (compare Fig. 1g with Fig. 1h). Addition of 0·02 % gluconate to minimal medium containing 0·2 % glucose caused a reduction in Pig (compare Fig. 1i with Fig. 1j). Furthermore, when LBA was supplemented with 0·02 % gluconate, Pig production was decreased (compare Fig. 1k with Fig. 1l). To examine further the gluconate-mediated repression of Pig, 0·02 % gluconate was added to glucose minimal medium, either at the start of growth (t0) or after 8 h (t8), and Pig was examined throughout growth (Fig. 5a). Gluconate repressed Pig to approximately 35 % of WT levels when added at t0. Gluconate added just prior to the onset of pigmentation (t8) decreased Pig to approximately 50 % of WT levels, after it had initially been produced (Fig. 5a). The level of pigA–O transcription was measured using a strain with a chromosomal pigA : : lacZ reporter (MCP2L) under identical conditions to those described above. Gluconate added at t0 and t8 resulted in approximately 12 and 35 %, respectively, of pigA–O transcription when measured by {beta}-galactosidase activity (Fig. 5b). Car and BHL/HHL levels were unaffected throughout growth in LB by the addition of 0·02 % gluconate (data not shown). These experiments showed that gluconate, even in the presence of glucose, could repress Pig biosynthesis, and that this physiological repression operates, at least in part, at the level of transcription of the pigA–O operon.



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Fig. 5. Addition of gluconate inhibits Pig production and transcription of the Pig biosynthetic genes, pigA–O. (a) Pig production throughout growth in glucose minimal medium by the WT after addition of 0·02 % (w/v) gluconate at the start of growth (t0, triangles), after 8 h (t8, diamonds) or with no added gluconate (squares). (b) {beta}-Galactosidase activity was measured from a chromosomal pigA : : lacZ fusion throughout growth in glucose minimal medium after addition of 0·02 % gluconate at the start of growth (t0, triangles), after 8 h (t8, diamonds) or with no added gluconate (squares). Solid symbols and lines represent either Pig or {beta}-galactosidase activity, whereas the open symbols and dashed lines represent the growth curves of the corresponding strains. Data shown are the mean±SD of at least three independent experiments.

 
Gluconate-mediated repression of Pig requires PigT
In E. coli, the GntR protein responds to gluconate and alters gene expression accordingly. Since the phenotypic consequences of a pigT mutation and addition of gluconate were similar, it was predicted that the gluconate response required functional PigT. Therefore, an epistasis test was performed by growing the WT and pigT mutant in LB with or without gluconate (0·02 %), and Pig was measured in stationary phase. The presence of gluconate resulted in a 50 % reduction in Pig in the WT strain (Fig. 6a). No additional decrease in Pig was observed in the pigT mutant in the presence of gluconate, implying that PigT was required for the gluconate-mediated repression (Fig. 6b). Furthermore, PigT-mediated activation of a pigA promoter lacZ fusion in an E. coli gntR mutant was inhibited by 0·02 % gluconate (Fig. 6c). Addition of gluconate, in the absence of PigT, had no effect on the expression of the pigA promoter in this E. coli background (Fig. 6c). The available data suggested that gluconate, by inhibiting the activity of PigT, was repressing Pig production.



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Fig. 6. PigT is required for the gluconate-mediated repression of Pig production. Pig production by (a) WT and (b) pigT mutant (HSPIG36) in stationary phase (12 h). Strains were grown in LB in the presence (open bars) or absence (black bars) of 0·02 % (w/v) gluconate added at the start of growth. Note that the scales of (a) and (b) are different. (c) {beta}-Galactosidase activity was measured from a pigA promoter lacZ fusion (plasmid pTA30) in an E. coli gntR mutant (YU563) containing either vector-only control (No PigT; pQE-80L) or a PigT expression plasmid (PigT; pTA38) in LB in the presence (open bars) or absence (black bars) of 0·02 % gluconate added at the start of growth. Data shown are the mean±SD of at least three independent experiments.

 

   DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
We have shown previously that in Serratia sp. ATCC 39006 (39006), production of prodigiosin (Pig) is regulated by a complex regulatory network involving an N-AHL QS system, PigP, Rap, GacAS and PhoBR two-component systems, and several other regulators (Fineran et al., 2005; Slater et al., 2003; Thomson et al., 1997, 2000) (Fig. 7). Furthermore, numerous studies have implicated a spectrum of physiological factors in the production of Pig in strains of S. marcescens (Cang et al., 2000; Giri et al., 2004; Rjazantseva et al., 1994; Silverman & Munoz, 1973; Sole et al., 1997; Williams et al., 1971a, b; Witney et al., 1977). Therefore, it is apparent that multiple environmental, physiological and genetic cues influence Pig production in Serratia species. In this paper, we have characterized the mechanism underlying gluconate-mediated repression of the biosynthesis of Pig: yet another regulatory input in the control of this red antibiotic in 39006.



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Fig. 7. Schematic model of physiological and genetic factors that affect Pig production in 39006. Gluconate ‘deactivates' PigT, which, in the absence of gluconate, acts via an independent pathway to activate transcription of pigA–O. Low phosphate results in activation of transcription of both smaI and pigA–O via the PstSCAB/PhoBR system (Slater et al., 2003). Pig production is also controlled by cell density via the SmaIR quorum-sensing (QS) system (Slater et al., 2003; Thomson etal., 2000). SmaIR also regulates transcription of pigQ of the PigQW (GacAS family) two-component system. Presumably, PigQ is activated in response to signals sensed by the histidine kinase PigW (Fineran et al., 2005). Transcription of rap (regulator of antibiotic and pigment), which is a member of the SlyA/MarR family, is also controlled by QS (Fineran et al., 2005). Both rap and pigQ are also activated by a master Pig regulator (PigP), which controls expression of numerous other Pig and Car regulators (not shown). A deeper discussion of all these regulatory inputs is described in Fineran et al. (2005).

 
Firstly, we demonstrated that mutation of a gene predicted to encode a GntR homologue (pigT) resulted in lower Pig production and a reduction in swimming motility (Figs 1 and 2). The reduction in Pig was the result of decreased transcription of the biosynthetic operon (pigA–O) observed both by chromosomal pigA : : lacZ fusion analysis and by primer extension of pigA–O transcript levels in the pigT mutant (Fig. 2). Furthermore, complementation of the pigT mutation with a plasmid encoding the pigT ORF restored Pig production (Fig. 1).

In E. coli, GntR negatively regulates the GntI group of genes involved in gluconate utilization. However, when gluconate is present in the medium, GntR is thought to bind gluconate (or one of its metabolic intermediates), and repression of the GntI genes is alleviated (Izu et al., 1997; Tong et al., 1996). The gluconate-sensing role of GntR, and the sequence similarity to PigT, prompted us to examine the effects of gluconate on secondary-metabolite biosynthesis in 39006.

Gluconate, even in the presence of glucose or in complex media (LB), could decrease Pig production in 39006 (Figs 1 and 5a). The gluconate-mediated decrease in Pig production was accompanied by a reduction in pigA transcription, even if gluconate was added after the onset of pigmentation (Fig. 5b).

These data suggested a model whereby PigT mediates gluconate repression of Pig production. Based on genetic and biochemical data from E. coli GntR, it is clear that GntR represses gene expression by binding to DNA in the promoter regions of GntI genes in the absence of gluconate (Peekhaus & Conway, 1998). However, in 39006, PigT-activated Pig production is hypothesized to be ‘deactivated’ by the addition of gluconate. It was possible that PigT was controlling transcription of pigA–O indirectly by repressing a repressor of Pig. Therefore, to try and identify a potential regulatory intermediate, the effects of a pigT mutation on the transcription of multiple known secondary metabolite regulators were examined. However, the effect of PigT on Pig production was independent of all the regulators investigated. PigT did not control transcription of any of those tested and, likewise, no regulator examined controlled expression of pigT. This result suggested that PigT might directly activate transcription of the pigA–O operon.

Interestingly, a putative PigT binding site, based on that demonstrated for GntR (Porco et al., 1997), was identified upstream of the –35 element in the pigA promoter (Fig. 3a). Furthermore, promoter delineation experiments in E. coli implied that PigT only activated pigA promoter constructs that contained a 150 bp region (from –193 to –44) which includes this putative PigT binding site (Fig. 3b). Based on this genetic evidence, our model, that PigT directly activates expression of pigA–O in the absence of gluconate, contradicts the conventional role of GntR, which functions as a repressor of gluconate metabolism genes in E. coli. However, in our model, the binding of PigT (like GntR) to DNA is dependent on the absence of gluconate. We suggest that the binding of gluconate to PigT results in PigT being unable to bind DNA, and therefore activation of transcription is prevented. Interestingly, we have evidence that, in E. coli, GntR can activate the cloned pigA promoter, and this activation can be inhibited by the presence of gluconate (P. Fineran, unpublished results). This suggests that, E. coli GntR can also be an activator of gene expression if the placement of the GntR binding site is permitting. It remains to be determined whether PigT functions analogously to GntR by regulating genes involved in uptake and utilization of gluconate. However, both the sequence context of pigT, with respect to genes potentially involved in gluconate utilization, and the presence of a putative PigT binding site in the promoter of the putative gluconokinase (data not shown), indicate that PigT may have a role in regulating gluconate metabolism.

The pigA promoter studies raised a few interesting points about the regulation of the pigA–O operon. Firstly, the transcriptional start site of pigA has been mapped by primer extension, and putative –10 and –35 elements identified (Slater et al., 2003). A TAAAGA->TAAAGG substitution in the predicted –10 element resulted in a greater than 50 % reduction in pigA promoter activity (data not shown), supporting this assignment. Secondly, interrogation of the sequence 5' of pigA has revealed a predicted CRP binding site centred at –64·5, which overlaps one half of the predicted PigT binding site (Fig. 3a). Preliminary data demonstrated an increase in pigA promoter expression in an E. coli crp mutant (P. Fineran, unpublished results). However, the role of CRP requires further investigation. Interestingly, the gntT promoter in E. coli is also regulated by both GntR and CRP (Peekhaus & Conway, 1998). Furthermore, it is interesting that, in a previous mutagenesis screen, we isolated a predicted novel adenylate cyclase mutant (pigR) with decreased Pig (Fineran et al., 2005).

Primer extension and expression of a chromosomal pigT : : lacZ fusion was used to assess the transcription profile of pigT throughout growth. pigT was maximally expressed during mid–late exponential phase, but was attenuated during the transition into stationary phase (Fig. 4a, b). This transcript profile is consistent with a role for PigT as a major activator of pigA, and hence Pig, levels of which increase dramatically in mid–late exponential phase (Fig. 2a, b).

Our model of PigT directly activating the pigA–O operon (and hence Pig production) and inhibition of this interaction in the presence of gluconate is based on the following lines of evidence: (1) PigT shares significant sequence similarity with E. coli GntR (77 % identity/81 % similarity), which has been shown biochemically to respond to gluconate and alter its DNA binding affinity accordingly (Peekhaus & Conway, 1998); (2) addition of gluconate or mutation of pigT had similar effects on Pig production and transcription of pigA–O (Figs 2 and 5), but no effect on Car and BHL/HHL synthesis; (3) epistasis experiments demonstrated that gluconate could not further repress pigmentation in a pigT mutant (Fig. 6); (4) finally, in an E. coli gntR mutant, PigT-mediated activation of the pigA promoter was inhibited by gluconate (Fig. 6).

The physiological benefit of 39006 repressing Pig in the presence of gluconate is unknown. Little is known of the true habitat(s) of 39006, but it is possible that gluconate is present in a niche where Pig is not required. GntR homologues have been associated with the regulation of various phenotypes in bacteria. For example, a GntR homologue regulates production of antibiotics and the onset of aerial mycelia and spore formation in Streptomyces coelicolor (Sprusansky et al., 2003).

The current investigation has increased our understanding of the physiological and genetic regulation of the biosynthesis of a tripyrrole antibiotic pigment, prodigiosin (Pig). We have identified a new physiological cue (gluconate) that represses Pig production and characterized the potential sensor of this signal (PigT). The gluconate/PigT pathway represents a system for control of pigmentation that is apparently completely independent of the regulatory network of secondary metabolite production that we have recently characterized in some detail (Fineran et al., 2005; Slater et al., 2003; Thomson et al., 1997, 2000). Fig. 7 highlights some of the important physiological and genetic factors that modulate Pig biosynthesis in 39006, including the PigT/gluconate system. Both the physiological rationale and molecular mechanism driving this gluconate-mediated regulation of the red, tripyrrole antibiotic warrant further investigation.


   ACKNOWLEDGEMENTS
 
The authors wish to thank all members of the Salmond and Welch groups, I. Foulds for excellent technical assistance, A. Barnard for critically reading the manuscript and T. Burr for useful discussions. We also would like to thank S. Busby for the generous gift of plasmid pRW50 and M. Yamada for E. coli strain YU563. This work was supported by the BBSRC, UK, and P. F. was supported by a Bright Futures Top Achiever Doctoral Scholarship from the Tertiary Education Commission of New Zealand.


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Received 3 June 2005; revised 21 September 2005; accepted 27 September 2005.



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