Differential Regulation of High-Affinity Phosphate Transport Systems of Mycobacterium smegmatis: Identification of PhnF, a Repressor of the phnDCE Operon

Bacterial strains and growth conditions.All strains and plasmids used in this study are listed in Table 1. E. coli strains were grown in Luria-Bertani (LB) medium at 37°C with agitation (200 rpm). The M. smegmatis mc²155 strain (32) and derived strains were routinely grown at 37°C with agitation at 200 rpm in LB medium containing 0.05% (wt/vol) Tween 80 (LBT) or in Middlebrook 7H9 medium (Difco) supplemented with 10% albumin-dextrose-catalase enrichment (ADC; Becton Dickinson) and 0.05% (wt/vol) Tween 80. M. smegmatis transformants were grown at 28°C for the propagation of temperature-sensitive vectors and at 40°C for allelic exchange mutagenesis. For P_i limitation studies, M. smegmatis was grown in modified minimal Sauton's medium. The composition of this medium per liter was 0.5 g MgSO₄, 2 g citric acid, 1 g l-asparagine, 0.3 g KCl, 1 g glycerol, 0.5 g Tween 80, 320 μl 0.5 M FeCl₃, and 100 μl of 1 M NH₄Cl. The high-phosphate medium contained 100 mM K₂HPO₄. For phosphate starvation experiments, cells were grown in high-phosphate medium to an optical density at 600 nm (OD₆₀₀) of 0.7 to 1.2, washed twice in sterile 0.85% (wt/vol) saline with 0.05% (wt/vol) Tween 80, resuspended to an OD₆₀₀ of 0.7 in phosphate-free Sauton's medium, and incubated at 37°C with agitation at 200 rpm for 2 h.

Selective media contained kanamycin (50 μg ml⁻¹ for E. coli; 20 μg ml⁻¹ for M. smegmatis), gentamicin (20 μg ml⁻¹ for E. coli; 5 μg ml⁻¹ for M. smegmatis), or hygromycin (200 μg ml⁻¹ for E. coli; 50 μg ml⁻¹ for M. smegmatis). Solid medium contained 1.5% agar.

OD₆₀₀ was measured using culture samples diluted in saline to bring the OD₆₀₀ to below 0.5 when measured in cuvettes of 1-cm length light path in a Jenway 6300 spectrophotometer.

DNA manipulation and cloning of constructs.All molecular biology techniques were carried out according to standard procedures (31). Restriction or DNA-modifying enzymes and other molecular biology reagents were obtained from Roche Diagnostics or New England Biolabs.

Genomic DNA of M. smegmatis was isolated by a method modified from that described by Gonzalez-y-Merchand et al. (7). In brief, cells grown on LBT agar were resuspended in 200 μl of lysis buffer (4 M guanidine thiocyanate, 1 mM β-mercaptoethanol, 10 mM EDTA, 0.1% [wt/vol] Tween 80), snap-frozen in dry ice-ethanol, and heated to 65°C for 10 min. Snap-freezing/heating was repeated, and the cells were cooled on ice for 5 min. The aqueous phase was extracted twice with chloroform, and DNA was precipitated by isopropanol. The pellet was washed once in 70% ethanol, air dried, and dissolved in deionized water.

To create a transcriptional fusion of phnF, a PCR product encompassing 420 bp of DNA upstream of phnF plus 110 bp of its coding region was amplified using the primers PphnFF (5′-AAATTTGGGCCCGCATAGTCGGAGGCCTGGACG-3′) and PphnFR (5′-AAATTTGGTACCGGATCCCCGATGCGCATACC-3′). The product was cloned into the ApaI and Asp718 sites of pJEM15 (34), creating the plasmid pSG18.

To create a construct for the deletion of phnF, the kanamycin resistance (Km^r) cassette, encoded by aphA-3, was amplified from pUC18K (19) by PCR, using the primers 3′mcspUC (5′-GTTTTCCCAGTCACGACGTT-3′) and 5′mcspUC (5′-CACACAGGAAACAGCTATGA-3′). The resulting 850-bp product was digested with EcoRI and BamHI. PCR products of approximately 850 bp flanking the phnF gene of M. smegmatis were amplified by using the primer phnFKO1 (5′-AAATTTACTAGTGGCCTCAGAACCCGACTTGA-3′) with primer phnFKO2 (5′-AAATTTGAATTCGGATCCCCGATGCGCATACC-3′) (left flank) and the primer phnFKO3 (5′-AAATTTGGATCCACGGCCGTCATGCACGCTAAG-3′) with primer phnFKO4 (5′-AAATTTACTAGTCGACGCATCCGAATGCGCAC-3′) (right flank). The left-flank PCR product was digested with SpeI and EcoRI, and the right-flank PCR product was digested with BamHI and SpeI. Both flanking products and the kanamycin cassette were ligated into the SpeI site of the pBluescript II KS plasmid (Stratagene). The resulting assembly, left flank/Km^r/right flank, was subcloned as an SpeI fragment into pX33 (pPR23 [26] carrying a constitutive xylE marker), generating the pSG16 plasmid. The expected double-crossover event would result in a nonpolar deletion-insertion at the phnF locus, eliminating 80% of the phnF coding sequence in exchange for the kanamycin resistance marker. Allelic replacement of phnF was carried out essentially as described by Pelicic et al. (26) and was achieved by growing a culture of M. smegmatis carrying pSG16 in Middlebrook 7H9-ADC medium at 28°C with agitation (200 rpm) to an OD₆₀₀ of approximately 0.6 to 0.8, followed by plating onto low-salt LBT plates (2 g NaCl liter⁻¹) containing gentamicin and 10% sucrose at 40°C, selecting for double-crossover events. Replacement of phnF with the kanamycin marker created strain SG62 (ΔphnF::aphA-3). For Southern hybridization analysis, SmaI-digested genomic DNA of the putative mutants was separated on a 1% agarose-Tris-acetate-EDTA gel and transferred onto a nylon membrane (Hybond-N⁺; Amersham) by vacuum blotting. Probes were labeled by random priming using [α-³²P]dCTP (Amersham) and Ready-To-Go DNA-labeling beads (Amersham).

The construct used for complementation of the allelic exchange mutant was cloned into the integrative E. coli/mycobacterium shuttle vector pUHA267 (16). A 1.2-kb PCR product encompassing the phnF gene plus 428 bp of upstream DNA was amplified by PCR using the primers cphnFF (5′-AAATTTAAGCTTCATAGTCGGAGGCCTGGACG-3′) and cphnFR (5′-AAATTTAAGCTTCAAGAATCCGGTGTTTCCGC-3′) and cloned into the HindIII site of pUHA267, creating the plasmid pSG41.

Site-directed mutagenesis of the putative PhnF binding sites (inverted repeat unit 1 [IRU-1] and IRU-2) was carried out using PCR overlap-extension (11). For the mutagenesis of IRU-1, a 158-bp PCR product was amplified with the primers PphnFR and SDMIRU1-1 (5′-ACGTCTGTGTCTATCACAGCGGACGGCCGTCTGACGAG-3′), and a 247-bp PCR product was amplified using the primers SDMIRU1-2 (5′-GTCCGCTGTGATAGACACAGACGTATTCGCTTGTTC-3′) and PphnDR (5′-AAATTTGGTACCGCTTGTCGGAGCCCGAACAG-3′). For the mutagenesis of IRU-2, a 216-bp PCR product was amplified with the primers PphnFR and SDMIRU2-1 (5′-GTGGGGTGTGCTATACCAACGGGTGCATCTCGGG-3′), and a 189-bp PCR product was amplified by using the primers SDMIRU2-2 (5′-ACCCGTTGGTATAGCACACCCCACAAGGTGTGTGG-3′) and PphnDR. These fragment pairs were then used in overlap-extension PCR, using the primers PphnFR and PphnDR, and the resulting products were digested with Asp718 and cloned into the Asp718 site of pJEM15 (34). The orientation of the insert determined whether the resulting construct represented a phnD-lacZ or a phnF-lacZ fusion. Mutation of both IRUs simultaneously was achieved by using the fragment containing the changes in IRU-1 as a template for the mutagenesis of IRU-2. A fragment containing the wild-type sequence was amplified by using the primers PphnFR and PphnDR.

RNA extraction and dot blot analysis.For RNA extraction, 5 to 10 ml of broth culture was grown to an OD₆₀₀ of 0.5 to 0.7 in high-phosphate Sauton's medium or was subjected to phosphate starvation as described above. Cells were harvested by centrifugation, resuspended in 1.5 ml GTC buffer (5 M guanidine thiocyanate, 25 mM sodium-citrate [pH 7], 0.05% [wt/vol] Tween 80, 0.05% [wt/vol] Sarkosyl), vortexed for 10 s, pelleted again by centrifugation, and stored at −80°C. Total RNA was extracted using TRIzol reagent (Invitrogen) according to the manufacturer's instructions. Cell lysis was achieved by two cycles of bead beating in a Mini-Beadbeater (Biospec) at 5,000 rpm for 1 min. DNA was removed from the RNA preparation by treatment with 2 U of RNase-free DNase, using a TURBO DNA-free kit (Ambion), according to the manufacturer's instructions. RNA concentrations were determined by using a NanoDrop ND-1000 spectrophotometer. For dot blot analysis, serial fourfold dilutions of RNA samples, from 800 ng to 12.5 ng, were prepared in RNase-free water, and RNA was denatured using glyoxal sample buffer (Cambrex Bio Science, Rockland, ME) according to the manufacturer's instructions. Samples were mixed with 2 volumes of 20× SSC (3 M NaCl, 300 mM sodium citrate, pH 7 [1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate]) and applied to a nylon membrane by vacuum, using a Bio-dot apparatus (Bio-Rad). For use as probes, a 980-bp internal fragment of phnC was PCR amplified using the primers phnCintF (5′-GCTCCGAGAAGTGCAAGGGCTGAC-3′) and phnCintR (5′-CGATCAGCGCGGTCGGTACGAAATC-3′), and a 1-kb fragment of pstC was amplified using the primers pstSKO-3 and pstSKO-4 (5). Probes were labeled by random priming, using [α-³²P]dCTP (Amersham) and Ready-To-Go DNA-labeling beads (Amersham).

Mapping of TSSs.The transcriptional start sites (TSSs) of phnD and phnF were mapped by 5′ rapid amplification of cDNA ends (RACE), using the components of a 3′/5′ RACE kit (Roche) according to the manufacturer's instructions. First-strand cDNA was synthesized from 4 μg of total RNA isolated from phosphate-starved cells of wild-type M. smegmatis with the phnD-specific primer phnD-RACE1 (5′-CTTGTCGATCATGGTCTTG-3′) or the phnF-specific primer phnF-RACE1 (5′-AATTCTTAGCGTGCATGAC-3′). The resulting cDNA was purified, and a deoxyribosyladenine tail was added by following the kit instructions. Purified deoxyribosyladenine-tailed cDNA was then used as a template for PCR using the oligo(dT) anchor primer and the phnD-specific primer phnD-RACE2 (5′-ACGAAGCACACCTTCTTG-3′) or the phnF-specific primer phnF-RACE2 (5′-GTTGAGCAGGAACATGGG-3′). The resulting PCR products were gel purified and used as templates for a second PCR using the PCR anchor primer and the nested phnD-specific primer phnD-RACE3 (5′-CGAGCCCGTAGGACTGGTAGC-3′) or the phnF-specific primer phnF-RACE3 (5′-GTGAAGGCGATCCCACGGCTG-3′). These PCR products were cloned into pGEM-T Easy (Promega) according to the manufacturer's instructions. Three clones containing the correctly sized insert for phnD (ca. 400 bp) or phnF (ca. 550 bp) were sequenced using the primer phnD-RACE3 or phnF-RACE3, respectively, giving consistent results for each gene.

β-Galactosidase and inorganic phosphate assays.To determine the threshold P_i concentration leading to the induction of phnF, cells carrying pSG18 were grown in medium containing 200 μM phosphate. The medium was modified from the standard Sauton's medium used in this study to contain higher concentrations of the carbon source (5 g of glycerol liter⁻¹) and the nitrogen source (4 g l-asparagine liter⁻¹), providing both nutrients in excess. At various time points, 2 ml to 4 ml of the culture was removed to determine the OD₆₀₀. Cells were then pelleted, and cell pellets and supernatants were stored at −20°C. β-Galactosidase activity for the 0-h time point was determined with cells of the inoculum culture. β-Galactosidase activities were determined as described previously (5) and were expressed as Miller units (MU) (23), calculated as the increase in A ₄₂₀ per min per 1 ml of cell suspension used (normalized to an OD₆₀₀ of 1.0) and multiplied by a factor of 1,000. Statistical analyses were performed using a two-tailed, unpaired t test. The P_i concentration in the supernatant was determined by inorganic phosphate assay as described previously (24).

DNA and protein sequence analysis.The provisional genome sequence of M. smegmatis strain mc²155 was accessed via The Institute for Genomic Research (TIGR) website (http://www.tigr.org ; GenBank accession number CP000480). Sequence data for M. tuberculosis were obtained from the Institut Pasteur website (http://genolist.pasteur.fr/TubercuList ). Protein sequence alignments were carried out using a ClustalW function and the BLOSUM62 matrix of BioEdit (9). Secondary-structure predictions were performed using JPred (4). Promoter areas were searched for conserved motifs, using motif discovery and search tool MEME software, available at the San Diego Supercomputer Center website (http://meme.sdsc.edu/meme/ ).

Identification of a putative regulator in the phn locus of M. smegmatis.The PhnDCE phosphate transport system of M. smegmatis is encoded by a three-gene operon (Fig. 1A) (5). A gene adjacent to and divergently transcribed from this operon, MSMEG_0650, has been annotated as a GntR family transcriptional regulator encoding a 244-amino-acid protein. According to the results of a conserved-domain search (18), the protein was called PhnF (36) (Fig. 1A). A search for TIGRFAM and Pfam matches for PhnF, using the search tools available via the TIGR website (http://cmr.tigr.org ), revealed that the N terminus (amino acids 5 to 65) of the protein contained a helix-turn-helix motif of a GntR-like transcriptional regulator. The C-terminal part of the protein (amino acids 85 to 224) was a UbiC transcriptional regulator-associated (UTRA) domain, which is common to members of the HutC subfamily of GntR-like regulators and has been proposed to function in ligand binding (2). The predicted secondary structure of this region of M. smegmatis PhnF (Fig. 1B) showed a fold similar to the typical tandem arrangement of two α2-β-α-β2 repeats, further confirming that like the PhnF protein of E. coli (27), the M. smegmatis PhnF protein also belongs to the HutC subfamily. While few residues of the ligand binding pocket are conserved among members of this subfamily, two large residues in sheet 2 and one polar residue in sheet 3 appear to be universally conserved (2). An alignment between the E. coli and the M. smegmatis PhnF proteins shows that with the exception of Leu135 (E. coli numbering), these residues are conserved in the M. smegmatis protein (Fig. 1B). The crystal structure of E. coli PhnF showed that five residues conserved among PhnF orthologues delineate the proposed binding cavity (8). Only two of these residues (R181 and S228 [E. coli numbering]) are also conserved in the M. smegmatis protein (Fig. 1B). E. coli PhnF is thought to respond to the presence of alkylphosphonates (2, 8), but this seems unlikely to be the signal recognized by M. smegmatis PhnF, because the PhnDCE system does not appear to transport such compounds and because M. smegmatis cannot utilize phosphonates as phosphorous sources for growth (5). Differences in the signaling molecule recognized by the proteins from E. coli and M. smegmatis could explain the poor conservation of binding site residues between them. No phnF homologues have been identified in any of the other sequenced mycobacterial genomes, but a locus containing homologues to all four of the phn genes of M. smegmatis has been annotated in the genome of another actinomycete, Nocardia farcinica IFM 10152 (14).

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FIG. 1.

Sequence analysis of M. smegmatis PhnF. (A) Map of the phn genes of M. smegmatis. The names of the loci, as annotated in the provisional genome sequence of M. smegmatis mc²155, are indicated below the arrows. (B) Alignment of PhnF proteins from E. coli (Eco) and M. smegmatis (Msm). Identities are shown in black, and similarities (threshold, 90%) are shown in a gray background. The predicted secondary structure of M. smegmatis PhnF is shown below the sequence, with H indicating an α helix and E indicating a β sheet. Triangles indicate the large (L) and polar (H or T) signature residues of HutC subfamily proteins (2). Diamonds indicate the residues conserved among HutC members, which delineate the binding site of E. coli PhnF (8). Solid and open symbols show residues conserved and not conserved, respectively, in M. smegmatis PhnF.

The expression of phnF is induced by phosphate limitation.Both of the operons for the high-affinity phosphate transport systems identified in M. smegmatis to date, phnDCE and pstSCAB, are induced when the culture enters phosphate limitation at a threshold phosphate concentration of 40 μM (5). If PhnF had a role in the regulation of one or both of these operons, then it was conceivable that the expression of the phnF gene also responded to the concentration of phosphate in the growth medium. To test this hypothesis, we created a transcriptional fusion of the putative promoter area (ca. 500 bp) of phnF to that of lacZ (pSG18). An M. smegmatis strain carrying this construct was grown in a phosphate-limited minimal medium as described previously (5), and the phosphate concentration, the β-galactosidase activity, and the OD₆₀₀ were monitored throughout growth. Activities of the phnF-lacZ fusion rapidly increased from between 25 and 40 MU under phosphate-replete conditions to approximately 150 MU when the phosphate concentration in the medium dropped to below 40 μM (Fig. 2). The expression of phnF-lacZ therefore followed the same expression pattern as that of phnD-lacZ and pstS-lacZ, even though the magnitude of induction (ca. 4-fold) was much smaller than those observed for phnD and pstS (26-fold and 17-fold, respectively) (5). These data strongly suggest that PhnF is involved in the adaptation of M. smegmatis to phosphate-limited conditions.

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FIG. 2.

Transcriptional activities of phnF-lacZ during phosphate-limited growth. Cells were grown in modified minimal Sauton's medium (5 g glycerol liter⁻¹, 4 g l-asparagine liter⁻¹, 200 μM P_i) and monitored for growth, expressed as OD₆₀₀ (▪), phosphate concentration in the medium (P_i) (▵), and β-galactosidase activity (β-Gal), expressed as MU (□). Representative results of two independent experiments are shown.

Construction of a phnF deletion mutant.To further elucidate the function of PhnF, we created a phnF deletion mutant by allelic exchange mutagenesis. A construct in which phnF was replaced with a kanamycin resistance cassette, aphA-3, was cloned into pX33, as described in Materials and Methods, and M. smegmatis mc²155 was transformed with the resulting plasmid. Knockout mutants were selected as described previously (26). The replacement of phnF with the antibiotic marker introduced two additional SmaI restriction sites, resulting in a band shift from 6.7 kb in the wild type to 3.2 kb in the deletion mutant (strain SG62) in Southern hybridization analysis of SmaI-digested genomic DNA probed with a radiolabeled PCR product of the left flank of the deletion construct (Fig. 3). For complementation, an integrative plasmid containing phnF plus ca. 500 bp of the region upstream of the predicted translational start site was introduced into strain SG62, creating strain SG111.

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FIG. 3.

Allelic replacement of phnF. (A) Schematic diagram (not drawn to scale) of allelic replacement of phnF with aphA-3. SmaI restriction sites and band sizes as detected in panel B are indicated. The bold line shows the fragment used as a probe. (B) Southern hybridization analysis of the replacement of phnF in strain SG62. SmaI-digested genomic DNA from the wild-type (WT) strain and from strain SG62 (ΔphnF) was probed with the radiolabeled left-flank PCR product of the deletion construct. Molecular sizes are indicated in kilobases.

Deletion of phnF leads to the overexpression of phnDCE but not pstSCAB.In order to determine any of the effects that the deletion of phnF would have on the expression of the phnDCE and pstSCAB operons, we analyzed the relative amounts of mRNA synthesized by both of the operons in the wild type and the phnF deletion strain (SG62) under high- and low-phosphate conditions. In the wild-type strain, both of the phnDCE and pstSCAB operons were expressed at low levels in the high-phosphate samples, and expression was increased strongly in phosphate-starved cells (Fig. 4A), as was expected from previous results obtained from transcriptional lacZ fusion analyses (5). The deletion of phnF had no effect on the expression of the pstSCAB operon, which showed the same expression patterns in the SG62 ΔphnF strain, the SG111 complemented strain, and the wild-type strain (Fig. 4A, top panel). In contrast, the expression of phnDCE was strongly increased in the phosphate-starved cells of the phnF deletion mutant strain SG62 compared to that in the cells of the wild-type strain (Fig. 4A, bottom panel). The expression level of phnDCE under high-phosphate conditions was not changed in the SG62 strain. Complementation of the phnF deletion completely restored expression patterns to the wild-type level, confirming that the increase observed for expression under phosphate-starved conditions was indeed due to the deletion of phnF. These data clearly show that PhnF acts as a repressor of phnDCE. PhnF appears to have no role in the regulation of pstSCAB. The fact that the deletion of phnF does not lead to full constitutive expression of phnDCE under high-phosphate conditions suggests that a further regulatory mechanism is required for the induction of the operon. This hypothesis is addressed below.

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FIG. 4.

Dot blot analysis of RNA from wild-type and mutant strains. Fourfold dilutions of total RNA isolated from cells grown in high-phosphate medium (100 mM; +) or subjected to phosphate starvation for 2 h (−) were spotted onto nylon membranes. Membranes were probed with radiolabeled PCR products from internal fragments of pstC (A, top panel) or phnC (A, bottom panel, and B). Amounts of total RNA per spot are shown in ng. Strains are indicated above the autoradiographs. (A) WT, wild-type; ΔphnF, phnF deletion strain SG62; cphnF, phnF-complemented strain SG111. (B) WT, wild-type; ΔpstS, pstS deletion strain SG95; cpstS, pstS-complemented strain SG120. Representative results of two to three independent experiments are shown. Below each panel, 16S and 23S rRNA bands from 300 ng total RNA per sample on an agarose gel stained with ethidium bromide are shown as controls. Lanes correspond to the samples in the rows above.

Deletion of pstS leads to the constitutive expression of pstSCAB, phnDCE, and phnF.As mentioned above, the expression of the phnDCE operon appears to be subject to dual regulation: repression by PhnF and activation by a second regulatory mechanism. The most probable candidate for this second regulatory mechanism is the SenX3-RegX3 two-component system, which is reported to regulate the expression of the pstSCAB operon (6). SenX3, like its functional counterpart PhoR of E. coli, lacks any significant extracellular loops, which led to the proposal that the Pst system of M. smegmatis functions as the actual sensor of extracellular phosphate concentration (6). Accordingly, mutations in either pstS, pstC, or pstA are sufficient for the derepression of phoA expression (15), which is known to be under the control of SenX3-RegX3 (6). We therefore utilized the M. smegmatis pstS deletion mutant (strain SG95) and the pstS-complemented strain (SG120) (5) to determine whether the expression of phnDCE was affected by pstS deletion and thus was likely to be under the control of SenX3-RegX3. Using RNA dot blot analysis, a small but reproducible increase in the expression level of phnDCE was observed with the high-phosphate samples of strain SG95 (ΔpstS) compared to that in the wild-type strain, and no differences in the wild-type expression levels were observed with phosphate-starved cells (Fig. 4B). Complementation of the pstS deletion restored phnDCE expression to the wild-type pattern.

To study this pattern of gene expression in more detail, we introduced the transcriptional phnD-lacZ construct (pSG10) (5) into strain SG95 (ΔpstS) and monitored the expression patterns in both the phosphate-replete and the phosphate-starved cells. As a control, the transcriptional pstS-lacZ construct, pSG42 (5), was introduced into strain SG95 (ΔpstS), because the pstS promoter is known to be regulated by RegX3 (6) and thus should be constitutively expressed in a pstS deletion background. In cells of the wild-type strain carrying the pstS-lacZ construct pSG42, phosphate starvation led to a five- to sixfold increase in β-galactosidase activity, from ca. 10 MU to 55 MU, compared to that in cells grown under phosphate-replete conditions (Fig. 5A). Cells of the pstS deletion strain SG95 harboring pstS-lacZ had β-galactosidase activities of ca. 80 MU, independent of the phosphate concentration available. In the complemented strain SG120, regulation was restored, and phosphate starvation led to a fivefold induction of β-galactosidase activity, although the absolute levels of activity were lower than in the wild-type strain. These data show that the deletion of pstS causes constitutive expression from the RegX3-dependent pst promoter. Cells of the wild-type strain carrying the phnD-lacZ construct pSG10 displayed a 20-fold induction of β-galactosidase activity in response to phosphate starvation (Fig. 5B). In contrast, the expression of phnD-lacZ in SG95 (ΔpstS) was constitutive at around 100 MU. Complementation of the pstS deletion restored the regulation of expression to that of wild-type levels.

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FIG. 5.

Expression levels of pstS-lacZ, phnD-lacZ, and phnF-lacZ in various genetic backgrounds of M. smegmatis. Cells of the wild type (WT), the pstS deletion strain (SG95), and the complemented strain (SG120) carrying various lacZ fusion constructs were grown in ST medium containing 100 mM P_i (open bars) or subjected to phosphate starvation for 2 h (gray bars). β-Galactosidase activities are given as MU. (A) Cells harboring the pSG42 plasmid (pstS-lacZ). (B) Cells harboring the pSG10 plasmid (phnD-lacZ). (C) Cells harboring the pSG18 plasmid (phnF-lacZ). Results are shown as the means and standard deviations of results from two to four independent experiments. Differences between cells grown in 100 mM P_i and phosphate-starved cells of the wild-type and those of the SG120 strain and differences between the wild-type cells grown in 100 mM P_i and the cells of the SG95 strain are statistically significant (P < 0.05).

To study the effect of pstS deletion on phnF expression, phnF-lacZ activity was measured in the wild-type strain and in the SG95 strain (ΔpstS). In the wild-type strain carrying the phnF-lacZ construct, phosphate starvation led to a 2.6-fold induction, from 35 MU to 90 MU, while in strain SG95, the expression of phnF-lacZ was constitutive at about 80 MU (Fig. 5C). In the complemented strain SG120, β-galactosidase activity was lower than that in the wild-type strain, but phosphate starvation-dependent induction was restored to a 3-fold level.

These data demonstrate that pstS deletion leads to constitutive expression of phnDCE and phnF under phosphate-replete and phosphate-starved conditions. However, RNA analysis of phnDCE expression in phosphate-replete cells of the pstS deletion mutant indicated that the derepression was only partial. We attribute this discrepancy to the differences in copy numbers of the phnDCE promoter region in the two experiments: for RNA analysis, PhnF is able to exert its repressive effect on the single copy of the phnDCE promoter. In contrast, phnD-lacZ is present as 3 to 10 copies per cell (34), and therefore the effect of PhnF is titrated out. Taken together, these results show that the expression from the phnDCE and phnF promoters, like that from the pstSCAB promoter, is increased in a pstS deletion background. The phnDCE operon and phnF thus appear to be likely new candidates for the SenX3-RegX3 regulon of M. smegmatis, in which phnDCE is under additional control by the repressor PhnF. Further work is required to confirm the involvement of SenX3-RegX3 in the regulation of phnDCE and phnF and whether there is direct interaction between RegX3 and these promoters.

The region between phnF and phnD contains two putative binding sites for PhnF.To gain further understanding of the involvement of the different transcription factors in the regulation of the phnDCE and phnF promoters, we determined the TSSs for both phnD and phnF, using 5′ RACE analysis (Fig. 6). The TSS for phnD was determined as the “G” located 65 bp upstream of the translational start. No consensus −10 or −35 sequences could be identified for phnD. The TSS for phnF was determined as the first nucleotide of the GTG start codon, suggesting that phnF is transcribed as a leaderless transcript, a feature which has also been observed for other GntR family transcriptional regulators from actinomycetes (12, 30). Putative −10 (5′-TACGTT-3′) and −35 (5′-TCTGAC-3′) boxes with some similarities to mycobacterial promoter elements (1, 29) could be identified upstream of the phnF TSS (Fig. 6).

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FIG. 6.

Sequence analysis of the intergenic region between phnF and phnD. Transcriptional start sites for phnF and phnD were determined by 5′ RACE analysis as shown in the top and bottom panels, respectively (traces are given as reverse sequences). The sequence given in the middle panel shows a region of the coding strand for phnD, encompassing the first three codons of phnF and phnD (the first three amino acids are indicated) and the intergenic region. It should be noted that the sequence for phnF is therefore given as a reverse sequence. Start codons are shown in bold; transcriptional start sites are shown in bold and are indicated as +1. Putative −10 and −35 regions for phnF are underlined. The two IRUs, which constitute presumptive PhnF binding sites, are boxed.

Next, we searched both promoter regions for conserved binding motifs. RegX3 of M. smegmatis has been shown to bind to a loosely conserved inverted repeat (GTGAAC) separated by seven nucleotides in the promoters of phoA, pstS, and senX3 (6). We analyzed 500-bp regions upstream of phnD and phnF but were unable to identify sequences with obvious similarity to the previously described RegX3 binding sites. Analysis of the same promoter areas, together with 500-bp regions with phosphate-responsive promoter activity for one of the M. tuberculosis pst operons (35), using a MEME motif discovery and search tool, also did not lead to the identification of potential RegX3 binding sites. It is likely that the number of mycobacterial genes known to respond to phosphate starvation is not yet large enough to identify common motifs in their promoter areas.

As discussed above, PhnF is a member of the HutC subfamily of GntR-like transcriptional regulators. A core recognition sequence for HutC-type regulators has been predicted (27), and more recently, the binding site of a member of this subfamily, DasR from Streptomyces coelicolor, has been identified as the 16-bp palindromic sequence ACTGGTCTACACCATT (28). Analysis of the intergenic region between M. smegmatis phnD and phnF revealed the presence of two inverted repeats with a similar sequence of TGGTATAGACCA, which we termed IRU-1 and IRU-2 (Fig. 6). The same sequence has recently been identified as a potential binding site for PhnF by an in silico analysis of the M. smegmatis GntR family regulators (38).

To further investigate the role of IRU-1 and IRU-2, we introduced site-directed changes on a 377-bp PCR product encompassing the 199-bp intergenic region between phnF and phnD plus 93 bp of the start of the phnF coding region and 85 bp of the start of the phnD coding region. IRU-1 was changed to TGTGATAGACAC (mutated nucleotides are underlined), changing the sequence of each half site as well as the palindromic structure. The 5′ half of IRU-2 reaches to nucleotide −31 relative to the phnD TSS. To avoid introducing changes into a region that might function as a −35 element of the phnD promoter, this part of the IRU-2 was therefore left unchanged, leading to a mutated sequence of TGGTATACCACA. A third fragment containing the changes in both regions was amplified. The three mutated fragments as well as a PCR product containing the wild-type sequence were then used to construct transcriptional lacZ fusions of either the phnF or the phnDCE promoters, depending on the orientation of the insert. The effect of the mutations on promoter activity were then determined with cells grown under phosphate-replete conditions (100 mM phosphate in minimal Sauton's medium) and after 2 h of phosphate starvation. β-Galactosidase activities in cells carrying constructs with the wild-type sequence were comparable to the results obtained previously for the phnF-lacZ and phnD-lacZ constructs, in which phosphate starvation led to ca. 20-fold induction for the phnD-lacZ construct (Fig. 7A) and 3- to 4-fold induction for the phnF-lacZ construct (Fig. 7B). We therefore concluded that the fragment used for site-directed mutagenesis was sufficient for the phosphate-dependent control of phnDCE and phnF expression. The mutation of IRU-1 led to a partial but significant (P < 0.0001) derepression (an 8-fold increase compared to that of the wild-type promoter) of the phnDCE promoter under phosphate-replete conditions (Fig. 7A). Phosphate starvation further induced expression by approximately 2.7-fold (P < 0.0001). The mutation of IRU-2 had no significant effect on the expression of the phnDCE promoter. The mutation of both putative binding sites simultaneously increased the expression of the phnDCE promoter even further than the mutation of IRU-1 alone, under both phosphate-replete and phosphate-starved conditions (P < 0.01). The expression of phnF was unaffected by the mutation of IRU-1, whereas the mutation of IRU-2 led to a 2.5-fold increase in β-galactosidase activity under phosphate-replete conditions compared to that of the wild-type promoter (P < 0.0001) (Fig. 7B). Phosphate starvation still led to a twofold induction of expression (P = 0.0001). As observed for the phnDCE expression level, the mutation of both IRU-1 and IRU-2 simultaneously further increased the expression of phnF under both conditions tested (P < 0.05). These data suggest that IRU-1 is required for the repression of phnDCE transcription, while IRU-2 is required to repress the transcription of phnF. IRU-1 is centered at position −83.5 relative to the phnD TSS. IRU-2 is centered at position −110.5 relative to the phnF TSS. It is likely that IRU-1 and IRU-2 are binding sites for PhnF, and phnF would therefore appear to be subject to autoregulation.

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FIG. 7.

Effects of site-directed mutagenesis of the IRUs in the intergenic region between phnF and phnD on gene expression. Changes were introduced into IRU-1 and IRU-2 (see text for details), and effects were monitored as the expression of phnD-lacZ (A) and phnF-lacZ (B) transcriptional fusions. Cells were grown in minimal Sauton's medium containing 100 mM phosphate (open bars) or starved in phosphate-free medium for 2 h (gray bars). The presence of wild-type sequences (+) or site-directed changes (−) in each IRU are shown below the graphs. Results are shown as the means and standard deviations from three independent experiments.

Conclusions.In the present study, we investigated the transcriptional regulation of two operons, pstSCAB and phnDCE, encoding high-affinity phosphate transport systems of M. smegmatis. We showed that the phn locus contains another gene, phnF, which encodes a functional transcriptional regulator belonging to the HutC subfamily of GntR-like regulators. Furthermore, we demonstrated that PhnF acts as a repressor of the phnDCE operon but does not affect the expression of pstSCAB. Two presumptive binding sites for PhnF were identified in the region between phnD and phnF, and these sites are required for the repression of phnD and phnF. The deletion of pstS leads to the constitutive expression of phnDCE and phnF, suggesting that these genes may be under the control, directly or indirectly, of the SenX3-RegX3 system of M. smegmatis.

Based on these findings, we propose a model for the regulation of the phnDCE and pstSCAB operons in M. smegmatis (Fig. 8). Both operons are strongly induced by phosphate limitation. The pstSCAB operon is under the sole and direct control of SenX3-RegX3. In contrast, phnDCE expression at a low-phosphate concentration requires both the derepression by PhnF and the activation by a second system, presumably SenX3-RegX3. Transcription of phnF also appears to be under such dual control of activation by SenX3-RegX3 and repression by PhnF itself. Increased expression of PhnF under phosphate-limited conditions may supply the cell with a means to reestablish the repression of the Phn system when phosphate availability improves. Such a phenomenon of induction of repressors for the Pho regulon, pstSCAB and phoU, has been reported for E. coli and seems to be required to terminate the phosphate starvation response (39). Further work is required to determine whether PhnF and RegX3 interact directly with the promoters studied here or indirectly via additional regulatory systems and to study how these different regulatory proteins interact with each other or with RNA polymerase. The differences in regulation between the Pst and Phn systems of M. smegmatis described here may shed light on the differences between the multiple phosphate transport systems present in other mycobacterial species. It has been proposed previously that the requirement of several such systems by mycobacteria may reflect subtle adaptations of the bacteria to a highly variable environment (17). In this case, the differential regulation of gene expression, as observed here, may be a key feature of mycobacterial phosphate transport systems.

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FIG. 8.

Model for the regulation of pstSCAB, phnDCE, and phnF in M. smegmatis. Genes are shown as open arrows; proteins are shown as gray ovals. Flat-headed arrows indicate negative regulation; a pointed arrowhead indicates positive regulation. Dotted lines indicate that the proposed regulation may be direct or indirect. The presumptive PhnF binding sites are shown as black diamonds.