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Journal of Bacteriology, March 2004, p. 1493-1502, Vol. 186, No. 5
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.5.1493-1502.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Residues Required for Bacillus subtilis PhoP DNA Binding or RNA Polymerase Interaction: Alanine Scanning of PhoP Effector Domain Transactivation Loop and {alpha} Helix 3

Yinghua Chen,1 Wael R. Abdel-Fattah,1 and F. Marion Hulett1*

Laboratory for Molecular Biology, Department of Biological Sciences, University of Illinois at Chicago, Chicago, Illinois 606071

Received 22 September 2003/ Accepted 18 November 2003


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Bacillus subtilis PhoP is a member of the OmpR family of response regulators that activates or represses genes of the Pho regulon upon phosphorylation by PhoR in response to phosphate deficiency. Because PhoP binds DNA and is a dimer in solution independent of its phosphorylation state, phosphorylation of PhoP may optimize DNA binding or the interaction with RNA polymerase. We describe alanine scanning mutagenesis of the PhoP {alpha} loop and {alpha} helix 3 region of PhoPC (Val190 to E214) and functional analysis of the mutated proteins. Eight residues important for DNA binding were clustered between Val202 and Arg210. Using in vivo and in vitro functional analyses, we identified three classes of mutated proteins. Class I proteins (PhoPI206A, PhoPR210A, PhoPL209A, and PhoPH208A) were phosphorylation proficient and could dimerize but could not bind DNA or activate transcription in vivo or in vitro. Class II proteins (PhoPH205A and PhoPV204A) were phosphorylation proficient and could dimerize but could not bind DNA prior to phosphorylation. Members of this class had higher transcription activation in vitro than in vivo. The class III mutants, PhoPV202A and PhoPD203A, had a reduced rate of phosphotransfer and could dimerize but could not bind DNA or activate transcription in vivo or in vitro. Seven alanine substitutions in PhoP (PhoPV190A, PhoPW191A, PhoPY193A, PhoPF195A, PhoPG197A, PhoPT199A, and PhoPR200A) that specifically affected transcription activation were broadly distributed throughout the transactivation loop extending from Val190 to as far toward the C terminus as Arg200. PhoPW191A and PhoPR200A could not activate transcription, while the other five mutant proteins showed decreased transcription activation in vivo or in vitro or both. The mutagenesis studies may indicate that PhoP has a long transactivation loop and a short {alpha} helix 3, more similar to OmpR than to PhoB of Escherichia coli.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
In Bacillus subtilis the response to phosphate deprivation is controlled by activation of genes previously silent and repression of other genes whose products are antithetical to cell survival in phosphate-limiting conditions. At least two global systems are responsible for these changes, sigma B (3) and PhoP-PhoR. PhoP and PhoR comprise a prototypical two-component system in that the transmembrane sensor histidine kinase, PhoR, is autophosphorylated on a conserved histidine residue in response to a signal. The phosphoryl group is then transferred to a conserved aspartate residue in the N-terminal domain of the response regulator, PhoP, thus activating PhoP for transcriptional regulation of Pho regulon genes. PhoP~P is known to directly regulate 31genes in seven operons (14) and is directly or indirectly required for eight additional genes (23, 24). Among the genes that are activated or repressed by PhoP~P are the genes that encode a high-affinity phosphate ABC transporter, phosphatases (PhoA, PhoB, and PhoD), and biosynthetic proteins for essential anionic cell wall polymers (teichoic acid or teichuronic acid).

PhoP is a member of the winged helix OmpR subfamily of response regulators based on sequence similarities of the C-terminal effector domains. The 240-amino-acid (aa) PhoP molecule is composed of two functional domains, an N-terminal receiver domain consisting of 119 aa (PhoPN) and a 102-aa C-terminal effector domain (PhoPC) involved in DNA binding and transactivation via RNA polymerase (RNAP). A long linker region (19 aa) connects the regulatory domain (PhoPN) to the output domain (PhoPC). The receiver domain of PhoP has been structurally characterized (5). The structure analysis revealed overall folds similar to those of two other OmpR family proteins, PhoB from Escherichia coli (7) and DrrD from Thermotoga maritima (8), but showed that there are remarkable differences in the ß4-{alpha}4 loop and {alpha}4 helix regions compared to either PhoB or DrrD despite the high levels of sequence similarity exhibited by the three proteins in these regions. The PhoPN structure analysis also showed that there is a novel asymmetric association between PhoPN protomers that supports the established DNA binding properties of PhoP. The dimer interface between two PhoP monomers involves nonidentical surfaces such that each monomer in a dimer has a second surface available for further oligomerization. DNA footprinting studies have shown that there is cooperative binding of PhoP dimers at PhoP-activated promoters and have revealed that both phosphorylated and nonphosphorylated dimers can bind to the four 6-bp direct repeats, TT(A/C/T)A(C/T)A, spaced 4 to 6 bp apart that constitute the core binding region for PhoP at PhoP-activated promoters (12). PhoP footprinting data for repressed promoters (6, 18) suggest that oligomerization of PhoP along the DNA extends well into the coding region. Support for the physiological relevance of the asymmetric dimerization interface of PhoP was obtained when mutations designed to disrupt this interface resulted in PhoP mutant proteins that could be phosphorylated by PhoR but could not dimerize or bind DNA in vitro and had no activity in vivo (9).

Our previous footprinting studies showed that there was cooperative binding between PhoP dimers at PhoP-activated promoters and that both nonphosphorylated and phosphorylated PhoP could bind to the four 6-bp repeats that form the core binding region for PhoP, suggesting that nonphosphorylated and phosphorylated PhoP dimers have structural relationships in common that allow similar binding of their effector domains to target DNA. The fact that phosphorylation is required for transcriptional activation or repression suggests that it may enhance cooperativity for DNA binding or interactions with components of the transcription machinery. Studies described here were initiated to determine which residues of the effector domain, PhoPC, are required for PhoP DNA binding and which residues are required for transcriptional regulation. For certain OmpR homologues the purified winged helix domain is sufficient for DNA binding to target DNA as a dimer (7, 15, 20). In fact, the DNA binding domain of PhoB, the presumed E. coli orothologue of B. subtilis PhoP, has a higher affinity for target DNA than the nonphosphorylated intact PhoB protein has (13). In contrast, the DNA binding domain of PhoP was isolated and was shown to be a monomer in solution that required more than fivefold more protein for DNA binding to target DNA than nonphosphorylated intact PhoP required. In addition, expression of PhoPC in B. subtilis could not induce Pho regulon genes, at least in part due to the instability of this domain in vivo (Chen and Hulett, unpublished data). For these reasons mutant proteins resulting from alanine scanning mutagenesis of the PhoP {alpha} loop and the {alpha} helix 3 region of PhoPC domain were constructed with the intact PhoP protein for both in vivo and in vitro functional analyses.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Strains and plasmids. E. coli DH5{alpha} (lab stock) and XL1-Blue (Stratagene) were used as the hosts for plasmid construction (Table 1). E. coli BL21(DE3)(pLysS) (Novagen) was used as the host for overexpressing the PhoP proteins. B. subtilis JH642 and MH6110, a derivative of JH642, were used for in vivo Pho induction experiments. When required, antibiotics were added at the following concentrations: for B. subtilis strains, 5 µg of chloramphenicol per ml, 10 µg of spectinomycin per ml, or 10 µg of tetracycline per ml; and for E. coli strains, 100 µg of ampicillin per ml or 50 µg of kanamycin per ml. 5-Bromo-4-chloro-3-indolyl-ß-D-galactopyranoside (X-Gal) was used at a concentration of 30 µg/ml, and isopropyl-ß-D-thiogalactopyranoside (IPTG) was used at a concentration of 1 mM.


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  		TABLE 1. Bacterial strains and plasmids

 
To construct MH6110 ({Delta}phoPR::Tetr amyE::phoA-lacZ Spr), the chromosomal DNA from MH5923 (amyE::phoA-lacZ Spr) was used to transform MH5913 ({Delta}phoPR::Tetr). Transformants were selected on plates containing tryptose blood agar base (Difco) supplemented with 0.5% glucose (TBABG) containing tetracycline and spectinomycin. The representative clones were confirmed by phenotypic screening.

To generate a B. subtilis strain with an IPTG-inducible phoPR operon, pWL29 was digested with ClaI, treated with the Klenow enzyme, and relegated, and the resulting plasmid was designated pCH101. This plasmid was digested with BpuI102I and SphI, which removed a 178-bp fragment of the phoP gene 3' region, and then ligated with the 2.9-kbp (2,875-bp) BpuI102I/SphI fragment obtained from pHT4phoPR that contained the 178 bp 3' of phoP, the complete phoR gene, and 750 bp 5' of the polA gene, yielding pCH102. pCH102 was transformed into MH6110 and integrated into the chromosome by homologous recombination. Transformants were selected for Cmr and screened for Spr and Tetr. The position of homologous recombination, 3' or 5' of the Tetr gene in chromosome, was determined by PCR to be in the phoR polA region downstream of the Tetr gene in MH6110. The resulting B. subtilis strain, MH6111, contains the complete phoPR operon under control of the Pspac promoter.

Plasmids similar to pCH102 but with a mutation at codons affecting PhoP amino acid residues 190 to 214 were constructed as follows. The 676-bp fragment containing the phoP gene 3' region and the phoR gene 5' region was released from pCH102 by digestion with ClaI and SacI and cloned into pBluescript KS(+) at the same sites in order to generate deltaPRKS+. Most of the PhoP codons from the V190 codon to the E214 codon in plasmid pdeltaPRKS+ were individually mutated to the alanine codon, GCG, by using a QuickChange site-directed mutagenesis kit (Stratagene) and the necessary primer pairs; the only exception was the A196 codon, GCC, which was mutated to the V196 codon, GAT. After sequence confirmation, the 676-bp fragment with the required mutation was released from plasmid pdeltaPRKS+ by digestion with ClaI and SacI and was used to replace the same region in pCH102, yielding the Pspac-controlled mutated phoPR operon with a ribosome binding site and a partial 5' polA gene. Plasmids pCH103 to pCH127 were transformed into MH6110, and representative transformants were used in in vivo experiments. The mutant strains were confirmed by PCR and DNA sequencing.

To construct a plasmid for overexpressing mutant PhoP proteins, the phoP gene was released from pCH01 (9) by digestion with NdeI and BamHI and cloned into pSKB3 at the same sites to generate pCH128. The PhoP codons from V190 to E214 were individually mutated as described above, yielding plasmids pCH129 to pCH153. For the PhoP codon substitution R113 to E113, plasmid pCH05 (9) was digested with NdeI and BamHI and cloned into pSKB3 at the same sites to generate pCH154. The required mutation was confirmed by DNA sequencing. These plasmids were transformed into E. coli BL21(DE3)(pLysS), and the representative transformants were used to overexpress PhoP proteins.

phoA-lacZ plate assay. Low-phosphate complex medium (LPM) or low-phosphate complex medium with 10 mM phosphate added (HPM) contained Noble agar (1.5%), X-Gal (30 µg/ml), and IPTG (1 mM) (26). The fresh transformants generated as described above (typically 100 transformants per construct) were selected and grown on TBABG agar plates with antibiotics for 16 h at 37°C. The clones were then grown on LPM or HPM agar containing X-Gal and IPTG for 16 h at 37°C.

Growth conditions and APase activity assay. Alkaline phosphatase (APase) activity was measured in cells that had been grown in low-phosphate defined medium (LPDM) as described previously (9) with IPTG present at a concentration of 1 mM throughout growth.

Western immunoblotting. Samples (50 ml) were taken from an LPDM culture at 11 h. The cells were separated from the medium by centrifugation at 10,000 x g for 10 min. The pellet fraction was suspended in 5 ml of Tris-EDTA containing 0.8 M sucrose, and then lysozyme was added to the cell suspension at a final concentration of 5 mg/ml. After the mixture was incubated at 37°C for 10 min, the cells were collected by centrifugation at 5,000 x g for 10 min and washed twice with Tris-EDTA containing 0.8 M sucrose. The pellet fraction was then suspended in 5 ml of Tris-EDTA containing 0.3 M NaCl, 1 mM phenylmethylsulfonyl fluoride, and 1 mM MgCl2 and subjected to sonication immediately. After centrifugation at 100,000 x g for 1 h, the supernatant fraction was used as the soluble protein fraction. Equal volumes of an MH5913 ({Delta}phoPR) cell extract containing different dilutions of purified wild-type PhoP (PhoPWT) were used to generate a standard curve. Sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) separation and immunodetection were performed as described previously (9).

Overexpression and purification of PhoP proteins. E. coli BL21(DE3)(pLysS) harboring a PhoP protein-overexpressing plasmid under control of a T7/lac promoter was incubated overnight at 37°C in Luria-Bertani medium containing ampicillin and was used to inoculate 1 liter of the same medium at a ratio of 1:100. The cells were grown at 30°C until the optical density at 600 nm of the culture reached about 0.4. IPTG was then added to the culture at a final concentration of 1 mM, and the culture was grown for another 3 h. The cells were harvested by centrifugation at 4°C and washed with buffer A (1 M NaCl, 5 mM MgCl2, 10 mM dithiothreitol, 50 mM Tris-HCl [pH 7.8]). The cell pellets were then suspended in 50 ml of prechilled buffer A on ice containing 1 mM phenylmethylsulfonyl fluoride and were immediately subjected to sonication. Disruption of the cells was confirmed by phase-contrast microscopy. After centrifugation at 40,000 x g for 1 h at 4°C, the supernatant fraction was filtered through a 0.45-µm-pore-size membrane (Amicon). After 1/50 volume of 0.5 M imidazole in buffer A was added, the clear cell lysate was mixed with 2 ml of Ni-nitrilotriacetic acid (Qiagen) affinity resin preequilibrated with buffer A. After gentle shaking at 4°C for 30 min, the mixture was loaded onto an Econo column (inside diameter, 2.5 cm; height, 10 cm; Bio-Rad). The column was washed with buffer A until the protein concentrations in the elute were not detectable by the Bio-Rad protein assay. The protein bound to the column was eluted by using 300 mM imidazole in buffer A. The protein fractions were dialyzed stepwise at 4°C against buffer A containing 20% glycerol with decreasing concentrations of NaCl (1, 0.8, 0.6, 0.4, 0.2, and 0.1 M). The PhoP proteins were more than 95% pure, as judged by SDS-PAGE.

Phosphotransfer assays. Glutathione S-transferase (GST)-*PhoR (30), prepared in phosphorylation buffer (50 mM KCl, 5 mM MgCl2, 50 mM HEPES [pH 8.0]), was used to phosphorylate each PhoP. Boiled glutathione beads (400 mg) were washed with phosphorylation buffer and incubated with 200 µg of GST-*PhoR on a rocker at room temperature for 10 min. The unbound component was washed off the beads with 20 volumes of phosphorylation buffer, and the extra buffer was removed by microcentrifugation for 10 s. Phosphorylation buffer (0.4 ml) with 20 µl of [{gamma}-32P]ATP (10 mCi/ml) was added to the beads, and autophosphorylation of GST-*PhoR was performed at room temperature for 20 min. The beads were thoroughly washed with phosphorylation buffer until there was no ATP in the flowthrough. The beads bound with GST-*PhoR~P were suspended in 0.4 ml of phosphorylation buffer containing 50 U of thrombin (Pharmacia) and incubated for 15 min at room temperature. The released *PhoR~P was recovered by microcentrifugation through a Micro Bio-Spin chromatography column (Bio-Rad). For phosphotrasfer reactions, *PhoR~P was mixed with each PhoP protein at a molar ratio of 1:15 in phosphorylation buffer. After incubation at room temperature for 10 or 60 s or 10 min, an equal-volume sample was taken and mixed with 4x SDS loading buffer containing 0.1 M EDTA (pH 8.0), and the sample was applied to an SDS—10% PAGE gel.

For native PAGE experiments, PhoP proteins were phosphorylated with GST-*PhoR~P as described previously (9).

Determination of protein concentration. The protein concentration was determined by the Bradford method by using a Bio-Rad protein assay kit as instructed by the manufacturer.

SDS-PAGE and native PAGE. SDS-PAGE was performed as described by Laemmli (17). A 10% polyacrylamide separating gel was used for detection of His10-PhoP. An 8% native PAGE gel was prepared as described previously (9).

Quantitation of radioactivity. Radioactivity of proteins on SDS-PAGE or native PAGE gels was detected with Fuji medical X-ray film (Fuji) and/or a PhosphorImager (Molecular Dynamics). When required, the image was quantitated with Imagequant, version 5.1.

Gel shift assays. Gel shift assays were done as described previously (19). The probe was a 159-bp fragment containing the phoB promoter released from pRC696 (28) by digestion with BamHI and EcoRI and labeled with the Klenow enzyme in the presence of 4 µl of [{alpha}-32P]dATP (6,000 Ci/mmol; 10 mCi/ml).

In vitro transcription assay. All components used for the transcription reaction were adjusted to the required concentrations and equilibrated in nuclease-free transcription buffer (NFTB) (10 mM Tris-HCl [pH 8.0], 50 mM KCl, 5 mM MgCl2, 1 mM CaCl2, 0.1 mM EDTA, 5% glycerol), unless indicated otherwise. To construct a template for the in vitro transcription assay, the phoB promoter region (located at positions -221 to 105 relative to the translation start at position 1) was amplified by PCR by using primers FMH745 and FMH746 and strain JH642 chromosomal DNA as the template. The PCR product was extracted from a 1.2% agarose gel by using a gel extraction kit (Qiagen) and was further purified by phenol extraction and ethanol precipitation. Finally, the phoB promoter was dissolved in NFTB and stored at -20°C. B. subtilis {sigma}A and core RNAP were prepared as previously described (27). To prepare E{sigma}A, core RNAP was incubated with {sigma}A at a molar ratio of 1:30 in NFTB on ice for 30 min. For each reaction, addition of 5 µl of template DNA (40 ng/µl) was followed by addition of 1 µl of ATP (1 mM), 1 µl of *PhoR (4 µM), and 1 µl of each PhoP protein (wild type or mutant; concentration, 6 µM), and the mixture was incubated at room temperature for 20 min. Then 2 µl of E{sigma}A (0.5 µM core RNAP) was added to the reaction mixture, followed by 5 µl of a nucleoside triphosphate mixture (500 µM ATP, 500 µM GTP, 500 µM CTP, 50 µM UTP, 0.33 µM [{alpha}-32P]UTP [3,000 Ci/mmol; 10 µCi/µl], and 4 U of RNasin per µl in NFTB), and the mixture was incubated at room temperature for another 30 min. The reactions were stopped by adding 7.5 µl of stop buffer (7 M urea, 100 mM EDTA, 0.05% xylene cyanol, 0.05% bromophenol blue, 5% glycerol) and heating the preparations at 75°C for 5 min. After electrophoresis on a sequencing gel, the transcripts were detected by PhosphorImager analysis.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Experimental design and rationale. In this study, we set out to identify the amino acid residues of the PhoP C-terminal effector domain required for DNA binding or transcription activation, as well as to identify residues that may be involved in other functions, including protein stability or interdomain interactions in PhoP. To do this, we used the QuickChange method of site-directed mutagenesis to obtain single-alanine substitutions of 24 nonalanine residues of PhoP and one valine substitution of alanine, which constituted the putative transactivation loop and the DNA recognition helix. We introduced the mutations into the phoPR operon residing in plasmid pDH88 under control of the Pspac promoter, and the plasmids were transformed into the chromosome of a phoPR deletion strain via Campbell insertion (Fig. 1). In the resulting strains, expression of the phoPR operon was exclusively controlled by an inducer, IPTG.



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  		FIG. 1. Chromosomal structure of the phoPR locus in various B. subtilis strains: phoPR loci in wild-type strain JH642 (A), null phoPR strain MH6110 (B), and IPTG-inducible phoPR wild-type and amino acid substitution mutant strains (C). The phoPR genes or portions of the phoPR genes are indicated by thick solid arrows. Other chromosomal genes (mdh or polA) are indicated by thick gray arrows. The pDH88 vector genes are indicated by thick windowpane arrows. The Tetr insertion into the phoPR EcoRI deletion is indicated by a cross-hatched box. The structure of pCH102-127 used for Campbell insertion-duplication into MH6110 is also shown. The SacI and ClaI restriction enzyme sites, used for shuttling the internal phoPR gene region containing the {alpha} loop and {alpha} helix 3 between deltaPRKS+ and pCH102, are indicated by arrows. The bent arrows indicate promoters.

 
Qualitative in vivo assessment of the effect of each PhoP mutation on PhoP function. The 36-fold increase in total APase activity upon phosphate starvation represented the sum of the effects on three APases, PhoA, PhoB, and PhoD, which account for approximately 65, 30, and <5% of the total activity, respectively. The promoter-lacZ fusion of the major APase, PhoA, was inserted at the amyE locus in our constructs and used as a reporter of PhoP~P transcriptional activity. Fresh transformants of each PhoP variant were picked and transferred to a TBABG plate and incubated for 16 h at 37°C. Then the single clones were grown on HPM or LPM with IPTG and X-Gal to assess the relative promoter activities. The colony colors were scored after incubation of the plates at 37°C for 16 h. Parent strain MH6110 ({Delta}phoPR) and strain MH6111 with an IPTG-inducible wild-type phoPR operon were included as phoA-lacZ controls. Figure 2A shows that under phosphate-sufficient conditions (HPM), neither the wild-type strain nor the PhoP mutant strains activated phoA promoter expression. Under phosphate starvation conditions (LPM), the expression in strains having substitutions at PhoP residues 192, 194, 196, 198, 201, 207, 212, and 213 was similar to the phoA-lacZ expression (colony color score) in the strain with the wild-type phoPR operon (MH6111), suggesting that substitutions at these residues had no effect on PhoP function. However, strains having substitutions at residues 191, 193, 195, 199, 200, 202, 203, 206, 208, 209, and 210 showed no phoA-lacZ activity, indicating that each of these substitutions was deleterious to the PhoP function. Strains having the remaining PhoP substitutions exhibited decreased phoA-lacZ activity, indicating that these residues may affect the PhoP function.



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  		FIG. 2. Assessment of the effect of each phoP mutation on PhoP function. (A) phoA-lacZ fusion plate assay under Pi-replete (HPM) or phosphate starvation (LPM) conditions. One hundred clones of strains containing a phoPR deletion, phoPWT, or a phoP mutation (affecting residues 190 to 214) were grown on HPM or LPM containing X-Gal and IPTG. Colony color was scored after 16 h. ++, blue, similar to PhoPWT strain color; -, white, similar to {Delta}phoPR strain color; +, light blue, between PhoPWT strain blue color and white. (B) Effect of each amino acid substitution in PhoP on APase specific activity under phosphate starvation conditions. The APase specific activities are averages of the values from 12 clones for each construct. The error bars indicate standard deviations. OD540, optical density at 540 nm; WT, wild type.

 
Quantitative in vivo assessment of the effect of each PhoP mutation on Pho induction. To assess the Pho induction phenotype of each phoP mutation further, total APase specific activity was measured for each strain under Pi starvation conditions. Previous data showed that in strain MH6101 expressing the phoPR operon constitutively from the Pspac promoter, the APase specific activity increased sharply upon phosphate starvation and reached a maximal value that was maintained as cells entered the stationary phase under phosphate starvation conditions (9). We took advantage of the plateau in total APase specific activity to compare the total APase specific activities exhibited by the strains. Samples were taken from cultures grown in LPDM at 12 to 14 h, when the total APase specific activity remained constant. In agreement with the plate assay data, strains having mutations that changed PhoP residues 192, 194, 196, 198, 201, 207, and 212 had levels of APase specific activity similar to those of strains with the wild-type phoPR operon (Fig. 2B), while strains having substitution mutations that affected residues 191, 193, 195, 199, 200, 202, 203, 204, 205, 206, 208, 209, and 210 each exhibited a very low level of APase specific activity, which was comparable to the level of the parental strain (MH6110 {Delta}phoPR::Tetr phoA-lacZ). The remaining mutant strains had reduced total APase specific activities, and the levels were less than 50% of the level of the wild-type phoPR operon strain (MH6111 Pspac phoPR). Discrepancies in the qualitative and quantitative expression assays were observed for strains containing K212, V204, and H205 PhoP variants.

Similar PhoP concentrations (wild-type or mutant proteins) were detected in the soluble fraction of strains with phoPR expression controlled by IPTG. Because the PhoP protein concentration in cells may affect phoA-lacZ and/or total APase expression during Pho induction, anti-PhoPC antiserum was used to immunodetect PhoP protein in the cells. Cells were grown in LPDM containing IPTG until the culture entered stationary growth due to limiting phosphate concentrations and induction of APase had plateaued. Cells were harvested, and the proteins were extracted as described in Materials and Methods. Soluble proteins (12 µg) from each phoP mutant strain and the strain containing the wild type phoPR operon under Pspac control (MH6111) were separated on an SDS-PAGE gel, blotted, and immunodetected with anti-PhoPC antiserum (Fig. 3A). The position on the PhoP standard curve of PhoP in 12 µg of soluble proteins from MH6111 is indicated in Fig. 3B. The concentrations of PhoP proteins induced with IPTG in strains having phoP mutations were similar (within 5%) to the concentration of wild-type PhoP in MH6111. The same results were obtained when lysed cell samples were analyzed similarly (data not shown). Thus, the variability in the phoA-lacZ activities and/or the total APase activities (Fig. 2) was used to identify which residue changes affected PhoP function.



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  		FIG. 3. Western immunoblot detection of PhoP proteins from B. subtilis strains. (A) The cells were grown for 11 h and collected by centrifugation, and soluble proteins were extracted as described in Material and Methods. The same amount (12 µg) of protein for each sample was separated by SDS-PAGE, transferred onto a polyvinylidene fluoride membrane, and immunodetected by using anti-PhoPC polyclonal sera. The arrow indicates the migration position of purified PhoPWT. The samples from strains expressing wild-type PhoP and individual amino acid substitutions of PhoP are indicated below the lanes. PhoPWT and PhoP variant densities were measured by ImageQuant. The densities of PhoP variants were within 5% of the PhoPWT density. (B) Quantification of purified PhoP protein in 12 µg of soluble protein from MH5913 ({Delta}phoPR). The polyvinylidene difluoride membranes were scanned, and the PhoP protein densities were measured by ImageQuant; the results are expressed in arbitrary units. The error bars indicate the standard deviations for four samples. The arrow indicates the average of density for four samples of 12 µg of soluble protein of MH6111 (PhoPWT) extracted as described above. The error range for the four PhoPWT samples was ±5%. WT, wild type.

 
In vitro phosphotransfer reactions were not altered in most PhoP mutated proteins, and three of these proteins exhibited a reduced phosphotransfer rate. To understand how PhoP function was affected by each residue change, PhoP mutant proteins were overexpressed in E. coli and purified by Ni-nitrilotriacetic acid affinity chromatography so that they were >95% pure, as judged by SDS-PAGE (data not shown). The ability to receive a phosphoryl group from the cognate histidine kinase PhoR was analyzed by using the functional N-terminal truncated form of PhoR, *PhoR (30). *PhoR~P, free of ATP, was used for phosphotransfer to each mutated PhoP. Samples were taken after 10 and 60 s and 10 min for analysis of phosphotransfer. In general, phosphoryl transfer from *PhoR~P to most of the mutant Pho proteins was maximal within 10 s and remained stable, and there was only a slight loss of ~P over time, which is typical of the wild-type PhoP protein (Fig. 4). However, two Pho mutants, mutants 202 and 203, exhibited reduced phosphotransfer rates from *PhoR~P; there was only 20% of the wild-type PhoP labeling after 10 s, but the level increased to nearly 80% of the wild-type labeling after 10 min. These two mutants identified residues located in the C-terminal domain that might cause a defect in phosphotransfer via domain-domain interaction. Howerver, most phoP variants behaved like the wild-type PhoP in phosphotransfer reactions, suggesting that the mutant proteins retained the overall structure of the wild-type PhoP.



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  		FIG. 4. Phosphotransfer rate from *PhoR~P to each PhoP. GST-*PhoR was phosphorylated by incubation with [{gamma}-32P]ATP and bound to glutathione-agarose. After excess [{gamma}-32P]ATP was removed, 30 U of thrombin was added to the beads and mixed at room temperature for 20 min. The *PhoR~P was separated from the beads and mixed with each PhoP variant at a molar ratio of 1:6.6. Equal-volumes samples were removed from each reaction mixture at three different times, and the reaction was stopped with SDS loading buffer. The samples were subjected to SDS-PAGE, the gels were dried, and the radioactivity was detected with a PhosphorImager. Lanes 1, 10 s; lanes 2, 60 s; lanes 3, 600 s. WT, wild type.

 
PhoP changes that affect DNA binding. To identify residues that might be directly involved in the protein-DNA interaction, PhoP-promoter DNA complexes were detected by a gel mobility assay. The phoB promoter was chosen as the target because it is the simplest of the well-characterized Pho regulon promoters, with PhoP binding solely to the core binding region between approximately positions-20 and -60 upstream of the Pho-regulated promoter. Previous data had shown that PhoP or PhoP~P can bind to this promoter, although lower concentrations of PhoP~P (almost 25% the concentration of PhoP) were needed to bind (19). Using PhoPWT, we confirmed that 5 µM PhoP was needed to form the protein-DNA complex, while 1.5 µM PhoP~P was sufficient (data not shown). PhoPWT- or mutant PhoP-DNA complexes were detected by native gel electrophoresis under the same conditions (Fig. 5). Variants with mutations that created residue substitutions at positions 202, 203, 204, 205, 206, 208, 209, and 210 did not form complexes at the concentrations used, while other PhoP variants were similar to PhoPWT. These residues may be involved in direct PhoP-DNA interactions.



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  		FIG. 5. Gel shift assays of the 159-bp phoB promoter bound by PhoP or each mutant PhoP. The amino acid substitution residues are indicated above the gels. Lane F, no PhoP in the reaction mixture; lanes 1, 1.38 µM PhoP in the reaction mixture; lanes 2, 2.76 µM PhoP in the reaction mixture; lanes 3, 5.51 µM PhoP in the reaction mixture. WT, wild type.

 
The fact that phosphorylation of PhoP in vitro increased the DNA binding efficiency in vitro (19) and the fact that in vivo a PhoR deletion strain did not express APase activity indicated that Pho induction required phosphorylation of the response regulator, PhoP. When the phosphorylated form of PhoP was used to assess DNA binding, as expected, most complexes formed at a concentration that was approximately 25% of the required PhoP concentration (Fig. 6). Surprisingly, phosphorylated forms of PhoP with residue substitutions at positions 204 and 205 formed complexes at PhoPWT concentrations, while nonphosphorylated forms of PhoP with residue substitutions at position 204 or 205 failed to bind DNA (Fig. 5). Taken together, the results of the gel shift assay suggest that residues 202, 203, 204, 205, 206, 208, 209, and 210 might be involved in protein-DNA interactions. Phosphorylation of PhoP proteins with substituted residues at positions 204 or 205 may affect protein-DNA formation via a mechanism that differs from the mechanism for other PhoP proteins.



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  		FIG. 6. Gel shift assays of the 159-bp phoB promoter bound by PhoP~P or each mutant PhoP~P. The 159-bp phoB promoter was incubated with PhoP or each mutant PhoP and GST-*PhoR in the presence of ATP. The amino acid substitution residues are indicated above the gels. Lane F, no PhoP in the reaction mixture; lanes 1, 0.368 µM PhoP in the reaction mixture; lanes 2, 0.735 µM PhoP in the reaction mixture; lanes 3, 1.47 µM PhoP in the reaction mixture. Each reaction mixture contained 2 µM GST-*PhoR. WT, wild type.

 
It has been reported that PhoP mutant proteins that cannot dimerize are defective in binding to target DNA (9). Native gel electrophoresis of nonphosphorylated or phosphorylated mutant proteins, the native PhoP protein (dimer), and PhoPR113E (monomer) was used to assess the possibility that dimerization was affected by the residue changes between I201 and K212 (Fig. 7). The phosphorylated or nonphoshorylated PhoP variants migrated at the rate of wild-type PhoP, not the higher rate of PhoPR113E, suggesting that dimerization was not affected and therefore not the reason for a DNA binding deficiency.



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  		FIG. 7. Native gel migration of PhoP variants, PhoPWT (dimer), and PhoPR113E (monomer): Coomassie blue staining (A) and radioactivity (B) of native PAGE gel. The positions of GST-*PhoR, PhoPWT, and PhoPR113E are indicated on the right. The amino acid residue substitutions are indicated at the bottom. GST-*PhoR and each PhoP protein were mixed with (+) or without (-) 10 µCi of [{gamma}-32P]ATP in phosphorylation buffer as described in Materials and Methods. After incubation at room temperature for 20 min, 0.25 volume of 40% glycerol-100 mM EDTA was added to each mixture, and the mixture was loaded onto the native PAGE gel. WT, wild type.

 
In vitro transcription assays with E{sigma}A resulted in identification of PhoP residue changes that affect transcriptional activity and potential RNAP interaction residues. To test the transcriptional activity of each PhoP mutant protein, runoff in vitro transcription assays were conducted with the phoB promoter as the template. Previous primer extension data showed that the phoB promoter has two transcriptional start sites, Pv and Ps (10), which have recently been shown to be dependent on {sigma}A and {sigma}E, respectively (W. R. Abdel-Fattah, Y. Chen, and F. M. Hulett, unpublished data). Only the E{sigma}A promoter is PhoP regulated. To simplify the reaction, only transcription from the Pv promoter was tested. B. subtilis core RNAP was purified and mixed with B. subtilis {sigma}A factor that had been overexpressed in and purified from E. coli to reconstitute active {sigma}A holo-RNAP. As expected, PhoP DNA binding-deficient substitutions at positions 202, 203, 206, 209, and 210 did not activate transcription at all, while a substitution at position 208 resulted in a decreased level of transcript (perhaps 25% that of PhoPWT). In contrast, although the PhoP proteins with residue substitutions at positions 190, 191, 193, 195, 197, 199, and 200 bound DNA efficiently, they activated transcription at moderately to severely reduced levels. PhoP proteins with substituted residues at position 191 or 200 yielded no detectable transcripts at all. Considering the in vivo (Fig. 2) and in vitro (Fig. 8) data together, residues 191 and 200 are probably two residues that directly interact with RNAP, while residues 190, 193, 195, 197, and 199 may contribute to this interaction.



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  		FIG. 8. Transcription activation of the phoB Pv promoter by wild-type PhoP~P or mutant PhoP~P proteins having amino acid substitutions. The position of the transcripts (142 bp) from the Pv promoter is indicated by an arrow. WT, wild type.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Functional analysis of PhoP mutant proteins derived from alanine scanning mutagenesis of the putative PhoPC transactivation loop ({alpha} loop) and DNA binding helix ({alpha} helix 3) showed that mutations involving certain amino acid residues between PhoPV190 and PhoPE214 were essential or important for PhoP function in vivo. In vitro functional analysis revealed that all mutated proteins defective for DNA binding had residue changes between PhoPI202 and PhoPE210, while mutations involving certain residues between PhoPV190 and PhoPI200 produced proteins that had reduced PhoP function in vivo and were unable or had reduced ability to activate transcription in vitro (Fig. 9).



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  		FIG. 9. Comparison of the amino acid sequences of the C-terminal effector domains of PhoP with those of PhoB and OmpR. The numbers refer to the residues of PhoP. The secondary structure elements of PhoB (7) and OmpR (21) are indicated above and below the sequences, respectively. Residues 190 to 214 of PhoP used for alanine mutagenesis are indicated in columns, and within this region the residues conserved in three proteins and in two proteins are indicated by red and blue, respectively. PhoB residues involved in the RNAP-{sigma}70 interaction and DNA binding (20) are indicated by red diamonds and red circles, respectively, and residues structurally predicted to contact DNA are indicated by green circles (7). PhoP residues that are essential for transcriptional activation (interaction with RNAP) are indicated by black diamonds, and residues predicted have a role in the RNAP interaction are indicated by open diamonds. Three classes (classes 1, 2, and 3) (see text for details) of PhoP residues involved in DNA binding are indicated by black circles, black circles with lines, and open circles, respectively. OmpR residues involved in the RNAP-{alpha} subunit interaction (25, 29) and DNA binding (1, 16, 29) are indicated by blue diamonds and blue circles, respectively.

 
Distribution of PhoP residues involved in DNA binding or transactivation appears more similar to the distribution determined for OmpR than to the distribution determined for PhoB of E. coli. The results of our analysis of mutations throughout the PhoP DNA binding-transactivation domain can be compared to the results of a mutational analysis and amino acid alignment of the domains of two structurally characterized E. coli response regulators, OmpR (21) and PhoB (7) (Fig. 8). The level of sequence identity within the DNA binding region of the three proteins is quite high, 47%, whereas the transactivation loop has no conserved residues in the three proteins. The eight mutations between V190 and E214 in PhoP that affect DNA binding are clustered between V202 and R210 in {alpha} helix 3, whereas the mutations affecting DNA binding of PhoB (20) or OmpR (1, 16, 29) are more broadly spread throughout the corresponding region, especially in PhoB, in which mutations in the {alpha} loop were shown to specifically affect DNA binding (20) and in which structural analysis (7) showed that transactivation loop residues Val183, Glu191, and R193 bind DNA. In contrast to mutations that affect PhoP DNA binding, mutations in PhoP that affect the transcription activation function of PhoP were broadly distributed throughout the transactivation loop, extending as far C terminal as R200, a conserved residue in the three proteins that is part of an extended {alpha} helix 3 in PhoB but is in the {alpha} loop of OmpR.

PhoP DNA binding mutants fall into three classes. One class of PhoP DNA binding mutants, which included PhoPI206A, PhoPR208A, PhoPL209A, and PhoPH210A, had a phosphotransfer rate similar to that of the native PhoP and could dimerize, but neither the phosphorylated nor nonphosphorylated PhoP mutants could bind DNA at concentrations appropriate for PhoPWT. In general, the members of this class could not activate transcription of the phoB promoter in vitro or of total APases or phoA in vivo; the only exception was PhoPH208A, which produced a weak transcript in vitro but exhibited no transcription in vivo. Higher concentrations of PhoP and/or E{sigma}A in vitro than in vivo may account for the detectable transcriptional activity of PhoPH208A observed in vitro. The phenotype of the mutant proteins in this class is consistent with residues that may directly interact with DNA. PhoB (E. coli) residues that correspond to PhoPR210 bind DNA via nonspecific interactions, while the residue corresponding to PhoPH208 binds via both specific and nonspecific interactions (7). In addition, especially for PhoPI206A, PhoPR210A, and PhoPL209A, the phenotype is also consistent with disruption of conserved residues that stabilize secondary structural elements involved in DNA recognition, a role determined for the analogous residues of the E. coli ortholog, PhoB (7).

A second class of PhoP DNA binding mutants, including PhoPV204A and PhoPH205A, also had a phosphotransfer rate equal to that of the native PhoP and could dimerize, and the nonphosphorylated PhoP mutant proteins could not bind DNA at concentrations appropriate for nonphosphorylated PhoPWT; however, the phosphorylated PhoPV204A and PhoPH205A proteins could. Transcription of phoA and the total APase activity in strains with these mutant phoP alleles were quite low compared to those of the wild-type strain, but in vitro phoB transcription by using PhoPH205A~Por PhoPV204A~P was more than 50% of the transcription observed when PhoPWT was used. These data suggest that unlike the native PhoP (19), phosphorylation of PhoPV204A and PhoPH205A is required to change the conformation of the mutant protein to allow DNA binding (compare Fig. 5 and 6) required for transcriptional activity, perhaps in a way similar to either that proposed for NarL (4) and CheB (11), in which functional regions of the effector domains are blocked by the regulatory domains in the nonphosphorylated state, or that proposed for PhoB (7) of E. coli, in which it has been proposed that the position of the DNA binding region in the nonphosphorylated state prevents binding of the response regulator to the tandem repeats of the Pho box. Both structural and genetic studies (2) implicated helix {alpha}5 of the receiver domain in the interdomain interactions in PhoB negative regulation. The phosphorylation-dependent phenotype for DNA binding of PhoPH205A or PhoPV204A implies that a mutation in the C-terminal DNA binding region of PhoP may alter the interaction of the domain with the PhoP N-terminal domain. A mutation in OmpR, OmpRV203 M, at the position analogous toV204A in PhoP, reduced DNA binding (29), and the protein exhibited functional properties that affected both DNA recognition and small-molecule phosphorylation of OmpR, leading to the conclusion that a single mutation in the OmpR DNA binding domain affects the N-terminal domain (31). The E. coli PhoB residue corresponding to PhoPV204 showed specific DNA interactions, while residues corresponding to PhoPH205 had both specific and nonspecific DNA binding interactions, as well as a structural role via hydrogen bonding to a conserved Glu residue of {alpha} helix 1 of the DNA binding domain (7).

A third class of PhoP DNA binding mutants, including PhoPV202A and PhoPD203A, have a reduced rate of phosphotransfer, can dimerize, and do not bind DNA or activate transcription in vitro or in vivo. The reduced rate of phosphotransfer may again indicate that mutations in this DNA binding class affect the PhoPN structure in addition to the C-terminal domain structure. Residues of PhoB (7) or OmpR (22) corresponding to PhoPV202 are residues of the central hydrophobic core that maintain the tertiary structure of that domain. Residues of PhoB and OmpR analogous to PhoPD203 also link secondary structural elements by hydrogen bonding to the invariant tyrosine in the C-terminal hairpin that corresponds to PhoPY231. All mutant PhoP proteins in this class were hypersensitive compared to native PhoP in limited proteolysis experiments, which is consistent with the importance of these residues in stabilization of the native PhoP conformation (data not shown).

Certain mutant PhoP proteins involving PhoP residues V190 to I201 are affected in in vivo and in vitro transcription. None of the PhoPV190 to PhoPI201 mutant proteins had phosphorylation or DNA binding defects, although a number had transcription defects. The PhoPW191A and PhoPR200A mutant proteins could not activate in vivo or in vitro transcription, while PhoPY193A and PhoPF195A abolished in vivo transcription but supported very reduced in vitro transcription compared to that of PhoPWT. PhoPV190A, PhoPG197A, and PhoPT199A showed reduced transcription in vivo or in vitro or both compared to the transcription of PhoPWT. These data indicate that PhoP residues V190, W191, Y193, F195, G197, T199, and R200 are candidates for RNAP interaction. PhoPW191A is conserved in E. coli PhoB, in which it has been shown to interact with the {sigma}70 subunit of RNAP (20). PhoPR200A is also conserved in PhoB, in which it is implicated in DNA binding but not in RNAP interaction.

PhoP variants that are DNA binding proficient but are defective in the RNAP interaction have Ala substitutions that are broadly distributed throughout 11 residues, from V190 to R200 (Fig. 9). Since residues in the response regulator that contribute to the RNAP interaction generally belong to the transactivation loop, secondary structure considerations suggest that PhoP has a longer transactivation loop and is more similar to OmpR than to PhoB of E. coli. Mutations in PhoP that affect DNA binding (eight of nine contiguous residues) are in the highly conserved sequence that is part of {alpha} helix 3 in either PhoB or OmpR. While OmpR mutations that affect DNA binding are confined to {alpha} helix 3, PhoB residues in both the {alpha} loop and {alpha} helix 3 have been shown by mutagenesis and/or PhoB DNA structure analysis to be important for binding. It is of interest to determine the relevance of these observations with respect to PhoPC structure and the PhoP-RNAP interaction.


   ACKNOWLEDGMENTS
 
This work was supported by National Institutes of Health grant GM-33471 to F.M.H.

We thank J.-P. Samama for helpful discussions and suggestions.


   FOOTNOTES
 
* Corresponding author. Mailing address: Laboratory for Molecular Biology, Department of Biological Sciences, University of Illinois at Chicago, 900 S. Ashland Avenue (M/C 567), Chicago, IL 60607. Phone: (312) 996-5460. Fax: (312) 413-2691. E-mail: Hulett{at}uic.edu. Back


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Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Journal of Bacteriology, March 2004, p. 1493-1502, Vol. 186, No. 5
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.5.1493-1502.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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