1 Department of Microbiology and Immunology, McGill University, Montréal, Québec, Canada H3A 2B4
2 ProteinProtein Interaction Facility, Sheldon Biotechnology Centre, McGill University, Montréal, Québec, Canada H3A 2B4
3 Faculty of Dentistry, McGill University, Montréal, Québec, Canada H3A 2B4
Correspondence
Hervé Le Moual
herve.le-moual{at}mcgill.ca
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ABSTRACT |
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Present address: Département de Génie Chimique, École Polytechnique de Montréal, Boîte Postale 6079, Station Centre-Ville, Montréal, Québec, Canada H3C 3A7.
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INTRODUCTION |
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Response regulators are classified into the OmpR/PhoB, NarL/FixJ and Ntrc/DctD subfamilies based on sequence similarity within the C-terminal effector domain (Stock et al., 1989). In members of the OmpR/PhoB subfamily, the C-terminal domain consists of a winged helixturnhelix motif (Kenney, 2002
). Although all members of this subfamily share a similar three-dimensional structure and are activated through phosphorylation, they appear to vary in their mechanism of activation. Phosphorylation of Escherichia coli PhoB has been shown to induce dimerization and, in turn, increase its affinity for DNA (Fiedler & Weiss, 1995
; McCleary, 1996
). In contrast, phosphorylation of OmpR was found to enhance its DNA-binding affinity without promoting dimerization of the protein in solution, suggesting that dimerization may occur upon DNA interaction (Aiba et al., 1989
; Jo et al., 1986
). The PhoPBSU response regulator of the Bacillus subtilis PhoP/PhoR two-component system (designated PhoPBSU to distinguish it from PhoP of the Salmonella enterica PhoP/PhoQ system) has been shown to be dimeric and bind target DNA independently of its phosphorylation state (Liu & Hulett, 1997
; Prágai et al., 2004
).
In S. enterica, PhoP is the response regulator of the PhoP/PhoQ two-component system, which responds to environmental Mg2+ (Garcia-Vescovi et al., 1996). It is another member of the OmpR/PhoB subfamily. It modulates the expression of more than 40 genes involved in adaptation to Mg2+-limiting environments, survival within macrophages, LPS modifications and resistance to antimicrobial peptides (Ernst et al., 2001
; Groisman, 2001
). Although some genes regulated by PhoP do not display a consensus DNA recognition sequence for PhoP, other PhoP-regulated genes, such as the mgtA gene, contain a single conserved PhoP box in their promoter region (Lejona et al., 2003
). This PhoP box is located approximately 30 bases upstream of the transcription start site and consists of a direct repeat of the heptanucleotide (G/T)GTTTA(A/T) (Groisman et al., 1989
; Lejona et al., 2003
). To date, the E. coli and S. enterica PhoP proteins that have been characterized in vitro harboured a C-terminal His tag (PhoPHis) (Lejona et al., 2003
, 2004
; Minagawa et al., 2003
; Yamamoto et al., 2002
). These studies led to a model in which phosphorylation promotes PhoP dimerization and, in turn, increases PhoP binding to the PhoP box.
In this study, we examined the effect of phosphorylation on the oligomeric state and DNA binding properties of the untagged S. enterica PhoP (PhoP). To compare our results with previous studies, we conducted similar experiments with the PhoP variant harbouring a C-terminal His tag (PhoPHis). Our data showed that phosphorylation of PhoP has no major effect on dimerization and DNA interaction. In contrast, we found that phosphorylation of PhoPHis promotes both dimerization and binding to DNA.
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METHODS |
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The DNA fragment corresponding to the PhoQ cytoplasmic domain (Arg 252Glu 487) was PCR amplified from plasmid pET-Q (Montagne et al., 2001) using primers PHOQCYTO-5' (5'-GTCAGAATTCCGCAACCTTAATCAACTGCTCAAAAG-3') and PHOQCYTO-3' (5'-CTGACTCGAGTTATTCCTCTTTCTGTGTGGGATGC-3'). PCR products were purified, digested with EcoRI and XhoI, and inserted into plasmid pGEX-4T1 (Amersham Biosciences) digested with the same enzymes, to generate plasmid pGEX-Qcyto.
Overexpression and purification of recombinant proteins.
For purification of the PhoP protein, E. coli BL21(DE3)/pLysE cells transformed with the pET-P plasmid were grown at 22 °C in LuriaBertani broth supplemented with ampicillin (100 µg ml1) and chloramphenicol (30 µg ml1). At OD600 0·8, transcription of the phoP gene was induced overnight with 1 mM IPTG. Cells were harvested by centrifugation and resuspended in buffer A (25 mM NaPO4, pH 6·0). After lysis by ultrasonic disruption, cell debris was removed by centrifugation at 216 000 g for 30 min at 4 °C. The supernatant was applied to a 1 ml Resource Q column (Amersham Biosciences) equilibrated with buffer A. After a 15 ml wash with buffer A, proteins were eluted by applying a 01·0 M NaCl gradient in the same buffer, at a flow rate of 1·5 ml min1. Fractions enriched in PhoP were pooled and dialysed overnight against buffer B (25 mM NaPO4, pH 7·0). The dialysed proteins were then applied to a 1 ml HiTrap Heparin HP column (Amersham Biosciences) equilibrated with buffer B. After a 15 ml wash with buffer B, proteins were eluted by applying a 01·0 M NaCl gradient in the same buffer. The PhoPHis protein was overexpressed as described above for PhoP, and purified by Ni2+-NTA (nitrilotriacetic acid) chromatography as described previously (Montagne et al., 2001). Both purified PhoP and PhoPHis were dialysed overnight against buffer C (20 mM Tris/HCl, pH 7·9, 50 mM NaCl). Proteins were kept at 4 °C and used within 4 days.
For purification of the PhoQ cytoplasmic domain fused to the glutathione S-transferase (GST) protein (GST-PhoQcyto), E. coli XL1-Blue cells transformed with the pGEX-Qcyto plasmid were grown at 37 °C in LB broth supplemented with ampicillin (100 µg ml1). At OD600 0·8, cultures were induced with 1 mM IPTG for 4 h. Cells were harvested and lysed as described above. The supernatant was loaded on a 1 ml GSTrap FF column (Amersham Biosciences) equilibrated in buffer D (20 mM Tris/HCl, pH 7·9, 200 mM NaCl). After extensive wash with buffer D, proteins were eluted with the same buffer supplemented with 10 mM glutathione. Fractions containing GST-PhoQcyto were pooled and dialysed overnight against buffer C. Protein concentrations were determined with the BCA protein assay kit (Pierce), using dilutions of BSA as standards.
Phosphorylation of PhoP and PhoPHis.
The purified PhoP or PhoPHis protein was phosphorylated with either the GST-PhoQcyto protein or E. coli membranes containing the overexpressed PhoQ-T48I protein. E. coli membranes enriched in the PhoQ-T48I protein were prepared as described previously (Sanowar et al., 2003). Reactions were performed in the presence of 11·5 mM ATP to maximize the efficiency of phosphorylation. For gel filtration experiments, PhoP or PhoPHis (10 µM) was phosphorylated with GST-PhoQcyto (2 µM) in a 2 ml volume of phosphorylation buffer (50 mM Tris/HCl, pH 7·5, 200 mM KCl, 0·1 mM EDTA, 5 % v/v glycerol) supplemented with 5 mM MgCl2. Reactions were initiated by the addition of 1 mM ATP and incubated for 1 h at 22 °C. Unphosphorylated PhoP or PhoPHis was subjected to the same procedure except that H2O was substituted for ATP. For fluorescence spectroscopy and surface plasmon resonance (SPR) experiments, PhoP or PhoPHis (15 µM) was incubated for 20 min at 22 °C with E. coli membranes containing the overexpressed PhoQ-T48I protein (approx. 1·5 µM) in phosphorylation buffer supplemented with 5 mM MgCl2 and 1·5 mM ATP. Reactions were stopped by the addition of EDTA to a final concentration of 10 mM. PhoP
P or PhoPHis
P was separated from the PhoQ-containing membranes by centrifugation at 216 000 g for 10 min at 4 °C. Unphosphorylated PhoP proteins were subjected to the same procedure except that H2O was substituted for ATP. For SPR experiments, the PhoP proteins were dialysed overnight at 4 °C against SPR buffer (10 mM HEPES, pH 7·4, 150 mM KCl, 10 mM EDTA, 0·005 % v/v Tween 20). The presence of EDTA in the SPR buffer prevented the autodephosphorylation of PhoP
P or PhoPHis
P.
To assess the extent of PhoP phosphorylation, reactions were performed with 1 mM [-32P]ATP [10 Ci mmole1 (37x1010 Bq mmole1)] and stopped by the addition of 4x Laemmli SDS sample buffer (250 mM Tris/HCl, pH 6·8, 8 % SDS, 40 % (v/v) glycerol, 0·02 % bromophenol blue, 4 %
-mercaptoethanol). Reaction products were heated at 37 °C for 5 min and applied to 10 % SDS-PAGE gels. Gels were dried and exposed to a PhosphorImager (Bio-Rad). The band corresponding to PhoP
P or PhoPHis
P was quantified using Quantity One software (Bio-Rad). This value was then compared to a standard curve generated by using known concentrations of [
-32P]ATP (5, 25, 50, 75 and 100 pM). The efficiency of the phosphorylation reaction was calculated using the ratio of 1 radiolabelled/100 000 non-radiolabelled phosphoryl groups.
Gel filtration chromatography.
Samples (2 ml) of PhoP, PhoPP, PhoPHis or PhoPHis
P (10 µM solution) were individually applied to a Superdex 200 HR 10/30 gel filtration column (Amersham Biosciences) equilibrated with running buffer (25 mM Tris/HCl, pH 7·5, 150 mM NaCl). Proteins were eluted in the same buffer at a flow rate of 0·5 ml min1 and collected in 1 ml fractions. Protein concentration was monitored by measuring the OD280 and fractions were analysed by SDS-PAGE. Blue dextran 2000 was used to determine the column void volume. A mixture of protein molecular mass standards, containing
-amylase (200 kDa), alcohol dehydrogenase (150 kDa), BSA (66 kDa), carbonic anhydrase (29 kDa) and cytochrome C (12·4 kDa), was applied to the column under similar conditions. The elution volumes and molecular masses of the protein standards were used to generate a standard curve from which we determined the apparent molecular mass of the various PhoP proteins.
Fluorescence spectroscopy.
Fluorescence spectra were obtained at 22 °C using a Varian Cary Eclipse fluorescence spectrophotometer. Phosphorylated and unphosphorylated proteins treated with 10 mM EDTA were diluted to the desired concentration using a buffer containing 20 mM Tris/HCl, pH 7·9, 50 mM NaCl. Each spectrum was the mean of four consecutive scans. All spectra were corrected by subtracting the blank spectrum corresponding to buffer alone. For intrinsic fluorescence, protein samples (4 µM) were excited at 295 nm with a slit width of 5 nm. Emission spectra were collected from 300 to 450 nm with a slit width of 5 nm. 1-Anilinonaphthalene-8-sulfonic acid (ANS), a fluorescent hydrophobic dye, was purchased from Fluka. For binding experiments, ANS (500 µM) was incubated with the various PhoP proteins (1·6 µM) in the absence or presence of 7·0 M urea for 30 min at 22 °C in the dark. Following incubation, spectra were recorded by exciting the samples at 372 nm, and measuring emission over the range of 400 to 600 nm using 10 nm slit widths for both excitation and emission.
Surface plasmon resonance.
SPR measurements were performed on a Biacore 2000 using a streptavidin sensor chip (Biacore AB). Each surface of the chip was conditioned as recommended by the manufacturer, and equilibrated with SPR buffer (10 mM HEPES, pH 7·4, 150 mM KCl, 10 mM EDTA, 0·005 % v/v Tween 20). The PhoP box of the mgtA promoter was prepared by annealing a 5'-biotinylated oligonucleotide (5'-biotin-GGGGTCTGGTTTATCGTTGGTTTAATTGGGG-3'; underlined nucleotides indicate the PhoP box) to a non-biotinylated complementary oligonucleotide (5'-CCCCAATTAAACCAACGATAAACCAGACCCC-3'). Equimolar amounts of both oligonucleotides were mixed, heated at 95 °C for 5 min and slowly cooled to room temperature. Similarly, a DNA duplex containing a randomized PhoP box was prepared by annealing the oligonucleotide 5'-biotin-GGGGTCTACGCTCTCGTTAACCCTATTGGGG-3' (underlined nucleotides indicate the randomized PhoP box) to a complementary oligonucleotide (5'-CCCCAATAGGGTTAACGAGAGCGTAGACCCC-3'). Biotinylated DNA duplexes were diluted to 20 nM in SPR buffer and injected at a flow rate of 5 µl min1 until approximately 700 resonance units (RUs) were immobilized on the chip. Measurements were performed at 22 °C at a flow rate of 25 µl min1. Phosphorylated and unphosphorylated proteins were diluted in SPR buffer, containing 10 mM EDTA, prior to injection. Following two 75 µl injections of SPR buffer, PhoP, PhoPP, PhoPHis or PhoPHis
P (0·1, 0·3 and 1 µM) was injected over the sensor chip surfaces on which the DNA duplexes had been immobilized. Following an injection of 180 s, the kinetics of dissociation were monitored by injecting SPR buffer for 300 s. The sensor surfaces were regenerated by performing two 50 µl injections of 3 M NaCl (100 µl min1) followed by an EXTRACLEAN procedure as recommended by the manufacturer. All measurements were performed within 24 h following phosphorylation. Specific binding was calculated by subtracting the sensorgrams obtained with the randomized PhoP box from the sensorgrams obtained with the mgtA PhoP box.
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RESULTS |
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Most response regulators possess an intrinsic autophosphatase activity that is Mg2+-dependent. The half-life of PhoPHisP has been reported to be approximately 60 min (Castelli et al., 2000
). To increase the stability of PhoP
P and PhoPHis
P, EDTA was added to a final concentration of 10 mM after the phosphorylation reaction. As shown in Fig. 1(c)
, the presence of EDTA prevented PhoP
P dephosphorylation for at least 24 h. Similar results were obtained with PhoPHis
P (data not shown).
Effect of phosphorylation on oligomerization of PhoP and PhoPHis
To determine the oligomeric state of PhoP, PhoPP, PhoPHis and PhoPHis
P, these proteins were subjected to gel filtration chromatography on a Superdex 200 column that was calibrated with molecular mass standards. PhoP and PhoP
P phosphorylated by GST-PhoQcyto were individually applied to the gel filtration column. PhoP and PhoP
P eluted as two peaks (Fig. 2a, b
). The earlier- (60 ml) and later-eluted (71 ml) peaks corresponded to masses of 55 and 24 kDa, respectively. These values are consistent with PhoP being in the monomeric (24 kDa) and dimeric (55 kDa) states, and suggest that both PhoP and PhoP
P are in dynamic equilibrium between these two oligomeric states. Strikingly, when PhoPHis was applied to the column, the elution profile differed greatly. Unphosphorylated PhoPHis was eluted essentially as a single peak at 67 ml (30 kDa), indicative of a monomer (Fig. 2c
). This peak presented a shoulder, towards the higher molecular masses at 57 ml (69 kDa), which may correspond to small amounts of dimer (Fig. 2c
). In contrast, phosphorylated PhoPHis was eluted as two peaks at 67 ml (30 kDa) and 57 ml (69 kDa), indicating that PhoPHis
P exists as a mixture of monomer and dimer in solution (Fig. 2d
). Overall, these data show that unphosphorylated PhoP and PhoPHis have different oligomeric states in solution, suggesting that the C-terminal His tag affects the stability of the dimer. They also indicate that phosphorylation has no major effect on dimerization of wild-type PhoP.
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DISCUSSION |
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Our data clearly showed that both wild-type PhoP and PhoPP exist as a mixture of monomer and dimer in solution (Fig. 2a, b
). In contrast to what was observed for the ArcA response regulator (Jeon et al., 2001
), higher-order oligomers were not detected under our experimental conditions. In addition, phosphorylation of PhoP did not appear to significantly increase the proportion of dimers (Fig. 2a, b
). These data indicate that the oligomeric state of PhoP is phosphorylation independent. They also suggest that PhoP is involved in a monomerdimer equilibrium that is concentration dependent rather than phosphorylation dependent. We determined the oligomeric state of PhoP and PhoP
P at a protein concentration of 10 µM (Fig. 2a, b
). It has been estimated that the cellular concentration of PhoP ranges between 2 and 11 µM, depending on the presence of Mg2+ in the growth medium (Lejona et al., 2004
). A cellular concentration in the same range (6 µM) was estimated for the OmpR response regulator (Cai & Inouye, 2002
). At such concentrations, it is most likely that PhoP dimers will form, in vivo, regardless of the PhoP phosphorylation state. Our results regarding the oligomeric state of PhoP are in contrast to those reported for many response regulators. For example, phosphorylation is required for dimerization of E. coli PhoB (Fiedler & Weiss, 1995
; McCleary, 1996
), whereas PhoPBSU has been reported to be dimeric independently of its phosphorylation state (Liu & Hulett, 1997
).
Our SPR experiments (Fig. 6) clearly showed that both PhoP and PhoP
P bind to the PhoP box of the mgtA promoter. This is in contrast to the OmpR and PhoB response regulators that poorly interact with DNA when unphosphorylated (Aiba et al., 1989
; Makino et al., 1989
). Recently, it has been shown that the unphosphorylated receiver domain of PhoB inhibits the activity of the DNA-binding domain, and that this inhibition is relieved upon phosphorylation (Ellison & McCleary, 2000
). Our data suggest that such a mechanism is most unlikely for PhoP. Overall, this study clearly showed that the S. enterica PhoP protein dimerizes and binds to the mgtA promoter regardless of its phosphorylation state. Thus, structural rearrangements that occur upon phosphorylation do not significantly affect the oligomeric state and the ability of PhoP to interact with DNA. Although S. enterica PhoP belongs to the same subfamily of response regulators as OmpR and PhoB, their mechanisms of activation appear to be different.
S. enterica or E. coli PhoP response regulators with a C-terminal His tag have been used in previous studies (Lejona et al., 2003, 2004
; Minagawa et al., 2003
; Yamamoto et al., 2002
). Hypothesizing that a His tag at the C-terminus of the DNA-binding domain may interfere with the ability of PhoP to interact with DNA, we systematically compared the in vitro biochemical properties of PhoP, PhoP
P, PhoPHis and PhoPHis
P. Strikingly, we found that the His tag specifically affects the properties of unphosphorylated PhoPHis. Both stability of the dimer (Fig. 2c
) and binding to target DNA (Fig. 6c
) were impaired for unphosphorylated PhoPHis. These results suggest that the C-terminal His tag may affect the function of PhoPHis. This possibility is strongly supported by our fluorescence experiments. Both the intrinsic tryptophan fluorescence measurements (Fig. 3
) and the ANS binding experiments (Fig. 4
) clearly indicated that unphosphorylated PhoP and PhoPHis have different conformations, and/or oligomerization states. Recently, the crystal structure of the full-length Thermotoga maritima DrrD response regulator has been solved (Buckler et al., 2002
). This is the first three-dimensional structure of a full-length member of the OmpR/PhoB subfamily. Examination of the DrrD structure suggests that extension of the protein at its C-terminus may introduce a steric hindrance with the four-stranded
-sheet located at the N-terminus of the DNA-binding domain. This
-sheet appears to be involved in interdomain interactions (Buckler et al., 2002
). It is possible that steric hindrance induced by the His tag reduces PhoPHis flexibility and/or affects interdomain communication. In turn, this would result in a locked conformation less permissive for PhoPHis dimerization. Interestingly, our results showed that PhoPHis forms stable dimers and interacts with target DNA upon phosphorylation (Fig. 2d
, Fig. 6d
). In addition, the tryptophan fluorescence spectrum of PhoPHis
P is very similar to that of PhoP
P (Fig. 3
). This suggests that a phosphorylation-induced conformational change released the steric hindrance imposed by the C-terminal His tag. These results are in good agreement with the proposal that the phosphorylation-induced conformational change is propagated up to the C-terminus of response regulators and promotes a repositioning of the N- and C-terminal domains (Stock & West, 2003
).
Using gel mobility-shift assays and DNase I footprinting, a previous study showed that phosphorylation of PhoPHis enhanced its binding to the promoters that contain a consensus PhoP box (Lejona et al., 2003). Another study used chemical cross-linking to show that self-association of PhoPHis is increased by phosphorylation (Lejona et al., 2004
). Our data obtained with PhoPHis strongly support these findings. However, our data obtained with PhoP indicate that these conclusions may not be relevant for wild-type PhoP. Thus, we propose that unphosphorylated PhoP binds to the PhoP box of target genes, most likely as a dimer. As in the case of OmpR (Ames et al., 1999
), the interaction of PhoP with DNA might stimulate PhoP phosphorylation by its cognate sensor kinase. Since there are only single, if any, PhoP boxes in the promoters of known PhoP-activated genes (Lejona et al., 2003
), phosphorylation of PhoP is unlikely to favour cooperative binding between PhoP dimers at the PhoP-activated promoters, as previously observed for OmpR (Harlocker et al., 1995
). With very few exceptions, response regulators are active in their phosphorylated forms. Thus, it is possible that the role of PhoP phosphorylation is to favour proteinprotein interactions with the transcriptional machinery. This possibility is supported by previous studies showing that PhoB interacts with
70 (Kumar et al., 1994
; Makino et al., 1993
), and that OmpR interacts with the C-terminal domain of the RNA polymerase
subunit (Slauch et al., 1991
).
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ACKNOWLEDGEMENTS |
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Received 27 May 2005;
revised 22 August 2005;
accepted 9 September 2005.
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