JB
Home Help [Feedback] [For Subscribers] [Archive] [Search] [Contents]
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Comolli, J. C.
Right arrow Articles by Donohue, T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Comolli, J. C.
Right arrow Articles by Donohue, T.
Journal of Bacteriology, January 2002, p. 390-399, Vol. 184, No. 2
0021-9193/01/$04.00+0     DOI: 10.1128/JB.184.2.390-399.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Transcriptional Activation of the Rhodobacter sphaeroides Cytochrome c2 Gene P2 Promoter by the Response Regulator PrrA

James C. Comolli, Audrey J. Carl, Christine Hall, and Timothy Donohue*

Department of Bacteriology, University of Wisconsin-Madison, Madison, Wisconsin 53706

Received 17 May 2001/ Accepted 11 October 2001


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Anoxygenic photosynthetic growth of Rhodobacter sphaeroides, a member of the {alpha} subclass of the class Proteobacteria, requires the response regulator PrrA. PrrA and the sensor kinase PrrB are part of a two-component signaling pathway that influences a wide range of processes under oxygen-limited conditions. In this work we characterized the pathway of transcription activation by PrrB and PrrA by purifying these proteins, analyzing them in vitro, and characterizing a mutant PrrA protein in vivo and in vitro. When purified, a soluble transmitter domain of PrrB (cPrrB) could autophosphorylate, rapidly transfer phosphate to PrrA, and stimulate dephosphorylation of phospho-PrrA. Unphosphorylated PrrA activated transcription from a target cytochrome c2 gene (cycA) promoter, P2, which contained sequences from -73 to +22 relative to the transcription initiation site. However, phosphorylation of PrrA increased its activity since activation of cycA P2 was enhanced up to 15-fold by treatment with the low-molecular-weight phosphodonor acetyl phosphate. A mutant PrrA protein containing a single amino acid substitution in the presumed phosphoacceptor site (PrrA-D63A) was not phosphorylated in vitro but also was not able to stimulate cycA P2 transcription. PrrA-D63A also had no apparent in vivo activity, demonstrating that aspartate 63 is necessary both for the function of PrrA and for its phosphorylation-dependent activation. The cellular level of wild-type PrrA was negatively autoregulated so that less PrrA was present in the absence of oxygen, conditions in which the activities of many PrrA target genes increase. PrrA-D63A failed to repress expression of the prrA gene under anaerobic conditions, suggesting that this single amino acid change also eliminated PrrA function in vivo.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Under oxygen-limiting conditions, the purple nonsulfur bacterium Rhodobacter sphaeroides induces the synthesis of a specialized set of photosystem gene products that allow it to grow photosynthetically. The two-component PrrBA system is one of the regulatory systems that ensure appropriate expression of R. sphaeroides photosystem genes (reviewed in references 39 and 54). Mutants lacking PrrB or PrrA exhibit moderate to severe photosynthetic growth defects (14, 15) and fail to appropriately activate expression of genes that encode components of the photosynthetic reaction center and light-harvesting complexes (the puc and puf operons and puhA) (15), energy-transducing proteins, such as cytochrome c2 (cycA) (15, 27), components of the carbon dioxide assimilation pathway (cbb), and proteins involved in nitrogen fixation (nif) (26, 40). Despite the important role that PrrBA plays in the different metabolic lifestyles of R. sphaeroides, details concerning the molecular mechanism by which this signal transduction pathway controls target gene expression are still not known.

Two-component systems analogous to PrrBA have been described in other members of the {alpha} subclass of the class Proteobacteria ({alpha}-proteobacteria), and it has been proposed that they play a similar role in controlling gene expression in response to oxygen limitation (2, 35, 48). The related response regulators of these bacteria exhibit more than 75% amino acid identity, and each regulator has an invariant 21-residue region in its C-terminal output domain that encodes a putative helix-turn-helix motif thought to be responsible for direct interaction with the promoters of target genes (32). Interestingly, this C-terminal domain does not exhibit significant homology with output domains of other DNA-binding response regulators, including those of the OmpR, FixJ, and NtrC subfamilies (19). In studies of response regulators that are closely related to PrrA, it has been shown that Bradyrhizobium japonicum RegR binds to the promoter region of the fixR-nifA operon (2, 12) and Rhodobacter capsulatus RegA directly interacts with the promoters of more than 20 target genes (3, 5, 9, 10, 48, 50). The apparent binding sites for these proteins at target promoters are located at different distances from their transcriptional start sites, making the mechanism of transcription activation an interesting question (3, 5, 11, 48).

Genetic evidence has suggested that phosphorylation of R. sphaeroides PrrA by PrrB stimulates transcription of target genes when oxygen becomes limiting. However, it also appears that unphosphorylated PrrA can influence gene expression, particularly under aerobic conditions, when the kinase activity of PrrB is presumably very low (13, 14, 27). We sought to address the properties of the PrrBA signal transduction pathway by developing a system to investigate the mechanism of transcriptional activation by R. sphaeroides PrrA at the P2 promoter of the cytochrome c2 gene (cycA P2).

Workers in our lab have previously demonstrated that PrrA is responsible for both aerobic expression and anaerobic induction of cycA P2 (27). Purified R. capsulatus RegA*, a RegA mutant protein that has increased activity in the presence and absence of phosphorylation (9), stimulated transcription from cycA P2 in vitro (27). Promoter deletion analysis and DNase I footprinting suggested that unlike the previously described target promoters of R. capsulatus RegA, the cycA P2 promoter contains only one binding site for PrrA. This suggestion was supported by the observation that a 4-bp mutation in the region protected by RegA* eliminated both cycA P2 expression in vivo and RegA*-dependent transcription activation in vitro (27). In the present work, we utilized purified R. sphaeroides PrrA and PrrB to gain insight into the function of this signaling system. Our results show that unphosphorylated wild-type PrrA can stimulate transcription from cycA P2 in vitro and that phosphorylation of PrrA enhances this activity. In addition, analysis of a mutant PrrA with a single amino acid change at the putative phosphoacceptor site (PrrA-D63A) revealed several important features of the PrrBA signaling pathway.

(A preliminary report of this work was presented at the 100th General Meeting of the American Society for Microbiology, Los Angeles, Calif., May 2000.)


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Strains and culture conditions. R. sphaeroides was grown at 30°C in Sistrom’s minimal medium A (44) containing 1 µg of tetracycline per ml or 25 µg of kanamycin per ml as needed. For aerobic growth culture flasks were vigorously shaken. For anaerobic growth filled and sealed tubes were incubated in the light (10 W/m2) for photosynthetic growth or filled and sealed tubes containing Sistrom’s medium supplemented with 0.625% dimethyl sulfoxide (DMSO), 0.2% yeast extract, and 20 mM glucose were incubated in the dark for growth by anaerobic respiration. Escherichia coli DH5{alpha} and XL-1 Blue were used for cloning procedures, E. coli S17-1 was utilized for conjugal transfer of plasmids to R. sphaeroides, E. coli ER2566 was used for intein/chitin-binding domain fusion expression, and E. coli M15(pREP4) was used for His6-tagged protein expression (Table 1). E. coli strains were propagated at 37°C in Luria-Bertani medium supplemented with 20 µg of tetracycline per ml, 100 µg of ampicillin per ml, or 25 µg of kanamycin per ml as needed.


View this table:
[in this window]
[in a new window]
  		TABLE 1. Bacterial strains and plasmids

 
Purification of cPrrB. The cytoplasmic transmitter domain of PrrB (cPrrB) was fused at its N terminus to a His6 tag; this hybrid protein was overexpressed in E. coli and purified by Ni2+-nitrilotriacetic acid (NTA) Sepharose chromatography. The expression plasmid pAC101 was constructed by PCR amplification of codons 189 to 461 of the prrB gene with primers PrrB-3 (5'-CCATCGGGATCCGTTCGATGTCGGATGCGC-3') and PrrB-4 (5'-TCCGGCGGTACCTCAGGTCTGGATCAGGACGTTC-3') and pUI1643. The PCR product was digested with BamHI and KpnI (at sites in the primers) and ligated into BamHI/KpnI-digested pQE32 (Qiagen Inc., Valencia, Calif.) to generate a His6-cPrrB fusion. After DNA sequencing verified that no mutations were present, this plasmid was transformed into M15(pREP4). To induce His6-cPrrB expression, 1 mM isopropyl-ß-D-thiogalactopyranoside (IPTG) was added to an 800-ml culture of the resulting strain and the preparation was incubated for 3 h at 37°C. The cells were harvested, suspended in lysis buffer (50 mM NaH2PO4 [pH 8.0], 300 mM NaCl, 10 mM imidazole, 1 µg of lysozyme per ml), and lysed by sonication. To purify His6-cPrrB, the cell extract was centrifuged (10,000 x g, 20 min, 4°C), and the supernatant was loaded onto a 4-ml Ni2+-NTA Sepharose column, which was washed with 3 volumes of lysis buffer and 10 volumes of column buffer (50 mM NaH2PO4 [pH 8.0], 300 mM NaCl) containing 20 mM imidazole before the bound protein was eluted with column buffer containing 250 mM imidazole. Fractions determined to contain cPrrB by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) were pooled and dialyzed against 20 mM HEPES (pH 8.0)-400 mM NaCl-5 mM MgCl2-20% glycerol. The protein concentrations of this and other preparations were determined by a protein assay (Bio-Rad, Hercules, Calif.).

Expression and purification of PrrA and PrrA-D63A. PrrA and PrrA-D63A were overexpressed in E. coli and were purified by using an intein/chitin-binding domain fusion system (New England Biolabs, Waltham, Mass.) like that used for purification of R. capsulatus RegA* (11). The plasmid used for purification of PrrA (pJC407) was constructed by amplifying prrA from pUI1643 with primers PrrA-4 (5'-CGAACACCATGGCTGAGGATCTGGTATTC-3') and PrrA-5B (5'-GCGACTCCCGGGGCGCGGGCTGCGCTTCGCC-3'); the product was digested with NcoI and SmaI (at sites in the primers) and ligated into NcoI/SmaI-digested pTYB4. The prrA gene of each plasmid was sequenced to ensure that no mutations were introduced by PCR.

The point mutation used to generate the aspartate-to-alanine change at PrrA residue 63 (PrrA-D63A) was an A-to-C change at nucleotide position 187 in prrA. Plasmid pJC402 was generated by cloning a 1.9-kb EcoRI/XbaI fragment of pUI1643 into EcoRI/XbaI-digested pBS-SK; this plasmid served as the template for the first of two PCR steps in which the T7 vector primer and primer PrrAD63A (5'-GAGCCGCAGGGCCACCACTGC-3') were used. The reaction product was used in combination with the T7 vector primer for a second PCR in which pUI1643 was used as the template. The ~2.0-kb product was digested with XbaI and HindIII and cloned into XbaI/HindIII-digested pBSII-KS to generate pJC410. The prrA region was sequenced to verify that only the desired nucleotide change was present. The plasmid used for purification of PrrA-D63A (pJC417) was generated by using the procedure for construction of pJC407 described above.

To purify PrrA or PrrA-D63A, cultures of E. coli ER2566 containing either pJC407 or pJC417 were incubated with 0.1 mM IPTG for 5 h at 28°C. Cells were harvested by centrifugation, suspended in 10 ml of column buffer (20 mM Tris-Cl [pH 8.0], 0.1 mM EDTA, 0.5 M KCl, 0.1% Triton X-100), and lysed by sonication. The cell extract was centrifuged at 13,000 x g and 4°C for 30 min, and the supernatant was loaded onto a 10-ml chitin column previously equilibrated with column buffer. The column was washed with 10 volumes of column buffer, flushed with 35 ml of 20 mM Tris-Cl (pH 8.0)-0.1 mM EDTA-0.5 M KCl-50 mM dithiothreitol (DTT), and stored overnight at 4°C to allow for intein cleavage. Released protein was collected in 3-ml fractions upon elution with chitin column buffer. Fractions that contained PrrA or PrrA-D63A were pooled and dialyzed against 50 mM HEPES (pH 7.8)-0.2 M KCl-10 mM MgCl2-1 mM DTT-50 µM phenylmethylsulfonyl fluoride and then against the same buffer containing 50% glycerol for storage at -80°C.

For in vivo analysis of PrrA-D63A, pJC410 was digested with SalI in order to delete a 1.1-kb fragment and generate pJC419. The 0.83-kb prrA-containing insert from pJC419 was then removed by digestion with XbaI and HindIII and ligated into broad-host-range vector pRK415 treated with the same enzymes. The resulting plasmid (pJC414) contained the 3' portion of prrC (containing a promoter for prrA [15]) and prrA-D63A in an orientation opposite that of the lac promoter on RK415. This plasmid was transformed into E. coli S17-1 and transferred to R. sphaeroides by conjugation.

Histidine kinase assays. cPrrB activity was analyzed in kinase buffer containing 50 mM Tris-Cl (pH 7.5), 50 mM KCl, and 5 mM MgCl2. To determine the ATP dependence of cPrrB autophosphorylation, 5 µM cPrrB was incubated for 15 min at 25°C with 25 µM to 1 mM ATP supplemented with [{gamma}-32P]ATP at a final specific activity of 4,000 cpm/pmol. The time course of cPrrB autophosphorylation was determined by using identical conditions in the presence of 600 µM ATP. The stability of phospho-cPrrB was assessed after autophosphorylation of cPrrB in kinase buffer for 60 min at 25°C with 200 µM ATP (4,000 cpm/pmol), followed by 200 mM ATP. In each of these experiments, reactions were terminated by adding 0.33 volume of 3x SDS-polyacrylamide sample buffer (188 mM Tris-Cl [pH 6.8], 6% SDS, 30% glycerol, 0.3 M DTT, 0.03% bromophenol blue), and the preparations were stored on ice. Samples of all radiolabeling assay mixtures were analyzed by electrophoresis on SDS-12% polyacrylamide gels which were dried and visualized by phosphor imager analysis (Molecular Dynamics, Sunnyvale, Calif.).

Phosphorylation of PrrA. Purified PrrA or PrrA-D63A (5 µM) was added to 7 µM phospho-cPrrB that was autophosphorylated in kinase buffer with 50 µM ATP (50,000 cpm/pmol) for 60 min at 25°C. Alternatively, PrrA or PrrA-D63A (final concentration, 25 µM) was incubated with 25 mM [32P]acetyl phosphate, synthesized as described by McCleary and Stock (33) by using 0.4 ml of H332PO4 in a solution containing 50 mM Tris-Cl (pH 7.0), 5 mM MgCl2, 0.1 mM DTT, and 0.1 mg of bovine serum albumin per ml.

To measure the stability of phospho-PrrA, 10 µM cPrrB was phosphorylated with 50 µM ATP (50,000 cpm/pmol) for 60 min at 25°C in a 300-µl reaction mixture, diluted with 200 µl of kinase buffer, and then incubated with 50 µl of a 50% slurry of Ni2+-NTA agarose washed in kinase buffer. This mixture was kept on ice for 60 min at 4°C with frequent agitation, and then the precipitate was pelleted by centrifugation at 16,000 x g for 5 min. The pellet was washed six times in cold kinase buffer and suspended in a solution containing 8 µM PrrA in kinase buffer. After 10 min of incubation at 25°C, the mixture was centrifuged three times to remove the agarose-bound cPrrB. The resulting supernatant was divided into two equal portions and added to equal volumes of kinase buffer with and without 4 µM purified cPrrB.

In vitro transcription assay. Crude R. sphaeroides RNA polymerase holoenzyme was prepared by heparin agarose chromatography (6). Briefly, 2 g (wet weight) of aerobically grown R. sphaeroides cells was suspended in 6 ml of a solution containing 10 mM Tris-Cl (pH 8.0), 50 mM NaCl, 10 mM MgCl2, 1 mM EDTA, 0.3 mM DTT, 7.5% glycerol, 0.5 mM phenylmethylsulfonyl fluoride, and 1.5 mg of lysozyme per ml and incubated on ice for 10 min. Following lysis by sonication, the cell extract was centrifuged at 10,000 x g for 40 min at 4°C, and the resulting supernatant was mixed with 3 volumes of 50% heparin-agarose equilibrated in heparin-agarose buffer (10 mM Tris-Cl [pH 8.0], 10 mM MgCl2, 1 mM EDTA, 0.3 mM DTT) for 60 min at 4°C. This slurry was poured into a column and washed with 5 volumes of heparin-agarose buffer containing 7.5% glycerol and 50 mM NaCl. Fractions were eluted from the column with heparin-agarose buffer containing 7.5% glycerol and 0.6 M NaCl and were analyzed on SDS-polyacrylamide gels. The elution fractions that contained RNA polymerase were pooled.

The in vitro transcription assay was performed as described previously (30) by using approximately 4.3 µg of the crude RNA polymerase preparation, PrrA (either 0.15, 0.45, or 2 µM PrrA, PrrA treated for 60 min at 25°C with 25 mM acetyl phosphate, or PrrA-D63A), and 20 nM cycA P2 template plasmid pRKK146 in a 20-µl reaction mixture containing transcription buffer (40 mM Tris-Cl [pH 7.9], 50 mM KCl, 5 mM MgCl2, 0.2 mM EDTA, 0.1 mM DTT, 0.1 mg of bovine serum albumin per ml). The components were incubated for 20 min at 30°C, and the transcription reaction was initiated by adding nucleotide triphosphates (0.5 mM ATP, 0.5 mM CTP, 0.5 mM GTP, and 50 µM UTP plus 10 µCi of [{alpha}-32P]UTP) and was allowed to proceed for 20 min at 30°C before 10 µl of loading buffer containing 95% formamide, 20 mM EDTA, 0.05% bromophenol blue, and 0.05% xylene cyanol was added. Samples were heated to 80°C for 5 min and loaded onto a 6% polyacrylamide-urea gel that was analyzed with a phosphor imager.

Analysis of reporter activity. The levels of ß-galactosidase expression in exponentially growing R. sphaeroides cultures were measured by using a protocol adapted for R. sphaeroides (41).

Western blot analysis. Antiserum to PrrA was generated in New Zealand White rabbits by using purified PrrA (Cocalico Biologicals, Reamstown, Pa.). Approximately 8.0 x 108 R. sphaeroides cells in the logarithmic growth phase were pelleted, resuspended in 100 µl of 1x SDS-PAGE sample buffer, and heated at 90°C for 10 min. Portions (10 µl) of the samples were electrophoresed on an SDS-12% polyacrylamide gel and transferred to a polyvinylidene difluoride membrane (Millipore Corp., Waltham, Mass.). The membrane was treated with 5% milk in TBS-T (10 mM Tris-Cl [pH 8.0], 150 mM NaCl, 0.05% Tween 20) and then incubated with a 1:3,000 dilution of rabbit anti-PrrA antiserum in 5% milk-TBS-T; this was followed by incubation in a 1:5,000 dilution of horseradish peroxidase-conjugated anti-rabbit immunoglobulin G (Pierce) in TBS-T. Secondary antibody was detected by chemiluminescence by using 2.5 mM luminol (3-aminophthalhydrazine) and 450 µM p-coumaric acid (Sigma) in 100 mM Tris-Cl (pH 8.5)-0.02% H2O2 and was visualized by exposure to X-AR5 film.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Autophosphorylation by the C-terminal transmitter domain of PrrB. As a first step in the analysis of the R. sphaeroides PrrBA signal transduction chain, we purified PrrB and PrrA and monitored their phosphorylation in vitro. We were unable to overexpress N-terminal histidine-tagged (His6) full-length PrrB in E. coli (data not shown), a difficulty previously encountered by other groups working with related membrane-bound sensor histidine kinases (12, 25), so instead we fused an N-terminal His6 tag to the C-terminal transmitter domain (amino acids 193 to 461) of PrrB, which has been shown to reside in the cytoplasm (38). This truncated protein, called cPrrB, was soluble in E. coli and was obtained at more than 98% homogeneity by Ni2+-NTA Sepharose chromatography (data not shown). Incubation of cPrrB with [{gamma}-32P]ATP and Mg2+ resulted in phosphorylation of this protein (Fig. 1), indicating that the C-terminal domain of PrrB contained the regions required for ATP binding and kinase activity.



View larger version (30K):
[in this window]
[in a new window]
  		FIG. 1. Autophosphorylation of the transmitter domain of PrrB (cPrrB). (A) ATP dependence of cPrrB autophosphorylation. cPrrB (5 µM) was incubated with different concentrations of ATP for 15 min. The amount of [{gamma}-32P]ATP was kept constant at 4,000 cpm/pmol. (B) Rate of cPrrB autophosphorylation. The time course of phosphorylation was monitored by using 5 µM cPrrB and 600 µM ATP. (C) Stability of phospho-cPrrB. Following a 60-min autophosphorylation of cPrrB, a 1,000-fold excess of unlabeled ATP was added to the reaction mixture, and the amount of phospho-cPrrB remaining was monitored over time. The amount of cPrrB phosphorylation in each experiment, quantified by SDS-PAGE and phosphor imager analysis, is also shown graphically.

 
To determine the optimal reaction conditions for cPrrB kinase activity, the concentration of ATP in the reaction mixture was varied at a fixed protein concentration. Within the range from 25 µM to 1 mM ATP, autophosphorylation of cPrrB exhibited typical saturation kinetics, and a half-maximal rate was obtained at an ATP concentration of about 170 µM. The maximal amount of phosphorylated cPrrB (phospho-cPrrB), estimated to be 30% of the total protein, was produced when >=600 µM ATP was used in a 60-min assay (Fig. 1A). To measure the rate of cPrrB autophosphorylation, the reaction when 600 µM ATP was present was monitored as a function of time. Under these conditions it took ~7 min to obtain half-maximal phosphorylation, and after 15 min no additional accumulation of phospho-cPrrB was detected (Fig. 1B). An additional control experiment demonstrated that the extent of cPrrB phosphorylation after 60 min in the presence of 600 µM ATP was directly related to the amount of cPrrB present in the reaction mixture for protein concentrations ranging from 0.2 to 10 µM (data not shown).

To assess the stability of phospho-cPrrB, cPrrB was phosphorylated with [{gamma}-32P]ATP, a 1,000-fold excess of unlabeled ATP was added to the reaction mixture, and the amount of phospho-cPrrB remaining was quantified over time. Under these conditions, phospho-cPrrB had a half-life (t1/2) of 5.5 to 6.0 h (Fig. 1C). In many respects, this property of purified cPrrB is similar to the properties of truncated transmitter domains of other sensor kinases, including the related proteins RegB of R. capsulatus and RegS of B. japonicum (1, 16, 18, 24, 31).

To assess the in vivo activity of cPrrB, a plasmid-derived (pJC415) gene encoding cPrrB under control of its own promoter was placed in the R. sphaeroides wild-type strain or PrrB1, a mutant strain in which the prrB gene is inactivated (14). The plasmid did not rescue the photosynthetic growth defect of the PrrB1 mutant, nor did it alter the aerobic expression of two different PrrBA-dependent reporter genes (containing the puc or cycA P2 promoter) fused to the E. coli ß-galactosidase gene in a wild-type or PrrB1 background (data not shown). Thus, while cPrrB had measurable in vitro activity, its activity in vivo relative to that of wild-type PrrB was minimal, suggesting that the truncated version of the protein is either unstable or inactive when it is expressed in R. sphaeroides. Unfortunately, it has been difficult to raise specific, high-titer polyclonal antiserum to cPrrB (Comolli and Donohue, unpublished data), so we could not determine if this soluble kinase domain accumulates in R. sphaeroides.

Phosphorylation of the response regulator PrrA. In order to observe phosphotransfer from phospho-cPrrB to its cognate response regulator, PrrA was obtained from E. coli at nearly 99% homogeneity by using an intein/chitin-binding domain fusion system similar to that used for purification of active R. capsulatus RegA (9; data not shown). The rate and extent of phosphotransfer were determined by mixing PrrA with 32P-labeled cPrrB and observing phospho-PrrA production. As shown in Fig. 2A, phosphate was rapidly transferred from phospho-cPrrB to PrrA; 98% of the phosphate of phospho-cPrrB was transferred to PrrA in 15 s or less. No detectable phospho-PrrA was generated when PrrA was incubated with [{gamma}-32P]ATP in the absence of cPrrB (data not shown). Rapid phosphotransfer from sensor kinase to response regulator is typical of two-component partners (46). When the rate of cPrrB phosphorylation was compared to the rate of phosphotransfer to PrrA, the data suggested that a rate-limiting step in the production of phospho-PrrA is the autophosphorylation of PrrB.



View larger version (43K):
[in this window]
[in a new window]
  		FIG. 2. Phosphorylation of PrrA. (A) Phosphotransfer from phospho-cPrrB (cPrrB~P) to PrrA. PrrA (5 µM) was added to 7 µM cPrrB that had been autophosphorylated for 60 min. The time point indicated as 0 s was immediately before PrrA was added. (B) PrrA can accept phosphate from acetyl phosphate. Radiolabeled acetyl phosphate at a concentration of 25 mM was incubated with 25 µM PrrA for different times. (C) Stability of phospho-PrrA. Isolated phospho-PrrA was monitored to determine the release of phosphate over time. The decay of phospho-PrrA without (•) or with ({circ}) cPrrB added at a 1:1 molar ratio is shown. t1/2 of phospho-PrrA were calculated from linear regression of the amount of remaining phospho-PrrA.

 
Many response regulators also can be phosphorylated with small phosphodonors such as acetyl phosphate (29, 33). To determine if this was also a property of PrrA, the protein was mixed with 25 mM [32P]acetyl phosphate, and the production of phospho-PrrA was measured. Labeled phospho-PrrA was detected as early as 5 min after the addition of [32P]acetyl phosphate, and the maximal level accumulated in 60 min (Fig. 2B). The maximum amount of phospho-PrrA, which was estimated to be 15 to 20% of the total protein, did not change when the concentration of [32P]acetyl phosphate was varied between 5 and 50 mM, indicating that the amount of phosphodonor was not limiting (data not shown).

The t1/2 of aspartyl phosphate on different response regulators can range from seconds to hours (22, 23, 31, 52, 53) and is often influenced by the presence of the cognate sensor kinase (45). When we measured the inherent stability of the phosphate on phospho-PrrA, isolated as described in Materials and Methods, we found that it had a t1/2 of ~330 min (Fig. 2C) and thus was relatively stable compared to other response regulators. To examine the ability of cPrrB to enhance dephosphorylation of phospho-PrrA, an equimolar amount of cPrrB was added to a preparation of 32P-labeled PrrA. The presence of cPrrB decreased the t1/2 of phospho-PrrA to 20 min, which represented a >16-fold reduction relative to the stability in the absence of the kinase (Fig. 2C). Although determined by a different method, the stability of phospho-PrrA is similar to that of B. japonicum phospho-RegR (t1/2, more than 162 min [12]) but is significantly greater than that of R. capsulatus phospho-RegA (t1/2, approximately 90 min [3]). In these two systems, the rate of response regulator dephosphorylation was also stimulated by the cognate sensor kinase.

Transcription activation by PrrA and phospho-PrrA in vitro. Several observations suggest that PrrA can stimulate target gene expression in R. sphaeroides under aerobic conditions when the kinase activity of PrrB and phosphorylation of this response regulator should be negligible (13, 14, 27). To assess the ability of PrrA to stimulate gene expression, we monitored transcription from the R. sphaeroides PrrA-dependent promoter cycA P2 in vitro. For these assays, we used a cycA P2 promoter that extends from -73 to +22 relative to the transcription initiation site, which was shown to contain a single binding site when it was tested with the mutant RegA* protein from R. capsulatus (27). In the absence of PrrA, R. sphaeroides RNA polymerase holoenzyme reproducibly produced no transcript from this promoter (Fig. 3). Addition of 0.15 to 1.5 µM PrrA to the in vitro reaction mixture resulted in dose-dependent increases in transcription from cycA P2 of up to 126-fold compared to the results obtained with the reaction mixture lacking PrrA (Fig. 3). The PrrA-dependent stimulation was specific to cycA P2 since the amount of transcript generated from the control RNA 1 promoter was not affected by addition of the activator. Additional experiments showed that a 4-bp mutation of the presumed PrrA-binding site eliminated transcription by PrrA in vitro (data not shown), as shown previously both in vivo and in vitro with RegA* (27). When these results were considered together, we concluded that PrrA is able to interact with R. sphaeroides RNA polymerase and activate transcription from a single binding site within cycA P2 even in the absence of phosphorylation. We do not believe that this activation is the result of a fraction of the PrrA being phosphorylated after purification from E. coli, since attempts to dephosphorylate purified PrrA by using cPrrB or alkaline phosphatase had no measurable effect on PrrA function at this promoter in vitro (data not shown). In addition, electrospray mass spectroscopy failed to detect evidence of phosphorylated PrrA in the preparations (data not shown).



View larger version (149K):
[in this window]
[in a new window]
  		FIG. 3. In vitro transcription of the cycA P2 promoter by PrrA, phospho-PrrA (PrrA~P), or PrrA-D63A. Transcription products from pRKK146, a template containing a cycA P2 promoter (positions -73 to +22), and a control promoter (RNA 1) were generated by using purified R. sphaeroides RNA polymerase holoenzyme. The transcription reaction mixtures contained no added PrrA or 0.15, 0.45, or 1.5 µM PrrA~P, PrrA, or PrrA-D63A as indicated. The relative transcript abundance indicates the phosphor imager density of each transcript after subtraction of the background value and standardization to the amount of RNA 1 produced in each lane.

 
To determine the effect of phosphorylation on PrrA function in vitro, we used acetyl phosphate as a phosphodonor since addition of cPrrB to a transcription reaction mixture prevented accumulation of transcripts from both the cycA P2 and control promoters. When PrrA was treated with 25 mM acetyl phosphate for 60 min before it was added to an in vitro transcription reaction mixture, it stimulated cycA P2 transcription to a greater extent than unphosphorylated PrrA. Transcriptional activation by acetyl phosphate-treated PrrA was 15.6-fold greater with 0.15 µM PrrA, 6.6-fold greater with 0.45 µM PrrA, and 2.5-fold greater with 1.5 µM PrrA compared to transcriptional activation in reaction mixtures containing the same concentrations of untreated PrrA (Fig. 3). Control experiments indicated that the differences in stimulation due to phosphorylation of PrrA reflected the fact that maximum cycA P2 promoter activity occurred at concentrations of phosphorylated PrrA greater than 0.5 µM (Carl and Donohue, unpublished results). To summarize, unphosphorylated PrrA was able to stimulate transcription from a cycA P2 promoter that extends from position -73 to position +22, but phosphorylation of this protein increased the ability of PrrA to activate this promoter.

Characteristics of a mutant PrrA-D63A protein. Since PrrA activated transcription in vitro in the absence of phosphorylation, we sought to further address the contribution of the unphosphorylated protein to target gene expression in vivo. Although PrrA-dependent target genes generally exhibit increased expression under anaerobic conditions when PrrA is thought to be phosphorylated in vivo (37), there is evidence which suggests that the unphosphorylated form of this protein can also stimulate transcription (13, 14, 27). To assess the contribution of PrrA phosphorylation to activity, we generated a mutant PrrA protein with the conserved putative phosphorylation site, aspartate residue 63, changed to an alanine in order to create a form of PrrA (PrrA-D63A) that should not be able to accept phosphate. When PrrA-D63A was overexpressed in E. coli in the same manner as wild-type PrrA, a similar quantity of soluble protein was produced, suggesting that the tertiary structure of PrrA-D63A was not drastically different from that of the wild-type molecule. PrrA-D63A could not be phosphorylated by cPrrB under conditions that lead to rapid phosphorylation of wild-type PrrA (Fig. 4A). PrrA-D63A also was not phosphorylated at a detectable level when it was incubated with [32P]acetyl phosphate for up to 90 min, even though wild-type PrrA was maximally phosphorylated within 45 min (Fig. 4B). These results demonstrate that the alanine at residue 63 prevented phosphorylation of PrrA-D63A, which implicates D63 as the site of PrrA phosphorylation. The failure to detect any phosphorylation of PrrA-D63A under conditions that lead to efficient labeling of its wild-type counterpart also suggests that an alternate residue does not serve as a phosphoacceptor in this mutant protein.



View larger version (40K):
[in this window]
[in a new window]
  		FIG. 4. Assay of PrrA-D63A phosphorylation. (A) Phosphotransfer from 5 µM 32P-labeled cPrrB to 5 µM wild-type PrrA or PrrA-D63A. Components were mixed for either 0.5 or 5 min. (B) Analysis of phosphorylation of PrrA or PrrA-D63A by [32P]acetyl phosphate. Protein (20 µM) was incubated with 25 mM radiolabeled acetyl phosphate for the indicated times.

 
When the activity of purified PrrA-D63A was analyzed in an in vitro transcription assay, the amount of cycA P2 transcript produced with up to 1.5 µM protein was comparable to the amount formed in a reaction mixture to which no PrrA was added (Fig. 3). Since PrrA-D63A was not competent in cycA P2 transcription activation in vitro and unphosphorylated wild-type PrrA was competent, it appears that the substitution at residue 63 not only prevented phosphorylation but completely eliminated PrrA activity. This suggests that an aspartate at position 63 may be essential both for the activity of PrrA as a transcription factor and for phosphorylation-dependent activation of PrrA.

To examine the properties of PrrA-D63A in vivo, the gene encoding PrrA-D63A was cloned into the broad-host-range plasmid pRK415 to obtain pJC414, which was placed in an R. sphaeroides strain in which the wild-type prrA gene was inactivated (PrrA1 [15]). PrrA1 cells containing pJC414 were not able to grow via photosynthesis, while a wild-type copy of prrA on a similar plasmid (pUI1621) restored the growth rate to a rate similar to that of a wild-type strain (data not shown). This indicated that PrrA-D63A was deficient in activation of one or more genes required for anaerobic photosynthetic growth. To more closely assess PrrA-D63A function in vivo, the activity of a lacZ transcriptional fusion to the PrrA-dependent cycA P2 promoter (pRKK232) was measured in R. sphaeroides strains containing different prrA-bearing plasmids. Reporter activity was quantified in cells either growing aerobically or growing anaerobically by using DMSO as an electron acceptor since the PrrA1 mutant is unable to grow photosynthetically (15). When PrrA-D63A served as the sole source of PrrA, the level of expression from the cycA P2 promoter was the same as the level of expression in the PrrA1 mutant grown in the presence or absence of oxygen (Fig. 5A). In contrast, a comparable plasmid containing the wild-type prrA gene increased the level of expression of cycA P2 to approximately the level found in wild-type cells. When the puc promoter was utilized as a second reporter of PrrA-induced gene expression, the results were similar to the results obtained with the cycA P2 promoter (data not shown), suggesting that the inability of PrrA-D63A to stimulate target gene expression in vivo was not linked to cycA P2.



View larger version (27K):
[in this window]
[in a new window]
  		FIG. 5. Activity of PrrA-D63A in vivo. (A) Level of cycA P2 expression from a lacZ reporter plasmid (pRKK232) harbored in wild-type (+) or PrrA1 (-) cells grown under aerobic (open bars) or anaerobic respiratory (solid bars) conditions. The strains also contained compatible plasmids that either carried a prrA gene (wild-type), prrA-D63A, or had no prrA gene (-). The error bars indicate the standard deviations based on three independent samples. ß-gal, ß-galactosidase. (B) Anti-PrrA immunoblot analysis of protein levels in the strains used in the experiment described in panel A. Samples were generated from equivalent numbers of cells growing under aerobic conditions (+) or in the absence of oxygen by DMSO respiration (-) or by photosynthesis (PS). chromo, chromosome.

 
To ensure that PrrA-D63A was expressed in vivo at levels sufficient to promote transcription, the amount of PrrA or PrrA-D63A present in cells was examined by performing a Western blot analysis with antiserum generated against purified PrrA. When this analysis was performed with cells expressing PrrA-D63A, the amount of protein produced under aerobic conditions was similar to the amount produced by cells expressing wild-type PrrA from a comparable plasmid (pUI1621) (Fig. 5B), indicating that PrrA-D63A was stable in vivo. From these results, it is clear that the inability of PrrA-D63A to activate transcription could not be attributed to a reduced intracellular level of the protein. These results show that PrrA-D63A does not activate the expression of target genes in vivo and that the D63A substitution prevents PrrA function both in vitro and in vivo.

In the course of these experiments, we also found that wild-type cells grown aerobically contained three- to fivefold more PrrA than cells grown anaerobically either photosynthetically or by respiration using DMSO (Fig. 5B). The quantity of PrrA produced from plasmid pUI1621 was greater than the quantity produced by the wild-type strain under identical conditions due to the multicopy nature of the plasmid, but a two- to threefold difference in the levels of this protein was also observed when cells grown in the presence of oxygen and cells grown in the absence of oxygen were compared. These observations show that expression of the prrA gene is negatively regulated under anaerobic conditions. This result was somewhat unexpected since PrrA-dependent expression from target promoters, such as puf, puc, and cycA P2, is greater under anaerobic conditions (14, 15, 27).

Western blot analysis also showed that cells expressing PrrA-D63A appeared to be unable to control the level of PrrA since similar amounts of protein were detected in the presence and in the absence of oxygen. This finding implies that the reduction in the PrrA levels in the absence of oxygen requires active PrrA protein. When considered together, our data indicate that the PrrA-D63A mutant is not able to activate transcription in vivo and apparently cannot negatively regulate its own expression.


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The sensor kinase PrrB and the response regulator PrrA are signaling molecules necessary for the anaerobic photosynthetic lifestyle of R. sphaeroides Because they influence numerous cellular processes, PrrBA and its R. capsulatus homologue, RegBA (10, 11, 42, 50), are considered global regulators of gene expression under oxygen-limiting conditions. Two-component signal transduction proteins related to PrrBA and RegBA also exist in numerous other {alpha}-proteobacteria, including nonphotosynthetic species like B. japonicum, Sinorhizobium meliloti, and Caulobacter cresentus, and in facultative anaerobes, such as Pseudomonas aeruginosa, Bordetella pertussis, and Shewenella putrefaciens (2, 32, 34, 47, 49). While in our studies we characterized transcription activation by R. sphaeroides PrrBA, the high degree of amino acid identity of the response regulators suggests that our findings may apply to the function of orthologous regulatory systems of other proteobacteria.

Our in vitro studies showed that purified PrrA was able to activate transcription in the absence of phosphorylation. This is the first demonstration of activation by a wild-type regulator in this family since in a previous transcription analysis with R. capsulatus RegA the workers used a gain-of-function mutant with altered in vitro properties (RegA*) (5). In this regard, it is significant that more than 10-fold more R. capsulatus RegA* protein than wild-type PrrA was needed to activate expression from cycA P2 (27). Since RegA and PrrA are 89% identical at the amino acid level, this implies that some of the amino acids that are different may be involved in species-specific RNA polymerase contacts.

A number of response regulators, including OmpR, SpoOA, and NtrC, have been shown to activate transcription of target promoters in vitro while they are unphosphorylated (4, 23, 35), so transcriptional activation by unphosphorylated PrrA cannot be considered unusual. After phosphorylation, PrrA activated the cycA P2 promoter up to 15-fold better than unphosphorylated PrrA in an in vitro transcription assay. Phosphorylation of DNA-binding response regulators often stimulates transcription by increasing affinity for target promoters (17, 20, 35). Additional studies are needed to determine whether phosphorylation-dependent activation of PrrA increases interaction of this response regulator with the transcriptional apparatus or stimulates DNA binding.

PrrA is capable of activating transcription in vitro even in the absence of phosphorylation, supporting the hypothesis that this form of the protein can stimulate target gene expression in vivo. However, a change in the conserved aspartate at position 63 to alanine that prevented phosphorylation in vitro produced a protein that failed to activate transcription of the cycA P2 promoter in vivo and in vitro. Because the in vitro activity of PrrA-D63A was less than that of unphosphorylated PrrA, we concluded that the D63A substitution significantly impairs the function of PrrA. One simple inference drawn from these results is that the aspartate at position 63 plays a crucial role in the structural integrity or activity of PrrA in addition to its predicted function in controlling phosphorylation-dependent activation by accepting phosphate. In this regard, it has been reported that the N-terminal receiver domains of R. capsulatus RegA and B. japonicum RegR, which contain a conserved D63 residue, are necessary for DNA binding, suggesting that these regions productively interact with their corresponding C-terminal DNA-binding domains (12, 21). If this were also true in PrrA, then it is possible that D63 is essential for a productive intramolecular interaction between the N- and C-terminal domains. Consistent with this notion, a D63N substitution in RegR greatly reduced its ability to form a specific protein-DNA complex (12). However, a similar substitution in R. capsulatus RegA (D63K) created a regulator which had significant DNA-binding activity (21), suggesting that the amino acid side chain at this position can have different effects on the biological activities of members of this class of response regulators.

Our results indicate that there are several mechanisms that manipulate the intracellular levels of PrrA and phospho-PrrA. One of these involves the antagonistic effects of the sensor kinase PrrB on PrrA phosphorylation. The cytoplasmic transmitter domain of PrrB phosphorylated PrrA at a rate significantly higher (t1/2 of seconds) than the rate of autophosphorylation of the kinase (t1/2 of minutes). This implies that PrrB autokinase activity is rate limiting and thus a possible point of in vivo regulation of phospho-PrrA production. Once phosphorylated, phospho-PrrA had a low intrinsic rate of dephosphorylation (t1/2, more than 300 min), but its stability was reduced more than 15-fold by an equimolar amount of cPrrB. This in vitro finding corroborates the in vivo observation that PrrB can negatively affect PrrA-dependent target gene expression (13). As is the case with other sensor kinases (1, 7, 28), the ability of cPrrB to dephosphorylate phospho-PrrA may be regulated by the phosphorylation state of the kinase (Comolli and Donohue, unpublished). Thus, it is possible that the signal received by PrrB can affect either its ability to phosphorylate PrrA or its ability to dephosphorylate PrrA or both, in order to control steady-state phosphorylation of this response regulator.

Regulation of the level of cellular PrrA appears to be another mechanism used to modify PrrBA signal propagation. Under aerobic conditions, the level of transcription of cycA P2 and other PrrA-dependent target genes is low and is affected only slightly by disruption of prrB (14, 27), implying that PrrA exists largely in a dephosphorylated state. However, our data showed that there was three- to fivefold more PrrA present per cell under aerobic conditions than under anaerobic conditions, when PrrA is thought to be phosphorylated and expression of PrrBA-dependent target genes is induced (13, 14, 27). In other words, PrrA levels were lower when target gene expression was stimulated in vivo. The most likely explanation for this observation is that there is a phosphorylation-dependent increase in the specific activity of PrrA under oxygen-limiting conditions. Consistent with this notion, phosphorylation increased the activity of PrrA up to 15-fold in vitro, suggesting that this form of the protein could be responsible for the higher specific activity observed in vivo.

These changes in the cellular amount of PrrA are likely due to negative autoregulation of PrrA expression since the level of this protein was not altered in cells expressing the inactive PrrA-D63A protein. Similarly, Du et al. (10) demonstrated that the roughly twofold decrease in expression from the R. capsulatus regB and regA operon promoters under anaerobic conditions was eliminated in cells lacking either RegB or RegA. Their model of direct anaerobic autoregulation of regA and regB gene expression in R. capsulatus is consistent with our observations regarding the cellular levels of R. sphaeroides PrrA. It also suggests that similar autoregulatory pathways may be present in these two photosynthetic bacteria and possibly other organisms that contain homologues of PrrBA.

In summary, our analysis of the in vitro properties of and interactions among PrrB, PrrA, and a target promoter, cycA P2, have provided considerable new insight into the factors that control the function of this signal transduction pathway in vivo. The multiple mechanisms that regulate the amount of cellular phospho-PrrA point towards the presence of a signaling system that is able to respond rapidly to changes in oxygen tension. Under aerobic conditions, PrrA is unphosphorylated and presumably less active as a result of decreased PrrB kinase activity and/or the increased ability of PrrB to dephosphorylate phospho-PrrA. As oxygen tension decreases, a signal sensed by R. sphaeroides PrrB, proposed to be generated by electron flow through the cbb3 respiratory oxidase (36), increases the kinase activity of PrrB to convert the pool of PrrA to phospho-PrrA. This causes rapid induction of transcription of PrrA-dependent target genes. When steady-state anaerobic conditions are established, there is less need for PrrA due to its higher activity when it is phosphorylated, so the level of phospho-PrrA is reduced by autoregulation of prrA. In this model, the signaling pathway is poised to rapidly respond to rising oxygen tensions since inactivation of phospho-PrrA is more easily accomplished when there is less available.


   ACKNOWLEDGMENTS
 
We acknowledge the Pseudomonas genome project (www.pseudomonas.com) and the Institute for Genomic Research (www.tigr.org) for providing sequence information and access to databases for finished and unfinished microbial genomes.

This work was supported in part by NRSA grant GM 20344 from the National Institutes of Health (NIH) to J.C.C., by grant GM 37509 from the NIH to T.J.D., and by Biotechnology Training Grant GM 08349 from the NIH to C.H.


   FOOTNOTES
 
* Corresponding author. Mailing address: Department of Bacteriology, University of Wisconsin-Madison, Room 312, Fred Hall, Madison, WI 53706. Phone: (608) 262-4663. Fax: (608) 262-9865. E-mail: tdonohue{at}bact.wisc.edu. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Aiba, H., T. Mizuno, and S. Mizushima. 1989. Transfer of phosphoryl group between two regulatory proteins involved in osmoregulatory expression of the ompF and ompC genes in Escherichia coli. J. Biol. Chem. 264:8563–8567.[Abstract/Free Full Text]
  2. Bauer, E., T. Kaspar, H. M. Fischer, and H. Hennecke. 1998. Expression of the fixR-nifA operon in Bradyrhizobium japonicum depends on a new response regulator, RegR. J. Bacteriol. 180:3853–3863.[Abstract/Free Full Text]
  3. Bird, T. H., S. Du, and C. E. Bauer. 1999. Autophosphorylation, phosphotransfer, and DNA-binding properties of the RegB/RegA two-component regulatory system in Rhodobacter capsulatus. J. Biol. Chem. 274:16343–16348.[Abstract/Free Full Text]
  4. Bird, T. H., J. K. Grimsley, J. A. Hoch, and G. B. Spiegelman. 1996. The Bacillus subtilis response regulator SpoOA stimulates transcription of the spoIIG operon through modification of RNA polymerase promoter complexes. J. Mol. Biol. 256:436–448.[CrossRef][Medline]
  5. Bowman, W. C., S. Du, C. E. Bauer, and R. G. Kranz. 1999. In vitro activation and repression of photosynthesis gene transcription in Rhodobacter capsulatus. Mol. Microbiol. 33:429–437.[CrossRef][Medline]
  6. Chamberlin, M., R. Kingston, M. Gilman, J. Wiggs, and A. deVera. 1983. Isolation of bacterial and bacteriophage RNA polymerases and their use in synthesis of RNA in vitro. Methods Enzymol. 101:540–568.[Medline]
  7. Dahl, M. K., T. Msadek, F. Kunst, and G. Rapoport. 1992. The phosphorylation state of the DegU response regulator acts as a molecular switch allowing either degradative enzyme synthesis or expression of genetic competence in Bacillus subtilis. J. Biol. Chem. 267:14509–14514.[Abstract/Free Full Text]
  8. Ditta, G., T. Schmidhauser, E. Yakobson, P. Lu, X. W. Liang, D. R. Finlay, D. Guiney, and D. R. Helinski. 1985. Plasmids related to the broad host range vector, pRK290, useful for gene cloning and for monitoring gene expression. Plasmid 13:149–153.[CrossRef][Medline]
  9. Du, S., T. H. Bird, and C. E. Bauer. 1998. DNA binding characteristics of RegA*. A constitutively active anaerobic activator of photosynthesis gene expression in Rhodobacter capsulatus. J. Biol. Chem. 273:18509–18513.[Abstract/Free Full Text]
  10. Du, S., J. L. Kouadio, and C. E. Bauer. 1999. Regulated expression of a highly conserved regulatory gene cluster is necessary for controlling photosynthesis gene expression in response to anaerobiosis in Rhodobacter capsulatus. J. Bacteriol. 181:4334–4341.[Abstract/Free Full Text]
  11. Elsen, S., W. Dischert, A. Colbeau, and C. E. Bauer. 2000. Expression of uptake hydrogenase and molybdenum nitrogenase in Rhodobacter capsulatus is coregulated by the RegB-RegA two-component regulatory system. J. Bacteriol. 182:2831–2837.[Abstract/Free Full Text]
  12. Emmerich, R., K. Panglungtshang, P. Strehler, H. Hennecke, and H. M. Fischer. 1999. Phosphorylation, dephosphorylation and DNA-binding of the Bradyrhizobium japonicum RegSR two-component regulatory proteins. Eur. J. Biochem. 263:455–463.[Medline]
  13. Eraso, J. M., and S. Kaplan. 1996. Complex regulatory activities associated with the histidine kinase PrrB in expression of photosynthesis genes in Rhodobacter sphaeroides 2.4.1. J. Bacteriol. 178:7037–7046.[Abstract/Free Full Text]
  14. Eraso, J. M., and S. Kaplan. 1995. Oxygen-insensitive synthesis of the photosynthetic membranes of Rhodobacter sphaeroides: a mutant histidine kinase. J. Bacteriol. 177:2695–2706.[Abstract/Free Full Text]
  15. Eraso, J. M., and S. Kaplan. 1994. prrA, a putative response regulator involved in oxygen regulation of photosynthesis gene expression in Rhodobacter sphaeroides. J. Bacteriol. 176:32–43.[Abstract/Free Full Text]
  16. Forst, S. A., J. Delgado, and M. Inouye. 1989. DNA-binding properties of the transcription activator (OmpR) for the upstream sequences of ompF in Escherichia coli are altered by envZ mutations and medium osmolarity. J. Bacteriol. 171:2949–2955.[Abstract/Free Full Text]
  17. Galinier, A., A. M. Garnerone, J. M. Reyrat, D. Kahn, J. Batut, and P. Boistard. 1994. Phosphorylation of the Rhizobium meliloti FixJ protein induces its binding to a compound regulatory region at the fixK promoter. J. Biol. Chem. 269:23784–23789.[Abstract/Free Full Text]
  18. Gilles-Gonzalez, M. A., G. S. Ditta, and D. R. Helinski. 1991. A haemoprotein with kinase activity encoded by the oxygen sensor of Rhizobium meliloti. Nature 350:170–172.[CrossRef][Medline]
  19. Hakenbeck, R., and J. B. Stock. 1996. Analysis of two-component signal transduction systems involved in transcriptional regulation. Methods Enzymol. 273:281–300.[Medline]
  20. Harlocker, S. L., L. Bergstrom, and M. Inouye. 1995. Tandem binding of six OmpR proteins to the ompF upstream regulatory sequence of Escherichia coli. J. Biol. Chem. 270:26849–26856.[Abstract/Free Full Text]
  21. Hemschemeier, S. K., U. Ebel, A. Jager, A. Balzer, M. Kirndorfer, and G. Klug. 2000. In vivo and in vitro analysis of RegA response regulator mutants of Rhodobacter capsulatus. J. Mol. Microbiol. Biotechnol. 2:291–300.[Medline]
  22. Hess, J. F., K. Oosawa, N. Kaplan, and M. I. Simon. 1988. Phosphorylation of three proteins in the signaling pathway of bacterial chemotaxis. Cell. 53:79–87.[CrossRef][Medline]
  23. Igo, M. M., A. J. Ninfa, and T. J. Silhavy. 1989. A bacterial environmental sensor that functions as a protein kinase and stimulates transcriptional activation. Genes Dev. 3:598–605.[Abstract/Free Full Text]
  24. Igo, M. M., and T. J. Silhavy. 1988. EnvZ, a transmembrane environmental sensor of Escherichia coli K-12, is phosphorylated in vitro. J. Bacteriol. 170:5971–5973.[Abstract/Free Full Text]
  25. Inoue, K., J. L. Kouadio, C. S. Mosley, and C. E. Bauer. 1995. Isolation and in vitro phosphorylation of sensory transduction components controlling anaerobic induction of light harvesting and reaction center gene expression in Rhodobacter capsulatus. Biochemistry 34:391–396.[CrossRef][Medline]
  26. Joshi, H. M., and F. R. Tabita. 1996. A global two component signal transduction system that integrates the control of photosynthesis, carbon dioxide assimilation, and nitrogen fixation. Proc. Natl. Acad. Sci. USA 93:14515–14520.[Abstract/Free Full Text]
  27. Karls, R. K., J. R. Wolf, and T. J. Donohue. 1999. Activation of the cycA P2 promoter for the Rhodobacter sphaeroides cytochrome c2 gene by the photosynthesis response regulator. Mol. Microbiol. 34:822–835.[CrossRef][Medline]
  28. Lois, A. F., M. Weinstein, G. S. Ditta, and D. R. Helinski. 1993. Autophosphorylation and phosphatase activities of the oxygen-sensing protein FixL of Rhizobium meliloti are coordinately regulated by oxygen. J. Biol. Chem. 268:4370–4375.[Abstract/Free Full Text]
  29. Lukat, G. S., W. R. McCleary, A. M. Stock, and J. B. Stock. 1992. Phosphorylation of bacterial response regulator proteins by low molecular weight phospho-donors. Proc. Natl. Acad. Sci. USA 89:718–722.[Abstract/Free Full Text]
  30. MacGregor, B. J., R. K. Karls, and T. J. Donohue. 1998. Transcription of the Rhodobacter sphaeroides cycA P1 promoter by alternate RNA polymerase holoenzymes. J. Bacteriol. 180:1–9.[Abstract/Free Full Text]
  31. Makino, K., H. Shinagawa, M. Amemura, T. Kawamoto, M. Yamada, and A. Nakata. 1989. Signal transduction in the phosphate regulon of Escherichia coli involves phosphotransfer between PhoR and PhoB proteins. J. Mol. Biol. 210:551–559.[CrossRef][Medline]
  32. Masuda, S., Y. Matsumoto, K. V. Nagashima, K. Shimada, K. Inoue, C. E. Bauer, and K. Matsuura. 1999. Structural and functional analyses of photosynthetic regulatory genes regA and regB from Rhodovulum sulfidophilum, Roseobacter denitrificans, and Rhodobacter capsulatus. J. Bacteriol. 181:4205–4215.[Abstract/Free Full Text]
  33. McCleary, W. R., and J. B. Stock. 1994. Acetyl phosphate and the activation of two-component response regulators. J. Biol. Chem. 269:31567–31572.[Abstract/Free Full Text]
  34. Nierman, W. C., T. V. Feldblyum, M. T. Laub, I. T. Paulsen, K. E. Nelson, J. Eisen, J. F. Heidelberg, M. R. Alley, N. Ohta, J. R. Maddock, I. Potocka, W. C. Nelson, A. Newton, C. Stephens, N. D. Phadke, B. Ely, R. T. DeBoy, R. J. Dodson, A. S. Durkin, M. L. Gwinn, D. H. Haft, J. F. Kolonay, J. Smit, M. B. Craven, H. Khouri, J. Shetty, K. Berry, T. Utterback, K. Tran, A. Wolf, J. Vamathevan, M. Ermolaeva, O. White, S. L. Salzberg, J. C. Venter, L. Shapiro, and C. M. Fraser. 2001. Complete genome sequence of Caulobacter crescentus. Proc. Natl. Acad. Sci. USA 98:4136–4141.[Abstract/Free Full Text]
  35. Ninfa, A. J., and B. Magasanik. 1986. Covalent modification of the glnG product, NRI, by the glnL product, NRII, regulates the transcription of the glnALG operon in Escherichia coli. Proc. Natl. Acad. Sci. USA 83:5909–5913.[Abstract/Free Full Text]
  36. Oh, J. I., and S. Kaplan. 1999. The cbb3 terminal oxidase of Rhodobacter sphaeroides 2.4.1: structural and functional implications for the regulation of spectral complex formation. Biochemistry 38:2688–2696.[CrossRef][Medline]
  37. Oh, J. I., and S. Kaplan. 2001. Generalized approach to the regulation and integration of gene expression. Mol. Microbiol. 39:1116–1123.[CrossRef][Medline]
  38. Ouchane, S., and S. Kaplan. 1999. Topological analysis of the membrane-localized redox-responsive sensor kinase PrrB from Rhodobacter sphaeroides 2.4.1. J. Biol. Chem. 274:17290–17296.[Abstract/Free Full Text]
  39. Pemberton, J. M., I. M. Horne, and A. G. McEwan. 1998. Regulation of photosynthetic gene expression in purple bacteria. Microbiology 144:267–278.[Medline]
  40. Qian, Y., and F. R. Tabita. 1996. A global signal transduction system regulates aerobic and anaerobic CO2 fixation in Rhodobacter sphaeroides. J. Bacteriol. 178:12–18.[Abstract/Free Full Text]
  41. Schilke, B. A., and T. J. Donohue. 1995. ChrR positively regulates transcription of the Rhodobacter sphaeroides cytochrome c2 gene. J. Bacteriol. 177:1929–1937.[Abstract/Free Full Text]
  42. Sganga, M. W., and C. E. Bauer. 1992. Regulatory factors controlling photosynthetic reaction center and light-harvesting gene expression in Rhodobacter capsulatus. Cell 68:945–954.[CrossRef][Medline]
  43. Simon, R., U. Priefer, and A. Puhler. 1983. A broad host range mobilization system for in vivo genetic engineering: transposon mutagenesis in gram negative bacteria. Bio/Technology. 1:784–791.[CrossRef]
  44. Sistrom, W. R. 1960. A requirement for sodium in the growth of Rhodopseudomonas sphaeroides. J. Gen. Microbiol. 22:778–785.[Medline]
  45. Stock, A. M., V. L. Robinson, and P. N. Goudreau. 2000. Two-component signal transduction. Annu. Rev. Biochem. 69:183–215.[CrossRef][Medline]
  46. Stock, J. B., M. G. Surette, M. Levit, and P. Park. 1995. Two-component signal transduction systems: structure-function relationships and mechanisms of catalysis, p.25–51. In J. A. Hoch (ed.), Two-component signal transduction. American Society for Microbiology, Washington, D.C.
  47. Stover, C. K., X. Q. Pham, A. L. Erwin, S. D. Mizoguchi, P. Warrener, M. J. Hickey, F. S. Brinkman, W. O. Hufnagle, D. J. Kowalik, M. Lagrou, R. L. Garber, L. Goltry, E. Tolentino, S. Westbrock-Wadman, Y. Yuan, L. L. Brody, S. N. Coulter, K. R. Folger, A. Kas, K. Larbig, R. Lim, K. Smith, D. Spencer, G. K. Wong, Z. Wu, and I. T. Paulsen. 2000. Complete genome sequence of Pseudomonas aeruginosa PAO1, an opportunistic pathogen. Nature 406:959–964.[CrossRef][Medline]
  48. Swem, L. R., S. Elsen, T. H. Bird, D. L. Swem, H. G. Koch, H. Myllykallio, F. Daldal, and C. E. Bauer. 2001. The RegB/RegA two-component regulatory system controls synthesis of photosynthesis and respiratory electron transfer components in Rhodobacter capsulatus. J. Mol. Biol. 309:121–138.[CrossRef][Medline]
  49. Tiwari, R. P., W. G. Reeve, M. J. Dilworth, and A. R. Glenn. 1996. Acid tolerance in Rhizobium meliloti strain WSM419 involves a two-component sensor-regulator system. Microbiology 142:1693–1704.[Abstract]
  50. Vichivanives, P., T. H. Bird, C. E. Bauer, and F. Robert Tabita. 2000. Multiple regulators and their interactions in vivo and in vitro with the cbb regulons of Rhodobacter capsulatus. J. Mol. Biol. 300:1079–1099.[CrossRef][Medline]
  51. Villarejo, M. R., and I. Zabin. 1974. Beta-galactosidase from termination and deletion mutant strains. J. Bacteriol. 120:466–474.[Abstract/Free Full Text]
  52. Weiss, V., and B. Magasanik. 1988. Phosphorylation of nitrogen regulator I (NRI) of Escherichia coli. Proc. Natl. Acad. Sci. USA 85:8919–8923.[Abstract/Free Full Text]
  53. Wright, G. D., T. R. Holman, and C. T. Walsh. 1993. Purification and characterization of VanR and the cytosolic domain of VanS: a two-component regulatory system required for vancomycin resistance in Enterococcus faecium BM4147. Biochemistry 32:5057–5063.[CrossRef][Medline]
  54. Zeilstra-Ryalls, J., M. Gomelsky, J. M. Eraso, A. Yeliseev, J. O’Gara, and S. Kaplan. 1998. Control of photosystem formation in Rhodobacter sphaeroides. J. Bacteriol. 180:2801–2809.[Free Full Text]

Journal of Bacteriology, January 2002, p. 390-399, Vol. 184, No. 2
0021-9193/01/$04.00+0     DOI: 10.1128/JB.184.2.390-399.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:


This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld