Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Li, L. Articles by Kehoe, D. M. Search for Related Content PubMed PubMed Citation Articles by Li, L. Articles by Kehoe, D. M.

 Previous Article  |  Next Article 

Journal of Bacteriology, November 2008, p. 7241-7250, Vol. 190, No. 21
0021-9193/08/$08.00+0     doi:10.1128/JB.00762-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Abundance Changes of the Response Regulator RcaC Require Specific Aspartate and Histidine Residues and Are Necessary for Normal Light Color Responsiveness

Lina Li and David M. Kehoe^*

Department of Biology, 1001 East Third Street, Indiana University, Bloomington, Indiana 47405

Received 28 May 2008/ Accepted 19 August 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
RcaC is a large, complex response regulator that controls transcriptional responses to changes in ambient light color in the cyanobacterium Fremyella diplosiphon. The regulation of RcaC activity has been shown previously to require aspartate 51 and histidine 316, which appear to be phosphorylation sites that control the DNA binding activity of RcaC. All available data suggest that during growth in red light, RcaC is phosphorylated and has relatively high DNA binding activity, while during growth in green light RcaC is not phosphorylated and has less DNA binding activity. RcaC has also been found to be approximately sixfold more abundant in red light than in green light. Here we demonstrate that the light-controlled abundance changes of RcaC are necessary, but not sufficient, to direct normal light color responses. RcaC abundance changes are regulated at both the RNA and protein levels. The RcaC protein is significantly less stable in green light than in red light, suggesting that the abundance of this response regulator is controlled at least in part by light color-dependent proteolysis. We provide evidence that the regulation of RcaC abundance does not depend on any RcaC-controlled process but rather depends on the presence of the aspartate 51 and histidine 316 residues that have previously been shown to control the activity of this protein. We propose that the combination of RcaC abundance changes and modification of RcaC by phosphorylation may be necessary to provide the dynamic range required for transcriptional control of RcaC-regulated genes.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The response regulator RcaC is part of a multistep phosphorelay-class system that controls the biogenesis of photosynthetic light-harvesting antennae in Fremyella diplosiphon (Calothrix sp. strain PCC 7601) in response to changes in the ratio of ambient red light (RL) to ambient green light (GL) (10). RcaC is a member of a family of large response regulators that range from 500 to 1,600 amino acids long and have complex and unusual arrangements of traditional and nontraditional domains (17, 29). Little is known about the functions of most members of this group or the reason(s) for their complex architecture. Their many domains may allow multiple, diverse signals to be integrated by a single regulator and/or efficiently transduce a signal from one type of domain to another.

Four domains within RcaC have been identified (10, 25). All of these domains have been characterized using a combination of in vivo and in vitro approaches (1, 28, 29). RcaC contains two receiver domains, one at each end of the peptide. While both of these domains contain an aspartate (D) at the position that typically serves as the phosphorylation site in such domains, the light color response is primarily mediated through D51 in the N-terminal receiver module. Adjacent to this module is an OmpR-class DNA binding domain that binds a direct repeat DNA sequence, the L box, within promoters of genes that are activated by RL and an operon that is derepressed in GL. Finally, there is a histidine phosphotransfer domain between the DNA binding region and the C-terminal receiver module. This domain contains a histidine (H) at position 316 that is also required for normal light-regulated gene expression.

In addition to RcaC, the Rca system consists of RcaE, a sensor histidine kinase belonging to the phytochrome-class photoreceptor superfamily, and RcaF, a single-domain response regulator that appears to act after RcaE and before RcaC (25, 26, 39). Site-directed mutant studies have indicated that RcaC is more phosphorylated in RL than in GL (29), leading to activation of the cpcB2A2H2D2 operon (hereafter referred to as cpc2), which encodes a pigmented, RL-absorbing protein called phycocyanin (PC), and repression of the cpeCDESTR operon (referred to as cpeC below), which is required for production of a GL-absorbing, pigmented protein called phycoerythrin (PE). During growth in GL, RcaC does not activate cpc2 transcription or repress cpeC transcription (28). The net result of these light-mediated changes in gene expression is that the pattern of accumulation of RL-absorbing PC or GL-absorbing PE precisely corresponds to the availability of these two colors of light for photosynthesis. This process is called complementary chromatic adaptation (CCA) (19, 27, 37). The Rca system is present in both freshwater and marine cyanobacteria capable of CCA (28), and variants of this pathway are likely used for light-regulated transcriptional control of many other cyanobacterial genes.

An intriguing feature of RcaC is that it is approximately six times more abundant in RL than in GL (29). This observation raises questions about the possible reason(s) for this difference. Since all available data support the hypothesis that RcaC interacts predominantly with the L boxes of both light-induced and -repressed promoters during growth in RL (29), it is reasonable to conjecture that increases in RcaC abundance in RL and/or decreases in RcaC abundance in GL may be needed for effective light regulation of transcription. This observation also raises questions about the level(s) at which these abundance changes are regulated and the mechanism(s) through which they are controlled.

Here, we determined that the kinetics of RcaC abundance changes after RL-GL transitions are very different from the kinetics of RNA accumulation from the cpc2 and cpeC operons, as well as the accumulation of PC and PE. We show that GL-grown cells containing levels of RcaC normally found in RL-grown cells were unable to undergo proper CCA, demonstrating that the reduced amount of RcaC in GL-grown cells is required for normal CCA regulation. We found that mutations at D51 and H316 of RcaC also eliminated the RcaC abundance changes, even in cells containing a functional CCA system, strongly suggesting that these two amino acids contribute to changes in RcaC abundance. Finally, we demonstrated that RcaC abundance changes are regulated at both the level of RNA and the level of protein stability. Overall, our data indicate that this system uses a novel approach for regulating major shifts in transcriptional activity of target promoters by fine-tuning RcaC levels during CCA. They also suggest that the abundance changes may operate in part through sensing of phosphorylation states of specific amino acids within RcaC that are needed for proper CCA function, a control mechanism that has not been described yet for a prokaryotic response regulator.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Strains, growth conditions, and transformation. A short-filament F. diplosiphon mutant designated SF33 (12) with a normal CCA response was used as the wild type. Cells were cultured and were spectrally analyzed using a Beckman DU640B spectrophotometer as previously described (36). Cells were grown using light intensities of approximately 15 µmol photons m^–2 s^–1 unless indicated otherwise and were transformed by electroporation as described previously (29).

Plasmid construction. RcaE H430 was changed to alanine (A) using primers 5' RcaEmutHbox (Table 1) and 3'RcaEmutHbox. Primers 5' RcaE and 3'RcaEmutHbox were used to amplify the 5' region of rcaE encoding a protein with the H at position 430 replaced by A, while primers 3' RcaE and 5'RcaEmutHbox were used to amplify the 3' region of rcaE with the same H-to-A mutation. The two PCR products were combined and used as a template to amplify the whole rcaE coding sequence along with 468 bp upstream of the start codon and 132 bp downstream of the stop codon using the 5' RcaE and 3' RcaE primers. The PCR product was cut with BamHI and inserted into pPL2.7 (11). A clone with rcaE transcribed in the same direction as the kanamycin resistance gene was selected, designated pPLrcaEA430, and used to transform the rcaE null mutant FdBk14 (26, 39) to create mutant RcaEH430A.

View this table: [in this window] [in a new window]

TABLE 1. Primers

Plasmids pPLrcaC and pPLN51/Q316 (29) were used as templates to PCR amplify either the wild-type rcaC coding sequence or the same coding sequence with D51 and H316Q mutations plus 768 bp of the upstream promoter sequence. The primers used in the PCRs were rcaC 3XFLAG 5 PstI and rcaC 3XFLAG 3 XmaI. The W03 plasmid contained a 3XFLAG tag sequence followed by a stop codon after the XmaI site. W03 was cut with PstI and XmaI, and the 4.7-kbp fragment containing the 3XFLAG tag was gel purified and ligated with the PCR products described above cut with the same enzymes to create W03rcaC 3XFLAG and W03N51/Q316 3XFLAG. In these plasmids, the 3XFLAG tag (N terminus DYKDHDGDYKDHDIDYKDDDDK C terminus) was fused in frame to the C terminus of the rcaC coding sequence with a proline and a glycine in between. Each of these plasmids was then used as a template with the same upstream primer, rcaC 3XFLAG 5 PstI, and another downstream primer, rcaC 3XFLAG 3 PstI, to PCR amplify the rcaC coding sequence, including the promoter region and the 3XFLAG tag sequence. The PCR products were cut with PstI and inserted into PstI-cut pPL2.7 to make the autonomously replicating plasmids pPL3XFrcaC and pPL3XFN51/Q316. Clones with rcaC transcribed in the same direction as the kanamycin resistance gene were used for transforming wild-type cells and further analysis.

Primers rcaC copper 5 and rcaC copper 3, as well as plasmids pPLrcaC and pPLN51/Q316/N576 (29), were used to PCR amplify the rcaC coding sequence (either the wild-type sequence or a sequence containing the D51N, H316Q, and D576N mutations). The PCR products were cleaved with NdeI and SacI and inserted into the same sites of pGEM-T-PpetE (8) to generate pGEMTrcaC and pGEMTN51/Q316/N576. Primers PstIPpetEUp and PstIPpetELO were used to PCR amplify the petE promoter followed by either the wild-type or mutated rcaC coding sequence, and the PCR products were inserted into PstI-cut pPL2.7 to make pPLPpetErcaC and pPLPpetEN51/Q316/N576. Clones with rcaC transcribed in the direction opposite the direction of the kanamycin resistance gene were selected for transformation into a rcaC null mutant (CR2) and analyzed further.

The DNA that was derived from PCR amplification of all of the constructs described above was sequenced to ensure that no mutations were present in the amplified sequences or at the ligation junctions. All plasmids and strains used in this study are shown in Table 2.

View this table: [in this window] [in a new window]

TABLE 2. Relevant strains and plasmids

Immunoblot analyses. Extraction of total cellular protein and Western analyses with anti-RcaC antiserum were performed as previously described (29). Western analyses using the anti-FLAG M2-horseradish peroxidase antibody (Sigma, St Louis, MO) were performed according to the manufacturer's instructions, with the following modifications. Membranes were blocked with 5% nonfat milk in TBS buffer (20 mM Tris-HCl [pH 7.5], 140 mM NaCl) containing 0.05% Tween 20 at room temperature for 1 h and then incubated with anti-FLAG M2-horseradish peroxidase antibody diluted 1:1,000 with TBS buffer containing 0.05% Tween 20 overnight at 4°C. The membranes were then washed six times for 5 min each time with TBS buffer containing 0.05% Tween 20. To detect the antibody, SuperSignal West Pico chemiluminescent substrate (Pierce, Rockford, IL) was used according to the manufacturer's instructions, and the images were viewed and quantified using a Kodak Image Station 440.

Kinetics of RcaC abundance change. Wild-type cells were grown in RL or GL to an optical density at 750 nm (OD₇₅₀) of approximately 0.7. Cultures were then switched from RL to GL or from GL to RL. Parallel cultures left in either RL or GL were used as controls. At 0, 4, 10, 16, 22, and 28 h after the light switch, three independent replicates of 50-ml cultures of cells exposed to either constant RL or constant GL or subjected to light color switches were harvested by centrifugation at 5,000 x g for 5 min at 4°C. The cell pellets were resuspended in 3 ml of buffer B (50 mM Tris-HCl [pH 7.5], 10% [wt/vol] glycerol, 0.3 M NaCl, 0.05% [wt/vol] Tween 20, 1 mM phenylmethylsulfonyl fluoride, 0.2 mM benzamidine, 50 µM 6-amino-n-hexanoic acid) and then lysed by two passages through a French pressure cell at 18,000 lb/in² at 4°C. The cell lysates were then centrifuged at 50,000 x g for 30 min at 4°C. For electrophoretic analyses, 100 µg of total protein per lane was loaded onto 12.5% sodium dodecyl sulfate-polyacrylamide gels. Two duplicate gels were loaded and electrophoresed. After electrophoresis, one gel was stained with Coomassie brilliant blue R-250, and the other was transferred to a nitrocellulose membrane and used for Western blot analysis using anti-RcaC antiserum.

Kinetics of phycobiliprotein abundance change. Wild-type cells were grown in RL or GL to an OD₇₅₀ of approximately 0.6 and then either switched from RL to GL, switched from GL to RL, or left in the original light. Phycobiliprotein and chlorophyll contents were determined essentially as previously described (4, 24, 38). For each treatment, six 1.5-ml samples of cells were collected at each time point from cells exposed to the four different light conditions and centrifuged at 13,000 x g for 1 min at room temperature. The pellets were then frozen in liquid nitrogen. For each replicate, three of the pellets were used to determine the chlorophyll concentration by adding 500 ml of 100% methanol, mixing the preparation, and incubating it in the dark for 1 h at 4°C, which was followed by centrifugation at 10,000 x g for 10 min. The extraction procedure was repeated once, the supernatants were pooled, and the absorbance at 665 nm was recorded. The remaining three pellets were each resuspended in 1 ml of STES (50 mM Tris-HCI [pH 8.0], 50 mM NaCl, 10 mM EDTA [pH 8.0], 250 mM sucrose) supplemented with 5 mg/ml lysozyme. Samples were incubated at room temperature for 30 min, and after centrifugation at 13,000 x g for 5 min, the supernatants were collected. The supernatant absorbance at 565 nm, 620 nm, and 650 nm were used to calculate PE, PC, and allophycocyanin (AP) concentrations as previously described (38). After the averages for three sample values were normalized on the basis of the chlorophyll concentrations, the PE/AP and PC/AP ratios were calculated.

Copper induction. rcaC null mutant (CR2) (29) cells transformed with pPLPpetErcaC or pPLPpetEN51/Q316/N576 were grown to an OD₇₅₀ of 0.7 in BG-11 containing 10 µg/ml kanamycin in the presence of 50 µmol photons m^–2 s^–1 GL. Cells were harvested by centrifugation at 5,000 x g for 5 min at room temperature, and then they were resuspended in fresh BG-11 that contained 10 µg/ml kanamycin and either lacked copper sulfate or contained various concentrations (see below) of copper sulfate and were grown in plastic culture tubes in RL and GL. Cells were harvested at an OD₇₅₀ of 0.8, and the total cellular protein was extracted and analyzed as described previously (29).

qRT-PCR amplification analyses. RNA was isolated from wild-type cells grown in RL and GL as described previously (36). The concentration and purity of the RNA were determined spectrophotometrically using a Beckman DU640B spectrophotometer. Purified RNA was treated with a Turbo DNA-free kit (Ambion, Austin, TX) to remove genomic DNA according to the manufacturer's instructions. Reverse transcription was performed by incubating 5 µg of RNA with 5 µg of random hexamers (Qiagen, Valencia, CA) for 10 min at 70°C and then with 200 U of Superscript II (Invitrogen, Carlsbad, CA) in the presence of 40 U of RNaseOUT (Invitrogen, Carlsbad, CA) for 2 h at 42°C. RNA was removed by RNase H (Invitrogen, Carlsbad, CA) treatment for 20 min at 37°C. Quantitative real-time PCR (qRT-PCR) amplification was performed separately with two sets of primers for each cDNA sample. Primers rcaC RT9 and rcaC RT10 were used to detect rcaC cDNA. Primers 23S RT1 and 23S RT2 were used to detect 23S rRNA cDNA as a control. A 25-µl PCR mixture containing 12.5 µl Platinum SYBR green qRT-PCR Supermix-UDG (Invitrogen, Carlsbad, CA), 0.5 µl of a 10 µM forward primer solution, 0.5 µl of a 10 µM reverse primer solution, 0.5 µl of a 500 nM fluorescein (Bio-Rad, Hercules, CA) solution, and 10 µl of a cDNA solution was amplified using an iCycler iQ multicolor real-time PCR detection system (Bio-Rad, Hercules, CA).

The PCR amplification program was 50°C for 2 min, followed by denaturation at 94°C for 10 min, 45 cycles of 94°C for 15 s, 50°C for 30 s, and 60°C for 30 s, and then 80 cycles of 55°C for 1 min with a 0.5°C increase in the temperature for every additional cycle that was performed to collect the melting curve data. The PCR products from one experiment were analyzed by agarose gel electrophoresis to ensure that a single product that was the correct size had been generated. In order to examine the efficiency of the qRT-PCR amplifications, standard curves were generated with a 1:10 serial dilution of sample cDNA for 23S rRNA and a 1:5 dilution for rcaC. A negative cDNA control, in which no Superscript II was added during the reverse transcription step, and another negative control, in which 10 µl of water was used instead of cDNA in the PCR amplification mixture, were added to each experiment to assess the degree of genomic DNA contamination, as well as the amount of contamination present in the reagents that were used. Eight independently isolated RNA samples from both RL- and GL-grown cultures were used in the qRT-PCR assay for analysis of both the rcaC and 23S RNA levels. The qRT-PCR results were analyzed using the method of Pfaffl (33).

Analysis of RcaC protein stability. [³⁵S]methionine was used to assess the efficiency of inhibition of protein synthesis by 20 µg ml^–1 chloramphenicol (CAP) in F. diplosiphon cells. Four 1.5-ml samples from a 50-ml culture of wild-type cells grown to an OD₇₅₀ of 0.5 in RL were used for each experiment. Three of these samples were treated with 20 µg ml^–1 CAP for 5 min, while no CAP was added to the fourth sample. After 5 min, 15 µCi of the Tran ³⁵S-Label metabolic labeling reagent containing [³⁵S]methionine (MP Biomedicals, Irvine, CA) was added to each tube. One sample containing 20 µg ml^–1 CAP was harvested immediately by centrifugation at 13,000 x g for 1 min to assess the extent of physical trapping of the ³⁵S label. The control containing no CAP and one sample containing 20 µg ml^–1 CAP were incubated in RL for 4 h, while the final sample containing 20 µg ml^–1 CAP was immediately moved to GL and incubated for 4 h. Cells from the three tubes were then harvested as described above. For all samples, supernatants were removed, and cell pellets were frozen in dry ice and then thawed three times before 50 µl of buffer B supplemented with 0.25 U/µl mutanolysin (Sigma-Aldrich, St. Louis, MO) was added. Samples were incubated for 30 min at room temperature and centrifuged at 13,000 x g for 1 min, and the supernatants were collected. Total protein was precipitated on ice for 30 min using 10% trichloroacetic acid. The precipitate from each sample was filtered onto a GF/C glass filter (Whatman, Brentford, Middlesex, United Kingdom), and total counts were recorded using an LS6500 scintillation system (Beckman, Fullerton, CA).

For analysis of RcaC stability, wild-type cells were grown in RL to an OD₇₅₀ of approximately 0.8. Four 50-ml cultures grown from a single starter culture were used in each experiment. One of these cultures was treated with 20 µg ml^–1 CAP and harvested immediately. Another culture was kept in RL for an additional 4 h without CAP before cells were harvested. The final two cultures were treated with 20 µg ml^–1 CAP and either kept in RL or switched to GL for an additional 4 h before cells were harvested. All cells were harvested by centrifugation at 5,000 x g for 5 min at 4°C. Total protein was then extracted from each culture and subjected to polyacrylamide gel electrophoresis and Western blot analysis to detect the RcaC protein (29). Equal protein loads were confirmed by Coomassie brilliant blue staining of equivalently loaded gels run in parallel with the gels used in Western blot analyses.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The kinetics of RcaC abundance changes are different from known CCA responses. The light-driven change in RcaC abundance might facilitate CCA-regulated alterations in the transcriptional activity and RNA accumulation from cpc2 and cpeBA, which change dramatically within 2 to 4 h after a light shift (30). Western blots were used to measure RcaC abundance over time after cells were shifted between RL and GL (Fig. 1A). Most changes in RcaC abundance occurred within 21 h after a change in the color of light in either direction, and the half-times for these changes were approximately 6 h for the shift from RL to GL and approximately 12 h for the shift from GL to RL (Fig. 1B).

View larger version (41K): [in this window] [in a new window]

FIG. 1. Kinetics of RcaC light-regulated abundance changes. (A) Western blot analyses of RcaC levels during growth under constant light conditions (0 h) or at various times after switches from RL to GL (upper blot) or from GL to RL (lower blot). The positions of molecular mass markers are indicated on the left. (B) Densitometric quantification of RcaC levels detected by Western blotting in cells grown in RL (filled squares), in GL (filled circles), in RL and then switched to GL (open squares), and in GL and then switched to RL (open circles) and harvested at 0, 4, 10, 16, 22, and 28 h after the light switch. The y axis indicates the amount of RcaC detected, which was normalized to the amount of RcaC under constant RL growth conditions. Each data point indicates the average for three independent replicates. The error bars indicate standard errors.

We then compared the kinetics of RcaC abundance changes shown in Fig. 1 to the timing of the shifts in the cellular PC/PE ratio after switches in the color of light during CCA. The kinetics of the shifts in PC and PE abundance were determined by extracting these phycobiliproteins and comparing their levels to the level of another phycobiliprotein, AP, whose abundance does not change significantly during CCA (13, 21), and the results were expressed as the ratio of PC to AP or the ratio of PE to AP. The complete CCA-driven shifts in PC and PE abundance took approximately 150 h, regardless of the direction of the shift in the light color (Fig. 2). The half-time for this process in both directions was close to 50 h. This is four to six times longer than the half-time for RcaC abundance changes during CCA (Fig. 1).

View larger version (31K): [in this window] [in a new window]

FIG. 2. Time course of the changes in the relative abundances of (A) PE and (B) PC in response to changes in the ambient light color. Levels of PC and PE were determined spectrophotometrically at various times after wild-type cells were shifted either from RL to GL or from GL to RL. The y axes indicate the cellular ratio of (A) PE to AP or (B) PC to AP. The experiments were repeated two or three times, and the error bars indicate the standard errors.

RcaC abundance changes require a functional RcaE photoreceptor. We have previously reported that RcaC is approximately six times more abundant in RL-grown cells than in GL-grown cells and that the abundance changes are eliminated if substitutions are made at either D51 or H316 of RcaC, two amino acids that are crucial for a normal CCA response (29). We further examined the mechanism(s) by which light color controls the level of RcaC by determining the abundance of RcaC in several different types of rcaE mutants.

The rcaE null mutant FdBk14 was grown in both RL and GL, and the RcaC abundance was checked (26, 39). In both light conditions, RcaC accumulated at intermediate levels relative to the levels found in wild-type cells grown in RL (Fig. 3A). This result suggested that RcaE could affect the differential accumulation of RcaC in RL and GL either through its physical presence or by its presumed ability to control the phosphorylation state of RcaC. We therefore examined RcaC levels in two additional types of rcaE mutants. The first mutant, the FdR9 mutant, was red during growth in RL and GL and contained an insertion between the H box and the ATP binding region of the histidine kinase domain. This insertion was predicted to lead to the synthesis of a truncated form of RcaE that dephosphorylated the Rca pathway in both RL and GL, since similar mutations in other histidine kinases result in the loss of kinase activity but retention of phosphatase activity (3, 9, 25). The second mutant, an RcaE site-directed mutant, RcaE-H430A, had an alanine substitution at the histidine at position 430 of RcaE, which is within the H box and is the likely site of phosphorylation during CCA signal transduction. This mutant was also red during growth in both RL and GL (Fig. 3B), indicating that RcaC was predominantly unphosphorylated in the cells (29). In both FdR9 and RcaE-H430A, the levels of RcaC were the same and relatively high in cells grown in RL and GL (Fig. 3A).

View larger version (35K): [in this window] [in a new window]

FIG. 3. RcaC levels in rcaE mutants. (A) Representative Western blot analysis of RcaC levels in RL- and GL-grown wild-type strain and rcaE null (FdBk14), rcaE truncation (FdR9), rcaE site-directed (pPLrcaEA430), and rcaC null (CR2) mutants. The positions of molecular mass markers are indicated on the left. Analyses were repeated either two or three times. (B) Representative whole-cell absorbance spectra of RL-grown (solid line) and GL-grown (dashed line) FdBk14 cells harboring the pPLrcaEA430 plasmid. PE and PC absorbance peaks are indicated.

RcaC abundance changes depend on the presence of the D51 and H316 residues themselves rather than on the CCA processes that they regulate. Previous work has shown that substitutions at both D51 and H316 lead to the production of forms of RcaC that are equally abundant during growth in RL and growth in GL (29). Based on this finding, together with the RcaE mutant results shown in Fig. 3, it is reasonable to hypothesize that the light regulation of RcaC abundance occurs via sensing of the phosphorylation state of RcaC, particularly at residues D51 and H316. However, the mutants are also not capable of normal CCA, making it possible that the loss of the light-regulated changes in RcaC abundance in the mutant cells were the indirect result of the loss of a CCA-regulated system that provided feedback control of RcaC abundance. This possibility was tested by producing C-terminally 3XFLAG-tagged versions of RcaC-D51N/H316Q and wild-type RcaC in wild-type cells. Whole-cell absorption spectral analyses of a number of transformed lines producing these proteins demonstrated that CCA, as measured by shifts in the PC/PE ratio during growth in RL versus growth in GL, functioned properly in all of these lines (Fig. 4A and 4B).

View larger version (19K): [in this window] [in a new window]

FIG. 4. RcaC abundance change requires D51 and H316. (A and B) Representative whole-cell absorbance spectra of wild-type (WT) cells transformed with (A) pPL3XFrcaC or (B) pPL3XFN51/Q316 and grown in RL (solid line) and GL (dashed line). PE and PC absorbance peaks are indicated. (C) Representative Western blot of wild-type cells transformed with either pPL3XFrcaC or pPL3XFN51/Q316 and grown in RL and GL. Anti-FLAG antibody was used to detect both of these forms of RcaC. (D) Quantification of 3XFLAG-tagged RcaC abundance, indicated by the ratio of the abundance of each protein in RL to the abundance of each protein in GL. The bars indicate the averages of six independent experiments, and the error bars indicate standard errors.

Transformed cells expressing either the wild-type or D51N/H316Q form of 3XFLAG-tagged RcaC were grown in RL and GL, and the levels of the proteins were detected using an anti-FLAG antibody (Fig. 4C and 4D). The cell lines producing wild-type RcaC-3XFLAG contained about 3.5-fold more of this protein in RL than in GL. Conversely, the cell lines producing RcaC-D51N/H316Q-3XFLAG contained approximately equal amounts of this protein during growth in RL and growth in GL. These results suggest that D51 and H316, rather than a downstream event that is part of the CCA response, are required for the light-mediated changes in RcaC abundance.

Both the D51, H316, and N576 residues and the abundance of RcaC regulate its activity. We assessed the importance of the levels of RcaC and its presumed phosphorylation state in the regulation of CCA by placing wild-type rcaC under the control of the copper-inducible petE promoter (6, 8) and transforming the resulting construct into the rcaC null mutant CR2. By controlling the copper concentration in the medium, we were able to manipulate the cellular level of RcaC in a light-independent fashion. Precise RcaC levels in the various transformed lines could be obtained consistently with specific concentrations of exogenously supplied copper (Fig. 5C).

View larger version (23K): [in this window] [in a new window]

FIG. 5. Effect of RcaC overexpression on CCA. (A and B) Representative whole-cell absorbance spectra for the CR2 mutant transformed with pPLPpetErcaC and grown in (A) RL and (B) GL with three different concentrations of copper (black line, 0 µM copper; dashed line, 0.3 µM copper; gray line, 1 µM copper). PE and PC absorbance peaks are indicated. Each scan is representative of the results for at least three independently transformed lines. (C) Representative Western blots showing RcaC abundance during growth in RL and GL for wild-type cells and cells of the CR2 mutant transformed with pPLPpetErcaC in the presence of three different concentrations of copper. Each Western blot analysis was repeated with at least two independently transformed lines.

For transformants expressing RcaC and grown in RL, increasing the RcaC level above the level present in wild-type cells had no discernible effect on the process of CCA (compare Fig. 5A and 5C to Fig. 4A). This suggests that in RL, the level of RcaC present in wild-type cells is sufficient to sustain the maximal CCA response. Because of the strength of the petE promoter, we were unable to produce RcaC at levels below those found in cells grown in RL and thus could not determine the effect of production of below-normal levels of RcaC in RL. Expression of RcaC in GL-grown cells at a level slightly above that found in wild-type RL-grown cells slightly inhibited the normal GL CCA response (compare Fig. 5B and C to Fig. 4A). Driving RcaC levels even higher strongly shifted the cells toward an RL phenotype, with decreased PE accumulation and increased PC accumulation (Fig. 5B and 5C). These results demonstrate that the lower abundance of RcaC present in wild-type cells grown in GL is required to obtain a normal CCA response in that color of light.

The RcaC levels in RL- and GL-grown cells were virtually the same in the absence of copper (Fig. 5C), but the CCA states were very different under these two conditions (Fig. 5A and 5B). This demonstrates that there must be differences in the activity of RcaC in these two light conditions that is controlled by a process that is independent of the abundance of this protein. These data, along with previous work, indicate that this additional process is the control of the phosphorylation state of the three amino acids in RcaC through which CCA is regulated, since cells expressing the RcaC-N51/Q316/N576 mutant at levels comparable to the levels observed in the absence of copper here (Fig. 5C) are not capable of undergoing CCA (29).

rcaC transcript levels are regulated by light. We have previously observed that RcaC abundance is greater during growth in RL than during growth in GL (29). We sought to determine whether this difference is partially or completely the result of corresponding changes in rcaC RNA levels under these two light conditions. RcaC abundance was quantified by Western blotting. Although the ratio of the RcaC level in RL to the RcaC level in GL was different in different experiments (data not shown), multiple measurements demonstrated that RcaC was 5.8 times more abundant in RL-grown cells than in GL-grown cells (Fig. 6A). We then performed qRT-PCR amplification of rcaC transcripts from RNA samples isolated from RL- and GL-grown cells. rcaC transcripts were slightly more than threefold more abundant in RL-grown cells than in GL-grown cells (Fig. 6B). Thus, the higher level of rcaC RNA in RL-grown cells than in GL-grown cells accounted for slightly more than one-half of the 5.8-fold difference in RcaC protein abundance measured under these two light conditions.

View larger version (17K): [in this window] [in a new window]

FIG. 6. (A) Relative RcaC abundance in RL- and GL-grown F. diplosiphon cells as determined by Western blot analyses. The RcaC protein level in GL-grown cells was defined as 1, while the RcaC level in RL-grown cells was expressed relative to the level in GL-grown cells. The averages for 29 Western blot analyses are indicated by the bars, and the error bars indicate standard errors. (B) Relative rcaC transcript abundance in RL- and GL-grown cells as determined by qRT-PCR amplification. The rcaC transcript level in cells grown in GL was defined as 1, and the level of rcaC RNA in RL-grown cells was expressed relative to the level in GL-grown cells. The bars indicate the averages for eight qRT-PCR experiments, and all values were normalized using 23S rRNA levels. The error bars indicate standard errors.

RcaC protein is less stable in GL than in RL. The differences in rcaC RNA levels (Fig. 6B) explained some, but not all, of the RcaC protein abundance differences (Fig. 6A) between growth in RL and growth in GL. Therefore, we examined the effect of growth in these two types of light on the stability of RcaC, utilizing CAP to inhibit protein synthesis. Although the method that we used has been reported to inhibit more than 90% of new protein synthesis (32), in our experiments an average of approximately 70% of new protein synthesis was inhibited, as measured by incorporation of [³⁵S]methionine (data not shown). Cultures were grown in RL and then either kept in RL or switched to GL. The cultures kept in RL were either treated or not treated with CAP, while those switched to GL were treated with CAP just prior to the switch. An RL-grown culture treated with CAP and then harvested immediately was also included as a control. Four hours after CAP treatment, cells were harvested, and RcaC protein abundance was measured using Western blot analysis. The RcaC levels in the cultures that were kept in RL and treated with CAP remained essentially unchanged (Fig. 7), suggesting that the RcaC protein is relatively stable under RL growth conditions. RcaC abundance in cells grown under the same light conditions and not treated with CAP also changed little over the same period. However, after 4 h, the level of RcaC protein found in cultures switched from RL to GL after CAP treatment was only approximately 65% of the level of RcaC protein found in cultures that were treated with CAP and kept in RL. These data strongly suggest that RcaC is less stable in cells that have been switched to GL from RL than in cells maintained in continuous RL.

View larger version (29K): [in this window] [in a new window]

FIG. 7. RcaC protein is less stable during growth in GL than during RL growth. Quantification of Western blot results provided relative RcaC levels in wild-type cells grown in the light conditions indicated. The bars indicate the means of four independent experiments. The error bars indicate standard errors of the means. CAP20, 20 µg ml–1 CAP.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
A central goal of the studies presented here was to determine whether light color-regulated changes in RcaC abundance (29) are important for proper control of CCA. By analyzing the kinetics of RcaC abundance changes after switches between RL and GL (Fig. 1), we determined that this process is essentially complete within 21 h and is much more rapid than light color-induced changes in PC and PE levels (Fig. 2), which require approximately 150 h. However, the rate of RcaC abundance changes was significantly lower than the rate of CCA-mediated changes in the cpc2, cpeBA, and cpeC RNA levels. During CCA, the changes in cpc2 mRNA levels after shifts to both RL and GL are complete within 2 h, while increases in cpeBA mRNA levels after a shift from RL to GL take approximately 4 h (31). There have been two reports on the time required for completion of cpeBA mRNA abundance changes after exposure to RL, 4 h (15) and more than 16 h (30). The decrease in the cpeCDE mRNA level is complete within 8 h after a shift from GL to RL (15). Even though several of the studies described above involved treating cells in the dark after the reversing light treatment, which may have affected RNA degradation rates or other factors that affect RNA abundance, the changes in CCA-driven RcaC abundance are clearly slower than the changes in the RNA levels of RcaC-controlled genes. This demonstrates that CCA regulation, as measured by PC and PE abundance changes, is not strictly tied to RcaC abundance and suggests that reversible modification of RcaC, possibly through phosphorylation, plays an important role in controlling RcaC activity. This fact is reinforced by the results shown in Fig. 5, which show that the PC/PE ratio obtained for transformed cultures when they were grown in RL was always greater than the PC/PE ratio obtained when they were grown in GL, even when RcaC levels were very high and comparable in the cultures. This, along with previous results (29), strongly suggests that RcaC likely requires phosphorylation for a complete CCA response in RL.

However, the data in Fig. 5 also show that, at least in GL, normal CCA requires the proper RcaC abundance. Increasing the level of RcaC in GL-grown cells to that normally present in RL-grown cells (0 µM) (Fig. 5B) prevented the cells from achieving the ratio of PE to PC normally achieved in wild-type cells (Fig. 4A), demonstrating that RcaC levels must be reduced in GL in order for the normal CCA response to occur. Since RcaC is apparently not significantly phosphorylated in GL (29) (Fig. 3B) and we obtained data comparable to the data shown in Fig. 3B using an RcaCN51/Q316/N576 triple mutant (L. Li and D. M. Kehoe, unpublished data), it follows that elevated levels of apparently nonphosphorylated RcaC can still activate CCA-responsive promoters. This conclusion is further supported by the results of in vitro footprinting studies showing that nonphosphorylated RcaC is capable of specifically binding the L box (28). Similar results have been obtained for OmpR, which in its nonphosphorylated form still binds target DNA with relatively high affinity (10% of the affinity of the phosphorylated form) (20). However, the change in OmpR abundance is less than twofold at low osmolarity compared with high osmolarity (7, 16), which is much less than the change in RcaC abundance during CCA (29) (Fig. 6A). Since we were unable to reduce the abundance of RcaC below the abundance normally found in RL-grown cells, we could not examine the effect of such a change. However, it was clear that the CCA response in RL was saturated at wild-type RcaC levels, since further increases in RcaC abundance had no effect on the pattern of PE or PC accumulation under these light conditions (Fig. 5A and C).

We also sought to determine the level at which RcaC light-regulated abundance changes were regulated and found that they are controlled at both the RNA and protein levels (Fig. 6). It is not yet clear if rcaC RNA abundance is controlled transcriptionally or posttranscriptionally or both. RcaC controls both RL- and GL-activated promoters by binding an L box (28), so feedback regulation of rcaC transcription could occur via an L box upstream of this gene. However, there is no identifiable L box in either orientation in the region of DNA upstream of rcaC (data not shown). This does not eliminate the possibility that there is a direct feedback control mechanism involving a degenerate L box or a different binding site.

Changes in RcaC protein abundance appear to be due at least in part to decreased stability during growth in GL (Fig. 7), suggesting that a light color-regulated protease activity is involved in this process. The photoreceptor/sensor RcaE clearly plays an important role in controlling such activity (Fig. 3 and 4) (29). There are three likely ways that RcaE controls RcaC abundance. The first way is through direct interactions between RcaE and RcaC. Several studies have suggested that in Caulobacter crescentus the sensor kinase CckA might interact with the essential response regulator CtrA, protecting it from proteolysis by ClpXP (22, 34). In our system, the absence of RcaE leads to an intermediate level of RcaC in both RL and GL (Fig. 3A). If interaction between RcaE and RcaC is an important determinant of RcaC levels, this result suggests that interactions between RcaE and RcaC may actually lead to the loss of RcaC.

The second way is by RcaE controlling the phosphorylation state of RcaC. Although phosphorylation of RcaC has not been demonstrated directly yet, this possibility is strongly supported by the lack of RcaC abundance changes in rcaE mutants FdR9 and FdBk14/pPLrcaEA430 (Fig. 3A) and by a similar lack in the 3XFLAGRcaCN51/Q316 mutant in the wild-type background (Fig. 4C and D). We propose a model in which RcaC is phosphorylated and therefore more abundant in RL and is not phosphorylated and is therefore less abundant in GL. Thus, an rcaE null mutant, in which RcaC has been proposed to be partially activated but not light responsive (25), would be expected to have intermediate, non-light-regulated levels of RcaC, the phenotype actually observed (Fig. 3A). Why are RcaC levels so high in the rcaE truncation mutant FdR9 and the site-directed mutant rcaEH430A, in which RcaC should be dephosphorylated (Fig. 3A)? It is possible that RcaE has multiple roles in regulating RcaC abundance. In this scenario, dephosphorylation of RcaC increases its interactions with RcaE, making it more likely to be degraded. Phosphorylation of RcaC by RcaE leads to decreased RcaC-RcaE interactions, thus stabilizing it. If such an RcaE-RcaC interaction is phosphorylation dependent, it is reasonable to expect that the phosphorylation sites are important for the interaction. The nuclear magnetic resonance structure of EnvZ suggests that the phosphorylation site H residue of this sensor probably interacts with both the ATP binding domain of EnvZ and the receiver domain of OmpR (41). Thus, for the two rcaE mutants described above, even though there is no apparent phosphorylation of RcaC, there also may be sufficiently reduced interactions between RcaE and RcaC to prevent the destabilization of RcaC, as observed. No significant degradation would occur without such interactions, leading to higher levels of RcaC. If this is correct, the effect of this RcaE-RcaC interaction on RcaC degradation would be opposite the effect proposed for the CckA-CtrA interaction (22, 34).

The third way that RcaE could regulate RcaC abundance is through an indirect mechanism, such as a CCA-controlled proteolysis pathway that degrades RcaC only during growth in GL, regardless of the RcaC phosphorylation state. We expect that such a mechanism, which could be RL repressible or GL inducible, would not be detectable using RcaE or RcaC mutants, since the effects on the phosphorylation of these proteins (29) (Fig. 3A) would be inseparable from the loss of subsequent CCA responses. However, we were able to separate these effects by expressing epitope-tagged versions of wild-type and mutant forms of RcaC in a wild-type background. We found that even when CCA was functioning properly (Fig. 4A and B), the abundance of the RcaC mutant having substitutions at D51 and H315 was not light responsive (Fig. 4C). This clearly demonstrates that it is the presence of D51 and H316 that directly affects RcaC abundance rather than an RcaC-controlled process that feeds back to regulate RcaC abundance. It is also noteworthy that the 3XFLAG-tagged RcaC/D51N/H316Q levels in both RL- and GL-grown cells were equivalent to the 3XFLAG-tagged RcaC levels in GL-grown cells. This result is consistent with our model indicating that in GL RcaC is not phosphorylated and is less abundant than it is in RL.

Overall, our data demonstrate that for RcaC, light color-driven abundance changes operate through D51 and H316, residues that are critical for CCA regulation, and they strongly suggest that the phosphorylation state of these residues affects the stability of this response regulator. Proteolysis plays an important regulatory role in bacteria (18, 23), but to date no prokaryotic response regulator whose stability is regulated by its phosphorylation state has been identified. In C. crescentus, the response regulator CtrA is proteolysed in a cell cycle-specific fashion just prior to the start of DNA replication (14). Phosphorylation of the D51 residue of CtrA was initially considered a possible trigger for the proteolytic event, but this possibility was eliminated when proteolysis of CrtA D51A and D51E mutants was found to occur at the correct time in the cell cycle (14, 34). However, there are examples of eukaryotic response regulators whose degradation is phosphorylation dependent (35, 40), so such a control mechanism for prokaryotic systems seems likely.

The results presented here suggest that the CCA regulatory activity of RcaC is controlled by both its phosphorylation state and abundance level. Why might these two levels of control be needed? It is possible that the difference in L box binding affinities between the hypothesized phosphorylated and nonphosphorylated forms of RcaC is not sufficient to regulate the transcriptional responses of some of the genes that it controls. This may be particularly true for the regulation of the RcaC-controlled cpc2 operon, which is activated 50- to 100-fold in RL (PC-containing light-harvesting antennae comprise 60% of the total soluble protein in the cell) and virtually shut off in GL (2, 5, 36). The light-regulated changes in RcaC abundance might be required to achieve the additional dynamic range needed in this transcriptional response. In RL, a higher concentration of RcaC would increase L box occupancy and transcription rates, while lower RcaC levels in GL would have the opposite effect.

 
This work was supported by National Science Foundation grant MCB-0519433 to D.M.K., which we gratefully acknowledge.

We thank Rick Alvey, Andrian Gutu, and Ryan Bezy for thoughtful discussions and comments on the manuscript. We also thank Wenchao Wang (Strome Laboratory, Indiana University) for providing the W03 plasmid and Sean M. Callahan for generously providing the pGEM-T-PpetE plasmid.

 
* Corresponding author. Mailing address: Department of Biology, Indiana University, 1001 E. Third Street, Bloomington, IN 47405. Phone: (812) 856-4715. Fax: (812) 855-6705. E-mail: dkehoe{at}indiana.edu

Published ahead of print on 29 August 2008.

Present address: School of Dentistry, CHS 20-118, University of California, Los Angeles, CA 90095.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Alvey, R. M., R. P. Bezy, N. Frankenberg-Dinkel, and D. M. Kehoe. 2007. A light regulated OmpR-class promoter element coordinates light harvesting protein and chromophore biosynthetic enzyme gene expression. Mol. Microbiol. 64:319-332.[CrossRef][Medline]
2
2. Alvey, R. M., J. A. Karty, E. Roos, J. P. Reilly, and D. M. Kehoe. 2003. Lesions in phycoerythrin chromophore biosynthesis in Fremyella diplosiphon reveal coordinated light regulation of apoprotein and pigment biosynthetic enzyme gene expression. Plant Cell 15:2448-2463.[Abstract/Free Full Text]
3
3. Atkinson, M., and A. J. Ninfa. 1993. Mutational analysis of the bacterial signal-transducing protein kinase/phosphatase nitrogen regulator II (NRII or NtrB). J. Bacteriol. 175:7016-7023.[Abstract/Free Full Text]
4
4. Balabas, B. E., B. L. Montgomery, L. E. Ong, and D. M. Kehoe. 2003. CotB is essential for complete activation of green light-induced genes during complementary chromatic adaptation in Fremyella diplosiphon. Mol. Microbiol. 50:781-793.[CrossRef][Medline]
5
5. Bogorad, L. 1975. Phycobiliproteins and complementary chromatic adaptation. Annu. Rev. Plant Physiol. 26:369-401.
6
6. Buikema, W. J., and R. Haselkorn. 2001. Expression of the Anabaena hetR gene from a copper-regulated promoter leads to heterocyst differentiation under repressing conditions. Proc. Natl. Acad. Sci. USA 98:2729-2734.[Abstract/Free Full Text]
7
7. Cai, S. J., and M. Inouye. 2002. EnvZ-OmpR interaction and osmoregulation in Escherichia coli. J. Biol. Chem. 277:24155-24161.[Abstract/Free Full Text]
8
8. Callahan, S. M., and W. J. Buikema. 2001. The role of HetN in maintenance of the heterocyst pattern in Anabaena sp. PCC 7120. Mol. Microbiol. 40:941-950.[CrossRef][Medline]
9
9. Cavicchioli, R., I. Schroder, M. Constanti, and R. P. Gunsalus. 1995. The NarX and NarQ sensor-transmitter proteins of Escherichia coli each require two conserved histidines for nitrate-dependent signal transduction to NarL. J. Bacteriol. 177:2416-2424.[Abstract/Free Full Text]
10
10. Chiang, G. G., M. R. Schaefer, and A. R. Grossman. 1992. Complementation of a red-light-indifferent cyanobacterial mutant. Proc. Natl. Acad. Sci. USA 89:9415-9419.[Abstract/Free Full Text]
11
11. Chiang, G. G., M. R. Schaefer, and A. R. Grossman. 1992. Transformation of the filamentous cyanobacterium Fremyella diplosiphon by conjugation or electroporation. Plant Physiol. Biochem. 30:315-325.
12
12. Cobley, J. G., E. Zerweck, R. Reyes, A. Mody, J. R. Seludo-Unson, H. Jaeger, S. Weerasuriya, and S. Navankasattusas. 1993. Construction of shuttle plasmids which can be efficiently mobilized from Escherichia coli into the chromatically adapting cyanobacterium, Fremyella diplosiphon. Plasmid 30:90-105.[CrossRef][Medline]
13
13. Conley, P. B., P. G. Lemaux, T. L. Lomax, and A. R. Grossman. 1986. Genes encoding major light-harvesting polypeptides are clustered on the genome of the cyanobacterium Fremyella diplosiphon. Proc. Natl. Acad. Sci. USA 83:3924-3928.[Abstract/Free Full Text]
14
14. Domian, I. J., K. C. Quon, and L. Shapiro. 1997. Cell type-specific phosphorylation and proteolysis of a transcriptional regulator controls the G1-to-S transition in a bacterial cell cycle. Cell 90:415-424.[CrossRef][Medline]
15
15. Federspiel, N. A., and A. R. Grossman. 1990. Characterization of the light-regulated operon encoding the phycoerythrin-associated linker proteins from the cyanobacterium Fremyella diplosiphon. J. Bacteriol. 172:4072-4081.[Abstract/Free Full Text]
16
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
17. Galperin, M. Y., A. N. Nikolskaya, and E. V. Koonin. 2001. Novel domains of the prokaryotic two-component signal transduction systems. FEMS Microbiol. Lett. 203:11-21.[CrossRef][Medline]
18
18. Gottesman, S. 2003. Proteolysis in bacterial regulatory circuits. Annu. Rev. Cell Dev. Biol. 19:565-587.[CrossRef][Medline]
19
19. Grossman, A. R. 2003. A molecular understanding of complementary chromatic adaptation. Photosynth. Res. 76:207-215.[CrossRef][Medline]
20
20. Head, C. G., A. Tardy, and L. J. Kenney. 1998. Relative binding affinities of OmpR and OmpR-phosphate at the ompF and ompC regulatory sites. J. Mol. Biol. 281:857-870.[CrossRef][Medline]
21
21. Houmard, J., V. Capuano, T. Coursin, and N. Tandeau de Marsac. 1988. Genes encoding core components of the phycobilisome in the cyanobacterium Calothrix sp. strain PCC 7601: occurrence of a multigene family. J. Bacteriol. 170:5512-5521.[Abstract/Free Full Text]
22
22. Jacobs, C., N. Ausmees, S. J. Cordwell, L. Shapiro, and M. T. Laub. 2003. Functions of the CckA histidine kinase in Caulobacter cell cycle control. Mol. Microbiol. 47:1279-1290.[CrossRef][Medline]
23
23. Jenal, U., and R. Hengge-Aronis. 2003. Regulation by proteolysis in bacterial cells. Curr. Opin. Microbiol. 6:163-172.[CrossRef][Medline]
24
24. Kahn, K., D. Mazel, J. Houmard, N. Tandeau de Marsac, and M. R. Schaefer. 1997. A role for cpeYZ in cyanobacterial phycoerythrin biosynthesis. J. Bacteriol. 179:998-1006.[Abstract/Free Full Text]
25
25. Kehoe, D. M., and A. R. Grossman. 1997. New classes of mutants in complementary chromatic adaptation provide evidence for a novel four-step phosphorelay system. J. Bacteriol. 179:3914-3921.[Abstract/Free Full Text]
26
26. Kehoe, D. M., and A. R. Grossman. 1996. Similarity of a chromatic adaptation sensor to phytochrome and ethylene receptors. Science 273:1409-1412.[Abstract]
27
27. Kehoe, D. M., and A. Gutu. 2006. Responding to color: the regulation of complementary chromatic adaptation. Annu. Rev. Plant Biol. 57:127-150.[CrossRef][Medline]
28
28. Li, L., R. M. Alvey, R. P. Bezy, and D. M. Kehoe. 2008. Inverse transcriptional activities during complementary chromatic adaptation are controlled by binding of the response regulator RcaC to red and green light responsive promoters. Mol. Microbiol. 68:286-297.[CrossRef][Medline]
29
29. Li, L., and D. M. Kehoe. 2005. In vivo analysis of the roles of conserved aspartate and histidine residues within a complex response regulator. Mol. Microbiol. 55:1538-1552.[CrossRef][Medline]
30
30. Oelmüller, R., P. B. Conley, N. Federspiel, W. R. Briggs, and A. R. Grossman. 1988. Changes in accumulation and synthesis of transcripts encoding phycobilisome components during acclimation of Fremyella diplosiphon to different light qualities. Plant Physiol. 88:1077-1083.[Abstract/Free Full Text]
31
31. Oelmüller, R., A. R. Grossman, and W. R. Briggs. 1988. Photoreversibility of the effect of red and green light-pulses on the accumulation in darkness of mRNAs coding for phycocyanin and phycoerythrin in Fremyella diplosiphon. Plant Physiol. 88:1084-1091.[Abstract/Free Full Text]
32
32. Oelmüller, R., A. R. Grossman, and W. R. Briggs. 1989. Role of protein synthesis in regulation of phycobiliprotein mRNA abundance by light quality in Fremyella diplosiphon. Plant Physiol. 90:1486-1491.[Abstract/Free Full Text]
33
33. Pfaffl, M. W. 2001. A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res. 29:e45.[Abstract/Free Full Text]
34
34. Ryan, K. R., E. M. Judd, and L. Shapiro. 2002. The CtrA response regulator essential for Caulobacter crescentus cell-cycle progression requires a bipartite degradation signal for temporally controlled proteolysis. J. Mol. Biol. 324:443-455.[CrossRef][Medline]
35
35. Sato, N., H. Kawahara, A. Toh-e, and T. Maeda. 2003. Phosphorelay-regulated degradation of the yeast Ssk1p response regulator by the ubiquitin-proteasome system. Mol. Cell. Biol. 23:6662-6671.[Abstract/Free Full Text]
36
36. Seib, L. O., and D. M. Kehoe. 2002. A turquoise mutant genetically separates expression of genes encoding phycoerythrin and its associated linker peptides. J. Bacteriol. 184:962-970.[Abstract/Free Full Text]
37
37. Tandeau de Marsac, N., and J. Houmard. 1993. Adaptation of cyanobacteria to environmental stimuli: new steps towards molecular mechanisms. FEMS Microbiol. Rev. 104:119-189.[CrossRef]
38
38. Tandeau de Marsac, N., D. Mazel, T. Damerval, G. Guglielmi, V. Capuano, and J. Houmard. 1988. Photoregulation of gene expression in the filamentous cyanobacterium Calothrix sp. PCC 7601: light-harvesting complexes and cell differentiation. Photosynth. Res. 18:99-132.[CrossRef]
39
39. Terauchi, K., B. L. Montgomery, A. R. Grossman, J. C. Lagarias, and D. M. Kehoe. 2004. RcaE is a complementary chromatic adaptation photoreceptor required for green and red light responsiveness. Mol. Microbiol. 51:567-577.[CrossRef][Medline]
40
40. To, J. P. C., J. Deruere, B. B. Maxwell, V. F. Morris, C. E. Hutchison, F. J. Ferreira, G. E. Schaller, and J. J. Kieber. 2007. Cytokinin regulates type-A Arabidopsis response regulator activity and protein stability via two-component phosphorelay. Plant Cell 19:3901-3914.[Abstract/Free Full Text]
41
41. Tomomori, C., T. Tanaka, R. Dutta, H. Y. Park, S. K. Saha, Y. Zhu, R. Ishima, D. J. Liu, K. I. Tong, H. Kurokawa, H. Qian, M. Inouye, and M. Ikura. 1999. Solution structure of the homodimeric core domain of Escherichia coli histidine kinase EnvZ. Nat. Struct. Biol. 6:729-734.[CrossRef][Medline]
42
42. Woodcock, D. M., P. J. Crowther, J. Doherty, S. Jefferson, E. Decruz, M. Noyerweidner, S. S. Smith, M. Z. Michael, and M. W. Graham. 1989. Quantitative evaluation of Escherichia coli host strains for tolerance to cytosine methylation in plasmid and phage recombinants. Nucleic Acids Res. 17:3469-3478.[Abstract/Free Full Text]

---

Journal of Bacteriology, November 2008, p. 7241-7250, Vol. 190, No. 21
0021-9193/08/$08.00+0     doi:10.1128/JB.00762-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Li, L. Articles by Kehoe, D. M. Search for Related Content PubMed PubMed Citation Articles by Li, L. Articles by Kehoe, D. M.