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J Mol Biol. Author manuscript; available in PMC Dec 21, 2009.
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PMCID: PMC2796631
NIHMSID: NIHMS161941

Interactions Between the Rhodobacter sphaeroides ECF Sigma Factor, σE, and its Anti-sigma Factor, ChrR

Abstract

Rhodobacter sphaeroides σE is a member of the extra cytoplasmic function sigma factor (ECF) family, whose members have been shown to regulate gene expression in response to a variety of signals. The functions of ECF family members are commonly regulated by a specific, reversible interaction with a cognate anti-sigma factor. In R. sphaeroides, σE activity is inhibited by ChrR, a member of a newly discovered family of zinc containing anti-sigma factors. We used gel filtration chromatography to gain insight into the mechanism by which ChrR inhibits σE activity. We found that formation of the σE:ChrR complex inhibits the ability of σE to form a stable complex with core RNA polymerase. Since the σE:ChrR complex inhibits the ability of the sigma factor to bind RNA polymerase, we sought to identify amino acid substitutions in σE that altered the sensitivity of this sigma factor to inhibition by ChrR. This analysis identified single amino acid changes in conserved region 2.1 of σE that either increased or decreased the sensitivity of σE for inhibition by ChrR. Many of the amino acid residues that alter the sensitivity of σE to ChrR are located within regions known to be important for interacting with core RNA polymerase in other members of the s70 superfamily. Our results suggest a model where solvent-exposed residues with region 2.1 of σE interact with ChrR to sterically occlude this sigma factor from binding core RNA polymerase and to inhibit target gene expression.

Keywords: sigma factor, anti-sigma factor, transcription, regulation, Rhodobacter sphaeroides

Introduction

Transcription is a key step in the regulation of prokaryotic gene expression, and is catalyzed by RNA polymerase. The sigma factor (σ) of this multi-subunit enzyme plays a key regulatory role in gene expression by recognizing specific promoter sequences.1,2 Almost all bacteria currently studied possess multiple sigma factors, including a primary or “housekeeping” sigma (s70-type) and several classes of alternative sigma factors. These alternative σ factors allow cells to regulate transcription of specific genes in response to stress or changing environmental conditions.3-5 Most of these alternative sigma factors are classified as members of the σ70 superfamily due to the conservation of amino acid sequence in regions that either interact with core RNA polymerase, promoter sequences, or facilitate the process of transcription initiation.6-8

Extra-cytoplasmic function (ECF) sigma factors are a group of alternative sigma factors whose target gene products often function outside the cytoplasm (in the membrane, periplasm of Gramnegative bacteria, or beyond).9 ECF sigma factors control cellular responses to diverse environmental demands including: periplasmic stress,10,11 resistance to cobalt and nickel,12,13 high levels of light,14 and oxidative stress.15-17 Analyses of bacterial genome sequences suggest that ECF sigma factors could play a major role in gene regulation, since a large number of the σ70 superfamily currently present in the NCBI database are predicted to be ECF sigma factors.

A common feature of ECF sigma factor family members is the means by which their activity is regulated. The ECF sigma factor is often co-transcribed with a gene coding for a negative regulator, or “anti-sigma factor.”18-20 The regulation of transcription by an anti-sigma factor occurs either by blocking sigma factor binding to core RNA polymerase (SpoIIAB,21 RseA,22 and RsrA23), by facilitating the dissociation of the sigma factor from RNA polymerase holoenzyme (FlgM),24 or by preventing promoter recognition by RNA polymerase holoenzyme (AsiA).25 To date, four ECF sigma factors have been shown to interact directly with their cognate anti-sigma factors: Rhodobacter sphaeroides σE:ChrR,26 Escherichia coli σE:RseA,22,27,28 Pseudomonas aeruginosa AlgU:MucA,29 and S. coelicolor σR:RsrA.16,30 While the molecular interactions between E. coli σE:RseA22 and S. coelicolor σR:RsrA23 have been analyzed, the lack of significant amino acid sequence similarity among anti-sigma factors makes it difficult to predict how each inhibitor will interact with its cognate ECF sigma factor.

We have been analyzing the interactions between the R. sphaeroides ECF sigma factor σE, and its inhibitor, ChrR. ChrR is predicted to be a soluble zinc-dependent anti-sigma factor that lacks significant amino acid sequence similarity to either E. coli RseA or other characterized membrane-bound inhibitors of ECF sigma factors.16,22 Previous work has determined that ChrR forms a heterodimeric complex with R. sphaeroides σE, but the process by which this anti-sigma factor blocks σE function is unknown.26 We show that ChrR can prevent σE from forming a stable complex with core RNAP. In addition, we characterize the effects of amino acid substitutions within region 2.1 of σE which alter the sensitivity of the sigma factor to inhibition by ChrR in vivo and in vitro. We propose that region 2.1 of σE defines a potential site of interaction between σE and ChrR by which the anti-sigma factor could prevent σE activity.

Results

ChrR prevents σE from binding R. sphaeroides core RNA polymerase

Previous work indicated that σE and ChrR form a heterodimeric complex,26 but little was known about the mechanism by which ChrR blocks σE function. To address how ChrR inhibits σE activity, gel filtration chromatography was used to monitor the interactions of σE or the σE:ChrR complex with core RNA polymerase. R. sphaeroides core RNA polymerase and the σE:ChrR complex (predicted molecular mass of 43 kDa) were resolved on a Superdex 200 column (Amersham Pharmacia, Piscataway, NJ), with core RNA polymerase eluting in the void volume and complex eluting with an apparent molecular mass of ~32 kDa (Figure 1a). In addition, when σE (predicted molecular mass of 19.2 kDa) was analyzed on this column, it eluted as a species of an apparent molecular mass of ~17 kDa (Figure 1a). The ability to resolve core RNA polymerase, the σE:ChrR complex, and σE suggested that gel filtration experiments would provide insight into how ChrR blocks σE function.

Figure 1
σE binding to ChrR prevents its ability to interact with RNA polymerase. (a) The elution profile of purified R. sphaeroides core RNA polymerase (0.25 μM), σE (4 μM), and σE:ChrR complex (4 μM) when passed ...

When σE was incubated with core RNA polymerase and passed over this column, a decrease was seen in the amount of UV-absorbing material eluting at ~17 kDa (Figure 1b). SDS-PAGE of TCA precipitated column fractions showed that σE was present in the void volume along with RNA polymerase subunits (Figure 1b). This shift in the σE elution profile indicated that σE was able to bind to RNA polymerase and form a stable complex under these conditions. The presence of σE in the ~17 kDa fraction could be the result of excess σE over core RNA polymerase in the experiment, or some of the sigma factor was inactive due to the purification process and unable to bind core RNA polymerase.

To test if ChrR prevents σE from binding core RNA polymerase, we observed what happened when pure σE:ChrR complex was incubated with core RNA polymerase. When a mixture of core RNA polymerase and the σE:ChrR complex was passed over the Superdex 200 column, there was no detectable change in the area under the σE:ChrR complex peak, nor was there an appearance of a species with an apparent molecular mass predicted for ChrR (~21 kDa, Figure 1c). In addition, SDS-PAGE of TCA precipitated column fractions showed no detectable σE or ChrR in the void volume fractions that contained core RNA polymerase subunits (Figure 1c). This suggests that the σE:ChrR complex does not interact with core RNA polymerase to form a stable complex under conditions where σE can bind to this enzyme. These results also suggest that core RNA polymerase does not remove σE from ChrR under these conditions.

Screen for σE mutants having increased activity in the presence of ChrR

Since formation of a σE:ChrR complex appears to play a critical role in inhibiting σE activity, we sought to identify amino acid residue substitutions in σE that altered the sigma factor’s sensitivity to inhibition by ChrR. To do this, we capitalized on the observation that R. sphaeroides σE and ChrR function in an E. coli tester strain that contains a chromosomal rpoE P1 :: lacZ reporter gene (ΦλJDN1).26 This strain is white on MacConkey’s lactose medium in the absence of R. sphaeroides rpoE, red when it contains rpoE on a plasmid under the control of its own promoter (rpoE P1), and pink when it contains the rpoEchrR operon on the same plasmid (data not shown). Thus, this tester strain provides a screen for rpoE mutations that alter the sensitivity of σE to ChrR.

To look for amino acid substitutions in σE that alter its sensitivity to ChrR, we screened a library of PCR-mutagenized rpoE genes for σE activity in this tester strain. After screening ~5500 colonies from eight independent mutagenesis experiments on MacConkey’s lactose media, ~81% were pink, indicating “wild-type” σE activity, ~16% were white, indicating a decrease in σE activity, and ~3% were red, indicating an increase in σE activity in the presence of ChrR. When the R. sphaeroides rpoE gene from 65 of the 151 red colonies was sequenced, four different single amino acid substitutions in σE were identified. Three of the amino acid substitutions (K38E, K38R, and F40S) were located in σE region 2.1, and one (F81I) was located in σE region 2.3 (Figure 2). The single amino acid substitutions in σE, K38E and K38R, were identified twice from independent mutagenesis screens. The rpoE genes in the remaining red colonies that were sequenced contained multiple mutations, including ~30 isolates which had K38E, K38R, F40S, or F81I as one of the multiple amino acid substitutions. These results suggest that amino acid residues in regions 2.1–2.3 of σE may be important for its interaction with ChrR.

Figure 2
Alignment of E. coli and R. sphaeroides σE proteins. The amino acid sequence alignment was generated using ClustalW and the indicated sequences: E. coli σE E. coli (GenBank accession no. P334086) and R. sph, σE R. sphaeroides (GenBank ...

The effects of rpoE mutations in vivo

To assess the effects of each amino acid substitution in σE on its function, we measured levels of β-galactosidase produced from our tester strain, E. coli VH1000 (ΦλJDN1), which contained either wild-type, K38E, K38R, F40S, or F81I σE proteins. Cells containing either the K38E, K38R, or F81I mutant σE proteins had between ~three and fivefold more β-galactosidase activity than wild-type σE in the presence of ChrR (Figure 3a). Cells containing the F40S σE protein showed a slight, but reproducible, increase in β-galactosidase activity in the presence of ChrR as compared to those cells containing wild-type σE (Figure 3a).

Figure 3
R. sphaeroides σE mutants have altered activity in vivo in the presence of the anti-sigma factor, ChrR. (a) β-galactosidase activity from an E. coli tester strain containing a chromosomal rpoE P1 :: lacZ transcriptional fusion and ...

To test if the amino acid substitutions in σE affected sigma factor function, we monitored β-galactosidase activity from strains that lacked ChrR but contained either wild-type, K38E, K38R, F40S, or F81I σE. If the amino acid substitutions in σE did not affect sigma factor function, we expected to find rpoE P1 :: lacZ reporter activity comparable to that seen in cells containing only wild-type σE. Cells containing the K38E, K38R, or F81I mutant σE proteins in the absence of ChrR had levels of β-galactosidase activity comparable to a strain containing wild-type σE in the absence of ChrR (Figure 3a), suggesting that these amino acid substitutions did not negatively affect sigma factor function. However, the F40S substitution in σE caused a slight (~33%) decrease in β-galactosidase activity in the absence of ChrR, suggesting that this amino acid substitution affects σE function. In addition, this analysis showed that the K38E, K38R, and F81I mutant σE proteins had a decrease in activity in the absence of ChrR when compared to activity in the presence of the anti-sigma factor. Possible explanations for the decreased σE activity in cells lacking the inhibitor will be presented in the Discussion.

To test the behavior of these mutant σE proteins in its native host, we expressed ChrR and either wild-type, K38E, K38R, F40S, or F81I σE (under the control of its own promoter, rpoE P1) from a stable low copy plasmid in a R. sphaeroides strain that contains a chromosomal deletion of rpoEchrR (TF18). To determine σE function, we placed a rpoE P1:lacZ reporter fusion on a compatible low copy plasmid (pJDN30) in this strain. β-galactosidase activity from this reporter fusion in TF18 cells containing a plasmid with the intact rpoEchrR operon is low (Figure 3b), because we know ChrR inhibits σE under these growth conditions.26 Cells containing K38E σE had an ~100-fold increase in β-galactosidase activity, cells containing F81I σE exhibited an ~12-fold increase in β-galactosidase activity, and cells containing K38R σE had ~eightfold more β-galactosidase activity than the control strain containing wild-type σE (Figure 3b). In contrast, cells containing F40S σE only had ~1.4-fold more β-galactosidase activity, which is similar to the behavior of this mutant sigma factor in E. coli. When taken together, the properties of these mutant σE proteins in E. coli and R. sphaeroides suggest that the K38E, K38R, and F81I substitutions alter the sensitivity of σE to inhibition by ChrR without reducing their ability to function as sigma factors.

Region 2.1 of R. sphaeroides σE contains additional amino acid residues that are important for inhibition by ChrR

We sought to determine if the amino acid residues identified in our screen mapped to a specific region of sigma factors. Aligning the amino acid sequences of R. sphaeroides σE and E. coli σE suggests that residues K38, F40, and F81 of R. sphaeroides σE correspond to amino acid residues A34, L36, and Y75 of E. coli σE (Figure 2). Mapping these residues of R. sphaeroides σE on the structure of E. coli σE,22 predicts that residue K38 of this protein is surface exposed within the α-helical domain of region 2.1; residues F40 and F81 of R. sphaeroides σE appear to be involved in stabilizing interactions between the α-helices of regions 2.1 and 2.3 (Figure 4). In addition, this model predicts that residues K38 and F40 are within a part of σE region 2.1 that makes protein–protein contacts with other parts of the sigma factor,31 core RNA polymerase,32,33 or anti-sigma factors.22,23 To determine if other amino acid residues within region 2.1 of R. sphaeroides σE are important for inhibition by ChrR, we individually substituted 21 amino acid residues within a 25 residue region (22DEAAFAELFQHFAPKVKGFLMKSGS46) with alanine (residues A24, A25, A27, and A34 are alanine in wild-type protein). These mutant σE proteins were expressed in the E. coli tester strain to determine if any of these alanine substitutions altered the sensitivity of σE to inhibition by ChrR or disrupted their function as sigma factors. β-galactosidase levels from the strains tested suggest that the single amino acid substitutions in region 2.1 of σE can be classified into several categories.

Figure 4
Amino acid substitutions in R. sphaeroides σE which alter activity in the presence of ChrR. Crystal structure of E. coli σE region 2,22 highlighting the side-chains of residues in R. sphaeroides σE that alter sensitivity to ChrR. ...

Cells containing the D22A, E23A, H32A, K36A, K38A, G39A, F40A, L41A, K43A, S44A, G45A, or S46A mutant σE proteins had β-galactosidase activity similar to that found in cells containing either wild-type σE or those containing both wild-type σE and ChrR (Figure 5a). This suggests that alanine substitutions at these positions of R. sphaeroides σE do not have measurable effects on either sigma factor function (in cells lacking ChrR), or on their sensitivity to inhibition by ChrR. The K38A σE protein falls into this category, which suggests that removing the lysine side-chain has no affect on σE activity, but replacing the lysine with an arginine or a glutamate (K38R or K38E) reduces the sensitivity of this mutant sigma factor to inhibition by ChrR (Figure 3a).

Figure 5
Effects of alanine substitutions on σE activity in vivo. β-galactosidase activity from E. coli tester strains containing either rpoE (□), or rpoE and chrR (■). All assays were performed in triplicate, with vertical bars ...

Cells containing M42A σE had a small, but reproducible (1.2-fold) increase in σE activity in the presence of ChrR, and an ~fourfold decrease in activity when compared to wild-type σE in the absence of ChrR (Figure 5b). Amino acid sequence alignments indicate that residue M42 of R. sphaeroides σE corresponds to residue S38 of E. coli σE (Figure 2). By mapping R. sphaeroides σE residue M42 on the E. coli σE structure,22 the side-chain of this amino acid appears to lie on the same face as that of residue K38. From the mutational studies of R. sphaeroides σE residues K38 and M42, it appears that amino acid changes in this region can reduce the sensitivity of mutant sigma factors for inhibition by ChrR (Figure 4).

Cells containing alanine substitutions at position E28, Q31, and P35 of σE had between ~three and sevenfold less activity in the presence of ChrR than cells containing wild-type σE (Figure 5b). In the absence of ChrR, the E28A, Q31A, and P35A mutant σE proteins have activity similar to or slightly lower than cells containing wild-type σE (Figure 5b). These results suggest that alanine substitutions at positions E28, Q31, and P35 increase σE sensitivity to inhibition by ChrR without affecting function. When the R. sphaeroides σE E28, Q31, and P35 side-chains were mapped onto the E. coli σE structure,22 these residues are predicted to lie on the same surface exposed face of region 2.1 as residues K38 and M42 (Figure 4). Therefore, it appears that single amino acid substitutions in a potential surface exposed face of R. sphaeroides σE region 2.1 can either decrease or increase the sensitivity of this sigma factor to inhibition by ChrR.

Finally, cells containing mutant σE proteins with amino acid substitutions at residues F26, L29, F30, F33, and V37 show a ~two to fourfold decrease in σE activity in the presence of ChrR, and a ~three to ninefold decrease in σE activity in the absence of ChrR when compared to cells containing wild-type σE (Figure 5c). It has been impossible to determine if the abundance of any mutant σE proteins in E. coli is significantly different from their wild-type counterparts (data not shown). Thus the behavior of this class of mutant proteins suggests that alanine substitutions at these positions affect the activity or stability of σE. If the R. sphaeroides σE F26, L29, F30, F33, and V37 side-chains are mapped on the structure of E. coli σE,22 they are predicted to stabilize helix–helix interactions between region 2.1 and the other parts of region 2 (data not shown). The negative effects of alanine substitutions in residues F26, L29, F30, F33, and V37 on R. sphaeroides σE activity in vivo also suggests that these residues are involved in stabilizing potential helix–helix interactions between region 2.1 and other regions of σE.

Sensitivity of mutant σE proteins to inhibition by ChrR in vitro

The behavior of these mutant σE proteins in vivo predicts that these amino acid substitutions should alter the sensitivity of the sigma factor to inhibition by ChrR in vitro. To test the sensitivity of mutant σE proteins to inhibition by ChrR in vitro, a His6-tagged version of each protein was purified and used for in vitro transcription reactions with the rpoE P1 template. We focused on several mutant σE proteins because they showed either altered sensitivity for ChrR (E28A, Q31A, P35A, K38E, K38R, and M42A), or because they provided controls which had essentially wild-type σE activity in vivo (G39A).

Before testing the sensitivity of each mutant σE protein to inhibition by ChrR, the amount of sigma factor required to produce maximal transcription from the rpoE P1 reporter was determined. By measuring the amount of rpoE P1 transcript produced as the concentration of wild-type or mutant σE was increased (0–100 nM), the relative activity of each mutant sigma factor was measured. Maximal transcript levels were seen with 50–100 nM concentrations of each protein (Figure 6), suggesting that the activity of each purified mutant σE protein was within twofold of wild-type σE. From this we conclude that none of the single amino acid substitutions dramatically reduce the ability of these mutant σE proteins to function in transcription.

Figure 6
Activity of wild-type and mutant σE proteins in vitro. Multiple round in vitro transcription assays performed with increasing amounts of σE proteins. Shown is the amount of rpoE P1 transcript produced with a constant amount of R. sphaeroides ...

To test the ability of ChrR to inhibit wild-type and mutant σE proteins in vitro, we measured the amount of rpoE P1 transcript remaining after the addition of increasing amounts of ChrR fused to maltose-binding protein (ranging from one to 20-fold molar excess compared to σE).26 For these assays, a concentration of σE protein was used that produced 50% of the maximal rpoE P1 transcript. This concentration increased our ability to monitor any alterations in the sensitivity of individual sigma factors to inhibition by ChrR. In assays containing wild-type σE, we saw a concentration-dependent decrease in the rpoE P1 transcript when ChrR was added. When ChrR concentrations were at fivefold excess over σE, an ~80% decrease in σE-dependent transcription was seen (Figure 7a and inset). As a control, a mutant σE protein which appeared to have wild-type sensitivity to ChrR in vivo (G39A) was also inhibited in a concentration-dependent manner by ChrR, with a fivefold excess of ChrR causing ~70% inhibition of sigma factor activity (Figure 7a and inset).

Figure 7
Amino acid substitutions in R. sphaeroides σE that affect the ability of ChrR to inhibit σE-dependent activity in vitro. The percent of rpoE P1 transcript produced in multiple round transcription assays with increasing amounts of ChrR. ...

When this assay was used to analyze the effects of ChrR on the other mutant σE proteins, transcription using the K38E, K38R, or M42A σE proteins was less sensitive to inhibition by ChrR. Only a 5–25% reduction in rpoE P1 transcript levels was seen when ChrR was present at fivefold excess over K38E, K38R, or M42A mutant σE proteins (Figure 7a). Therefore, the K38E, K38R, and M42A substitutions in σE result in mutant sigma factors which are less sensitive to ChrR in vivo and in vitro, reinforcing the conclusion that residues K38 and M42 could define a site of interaction with the anti-sigma factor.

In contrast, σE-dependent transcription from the E28A and Q31A mutant σE proteins reproducibly required only twofold excess ChrR to obtain maximal inhibition (80%), instead of the fivefold excess required for maximal inhibition of either wild-type or G39A σE proteins (Figure 7b). These results are consistent with the in vivo behavior of the E28A and Q31A mutant σE proteins, and they suggest that alanine substitutions at these positions increase the sensitivity of this sigma factor to ChrR.

The last mutant σE protein tested, P35A, also appeared to be more sensitive to inhibition by ChrR in vivo (Figure 5b). However, when P35A σE was tested in vitro for sensitivity to ChrR, its activity was only reduced ~25% in the presence of fivefold excess ChrR (Figure 7a). This level of inhibition is similar to that observed with other mutant σE proteins that appear less sensitive to ChrR, suggesting that the P35A substitution in σE reduces the sensitivity of this sigma factor to its inhibitor. Possible explanations for the different behaviors of the P35A σE mutant protein in the presence of ChrR in vivo and in vitro will be presented in the Discussion.

Discussion

In previous studies, R. sphaeroides σE was shown to form a 1 : 1 complex with ChrR, and it was suggested that binding of ChrR to σE was sufficient to inhibit σE-dependent transcription.26 However, it was not known how ChrR binding inhibited σE activity and what regions of the σ factor were important for inhibition by ChrR. In this work we determined a possible mechanism of σE inhibition by ChrR and identified amino acid substitutions in σE which alter sensitivity of this sigma factor to inhibition by its anti-sigma factor, ChrR.

ChrR prevents σE from interacting with core RNA polymerase

Previous work has shown that the σE:ChrR complex is unable to transcribe σE target genes when added to core RNA polymerase.26 One possible explanation for this observation is that ChrR binding to σE prevents this sigma factor from binding to core RNA polymerase. A similar situation occurs between E. coli σE and its anti-sigma factor RseA, as well as between S. coelicolor σR and RsrA. For E. coli RseA, it is proposed that interactions with σE regions 2 and 4 sterically occlude binding sites for the RNA polymerase β’ and β subunits.22 In the case of S. coelicolor RsrA, an ~10 kDa fragment of the sigma factor that contains region 2 has been shown to interact with RNA polymerase.23 In addition, the interaction between this fragment of σR and core RNA polymerase can be prevented by the presence of RsrA, suggesting that region 2 of this sigma factor is involved in forming a complex with RNA polymerase and with RsrA.23 However, there are other examples of anti-sigma factors that allow the σ factor to interact with RNA polymerase but prevent the resulting holoenzyme from initiating transcription.34-37 Given the lack of significant amino acid similarity between ChrR and other characterized anti-sigma factors, it was important to understand how this protein prevents σE activity.

We found that the σE:ChrR complex was unable to form a stable complex with core RNA polymerase under conditions where σE was able to bind to this enzyme to form EσE. In addition, the σE:ChrR complex did not appear to dissociate when incubated with core RNA polymerase, suggesting that ChrR binding to σE is mutually exclusive with core RNA polymerase binding to σE. These observations, when considered with previous work, provide strong evidence that formation of the σE:ChrR complex is key to inhibiting R. sphaeroides σE-dependent transcription.

Region 2.1 of R. sphaeroides σE is important for sensitivity to ChrR

One possible explanation for the inability of σE to bind to core RNA polymerase when this sigma factor interacts with ChrR is that the anti-sigma factor masks RNA polymerase-binding determinants on σE. Analysis of members of the σ70 superfamily of sigma factors suggests that regions 2.1, 2.2 and 4.1 are principal sites for RNA polymerase binding.33,38-40 If ChrR were to inhibit σE holoenzyme formation by blocking RNA polymerase-binding determinants, one might expect that residues within one or more of these regions of σE are also important for sensitivity to this anti-sigma factor.

Our results implicate region 2.1 of σE as being important for inhibition by the anti-sigma factor, ChrR. Specifically, we identified amino acid changes in region 2.1 of R. sphaeroides σE that either decrease (K38E, K38R, and M42A) or increase (E28A and Q31A) the sensitivity of σE to inhibition by ChrR. When these residues are modeled on the E. coli σE structure,22 they appear to be located on a solvent-exposed face of region 2.1. Thus it is possible that one or more of these amino acid side-chains constitute a site on σE that is involved in making direct interactions with ChrR. If this hypothesis is correct, ChrR binding to σE could sterically occlude a major RNA polymerase-binding determinant on σE and thereby prevent formation of σE holoenzyme. Each of the mutant σE proteins studied (E28A, Q31A, P35A, K38E, K38R, and M42A) were able to form transcriptionally competent complexes with core RNA polymerase, suggesting that these amino acid residues are not essential for RNA polymerase holoenzyme formation. We propose that these determinants in region 2.1 of σE are part of a larger domain used by σ70 family members to interact with core RNA polymerase, and that ChrR interactions within this domain interfere sterically with RNA polymerase holoenzyme formation.22,23

The crystal structure of the E. coli σE:RseA complex shows that the N terminus of RseA makes both van der Waals and hydrogen bond contacts with residues of E. coli σE region 2.1, including Leu24 and Val27.22 The corresponding R. sphaeroides σE region 2.1 residues, Glu28 and Gln31, were identified as important for sensitivity to ChrR since alanine substitutions at these two positions increased inhibition of σE by this anti-sigma factor in vivo and in vitro. It is interesting to note that the loss of a large charged side-chain (Glu and Gln) at each position in R. sphaeroides σE results in increased sensitivity to ChrR, perhaps by allowing additional main-chain interactions between the sigma factor and its anti-sigma factor. Experiments are in progress to probe the nature of the interactions between σE and ChrR, and to test if these and other changes in region 2.1 directly alter the interactions of these two proteins.

The organization of region 2.1 of R. sphaeroides σE

Our data suggest that the overall structure of region 2.1 in R. sphaeroides σE is α-helical as is the case of other members of the σ70 superfamily.22,23,31,41 If we map the amino acid side-chains in region 2.1 of R. sphaeroides σE onto a helical wheel model, all of the alanine substitutions that reduce sigma factor activity map to one face of the predicted helix (Figure 8). This face is predicted to make helix–helix interactions with regions 2.2 and 2.3 based on the analysis of other σ70 family members (see below).22,23,31,41 In addition, residues identified by our studies which alter the sensitivity of σE to inhibition by ChrR map to the opposite face of these helical wheel projections, and could be surface exposed (Figure 8, see below).

Figure 8
Helical wheel model of region 2.1 of R. sphaeroides σE. Due to the presence of a proline at position 35 in σE we modeled region 2.1 of R. sphaeroides σE as two helical domains. Residues in σE are highlighted as follows: ...

The position of region 2.1 in E. coli σ70 and σE appears to be stabilized by hydrophobic side-chain interactions with residues in regions 2.3 and 2.4.22,31,41 The properties of R. sphaeroides σE mutant proteins, F40S and F81I, could be explained if these side-chains made Van der Waals interactions with amino acid side-chains in regions 2.3 and 2.1, respectively. If this is true, interrupting these interactions could both alter the sensitivity of σE proteins for ChrR, and reduce their ability to function as sigma factors.

In the case of E. coli σ70, E. coli σE and S. coelicolor σR, the α-helical nature of region 2.1 is disrupted at the position equivalent to the proline at residue 35 in R. sphaeroides σE.22,23 For this reason, we have chosen to model region 2.1 of R. sphaeroides σE as two helices extending from residue D22 to P35 and K36 to S44 (Figure 8). A search of the protein database indicates that a proline at this position is present in several ECF sigma factors with a high degree of amino acid sequence identity to R. sphaeroides σE (unpublished data). In addition, the genes encoding for each of these σE homologues are linked to genes predicted to encode proteins related to ChrR. Therefore, it is possible that a structural feature created by a proline at this position in σE may be important for recognition of this particular group of ECF sigma factors by their cognate anti-sigma factors.

Properties of mutant σE proteins in vivo and in vitro

Somewhat surprisingly, the in vivo analysis of the K38E, K38R, and F81I mutant σE proteins revealed that the absence of ChrR led to a decrease in σE activity when compared to that found in the presence of the anti-sigma factor. Western blot analysis with antiserum to σE showed a significant decrease in the amount of each of these three mutant σE proteins in cells lacking ChrR (unpublished data), suggesting that σE turnover could influence the amount of target promoter activity measured in vivo. Protein turnover is a critical part of the regulatory circuit which controls σE activity in E. coli.42,43 By analogy, proteolysis of free R. sphaeroides σE may influence the activity of wild-type and mutant sigma factors in vivo. If this were true, turnover of K38E, K38R, and F81I mutant σE proteins might account for the decreased target promoter activity observed in the absence of ChrR.

Proteolysis may also explain why the P35A mutant σE protein appeared to have increased sensitivity to ChrR in vivo, while in vitro studies showed this protein was less sensitive to inhibition by this anti-sigma factor (compare Figures Figures5b5b and and7a).7a). We propose that P35 is important for the structure of R. sphaeroides σE region 2.1 and the ability of the ECF sigma factor to interact with its cognate anti-sigma factor. Experiments are underway to determine whether residue P35 of σE makes direct interactions with ChrR, or if the proline at this position is important for maintaining a structure that is necessary for recognition of σE by ChrR.

Information available from the analysis of several anti-sigma factors suggests that these proteins share little sequence similarity, and have different structures when bound to their cognate σ factors.21,22,30 ChrR also shares little sequence similarity with any of the anti-sigma factors that have been structurally characterized to date, so we believe that its structure when bound to σE will be novel. Thus, it will be interesting to determine if R. sphaeroides σE makes additional contacts with ChrR that were not identified in our analysis. Indeed, ChrR belongs to a recently discovered class of zinc-binding anti-sigma factors common among various α and γ proteobacteria,15,17,26,44,45 so understanding more about its structure, mechanism of inhibition, and the factors that control its regulation will shed light on a process that is likely to be conserved in the microbial world.

In summary, we have identified one mechanism by which ChrR inhibits R. sphaeroides σE activity, and have identified region 2.1 as a likely site of interaction on this ECF sigma factor for its cognate anti-sigma. Studies are under way to identify the region(s) of the anti-sigma factor that are required for inhibition of σE, to determine the structure of this complex, and to identify the signal which controls the interaction between ChrR and σE.

Materials and Methods

Bacterial strains, plasmids, and growth conditions

E. coli strains (Table 1) were grown at 37 °C in Luria–Bertani medium.46 R. sphaeroides strains (Table 1) were grown at 30 °C in Sistrom’s succinate-based minimal medium A.47 When necessary, media was supplemented with 100 μg/ml ampicillin, 25 μg/ml kanamycin, 25 μg/ml spectinomycin, or 1–10 μg/ml tetracycline to maintain plasmids. Sequences of primers used in this study are available upon request.

Table 1
Bacterial strains and plasmids

Purification of R. sphaeroides core RNA polymerase, σE:ChrR complex, MBP-ChrR, and σE

R. sphaeroides core RNA polymerase was purified via a combination of affinity chromatography using the polyol-responsive 4RA2 monoclonal antibody (Dr Richard Burgess, Madison, WI) and anion exchange chromatography.48 A σE:ChrR complex, containing either wild-type or mutant σE proteins, was purified by Ni2+ affinity chromatography from E. coli cells containing an intact rpoEchrR operon behind an inducible promoter.48 MBP-ChrR was purified by affinity chromatography as described.26

To purify σE proteins, we constructed expression plasmids by digesting pBS16 derivatives containing either wild-type σE or the various σE mutants with BsmFI and BamHI. The resulting 430 bp fragment containing the appropriate rpoE was cloned into pJDN14, and introduced into M15pREP4 (QIAGEN, Valencia, CA). Wild-type and mutant σE proteins were overexpressed and purified as described.48

Gel filtration chromatography

σE (4 μM), R. sphaeroides core RNA polymerase (0.25 μM), σE:ChrR complex (4 μM), σE incubated with core RNA polymerase, or σE:ChrR complex incubated with core RNA polymerase was mixed in HPLC buffer (25 mM Tris–HCl, pH 7.9, 150 mM NaCl) at room temperature for 30 minutes. Samples were loaded onto a Superdex 200 column (Amersham Pharmacia, Piscataway, NJ) that was equilibrated with HPLC buffer at a flow rate of 0.5 ml/minute for 50 minutes using a System Gold 125NM solvent module connected to a model 168 diode array detector (Beckman Coulter, Fullerton, CA). Apparent molecular masses were estimated by comparison to elution of low and high molecular mass standards (Amersham Pharmacia, Piscataway, NJ). Column eluates were TCA precipitated, separated on a 4–12% Bis-Tris polyacrylamide gel (Invitrogen, Carlsbad, CA), and visualized using the Gelcode Blue system (Pierce, Rockford, IL).

Mutagenesis of rpoE

Error-prone PCR was carried out in EasyStart PCR tubes (Molecular BioProducts, San Diego, CA) with 1% (w/v) Triton X-100, using 10 pmol of primers 1212 and 1233 (New England BioLabs, Beverly, MA), 20 ng of pUC19rpoE plasmid (Table 1), 0.5 mM of each dNTP, 1 × Taq DNA polymerase buffer, and 2.5 units Taq DNA polymerase (Promega Corp., Madison, WI). Reactions were allowed to proceed for 35 cycles of 95 °C for 30 seconds, 60 °C for 30 seconds, and 72 °C for two minutes. PCR products were cloned into pBS16 using restriction sites NdeI and StuI, which were within rpoE.

Alanine substitutions within σE region 2.1 (22DEAAFAELFQHFAPKVKGFLMKSGS46) were generated via sitedirected mutagenesis using the QuickChange Mutagenesis kit (Stratagene, La Jolla, CA) with pBS16 or pJDN48 templates. Genes encoding mutant σE proteins were placed in plasmids lacking ChrR by digesting pJDN48 derivatives with EcoRI. All plasmids encoding mutant rpoE were confirmed by DNA sequencing, and transformed into VH1000 (ΦλJDN1).

Screening of σE proteins for activity in the presence of ChrR

pBS16 plasmids containing wild-type or mutant rpoE genes were transformed into the VH1000 tester strain that contains an rpoE P1 :: lacZ fusion as a lambda lysogen (ΦλJDN1).26 Strains were plated on MacConkey’s media supplemented with 0.5% (w/v) lactose. Colonies that became red after overnight incubation at 37 °C, indicating lactose utilization (and increased σE-dependent transcriptional activity) had the rpoE gene from their plasmid sequenced using specific primers to determine the nature of the mutation(s).

Mutant rpoE genes encoding σE proteins with single amino acid changes were cloned into pJDN48 using BsmFI and BamHI restriction digests. This pBS16-derived plasmid increases the abundance of ChrR by placing a strong E. coli ribosome-binding site in front of chrR.26 Additional plasmids lacking chrR were created by EcoRI digestion of the appropriate pJDN48 derivatives.

Placing σE mutant proteins in R. sphaeroides

Wild-type or mutant rpoEchrR operons were excised from pBS16 via an EcoRI and HindIII restriction digest. The resulting 1.8 kb fragments were cloned into pJDN18, a pRK415 derivative that is stable in R. sphaeroides. Plasmid constructs were confirmed by DNA sequencing, transformed into E. coli S17-1, and placed in R. sphaeroides TF18 (pJDN30) by conjugation.49 R. sphaeroides TF18 has a deletion in the rpoEchrR operon that renders σE and ChrR inactive,50 so σE activity is dependent on the rpoE gene within pJDN18. Plasmid pJDN30 contains a rpoE P1:lacZ transcriptional fusion on a compatible, low copy plasmid used to quantify σE activity in R. sphaeroides.51

β-galactosidase assays

β-galactosidase activity assays were performed in triplicate as described.52 Results from β-galactosidase assays are presented in Miller units.53

Modeling of R. sphaeroides σE region 2.1

The structure of E. coli σE (PDB accession no. 1OR7) was used as a scaffold to model the position of residues of R. sphaeroides σE analyzed in this study.22 The program Deep View—Swiss PDB Viewer54 was used to replace the appropriate E. coli residues with the corresponding R. sphaeroides residues. For reference, R. sphaeroides σE Glu28, Gln31, Pro35, Lys38, Phe40, Met42, and Phe81 correspond to E. coli σE Leu24, Val27, His31, Ala34, Leu36, Ser38 and Tyr75.

Figure 4 was generated using DS ViewerPro 5.0 from Accelrys. Figure 8 was constructed using helical wheel software§.

In vitro transcription assays

To determine the relative activity of wild-type and mutant σE proteins, in vitro transcription assays were performed. Increasing concentrations (0–100 nM) of wild-type and mutant His6E proteins were added to 50 nM R. sphaeroides core RNA polymerase in transcription buffer (40 mM Tris–HCl, pH 7.9, 200 mM KCl, 10 mM Mg acetate, 1 mM DTT, 62.5 μg/ml acetylated BSA). Plasmid pJDN34, containing rpoE P1 cloned upstream of a known transcription terminator,39 was added to a final concentration of 20 nM and the reactions were incubated at 30 °C for 30 minutes. Transcription was initiated with the addition of ribonucleotides at final concentrations of 250 μM GTP, CTP, ATP; 25 μM UTP; and 1 μCi [α-32P]UTP (3000 Ci/mmol). Reactions were incubated at 30 °C for 30 minutes, and terminated with the addition of 95% (w/v) formamide loading buffer.55 RNA products were analyzed for the amount of σE-dependent transcript using 6% (w/v) denaturing PAGE and the Molecular Dynamics phosphorimaging system (Sunnydale, CA).

To determine the effects of MBP-ChrR on the activity of wild-type and mutant σE proteins, in vitro transcription assays were performed. Increasing amounts of MBP-ChrR (0–500 nM) were added to an assay containing an amount of His6E (~25–100 nM) that produced 50% of the maximal activity (see above) in transcription buffer. This mixture was allowed to incubate for 30 minutes at 30 °C. Template (pJDN34, 20 nM) and R. sphaeroides core RNA polymerase (50 nM) were added and the mixture was incubated at 30 °C for an additional 30 minutes. Transcription was initiated with the addition of ribonucleotides (see above) and allowed to proceed for 30 minutes at 30 °C. RNA products were analyzed as described above. The amount of σE-dependent transcript for each reaction was analyzed using ImageQuant software (Molecular Dynamics, Sunnydale, CA), and data were plotted using Origin 7.0 (OriginLab Corp., Northampton, MA).

Acknowledgements

This work was supported by NIH grant GM37509 to T.J.D. During a portion of this work, J.R.A. was supported by the Jerome J. Sterfaniak PreDoctoral Fellowship from UW-Madison Department of Bacteriology. We thank Larry Anthony, Liz Campbell, Heather Green, Tricia Kiley, and Ruth Saecker for their assistance during this project.

Abbreviations used

ECF
extra-cytoplasmic function.

References

1. Burgess RR, Travers AA, Dunn JJ, Bautz EK. Factor stimulating transcription by RNA polymerase. Nature. 1969;221:43–46. [PubMed]
2. Burgess RR, Anthony LC. How sigma docks to RNA polymerase and what sigma does. Curr. Opin. Microbiol. 2001;4:126–131. [PubMed]
3. Helmann JD, Marquez LM, Chamberlin MJ. Cloning, sequencing, and disruption of the Bacillus subtilis σ28 gene. J. Bacteriol. 1988;170:1568–1574. [PMC free article] [PubMed]
4. Gross CA, Chan CL, Lonetto MA. A structure/function analysis of Escherichia coli RNA polymerase. Phil. Trans. Roy. Soc. Lond. B Biol. Sci. 1996;351:475–482. [PubMed]
5. Wosten MM. Eubacterial sigma-factors. FEMS Microbiol. Rev. 1998;22:127–150. [PubMed]
6. Gribskov M, Burgess RR. Sigma factors from E. coli, B. subtilis, phage SP01, and phage T4 are homologous proteins. Nucl. Acids Res. 1986;14:6745–6763. [PMC free article] [PubMed]
7. Stragier P, Parsot C, Bouvier J. Two functional domains conserved in major and alternate bacterial sigma factors. FEBS Letters. 1985;187:11–15. [PubMed]
8. Lonetto M, Gribskov M, Gross CA. The σ70 family: sequence conservation and evolutionary relationships. J. Bacteriol. 1992;174:3843–3849. [PMC free article] [PubMed]
9. Lonetto MA, Brown KL, Rudd KE, Buttner MJ. Analysis of the Streptomyces coelicolor σE gene reveals the existence of a subfamily of eubacterial RNA polymerase sigma factors involved in the regulation of extracytoplasmic functions. Proc. Natl Acad. Sci. USA. 1994;91:7573–7577. [PMC free article] [PubMed]
10. Rouviere PE, De Las Penas A, Mecsas J, Lu CZ, Rudd KE, Gross CA. rpoE, the gene encoding the second heat-shock sigma factor, σE,in Escherichia coli. EMBO J. 1995;14:1032–1042. [PMC free article] [PubMed]
11. Raina S, Missiakas D, Georgopoulos C. The rpoE gene encoding the σE24) heat shock sigma factor of Escherichia coli. EMBO J. 1995;14:1043–1055. [PMC free article] [PubMed]
12. Tibazarwa C, Wuertz S, Mergeay M, Wyns L, van Der Lelie D. Regulation of the cnr cobalt and nickel resistance determinant of Ralstonia eutropha (Alcaligenes eutrophus) CH34. J. Bacteriol. 2000;182:1399–1409. [PMC free article] [PubMed]
13. Grass G, Grosse C, Nies DH. Regulation of the cnr cobalt and nickel resistance determinant from Ralstonia sp. strain CH34. J. Bacteriol. 2000;182:1390–1398. [PMC free article] [PubMed]
14. Gorham HC, McGowan SJ, Robson PR, Hodgson DA. Light-induced carotenogenesis in Myxococcus xanthus: light-dependent membrane sequestration of ECF sigma factor CarQ by anti-sigma factor CarR. Mol. Microbiol. 1996;19:171–186. [PubMed]
15. Paget MS, Kang JG, Roe JH, Buttner MJ. σR, an RNA polymerase sigma factor that modulates expression of the thioredoxin system in response to oxidative stress in Streptomyces coelicolor A3(2) EMBO J. 1998;17:5776–5782. [PMC free article] [PubMed]
16. Kang JG, Paget MS, Seok YJ, Hahn MY, Bae JB, Hahn JS, et al. RsrA, an anti-sigma factor regulated by redox change. EMBO J. 1999;18:4292–4298. [PMC free article] [PubMed]
17. Bae JB, Park JH, Hahn MY, Kim MS, Roe JH. Redox-dependent changes in RsrA, an anti-sigma factor in Streptomyces coelicolor: zinc release and disulfide bond formation. J. Mol. Biol. 2004;335:425–435. [PubMed]
18. Helmann JD. Anti-sigma factors. Curr. Opin. Microbiol. 1999;2:135–141. [PubMed]
19. Hughes KT, Mathee K. The anti-sigma factors. Annu. Rev. Microbiol. 1998;52:231–286. [PubMed]
20. Brown KL, Hughes KT. The role of anti-sigma factors in gene regulation. Mol. Microbiol. 1995;16:397–404. [PubMed]
21. Campbell EA, Masuda S, Sun JL, Muzzin O, Olson CA, Wang S, Darst SA. Crystal structure of the Bacillus stearothermophilus anti-sigma factor SpoIIAB with the sporulation sigma factor σF. Cell. 2002;108:795–807. [PubMed]
22. Campbell EA, Tupy JL, Gruber TM, Wang S, Sharp MM, Gross CA, Darst SA. Crystal structure of Escherichia coli σE with the cytoplasmic domain of its anti-sigma RseA. Mol. Cell. 2003;11:1067–1078. [PubMed]
23. Li W, Stevenson C, Burton N, Jakimowicz P, Paget M, Buttner M, Lawson D, Kleanthous C. Identification and structure of the anti-sigma factor-binding domain of the disulphide-stress regulated sigma factor σR from Streptomyces coelicolor. J. Mol. Biol. 2002;323:225. [PubMed]
24. Chadsey MS, Karlinsey JE, Hughes KT. The flagellar anti-sigma factor FlgM actively dissociates Salmonella typhimurium σ28 RNA polymerase holoenzyme. Genes Dev. 1998;12:3123–3136. [PMC free article] [PubMed]
25. Adelman K, Orsini G, Kolb A, Graziani L, Brody EN. The interaction between the AsiA protein of bacteriophage T4 and the σ70 subunit of Escherichia coli RNA polymerase. J. Biol. Chem. 1997;272:27435–27443. [PubMed]
26. Newman JD, Anthony JR, Donohue TJ. The importance of zinc-binding to the function of Rhodobacter sphaeroides ChrR as an anti-sigma factor. J. Mol. Biol. 2001;313:485–499. [PubMed]
27. De Las Penas A, Connolly L, Gross CA. The σE-mediated response to extracytoplasmic stress in Escherichia coli is transduced by RseA and RseB, two negative regulators of σE. Mol. Microbiol. 1997;24:373–385. [PubMed]
28. Missiakas D, Mayer MP, Lemaire M, Georgopoulos C, Raina S. Modulation of the Escherichia coli σE (RpoE) heat-shock transcription-factor activity by the RseA, RseB and RseC proteins. Mol. Microbiol. 1997;24:355–371. [PubMed]
29. Schurr MJ, Yu H, Martinez-Salazar JM, Boucher JC, Deretic V. Control of AlgU, a member of the sigma E-like family of stress sigma factors, by the negative regulators MucA and MucB and Pseudomonas aeruginosa conversion to mucoidy in cystic fibrosis. J. Bacteriol. 1996;178:4997–5004. [PMC free article] [PubMed]
30. Li W, Stevenson CE, Burton N, Jakimowicz P, Paget MS, Buttner MJ, Lawson DM, Kleanthous C. Identification and structure of the anti-sigma factor-binding domain of the disulphide-stress regulated sigma factor σR from Streptomyces coelicolor. J. Mol. Biol. 2002;323:225–236. [PubMed]
31. Campbell EA, Muzzin O, Chlenov M, Sun JL, Olson CA, Weinman O, Trester-Zedlitz ML, Darst SA. Structure of the bacterial RNA polymerase promoter specificity σ subunit. Mol. Cell. 2002;9:527–539. [PubMed]
32. Murakami KS, Masuda S, Darst SA. Structural basis of transcription initiation: RNA polymerase holoenzyme at 4 Å resolution. Science. 2002;296:1280–1284. [PubMed]
33. Vassylyev DG, Sekine S, Laptenko O, Lee J, Vassylyeva MN, Borukhov S, Yokoyama S. Crystal structure of a bacterial RNA polymerase holoenzyme at 2.6 Å resolution. Nature. 2002;417:712–719. [PubMed]
34. Colland F, Orsini G, Brody EN, Buc H, Kolb A. The bacteriophage T4 AsiA protein: a molecular switch for sigma 70-dependent promoters. Mol. Microbiol. 1998;27:819–829. [PubMed]
35. Ouhammouch M, Adelman K, Harvey SR, Orsini G, Brody EN. Bacteriophage T4 MotA and AsiA proteins suffice to direct Escherichia coli RNA polymerase to initiate transcription at T4 middle promoters. Proc. Natl Acad. Sci. USA. 1995;92:1451–1455. [PMC free article] [PubMed]
36. Severinova E, Severinov K, Darst SA. Inhibition of Escherichia coli RNA polymerase by bacteriophage T4 AsiA. J. Mol. Biol. 1998;279:9–18. [PubMed]
37. Minakhin L, Niedziela-Majka A, Kuznedelov K, Adelman K, Urbauer JL, Heyduk T, Severinov K. Interaction of T4 AsiA with its target sites in the RNA polymerase σ70 subunit leads to distinct and opposite effects on transcription. J. Mol. Biol. 2003;326:679–690. [PubMed]
38. Sharp MM, Chan CL, Lu CZ, Marr MT, Nechaev S, Merritt EW, et al. The interface of sigma with core RNA polymerase is extensive, conserved, and functionally specialized. Genes Dev. 1999;13:3015–3026. [PMC free article] [PubMed]
39. Lesley SA, Burgess RR. Characterization of the Escherichia coli transcription factor σ70: localization of a region involved in the interaction with core RNA polymerase. Biochemistry. 1989;28:7728–7734. [PubMed]
40. Owens JT, Miyake R, Murakami K, Chmura AJ, Fujita N, Ishihama A, Meares CF. Mapping the σ70 subunit contact sites on Escherichia coli RNA polymerase with a σ70-conjugated chemical protease. Proc. Natl Acad. Sci. USA. 1998;95:6021–6026. [PMC free article] [PubMed]
41. Malhotra A, Severinova E, Darst SA. Crystal structure of a σ70 subunit fragment from E. coli RNA polymerase. Cell. 1996;87:127–136. [PubMed]
42. Ades SE, Grigorova IL, Gross CA. Regulation of the alternative sigma factor σE during initiation, adaptation, and shutoff of the extracytoplasmic heat shock response in Escherichia coli. J. Bacteriol. 2003;185:2512–2519. [PMC free article] [PubMed]
43. Alba BM, Leeds JA, Onufryk C, Lu CZ, Gross CA. DegS and YaeL participate sequentially in the cleavage of RseA to activate the σE-dependent extracytoplasmic stress response. Genes Dev. 2002;16:2156–2168. [PMC free article] [PubMed]
44. Paget MS, Bae JB, Hahn MY, Li W, Kleanthous C, Roe JH, Buttner MJ. Mutational analysis of RsrA, a zinc-binding anti-sigma factor with a thiol-disulphide redox switch. Mol. Microbiol. 2001;39:1036–1047. [PubMed]
45. Li W, Bottrill AR, Bibb MJ, Buttner MJ, Paget MS, Kleanthous C. The role of zinc in the disulphide stress-regulated anti-sigma factor RsrA from Streptomyces coelicolor. J. Mol. Biol. 2003;333:461–472. [PubMed]
46. Maniatis T, Fritsch EF, Sambrook J. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press; Cold Spring Harbor, NY: 1989.
47. Sistrom WR. A requirement for sodium in the growth of Rhodopseudomonas spheroides. J. Gen. Microbiol. 1960;22:778–785. [PubMed]
48. Anthony JR, Green HA, Donohue TJ. Purification of Rhodobacter sphaeroides RNA polymerase and its sigma factors. Methods Enzymol. 2003;370:54–65. [PubMed]
49. Rott MA, Witthuhn VC, Schilke BA, Soranno M, Ali A, Donohue TJ. Genetic evidence for the role of isocytochrome c2 in photosynthetic growth of Rhodobacter sphaeroides Spd mutants. J. Bacteriol. 1993;175:358–366. [PMC free article] [PubMed]
50. Schilke BA, Donohue TJ. ChrR positively regulates transcription of the Rhodobacter sphaeroides cytochrome c2 gene. J. Bacteriol. 1995;177:1929–1937. [PMC free article] [PubMed]
51. Newman JD, Falkowski MJ, Schilke BA, Anthony LC, Donohue TJ. The Rhodobacter sphaeroides ECF sigma factor, σE, and the target promoters cycA P3 and rpoE P1. J. Mol. Biol. 1999;294:307–320. [PubMed]
52. Schilke BA, Donohue TJ. δ-Aminolevulinate couples cycA transcription to changes in heme availability in Rhodobacter sphaeroides. J. Mol. Biol. 1992;226:101–115. [PubMed]
53. Miller JH. Experiments in Molecular Genetics. Cold Spring Harbor Laboratory Press; Cold Spring Harbor, NY: 1972.
54. Guex N, Peitsch MC. SWISS-MODEL and the Swiss-PdbViewer: an environment for comparative protein modeling. Electrophoresis. 1997;18:2714–2723. [PubMed]
55. Barker MM, Gaal T, Josaitis CA, Gourse RL. Mechanism of regulation of transcription initiation by ppGpp. I. Effects of ppGpp on transcription initiation in vivo and in vitro. J. Mol. Biol. 2001;305:673–688. [PubMed]
56. Gruber TM, Bryant DA. Molecular systematic studies of eubacteria, using σ70-type sigma factors of group 1 and group 2. J. Bacteriol. 1997;179:1734–1747. [PMC free article] [PubMed]
57. Gaal T, Bartlett MS, Ross W, Turnbough CL, Jr, Gourse RL. Transcription regulation by initiating NTP concentration: rRNA synthesis in bacteria. Science. 1997;278:2092–2097. [PubMed]
58. Simon R, Piefer U, Puhler A. A broad host range mobilization system for in vivo genetic engineering: transposon mutagenesis in Gramnegative bacteria. Bio/Technology. 1983;1:784–791.
59. Villarejo MR, Zabin I. Beta-galactosidase from termination and deletion mutant strains. J. Bacteriol. 1974;120:466–474. [PMC free article] [PubMed]
60. Yanisch-Perron C, Vieira J, Messing J. Improved M13 phage cloning vectors and host strains: nucleotide sequences of the M13mp18 and pUC19 vectors. Gene. 1985;33:103–119. [PubMed]
61. Ditta G, Schmidhauser T, Yakobson E, Lu P, Liang XW, Finlay DR, Guiney D, Helinski DR. Plasmids related to the broad host range vector, pRK290, useful for gene cloning and for monitoring gene expression. Plasmid. 1985;13:149–153. [PubMed]
62. Karls RK, Wolf JR, Donohue TJ. Activation of the cycA P2 promoter for the Rhodobacter sphaeroides cytochrome c2 gene by the photosynthesis response regulator. Mol. Microbiol. 1999;34:822–835. [PubMed]
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