IraP inhibits intracellular degradation of σ^S

The insert carried by the clone containing iraP corresponds to the region located between 399,461 and 401,605 base pairs on the E. coli K12 chromosome. This region contains the coding sequence of iraP flanked by intergenic regions and part of the flanking genes ddlA and phoA, encoding, respectively, D-alanine-D-alanine ligase A and alkaline phosphatase. A series of deletions in the plasmid defined the iraP coding sequence as the active DNA sequence within this plasmid (data not shown). This assignment was confirmed by subcloning the ORF of iraP with its upstream region (387 nucleotides [nt] upstream of the iraP start codon) into the vector plasmid used for the library, pHDB3, yielding the pIraP plasmid, as described in Materials and Methods.

The effect of multicopy iraP (pIraP) on σ^S stability was measured in cells grown at 37°C in Luria-Bertani broth (LB). As expected, in exponentially growing cells with the vector control (pHDB3), σ^S was rapidly degraded, whereas in cells carrying pIraP, the σ^S half-life was increased approximately threefold (Fig. ​(Fig.1A,1A, exponential phase). In stationary phase, the half-life of σ^S was relatively short, ~3 min in the presence of the vector control (Fig. ​(Fig.1A).1A). However, in stationary phase cells carrying iraP in multicopy, the σ^S half-life increased to ~21 min, a sevenfold increase over the vector control (Fig. ​(Fig.1A).1A). The presence of a multicopy plasmid carrying iraP led to a modest (approximately twofold) increase in the intracellular amount of σ^S, presumably owing to decreased σ^S degradation (Fig. ​(Fig.1A,1A, cf. vector and pIraP at 0 min). This decreased degradation permitted the accumulation and, consequently, the detection of σ^S-LacZ (Fig. ​(Fig.1A,1A, stationary phase), and provides confirmation of the basis for the effect of IraP overproduction on the β-galactosidase activity of the P_BAD-rpoS990’-‘lacZ fusion.

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Figure 1.

Effects of multicopy iraP and deletion of iraP on σ^S stability. The half-life of σ^S was measured in cells grown at 37°C in LB. Protein synthesis was inhibited with chloramphenicol at OD₆₀₀ of ~0.3 and ~3 for the determination of σ^S half-life in exponential phase and stationary phase, respectively. Samples were removed at specific time points and analyzed by immunoblot with an anti-σ^S antiserum. (A) σ^S half-life was determined in the strain CRB316 (P_BAD-rpoS990’-‘lacZ chromosomal fusion) containing either pHDB3, as vector control, or pIraP. The accumulation and the degradation of σ^S-LacZ are shown in the top panel and chromosomally-encoded σ^S in the bottom panel. Quantification of the σ^S signals is shown in the graph. The density for samples at time 0 was set to 100% for each of the chase experiments. Half-lives (t_½) were calculated by regression analysis of the exponential decay of σ^S. (B) Effect of iraP expression from a foreign inducible promoter on σ^S stability in the strain AB009 (ΔiraPkn) carrying either the vector control pP_LlacO or pP_LlacO-iraP. The cells were grown until exponential phase in the presence or absence of 1 mM IPTG, as indicated. (C) Comparison of σ^S half-lives in the ΔiraP (AB006) and wild-type (MG1655) strains.

To confirm that the iraP ORF, rather than any sequences from the regulatory region, was responsible for the stabilization of σ^S, iraP was cloned under the control of the inducible promoter P_LlacO (described in Materials and Methods). The chromosomal copy of iraP was deleted (see Materials and Methods). σ^S half-life was monitored in exponential phase in a ΔiraP strain complemented by the pP_LlacO-iraP plasmid. σ^S was dramatically stabilized when iraP expression was induced with isopropyl-β-D-thiogalactopyranoside (IPTG) in comparison to the uninduced cultures or cells with the vector control (Fig. ​(Fig.1B).1B). σ^S was partially stabilized in strains carrying the uninduced pP_LlacO-iraP plasmid, probably reflecting leaky transcription from the repressed P_LlacO promoter. Furthermore, iraP induction also influenced the steady-state levels of σ^S (Fig. ​(Fig.1B,1B, cf. the time 0 min for each set of samples), presumably due to the effect of IraP on σ^S stability. σ^S levels in cells containing the uninduced or induced P_LlacO-iraP plasmid were increased 10- and 40-fold, respectively, compared with the vector control.

Overexpression of IraP did not affect the activity of either a rpoS’-lacZ transcriptional fusion or a rpoS477’-‘lacZ translational fusion, neither of which is degraded by ClpXP and RssB (data not shown). Thus, IraP does not affect transcription from the rpoS promoter or expression and folding of stable σ^S fusion proteins. Together with our observations of the effect of multicopy iraP on the P_BAD-rpoS990’-‘lacZ fusion, containing a foreign promoter and no upstream sites for translational control, these results strongly suggest that IraP acts at the level of σ^S protein stability.

IraP is required for σ^S stabilization during some but not all starvation conditions

To determine whether the iraP gene is required for the stabilization of σ^S in stationary phase, the half-life of σ^S was measured in both the wild-type strain and a strain deleted for iraP. In exponentially growing cells, the deletion of iraP had only a modest effect on σ^S stability. However, in stationary phase cells growing in rich broth, σ^S half-life was approximately threefold reduced in the ΔiraP strain (Fig. ​(Fig.1C).1C). In cells lacking iraP, σ^S is almost as unstable in stationary phase as in exponential phase in the wild-type strain; therefore, IraP contributes significantly to stationary phase stabilization of σ^S.

Expression of a σ^S-dependent fusion was also examined; as expected from the modest effect on accumulation and stability in LB, there was a slight decrease (twofold or less) in reporter gene expression in the iraP mutant and a slight increase (twofold or less) with iraP on a multicopy plasmid (pIraP) (data not shown).

Stationary phase growth in complex media such as LB involves multiple changes in the available nutrients and the environment. Many specific stress responses lead to stabilization of σ^S, including two well-defined conditions, carbon starvation (Lange and Hengge-Aronis 1994; Zgurskaya et al. 1997) and phosphate starvation (Ruiz and Silhavy 2003; Peterson et al. 2004). To evaluate the role of iraP under these conditions, we assayed σ^S stability following glucose or phosphate limitation in both wild-type and ΔiraP strains. Exponentially growing cells in minimal medium were harvested and resuspended in fresh medium lacking either phosphate or the carbon source. After 1 h of starvation, chloramphenicol was added and σ^S half-life was monitored. Prior to nutrient starvation, σ^S was very unstable, with a half-life of ~2 min (Fig. ​(Fig.2A).2A). Only a small effect of the ΔiraP mutation on σ^S degradation was noticed in exponentially growing cells in minimal media, as was seen in LB (Fig. ​(Fig.1C).1C). After 1 h of glucose starvation, σ^S was dramatically stabilized, as previously reported; σ^S stabilization during carbon starvation was not significantly affected by the absence of IraP (Fig. ​(Fig.2B2B).

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Figure 2.

Effect of ΔiraP on σ^S stability during carbon and phosphate starvation. The half-life of σ^S was measured in cells grown at 37°C in MOPS medium as described in Materials and Methods. σ^S half-life was determined in exponentially growing cells and after 1 h of glucose or phosphate starvation as indicated. Protein synthesis was inhibited with chloramphenicol at an OD₆₀₀ ~0.3. Samples were removed at specific time points and analyzed by immunoblot with an anti-σ^S antiserum. To obtain the same amount of σ^S at time 0 of each of the chase experiments, the amount of cell extract used for exponential phase cultures was twice the volume used for starved cells. σ^S half-lives (t_½) were determined from plots of intensity versus time. (A–C) σ^S degradation was monitored in the wild-type (MG1655) and ΔiraPkn (AB006) strains before (A, exponential phase) and after glucose (B) or phosphate (C) starvation. (D) Complementation of ΔiraP by pIraP. σ^S half-life was determined in phosphate-starved cells with the strain AB006 (ΔiraPkn) containing either the vector control pHDB3 or pIraP. (E) The half-life of σ^S was measured in strains YN868 (rssBD58P) and AB010 (rssBD58P, ΔiraPkn) in both exponential phase and after 1 h of phosphate starvation.

Very different results were obtained after phosphate starvation. As previously observed (Peterson et al. 2004), σ^S became stable when the cells were starved for phosphate. However, in the ΔiraP strain, σ^S remained unstable in phosphate-starved cells (Fig. ​(Fig.2C).2C). Complementation of ΔiraP with pIraP restored stabilization of σ^S during phosphate starvation (Fig. ​(Fig.2D),2D), confirming that IraP is required for full σ^S stabilization during phosphate limitation, although there is also an IraP-independent effect of phosphate starvation (4-min half-life, cf. the 1-min half-life without starvation). These data show that requirements for σ^S stabilization are different during phosphate starvation and glucose starvation; IraP is required for the former but not the latter.

To test whether IraP affects σ^S stabilization only in phosphate starvation, we measured the σ^S half-life in the wild-type and ΔiraP strains during nitrogen starvation, a condition in which σ^S is also stabilized (Mandel and Silhavy 2005). σ^S levels increased only modestly and the protein was only partially stabilized (half-life of ~15 min) after 1 h of nitrogen starvation with the wild-type strain (data not shown), consistent with the data of Mandel and Silhavy (Mandel and Silhavy 2005). In the ΔiraP strain, the σ^S half-life was reduced from 15 to 4 min, indicating that IraP also participates in σ^S stabilization during nitrogen starvation (data not shown).

Our results highlight the differences between different starvation conditions in affecting σ^S stability. IraP is critically important in phosphate starvation and plays a significant role in stationary phase and nitrogen starvation, but has no measurable effect in glucose starvation.

IraP regulates RssB activity by a phosphorylation-independent mechanism

While σ^S stability is controlled by RssB and ClpXP (Muffler et al. 1996; Pratt and Silhavy 1996; Zhou and Gottesman 1998), it was theoretically possible that another system was responsible for σ^S degradation in the absence of iraP. If so, σ^S might be unstable in an iraP mutant host, even in the absence of rssB. This was tested by comparing the σ^S half-life in an rssB-null strain (AB012) with that in a rssB iraP double mutant (AB013). The σ^S half-life was longer than 30 min in both strains (data not shown), consistent with IraP acting within the RssB pathway.

If IraP acts directly within the RssB pathway, does it function to change RssB phosphorylation status? To determine this, we measured σ^S half-life in cells containing a chromosomal rssB allele unable to be phosphorylated. In this strain, the rssB codon for asp58, the location of phosphorylation (Bouche et al. 1998), was mutated to encode proline (rssBD58P, Y. Zhou and S. Gottesman, in prep.), a change previously shown to be partially active for σ^S degradation (Becker et al. 2000). Peterson et al. (Becker et al. 2000) previously observed that cells carrying a chromosomal copy of rssB in which asp58 was mutated to ala retained the ability to promote σ^S degradation in exponentially growing cells, and that σ^S was stabilized after phosphate starvation. Our results agree; σ^S was degraded in exponential phase, albeit at a slightly slower rate than with the wild-type rssB, in both iraP⁺ and ΔiraP cells carrying rssBD58P (Fig. ​(Fig.2,2, cf. A and E, before starvation). After 1 h of phosphate starvation, σ^S was stabilized in the iraP⁺ rssBD58P strain (Fig. ​(Fig.2E).2E). Thus, the rssBD58P mutation did not abolish the ability of the cell to down-regulate σ^S degradation in response to environmental signals, although the stabilization was significantly less than in the rssB⁺ host.

As in the rssB⁺ strain, phosphate starvation did not lead to σ^S stabilization in the ΔiraP derivative of rssBD58P strain (Fig. ​(Fig.2E,2E, phosphate starvation). The effect of IraP overproduction in stabilizing σ^S (Fig. ​(Fig.1A)1A) and of the loss of IraP (ΔiraP strain) in destabilizing σ^S (Fig. ​(Fig.1C)1C) in LB cultures could also be detected in the rssBD58P background (data not shown). Thus, IraP still affects σ^S stability in a strain in which RssB cannot be phosphorylated. These data, taken together, indicate that IraP inhibits σ^S proteolysis by regulating RssB activity in a manner that does not require the potential to phosphorylate or dephosphorylate the asp58 residue.

Identification and characterization of a mutant of IraP

RssB is the limiting protein governing σ^S stability; it is generally made at low levels, and increasing the level of RssB increases σ^S degradation (Pruteanu and Hengge-Aronis 2002). We used a strain, referred to here as rssB⁺⁺⁺, carrying the P_BAD-rpoS990’-‘lacZ fusion as well as a transposon insertion upstream of rssB (rssA2cm) that increases levels of RssB and therefore decreases σ^S levels below our detection limit (Fig. ​(Fig.3A,3A, vector; Ruiz et al. 2001; Mandel and Silhavy 2005) to develop a simple screen for point mutants in iraP that could be used both in vivo and in vitro as controls to further examine IraP mechanism of action.

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Figure 3.

IraP, but not IraPL9S, overcomes RssB overexpression. (A) The half-life of σ^S was measured in exponentially growing cells at 37°C in LB after 15 min of induction with 1 mM IPTG. Protein synthesis was inhibited with tetracycline at an OD₆₀₀ of ~0.3, and samples were removed at specific time points and analyzed by immunoblot with an anti-σ^S antiserum. The strain AB011 (rssA2cm background) containing the vector control pQE-80L, pQE-IraP-His₆, or pQE-IraPL9S-His₆ was used. In the absence of IPTG, σ^S was not detected with any of the plasmids listed above (data not shown). The levels of His-tagged IraP protein expressed were analyzed by immunoblot using an anti-RGS-His₄ antibody. RssB levels were determined by immunoblot with anti-RssB antiserum. (B) Quantification of the levels of σ^S in panel A. σ^S half-lives (t_½) were determined from plots of the log of intensity versus time.

A plasmid carrying iraP-his₆ under control of the Lac repressor (pQE-IraP-His₆) was introduced into the rssB⁺⁺⁺ strain and induced with IPTG for 15 min. The stability of σ^S was then monitored by addition of tetracycline followed by sampling at various times. Figure ​Figure3A3A shows that after 15 min of induction, σ^S was highly expressed and was stable for >30 min, demonstrating both that the IraP-His₆ protein is active and that it can overcome the somewhat higher RssB levels in the rssB⁺⁺⁺ strain. In addition, the somewhat higher RssB levels allowed us to monitor the RssB protein by immunoblot. No change in intracellular levels of RssB was observed in cells overexpressing IraP, and RssB levels remained stable during the tetracycline chase (Fig. ​(Fig.3A).3A). Therefore, IraP does not appear to affect RssB levels or stability.

The effect of IraP-His₆ overexpression on σ^S levels in the rssB⁺⁺⁺ strain can easily be monitored on indicator plates with the P_BAD-rpoS990’-‘lacZ fusion. A strain carrying this fusion and the rssB⁺⁺⁺ mutation is white on MacConkey lactose plates (fusion protein degraded); colonies are dark red on McConkey lactose plates (fusion protein stabilized) when IraP is induced. This change in phenotype allowed us to screen for loss-of-function mutants of iraP using random mutagenesis, as described under Materials and Methods. Several mutants of iraP-his₆, each containing a single amino acid substitution and each able to synthesize significant levels of the protein, were obtained; one, iraPL9S-his₆, has been studied. IraPL9S-His₆ was much less effective at stabilizing σ^S than was IraP-His₆ (Fig. ​(Fig.33).

The levels of the His₆-tagged mutant and wild-type proteins were analyzed by immunoblot. Unexpectedly, the IraPL9S-His₆ mutant protein was more abundant than was IraP-His₆ (Fig. ​(Fig.3A);3A); we do not currently have an explanation for this difference. The levels of IraP (either mutant or wild type) did not change during an antibiotic chase, suggesting that IraP is not itself a substrate for degradation.

IraP interferes directly and specifically with the RssB-mediated degradation of σ^S

To investigate the mechanism of action of IraP and whether it acts directly to stabilize σ^S, we assayed σ^S degradation in vitro with purified proteins. As previously reported (Zhou et al. 2001), σ^S was degraded in the presence of ClpXP, RssB, and ATP, as measured by SDS-PAGE analysis (Fig. ​(Fig.4A).4A). Acetyl phosphate, which is a known phosphate donor for the phosphorylation of RssB (Bouche et al. 1998), also stimulated σ^S degradation, as previously described (Zhou et al. 2001). To test whether IraP acts directly to inhibit σ^S degradation, IraP-His₆ was purified under native conditions (see Materials and Methods) and included in the σ^S degradation reaction mixtures. In the complete reaction mixture, in the absence of IraP, ~80% of σ^S was degraded after 30 min of incubation (Fig. ​(Fig.4A).4A). When IraP was present in the reactions, σ^S was not detectably degraded. Thus, IraP is a direct inhibitor of σ^S degradation.

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Figure 4.

IraP inhibits σ^S degradation by ClpXP and RssB in vitro. The degradation reactions were carried out as described in Materials and Methods with purified components. Reaction products were separated by SDS-PAGE and stained with Coomassie blue. (A) The reactions were performed using 20 pmol of σ^S in the presence or absence of IraP-His₆. RssB, acetyl phosphate, and ClpX were omitted where indicated. (B) The reactions were performed as above, in the presence or absence of either IraP-His₆ or IraPL9S-His₆. (C) Ten picomoles of σ^S was mixed with 10 pmol of GFP-SsrA, in the presence or absence of IraP-His₆, and degradation reactions were performed as above.

We then asked whether the L9S mutation in IraP affects its in vitro activity. Figure ​Figure4B4B shows that IraPL9S-His₆ did not inhibit σ^S degradation. Thus, both in vivo and in vitro the L9S mutant protein is inactive. Figure ​Figure4B4B also shows that IraP-His₆ or IraPL9S-His₆ is not detectably degraded in vitro by ClpXP, confirming that IraP is not itself a substrate of ClpXP.

Given that IraP directly affects σ^S degradation, IraP might act by interfering with the activity of either the ClpXP protease or the adaptor protein RssB, or by protecting σ^S from interacting with either of these factors. To test the first possibility, we examined the effect of IraP on the degradation of another ClpXP substrate, GFP-SsrA (Kim et al. 2000; Singh et al. 2000; Farrell et al. 2005). In the absence of IraP-His₆, when σ^S and GFP-SsrA were incubated together with ClpXP and RssB, both were degraded (Fig. ​(Fig.4C).4C). In the presence of IraP-His₆, σ^S was protected from degradation but GFP-SsrA was not (Fig. ​(Fig.4C,4C, last lane). Similarly, degradation of RepAΔ25-SsrA, another known ClpXP substrate (Sharma et al. 2005), was not inhibited by IraP-His₆ (data not shown). These results indicate that IraP acts specifically on the σ^S degradation pathway and, importantly, that IraP does not inactivate ClpXP.

The physiological function of IraP during phosphate starvation

IraP is a small protein (86 amino acids) predicted to be highly helical. The only homologs are found in other Enterobacteria that also contain rpoS and rssB (Escherichia species, Shigella, Salmonella species, and Erwinia species); the N-terminal region where point mutants were identified is particularly well conserved. In E. coli, S. flexneri, and Salmonella, iraP is located upstream of phoA or psiF, genes induced under phosphate starvation. It was this linkage to phosphate-regulated genes that led us to test the effect of iraP mutations after phosphate starvation; our results show that IraP is particularly important under this stress condition (Fig. ​(Fig.2C2C).

However, while the expression of phoA and other members of the Pho regulon are regulated by the PhoR/PhoB two-component regulatory system in response to phosphate deprivation (Wanner 1996), the IraP-dependent stabilization of σ^S was not affected by a phoB mutation (data not shown). Therefore, the phosphate starvation must affect IraP expression and/or activity by another mechanism. Our data are consistent with the previous observation that RssB is inactivated upon phosphate starvation, in a PhoBR-independent manner (Ruiz and Silhavy 2003). We do not currently have an explanation for how the phosphate starvation signal is sensed through iraP. Northern blots show a clear but transient induction of iraP transcript accumulation (~25-fold) upon phosphate starvation, suggesting transcriptional regulation of iraP expression as at least one component of the regulation (data not shown). Both Northern blots and transcriptional fusions show an increase in iraP expression upon entry to stationary phase (data not shown). If increased transcription of iraP is the primary event leading to stabilization upon starvation, it still remains to be determined what happens when cells exit from phosphate starvation, and whether there is an active process to overcome IraP-dependent stabilization.

Regulated degradation of σ^S is only one of the levels of control of this important protein. It is known, for instance, that σ^S is also up-regulated at the level of synthesis upon phosphate starvation, most probably by a currently undefined small RNA (Ruiz and Silhavy 2003). Possibly because of these complex additional levels of regulation, the changes in the steady-state level of σ^S in iraP mutants are modest. This provides a simple explanation for why mutant searches using screens that reflect σ^S degradation have not previously been successful in identifying targets other that rssB. It is possible, as well, that mutations that impair the stabilization pathway are compensated by an increase in rpoS expression. This increase in synthesis of σ^S may itself also partially titrate RssB (Zhou and Gottesman 1998; Zhou et al. 2001; Pruteanu and Hengge-Aronis 2002). Such an increase in σ^S synthesis might be the explanation for the IraP-independent stabilization of σ^S after phosphate starvation (Fig. 2A,C, cf. exponential phase and phosphate starvation in the iraP mutant), although possibly another anti-adaptor could play a role here as well.

The increase in synthesis and decrease in degradation of σ^S suggest that its accumulation during phosphate starvation is important, but the function of increased σ^S under these conditions is not known. Increased σ^S is found after many stress treatments, including those that induce other, dedicated repair and recovery pathways (such as oxidative stress). σ^S induction may provide a general stress response, before committing the cell to the full-fledged specific response. In some agreement with this, a recent report suggests that high levels of σ^S negatively affect most of the phosphate starvation genes, possibly by competition with the primary σ factor, σ⁷⁰, for core RNA polymerase (Taschner et al. 2004); this paradoxical effect might reflect a need to delay the PhoB-dependent response. σ^S positively controls the expression of pstS, which encodes the high-affinity inorganic phosphate transporter Pst; transport of phosphate through this system may contribute to the block in induction of the Pho system (Taschner et al. 2004).

IraP-independent control of σ^S stability

IraP regulates σ^S turnover during phosphate starvation and participates in the stabilization of σ^S during nitrogen starvation and stationary phase (Figs. ​(Figs.1C,1C, ​,2C;2C; data not shown). However, IraP is not needed for σ^S stabilization during glucose starvation (Fig. ​(Fig.2B).2B). These results indicate that other mechanisms or pathways, independent of IraP, participate in the control of σ^S turnover.

It is unlikely that RssB activity is titrated by a high amount of σ^S in glucose-starved cells because the rate of σ^S synthesis is very low in these conditions (McCann et al. 1993; Lange and Hengge-Aronis 1994; Zgurskaya et al. 1997; Mandel and Silhavy 2005). The simplest explanation would be the existence of another anti-RssB regulator, which would be induced or become active specifically during glucose starvation and would play a similar role to IraP (Fig. ​(Fig.6).6). Additional evidence for other regulators is suggested by the stabilization of σ^S in cells carrying an hns mutation (Yamashino et al. 1995; Y. Zhou and S. Gottesman, in prep.), encoding the histone-like protein H-NS, which represses expression of many genes. The stabilization of σ^S in the hns mutant is not affected by ΔiraP (data not shown), suggesting that at least one other anti-RssB regulator is repressed by H-NS.

The regulation of σ^S degradation results from the integration of many input signal pathways (Hengge-Aronis 2002). IraP clearly controls σ^S stabilization, and its overexpression leads to a significant accumulation of σ^S. Mandel and Silhavy (Hengge-Aronis 2002) pointed out that starvations for different nutrients do not trigger identical responses. We believe that starvation for different nutrients and different stress conditions elicit specific signal transduction pathways converging to regulate σ^S stability. These pathways may overlap to some extent because we observed that IraP stabilizes σ^S in more than one condition, although the most pronounced effect was upon phosphate starvation.

Media and growth conditions

Cells were grown in LB or morpholinepropanesulfonic acid (MOPS) minimal medium (Teknova) supplemented with 0.4% glucose, 0.2% (NH₄)₂SO₄, 1.32 mM K₂HPO₄, and 1 μg/mL thiamine. MacConkey agar plates with 1% lactose, containing ampicillin when used with plasmids, were used in analyses of strains carrying the P_BAD-rpoS990’-‘lacZ chromosomal fusion. All liquid cultures were grown under aerobic conditions at 37°C, and growth was monitored by measuring the optical density at 600 nm (OD₆₀₀). Exponential phase and stationary phase correspond to an OD₆₀₀ of ~0.3 and ~3, respectively. Stationary phase cells were obtained 2 h after the break in the exponential growth curve.

For starvation experiments, MOPS was made without K₂HPO₄ (phosphate starvation medium), without (NH₄)₂SO₄ (nitrogen starvation medium), or without glucose (carbon starvation medium). Cells were first grown overnight in complete MOPS minimal medium, then subcultured into fresh minimal glucose medium at an OD₆₀₀ of 0.01 (~1:400 dilution) and grown to mid-logarithmic phase (OD₆₀₀ of ~0.3) for the following treatments. Cells were washed twice by filtration with prewarmed (37°C) starvation medium, resuspended in the same volume of starvation medium, and incubation at 37°C was continued for 1 h.

Assay for σ^S degradation in vivo

Cells were grown in LB or in MOPS media as described above. Exponential phase refers to an OD₆₀₀ of ~0.3 and stationary phase to an OD₆₀₀ of ~3. Cells were treated with chloramphenicol (200 μg/mL) or tetracycline (100 μg/mL), and 1-mL samples were removed at the indicated time points and treated as described below.

Electrophoresis and immunoblot analysis of proteins

Whole-cell extracts were prepared as follows: Cells grown in LB or MOPS media were assayed for OD₆₀₀, and 1-mL samples were removed and precipitated with 5% ice-cold tricarboxylic acid (TCA). Precipitated pellets were washed with 500 μL of 80% cold acetone and then resuspended in a volume of sodium dodecyl sulfate (SDS) sample buffer normalized to the OD₆₀₀. Samples were analyzed using NuPAGE 12% Bis-Tris gels (Invitrogen), transferred, and probed with a 1:4000 dilution of anti-σ^S antiserum, a 1:1200 dilution of anti-RssB antiserum, or a 1:1000 dilution of anti-RGS-His₄ antiserum (Qiagen). The blots were developed with the ECL system (Amersham) and quantified with ImageJ Software (National Institutes of Health).

Library screening

The multicopy plasmid DNA library screen was performed as described previously (Majdalani et al. 2001). Briefly, transformations were performed by electroporation of a pBR322-based library (Ulbrandt et al. 1997) into CRB316, plated on MacConkey lactose plates containing 50 μg/mL ampicillin to give ~500 colonies per plate, and incubated at 37°C. We screened ~15,000 colonies and isolated six red colonies and seven white colonies. Plasmid DNA was purified using Qiagen’s Minipreps kit, and insert junctions were sequenced using primers pBRlib.for (5′-CCTGACGTCTAAGAAACCATTATTATC-3′) and pBRlib.rev (5′-AACGACAGGAGCACGATCATGCG-3′).

Isolation of point mutants in iraP by random mutagenesis

Random mutagenesis of iraP was performed at a low mutation rate (three to 16 mutations per kilobase) using the GeneMorph II EZClone Domain Mutagenesis Kit (Stratagene) according to the manufacturer’s specifications. A mutant plasmid library of iraP was generated using pQE-IraP-His₆ as template and the primers RMiraP-HisF (5′-CAGAATTCATTAAAGAGGAGA AATTAACTATG-3′) and RMiraP-HisR (5′-GTGATGCGATC CTCTTCC-3′). XL10-Gold ultracompetent cells were transformed with 1.5 μL of EZclone reaction mixture. The transformed cells were cultured in a total volume of 150 mL of LB supplemented with 100 μg/mL ampicillin for 10 h at 37°C. Plasmid library DNA was isolated using HiSpeedPlasmid Maxi Kit (Qiagen) and eluted with 600 μL of 1× TE buffer.

The screen for nonfunctional mutants of iraP was done by electroporation of the iraP mutant plasmid library (see above) into AB011, a strain carrying the chromosomal P_BAD-rpoS990’-‘lacZ fusion and a rssA2cm mutation, which leads to overexpression of rssB from the chromosome (Ruiz et al. 2001; Mandel and Silhavy 2005). When transformed with pQE-80L, AB011 gave white colonies (Lac⁻), whereas transformation with pQE-IraP-His₆ gave red colonies (Lac⁺) on MacConkey plates supplemented with 1% lactose and 50 μg/mL ampicillin. We used this phenotype to isolate mutations within the iraP coding sequence. Among ~10,000 red colonies, 14 white clones were identified. The plasmids isolated were transformed again into AB011, and all gave the expected Lac⁻ phenotype. To discriminate between full-length nonfunctional mutants of IraP and truncated proteins due to the introduction of stop codons, colony-immunoblot against the C-terminal RGS-His₆ tag was performed using an anti-RGS-His₄ antibody (Qiagen) according to the manufacturer’s instructions. The RGS-His₆ tag was detected for five clones out of the 14 white colonies isolated. Sequence analysis showed that four plasmids each contained a single substitution; all were in the 5′ region of the coding sequence of iraP (the most conserved region); three of these, L4P, L9S, and A13D, were in fully conserved residues, and one, A6T, was in a partially conserved residue. Only the L9S substitution, which seemed to be the tightest mutant, was further studied; this mutation changes a TTA codon to TCA.

Proteins

Overproduction and purification of IraP-His₆ and IraPL9S-His₆ were performed as follows: The DH5α strain (Invitrogen), transformed with pQE-IraP-His₆ or pQE-IraPL9S-His₆, was grown in LB medium containing 100 μg/mL ampicillin to an OD₆₀₀ of 0.6. IPTG was added to 1 mM. After 5–6 h of induction, cells were harvested and stored at −80°C until use. Cell pellets were resuspended in lysis buffer (50 mM NaH₂PO₄, 300 mM NaCl, 10 mM imidazole at pH 8.0) and sonicated. The lysates were centrifuged at 10,000 × g for 30 min at 4°C, and the soluble fractions were applied to a Ni²⁺-NTA column (Invitrogen) equilibrated with lysis buffer. The column was washed with 10 column volumes of buffer A (50 mM NaH₂PO₄, 300 mM NaCl, 20 mM imidazole at pH 8.0), followed by elution with 250 mM imidazole, in buffer A. The eluates were desalted using an Amersham Biosciences PD-10 column equilibrated with 50 mM Tris-HCl (pH 7.5), 0.1 mM EDTA, 100 mM KCl, and 0.1 mM DTT. Glycerol was added to a final concentration of 10% (v/v), and the samples were stored at −80°C. Final preparations of IraP-His₆ and IraPL9S-His₆ were >90% pure as judged by SDS-PAGE.

ClpX (Grimaud et al. 1998), ClpP (Maurizi et al. 1994), and RepA(Δ25)-SsrA (Sharma et al. 2005) were isolated as described. σ^S was isolated from cells carrying pLHN14 and expressing σ^S (Nguyen et al. 1993) using a described procedure (Tanaka et al. 1993) with some modifications. RssB was purified as described, with minor modifications (Zhou et al. 2001). His₆-GFP-SsrA was isolated by Talon resin (Clontech) column chromatography as described by the manufacturer. All proteins were>90% pure as determined by Coomassie staining following SDS-PAGE. Protein concentrations were determined using the Bradford protein assay kit (Bio-Rad) and bovine serum albumin as a standard. Throughout, proteins are expressed as moles of RssB monomers, ClpX hexamers, ClpP tetradecamers, σ^S monomers, His₆-GFP-SsrA monomers, IraP-His₆ monomers, and IraPL9S-His₆ monomers.

Assay for σ^S degradation in vitro

Reaction mixtures were assembled in a 20-μL final volume of buffer (20 mM Tris-HCl at pH 7.5, 10 mM MgCl₂, 140 mM KCl, 1 mM DTT, 0.1 mM EDTA, 5% glycerol [v/v], 0.005% Triton X-100 [v/v]) containing 5 mM ATP, 10 mM acetyl phosphate, σ^S (20 pmol), IraP (20–80 pmol), RssB (1 pmol), ClpX₆ (2 pmol), and ClpP₁₄ (2 pmol), unless otherwise indicated. All of the proteins were diluted into the reaction buffer containing 0.05% Triton X-100 before use. The mixtures were incubated at room temperature for 30 min, and the reactions were stopped by the addition of 20 μL of SDS sample buffer. Protein samples were analyzed by SDS-PAGE. SeeBlue Plus2 (Invitrogen) prestained protein standards were used for molecular weight estimation.

We thank M. Mandel, N. Ruiz, and T. Silhavy for providing the rssA2cm mutation, and N. Majdalani for providing strain NM1100. We are grateful to C. Ranquet for making the P_BAD-rpoS-lacZ translational fusion used in this study. We thank Y. Zhou for making the rssBD58P chromosomal mutation. We thank M. Guillier for providing the pP_LlacO plasmid. We thank members of the Gottesman laboratory and Michael Maurizi for useful discussions and suggestions, and Carin Vanderpool and Michael Maurizi for their comments on the manuscript. This research was supported by the Intramural Research Program of the NIH, National Cancer Institute, Center for Cancer Research.