Present addresses: CNRS-UMR 5163, Laboratoire Adaptation et Pathogénie des Micro-organismes, Université Joseph Fourier GRENOBLE 1, BP 170, F-38042 Grenoble cedex 9, France;
Multiple pathways for regulation of σS (RpoS) stability in Escherichia coli via the action of multiple anti-adaptors
Article first published online: 23 MAR 2008
Journal compilation © 2008 Blackwell Publishing Ltd; No claim to original US government works
Volume 68, Issue 2, pages 298–313, April 2008
How to Cite
Bougdour, A., Cunning, C., Baptiste, P. J., Elliott, T. and Gottesman, S. (2008), Multiple pathways for regulation of σS (RpoS) stability in Escherichia coli via the action of multiple anti-adaptors. Molecular Microbiology, 68: 298–313. doi: 10.1111/j.1365-2958.2008.06146.x
- Issue published online: 23 MAR 2008
- Article first published online: 23 MAR 2008
- Accepted 25 January, 2008.
- Top of page
- Experimental procedures
- Supporting Information
σS, the stationary phase sigma factor of Escherichia coli and Salmonella, is regulated at multiple levels. The σS protein is unstable during exponential growth and is stabilized during stationary phase and after various stress treatments. Degradation requires both the ClpXP protease and the adaptor RssB. The small antiadaptor protein IraP is made in response to phosphate starvation and interacts with RssB, causing σS stabilization under this stress condition. IraP is essential for σS stabilization in some but not all starvation conditions, suggesting the existence of other anti-adaptor proteins. We report here the identification of new regulators of σS stability, important under other stress conditions. IraM (inhibitor of RssB activity during Magnesium starvation) and IraD (inhibitor of RssB activity after DNA damage) inhibit σS proteolysis both in vivo and in vitro. Our results reveal that multiple anti-adaptor proteins allow the regulation of σS stability through the regulation of RssB activity under a variety of stress conditions.
- Top of page
- Experimental procedures
- Supporting Information
In both eukaryotic and prokaryotic organisms, regulated proteolysis plays important roles in a variety of physiological processes such as apoptosis, cell differentiation and stress responses. In addition to the function of proteolysis in quality control, cytoplasmic energy-dependent proteases play a substantial regulatory role by controlling intracellular levels of regulatory proteins (Gottesman, 2003). In eukaryotic cells, the polyubiquitin ligase machinery targets most short-lived regulatory proteins to the proteasome system (reviewed in Hershko and Ciechanover, 1998). In bacteria, ubiquitin tagging does not exist and specificity for protein breakdown depends on the primary sequence motifs of substrate proteins that are directly recognized by proteases or, in some cases, on adaptor proteins (for reviews see Gottesman, 2003; Jenal and Hengge-Aronis, 2003; Ades, 2004). In Escherichia coli, one of these adaptor proteins is RssB, also called MviA in Salmonella (Bearson et al., 1996; Muffler et al., 1996; Pratt and Silhavy, 1996; Zhou and Gottesman, 1998). RssB is specifically required for the proteolysis of the σS subunit of RNA polymerase by the ClpXP protease (Bearson et al., 1996; Muffler et al., 1996; Pratt and Silhavy, 1996; Zhou and Gottesman, 1998; Zhou et al., 2001). RssB binds directly to σS and targets it to the protease (Zhou et al., 2001).
σS (σ38 or RpoS) is the master transcriptional regulator of stationary phase and the general stress response (Loewen and Hengge-Aronis, 1994; Loewen et al., 1998). σS controls the expression of ∼100 genes involved in different physiological functions that help the cells to cope with starvation and stressful conditions (Loewen et al., 1998; Hengge-Aronis, 2002; Lacour and Landini, 2004). A complex regulatory network controls σS expression, stability and activity, allowing integration of different environmental parameters (reviewed in Hengge-Aronis, 2002; Bougdour et al., 2004; Gaal et al., 2006). In exponentially growing cells, σS is maintained at a very low level owing in large part to active degradation by the ClpXP protease (Lange and Hengge-Aronis, 1994; Schweder et al., 1996). σS becomes stable in stationary phase or under starvation conditions, allowing a rapid accumulation of σS in the cells (reviewed in Hengge-Aronis, 2002). How this change in environmental and growth conditions is sensed and transmitted to affect σS turnover has been the subject of much interest.
RssB belongs to the two-component response regulator family of proteins and like the other members of this family, its activity can be modulated by phosphorylation of a highly conserved residue, asp58 in the case of RssB (Bouchéet al., 1998). Although effects of phosphorylation of RssB have been observed in purified in vitro degradation assays of σS (Zhou et al., 2001), a non-phosphorylatable RssB protein, owing to a mutation of the phosphorylation site, is still active and regulated in vivo (Peterson et al., 2004; Bougdour et al., 2006).
Identification of the anti-adaptor protein IraP provided an explanation for some of these observations. IraP inhibits σS proteolysis through a direct interaction with RssB in a manner that is independent of the phosphorylation of RssB (Bougdour et al., 2006). The promoter of iraP requires ppGpp, made in higher amounts after phosphate starvation, and the synthesis of IraP is induced by phosphate starvation dependent upon ppGpp (Bougdour and Gottesman, 2007); as a result, IraP stabilizes σS in response to phosphate starvation in E. coli and Salmonella (Bougdour et al., 2006; Tu et al., 2006). However, in Salmonella, Mg2+ starvation also induces IraP; the promoter of iraP is upregulated directly by the transcriptional regulator PhoP under this starvation condition, from an upstream promoter that is not conserved in E. coli (Tu et al., 2006). Consistent with stabilization of σS under inducing conditions, σS is stabilized in the presence of low Mg2+ concentrations in Salmonella, dependent upon IraP, while partial stabilization is seen in E. coli after Mg2+ starvation, this is independent of IraP (Tu et al., 2006).
IraP allows phosphate-starvation signal transduction to σS through RssB. However, σS is stabilized in response to glucose starvation or in an hns mutant strain in a manner that is independent of IraP (Bougdour et al., 2006; Zhou and Gottesman, 2006). Therefore, σS degradation is inhibited by other mechanisms; we hypothesized that other anti-adaptor proteins exist, each regulating RssB activity under a specific stress condition.
The work described here was undertaken in order to identify regulators other than IraP involved in the regulation of σS proteolysis. We report the isolation and characterization of two new regulators of σS turnover encoded by the previously uncharacterized yjiD and ycgW genes, and the identification of a role for the transcriptional regulator AppY, possibly by regulating an unidentified anti-adaptor. YjiD and YcgW modulate the stability of σSin vivo and in vitro by counteracting the activity of RssB, resulting in the stabilization of σS. Because YcgW is critically important for stabilization of σS during magnesium starvation, we have renamed this gene IraM for inhibitor of RssB activity in response to magnesium starvation. Because YjiD was shown by H. Merrikh and S. Lovett (pers. comm.) to be critically important for survival during DNA-damaging conditions (see Discussion), this gene was renamed IraD for inhibitor of RssB activity in response to DNA damage.
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- Experimental procedures
- Supporting Information
Identification of novel regulators of σS stability in E. coli and Salmonella typhimurium
As discussed above, there are still unidentified com ponents involved in the regulation of σS degradation. Previous work from our laboratory has shown that the use of a translational fusion PBAD–rpoS990′–′lacZ (Ranquet and Gottesman, 2007) that bypasses the normal signals controlling rpoS transcription and translation allowed an efficient identification of novel cellular regulators of σS degradation (Bougdour et al., 2006). The strain carrying this fusion, CRB316, gave slightly Lac+ (red) colonies on MacConkey lactose plates without arabinose but became Lac– (white) when deleted for iraP, consistent with increased degradation of σS in the absence of IraP. In order to identify regulators other than IraP that stabilize σS, the strain lacking IraP (AB007) was transformed with a pBR322-based E. coli genomic DNA library (Ulbrandt et al., 1997), with the expectation that overproduction of negative regulators of σS degradation from the plasmid would give red (Lac+) colonies on MacConkey lactose indicator plates. Seventeen Lac+ colonies were isolated out of ∼30 000 colonies screened, including some that contained the ara regulators (see Table 1 and Experimental procedures). One plasmid contained iraP, expected to complement the iraP deletion and increase σS stability. The other plasmids identified three additional regions of the E. coli chromosome; one of these, with the weakest effect, was not further studied (Table 1, plasmid MR′2). Another class, containing eight overlapping plasmids with varying end-points, all contained yjiD; the smallest of these plasmids, MR′14, was chosen for further study. The final class, with one isolate (MR′22), contained appY and fragments of flanking genes. Both MR′14 and MR′22 increased expression of the rpoS750′–′lacZ fusion and therefore were expected to act on either σS translation or stability. These two plasmids were directly tested for their effects on σS turnover, as described in Experimental procedures, and were found to stabilize it.
|Plasmids||Fragment boundaries||Genes presentb||Description of genesc||Frequency of isolates||Effect on PBAD–rpoS990′–′lacZ||Effect on rpoS750′–′lacZ|
|MR′7||69361–72447||< araB′||araB: ribulokinase.||6/17||Yes||No|
|> araC||araC: regulatory gene: activator and repressor, known regulator of the PBAD promoter.|
|> yabI||yabI: unknown function.|
|< thiQ′||thiQ: homologue to Salmonella thiamine transporter.|
|MR′14||4553207–4555503||< yjiC′||yjiC: unknown function.||8/17||Yes||Yes|
|> yjiD||yjiD: unknown function (small ORF, 130 aa).|
|> ′yjiE||yjiE: unknown function.|
|MR′22||582236–584460||> ylcE′||ylcE: unknown function.||1/17||Yes||Yes|
|> appY||appY: transcriptional regulator, regulates hya and appA operons; induced by PO4 starvation and stationary phase.|
|< ′ompT||ompT: outer membrane protease VIII.|
|MR′23||399520–401121||< ′ddlA||ddlA: d-alanine:d-alanine ligase, cell wall biosynthesis.||1/17||Yes||Yes|
|> iraP||iraP: inhibitor of RssB activity in response to Phosphate starvation, small ORF, 86 aa.|
|> ′phoA||phoA: alkaline phosphatase.|
|MR′2||2190932–2192713||< mrp||mrp: putative ATPase; in Salmonella, homologue is part of alternative pyrimidine pathway to thi.||1/17||Yes||ND|
|> ′metG||metG: methionyl-tRNA synthetase.|
While the screen described above was for higher levels of an unstable rpoS–lacZ hybrid protein, an alternative approach to identifying genes that may stabilize σS is to screen for increased σS activity, using a σS-dependent reporter system. Such a screen was performed in Salmonella typhimurium in strain TE6756, carrying a katE′–lac transcriptional fusion. An otherwise wild-type strain gave slightly Lac+ (red) colonies on MacConkey lactose plates. A multicopy plasmid library of S. typhimurium-DNA fragments cloned in the plasmid pBR328 (Hmiel et al., 1986) was introduced into the S. typhimurium strain and a single colony with increased expression of the reporter was found among ∼10 000 colonies screened; the corresponding plasmid was named pTE697. This plasmid was then introduced into four strains, each carrying a different σS-dependent transcriptional reporter fusion (proV′–lac, o186′–lac and otsA′–lac) (Cunning and Elliott, 1999). In all four strains, the presence of pTE697 resulted in a 5- to 10-fold increase in β-galactosidase activity compared with the vector control (Fig. 1A). The pTE697 plasmid contained the complete coding sequences of STM1110, renamed rssC (regulator of sigma SC), encoding a previously uncharacterized protein, and STM1109, a gene encoding a putative hydrolase (Fig. 1B). The smallest plasmid in a series of deletion derivatives of pTE697 that still gave full induction of katE′–lac was pTE698, which contains ∼1 kb of DNA extending from the right end of the original clone to an ApaI site. The DNA sequence of the insert in pTE698 was determined (GenBank Accession AF 214144). The sequence contains a single predicted ORF of 363 nt encoding for a putative protein of 120 amino acids. The rssC gene carried on pTE698 was disrupted by insertion of a Kn cassette into the PstI site at codon 94 of the gene; this insertion eliminated the effect of the plasmid on katE–lac expression (data not shown), indicating that the rssC coding sequence was the active DNA sequence within the plasmid responsible for stabilizing σS. The effect of RssC on σS turnover was directly assayed in vivo by pulse labelling and immunoprecipitation of σS. The half-life of σS was less than 3 min in the strain carrying the vector control and was increased 10-fold (half-life of ∼30 min) in the strain overexpressing RssC (data not shown), indicating that RssC overproduction inhibits σS degradation in S. typhimurium.
IraD, IraM and AppY inhibit intracellular degradation of σS in E. coli
The inserts carried by the clones MR′14 and MR′22, obtained from the screen in E. coli, contained the coding sequences of yjiD and appY respectively, flanked by intergenic regions and part of the flanking genes (Table 1). Both appear to be independent transcription units (H. Merrikh and S. Lovett, pers. comm.; Atlung et al., 1996).
yjiD (to be called iraD, see below) encodes a protein of 130 amino acids of unknown function. appY encodes a transcriptional regulator of 249 amino acids. iraD and appY were cloned under the control of the inducible promoter PT5/lacO (described in Experimental procedures) to create pQE-iraD and pQE-appY.
rssC is found only in very closely related species, but not in E. coli. The most closely related protein to RssC in E. coli is YcgW (also called ElbA), a 107 amino acid protein 40% identical to RssC; YcgW is found only in Shigella and E. coli. This pattern and the low GC content (38% and 34% GC respectively) suggest recent acquisition by horizontal gene transfer. The characterization of RssC was not further explored in Salmonella. Instead, the E. coli protein was further examined and proved to have a similar effect to RssC. ycgW (renamed iraM, see below) was cloned under the control of the same inducible promoter (PT5/lacO) to create pQE-iraM.
σS half-life was measured in exponential phase in a strain with iraP deleted and carrying the vector or one of the plasmids pQE-iraD, pQE-appY or pQE-iraM. σS was dramatically stabilized, in comparison to the uninduced cultures or cells with the empty vector control, when any of these three plasmids was induced with isopropyl-β-d-thiogalactopyranoside (IPTG) (Fig. 2A and B). σS was partially stabilized in strains harbouring the uninduced pQE-iraD, pQE-appY or pQE-iraM plasmids, probably a consequence of leaky transcription from the repressed T5/lacO promoter. Furthermore, induction also influenced the steady-state amounts of σS (Fig. 2A and B, cf. the time 0 min for each set of samples), presumably owing to the effect of IraD, AppY and IraM on σS stability before the chase was begun. All three of these plasmids also increased expression of PBAD–rpoS990′–′lacZ fusion (data not shown), containing a foreign promoter and none of the known cis-regulatory sequences of rpoS. Thus, these results strongly suggest that these regulators act directly or indirectly at the level of σS protein stability.
In theory, any of these genes might act through one of the others, rather than on its own to stabilize σS. Both IraD and IraM are small proteins, as is IraP, possibly suggesting they are directly acting as anti-adaptors, although they share no sequence similarity to IraP. AppY, on the other hand, is a known transcriptional regulator, suggesting that it may act by increasing the transcription of one or more anti-adaptors.
Because IraD and AppY were identified as stabilizing σS in an iraP mutant, their action must be independent of IraP. The epistasis of the plasmids was further explored by creating cells mutant in iraD, iraM or iraP, as well as a triple mutant iraD iraM iraP. The effects of pQE-iraD, pQE-appY and pQE-iraM plasmids on σS was monitored in strains carrying an rpoS750′–′lacZ translational fusion as well as a transposon insertion upstream of rssB (rssA2::cm) that increases levels of RssB and therefore decreases levels of the rpoS fusion protein (Ruiz et al., 2001; Mandel and Silhavy, 2005). Table 2 shows that IraD and IraM both act independently of each other, and AppY action is independent of IraD, IraM or IraP. A direct assay of σS degradation in an iraD iraM iraP triple mutant (strain AB049) showed that AppY was able to stabilize σS (> 30 min half-life) even when all three of the other proteins are absent, suggesting that it either acts directly or, more likely, via a yet unidentified anti-adaptor (Fig. 2C).
IraD and IraM interfere directly and specifically with the RssB-mediated proteolysis of σS
A direct test for the mode of action of IraD and IraM was carried out by purifying the proteins and testing them in an in vitroσS degradation system. As previously reported (Zhou et al., 2001; Bougdour et al., 2006), σS was rapidly degraded in the presence of ClpXP, RssB, Acetyl-phosphate and ATP, as measured by sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) analysis (Fig. 3A). RepA-SsrA, an RssB-independent substrate of ClpXP (Sharma et al., 2005), was included in the reaction, and was also rapidly degraded (Fig. 3A). To test whether IraD and IraM act directly to inhibit σS proteolysis, IraD-His6 and His6-IraM (both still active when expressed from a plasmid in stabilization of an RpoS–LacZ fusion, as judged by colour on MacConkey lactose agar) were purified under native conditions (see Experimental procedures) and included in the σS degradation reaction mixtures. In the complete reaction mixtures, in the absence of IraD and IraM, ∼90% of σS was degraded after 30 min of incubation (Fig. 3A). When IraD or IraM were present in the reactions, σS was poorly degraded, indicating that IraD and IraM are both direct inhibitors of σS degradation; RepA-SsrA was still degraded efficiently. Therefore, IraD and IraM do not interfere with general ClpXP activity, but specifically block the RssB-dependent degradation of σS. Fig. 3A also shows that neither IraD-His6 nor His6-IraM were detectably degraded in vitro by ClpXP, suggesting that IraD and IraM are not themselves direct substrates of ClpXP.
IraD interacts directly with the adaptor protein RssB
The results presented above showed that IraD and IraM control the half-life of σS and act within the RssB/ClpXP pathway to protect σS from degradation. These results are strikingly similar to those observed with the anti-adaptor protein IraP (Bougdour et al., 2006). IraP stabilizes σS by interaction with RssB, probably blocking RssB interaction with σS (Bougdour et al., 2006). Purified IraD-His6 or His6-IraM were tested for their ability to bind to RssB. The His-tagged proteins were bound to Ni2+-Talon-magnetic beads and incubated with purified RssB. Following elution with imidazole buffer and analysis by immunoblot, ∼80% of the RssB introduced to the IraD-His6-Ni2+-Talon-magnetic bead matrix was retained and coeluted with IraD-His6[Fig. 3B, compare lane 1 (Input) with lane 3 (protein eluted)]. With the His6-IraM-Ni2+-Talon-magnetic beads matrix, ∼30% of the RssB introduced (input) was coeluted with His6-IraM. Note that low levels of His6-IraM were detected in the eluted fraction with the anti-RGS-His4 antibody (Fig. 3B, lane 4), probably reflecting a poor binding efficiency of the His6-IraM protein to the Ni2+-Talon-magnetic beads. RssB did not bind to the Ni2+-Talon-magnetic beads in the absence of IraD-His6 and His6-IraM (Fig. 3B, lane 2).
A similar experiment was performed to probe for association of σS and IraD-His6 or His6-IraM. Figure 3C shows that σS was not retained by the IraD-His6-Ni2+-Talon-magnetic beads matrix and therefore was not coeluted with IraD-His6. A somewhat different result was observed with His6-IraM. Despite low levels of His6-IraM bound to the Ni2+-Talon-magnetic beads, ∼10% of the σS introduced (input) was coeluted with His6-IraM. These data indicate that IraD interacts directly with RssB, but not with σS. No clear-cut conclusion can be drawn for IraM; the protein could be interacting with both RssB and σS.
IraM is required for σS stabilization during Mg2+/Ca2+ starvation and is regulated by PhoP/PhoQ
Several stress responses have been shown to lead to stabilization of σS in E. coli, including carbon starvation (Lange and Hengge-Aronis, 1994; Zgurskaya et al., 1997), phosphate starvation (Ruiz and Silhavy, 2003; Peterson et al., 2004; Bougdour et al., 2006), and, to some extent, Mg2+ starvation (Tu et al., 2006). Given that AppY is a known transcriptional regulator, we focused our interest on the two small ORFs, iraD and iraM. To determine whether iraM or iraD are responsible for the stabilization of σS in these conditions, the half-life of σS was measured in wild-type and in strains deleted for either iraM or iraD after various starvation treatments. Exponentially growing cells in minimal medium were harvested and re-suspended in fresh medium lacking either the carbon source or Mg2+. After 1 h of starvation, chloramphenicol was added and σS half-life was monitored by Western blot. Prior to nutrient starvation, σS was very unstable in all strains tested, with half-lives from less than 1 min to ∼2 min (Fig. 4A). In glucose-starved cells, σS was dramatically stabilized as previously reported (Lange and Hengge-Aronis, 1994; Zgurskaya et al., 1997; Mandel and Silhavy, 2005; Bougdour et al., 2006); however, none of the deletions had any detectable effect on the stabilization of σS during glucose starvation (data not shown). Very different results were obtained during Mg2+ starvation. As previously described (Tu et al., 2006), σS was stabilized in the wild-type cells with a half-life of ∼10 min. σS remained unstable in cells lacking iraM (Fig. 4B); deletion of iraP or iraD had no effect on σS stability under these conditions. These data show that IraM is essential for σS stabilization during Mg2+ starvation in E. coli.
The above data indicate that IraM specifically controls σS turnover in response to Mg2+ starvation. Therefore, we hypothesized that iraM was transcriptionally upregulated in this condition, as iraP is under phosphate starvation. To study iraM expression, a transcriptional fusion, PiraM–lacZ, fusing the promoter region of iraM (−700 to +287 nt relative to the ATG start codon of iraM) to lacZ (see Experimental procedures) was used to monitor iraM expression after cells were grown in the presence or absence of Mg2+/Ca2+. The activity of the PiraM′–lacZ transcriptional fusion was ∼26-fold higher after 1 h of starvation for Mg2+/Ca2+ compared with cells growing continuously in the presence of Mg2+/Ca2+, indicating that the iraM promoter was induced during Mg2+/Ca2+ starvation (Fig. 4C).
Magnesium starvation is sensed by the PhoQ histidine kinase and the signal is transduced to the response regulator PhoP, which controls the transcription of a set of genes involved in the adaptation to low Mg2+/Ca2+ environments (Garcia-Vescovi et al., 1996; Groisman, 2001; Minagawa et al., 2003; Zwir et al., 2005). In Salmonella, iraP is regulated by PhoP during Mg2+/Ca2+ starvation, and σS is stabilized in a PhoP-dependent manner under these conditions (Tu et al., 2006). Therefore, an obvious candidate for regulating IraM expression was the two-component regulatory system PhoP/PhoQ. Figure 4C shows that in the absence of PhoP, the activity of an PiraM′–lacZ fusion in cells starved for Mg2+/Ca2+ remained as low as cells grown in the absence of starvation (exponentially growing cells). Note that no increase in the OD600 was observed after removal of Mg2+/Ca2+ from the culture by filtration in either phoP+ or phoP– strains (data not shown). These data indicate that PhoP regulates directly or indirectly the transcription of iraM. Analysis of the promoter region of the iraM gene revealed the presence of at least two putative PhoP binding sites (Fig. S1).
IraD and DNA damage
Although there was not much previously published on yjiD (iraD), a microarray study showed that it was one of the most strongly induced genes following hydrogen peroxide treatment (Zheng et al., 2001). To determine whether iraD was involved in a hydrogen peroxide stress response leading to σS stabilization, the half-life of σS was measured in wild-type and yjiD mutant strains. σS was fully stabilized 15 min after addition of hydrogen peroxide (1 mM final concentration) (Fig. 5). Furthermore, hydrogen peroxide treatment also affected the intracellular amount of σS (Fig. 5, cf. the time 0 min for each set of samples), presumably at least partially as a consequence of the decrease in σS degradation. σS levels were consistently higher in stressed bacteria than in an untreated culture. Therefore, as for many other stresses, oxidative stress leads to stabilization of σS.
We had irreproducible results in measuring σS turnover after hydrogen peroxide treatment in cells lacking iraD, possibly reflecting either redundancy in anti-adaptors for DNA damage, or too much damage to σS to allow its degradation even in the absence of IraD. We have renamed YjiD IraD, as an anti-adaptor for DNA damage, based on its induction after hydrogen peroxide treatment and on genetic experiments by H. Merrikh and S. Lovett (pers. comm.; see Discussion).
IraD and IraM regulate σS degradation by RssB phosphorylation-independent mechanisms
Given that IraD and IraM interfere directly within the RssB-mediated σS degradation pathway, it is theoretically possible that they act by changing the phosphorylation status of RssB. To determine this, σS half-life was measured in cells carrying rssBD58P, a chromosomal rssB allele unable to be phosphorylated but capable of stimulating degradation of σS (Zhou and Gottesman, 2006; Becker et al., 2000; Mika and Hengge, 2005; Bougdour et al., 2006). In exponentially growing cells carrying the rssBD58P allele and overexpressing IraM or IraD, σS was highly expressed and was stable for > 30 min, compared with the ∼2 min half-life in cells containing the vector control (Fig. 2D). Thus, the rssBD58P mutation did not abolish the ability of IraD or IraM to inhibit σS proteolysis. These data indicate that both IraD and IraM downregulate σS degradation by a mechanism that does not require the potential to phosphorylate or dephosphorylate the asp58 residue of RssB.
- Top of page
- Experimental procedures
- Supporting Information
Regulation of RssB activity by multiple anti-adaptor proteins
The discovery of IraP and its role as an anti-adaptor for stabilization of σS after phosphate starvation provided a new paradigm for understanding how environmental signals modulate the degradation and therefore the accumulation of the stress sigma factor σS. IraP acts by interacting with RssB, blocking delivery of σS to ClpXP. In this study, we asked if IraP was unique and found that it was not. We identified and characterized two new anti-adaptors, IraD and IraM. Overexpression of either stabilizes σS, and each can act in a purified in vitro system (Figs 2 and 3). In addition, we identified a transcriptional regulator, AppY, that is likely to regulate synthesis of yet another potential anti-adaptor; overproduction of AppY stabilizes σS independently of IraP, IraD and IraM. IraM is required for stabilization of σS during Mg2+ starvation (Fig. 4), and IraD is required for cell survival after DNA damage, most likely through its role in stabilizing σS (H. Merrikh and S. Lovett, pers. comm.). The identification of IraD and IraM as direct regulators of σS proteolysis demonstrates that multiple pathways leading to σS stabilization exist, each involving a specific anti-adaptor protein inhibiting RssB activity in response to a specific stress signal (Fig. 6).
The genes iraD (yjiD), iraM (ycgW) and appY have been identified previously in a number of other genetic screens, always along with σS or known regulators of σS. In a screen for multicopy plasmids that led to a mutator phenotype, rpoS, appY, iraD and iraM were found (Yang et al., 2004). Based on our findings, we suggest that the mutator effects of appY, iraD and iraM overexpression are via increased accumulation of RpoS, previously shown to lead to higher levels of frameshift mutations. A number of studies aimed at increasing production of the antioxidant lycopene in E. coli also led to identification of genes affecting the RpoS regulon, including identification of some of the anti-adaptors. In fact, iraM was named elbA after it was isolated as a multicopy suppressor of a defect in lycopene synthesis (Hemmi et al., 1998); we suggest that it be renamed to more properly reflect its function. In one study, screening of a multicopy library for increased lycopene synthesis identified appY, rpoS and crl, encoding a σS regulator (Pratt and Silhavy, 1998; Bougdour et al., 2004; Kang et al., 2005); in another, appY, rpoS, iraD (yjiD) and iraM (ycgW) were all found, and the possible connection to the mutator study and a role of RpoS was noted (Jin and Stephanopoulos, 2007). In a study in which strains carrying transposon insertions were screened, an insertion within rssB and another insertion upstream of iraD (yjiD), subsequently found to increase yjiD transcription, were found (Alper et al., 2005; Jin and Stephanopoulos, 2007). Our data strongly suggest that these genes and mutations are all acting in the σS pathway, and that increasing σS leads to both a higher mutation rate and enhanced lycopene biosynthesis; possibly other genes identified in these searches affect σS synthesis or stability as well.
Regulation of σS turnover in response to Mg2+ and phosphate starvation in E. coli and Salmonella
Salmonella typhimurium and E. coli respond to phosphate and Mg2+ starvation by stabilizing σS (Fig. 4; Bougdour et al., 2006; Tu et al., 2006). In both organisms, IraP is responsible for stabilization of σS after phosphate starvation; in E. coli, the promoter of iraP was shown to be positively regulated by ppGpp after phosphate starvation (Bougdour and Gottesman, 2007). S. typhimurium also uses IraP for stabilization of σS after Mg2+starvation, whereas in E. coli stabilization is IraP independent (Tu et al., 2006). Consistent with this, S. typhimurium but not E. coli has a second set of upstream promoters for iraP that respond to PhoP/PhoQ. However, the very low levels of σS detected in phoP mutant cells suggested that the PhoP/PhoQ system was involved in the regulation of σS during Mg2+starvation (Tu et al., 2006). Indeed, we show here that E. coli uses IraM to stabilize σS in Mg2+-starved cells; the induction of iraM transcription during Mg2+ starvation requires PhoP (Fig. 4).
RssC, which shares 40% identity with IraM, is capable of stabilizing σS when overexpressed (Fig. 1, data not shown). However, the promoter of rssC diverges significantly from that for iraM, and previous work has shown that σS stabilization in response to Mg2+ starvation in Salmonella is dependent upon IraP. Therefore, it seems likely that RssC responds to a thus-far unidentified stress signal in Salmonella.
Interestingly, both RssC and IraM have some homology with the PmrD protein of S. enterica (overall identity of 20–25%, Fig. S2). PmrD is a small connector protein which links the signal activating the PhoP/PhoQ regulatory system with expression of genes under the control of the PmrA/PmrB two-component regulatory system that senses extracellular concentrations of Fe3+ or Al3+ (Kox et al., 2000; Kato and Groisman, 2004). PmrD transcription is activated by PhoP in the presence of limited amount of Mg2+ and activates the response regulator PmrA through a direct protein–protein interaction with the phosphorylated form of PmrA (Kato and Groisman, 2004). While the exact mechanism of action of IraM remains to be determined, IraM plays the role of a connector protein between the PhoP/PhoQ and the RssB/RpoS regulatory systems. Although there are clearly differences between the mode of action of PmrD and IraM (PmrD affects phosphorylation of its target, while IraM acts independently of phosphorylation), the sequence similarity may suggest the existence of yet other proteins that act to modulate the function of response regulators.
σS and oxidative stress
IraD (previously YjiD) stabilizes σS both in vivo and in vitro (Fig. 2A and 3A). The direct interaction of IraD with RssB but not with σS strongly suggests that IraD interferes with σS degradation by sequestering RssB, thereby preventing it from interacting with σS (Fig. 6). The in vivo analysis of IraD function derives from two sources. The gene is highly induced upon treatment of cells with hydrogen peroxide, independent of OxyR (Zheng et al., 2001). In an independent study, mutations in iraD were identified in a screen for hypersensitive mutants to DNA damaging agents such as AZT and phleomycin (H. Merrikh and S. Lovett, pers. comm.). The hypersensitivity is apparently due to degradation of σS, as predicted from our results. Thus, deletion of rssB rescues iraD mutant cells treated with either hydrogen peroxide or AZT. In agreement with this, after hydrogen peroxide treatment, σS became fully stable and accumulated to higher levels and, in an iraD mutant, σS intracellular levels remained low (data not shown; H. Merrikh and S. Lovett, pers. comm.). Still unidentified is the signal transduction pathway for expressing IraD after DNA damage. Possibly these agents are causing multiple forms of cell damage, each leading to σS stabilization, complicating the picture when iraD alone is examined. For instance, Fredriksson et al. suggested a passive mechanism for σS stabilization by oxidized proteins that titrate the activity of the ClpP protease (Fredriksson et al., 2007).
AppY and σS
AppY, a transcriptional regulator, causes σS stabilization when overexpressed, and the effect is seen even in cells deleted for iraP, iraD and iraM (Fig. 2C). Therefore, it seems likely that AppY regulates an unidentified regulator, possibly an additional anti-adaptor, for σS turnover. appY is encoded within the DLP12 prophage (Lindsey et al., 1989), and is not found in organisms beyond E. coli. It has been identified as the transcriptional regulator for the appCBA (or cyxAB-appA) operon, encoding a cytochrome and acid phosphatase and a hydrogenase operon expressed anaerobically (Atlung et al., 1989; Atlung and Bronsted, 1994; Brondsted and Atlung, 1994). The appY promoter is expressed in stationary phase, under anaerobic growth, and after phosphate starvation, and is negatively regulated by HNS (Atlung et al., 1996; Brondsted and Atlung, 1996). However, cells mutant in appY did not affect σS stabilization in stationary phase or under low pH stress (data not shown), and the stabilization of σS in an hns mutant (Zhou and Gottesman, 2006) was not blocked by an appY mutation (data not shown). Anaerobic conditions have not been tested. Examination of the full set of genes regulated by AppY may lead to the identification of the proposed regulatory protein.
How many anti-adaptor proteins exist to control σS stability?
IraD and IraM (or RssC) are found in Enterobacteria that also contain σS and RssB, as is IraP. While IraD, IraM, and IraP all act directly to block RssB-mediated degradation of σS by ClpXP (Fig. 3A), they do not share any sequence similarity. It is therefore not currently possible to predict other potential anti-adaptor proteins based on sequence analysis alone. However, the characteristics of IraD, IraM and IraP suggest that there is a specific anti-adaptor protein for each stress condition that requires the stabilization of σS, and at least a few of these conditions do not yet have an identified regulatory mechanism. In addition, new conditions that lead to stabilization continue to be identified. It seems likely that yet other anti-adaptors will be found that mediate these stress responses. In addition, the finding that these small proteins modulate the activity of a response regulator in a way that is independent of the phosphorylation signals suggests the possibility that response regulators other than RssB may also be subject to regulation by small proteins, independent of the phosphorylation status.
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- Experimental procedures
- Supporting Information
Bacterial strains and plasmids
The bacterial strains and plasmids used in this study are listed in Table 3. The wild-type S. enterica typhimurium strain LT2 from our collection has been shown to be an mviA mutant and defective in σS turnover (Cunning and Elliott, 1999). For most experiments, we used a closely related mviA+ derivative of S. typhimurium LT2. The latter strain was obtained from W. Benjamin (WB335 in Table 3; (Benjamin et al., 1991). For convenience we designate this strain LT2A.
|Strain or plasmid||Genotype||Reference or source|
|MG1655||Wild-type E. coli|
|DJ480||MG1655 Δlac X74||D. Jin (NCI)|
|NM1100||MG1655 mini-λ-Red-tet||N. Majdalani (NCI)|
|NM1200||MG1655 mini-λ-Red-cm||N. Majdalani (NCI)|
|YN868||DJ480 rssBD58P||Zhou and Gottesman (2006|
|EG12976||MG1655 ΔphoP::kn||E. Groisman (Washington University)|
|AB006||MG1655 ΔiraP::kn||Bougdour et al. (2006)|
|AB007||CRB316 ΔiraP::cm||Bougdour et al. (2006)|
|AB012||MG1655 rssB::Tn10||Bougdour et al. (2006)|
|CRB316||MG1655 ΔlacI–lacZ::PBAD–rpoS990′–′lacZ||Ranquet and Gottesman (2007)|
|AB038||NM1100 ΔiraP::cat-sacB, mini-λ-Red-tet||Bougdour and Gottesman (2007)|
|AB041||MG1655 ΔiraP::cm||This studya|
|AB042||DJ480 PiraM′–lacZ||This study|
|AB043||DJ480 PiraM′–lacZ ΔphoP::kn||This studya|
|AB044||MG1655 ΔappY::kn||This studya|
|AB045||MG1655 ΔiraM::kn||This studya|
|AB046||MG1655 ΔiraD::tetA||This studya|
|AB047||MG1655 ΔiraP||This study|
|AB048||MG1655 ΔiraPΔiraM::kn||This studya|
|AB049||MG1655 ΔiraPΔiraM::knΔiraD::tetA||This studya|
|PJB001||DJ480, rpoS750′–′lacZ, rssA2::cm||This studya|
|PJB002||PJB001 ΔiraD::kn||This studya|
|PJB003||PJB001 ΔiraM::kn||This studya|
|S. enterica serovar Typhimurium|
|LT2A||Wild-type S. enterica (WB335)||Benjamin et al. (1991)|
|TE6756||LT2A putPA1303::kn-katE′–lac||Cunning and Elliott (1999)|
|TE7119||LT2A putPA1303::kn-proV′–lac||Cunning and Elliott (1999)|
|TE7179||LT2A orf o186::MudJ||Cunning and Elliott (1999)|
|TE7180||LT2A otsA::MudJ||Cunning and Elliott (1999)|
|pHDB3||AmpR||Ulbrandt et al. (1997)|
|pRS1553||AmpR||Simons et al. (1987)|
|pBR328||AmpR||Hmiel et al. (1986)|
All the E. coli strains used are derivatives of MG1655, made by P1 transduction as indicated, selecting the appropriate antibiotic-resistant marker. To construct the deletion/insertion mutants of iraM, iraD and appY, the λ Red recombination system was used as described (Yu et al., 2000). Briefly, PCR fragments were obtained by amplifying either the kanamycin or tetracycline (tetA) resistance cassettes of the strains MC4100, ybeW::GBKn cassette (J.M. Ghigo, Institut Pasteur, Paris, France), and AB012 (rssB::Tn10) respectively. The antibiotic resistance cassettes were amplified by PCR using primers ycgWHRL-Kn F (5′-taatgaaccatattaaataccgtgggataagacataacaaAAAGCCACGTTGTGTCTCAA-3′) and ycgWHRR-Kn R (5′-tattattaatagcatatcgagcatatttatatgaagcccaTTAGAAAAACTCATCGAGCA-3′), with primers appYHRL-Kn F (5′-gactactatcaacttgttttaattttatgataggtgcaagAAAGCCACGTTGTGTCTCAA-3′) and appYHRR-Kn R (5′-atattataattaacatgtagacaacttgtaataaacattaTTAGAAAAACTCATCGAGCA-3′) for amplification of the kn cassette; with primers yjiDHRL-Tet F (5′-ggcaagccgcatatgttgctaaacgtcaggagtgcgcaaaCTAGACATCATTAATTCCTA-3′) and yjiDHRR-Tet R (5′-gaacgcttctggcgtgaactgcgcgagctggagattgtgcGAAGCTAAATCTTCTTTATCG-3′) for amplification of the tetA gene. The capital letters of the primers above are identical to the ends of the respective antibiotic resistance cassettes. The rest of the primers are homologous to the regions flanking the coding sequences of iraM, appY and iraD respectively. The resulting PCR products were recombined into the chromosome of strain NM1100 for the constructions with the kn cassettes and with the strain NM1200 for constructions with the tet cassettes; NM1100 and NM1200 (N. Majdalani, NIH, NCI) are MG1655 derivatives carrying the mini-λ prophage encoding the λ Red functions (Court et al., 2003). The transformed cells were selected on LB plates containing either 50 μg ml−1 kanamycin or 25 μg ml−1 tetracycline at 32°C. Recombinant products were verified by PCR and the mutations were transferred into MG1655 by P1 transduction.
To construct the strain AB047 containing a clean deletion of the iraP ORF (ΔiraP unmarked), the λ Red system was used as described (Yu et al., 2000). The strain AB038 (ΔiraP::cat-sacB) (Bougdour and Gottesman, 2007) was transformed with the oligonucleotide Delta yaiB lagging (5′- ccgtgacaactttatgacagctgttgaaaagattaactttgtctgtatttccttatccaaagtatggagaagttaacatt-3′), encompassing the flanking regions of iraP ORF, to replace the cat-sacB cassette by counterselecting for sucrose resistance as previously described (Majdalani et al., 2005).
To construct the PiraM′–lacZ transcriptional fusion, a fragment encompassing the promoter region of the gene iraM from −700 to +287 bp relative to the ATG start codon of iraM ORF was amplified by PCR using primers EcoRI-ycgW PReg.F1 (5′-atcggaattcCTAAAGTTGGATACTTAAGAAATGCTT-3′) and BamHI-ycgW R (5′-tgacggatcCTTGAGCCCATATGGGCATATTTTTATAATGC-3′) and genomic DNA of strain MG1655 as template. The capital letters are homologus to the promoter region and the 3′ end of iraM. The resulting PCR product was digested and cloned into the EcoRI and BamHI sites of pRS1553 (Simons et al., 1987) to yield the pRS1553::PiraM plasmid. The strain AB042 was constructed by crossing the λRS468 phage with the pRS1553::PiraM plasmid to isolate λ lac transducing phages; these phages were used to lysogenize DJ480, a Δlac derivative of MG1655, obtained from D. Jin, NCI. Transcriptional fusions were delivered in single copy to the λatt site as described previously (Powell et al., 1994). The same PCR fragment was cloned into plasmid pRS1551 in an attempt to construct a PiraM′–′lacZ translational fusion; for unknown reasons, no blue plaques could be isolated.
Plasmids pQE-IraD, pQE-IraM and pQE-AppY were constructed by amplifying the coding sequences of iraD, iraM and appY by PCR using the primers EcoRI-rbs-yjiD F (5′-atcgGAATTCATTAAAGAGGAGAAATTAACTatgatgcgacaatcacttcaggctg-3′); PstI-yjiD R (5′-atcgCTGCAGttagctgacattctccagcgtcgcac-3′) for iraD, EcoRI-rbs-ycgW F (5′-atcgGAATTCATTAAAGAGGAGAAATTAACTatgaagtggatagtaattgacacggtaat-3′) and PstI-ycgW R (5′-atcgCTGCAGttacttgagcccatatgggcat-3′) for iraM; EcoRI-rbs-appY F (5′-atcgGAATTCATTAAAGAGGAGAAATTAACTatggattatgtttgctccgtagttttcat-3′) and PstI-appY R (5′-atcgCTGCAGTCAGTCAATTGTTTTGTTTATTCCATC-3′) for appY. The capital letters of these primers correspond to the pQE80L vector sequence. These primers amplify the entire ORF iraD, iraM and appY, adding the optimal ribosome-binding site (bold) of the expression vector pQE-80L (Qiagen) and two restriction sites, EcoRI and PstI. The start codons of IraD, IraM and AppY are underlined. After digestion with EcoRI and PstI, the PCR products were introduced into the EcoRI and PstI sites of pQE-80L (Qiagen). The resulting plasmids, called pQE-IraD, pQE-IraM and pQE-AppY respectively, contain a T5 promoter under the control of LacI (Qiagen) driving the transcription of the downstream sequences. By digesting the pQE-80L plasmid with EcoRI and PstI, the His6 tag was lost; therefore untagged proteins are expressed from these plasmids.
To construct the plasmids pQE-IraD-His6 and pQE-His6-IraM, the coding sequences of the genes iraD and iraM were cloned in a His6-tag pQE-80L, as follows. The coding sequences of iraD and iraM were PCR-amplified using chromosomal DNA from MG1655 as template and the primers EcoRI-rbs-yjiD F (see above) and PstI-3*-6HisSGRG-yjiD R (5′-ATAACTGCAGTCAGCTAATTAAGCTTAGTGATGGTGATGGTGATGCGATCCTCTTCCgctgacattctccagcgtcgcactgcg-3′) for IraD-His6, and primers BamHI-ycgW F (5′-TATCGGATCCaagtggatagtaattgacacggtaat-3′) and PstI-ycgW (5′-ATCGCTGCAGttacttgagcccatatgggcat-3′) for His6-IraM. The sequences corresponding to iraD or iraM are indicated in lowercase letters and the sequence coding for GRGSHis6 added directly to the C-terminal end of IraD is underlined. For the His6-IraM construction, the GRGSHis6-tag sequence carried by the pQE-80L vector was used to fuse the GRGSHis6-tag to the N-terminal end of IraM. The PCR fragments were digested by EcoRI and PstI for IraD-His6, and BamHI and PstI for His6-IraM and cloned into pQE-80L cut by the same enzymes respectively, yielding the pQE-IraD-His6 and pQE-His6-IraM vectors. Constructs in which the tag was put on the N-terminus of IraD failed to complement in a plate assay.
All plasmid sequences were checked by sequencing.
Media and growth conditions
Cells were grown in Luria–Bertani (LB) broth or morpholinepropanesulfonic acid (MOPS) minimal medium (Teknova) supplemented with 0.4% glucose, 0.2% (NH4)2SO4, 1.32 mM K2HPO4 and 1 μg ml−1 thiamine, or M9 minimal medium containing 2 mM MgSO4 and 100 μM CaCl2 and supplemented with 10 μM FeSO4. MacConkey agar plates supplemented with 1% lactose and 50 μg ml−1 ampicillin were used in analyses of strains carrying the PBAD–rpoS990′–′lacZ chromosomal fusion. All liquid cultures were grown under aerobic conditions (250 r.p.m) 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 (∼9 × 107 cfu ml−1) and ∼3 (∼9 × 108 cfu ml−1) respectively.
For starvation experiments, MOPS was made without glucose for carbon starvation and M9 medium was made without MgSO4 and CaCl2 for Mg2+/Ca2+ starvation. Cells were first grown overnight in complete MOPS or M9 minimal media, then subcultured into fresh complete media 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, re-suspended in the same volume of starvation medium, and incubation at 37°C was continued as indicated.
Library screening in E. coli. 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 AB007, plated on MacConkey lactose plates containing 50 μg ml−1 ampicillin to give about 500 colonies per plate, and incubated at 37°C. About 30 000 colonies were screened and 17 red (Lac+) colonies isolated (Table 1). 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′). Six plasmids contained araC and probably affect the expression of the PBAD promoter driving rpoS–lacZ, as shown by the lack of an effect on an rpoS750′–′lacZ translational fusion, which encodes a hybrid protein that is subject to the same transcriptional, translational and proteolytic regulation as σS itself and therefore is independent of PBAD.
Library screening in S. typhimurium. The pBR328-based plasmid DNA library (Hmiel et al., 1986) was introduced into S. typhimurium strain TE6756 carrying a katE′–lac transcriptional fusion (Brown and Elliott, 1996) by electroporation, plated on MacConkey lactose plates containing 100 μg ml−1 ampicillin and incubated at 37°C. About 10 000 colonies were screened and a single colony with increased β-galactosidase activity was isolated; the single confirmed plasmid that led to higher expression of the fusion was tested further. Primers Bamcw (5′-CACTATCGACTACGCGATCA-3′), Bam ccw (5′-GATATAGGCGCCAGCAACCGCAC-3′), and Sph ccw (5′-CAGTAGTAGGTTGAG-3′) were used to determine the sequence of the insert.
Assay for σS degradation in vivo
Cells were grown in LB, or in MOPS, or in M9 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−1) 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, MOPS or M9 media were assayed for OD600, and 1 ml samples were removed and precipitated with a final concentration of 5% ice-cold tricarboxylic acid (TCA). Precipitated pellets were washed with 500 μl of 80% cold acetone and then re-suspended in a volume of SDS sample buffer normalized to the OD600.
Samples were analysed using NuPAGE 10% or 12% Bis-Tris gels (Invitrogen), transferred and probed with a 1:4000 dilution of anti-σS antiserum or 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).
Overproduction and purification of IraD-His6 and His6-IraM were performed as follows: The MG1655 strain containing a ColE1-compatible pACYC-based plasmid (CmR) carrying extra copies of the argU, ileY and leuW tRNA genes (from BL21-CodonPlus-RIL strain, Stratagene), was transformed with pQE-IraD-His6 or pQE-His6-IraM. Fresh colonies from an LB plate supplemented with 100 μg ml−1 Amp and 25 μg ml−1 Cm were used to inoculate 500 ml of super broth medium supplemented with 100 μg ml−1 carbenecillin and 40 μg ml−1 Cm. Cells were grown at 25°C to an OD600 of ∼1. The temperature was cooled to 15°C and IPTG was added to 200 μM final concentration. After 16–18 h of induction, cells were harvested and stored at −80°C until use. Cell pellets containing IraD-His6 recombinant proteins were re-suspended in lysis buffer (20 mM Tris-HCl, pH 7.9, 500 mM NaCl, 10% glycerol (v/v), 10 mM imidazole, 1 mM PMSF) and cell pellets containing His6-IraM recombinant proteins were re-suspended in lysis buffer supplemented with 1% (v/v) Triton X-100. Bacteria were lysed using a French pressure cell and the lysates were centrifuged at 10 000 g for 30 min at 4°C. The soluble fractions were applied to a Ni2+-NTA column (Invitrogen) equilibrated with lysis buffer. The columns were washed successively with 20, 40, 60, 80 and 100 mM imidazole. Proteins were eluted with washing buffer containing 250 mM imidazole. The His-tagged proteins, eluted with 250 mM imidazole, were dialysed against a buffer composed of 50 mM Tris-HCl (pH 7.5), 0.1 mM EDTA, 150 mM KCl, 0.1 mM DTT and 10% glycerol (v/v), and the samples were stored at −80°C. Final preparations of IraD-His6 and His6-IraM were >90% and ∼70% pure, respectively, as judged by Coomassie staining following SDS-PAGE. Note that no IraD and less IraM were seen in the absence of the tRNA plasmid; both genes contain rare codons.
σS (Bougdour et al., 2006), RssB (Bougdour et al., 2006), ClpX (Grimaud et al., 1998), ClpP (Maurizi et al., 1994) and RepA(Δ25)-SsrA (Sharma et al., 2005) were isolated as described. 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, σS monomers, IraD-His6 monomers and His6-IraM monomers.
Assay for σS degradation in vitro
In vitroσS degradation assays were performed as described previously (Bougdour et al., 2006). Briefly, 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, σS (20 pmol), RepA(Δ25)-SsrA 20 pmol, IraD (6–60 pmol), IraM (6–60 pmol), RssB (2 pmol), ClpX6 (2 pmol) and ClpP14 (2 pmol), unless otherwise indicated. All of the proteins were diluted into the reaction buffer containing 0.05% (v/v) 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 analysed by SDS-PAGE. SeeBlue Plus2 (Invitrogen) prestained protein standards were used for molecular weight estimation.
Salmonella cells were grown as described in the text, centrifuged and re-suspended in Z-buffer (100 mM NaPO4, pH 7.0, 10 mM KCl, 1 mM MgSO4). They were permeabilized by treatment with SDS and chloroform (Miller, 1972), and assays were performed in Z-buffer containing 50 mM β-mercaptoethanol by a kinetic method using a plate reader (Molecular Dynamics). Activities (ΔOD420 per minute) are normalized to actual cell density (OD650) and were always compared with appropriate controls assayed at the same time. Each set of results shown is from a single experiment; each experiment was repeated several times with similar results (< 15% variation).
Escherichia coli cells were grown overnight in M9 minimal medium containing 2 mM MgSO4 and 100 μM CaCl2 (complete M9 medium), then subcultured into fresh complete medium at an OD600 of 0.01 (∼1:400 dilution) and grown to mid-logarithmic phase (OD600 of ∼0.3) for the following treatments. For starvation experiments (without Mg2+/Ca2+), cells were washed twice by filtration with prewarmed (37°C) starvation medium, re-suspended in the same volume of starvation medium, and incubation at 37°C was continued as indicated. As a control, a sample of the culture was kept in complete M9 medium and grown as indicated (with Mg2+/Ca2+). Kinetic assays for β-galactosidase activity were performed using a SpectraMax 250 microtitre plate reader (Molecular Devices) as described previously (Majdalani et al., 1998) with the following modification. The solubilization buffer was supplemented with 20 mM MgSO4 to saturate the reaction solution with Mg2+. Specific activities were calculated using the formula Vmax/OD600. The results reported represent data typical of at least three experimental trials.
- Top of page
- Experimental procedures
- Supporting Information
We thank Houra Merrikh and Sue Lovett for sharing their results on yjiD prior to publication, N. Majdalani, C. Ranquet, and Y. Zhou for providing essential strains, E. Groisman for providing the phoP mutant, and S. Wickner for purified ClpXP, RpoS, RssB, RepA-SsrA as well as anti-σS antibody. This research was supported in part by the Intramural Research Program of the NIH, National Cancer Institute, Center for Cancer Research. A.B. was also supported by the Philippe Foundation.
- Top of page
- Experimental procedures
- Supporting Information
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