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Genes Dev. Mar 1, 2001; 15(5): 627–637.
PMCID: PMC312640

The RssB response regulator directly targets σS for degradation by ClpXP

Abstract

The σS subunit of Escherichia coli RNA polymerase regulates the expression of stationary phase and stress response genes. Control over σS activity is exercised in part by regulated degradation of σS. In vivo, degradation requires the ClpXP protease together with RssB, a protein homologous to response regulator proteins. Using purified components, we reconstructed the degradation of σS in vitro and demonstrate a direct role for RssB in delivering σS to ClpXP. RssB greatly stimulates σS degradation by ClpXP. Acetyl phosphate, which phosphorylates RssB, is required. RssB participates in multiple rounds of σS degradation, demonstrating its catalytic role. RssB promotes σS degradation specifically; it does not affect degradation of other ClpXP substrates or other proteins not normally degraded by ClpXP. σS and RssB form a stable complex in the presence of acetyl phosphate, and together they form a ternary complex with ClpX that is stabilized by ATP[γ-S]. Alone, neither σS nor RssB binds ClpX with high affinity. When ClpP is present, a larger σS–RssB–ClpXP complex forms. The complex degrades σS and releases RssB from ClpXP in an ATP-dependent reaction. Our results illuminate an important mechanism for regulated protein turnover in which a unique targeting protein, whose own activity is regulated through specific signaling pathways, catalyzes the delivery of a specific substrate to a specific protease.

Keywords: ATP-dependent proteolysis, molecular chaperones, ClpA, RpoS, sigma S, acetyl phosphate

All organisms use rapid degradation of specific proteins as one way to tightly control important regulatory factors and developmental switches. Regulation of biological functions by proteolysis requires accurate substrate recognition at a precise time. In eukaryotes, fidelity of protein degradation depends on ubiquitin-tagging, a process that occurs in multicomponent complexes involving regulatory factors and various ubiquitin pathway enzymes. The ubiquitinated proteins are then targeted to the proteasome for degradation (for review, see Voges et al. 1999). In Escherichia coli, an example of substrate tagging as a means for regulating proteolysis is the degradation of incomplete proteins made from truncated mRNAs. In this case, short peptides coded by the SsrA RNA are added cotranslationally to polypeptides that have become stalled on ribosomes, and the tagged proteins are then degraded (Keiler et al. 1996; Gottesman et al. 1998; Roche and Sauer 1999). In general, E. coli proteins regulated by degradation interact directly with the proteases themselves, with each of the five known E. coli ATP-dependent proteases degrading a different but somewhat overlapping set of substrates (Gottesman 1996; Wickner et al. 1999). For many of the highly unstable E. coli proteins, degradation is rapid under all conditions, and synthesis is tightly controlled.

However, the activity of some proteins in E. coli is controlled by regulated degradation. For example, σ32 (RpoH), the heat shock sigma factor, is rapidly degraded under normal growth conditions by the AAA protease, FtsH, in a reaction modulated by the DnaJ/DnaK/GrpE chaperone system. During heat shock, σ32 is transiently stabilized, and this stabilization results in the rapid increase in the synthesis of the heat shock proteins (Yura and Nakahigashi 1999).

The stationary phase sigma factor, σS (RpoS), is another example of a protein whose activity is controlled by regulated proteolysis. σS promotes expression of more than 50 genes involved in responses to many stresses, including starvation, osmotic stress, acid shock, cold shock, heat shock, and oxidative damage, as well as the transition to stationary phase (Loewen and Hengge-Aronis 1994; Hengge-Aronis 2000). Although σS is present at very low levels during exponential cell growth, owing largely to its rapid degradation (half-life of ~2 min), its stability increases ~10-fold following transition to stationary phase or other stress treatments. Regulated degradation plays a major role in determining the amount of σS in the cell, but σS accumulation is also regulated at the transcriptional and translational levels (Lange and Hengge-Aronis 1994). The protease responsible for σS turnover in exponentially growing cells is ClpXP (Schweder et al. 1996), an ATP-dependent protease consisting of a regulatory component, ClpX, and a proteolytic component, ClpP (Gottesman et al. 1993; Wojtkowiak et al. 1993).

Degradation of σS requires an additional protein, RssB (Regulator of Sigma S; also referred to as SprE in E. coli, MviA in Salmonella, and ExpM in Erwinia) that is homologous to response regulator proteins (Bearson et al. 1996; Muffler et al. 1996b; Pratt and Silhavy 1996; Andersson et al. 1999). Genetic evidence shows that RssB is required for σS degradation but not for another ClpXP substrate, λ O, which indicates that RssB specifically targets σS for degradation (Zhou and Gottesman 1998). One of the hallmarks of the response regulator component of two-component signal transduction systems in prokaryotes is the presence of a conserved aspartate that is phosphorylated by the cognate sensor component. The N-terminal domain of RssB contains this conserved aspartate, although a cognate sensor protein has not been identified. RssB, like many other response regulators, is phosphorylated by acetyl phosphate in vitro (Bouche et al. 1998). In addition, phosphorylated RssB forms a stable complex with σS in vitro (Becker et al. 1999).

In this report we investigate RssB-regulated degradation of σS by reconstituting the pathway of σS degradation in vitro with purified RssB and ClpXP. We discovered that RssB acts directly and catalytically in stimulating degradation of σS by ClpXP in a reaction requiring acetyl phosphate and ATP. We have isolated and characterized subassemblies of the degradation machinery including σS–RssB, σS–RssB–ClpX, and σS–RssB–ClpXP complexes, and suggest a probable pathway for the degradation of σS.

Results

RssB promotes σS degradation by ClpXP in vitro

Although genetic evidence pointed to a central role for RssB in σS degradation, it was not clear how RssB modulated σS degradation and whether or not it acted directly. To study the mechanism of RssB regulation of σS degradation by ClpXP, we first developed an in vitro assay to measure σS degradation. Purified ClpXP alone degraded σS poorly, as measured by acid solubilization of radioactively labeled σS (Fig. (Fig.1A).1A). Degradation was greatly stimulated by the addition of a crude extract prepared from cells overproducing RssB from a multicopy pACYC plasmid (Fig. (Fig.1A).1A). A similar amount of crude extract of cells carrying the vector alone did not stimulate degradation significantly, very likely because the chromosomal copy of rssB provided only a small amount of RssB. These observations indicate that RssB is involved in σS degradation in vitro.

Figure 1
σS degradation by ClpXP and RssB in vitro. (A) Stimulation of ClpXP-dependent σS degradation by extracts of cells overproducing RssB. Degradation of [3H]σS by ClpXP was measured as described in Materials and Methods ...

We purified the protein that promoted degradation of σS by ClpXP from an extract of cells expressing RssB from a multicopy plasmid and later from cells producing RssB from a T7 expression system. The identification of RssB as the stimulatory factor was confirmed by N-terminal amino acid sequencing, and the final preparations were >90% pure as judged by SDS-PAGE. Purified RssB stimulated σS degradation by ClpXP more than 10-fold (Fig. (Fig.1B).1B). ClpX, ClpP, and ATP were essential for σS degradation (Fig. (Fig.1B).1B). Thus, as seen in vivo (Muffler et al. 1996a; Pratt and Silhavy 1996), σS degradation by ClpXP in vitro is dependent upon RssB. Importantly, these results show that the role of RssB in σS degradation is direct.

Because acetyl phosphate is a phosphate donor for the phosphorylation of several response regulators in vitro, including RssB (Bouche et al. 1998), it seemed likely that it might be required for RssB activity and was therefore included in the reaction mixtures. When acetyl phosphate was omitted, σS degradation decreased 10-fold (Fig. (Fig.1B).1B). Half-maximal activation of degradation occurred with ~7 mM acetyl phosphate. Other potential phosphate donors, including carbamyl phosphate and phosphoenol pyruvate, were ineffective in stimulating σS degradation. The observation that RssB is phosphorylated in vitro by acetyl phosphate (Bouche et al. 1998), in combination with our finding that σS proteolysis is stimulated by acetyl phosphate, indicates that the phosphorylated form of RssB is required to activate degradation.

We next addressed the question of whether RssB acts stoichiometrically or catalytically in the degradation reaction. Degradation of σS was linear with time in reaction mixtures containing a 60-fold molar excess of σS relative to RssB, ClpX, and ClpP (Fig. (Fig.2A).2A). After 40 min, 25 pmoles of σS had been degraded per pmole of RssB added (Fig. (Fig.2A).2A). Thus, RssB functions catalytically in σS degradation. By measuring the rate of degradation of σS by ClpXP in the presence and absence of RssB at various concentrations of σS, we found that RssB increased the apparent affinity for σS by >10-fold and increased the rate of the reaction by at least 6-fold (Fig. (Fig.2B).2B). Therefore, RssB facilitates the interaction of σS with ClpXP.

Figure 2
RssB functions catalytically and specifically. (A) RssB acts catalytically. [3H]σS degradation was measured as a function of time as described in Materials and Methods in reaction mixtures with 2 pmoles of RssB, 120 pmoles of [ ...

RssB is specific for σS and for ClpXP

In vivo, rssB null mutants do not degrade σS, but continue to degrade another ClpXP substrate, λ O protein (Gottesman et al. 1993; Wojtkowiak et al. 1993), which indicates that RssB acts on the specific substrate rather than on the protease (Zhou and Gottesman 1998). To test the substrate specificity of RssB in vitro, we examined the effect of RssB on λ O degradation by ClpXP. RssB stimulated σS degradation at all concentrations of ClpX; in contrast, RssB had no effect on λ O degradation by ClpXP (Fig. (Fig.2C,D).2C,D). Thus the in vitro results mimic the in vivo substrate specificity.

We asked whether RssB increased the promiscuity of ClpXP, allowing it to degrade proteins that are not normally recognized by ClpX. σ70, the vegetative sigma subunit of RNA polymerase, was tested as a potential substrate because the C-terminal half of σ70 is homologous to σS. ClpXP did not degrade σ70 in the presence or absence of RssB, using conditions in which σS degradation required ClpXP and RssB (Table (Table1).1). This result is consistent with the in vivo observation that σ70 is stable (Grossman et al. 1983). Similarly, RssB did not enable ClpXP to degrade α-casein or RepA, which are poor substrates for ClpXP but good substrates for another Clp protease, ClpAP (Table (Table1;1; Wojtkowiak et al. 1993; Wickner et al. 1994). We also tested whether RssB could stimulate degradation by ClpAP. There was no effect of RssB on the small but significant amount of σS degradation by ClpAP (Table (Table1).1). There was also no effect of RssB on the rate of degradation of α-casein or RepA by ClpAP (Table (Table1).1). These results indicate that RssB acts specifically with ClpX to promote efficient degradation of σS. Thus, in vitro as well as in vivo, RssB shows a high degree of specificity both for the substrate σS and for the protease to which σS is targeted, ClpXP.

Table 1
Substrate and protease specificity of RssB

Interaction of RssB and σS

To study the mechanism of action of RssB, we designed experiments to dissect the degradation pathway into partial reactions and to determine the role of acetyl phosphate and ATP at the various steps. Evidence for an interaction of σS and RssB was obtained by showing that σS protects RssB from inactivation by N-ethylmaleimide (NEM), a reagent that reacts with accessible sulfydryl groups. NEM treatment of RssB inhibited the ability of RssB to promote σS degradation by ClpXP in vitro by >10-fold (data not shown). When σS and RssB were incubated together, σS protected RssB from NEM inactivation, and protection required acetyl phosphate and Mg2+. These results suggested that phosphorylated RssB interacts with σS, and this intermediate is functional in the degradation pathway.

Interaction between RssB and σS was further assessed using gel filtration. σS, a 38-kD protein, alone eluted earlier than expected, at the position predicted for a globular protein of 48 kD. RssB, a 37-kD protein, alone eluted later than expected, at the position predicted for a globular protein of 25 kD (the positions where σS and RssB eluted when chromatographed separately are indicated by arrows at the top of Fig. Fig.3).3). The possibility that RssB interacts with the column matrix has not been ruled out. Acetyl phosphate did not affect the elution of either protein. When σS and RssB were incubated together in the absence of acetyl phosphate and analyzed by gel filtration, they eluted separately and at the same positions obtained when each was chromatographed separately (Fig. (Fig.3A).3A). Following incubation of σS and RssB with acetyl phosphate and Mg2+, the two proteins coeluted at a position indistinguishable from the position of σS alone (Fig. (Fig.3B).3B). Both acetyl phosphate and Mg2+ were required for the altered elution pattern. Because the elution profiles of σS and RssB alone were not as expected, we confirmed the interaction of σS and RssB by affinity chromatography using either His6-RssB or His6S (data not shown). The interaction was also detected by Far-Western blotting of His6-RssB associated with membrane-bound σS using RssB antibody (data not shown). Physical interaction between σS and tagged RssB was also shown by Hengge-Aronis and coworkers by affinity chromatography (Becker et al. 1999) and by Foster and coworkers using a two-hybrid system (Moreno et al. 2000). In light of independent demonstrations of a σS–RssB complex, the gel filtration experiments further indicate that the σS–RssB interaction results in significant conformational changes in σS.

Figure 3
Interaction of σS and RssB. (A) RssB (150 pmoles) and [3H]σS (320 pmoles) were incubated in the absence of acetyl phosphate and analyzed by gel filtration in the absence of acetyl phosphate as described in Materials and ...

To test whether or not the σS–RssB complex could be dissociated, we formed σS–RssB complexes in the presence of acetyl phosphate and Mg2+ and then added EDTA to chelate Mg2+, a metal ion essential for the interaction of σS and RssB. After 60 min, the mixture was analyzed by gel filtration. σS eluted free from RssB (Fig. (Fig.3C).3C). Taken together, these observations are consistent with a reversible acetyl phosphate-dependent physical interaction between RssB and σS.

σS, RssB, and ClpX form a ternary complex

Because ClpX is responsible for unfolding and delivering substrates to ClpP, we expected that σS alone or the σS–RssB complex interacts with ClpX. Using ultrafiltration membranes with a molecular weight cutoff of 100,000, it was observed that σS was retained on the membranes with ClpX when RssB was present (Fig. (Fig.4).4). σS was not retained when ClpX was omitted.

Figure 4
Binding of σS to ClpX promoted by RssB. Reaction mixtures containing 9 pmoles of [3H]σS, 60 pmoles of ClpX, and the indicated amounts of RssB were incubated with ATP[γ-S] and acetyl phosphate as ...

To distinguish whether RssB transferred σS to ClpX or whether σS and RssB formed a ternary complex with ClpX, mixtures of σS, ClpX, and RssB were incubated in the presence of acetyl phosphate, adenosine-5′-O-(3-thiotriphosphate) (ATP[γ-S]), and Mg2+ and analyzed by gel filtration. The three proteins eluted together in the excluded volume, showing that RssB assembles into a ternary complex with σS and ClpX (Fig. (Fig.5A).5A). In control experiments, ClpX, σS, and RssB alone eluted separately (at the positions indicated by arrows at the top of Fig. Fig.5).5). Complex formation required ATP[γ-S] and Mg2+. When acetyl phosphate was omitted, ~20% as much σS and RssB eluted with ClpX as when acetyl phosphate was present. A very small amount of RssB associated with ClpX in the absence of σS, and a small amount of σS associated with ClpX in the absence of RssB, indicating weak interactions between the individual proteins and ClpX (Fig. (Fig.5B,C).5B,C). RssB and σS were released from ClpX following the exchange of ATP[γ-S] with ATP by the addition of a 10-fold molar excess of ATP to ATP[γ-S] (data not shown). These data indicate that RssB is an integral part of the degradative complex and has a role beyond simply promoting a conformation of σS that is recognized by ClpX.

Figure 5
Formation of ternary σS–RssB–ClpX complexes. (A) [3H]σS (215 pmoles), 150 pmoles of RssB, and 250 pmoles of ClpX were incubated in the presence of acetyl phosphate and ATP[γ-S] and ...

σS, RssB, ClpX, and ClpP form a quaternary complex

Knowing that ClpX and ClpA form stable complexes with their specific substrates and with ClpP, we anticipated that larger proteolytic complexes containing σS and ClpXP might also be stable. To show this, σS, RssB, ClpX, and ClpP were incubated with a nonhydrolyzable ATP analog, adenylyl-imidodiphosphate (AMP-PNP), acetyl phosphate, and Mg2+ and applied to a gel filtration column. σS and RssB eluted together with ClpXP in the excluded volume of the column, demonstrating a stable quaternary complex (Fig. (Fig.6A).6A).

Figure 6
Formation of quaternary σS–RssB–ClpXP complexes and release of RssB with σS degradation. (A) [3H]σS (215 pmoles), 150 pmoles of RssB, 250 pmoles of ClpX, and 250 pmoles of ClpP were incubated in ...

In the absence of ClpP, ClpX did not associate with σS and RssB in the presence of AMP-PNP, in contrast to the ternary complex formed in the presence of ATP[γ-S] (cf. Figs. Figs.6B6B and and5A).5A). When we attempted to form σS–RssB–ClpXP complexes in the presence of ATP[γ-S], we found that σS was degraded (data not shown). Kinetic experiments showed that the rate of σS degradation was 8-fold slower with ATP[γ-S] than with ATP (data not shown). ClpX hydrolyzes ATP[γ-S] slowly, and ClpXP degrades another substrate, λ O, in the presence of ATP[γ-S], indicating that hydrolysis of ATP[γ-S] serves as the energy source for proteolysis (J. Rozycki and M.R. Maurizi, unpubl.). These results indicate that ClpP, when bound to ClpX, promotes conformational changes in the nucleotide-binding cleft of ClpX that allow AMP-PNP and ATP[γ-S] to bind.

RssB is released from ClpXP as σS is degraded

We wanted to know whether RssB was released from ClpXP as σS was degraded or whether it remained associated with ClpXP for multiple cycles of σS degradation. Therefore, σS, RssB, and ClpXP were incubated in the presence of AMP-PNP, acetyl phosphate, and Mg2+, and quaternary complexes were isolated by gel filtration as described above. The isolated complexes were incubated with a 10-fold molar excess of ATP over AMP-PNP to allow degradation, and then the reaction mixture was reapplied to the gel filtration column (Fig. (Fig.6C).6C). As expected, σS was degraded; the radioactively labeled σS peptides eluted in the fully included volume of the column and were acid-soluble. We found that RssB was released from ClpXP. It eluted as expected for free RssB and was able to stimulate σS degradation by ClpXP in the RssB assay (Fig. (Fig.6C).6C). Thus, RssB is not degraded by ClpXP concomitantly with σS degradation, nor is it inactivated upon release. When [3H]σS was added to aliquots of pooled fractions containing ClpXP (fractions 30–31), <1 pmole of σS was degraded in the absence of added RssB, and 6 pmoles was degraded in the presence of RssB. Thus, prior interaction of ClpXP with RssB does not activate ClpXP to degrade additional σS.

These results, together with the results showing that RssB acts catalytically (Fig. (Fig.2A),2A), indicate that RssB shuttles σS to ClpXP; the RssB–σS complex has high affinity for ClpX, whereas RssB and σS each alone has low affinity for ClpX.

σS is protected from degradation when associated with core RNA polymerase

In vivo, σS exists either free or in a complex with core RNA polymerase, forming an active holoenzyme. To determine if the interaction of σS with core affects RssB-promoted degradation of σS, we mixed core with an equimolar amount of σS, and, after a short incubation, measured degradation of σS by ClpXP in the presence of RssB. We found that core RNA polymerase inhibited σS degradation ~90%, demonstrating that σS is not a substrate for proteolysis when associated with core (Fig. (Fig.7A).7A). Because σS and σ70 are known to compete for binding core, we tested whether the addition of σ70 would allow σS degradation in the presence of core. Addition of σ70 in an equal or greater molar ratio to core resulted in degradation of the σS. This result indicates that RssB recognition sites in σS or ClpX recognition sites in the σS–RssB complex are inaccessible when σS is associated with core RNA polymerase.

Figure 7Figure 7
Competition between RssB and core RNA polymerase for σS binding. (A) Protection of σS from degradation by core RNA polymerase. [3H]σS (18 pmoles) alone (columns 1 and 2) or with 18 pmoles of σ70 (column ...

To determine if the complex of σS–RssB can be bound by core RNA polymerase, σS and RssB were first incubated with acetyl phosphate to generate σS–RssB complexes and then core RNA polymerase was added. Following incubation, the mixture was analyzed by gel filtration column chromatography (Fig. (Fig.7B),7B), and we found that >90% of the σS eluted with core RNA polymerase, whereas >90% of the RssB eluted as expected for monomeric RssB. Therefore, σS interacts with either RssB or with core RNA polymerase, but not with both together. These experiments also show that with equimolar concentrations of core and RssB, core displaces RssB from σS–RssB complexes, indicating that σS has a higher affinity for core than for RssB.

Discussion

In E. coli, regulating σS degradation quickly and reversibly in response to environmental signals is a key aspect of transitions between growth states, such as from exponential to stationary phase or between normal and stress response states. Mutations that block σS degradation lead to inappropriate expression of σS-dependent genes during exponential growth (Muffler et al. 1996a) and to decreased virulence in Salmonella and Erwinia (Swords et al. 1997; Andersson et al. 1999).

We have shown that in vitro, as in vivo, σS is degraded by the ClpXP protease. Degradation is greatly stimulated by RssB, and the stimulation by RssB depends on acetyl phosphate. RssB promotes σS degradation while having no effect on ClpXP activity with other substrates. Moreover, RssB itself is not consumed during the reaction. Thus, RssB is a specificity factor that delivers σS to ClpXP for degradation.

Our model for the pathway of degradation of σS is shown in Figure Figure8.8. σS is able to interact with both core RNA polymerase and RssB, but not with both simultaneously (Fig. (Fig.7).7). When bound by core, σS is protected from degradation. In the degradation pathway, σS initially binds the phosphorylated form of RssB with acetyl phosphate serving as the phosphate donor (Fig. (Fig.3;3; Becker et al. 1999). In vivo, acetyl phosphate contributes to phosphorylation of RssB (Bouche et al. 1998), but other phosphate donors must exist that have not been identified yet. The σS–RssB complex is bound by ClpX in the presence of ATP[γ-S] with high affinity, although neither RssB nor σS alone binds ClpX with high affinity (Fig. (Fig.5).5). These observations indicate that motifs on both RssB and σS contribute to the tight binding seen in the ternary complex and/or that complex formation exposes a motif on σS that ClpX recognizes and binds with high affinity. Because RssB has been found associated with ClpX by other methods (Moreno et al. 2000; R. Hengge-Aronis, unpubl.), RssB may have a direct role in interacting with ClpX in the ternary complex, possibly orienting σS to favor its binding to ClpX. When ClpP is present, a quaternary complex of σS–RssB–ClpXP is formed in the presence of a nonhydrolyzable ATP analog, AMP-PNP (Fig. (Fig.6).6). This proteolytic complex catalyzes σS degradation upon the exchange of AMP-PNP with ATP. ClpX most likely catalyzes ATP-dependent unfolding of σS, as has been shown for other ClpX substrates (Kim et al. 2000; Singh et al. 2000). The unfolded σS is then translocated through the narrow opening of ClpP into the interior chamber, where the protease active sites reside (Wang et al. 1997). Concurrent with or following σS degradation, RssB is released and is able to carry out multiple cycles of σS binding and delivery to ClpXP (Fig. (Fig.6).6).

Figure 8
Model for the mechanism of action of RssB in regulating σS degradation by ClpXP. See text for discussion.

In vitro, acetyl phosphate greatly stimulates σS degradation by ClpXP and RssB and is required for the interaction of RssB and σS (Figs. (Figs.1,1, ,3;3; Becker et al. 1999). These observations suggest that signals that affect the phosphorylation state of RssB modulate the activity of RssB. Phosphorylation very likely stabilizes a conformation of RssB that has high affinity for σS. However, phosphorylation may not be essential. We observed that in the absence of acetyl phosphate some σS and RssB associated with ClpX, although the amount was fivefold less than in the presence of acetyl phosphate. Moreover, when overexpressed, a mutant of RssB that cannot be phosphorylated can promote σS degradation in vivo (Bouche et al. 1998; unpubl.). Nonetheless, the higher affinity of σS for the phosphorylated form of RssB and the acetyl phosphate requirement for degradation support the critical role for RssB phosphorylation in promoting σS degradation under normal in vivo conditions.

The observation that core RNA polymerase protects σS from degradation suggests a second possible level of regulation of σS degradation (Fig. (Fig.7).7). Under the unusual conditions of high RssB expression and absence of ClpXP protease, RssB interferes with σS activity in vivo (Zhou and Gottesman 1998; Becker et al. 2000). These observations are consistent with a role for competition between RssB and core RNA polymerase for σS under some conditions and provide support for the direct interaction between σS and RssB that has been detected in vitro. If the ability of σ70 or other sigma factors to interact with core were diminished under some conditions, core would be more available to bind σS, protecting it from rapid degradation. Core RNA polymerase appears to be limiting during stationary phase growth, because an rpoS mutant allows better in vivo use of σ70 (Farewell et al. 1998). Thus, sigma factor competition for core may play a significant regulatory role in vivo. The identification of a stationary phase-specific factor, Rsd, that interacts with σ70 and interferes with the activity of σ70 (Jishage and Ishihama 1998) is consistent with the idea that sigma–core interactions may be regulated and may in turn regulate the susceptibility of σS to degradation. Degradation of σ32 also appears influenced by its interactions with core polymerase (Blaszczak et al. 1999).

RssB is an example of a trans-targeting protein, specifically enabling the degradation of σS by ClpXP. Use of such specific and regulatable targeting proteins may be essential for the regulated degradation of physiologically important substrates. The concept of trans-recognition was proposed by Varshavsky and coworkers (Johnson et al. 1990) for targeting of certain proteins for degradation in Saccharomyces cerevisiae. They showed that the presence of an amino acid that acts as a degradation signal at the amino terminus of one subunit of an oligomeric protein was sufficient to target the degradation of another subunit, which lacked its own degradation signal but carried a proximal site for ubiquitin conjugation. Recognition of the unusual N terminus was necessary to attract the ubiquitination machinery, but ubiquitin was conjugated to the subunit carrying the ubiquitination site, and only that subunit was degraded when the complex was presented to the 26S proteasome.

In addition to the trans-targeting of σS by RssB to ClpXP reported here, two other examples have been recently described for the trans-targeting of substrates to ClpXP in E. coli. UmuD‘, a processed and active form of the UmuD error-prone mutagenesis protein of E. coli, is degraded by ClpXP only when in a heteromeric complex with UmuD (Gonzalez et al. 2000). The unprocessed N terminus of UmuD is an essential element for recognition of UmuD‘ by ClpXP, but only UmuD‘ is degraded. Degradation of UmuD‘ acts to down-regulate mutagenesis as the cell recovers from DNA damage. Another example of trans-targeting is the stimulation of ClpXP degradation of SsrA-tagged proteins by SspB. It has recently been found that a ribosome-associated protein, SspB, binds specifically to SsrA-tagged proteins and enhances the recognition of these substrates by ClpXP without itself being degraded (Levchenko et al. 2000). SspB stimulates degradation of SsrA-tagged proteins both in vivo and in vitro.

It is intriguing that both RssB and SspB act by increasing the affinity of a specific substrate for ClpX. If, alternatively, the trans-targeting proteins modified the protease itself, degradation of many substrates would be affected. In eukaryotic cells, the diversity of ubiquitin ligase complexes provides a similar means for regulating access of specific substrates under given conditions without perturbing other degradative pathways. Possibly the small number of examples of regulated proteolysis in prokaryotes reflects the absence of a generalized regulatory network such as ubiquitin tagging, and therefore the evolution of novel solutions for each situation.

Materials and methods

Materials

ATP, AMP-PNP, and ATP[γ-S] were obtained from Boehringer Mannheim. Sephacryl S-100 and S-200 High Resolution gel filtration media were obtained from Pharmacia.

Proteins

ClpX (Grimaud et al. 1998), ClpP (Maurizi et al. 1994), ClpA (Maurizi et al. 1994), P1 RepA (Wickner 1990), and λ O (Wickner and Zahn 1986) were purified as described. σ70 was a gift from Jeffery Roberts (Cornell University). The core RNA polymerase used in the experiments described in Figure Figure7A7A was a gift from Ding Jin (NIH); the core used in Figure Figure7B7B was purchased from Epicentre Technologies. σS was isolated from cells carrying pLHN14 and expressing σS (Nguyen et al. 1993) using a described procedure (Tanaka et al. 1993) with some modifications. σS, λ O, RepA, and α-casein were labeled in vitro as previously described (Hoskins et al. 1998). The final specific activities of [3H]σS, [3H]λ O, [3H]RepA, and [3H]α-casein were 1150, 450, 22,000, and 5500 cpm/pmole, respectively.

Throughout, proteins are expressed as moles of RssB monomers, ClpX hexamers, ClpP tetradecamers, ClpA hexamers, σS monomers, σ70 monomers, λ O dimers, RepA dimers, and α-casein monomers.

Purification of RssB

For the preparation of RssB, the rssB gene was amplified by the polymerase chain reaction and inserted in a T7 expression vector, plasmid pVex11 (Studier et al. 1990). The sequence of rssB was verified by DNA sequencing. Cultures of E. coli BL21 lysS, rssB (YN305) carrying the plasmid pT7-RssB were grown to an O.D.600 of 0.5, induced by the addition of 1 mM IPTG, incubated for 2 h, collected, and frozen. Thawed cells were adjusted to 20 mM Tris-HCl, 0.2 M NaCl, 1 mM dithiothreitol (DTT), and 0.1 mM EDTA, and passed through a French pressure cell (Aminco) at 20,000 psi at 0°C. RssB activity was followed by measuring the stimulation of σS degradation by ClpXP in the assay described below. The lysate was centrifuged at 30,000g for 60 min at 4°C, and the pellet was resuspended in 8 M urea to give 12 mg/mL protein. After centrifuging as above, the supernatant was dialyzed against Buffer A (20 mM Tris-HCl, 10% [v/v] glycerol, 1 mM DTT, and 0.1 mM EDTA) for 2 h and diluted to a final NaCl concentration of 0.05 M. The sample was applied to a Q-sepharose column that was equilibrated with Buffer A. The column was washed with Buffer A, and protein was eluted with a linear gradient from 0 to 0.5 M NaCl in Buffer A. RssB eluted with 0.2–0.3 M NaCl. The protein was further purified by Sephacryl S-100 high resolution gel filtration in Buffer A containing 0.2 M NaCl, where it eluted as expected for a monomer. The RssB fractions were stored at −70°C. The final preparation was >90% pure as determined by commassie staining following SDS-PAGE.

σS in vitro degradation assay

Reaction mixtures were assembled in 20 μL of Buffer B (20 mM Tris-HCl at pH 7.5, 10 mM MgCl2, 140 mM KCl, 1 mM DTT, 0.1 mM EDTA, and 5% glycerol [v/v]) containing 5 mM ATP, 25 mM acetyl phosphate, 2 pmoles ClpX, 2 pmoles ClpP, 19 pmoles [3H]σS, and 1 pmole RssB, unless otherwise indicated. The mixtures were incubated at 24°C for 20 min unless otherwise indicated. TCA (trichloro acetic acid) was added to 20% (w/v), and [3H]σS degradation was quantitated by measuring acid soluble radioactivity. In the absence of ClpX, ClpP, and RssB, <1% of the [3H]σS was TCA-soluble. One unit (U) of RssB stimulates the degradation of 1 pmole of σS in 20 min with the standard assay conditions. When crude fractions were used as the source of RssB, 25 mM phosphocreatine and 1 μg of creatine kinase were added to reaction mixtures. Degradation by ClpAP was measured similarly in 20-μL reaction mixtures containing 1 mM ATP, 1 pmole ClpA, 1 pmole ClpP, 1 pmole of RssB (where indicated), and 1 pmole of [3H]α-casein, 1 pmole of [3H]RepA, or 18 pmoles of [3H]σS in Buffer B.

Isolation of RssB–σS, RssB–σS–ClpX and RssB–σS–ClpXP complexes by gel filtration chromatography

To isolate σS–RssB complexes, 100-μL mixtures containing 320 pmoles of [3H]σS, 150 pmoles of RssB, and 25 mM acetyl phosphate in Buffer B were incubated for 15 min at 23°C. The mixtures were applied onto a Sephacryl S-100 HR column (0.7 cm × 7 cm) equilibrated with Buffer B containing 25 mM acetyl phosphate and 50 μg/mL of bovine serum albumin. Fractions (120 μL) were collected, and σS was determined by measuring radioactivity by scintillation counting. RssB was determined using the σS degradation assay described above.

For the isolation of σS–RssB–ClpX complexes, 215 pmoles of [3H]σS, 250 pmoles of ClpX, 150 pmoles of RssB, 5 mM ATP[γ-S], and 25 mM acetyl phosphate in Buffer B were incubated in 100-μL reaction mixtures for 15 min at 23 °C. The reaction mixtures were then applied onto a Sephacryl S-200 HR column (0.7 cm × 7 cm) equilibrated with Buffer B containing 1 mM ATP[γ-S] and 25 mM acetyl phosphate. Fractions (120 μL) were collected. σS and RssB were determined as above. ClpX was measured by densitometry of SDS-PAGE of the fractions. To isolate σS–RssB–ClpXP complexes, 215 pmoles of [3H]σS, 250 pmoles of ClpX, 150 pmoles of RssB, 250 pmoles of ClpP, 5 mM AMP-PNP, and 25 mM acetyl phosphate in Buffer B were incubated in 100-μL reaction mixtures for 15 min at 23°C. The reaction mixtures were then applied onto a Sephacryl S-200 HR column as described above for S-100 column chromatography and equilibrated with Buffer B containing 1 mM AMP-PNP and 25 mM acetyl phosphate, and 120-μL fractions were collected. σS, RssB, and ClpX were determined as above, and ClpP was measured by densitometry after SDS-PAGE of the fractions.

Ultrafiltration of σS, RssB, and ClpX

In 100-μL reaction mixtures, 9 pmoles of [3H]σS was incubated with 60 pmoles of ClpX, various amounts of RssB, 1 mM ATP[γ-S], 20 mM acetyl phosphate, 0.01% (v/v) Triton X-100, and 25 μg/mL bovine serum albumin in Buffer B for 30 min at 24°C. NaCl (1 M) was added, and the mixtures were incubated for 10 min at 24°C. Reaction mixtures were centrifuged through Microcon 100 ultrafiltration devices (Amicon) for 10 min at 3200g. Retentates were recovered with 100 μL of 10% (w/v) SDS, and the radioactivity was measured.

Acknowledgments

We thank Jeffrey Roberts for σ70, Regis Grimaud for ClpX, Ding Jin for core RNA polymerase, and Richard Burgess for pLHN14.

The publication costs of this article were defrayed in part by payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 USC section 1734 solely to indicate this fact.

Footnotes

E-MAIL vog.hin.xileh@gnasus; FAX: (301) 496-3875.

E-MAIL vog.hin.xileh@kciweus; FAX: (301) 402-1344.

Article and publication are at www.genesdev.org/cgi/doi/10.1101/gad.864401.

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