INTRODUCTION

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Bacteria are constantly sampling their surroundings and regulating gene expression accordingly. Since many of the environments they encounter often have hazardous conditions (for example, limiting nutrients, high osmolarity, extreme pH, or extreme temperature), bacteria have evolved to survive in such hostile habitats. In particular, the gram-negative bacterium Escherichia coli enters a state in its life cycle known as the stationary phase, which renders it highly resistant to unfavorable environmental conditions.

When cells enter stationary phase, they undergo dramatic changes in their morphology and physiology that increase their chance for survival in a wide variety of stresses. This cross-protection results from the global control system regulated by RpoS. RpoS, encoded by the rpoS gene, is the second primary  factor of E. coli, and it is required for the transcription of stationary-phase-specific genes. Due to the drastic consequences (i.e., slowed metabolism) of entering stationary phase, RpoS is tightly regulated. In fact, RpoS is regulated at all levels: transcription, translation, protein stability, and activity (for a recent review, see reference 13).

Among the possible stresses that can induce RpoS (the stationary-phase response), starvation of an essential nutrient is perhaps the most widely studied. When nutrients are readily available, the levels of RpoS are very low, mainly due to its efficient degradation by the ATP-dependent ClpXP serine protease. Conversely, when nutrients become limiting for growth, this ClpXP-dependent proteolysis stops and, consequently, RpoS levels increase significantly (26). This mode of RpoS regulation has been shown to occur in response to carbon starvation as well as during growth in Luria-Bertani (LB) medium, although the specific signals sensed in the latter medium remain to be determined (25, 32).

The regulation of RpoS proteolysis is not mediated by controlling either the levels of the ClpXP protease itself or its activity (33). Instead, it is orchestrated by the response regulator SprE (named RssB in E. coli, MviA in Salmonella enterica serovar Typhimurium, and ExpM in Erwinia carotovora) (1, 2, 22, 25). In a recent report, Zhou et al. demonstrated in vitro that SprE plays a catalytic role in the delivery of RpoS to ClpX, the regulatory component of ClpXP that is believed to unfold RpoS and eventually feed it to ClpP, the proteolytic component (34). ClpP then degrades RpoS and SprE is released from the proteolytic complex. Furthermore, Zhou et al. also showed that this in vitro degradation is greatly enhanced upon SprE phosphorylation (34).

To date, it remains unknown how SprE is phosphorylated in vivo; therefore, SprE is an orphan response regulator. Moreover, unphosphorylated SprE can still promote, although less efficiently, RpoS degradation (reference 6 and unpublished results cited in reference 34). This raises the possibility that SprE might be regulated by a mechanism(s) other than phosphorylation.

A possible mechanism for regulating SprE-mediated degradation of RpoS is to control the levels of SprE itself. Paradoxically, the levels of SprE (and MviA) have been shown to increase when cells enter stationary phase (11, 21). Specifically, it has been shown that the translation of sprE increases in an RpoS-dependent manner (11). In addition, it was reported that sprE (and mviA) transcription is also upregulated during the stationary phase (11, 21). Although those studies did not prove the exact location of the sprE promoter, their reporter fusion data showed that sprE transcription can occur independently from that of its upstream gene, rssA (11). Prior to these studies it was assumed, based on DNA sequence analysis, that rssA and sprE constitute an operon (5, 22). In addition, RssA itself was implicated in the SprE pathway regulating RpoS, although its function has never been clearly demonstrated (unpublished results cited in references 13 and 22).

Since it is unclear how sprE transcription is regulated and what role RssA plays in RpoS regulation, we have constructed a series of reporter fusions and rssA null alleles that allow us to address these issues. In this report we present data demonstrating that although rssA and sprE constitute an operon, sprE can also be transcribed from an RpoS-regulated promoter located in the rssA-sprE intergenic region. In addition, while RpoS controls RssA levels, we found no role for RssA in the regulation of either SprE or RpoS under the conditions tested.

	MATERIALS AND METHODS

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Bacterial strains and bacteriophages. E. coli DH5 (Invitrogen Life Technologies) was used as the host strain for all plasmid constructions. All other bacterial strains used (Table 1) are derivatives of MC4100 (7). Standard microbial techniques were used for strain construction (27). All fusions were recombined with RZ5 (23).

Media and growth conditions. LB medium was prepared as described previously (27). Unless indicated, all bacterial strains were grown under aeration at 37°C and their growth was monitored by measuring the optical density at 600 nm (OD₆₀₀).

DNA manipulations. Plasmid DNA was purified by standard techniques and was introduced into the appropriate strains by the method of Kushner (17). All restriction endonucleases (New England Biolabs), Taq and Pfu DNA polymerases (Roche and Stratagene, respectively), T4 polynucleotide kinase (T4 PNK) (New England Biolabs), and T4 DNA ligase (New England Biolabs) were used according to the recommendations of their respective manufacturers. Primers were synthesized by the Princeton University Department of Molecular Biology Synthesis and Sequencing Facility.

Plasmid construction. The plasmids used in complementation studies were constructed as follows. A 3.06-kbp fragment containing the ychJ-rssA-sprE region was amplified from MC4100 chromosomal DNA using the primers RSSAUP (5'-CGGGATCCCGCCAGAGAACGTAAAGTATCG) and 3SPREMBP2 (5'-CCACCAGCCAAGCTTAGCAGG), which introduce BamHI and HindIII restrictions sites (underlined) at the 5' and 3' ends of the PCR products, respectively. After digestion with these restriction enzymes, the PCR fragment was inserted between the BamHI and HindIII sites of pBluescript KS(+) (Stratagene) and pBBR1MCS (15) to yield pRSSA5 and pRSSA6, respectively.

Insertional inactivation of rssA. A null rssA allele was constructed by insertion of a kanamycin cassette into the PstI site in the RssA-coding region (580 bp after the first guanine of the GTG start codon). The RssA open reading frame was amplified from MC4100 chromosomal DNA by PCR using primers 5RSSAKPN1 (5'-GCGATATGGGTACCATTGCATTCC) and 3RSSAHIND3 (5'-TTCTGGTCAAGCTTCGGTGCGTACC), which introduce KpnI and HindIII restriction sites (underlined), respectively. The resulting 0.94-kbp fragment was digested with KpnI and HindIII and ligated with KpnI-HindIII-digested pRSETB (Invitrogen Life Sciences), and the new plasmid was named pRSSA2. Then, the 794-bp EcoRV-HindIII fragment containing the 3' end of rssA was introduced into SmaI-HindIII-digested pBluescript KS(+) to generate pRSSA3. The PstI fragment containing the kanamycin cassette from pUC4K (30) was then inserted into the PstI site of pRSSA3 to create pRSSA3::kan. The kanamycin cassette was inserted in an orientation opposite to that of rssA transcription (confirmed by DNA sequencing). The XbaI-KpnI fragment from pRSSA3::kan, containing rssA::kan, was ligated with XbaI-KpnI-digested pAMPTS (24), and the resulting plasmid, pRSSA::kanTs, was used for the allelic exchange in MC4100 as previously described (12).

Introduction of an in-frame deletion into rssA. A defined in-frame deletion in rssA was generated in pRSSA5 (which carries the ychJ-rssA-sprE region; see above) by using the method of inside-out PCR previously described (14). Briefly, pRSSA5 served as the template for an inside-out PCR with primers RTRSSA1 (5'-CCTCTCGCCGCGCCAGATCCC) and RTRSSA2 (5'-GCACGATGCTCATTTGATGC), which were designed to create an in-frame deletion of amino acids 30 to 194 of RssA. The 5.47-kbp PCR product was phosphorylated using T4 PNK and self-ligated to yield pRSSA5, which contains the 492-bp in-frame deletion allele of rssA named rssA1. The pRSSA5 plasmid was then digested with the BamHI and HindIII restriction endonucleases and the fragment containing the ychJ-rssA1-sprE region was inserted between the BamHI and HindIII sites of pBBR1MCS to generate pRSSA6.

Construction of rssA-lacZ and sprE-lacZ fusions. An rssA'-`lacZ translational fusion containing the ychJ-rssA' region was constructed as described below. Two sprE'-lacZ<sup>+</sup> transcriptional fusions were also made: one containing the ychJ-rssA-sprE' region (rssA-sprE'-lacZ<sup>+</sup>) and the other containing only an `rssA-sprE' fragment (`rssA-sprE'-lacZ<sup>+</sup>). The appropriate regions (see below) were inserted into either pRS414 (for the rssA'-`lacZ fusion) or pRS415 (for both sprE'-lacZ⁺ fusions) (28). All fusion-containing plasmids were recombined into the phage RZ5 (23) as described by Simons et al. (28). For integration of the fusions at the  attachment (att) site, MC4100 was infected with the appropriate recombinant  phage. For integration of the fusions at the chromosomal sprE locus, att deletion MC4100 derivative strains (Table 1) were infected with the appropriate recombinant  phage.

-Galactosidase assays. After growing overnight in LB broth, cells were diluted 1:100 into fresh LB broth and grown to an OD₆₀₀ of ~0.3 to 0.4 for logarithmic-phase samples or to an OD₆₀₀ of ~3.0 for stationary-phase samples. -Galactosidase assays were performed using a microtiter plate assay as described previously (29). The -galactosidase activities were expressed as OD₄₂₀/(OD₆₀₀ × volume), where volume refers to the amount (in milliliters) of cell lysate used. For each experiment, every sample was assayed three times and the average activity and standard deviation (SD) were obtained. The data shown resulted from a single experiment representative of at least three other independent experiments.

Western blot analysis. Cells were grown as indicated above for the -galactosidase assays to obtain both logarithmic- and stationary-phase samples. Once cells reached the indicated OD₆₀₀, 1-ml samples were pelleted. To standardize samples, the pellets were resuspended in a volume of sodium dodecyl sulfate (SDS) sample buffer (18) equal to the OD₆₀₀/6. Samples were boiled for 5 min and equal volumes were subjected to SDS-12% polyacrylamide gel electrophoresis as described by Laemmli (18). The proteins were transferred to nitrocellulose membranes (Schleicher & Schuell), and Western blot analyses were performed as previously described (10). When appropriate, polyclonal sera against RpoS or SprE were used as primary antibodies at a dilution of 1:6,000 and 1:4,000, respectively. Donkey anti-rabbit immunoglobulin G-horseradish peroxidase conjugate (Amersham Pharmacia Biotech) was used as secondary antibody at a 1:6,000 dilution. For visualization of bands, the ECL antibody detection kit (Amersham Pharmacia Biotech) and X-Omat film (Kodak) were used.

Primer extension analysis. AF633 cells were grown to stationary phase as described above for the -galactosidase assays. Total RNA was extracted using Trizol (Invitrogen Life Sciences). Primer RTSPRE1 (5'-AGCCGCCAGTACCGTTGTCGC) was labeled with [-³³P]ATP (ICN) using T4 PNK prior to primer extension. For the primer extension reaction, 5 µg of total RNA, labeled primer, and 100 U of Moloney murine leukemia virus reverse transcriptase (U.S. Biochemicals) were used as directed by the manufacturer. For the sequencing reaction, 3 µg of pRSSA5 plasmid was digested with HindIII for 15 min. After the digestion, pRSSA5 was mixed with 0.5 pmol of unlabeled RTSPRE1 primer and denatured by boiling for 3 min. After cooling on ice for 20 min, the sequences were determined by using a Sequenase DNA sequencing kit (version 2; U.S. Biochemicals) with [-³³P]dATP (ICN) according to the directions of the manufacturer. All reactions were subjected to electrophoresis in an 8.3 M urea-6% polyacrylamide gel. The reaction products were visualized on X-Omat film (Kodak).

	RESULTS

Top Abstract Introduction Materials and Methods Results Discussion References

Disruption of rssA increases RpoS-mediated transcription. Sequence homology suggests that RssA belongs to a family of serine esterases/proteases found from bacteria to humans (20). Unfortunately, the only members of this family with a characterized function are those found in Drosophila and humans; their function pertains to neuronal development and they contain additional domains not found in their bacterial counterparts (16, 20).

Disruption of rssA increases RpoS levels by altering SprE levels. Since SprE regulates RpoS at the level of protein stability, it is likely that the increase in RpoS-mediated transcription caused by the rssA1::kan allele reflects increased RpoS levels rather than increased specific activity of RpoS. To test this, we determined the relative levels of RpoS by Western blot analysis in both logarithmic- and stationary-phase cultures of various strains grown in LB medium.

View larger version (50K): [in this window] [in a new window]   FIG. 1.   Disruption of rssA increases the levels of RpoS by reducing those of SprE, owing to polarity. (A and B) Western blot analysis was used to monitor the levels of RpoS in total cell lysates recovered during the logarithmic (A) and stationary (B) phases of growth in LB medium. (C) The same stationary-phase samples were used to determine the levels of SprE. The bands corresponding to RpoS and SprE are marked with labeled arrows. Lanes in each panel: wt, strain AF633; rpoS::kan, strain NR247; sprE19::cam, strain NR252; sprE::tet, strain NR253; rssA1::kan, strain NR260; rssA1::kan sprE19::cam, strain NR270; rssA1::kan sprE::tet, strain NR286. The sample orders are the same in all panels.

The decreased levels of SprE in the rssA1::kan null strain are the result of polarity. The decreased levels of SprE reported above that occurred upon disruption of rssA can be explained as follows. If rssA and sprE are cotranscribed, disruption of rssA would result in lowered sprE expression. Alternatively, RssA could be a positive regulator of SprE levels. In addition, both of these scenarios could be true.

View larger version (51K): [in this window] [in a new window]   FIG. 2.   RssA does not regulate SprE or RpoS. Complementation studies were done in stationary-phase cells grown in LB medium. RpoS and SprE levels (corresponding bands marked with labeled arrows) were monitored by Western blot analysis. (A) RpoS levels present in wild-type (wt) AF633 and rssA1::kan NR260 strains carrying the vector plasmid (pBBR1MCS), a plasmid encoding the ychJ-rssA-sprE region (pRSSA6), or one carrying the ychJ-rssA region (pRSSA9) (as marked above the lanes). (B) SprE levels present in wild-type AF633 and rssA1::kan NR260 strains carrying either pBBR1MCS, pRSSA6, or pRSSA9. The sample order is the same as for panel A. (C) RpoS levels in wild-type (wt) AF633 and rssA1::kan NR260 strains carrying either the vector plasmid pMAL-c2 (vector) or a plasmid encoding the MBP-SprE hybrid protein pMBPSprE (SprE), respectively.

rssA and sprE constitute an operon. To further support that polarity alone is responsible for the decreased levels of SprE in the rssA1::kan strain, we examined the levels of SprE in a cell depleted of RssA by an in-frame deletion in rssA. We introduced an allele (rssA1) that carries an internal in-frame deletion (encompassing amino acids 30 to 194) in rssA into a low-copy-number plasmid (pRSSA6) and determined its effects on SprE levels by Western blot analysis.

rssA1

rssA1

View larger version (15K): [in this window] [in a new window]   FIG. 3.   In-frame deletion in rssA has no effect on SprE levels. A plasmid carrying the in-frame deletion allele rssA1 was introduced into the rssA1::kan strain NR260 and the levels of SprE in logarithmic (L) and stationary (S) phase cells were monitored by Western blot analysis. The relevant genotype from the plasmid is shown above the lanes. Vector, samples from strain NR260(pBBR1MCS); wt, NR260 (pRSSA6); rssA1, samples from NR260(pRSSA6).

The levels of SprE are growth phase regulated at the level of transcription. Previous reports have shown that in both E. coli and S. enterica serovar Typhimurium, SprE (MviA) is growth phase regulated (11, 21). Consistent with this, we find that there is a significant increase in SprE levels between samples prepared from logarithmic- versus stationary-phase cells (Fig. 3). Paradoxically, this increase in SprE is RpoS dependent (11). This explains why an rpoS::kan null strain contains less SprE than its parent wild-type strain (Fig. 1C, compare wt and rpoS::kan lanes).

View larger version (19K): [in this window] [in a new window]   FIG. 4.   RssA levels are growth phase regulated by RpoS. The -galactosidase levels of an rssA'-`lacZ fusion were measured in a wild-type (NR501) and an rpoS::kan (NR502) strain during logarithmic (hatched bars) and stationary (black bars) phases of growth in LB medium. The relative levels of -galactosidase activity are shown as the average and SD of three measurements per sample and they are representative of at least three independent experiments.

View larger version (16K): [in this window] [in a new window]   FIG. 5.   RpoS regulates SprE at the transcriptional level. The -galactosidase levels of an rssA-sprE'-lacZ+ fusion were measured in wild-type strains (NR507 carries the fusion at the att site and NR509 carries the fusion at the sprE chromosomal locus) and rpoS::kan strains (NR508 carries the fusion at the att site and NR510 carries the fusion at the sprE chromosomal locus) during logarithmic (hatched bars) and stationary (black bars) phases of growth in LB medium. The location where the fusion was integrated (att site or sprE locus) and the rpoS genotype are indicated. The relative levels of -galactosidase activity are shown as the average and SD of three measurements per sample, and they are representative of at least three independent experiments.

sprE is transcribed from an RpoS-dependent promoter located in the rssA-sprE intergenic region. The last result described above confirms that although rssA and sprE constitute an operon, there is an additional promoter(s) from which sprE can be transcribed. This promoter must lie between the location of the kan cassette insertion and the translational start of sprE, since SprE is still made in the presence of the rssA1::kan allele. We have also observed that the already-decreased levels of SprE present in an rssA1::kan strain are growth phase regulated, since they are undetectable in the logarithmic growth phase by Western blot analysis (data not shown), suggesting that this second sprE promoter is also growth phase regulated. To better understand the nature of this promoter, we constructed an additional sprE transcriptional fusion and conducted primer extension analysis.

View larger version (13K): [in this window] [in a new window]   FIG. 6.   The intergenic rssA-sprE region contains an RpoS-dependent promoter for sprE. The -galactosidase levels of an `rssA-sprE'-lacZ+ fusion were measured in a wild-type strain (NR511) and an rpoS::kan strain (NR512) during logarithmic (hatched bars) and stationary (black bars) phases of growth in LB medium. The relative levels of -galactosidase activity are shown as the average and SD of three measurements per sample, and they are representative of at least three independent experiments.

View larger version (29K): [in this window] [in a new window]   FIG. 7.   Location of the sprE promoter in the rssA-sprE intergenic region. (A) Primer extension analysis on RNA prepared from stationary-phase wild-type cells (AF633) showed that there is a transcriptional start site located 21 bp upstream of the adenine of the ATG start codon of sprE. The primer extension reaction (shown on the right) was performed using the RTSPRE1 primer as described in Materials and Methods. A sequencing reaction was also performed using pRSSA5 as the template and RTSPRE1 as the primer. The section of the gel corresponding to the sequence of the noncoding strand for the relevant region is shown. The order of loading in the sequencing gel is marked above the lanes and the corresponding sequence of the coding strand appears on the right, with the transcriptional start site marked with an arrow pointing to the direction of transcription. (B) Sequence of the putative promoter (coding strand alone) and comparison with the consensus sequence (similarities appear underlined) of RpoS-dependent promoters (top line) as described by Becker and Hengge-Aronis (3). The location of the transcriptional start is marked with an arrow and it corresponds to +1. The sprE start codon is shown in lower case. The locations of the putative 10 and 35 regions are marked, but because RpoS-dependent promoters do not have a recognizable 35 consensus region, none is shown. The conserved 13 position is also marked with an asterisk. In RpoS-dependent promoters, the 15-bp-long sequence between the 10 and 35 regions is AT rich. K, G or T; W, A or T; R, A or G.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

The developmental commitment that E. coli makes when RpoS levels increase is immense; therefore, the cellular content of RpoS is tightly regulated. RpoS is controlled at multiple levels and regulation of its degradation by the ClpXP protease is considered a major level of control (13, 32). Although it has been known for several years that the response regulator SprE orchestrates this degradation of RpoS (22, 25), a critical question remains to be answered: how is SprE's activity regulated? We believe that the knowledge gained from understanding how SprE expression is regulated will help us to answer this question.

The studies presented here were aimed at examining sprE expression. By using rssA null alleles as well as reporter fusions and primer extension analysis, we have demonstrated that rssA and sprE are cotranscribed from a promoter, P₁, that is regulated by RpoS. We have also identified an additional promoter, P₂, located in the rssA-sprE intergenic region from which sprE is transcribed in an RpoS-dependent fashion. Furthermore, we have shown that under the conditions tested in this report, RssA is not involved in the regulation of either SprE or RpoS.

Interestingly, the P₂ promoter has features found in the consensus for promoters recognized by RpoS recently proposed by Becker and Hengge-Aronis (3). Specifically, it lacks a recognizable 35 region but it contains a run of A/T between the 30 and 14 positions. It also contains the GC motif at the 14 and 13 positions and the highly conserved T at position 6, besides a 10 region partially homologous to the proposed TATACT consensus. Although the existence of an RpoS-specific consensus is somewhat controversial (see below), it is important to emphasize that Becker and Hengge-Aronis have shown both allelic suppression between a C at position 13 and residue 173 of RpoS (i.e., proving direct interaction) and the necessity of having either a G or a T at position 14 for maximal expression of RpoS-dependent promoters (3). The sprE P₂ promoter fulfills both criteria, having the highly conserved C at the 13 position and a G at the 14 position. More experiments are required, though, to demonstrate the specific role of these positions in sprE expression.

Although many RpoS-dependent promoters have been identified through the years, deriving a consensus from them has not been an easy task. This difficulty arises because RpoS and ⁷⁰ are so similar (19). In a recent report, in vitro selection studies searching for the promoter sequences best recognized by RpoS showed that both RpoS and ⁷⁰ prefer the same consensus sequences. These studies propose that the specificity of these sigma factors is dictated by how they differ in tolerating binding to nonpreferred sequences (i.e., tolerance to binding to sites with deviations from the well-characterized ⁷⁰ consensus sequences) (T. Gaal, W. Ross, S. T. Estrem, L. H. Nguyen, R. R. Burgess, and R. L. Gourse, submitted for publication). Regardless of whether there is a clear RpoS promoter consensus sequence or not, depletion of RpoS significantly decreases transcription of sprE from both P₁ and P₂ promoters, proving RpoS-dependent transcription of sprE.

Previously, it was reported that although transcription from a fusion containing only the P₂ promoter (i.e., it did not include all of rssA) was upregulated in stationary phase, this growth phase regulation was RpoS independent (11). Furthermore, an analogous translational fusion was shown to be RpoS regulated, suggesting translational but not transcriptional control of sprE by RpoS. Additional evidence supporting this RpoS-dependent translational regulation of sprE showed that the levels of SprE present in a strain carrying the sprE19::cam allele were also growth phase regulated (i.e., they increased in stationary phase) (11). Note that the sprE19::cam allele carries a mini-Tncam cassette inserted 27 bp upstream of the translational start of sprE and it is believed to be constitutively transcribed (25).

In contrast, we found that the sprE P₂ promoter is RpoS regulated. We have recently isolated a strain that contains a mini-Tncam cassette inserted 95 bp from the translational start of sprE and, in this strain, sprE is constitutively expressed regardless of the growth phase or the presence or absence of RpoS (data not shown), which argues against an RpoS-dependent translational control. To resolve this controversy, we sequenced the region upstream of sprE and found that several strains previously used (11) contain an IS1E element in the rssA-sprE intergenic region which interferes with native sprE expression and regulation.

The paradox of RpoS being necessary for the expression of its negative regulator SprE remains (11). We speculate that by having this feedback loop, the cell ensures a proper amount of RpoS at all times. It is known that under certain conditions, too much RpoS is not beneficial to the cell and can even be fatal. For example, Zambrano et al. showed that during prolonged starvation, cells with an altered form of RpoS, which is less active as sigma factor, results in growth advantage (31). In addition, cells that have exceptionally high levels of RpoS (i.e., sprE null strains) cannot grow in media containing either succinate or acetate as the sole carbon source (24). Thus, coupling the levels of SprE to those of RpoS might serve as a safety mechanism to ensure that the levels of RpoS are appropriate in the cell at all times. Interestingly, it has been reported that in addition to its role in orchestrating RpoS degradation, SprE has anti-sigma factor activity (4, 33). Thus, it is possible that in stationary-phase cells, when SprE-mediated degradation of RpoS does not occur, SprE itself might be acting as an anti-RpoS factor. This could explain, at least in part, why SprE levels need to increase when ClpXP is not degrading RpoS.

In addition, as previously proposed, having SprE already present in stationary phase might be beneficial to cells once they encounter more favorable conditions, thus providing a growth advantage (11). High levels of SprE could ensure rapid degradation of RpoS when nutrients become available. RpoS-dependent transcription would cease, and the cell would then focus its transcriptional and translational machinery on producing proteins that are necessary for rapid growth (⁷⁰-dependent promoters). Reports showing that cells deficient in ClpP suffer a growth disadvantage during competition experiments (i.e., repeated rounds of glucose starvation and recovery) support this idea (8). Alternatively, growth phase regulation of SprE might be necessary if SprE plays a role, as yet to be identified, during stationary phase.