Stationary-Phase Gene Regulation in Escherichia coli?

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Stationary-Phase Gene Regulation in Escherichia coli^?

doi: 10.1128/ecosal.5.6.3

Regine Hengge

[Section Editor: Steve Finkel]

Posted 16 December, 2011

Institut für Biologie—Mikrobiologie, Freie Universität Berlin, 14195 Berlin, Germany.

Mailing address: Institut für Biologie-Mikrobiologie, Freie Universität Berlin, Königin-Luise-Str. 12-16, 14195 Berlin, Germany. Phone: (+49) 30–838-53119, Fax: (+49) 30–838-53118, E-mail: This e-mail address is being protected from spambots. You need JavaScript enabled to view it

^§This review is an updated version of a book chapter: Hengge, R. 2011. The general stress reponse in gram-negative bacteria, p. 251–289. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses, 2nd ed. ASM Press, Washington, DC.

INTRODUCTION

In their natural environments, bacteria usually have to cope with rather stressful conditions. These include the limitation of diverse nutrients and the intense competition for resources, which often do not support more than maintenance and survival in the stationary phase rather than growth (192, 251, 358), with maintenance of cell mass and numbers at the population level being achieved by "cryptic" growth of cells on the debris of others (139). In addition, bacteria have to adapt to a variety of abiotic stresses, such as nonoptimal temperature, pH, or osmolarity, oxygen-derived radicals, toxic metal ions and radiation, and predation by bacteriophages, other bacteria, or protozoa, as well. Bacteria use a combination of strategies to counteract these potentially life-threatening assaults. One strategy is to induce highly stress-specific responses that can eliminate the stress agent (e.g., reactive oxygen species) and repair damage that has occurred at the DNA, proteins, and membranes (460). On the other hand, many different stress conditions as well as entry into stationary phase also induce the complex general stress response, which renders bacteria resistant to diverse stress conditions, such that damage is prevented rather than has to be repaired (190, 194, 195). This includes "cross-protection," meaning that cells exposed to one stress also become resistant to other stresses not yet experienced (224, 225, 267, 318). In addition, the general stress response not only serves as a fine-tuned, long-term adaptation to adverse conditions that support only slow or even no growth, but also provides an emergency reaction to diverse stresses that can be rapidly activated in parallel to specific stress responses (193). When the specific and more limited responses turn out to be sufficient to cope with the actual problem, the general stress response is turned off again, as has been shown, for instance, for the classical diauxic shift (140, 332, 482).

The molecular processes underlying stationary-phase gene regulation and the general stress response have been intensively studied in two model organisms, Escherichia coli and Bacillus subtilis. With respect to multiple signal inputs and physiological outputs, these responses are similar in gram-negative and gram-positive bacteria. In addition, the use of alternative sigma subunits of RNA polymerase (RNAP) as master regulators is a common feature (460). However, the actual control circuitry used for signal transduction and gene regulation is radically different. In most gram positives the master regulator is σ^B, an alternative sigma subunit of RNAP that is controlled by a partner-switching mechanism involving anti- and anti-anti-sigma factors (385). In gram-negative bacteria, regulatory mechanisms are more diverse. E. coli and Salmonella as well as other γ-proteobacteria use the sigma subunit σ^S (RpoS) as a master regulator of the stationary phase and general stress response, with σ^S being under complex transcriptional and posttranscriptional control. In β-proteobacteria, σ^S appears to have a similar function. Other gram-negative bacteria, however, seem to use different mechanisms, but, with the exception of α-proteobacteria, which rely on an anti-sigma/sigma factor system (143), hardly anything is known about their stationary-phase gene regulation and a general stress response.

PHYSIOLOGICAL CONTEXT OF THE GENERAL STRESS RESPONSE OF E. COLI

E. coli is an environmentally highly versatile bacterium that comes in a wide variety of strains that include commensals as well as important pathogens. As an enteric commensal or pathogen, it is an inhabitant of the mammalian intestine, but certain pathogenic strains are specialized to living in other locations in their hosts (240). Moreover, E. coli thrives in freshwater and marine aquatic environments as well as in the soil, where it is exposed to a wide variety of abiotic and biotic stresses (10, 101, 105, 344). It is also accustomed to a feast-or-famine existence, because it can efficiently transit between rapid-growth phases, slow growth, and stationary phase depending on the availability of resources (194, 251). Thus, E. coli is a true generalist, with its stationary phase and general stress response being crucial for adaptation to these very different and variable environmental conditions.

From the Exponential to the Stationary Phase of Growth

The stationary-phase response (267) is tightly interconnected with the other phases of the E. coli growth cycle and the corresponding regulatory circuits (Fig. 1). Key players in these circuits are at least five sigma subunits of RNAP (σ⁷⁰, σ^FliA, σ^S, σ^E, σ⁵⁴) (84, 95, 114, 170, 195, 196), the flagellar master regulator FlhDC (450, 511), as well as the second messengers cAMP, (p)ppGpp, and c-di-GMP (50, 301, 373, 382). The interplay of all these factors is crucial for metabolic adaptation to available resources, the control of motility, cell morphology, stress resistances, and also biofilm functions during the exponential, postexponential, and stationary phases of the growth cycle. While these regulatory factors control the expression of distinct genes, they also influence each other and operate in the molecular environment of a dynamic nucleoid, in which accessibility and supercoiling of the DNA is modulated as cellular levels of nucleoid-associated proteins (such as Fis, HU, IHF, and Dps) change during the transition from exponential to stationary phase (15). These global and local layers of control are tightly interwoven and can even homeostatically compensate for each other (41).

Fig. 1Growth phases and corresponding master regulators in E. coli K-12. Using the E. coli K-12 strain W3110 growing in complex Luria-Bertani medium (LB) as a model system, three growth phases are shown: (I) exponential or “log“ phase, (II) postexponential phase, and (III) stationary phase. In the first part of the postexponential phase (starting at an OD of approximately 0.3), the flagellar master regulator FlhDC, the flagellar sigma factor sF (s28 or sFliA) and therefore flagella are expressed, but later on further expression of FlhDC is shut down and existing FlhDC is degraded. As a consequence, the expression of sF (and other proteins under FlhDC control) ceases, excess sF (not bound by its anti-sigma FlgM) is degraded, and synthesis of flagella comes to an end. However, assembled flagella are active and cells are highly motile. In parallel, the master regulator of the stationary phase, sS, begins to accumulate but initially is only inefficiently assembling into active RNAP holoenzyme (EsS). During transition into stationary phase, EsS-dependent gene expression is strongly stimulated. Note that only relative amounts of the various regulatory proteins or complexes are indicated that cannot be compared directly. The dynamics of the total cellular levels of sS and sF was measured as described in references 24 and 272, respectively, and the relative amounts of active regulators (FlhDC, EsF and EsS) reflect an average of the expression patterns of many of their respective direct target genes. OD(578 nm), optical density of the culture measured at 578 nm; ON, overnight, i.e., approximately 24 h. For further details and references, see main text. This figure is a modified version of a figure published in The Second Messenger Cyclic-Di-GMP (ASM Press) (189) and is used here with permission.

From the exponential to the postexponential phase of the growth cycle

Cells that grow exponentially in complex medium (the classical Luria-Bertani or LB medium; state I in Fig. 1) devote most of their resources to the synthesis of ribosomes, i.e., the machinery necessary for driving rapid growth and proliferation. This is reflected by rather condensed nucleoids and the concentration of RNAP in a few cellular foci at the ribosomal gene clusters (226). When carbon/energy sources become less optimal, cells enter into the postexponential growth phase. In a LB-grown E. coli K-12 culture, this happens at an optical density of approximately 0.3 (Fig. 1; optical density determined at 578 nm) (436). Physiologically, this phase of the growth cycle (state II in Fig. 1) appears to correspond to the "hungry" state of bacteria growing under carbon source limitation (134, 483). Now, cAMP begins to accumulate and, via cAMP receptor protein (CRP) as its effector protein, activates numerous genes, many of which are involved in alternative carbon source scavenging (134, 205, 291). In parallel, the global regulator Lrp modulates the expression of many genes (483). Overall, gene expression diversifies, the nucleoid becomes less compact, and RNAP becomes more dispersed (226, 291). One of the many operons activated by cAMP-CRP encodes the master regulator for flagella expression, FlhDC (450). Flagellar structural proteins and secondary regulators (including the flagellar sigma factor σ^FliA and its anti-sigma factor FlgM) are expressed in a fine-tuned temporal order (23, 84, 239). This process culminates in the assembly of several flagella per cell and entry into the motile "foraging" phase of the growth cycle (291, 540).

In parallel to the reduced growth rate and as a consequence of nutrient limitation, RelA/SpoT-synthesized (p)ppGpp begins to accumulate, which in a gradually fine-tuned manner results in reduced ribosomal gene expression and increased availability of vegetative RNAP for alternative gene expression (56, 301, 382, 483), including that of rpoS (153, 203, 270). However, σ^S does not yet seem to gain significant access to RNAP core , because the large majority of σ^S-dependent genes are still not expressed. Rather, the sigma subunits that determine overall gene expression during the postexponential phase are the vegetative σ⁷⁰, the flagellar sigma factor σ^FliA (also known as σ²⁸), which is specialized in expressing flagellar class 3 genes, and σ⁵⁴, which controls genes involved in nitrogen source scavenging as well as flagellar genes (114, 148, 158, 359, 540). Moreover, the regulatory factor FliZ, which is expressed under the control of FlhDC and thus belongs to the flagellar gene hierarchy, interferes with the expression of many σ^S-dependent genes by binding to and blocking their −10 promoter regions. FliZ thereby gives priority to FlhDC-driven motility during the initial phase of σ^S accumulation (374, 377).

Transition into stationary phase

This picture changes at a distinct point of the growth cycle that corresponds to an optical density of approximately 2 in a LB-grown E. coli culture (Fig. 1; optical density determined at 578 nm). At this point, σ^S is approaching the level typical for stationary-phase cells (268) and accessory factors such as Crl (54, 125, 333, 384, 492), the anti-σ⁷⁰ factor Rsd (228, 229, 328, 380), and (p)ppGpp (230, 483) influence sigma factor competition for RNAP core in favor of σ^S-containing RNAP holoenzyme (Eσ^S) formation. This results in the activation of σ^S-dependent genes in a fine-tuned temporal order that probably reflects the affinity of their promoters for Eσ^S (374). Because of RNAP core limitation (168, 169, 300), the cellular levels of vegetative and flagellar RNAP holoenzymes are reduced in parallel, and this process may actually be further enhanced by the equally (p)ppGpp-driven induction of the stress sigma subunit σ^E (95).

As a consequence of the now reduced levels of vegetative RNAP holoenzyme, the expression of various nutrient-scavenging systems is diminished (355, 356) and, furthermore, supported by also reduced σ⁵⁴-containing RNAP levels, the expression of the flagellar master regulator FlhDC comes to an end (23, 114, 374). The existing FlhDC protein complex is degraded by ClpXP (374, 477, 475). Together with FlgM-dependent inactivation (84, 87) and Lon-mediated degradation of excess flagellar sigma factor σ^FliA not bound by FlgM (23), this results in a cessation of flagellar gene expression (374). Moreover, cells begin to reduce their swimming speed, i.e., modify the activity of their flagella by a c-di-GMP-dependent process, and finally become nonmotile later in the stationary phase, which probably also reflects energy limitation (3, 9, 43, 374).

In parallel, σ^S-driven gene expression results in (i) morphological changes (cells become shorter and ovoid rather than rod-shaped) (266, 424, 426), (ii) pronounced metabolic alterations (toward a more anaerobic/fermentative energy metabolism) (371, 519), (iii) membrane alterations (234, 509), (iv) the development of multiple and strong stress resistances (267, 318), and (v) cellular adhesion via curli fimbriae, which prepares the cells for biofilm formation (180, 363, 388) (Fig. 2). Thus, the foraging strategy directed toward the search and optimal use of remaining carbon and energy sources for growth is replaced by a strategy that invests the remaining resources in maintenance and survival (state III in Fig. 1). It should be noted, however, that the term "stationary phase " refers to a cell population with a more or less constant amount of cells or total cell mass, which does not exclude slow "cryptic" growth of some cells on the debris of others (139, 251). In addition, during the transition from the postexponential to the stationary phase, cultures may also be heterogeneous with subpopulations either still being in the foraging state or already in the stationary-phase state.

Fig. 2Signal input and phenotypic output of the general stress response. Stress conditions that result in cellular σ^S accumulation and phenotypic alterations induced by high σ^S levels are depicted.

Induction of the General Stress Response by Diverse Stress Conditions

During transition into stationary-phase induction of σ^S and the general stress response as described above is a precisely fine-tuned process that continuously monitors decreasing nutrient availability (195, 267, 268, 483) and probably also cell density (268, 529). However, when rapidly growing cells are suddenly exposed to potentially life-threatening stress conditions, "emergency shortcuts" can rapidly and strongly induce the general stress response. Thereby, exponentially growing E. coli cells can accumulate σ^S and activate σ^S-dependent gene expression in response to sudden starvation for a specific nutrient source (268, 308), upon shifts to high osmolarity (193, 337) or low pH (28, 199), or in response to heat shock (340) or DNA damage (322). As a consequence, not only do stationary-phase cells survive diverse harsh stress conditions that would be lethal to growing cells, but this multiple stress resistance is also induced by other stressful treatments that result in σ^S accumulation and therefore generate "cross-protection" (224, 225, 267, 318) (Fig. 2).

In addition, σ^S is present at higher levels in cells that grow at reduced temperature, i.e., below 30°C (396, 443), but at a temperature range above the typical cold shock conditions. In contrast to the rapid emergency reactions mentioned above, σ^S induction upon the shift to lower temperature is a relatively slow process, probably reflecting the fact that reducing the temperature is not immediately life-threatening to the bacteria.

THE σ^S SUBUNIT OF RNA POLYMERASE AS THE MASTER REGULATOR OF THE GENERAL STRESS RESPONSE

From Discovery to Our Current View of σ^S

The discovery of σ^S (RpoS) as a master regulator for stationary-phase gene expression (267) was based on the observations that σ^S itself is induced during entry into stationary phase, and that an rpoS mutant was pleiotropically defective for typically stationary phase-associated functions, such as thermotolerance, resistance against high concentrations of salt or hydrogen peroxide, and glycogen production. Moreover, in this study (267), it was recognized that rpoS and several genes independently described as regulatory gene loci for a catalase (292), an acidic phosphatase (478), and exonuclease III (422, 489), in fact represented a single gene (which was not completely trivial at a time when no bacterial genome sequences were available and PCR was just slowly making its way into laboratory routine). On the basis of its sequence similarity with the vegetative sigma factor σ⁷⁰ (RpoD), this gene was suggested to encode a sigma factor (342), which was experimentally confirmed later (349, 470). The names "rpoS" and "σ^S " were proposed because of the function in stationary phase and stress responses (267).

Basic structural properties of σ^S

In vitro, σ^S has a lower affinity for RNAP core (E) than all other sigma subunits in E. coli (349, 470). Yet, in vivo its efficiency in competing for RNAP core and, therefore, its in vivo activity is positively modulated by several accessory components (see below). Among the sigma factors of E. coli, σ^S is the closest relative of the σ⁷⁰. It is a group 2 sigma factor featuring all four regions typically conserved in σ⁷⁰-like sigmas (for the classification and general principles of structure and function of sigma factors, see reference 184) . With its regions 2.4 and 4.2 being particularly similar to the corresponding regions of σ⁷⁰, the two sigmas in fact recognize similar −35 and −10 promoter regions when present within the RNAP holoenzyme (see below for details).

However, minor but significant differences in the molecular structure and function of the two holoenzymes contribute to their control of separate regulons and complementary physiological functions. Unlike Eσ⁷⁰, which prefers negatively supercoiled promoter DNA, Eσ^S can also activate gene expression from more relaxed DNA (48, 91, 258). This is in line with overall chromosomal supercoiling being reduced in stationary phase (41, 258). At the molecular level, this may have to do with a difference between the two sigmas in their interaction with the β-flap domain of RNAP (259). Moreover, potassium glutamate (which accumulates in hyperosmotically shifted cells) and possibly acetate seem to play important roles in σ^S-dependent gene expression. Again in contrast to Eσ⁷⁰, the activity of Eσ^S is positively modulated by these molecules by a mechanism that involves the C-terminal region of σ^S (106, 165, 242, 285, 360, 406, 408). This region is a 16-amino-acid extension not present in σ⁷⁰ that may be involved in the interactions with the core β-flap domain and with activator proteins bound adjacently to the DNA (259, 494, 501). Recently, this C-terminal region and the σ^S-β-flap interaction have been implicated in the formation of a "poised" state of promoter-bound Eσ^S from which Eσ^S can productively escape in response to certain stress conditions that result in DNA relaxation and/or accumulation of potassium glutamate (166, 208, 407).

Specific Promoter Recognition by σ^S-Containing RNAP

The σ^S promoter consensus sequence and the "σ^S promoter selectivity paradox"

Initial sequence comparisons of σ^S-controlled promoters left researchers puzzled with the finding that Eσ^S obviously recognizes promoter regions hardly distinguishable from those recognized by the vegetative Eσ⁷⁰ (194, 293, 469). Moreover, both holoenzymes were observed to transcribe the same DNA templates in vitro (349, 470), and in vitro selection of a sequence optimally bound by Eσ^S revealed −35 and −10 hexamers basically identical to the σ⁷⁰ consensus promoter (TTGACA and TATACT, respectively) (152). Nevertheless, Eσ^S specifically and efficiently activates a large regulon of its own and plays a physiological role complementary to that of Eσ⁷⁰, which seemed completely at odds with the simple concept of each sigma recognizing its own specific consensus promoter. The solution of this "σ^S promoter selectivity paradox " has not only contributed to understanding the general stress response at the molecular level, but has also refined our concepts of function and evolution of sigma factors in general (reviewed in references 197 and 493).

Unlike the in vitro SELEX experiment that selected just for optimal Eσ^S binding (152), natural σ^S-controlled promoters have been selected during evolution for a complex combination of properties: (i) an optimal balance of Eσ^S binding, open complex formation, and promoter clearance ; (ii) a distinct degree of σ^S selectivity; and (iii) a certain expression level, depending on the physiological roles of the respective genes. In vivo many genes are exclusively σ^S-dependent, but there are also promoters that can be activated by both holoenzymes such that Eσ⁷⁰ is responsible for expression during growth, and Eσ^S "takes over " during entry into the stationary phase (365, 392, 508). Physiological expression levels may be very different for different genes, but, as a general rule, the activities of genes in slowly growing or nongrowing cells do not have to be as strong as for genes expressed during rapid exponential growth.

Overall, natural selection of σ^S-selective promoters thus follows a complex mixed strategy that relies upon a series of special properties of Eσ^S and its interactions with promoter sequences. As a consequence, σ^S-dependent promoters have modular structures with each promoter using a specific combination of these properties. Features that can contribute to σ^S selectivity of a promoter are summarized in the following (for a more detailed account, the reader is referred to reference 493).

Eσ^S is more tolerant than Eσ⁷⁰ with respect to deviations from the "optimal" consensus promoter

Eσ^S-mediated transcriptional activity is less compromized by partially degenerate −35 regions (49, 152, 261, 284, 491, 524). Moreover, Eσ^S activity is also less affected when the distance between the −35 and −10 regions is one or even two nucleotides longer or shorter than the optimal 17-bp spacer (491). Because it is less deleterious to transcription driven by Eσ^S than by Eσ⁷⁰, this "deviation from the optimum" strategy can increase σ^S selectivity of a promoter, but comes at the price of reduced absolute promoter activity. With a promoter that has to function in nongrowing cells, this may not necessarily be a disadvantage. Yet, in cases where a particular promoter has to be relatively strong, it can also be compensated by non-sigma-discriminatory activity-enhancing elements such as AT richness in certain positions of the spacer region and/or a TGTG(−17 to −14) extended −10 element (24, 493). Recently, Eσ⁷⁰ was also found to have a preference for a T at the −18 position in the spacer region, which is due to a specific contact via its R451 residue, while Eσ^S is indifferent to the −18 nucleotide (446). Because of its reduced stringency with respect to the −35 sequence and spacer length and sequence, Eσ^S can also use more than one, overlapping −35 elements in some promoters, which offers the additional advantage of probably making these promoters more resistant against changes in local or global DNA superhelicity (491). In structural terms, the increased flexibility of Eσ^S with respect to the −35 position is probably related to the domains 4 of σ^S and σ⁷⁰ interacting differently with the β-flap domain of the RNAP core enzyme (259, 491).

Eσ^S makes additional contacts to DNA sites outside the core −10 hexamer in a manner distinct from Eσ⁷⁰

A hallmark of σ^S-controlled promoters is the presence of an extremely conserved C at position −13, often also preceded by a T(−14) (29, 152, 519). These two bases, which are located immediately upstream of the core −10 element, are specifically contacted by a lysine residue in σ^S (K173). By contrast, σ⁷⁰ has a glutamate at the corresponding amino acid position (E458), and operates best with a G at the −13 promoter position (29). This difference is reflected by slight differences in footprints obtained with both holoenzymes in this region (348). Besides increasing σ^S selectivity, the presence of C(−13) also enhances absolute promoter strength with Eσ^S in comparison with all three other bases at this position. T(−14), however, provides higher σ^S selectivity, but lower absolute activity than a G(−14) (which is part of the standard extended −10 element) (29). It also seems that C(−13) requires the natural supercoiling of the DNA in order to exert its positive effect on Eσ^S-mediated transcription initiation. Thus, σ^S-dependent promoters identified by analyzing runoff transcripts on microarrays ( ROMA), which were obtained with Eσ^S and linearized chromosomal DNA fragments in vitro, represented a smaller subset of the σ^S regulon in which promoters with a C(−13) were strongly underrepresented (301).

Moreover, σ^S-dependent promoters are often A/T-rich (often TAA) in the "discriminator" region immediately downstream of the −10 hexamer (284, 519), which is in striking contrast to the G/C-rich stringently controlled promoters that are inactivated during entry into stationary phase (181, 232). This A/T-rich element stimulates Eσ^S-mediated transcription (361, 390), and probably optimizes promoter melting at these promoters, because Eσ^S binds untwisted but not melted promoters more weakly than Eσ⁷⁰ (284). Taking together all these promoter elements, the consensus sequence for a σ^S-dependent promoter between positions −14 and −4 is in fact the endecamer TCTATACTTAA (with the underlined positions showing particularly strong conservation) (519). Promoters without any recognizable −35 element, in which this consensus endecamer is (almost) completely present, can have reasonable activity, exhibit complete σ^S dependence, and are therefore activated exclusively during entry into the stationary phase (91, 202, 374, 518, 519). Because of the expansion of the −10 element, this type of "compact" σ^S-dependent promoter also has an information content similar to that of standard promoters with typical −35 and −10 hexamers (493).

Eσ^S uses UP-elements in a manner distinct from Eσ⁷⁰

There is another class of σ^S-dependent promoters that do show recognizable (though usually not perfect) −35 regions, but, despite this similarity to vegetative promoters, they are σ^S-selective. These promoters operate in conjunction with a specific UP-element configuration (490). The UP element, which was initially identified in strongly transcribed ribosomal promoters, is an AT-rich segment located approximately between positions −56 and −40. It is composed of slightly asymmetric distal and proximal half-sites, which are contacted each by one C-terminal domain of the α-subunit of RNAP (α-CTD) (126, 163, 409). A distal UP-element half-site alone strongly stimulates Eσ^S-mediated expression, but interferes with Eσ⁷⁰ activity, whereas either a proximal half-site or a full UP-element strongly favors Eσ⁷⁰-dependent expression. Eσ^S-mediated promoter activation by a distal half-site obligatorily requires the presence of a −35 element, i.e., fixation of domain 4 of σ^S at this position of the promoter (490). The striking discrimination between the two sigma factors is probably based on the failure of σ^S to specifically interact with the α-CTD positioned in immediate proximity when bound to a proximal half-site. In σ⁷⁰, this interaction uses a positively charged surface area of domain 4 (involving K593, H600, and R603, which in σ^S are substituted by E308, E315, and Q318) and is strong enough to position one α-CTD next to σ⁷⁰ domain 4 even in the absence of a proximal UP-element subsite (163, 493). Time-resolved footprinting experiments suggest that at a promoter with a distal UP-element half-site only (which interacts with the other α-CTD), upstream DNA "traps" Eσ⁷⁰ by wrapping around it, whereas this does not happen to Eσ^S (493). In conclusion, a second type of σ^S-selective promoter (besides the compact "expanded −10 only " promoter) features a combination of a distal UP-element half-site, a −35 region, and at least a core −10 hexamer. In addition, most promoters of this type usually also include a C(−13).

Eσ^S may be less "trapped" by −10 element-like pausing sites during early elongation of transcription

Early transcription elongation complexes can retain their sigma subunit, which can interact with −10 element-ressembling sequences often located downstream of transcriptional start sites that cause temporary transcriptional pausing (58, 102, 350, 399). Such −10 element-like putative pausing sites also occur downstream of σ^S-controlled promoters, and preliminary data indicate that transcription by Eσ^S may be less affected than by Eσ⁷⁰ at these sites (493). In addition, σ^S seems more prone to leave the early elongation complex than σ⁷⁰ (392), which not only increases the cellular pool of free σ^S available for sigma factor competition for RNAP core, but can make Eσ^S-mediated transcription less responsive to −10-like pausing signals at promoters where both RNAP holoenzymes can initiate transcription. Thus, early transcription elongation events may contribute to σ^S selectivity of a promoter.

Cooperation of σ^S-Containing RNAP with Transcription Factors

In contrast to "simple" alternative RNAP holoenzymes, as, for instance, Eσ³², which is responsible for the heat shock response (289), Eσ^S resembles the vegetative Eσ⁷⁰ because its activity can be modulated by additional regulatory proteins that bind to promoter regions. As a consequence, expression of σ^S-dependent genes does not necessarily just follow σ^S accumulation in the cell. Rather, subsets of the σ^S regulon require additional signal input. Thus, among the 481 genes that showed reduced expression in a rpoS mutant in a microarray study, only a core set of 140 genes were detected under all three different conditions that result in σ^S accumulation (519). The remaining genes exhibited σ^S-dependent expression under one or two of these conditions only, which suggests an involvement of additional regulators. Such coregulation can be a global phenomenon as suggested by the strikingly high number of σ^S-dependent genes (approximately 50%) that exhibit known or putative binding sites for cAMP-CRP in their control regions, which can occur at typically activating , repressing, or noncanonical positions (519).

"Classical" transcription factors

Because modulation by a specific transcription factor can be different for the two holoenzymes at a given promoter, these accessory regulators can also contribute to σ^S selectivity. At class II promoters, differential activation (e.g., by cAMP-CRP, Fis, XylS, or λcI) is based on differences in activator-sigma interaction, because σ^S and σ⁷⁰ expose oppositely charged surface areas to the adjacently bound activators (249, 351, 423, 494). However, differential activation (e.g., by cAMP-CRP) also occurs at promoters, where the activator binds further upstream at a class I position and interacts with α-CTD. Here, the two holoenzymes prefer different locations of the activator, as shown by shifting the CRP binding site to a series of typical and nontypical class I positions upstream of the csiD promoter (156). Another interesting, but not yet understood, case is the csgD promoter, which absolutely requires the MerR-like regulator MlrA as well as Eσ^S for activation (66). Because MerR-like activators act by distorting the promoter DNA and thereby compensate for nonoptimal −35/−10 spacing (in σ⁷⁰-dependent promoters) (65), MlrA may also exert a topological effect on the csgD promoter. Also, the product of this gene, CsgD, is a regulatory protein that obviously can cooperate both with Eσ^S and Eσ⁷⁰, and thereby activates the two target promoters yaiC_p and csgB_p, respectively (518).

Abundant histone-like proteins

In addition to "classical" regulators, major nucleoid-associated or "histone-like" proteins can differentially affect promoter activation by Eσ^S and Eσ⁷⁰ and thereby contribute to σ^S selectivity. The regulons controlled by the leucine-responsive regulatory protein, Lrp, and σ^S show a large overlap (471), and, in an lrp mutant background, numerous genes exhibit altered ratios of σ^S-dependence (519). Similarly, integration host factor, IHF, modulates the expression of many σ^S-controlled genes (310). Depending on the specific promoter, activation or repression by these factors is more strongly imposed upon Eσ^S or Eσ⁷⁰ and thus improves the selectivity for one or the other holoenzyme (57, 90, 264, 519).

The global repressor/silencer protein H-NS, which can oligomerize on DNA (117, 127), plays a modulatory role in the expression of many σ^S-dependent genes by selectively interfering with Eσ⁷⁰-mediated activation (12, 26, 400, 438, 515, 518). For instance, during log-phase growth, H-NS, together with Fis protein, strongly inhibits the dps promoter by trapping and displacing Eσ⁷⁰. In stationary-phase cells, however, Fis is no longer present and H-NS does not interfere with Eσ^S-mediated expression of dps (164). Alternatively, the log-phase inhibition of Eσ⁷⁰-mediated expression of dps can be overcome by the activator OxyR, which responds to hydrogen peroxide stress (8). Also at the hdeAB promoter, H-NS can generate a complex nucleoprotein structure in which Eσ⁷⁰ becomes trapped, but not Eσ^S (438). However, Eσ^S does not always need to act directly on the H-NS-repressed promoter. Thus, at the gadA promoter, interference of H-NS with Eσ⁷⁰-dependent expression is relieved by the regulator GadX, which in turn is expressed under σ^S control, thus making gadA expression indirectly dependent on σ^S (157).

In apparently rare cases, H-NS can exert a direct positive effect on σ^S-dependent promoters. H-NS binds to the csiD promoter region, where it acts as a coactivator for the major activator cAMP-CRP. Here, H-NS seems to reduce torsional flexibility of the local DNA because it increases stringency with respect to the optimal position for cAMP-CRP (in an hns mutant, cAMP-CRP activates less well from its optimal position, but can activate to some extent from positions that are unproductive in the presence of H-NS). Thereby, H-NS also contributes to σ^S selectivity, which, in the csiD promoter, depends on the position of cAMP-CRP binding (156, 324).

CONTROL CIRCUITRY OF THE GENERAL STRESS RESPONSE: REGULATION OF σ^S EXPRESSION, PROTEOLYSIS, AND ACTIVITY

The primary requirement for inducing the general stress response in E. coli is σ^S accumulation, with expression patterns of many σ^S-dependent genes then being fine-tuned by additional signal input. Because σ^S accumulation can be triggered by a plethora of different stress conditions, it is not surprising that regulation, and therefore signal integration, occurs at all possible levels, with different stress conditions having an impact at different levels of control. The regulatory mechanisms affected include rpoS transcription, rpoS mRNA turnover and translation, as well as σ^S proteolysis (Fig. 3; reviewed in reference 196) . Moreover, the efficiency of σ^S in sigma factor competition for RNAP core, and therefore σ^S activity, is subject to regulation (reviewed in reference 493).

Fig. 3Diverse stress conditions affect different levels of σ^S control. Regulation of σ^S occurs at the levels of transcription, translation, proteolysis, and association with RNAP core enzyme. Stress conditions affecting this regulation are indicated as well as regulatory proteins, small RNAs, and second messengers that participate in the underlying regulatory mechanisms (see text for details).

The highly fine-tuned gradual accumulation of σ^S during the transition from exponential to postexponential and finally to stationary phase mostly relies on the control of σ^S expression and combines mechanisms of transcriptional and translational regulation (268, 270). However, rapid "emergency" accumulation of σ^S in response to potentially life-threatening stress conditions is mainly dependent on a rapid inhibition of σ^S proteolysis (28, 53, 188, 222, 268, 308, 340). In addition, σ^S stabilization against proteolytic degradation also allows σ^S to accumulate further in starved cells, when expression rates for σ^S (as for most proteins) are reduced again (268, 535).

Regulation of rpoS Transcription

The rpoS promoter region

The rpoS gene is the second gene in an operon with nlpD, which codes for a lipoprotein that activates one of several cell wall hydrolases involved in cell division (269, 496). A bicistronic mRNA is expressed from two closely adjacent promoters upstream of nlpD, which provide basal expression levels apparently not further regulated (269). The major rpoS promoter (rpoS_p1) is located within the nlpD gene and generates a monocistronic mRNA with a 5′-untranslated region ( 5′-UTR) of 565 nucleotides (270, 468). The existence of additional start sites just upstream of the rpoS gene is uncertain because they have been observed by some researchers (468), but not by others (270). Transcription from rpoS_p1 is strongly activated in a gradual manner during the postexponential phase of the growth cycle, when cells are grown in complex medium (267, 268). In minimal medium, cells contain higher levels of transcript and σ^S protein already during their slower but exponential growth (267, 268, 308). When cells are grown in chemostats, an inverse correlation is observed between the growth rate and σ^S levels (which probably reflect rpoS transcription) (211, 472).

Regulatory factors involved in rpoS transcription

Two transcription factors directly involved in rpoS transcriptional control are the response regulator ArcA and cAMP-CRP. ArcA represses rpoS expression by binding to two sites in the rpoS_p1 region, one of which overlaps with a cAMP-CRP binding site located at a typical class I activator position, from which cAMP-CRP activates rpoS expression. Thus, ArcA seems to act as an antiactivator (325). Because ArcA is phosphorylated by the sensor kinase ArcB, which in turn is inactivated by oxidized quinones (154, 306), the observation that a high NADH-to-NAD⁺ ratio reduces rpoS transcription (435) may be linked to rpoS repression by ArcA.

cAMP-CRP plays a complex dual role that can be positive or negative depending on the actual conditions (267, 268, 317). By binding to its class I binding site (at -61.5) it contributes to increased transcription of rpoS during the late postexponential phase (a second site at +56.5 binds cAMP-CRP in vitro but does not affect regulation of rpoS in vivo) (325). However, mutants deficient for CRP or adenylate cyclase already have strongly increased rpoS expression during the exponential and early postexponential phases (267, 268); this negative effect of cAMP-CRP is indirect because it is independent of the presence of the cAMP-CRP binding sites in the rpoS_p1 region (325).

A physiologically very important regulatory input into rpoS expression is provided by the general stress alarmone (p)ppGpp, which inversely correlates with growth rate and accumulates in cells exposed to various kinds of starvation and other stresses (56, 301, 382, 483). (p)ppGpp-free relA spoT mutants of E. coli show strongly reduced σ^S levels under all conditions, but still exhibit an induction of σ^S during entry into the stationary phase, albeit at a lower absolute level and with a reduced factor of induction (153, 203, 270). At different growth rates, cellular levels of (p)ppGpp and σ^S closely correlate both during steady-state growth and upon sudden growth rate transitions (472). While (p)ppGpp seems to also contribute to stabilization of σ^S protein during entry into the stationary phase (230) (and see below), a major part of its positive effect on σ^S is due to a role in rpoS transcription, as shown with transcriptional rpoS::lacZ fusions. When in these fusion constructs the original rpoS_p1 was replaced by tac_p or lacUV5_p, expression was still clearly reduced in relA spoT mutants (albeit somewhat less than when rpoS_p1 was present) (203, 270). Thus, it appears that (p)ppGpp affects both transcription initiation at rpoS_p1 and elongation of the rpoS transcript, but the actual molecular mechanisms remain to be elucidated.

At least two additional factors seem to play a positive modulatory role in rpoS transcription during the postexponential phase. One is the BarA/UvrY two-component system, with the UvrY response regulator directly binding to the rpoS promoter region (186, 341). The other seems to be an unidentified signaling molecule that is excreted by the AcrAB/TolC secretion system and can affect the timing of rpoS expression during the postexponential phase (529). Thus, rpoS transcription may also be sensitive to cell density signaling, but details have not been reported.

Regulation of rpoS mRNA Turnover and Translation

Degradation of rpoS mRNA

With half-lifes in the range of several minutes, decay rates of rpoS mRNA are not significantly different from those of average E. coli mRNAs. Both RNase E and RNase III are important for rpoS mRNA degradation, with RNase E-mediated turnover being subject to regulation, whereas RNase III activity on rpoS mRNA seems unregulated (27, 320). During entry into the stationary phase, rpoS mRNA turnover even slightly accelerates, i.e., changes in rpoS mRNA stability do not seem to contribute to accumulation of σ^S under these conditions (27). Nevertheless, overproduction of the small RNAs DsrA and RprA, which directly base pair to rpoS mRNA and stimulate translation (see below), stabilizes rpoS mRNA, most likely by inducing higher ribosome coverage of the mRNA that protects against RNases (320). In wild-type strains, however, regulation of rpoS mRNA turnover seems to play a minor role in the regulation of rpoS.

Role of rare codons in the rpoS mRNA

The rpoS gene contains a significantly higher frequency of rare codons than the homologous rpoD gene that encode the vegetative σ⁷⁰, including 11 synonymous codons (encoding the same amino acid) at corresponding positions, which are rare codons in rpoS, but frequent codons in rpoD. Substituting these rare codons in rpoS by the more frequent synonymous rpoD codons resulted in lower levels of rpoS mRNA and σ^S protein because of more rapid RNase E-mediated degradation. This indicates that slowing down translational speed by ribosomal pausing at many rare codons along the rpoS transcript reduces ribosome spacing and thereby protects the transcript against ribonucleolytic attack by RNase E (250). This rare codon-based mRNA stabilization may also be important for other translationally controlled genes, where secondary structure formation in the translational initiation region and therefore inefficient ribosome binding may compete. Moreover, a striking difference in codon usage in pairs of homologous isoenzyme genes expressed in log phase and stationary phase indicates that this role of rare codons might be more general (250).

Role of rpoS mRNA secondary structure in translational initiation

In contrast to rpoS mRNA turnover, translational initiation from rpoS mRNA is a major target for regulation and signal input. This mRNA folds into a complex structure in which the translational initiation region (TIR, consisting of the ribosome-binding or Shine-Dalgarno site and the initiation codon) is base-paired to a region further upstream in the long 5′-UTR (64, 98, 275, 277, 302). Efficient translational initiation requires unfolding of this inhibitory structure and ribosome binding to the TIR. Strong and very rapid activation of rpoS translation has been shown in cells shifted to high osmolarity (271, 337) or low pH (28, 199). In addition, rpoS translation is stimulated beyond a distinct cell density in the postexponential phase (268) and in cells exposed to reduced temperatures (396, 443).

Role of the RNA chaperone Hfq

Unlike the activation of rpoH mRNA translation, which is due to direct thermal melting of a similar inhibitory mRNA secondary structure (289), accessory factors are required to stimulate rpoS mRNA translation. A major prerequisite is the RNA-binding protein Hfq (63, 338, 444). Hfq was originally identified as a host factor for replication of the RNA phage Qβ (144), but the phenotypes of a hfq mutant suggested that it was a pleiotropic regulator (485). Its requirement for efficient rpoS translation was the first molecular role observed in non-phage-infected E. coli cells (338), but it was rapidly recognized that rpoS was not the only target (336, 507, 536). Hfq forms ring-like hexameric structures that highly dynamically interact with RNAs (132, 345, 427, 431). By simultaneously binding mRNAs and partially complementary small regulatory RNAs (sRNAs) Hfq hexamers provide a platform that facilitates structural rearrangements that modulate mRNA translation and/or degradation of the RNAs involved (326, 362, 393, 499, 536).

Role of small regulatory RNAs in rpoS regulation

Four small regulatory RNAs, i.e., DsrA, RprA, ArcZ, and OxyS, have been found to affect rpoS translation. As rpoS control has become the paradigm of positive translational regulation by small regulatory RNAs, a series of excellent reviews have dealt in detail with the mechanisms involved (6, 161, 162, 276, 396, 463, 462). DsrA expression increases and stimulates rpoS mRNA translation at low temperature (302, 442, 443). RprA, especially when overproduced, can also stimulate rpoS translation (304, 303). It plays a minor role in the hyperosmotic induction of rpoS translation and accumulates during entry into the stationary phase, but its physiological role in rpoS control has not been fully clarified. ArcZ (or rather, a processed form of only 56 nucleotides) can also stimulate rpoS translation, is itself repressed by the ArcB/ArcA two-component system (which is activated by low-oxygen/high-energy supply conditions), and in turn downregulates arcB (309). Interestingly, the sensor kinase ArcB also downregulates rpoS transcription (via ArcA) and stimulates σ^S degradation (via RssB, see below) (325). Thus, the inverse coordination of ArcB/ArcA and ArcZ seems to establish a positive feedback control loop, which may contribute to robust switching between low σ^S and high σ^S states and provides a link between different levels of rpoS regulation.

DsrA, RprA , and ArcZ can all base-pair with the region in the 5′-UTR of rpoS mRNA that occludes the TIR and thereby promote accessibility of the TIR for ribosome entry (11, 275, 303, 497). In order to efficiently bind to and act upon rpoS mRNA, all three sRNAs require Hfq, but to different degrees, with DsrA being least dependent on Hfq, especially at 37°C. The stability of the sRNA/rpoS mRNA complexes, rather than the rate of complex formation, best predicts in vivo activities (448). All three sRNAs stimulate rpoS translation (320, 448), with structural rearrangements preceding ribosome binding, as shown for DsrA (503). At least when expressed from plasmids, DsrA and RprA also protect rpoS mRNA from degradation (320).

The fourth small RNA affecting rpoS regulation, OxyS, inhibits rpoS translation by a mechanism that does not seem to involve direct base-pairing with rpoS mRNA (537). OxyS expression (and therefore a reduction of rpoS translation) is stimulated by the OxyR regulator that responds to H₂O₂ stress (7). Interestingly, the same stress conditions also stabilize σ^S protein (322) (and see below), with the net result that σ^S levels are hardly changed (537), but are maintained in a less dynamic but probably more economic manner.

Role of cold shock proteins and other factors acting at the level of rpoS mRNA

At low temperatures, several cold shock proteins become important for the mRNA structural rearrangements required for rpoS translation. These are (i) the CsdA DEAD-box helicase that contributes to melting of the rpoS mRNA TIR, which allows Hfq-presented DsrA to base pair (397); (ii) CspC, which binds to and stabilizes rpoS transcripts and can partly compensate for a loss of Hfq (89); and (iii) CspE, which induces σ^S when overproduced from a plasmid (379).

Other components that control rpoS translation include the histone-like proteins HU (20) and H-NS (26, 527). HU is active as a dimer with α- and β-subunits being encoded by two highly similar genes, hupA and hupB, respectively (46). In rapidly growing cells, HU is present as α₂ -homodimers. Upon the induction of β-subunits during the postexponential phase, αβ-heterodimers are formed and occur in parallel to the α₂-homodimers (88). In particular , αβ-heterodimers efficiently bind to rpoS mRNA (20). Mutants that are deficient for H-NS exhibit highly activated rpoS translation (26, 527), but it is unknown whether H-NS directly interacts with rpoS mRNA in vivo or whether its effects are indirect. A potential regulation of DsrA or RprA expression by H-NS has been excluded (543). By contrast, inhibition of rpoS translation by the LysR-like transcription factor LeuO has been shown to occur by its repression of DsrA expression (245). Other factors that seem to positively modulate rpoS translation are (i) the LysR-like transcription factor LrhA (377) ; (ii) the SsrA (tmRNA) system, which releases stalled ribosomes from incomplete mRNAs and tags the corresponding truncated polypeptides for degradation (395) ; and (iii) the chaperone DnaK (340, 405, 404). Components that somehow downregulate rpoS translation include (i) Lon protease (at low temperature only) (395), (ii) the PTS component EIIA(Glc) (495), and (iii) high cellular levels of UDP-glucose, which may reflect good nutritional conditions (69).

For many of these factors, mechanisms of action have remained undefined. Moreover, potential small regulatory RNAs required for the drastic and rapid physiological activation upon hyperosmotic or acidic shifts have not been identified so far. Thus, it is clear that translational control of rpoS is a field in need of further research.

Regulation of σ^S Proteolysis

Although rapidly growing cells do express σ^S at a basal rate, it is hardly detectable because of efficient proteolytic degradation (with half-lifes in the 1- to 5-min range, depending on the actual growth conditions) (268, 468). Yet, σ^S is instantaneously stabilized in response to a variety of stress conditions. These include sudden starvation for carbon or phosphorus sources, the diauxic lag phase, shifts to hyperosmotic or acidic conditions, or the addition of hydrogen peroxide (28, 140, 193, 268, 308, 322, 337, 375). In addition, σ^S proteolysis is strongly reduced upon heat shock and during entry into the stationary phase in complex medium, but in a more gradual manner (268, 340, 535). The molecular mechanisms of σ^S proteolysis and its control by stress signals have recently been reviewed in detail (188).

Role of the proteolytic targeting factor RssB in ClpXP-mediated σ^S turnover

σ^S is degraded by the ATP-dependent complex ClpXP protease (434). However, ClpXP alone cannot bind to σ^S, but a specific recognition or targeting factor, the response regulator RssB (also termed SprE or MviA in Salmonella), is required and sufficient for delivery of σ^S to ClpXP (28, 339, 383, 542). RssB directly binds to σ^S with a 1:1 stoichiometry (30, 31, 246, 335, 544). Within σ^S, the first α-helix in domain 3, with its crucial amino acid K173, is essential and sufficient for this interaction (30, 466). Interestingly, K173 is also the amino acid in σ^S that makes the direct contact to the C(−13) position that is highly conserved in σ^S-dependent promoters (29) (and see above). By contrast, both domains of RssB are required to bind σ^S in vitro and to downregulate σ^S in vivo (246).

RssB consists of a N-terminal receiver domain (REC) linked to a C-terminal domain that seems distantly related to the PP2C-like Ser/Thr phosphatase domain (186) known from regulation by "partner-switching modules " in gram-positive bacteria (385). Phosphorylation of the REC domain (at D58) increases the affinity of RssB for σ^S (for a detailed discussion, see reference 188). Thus, in vitro binding and turnover of σ^S are stimulated by phosphorylation (30, 544). Interestingly, RssB^D58P or RssB^D58A mutant variants seem to support activities between those of nonphosphorylated and phosphorylated wild-type RssB (188, 246, 376). Because of the low cellular level of RssB (246, 390), RssB phosphorylation is required for efficient turnover of wild-type σ^S in vivo (52, 246, 335, 376). In vivo phosphodonors for RssB phosphorylation are acetyl phosphate (52) and the hybrid histidine sensor kinase ArcB (325). Residual σ^S degradation in acetyl phosphate-free and ArcB-deficient ackA pta arcB mutants (325) may be due to additional unidentified sensor kinases or to residual low-affinity interaction of unphosphorylated RssB with σ^S. RssB does not show autophosphatase activity nor reverse phospho-flow from RssB to ArcB in vitro (246, 325).

The interaction with RssB triggers a conformational opening of σ^S that exposes a binding site for the hexameric ClpX₆ ring, which is located close to the N terminus and, in the closed conformation, seems occluded by interaction with a region in the C-terminal half of σ^S (188, 466). In contrast to σ^S, RssB does not or only weakly interact with ClpX₆ in the absence of its substrate σ^S (246, 335, 544). Therefore, RssB should be considered a substrate-targeting factor rather than a protease adapter that initially acts upon the chaperone-protease complex and modulates the substrate-binding properties of the latter (188, 185). The binary RssB-σ^S complex then associates with ClpX₆, with the ClpP subunits either joining in then or being present already in association with the ClpX₆ rings. Using ATP hydrolysis, the ClpX₆ chaperone (a member of the AAA+ ATPase family) then actively and progressively unfolds σ^S and threads it into the proteolytic chamber formed by the double heptameric rings of ClpP₁₄, as observed for other Clp protease substrates (19, 254). During this process, RssB is not degraded (246, 544), and apparently not even dephosphorylated (188). This means that RssB acts catalytically in the degradation of σ^S, which is consistent with σ^S being present in vivo in an approximate 20-fold excess over RssB (31). However, when ClpXP protease is absent (in a clpP mutant) or is less active (apparently in starved cells that contain less ATP), RssB remains stably bound to σ^S. RssB then inhibits activity of σ^S, as it interferes with σ^S binding to RNAP core; when overproduced in stoichiometric amounts with σ^S, it can act like an anti-sigma factor (31, 246, 335, 542).

Role of ArcB-mediated RssB phosphorylation in the control of σ^S proteolysis

Proteolysis of σ^S is regulated by an astonishing multitude of different stress signals (Fig. 4; for details beyond the concise summary given here, see reference 188). Much of this regulation occurs at the step of substrate recognition, i.e., affects the interaction of σ^S and RssB in one way or another. This allows degradation of other ClpXP substrates to continue, while σ^S proteolysis becomes inhibited and σ^S accumulates. With RssB being a response regulator, its phosphorylation state seems an obvious target for regulation. However, phosphorylated RssB does not have detectable autophosphatase activity, it is not dephosphorylated during its catalytic cycle, and no specific phosphatases have been found (see above). As a consequence, control of RssB phosphorylation via the ArcB sensor kinase (see above) may mainly serve to activate newly synthesized RssB and therefore changes in ArcB activity may affect σ^S turnover rates only gradually. This seems to occur during entry into the stationary phase in complex medium or in other situations where the nutritional situation gradually deteriorates. Under these conditions, cellular levels of acetyl phosphate decline (247, 319). In addition, in an aerobic environment, reduced electron influx into the respiratory chain will generate oxidized quinones, which inactivate the ArcB sensor kinase (154, 306, 307). This will result in less phosphorylated RssB and ArcA, which in turn reduces σ^S proteolysis and relieves rpoS transcriptional repression, respectively, with both processes integrating into a gradual accumulation of σ^S (325).

Fig. 4Molecular mechanisms in the regulation of σ^S. The homeostatic σ^S/RssB/σ^S feedback cycle is shown as well as several competition, titration, and sequestration mechanisms that integrate numerous stress signals in the control of RssB/ClpXP-mediated σ^S proteolysis. Key features are the maintenance of a distinct σ^S:RssB ratio by the homeostatic feedback loop (up to a certain threshold where the rssAB promoter is maximally activated by Eσ^S) and the competition of RssB and RNAP core enzyme (E) for σ^S. For the detailed functions of all components and the regulatory consequences, see the main text. This figure is a modified version of a figure originally published in Current Opinion in Microbiology (222) and is used here with permission from Elsevier.

Role of the σ^S-RssB feedback cycle in the control of σ^S proteolysis

Interestingly, the slow and fine-tuned regulation of RssB phosphorylation does not contribute to the rapid "emergency" stabilization of σ^S triggered by many suddenly arising acute stress conditions (325, 376). Rather, sudden inhibition of σ^S proteolysis depends on rapid alterations of the ratio between cellular σ^S and RssB available for interaction. This requires that (i) RssB is the rate-limiting factor for σ^S degradation in vivo, which was experimentally demonstrated (390); and (ii) an appropriate σ^S:RssB ratio is maintained under nonstress conditions, i.e., during growth when σ^S is efficiently degraded. This ratio is approximately 20:1 (31) and is maintained by homeostatic feedback regulation, because the expression of RssB (from the rssAB operon) depends on σ^S (390, 412). Thus, σ^S activates the expression of its own "destruction factor" (Fig. 4). This homeostatic negative feedback allows maintenance of the same high σ^S degradation rates under conditions of changing σ^S expression rates (up to a threshold where the relatively weak rssA_p promoter becomes saturated) (18). In addition, this homeostatic feedback loop sets the stage for controlling the rate of σ^S proteolyis by rapidly acting competition, titration, and sequestration mechanisms that alter the ratio between σ^S and RssB available for interaction. Moreover, the more slowly operating homeostatic feedback loop can confer adaptation, i.e., can result in rapid σ^S stabilization being more or less transient (188).

Role of RssB sequestration for the control of σ^S proteolysis

Many mechanisms can affect the σ^S:RssB ratio, which provides a basis for multiple-signal integration in the control of σ^S degradation (Fig. 4). On the one hand, RssB can be titrated by sudden strong σ^S synthesis, which means that σ^S stabilization can actually be a passive consequence of increased σ^S expression; this happens when rpoS translation is stimulated by a shift to hyperosmotic or acidic conditions (199, 337). Furthermore, RssB can be sequestered or "trapped" in nonproductive complexes (Fig. 4). To date, three specific and directly acting RssB antagonists termed Ira proteins (for inhibitor of RssB activity) have been identified: (i) IraP, which is induced in a (p)ppGpp-dependent manner by phosphate starvation in E. coli and via the magnesium starvation-sensing PhoQ/P system in Salmonella (53, 55, 487); (ii) IraM, which is magnesium starvation-induced and regulated by a cascade of the EvgS/A and PhoQ/P two-component systems in E. coli (53, 121); and (iii) IraD, which is induced by DNA damage or hydrogen peroxide by mechanisms that are independent of the SOS and OxyR control systems (53, 322). In vitro, all three Ira proteins directly interact with RssB, IraM in addition also with σ^S. While not being related in primary sequence, the Ira proteins belong to the functionally defined group of small "connector" proteins that modulate the output of two-component systems in addition to the classical phospho-transfer signal transduction mechanism (329). RssB is thus an example of a response regulator, where rapid regulation of output activity may actually be dominated by connector proteins instead of the classical phosphotransfer pathway.

In addition, RssB can also be trapped in proteolytically inactive complexes with σ^S or even with σ^S and ClpXP protease (Fig. 4). This could happen when ClpXP is sequestered by alternative substrates, possibly during heat shock (340) or during starvation, when mistranslated or oxidized proteins accumulate (145); alternatively, ClpXP may still be available, but because of reduced ATP levels in carbon/energy-starved cells, may be unable to efficiently unfold σ^S (311). Inactivation of a fraction of the (now stable) σ^S by RssB and ClpXP has indeed been observed in starved cells (31, 542). Because of the excess of σ^S over RssB, only a fraction of σ^S can be "co-trapped" and therefore inactivated, and the large remainder would be stable and free to interact with RNAP core enzyme.

Role of Eσ^S holoenzyme formation in the control of σ^S proteolysis

Finally, RssB competes with RNAP core enzyme for σ^S binding (544) (Fig. 4). As a consequence, the cellular level of free σ^S available to interact with RssB, and thus the rate of σ^S degradation, is also modulated by processes that affect binding of σ^S to RNAP core enzyme. Therefore, Crl, a protein that binds to σ^S and stimulates its association with RNAP, has a stabilizing effect on σ^S (54, 492). The anti-σ⁷⁰ factor Rsd, which reduces the level of σ⁷⁰ available for competition with σ^S for RNAP core (228), could have a similar effect on σ^S turnover. Overall, a complex protein-protein interaction network connects the σ^S degradation and σ^S activity cycles, such that stress-induced changes in many of these interactions can indirectly affect σ^S proteolysis. Moreover, this network can exert several homeostatic functions: increased Eσ^S formation will be compensated for by increased RssB expression in the longer run; also, increased Eσ^S formation at the expense of Eσ⁷⁰ formation will result in decreased σ^S expression, since rpoS is transcribed from Eσ⁷⁰- controlled promoters.

Open questions in the regulation of σ^S proteolysis

In conclusion, we now begin to understand the molecular mechanisms by which σ^S is stabilized in response to a variety of stress conditions. However, this picture is probably still incomplete. There may be additional Ira-like proteins that sequester RssB under still unknown conditions (53). In addition, σ^S itself could be sequestered in a form that is stable and inactive at the same time, such that it is "saved" for later, e.g., in stationary phase, where it may be released but is no longer degraded (188). Also, the mechanisms that allow σ^S proteolysis to instantaneously resume, e.g., upon refeeding of starving cells (390), remain to be elucidated. Furthermore, a small excreted molecule (probably a peptide), which seems to be produced during the postexponential phase and somehow reduces σ^S turnover, has yet to be identified (204).

Finally, σ^S proteolysis seems connected to general RNA degradation in a nonunderstood manner. Thus, a deficiency in the mRNA poly-adenylating enzyme poly(A)-polymerase I (PcnB or PAPI), which is inversely growth-rate-regulated (220), enhances RssB-mediated turnover of σ^S specifically in starved cells (425). On the other hand, strong overproduction of RssB had a growth-reducing effect that was independent of the presence of σ^S, but that could be suppressed by eliminating PAPI. A rssB (ΔsprE) mutant also showed some reduction of intracellular poly(A) levels in exponentially growing cells, while PAPI levels were not affected (74). In stationary-phase cells of a rssB mutant, PAPI seems to localize to the membrane (74), and the composition of PAPI-associated proteins (many of which are part of the degradosome, but do not include RssB itself) may be slightly altered (75). Unfortunately, it was not tested whether these rssB mutant phenotypes were due to increased σ^S levels or not (i.e., rssB rpoS double mutants were not analyzed). Nevertheless, the authors speculate that RssB affects PAPI activity and therefore mRNA poly-adenylation and turnover by some pathway that is independent of the well-characterized activity of RssB in σ^S proteolysis (74, 75), but further work is clearly required here.

Regulation of σ^S Activity

Components affecting sigma factor competition for RNAP core enzyme

In order to become active, σ^S has to associate with the RNAP core enzyme, with this process of holoenzyme formation serving as yet another target for regulation. E. coli contains limiting amounts of RNAP core for which different sigma factors compete (168, 169, 214, 358). During exponential growth, the vegetative σ⁷⁰, which is present in excess over RNAP core, predominates (168, 227). In vitro, σ⁷⁰ has the highest affinity of all sigmas for RNAP core (300). Consequently, Eσ⁷⁰ is the major form of RNAP holoenzyme in rapidly growing cells, with about 70% of it being involved in transcribing rRNA operons (392).

During the transition into post-exponential and finally stationary phase or upon sudden exposure to stress, alternative sigma factors such as σ^S, σ^FliA, σ ³², and/or σ^E are induced and compete for RNAP core. In parallel, rRNA transcription is reduced, which releases free RNAP core available for alternative sigmas to use. A key regulator of these processes is (p)ppGpp, which (i) reduces rRNA transcription (56), (ii) activates the expression of σ^S (see above) and σ^E (95), (iii) may contribute to σ^S stabilization by inducing IraP (53), and (iv) affects sigma factor competition in favor of alternative sigmas (230, 272). In addition, the formation of alternative holoenzymes is promoted by the expression of Rsd, which sequesters a significant fraction of σ⁷⁰ in stationary-phase cells (213, 228, 380). Overproduction of Rsd mimics the effects of (p)ppGpp on holoenzyme switching (230, 272). Moreover, when cells enter into stationary phase, the small conserved 6S RNA sequesters a fraction of the Eσ⁷⁰ holoenzyme in an inactive form by mimicking an open promoter complex that is specifically bound by Eσ⁷⁰ only (513).

Role of Crl protein in Eσ^S formation and activity

While all these mechanisms promote alternative RNAP holoenzyme formation, they do not favor Eσ^S formation specifically. Nevertheless, Eσ^S transcribes more genes than any other RNAP holoenzyme during entry into the stationary phase (519) and even takes over some housekeeping duties from Eσ⁷⁰ (365, 392). Despite its induction and obvious performance, σ^S levels reach only about one-third of those of σ⁷⁰ (227) and the in vitro affinity of purified σ^S for RNAP core is the lowest of all sigmas in E. coli (300). Therefore, additional factors are required for σ^S to assume its role as a master regulator in stationary-phase gene expression. One such factor, Crl, was originally found to activate curli fimbriae expression (13), and then defined as a factor that stimulates σ^S activity at certain target genes (384). Crl binds to domain 2 of free and RNAP-associated σ^S (54, 125, 333) and stimulates the expression of a large set of σ^S-dependent genes independently of any specific promoter motif (492). In vivo, where σ^S has to compete with other sigmas for RNAP core, Crl is a dedicated σ^S-auxiliary factor, in particular, when σ^S levels are low (402, 403, 492). In vitro, Crl stimulates gene expression by Eσ^S more than by Eσ⁷⁰ (151), and promotes Eσ^S formation and activation under conditions where σ^S and σ⁷⁰ compete for RNAP core (492). It does so by forming a 1:1 complex with σ^S, which increases the association rate with RNAP core without changing the dissociation rate. Moreover, Crl can stimulate binding and open complex formation at certain promoters, when bound to the Eσ^S holoenzyme (125).

Role of K⁺ in stimulating σ^S activity

High intracellular levels of K⁺ (as found in a mutant devoid of enzyme IIA(Ntr) of the nitrogen PTS) increase σ^S-dependent gene expression without affecting σ^S levels. In vitro, K⁺ stimulates σ^S binding to RNAP core at the expense of Eσ⁷⁰ formation (278). Moreover, K⁺ also directly stimulates Eσ^S activity on certain promoters in a process that requires the C-terminal 16-amino-acid extension of σ^S not present in σ⁷⁰ (106, 165, 166, 208, 242, 285, 360, 406, 408). Since hyperosmotically shifted cells rapidly accumulate K⁺, the increased efficiency of formation and activity of Eσ^S may add to enhanced synthesis and stabilization of σ^S, thus explaining the very strong osmoregulation of many σ^S-dependent genes (519).

Role of FliZ protein as an antagonist of σ^S activity

Finally, FliZ was recently identified as a factor that negatively affects the expression of many σ^S-dependent genes (Fig. 5) (374). FliZ is transiently expressed during the postexponential phase from the FlhDC-activated operon fliAZY, which also specifies the flagellar sigma factor σ^FliA (σ²⁸). FliZ downregulates a spectrum of genes that resembles the regulon activated by Crl. Even slight overproduction of FliZ can strongly repress these genes. Mechanistically, FliZ acts as a global repressor that antagonizes σ^S by recognizing an operator sequence that resembles the extended −10 region of σ^S-dependent promoters. FliZ does so with a structural element that mimics the first α-helix (α3.0) in domain 3 of σ^S, which makes contact to the extended−promoter element, and, in fact, the C(−13) promoter nucleotide is of key importance for interaction with both Eσ^S and FliZ (377).

Fig. 5Inverse coordination of the motile and adhesive lifestyles in E.coli K-12. This coordination is part of the general transition from the foraging lifestyle of postexponentially and therefore slowly growing cells (with cAMP-CRP and Eσ⁷⁰ acting as master regulators; phase II in Fig. 1) to the stationary-phase lifestyle characterized by maintenance metabolism, stress resistance, high cell density, and cellular adherence (where Eσ^S acts as a master regulator; phase III in Fig. 1). The flagellar control cascade (FlhDC/FliA) interferes with the activity of the σ^S/CsgD/curli control cascade at two levels: (i) FliZ, which is expressed from a class 2 gene in the flagellar hierarchy, is an inhibitor of σ^S activity at many σ^S-dependent promoters, including those of ydaM, mlrA, and csgD; and (ii) the phosphodiesterase (PDE) YhjH, which is expressed from a class 3 gene (under σ^FliA control), degrades c-di-GMP and thereby keeps motility going, while not allowing the activation of the transcription of csgD and therefore curli expression. When the flagellar control cascade (including yhjH expression) shuts down in mid-postexponential phase, the diguanylate cyclases (DGCs) YegE and YedQ, which are increasingly expressed because of now accumulating σ^S, outbalance the PDE activity of YhjH, and c-di-GMP can accumulate. Via YcgR, c-di-GMP interferes with flagellar activity, and, via an unknown effector, stimulates csgD transcription. In essence, this c-di-GMP control module acts as a checkpoint that allows curli expression only after flagellar gene expression has been shut down. In parallel, a second DGC/PDE system, YdaM/YciR, is expressed under σ^S control. This system exclusively acts on csgD transcription in a way, which is not additive with the effect of the YegE/YhjH system (but both systems are essential for activation). An additional c-di-GMP control module operates downstream of CsgD expression and affects the expression of cellulose biosynthesis. The activity of all DGCs and PDEs (probably with the exception of YhjH, which basically consists of an EAL domain only) is likely to be modulated by additional unknown signals (bolts) perceived by their N-terminal sensor domains. Note that only relevant genes or proteins under FlhDC and σ^S control are shown here; overall, FlhDC and σ^S activate more than 60 and 500 genes, respectively. For further details and references, see the main text. This figure is a modified version of a figure that was previously published in The Second Messenger Cyclic-di-GMP (189), ASM Press.

FliZ is a highly abundant protein (>30,000 molecules per cell during the postexponential phase) and thus may be regarded as a histone-like protein (377). Physiologically, FliZ imposes a temporal delay on the activation of various σ^S-dependent genes (i.e., in fliZ mutants these genes are activated earlier during the transition into the stationary phase), which indicates that FliZ gives priority to the expression of flagellar genes and motility (over σ^S-dependent genes) during the postexponential phase of the growth cycle. However, this priority is transient, since FliZ levels decline during entry into the stationary phase, since its expression (like that of other FlhDC-controlled genes) already ceases in the second half of the postexponential phase (374).

THE σ^S-CONTROLLED DOWNSTREAM NETWORK: REGULATORY CASCADES, FEEDFORWARD AND FEEDBACK LOOPS

Regulatory Cascades and Feedforward Loops

By transcriptional profiling, more than 500 genes of E. coli (corresponding to roughly 10% of the genes in the genome) were found to be under positive control of σ^S (112, 262, 371, 519). Approximately 8 to 10% of these genes specify signal-transducing and/or DNA-binding regulatory proteins. This opens a wide field for indirect σ^S control, with σ^S being the top regulator in regulatory cascades. Moreover, many target genes are under direct as well as indirect σ^S regulation in feedforward control patterns. Regulatory cascades or feedforward arrangements allow for complex regulatory behavior such as additional signal input at lower levels, transient, delayed, or temporal ordering of gene expression, and noise filtering (for a discussion of different regulatory motifs, see reference 60). Transcriptional cascades in bacteria rarely seem to have more than three transcription factors acting in a row, which has been attributed to the need for rapid reaction because bacteria can rapidly proliferate (327, 437). However, recent research has shown that regulatory networks downstream of σ^S can be much more complex, because they combine transcriptional and posttranscriptional control mechanisms in intricate ways that are missed when network analysis is restricted to the transcriptional level. Thus, σ^S not only controls the expression of secondary transcription factors, but also that of signaling enzymes, which, for instance, produce or degrade the second messenger c-di-GMP, and of proteins that modulate the activities of transcription factors. Moreover, many of these components can cooperate in systems of nested feedforward loops to precisely control specific outputs.

A Biofilm Control Network

The currently best studied example is the network by which σ^S controls the formation of certain biofilm structures, such as adhesive curli fimbriae and the matrix components cellulose and, to some extent, colanic acid (Fig. 5; recently reviewed in reference 188) . The secondary transcriptional activator in this system is CsgD, an important regulator of biofilm formation (60, 61, 387, 417, 419, 421). Eσ^S directly controls csgD transcription, but does so in conjunction with a multitude of regulatory factors, many of which are themselves under σ^S control (Fig. 5). These include the MerR-like transcription factor MlrA (66, 374, 518) and several diguanylate cyclases (DGCs) and phosphodiesterases (PDEs) (374, 447, 518).

These signaling enzymes belong to the families of GGDEF and EAL domain proteins, respectively, that antagonistically control the second messenger c-di-GMP (187, 188, 221, 223, 415, 416). Two such antagonistic DGC/PDE pairs (YdaM/YciR and YegE/YhjH, with the DGC YedQ acting as a minor backup for YegE) control csgD transcription in a nonadditive manner (374, 518). The ydaM, yciR, yegE, and yedQ genes are all under positive control of σ^S (374, 447, 518), whereas yhjH is a class III gene in the flagellar hierarchy (148, 158, 248, 374, 414). Since σ^S, via sigma factor competition, is involved in downregulating FlhDC and the expression of the flagellar cascade, it also exerts an indirect negative effect on yhjH expression. In fact, this downregulation of the PDE YhjH is the triggering signal that allows the DGCs YegE and YedQ to accumulate sufficient c-di-GMP to activate csgD expression in conjunction with the YdaM/YciR system and MlrA (374). How two distinct DGC/PDE systems are integrated in this control, with only one also affecting motility [YegE(YedQ)/YhjH; see Fig. 5], is still unclear. In addition, csgD transcription is fine-tuned by the global regulators OmpR and CpxR, which allows for the integration of additional stress signals (233, 504).

Once CsgD accumulates, it activates the expression of the curli operon csgBAC and the yaiC gene, which encodes another DGC involved in activating cellulose biosynthesis, as well as of several other genes (60, 61, 419). Moreover, σ^S is required for the expression of the ycgZ-ymgABC operon, whose gene products (in particular, YmgB) modulate the activity of the Rcs phosphorelay system, with the response regulator RcsB inhibiting CsgD expression and stimulating colanic acid production (Fig. 5) (484). This may play a role later during biofilm formation when the composition of the matrix, consisting of cellulose and colanic acid content, is determined.

In summary, Eσ^S directly initiates transcription at the csgD promoter, but in addition deploys (i) a complex signal-integrating system of no less than seven factors that precisely control this initiation of transcription in response to numerous signals and (ii) several factors that differentially modulate the regulatory output of CsgD in a temporal manner.

The Acid Resistance Regulatory Network

Another σ^S-controlled network consisting of nested feedforward loops is responsible for acid resistance (141). Eσ^S, which is rapidly but transiently induced in response to pH downshift, is required for the transcription of the regulatory genes gadX, gadW, gadY, and gadE. The small regulatory GadY RNA controls the cellular level of GadX (366, 479). GadX is an AraC-type regulator, which may sense an alteration of the cellular Na⁺ concentration associated with exposure to acid and activates the transcription of gadE, gadY (thereby setting up a positive feedback loop), and other genes (366, 398, 439, 479, 480, 481, 488, 519). GadW is a GadX paralog with a complex, conditionally positive or negative function in the system (297, 439). The LuxR-like regulator GadE is required for the expression of the core acid resistance genes (including gadA, gadBC, hdeAB, hdeD), with these final target genes apparently being transcribed by vegetative RNAP (141, 299, 519). Positive autoregulation of GadE allows very strong expression of these target genes (141), with rapid shutoff of the system being possible because of Lon-mediated continuous degradation of GadE (199). Interestingly, GadE also downregulates many genes located in the Lee pathogenicity island of E. coli O157:H7 (500).

Expression of the acid resistance genes is further modulated by the Rcs phosphorelay system, with the response regulator RcsB acting directly in a complex with GadE (76, 77, 255, 256). Via the Rcs system, acid resistance genes also respond to the σ^S-controlled YmgB protein (282, 484).

Taken together, a similar complex pattern as in curli/cellulose control appears in the acid resistance network: σ^S deploys a battery of signal-integrating and interconnected regulatory factors, which then, in conjunction with vegetative RNAP, control the expression of the downstream target genes, whose gene products generate the phenotypic output. In addition, there are connections to σ^S-independent global regulatory circuits, because GadE expression can also be activated by a pathway that involves the EvgS/EvgA two-component system and the AraC-like regulator YdeO (298, 314, 315).

Feedback Loops within the σ^S Network

Finally, some σ^S-controlled genes are involved in regulatory feedback loops that affect σ^S itself. The most important one is certainly σ^S-dependent expression of its own "destruction" factor RssB, which sets up the negative and homeostatic feedback that is the center piece of the regulation of σ^S proteolysis by numerous stress signals (as described above). In addition, positive feedback loops probably assist robust accumulation of σ^S in stationary phase or upon stress exposure: (i) via sigma factor competition, σ^S negatively affects the expression of its own antagonist, FliZ (374) (and see above); (ii) the accumulation of the anti-σ⁷⁰ factor Rsd during entry into the stationary phase is partly σ^S-dependent (229); (iii) σ^S further stimulates the expression of ArcA, which by efficiently competing with RssB for being phosphorylated by ArcB, can reduce RssB activity and therefore contributes to σ^S stabilization (accordingly, an arcA mutant stabilizes σ^S to a lesser extent during entry into the stationary phase and shows reduced survival and stress resistance) (325, 357); and (iv) the σ^S-dependent biofilm regulator CsgD may be involved in further stabilizing σ^S, because it has been reported to stimulate the expression of IraP, which can sequester and thereby inactivate RssB (171) (and see above).

In conclusion, the σ^S-controlled network of target gene expression is beginning to reveal a strikingly complex organization. To understand the dynamic behavior of these multiple nested feedforward and feedback loops will be a major challenge for the future.

PHYSIOLOGICAL FUNCTIONS OF σ^S-CONTROLLED GENES

The σ^S Regulon

The σ^S-dependent general stress response is required for long-term survival in the stationary phase and adaptation to a variety of adverse conditions, but it also triggers rapid emergency responses to diverse suddenly arising stress conditions. Thus, rpoS mutants die off more rapidly when starved for various nutrients and do not develop resistance and cross-protection under different stress conditions (177, 191, 193, 237, 267, 318, 445). Not surprisingly, σ^S is also required for survival in natural environments such as the mammalian host or seawater and under conditions used in food preservation (85, 107, 159, 344, 386, 413).

σ^S positively controls more than 500 genes in E. coli K-12 laboratory strains and even more genes in pathogens like E. coli O157:H7 that have larger genomes (111, 112, 237, 262, 371, 519). In addition, negative control by σ^S can be observed, which in most cases probably reflects sigma factor competition for RNAP core enzyme, with rpoS mutants containing more of the vegetative and flagellar RNAP holoenzymes, Eσ⁷⁰ and Eσ²⁸, respectively (111, 129, 519). It should also be noted that σ^S-mediated expression of a gene does not necessarily mean that the gene product is there and active, since there might be posttranscriptional control and, in particular for enzymes, control at the metabolic level. Thus, σ^S-mediated gene expression sets the stage, on which additional and often unknown signals can then determine the actual activity of the gene products. For instance, the trehalose synthesis operon is activated under various conditions that induce σ^S, but high levels of trehalose accumulate only in growing cells shifted to high osmolarity (155, 191, 236). Another example is σ^S-controlled expression of mechanosensitive channels, which become active only in response to osmotic downshift (457).

Because of the large size of the σ^S regulon, it is often difficult to pinpoint the specific genes required for particular σ^S-controlled phenotypes. Consequently, complex phenotypes, as for instance σ^S-dependent thermotolerance in stationary phase, are still not understood on a mechanistic basis, whereas, on the other hand, many σ^S-controlled genes have remained functionally uncharacterized. This review can just give a cursory overview of a limited number of σ^S-dependent genes with either well-characterized or potentially interesting physiological functions in order to illustrate the functional diversity and power of the general stress response that orchestrates stationary-phase gene regulation.

Genes Involved in Multiple Stress Resistance and Cross-Protection

Stationary-phase cells of E. coli survive a variety of harsh stress conditions much better than growing cells. This multiple stress resistance is strongly dependent on a functional rpoS allele and its induction during entry into stationary phase (267, 318).

Osmotic resistance

A typical example is stationary-phase survival upon hyperosmotic shift (e.g., with 3 M NaCl). However, σ^S is also induced upon osmotic upshift in growing cells (193, 337), and σ^S-dependent expression was observed for 326 genes under these conditions. These included 186 genes not found under other conditions where σ^S accumulates, indicating that σ^S also serves as a major osmoregulator (519). Proteomic analysis corroborated the induction of many σ^S-controlled proteins upon osmotic upshift (517). σ^S activates the otsBA operon, which encodes the enzymes that produce the compatible solute and osmoprotectant trehalose, as well as the genes for the hyperosmotically induced periplasmic trehalase, TreA, and its cytoplasmic counterpart, TreF, which is involved in degrading trehalose upon osmotic downshift (47, 191, 235, 236, 465). A series of genes originally identified for their strong hyperosmotic activation has been found to be σ^S-controlled and also stationary phase-induced (49, 57, 175, 234). These include osmY, which encodes a periplasmic protein required for osmotolerance (25, 271, 520, 531, 532), and osmF (yehZ), which is part of the yehZVXW operon that codes for an ABC transporter probably involved in the uptake of an unidentified osmoprotectant (80). Another example is osmC, which encodes a peroxidase with a preference for organic hydroperoxides, the loss of which results in increased sensitivity to oxidative stress (57, 93, 160, 174, 287).

Oxidative stress and UV resistance

σ^S also confers pronounced oxidative stress resistance in the stationary phase (267, 318). Moreover, several other stress conditions such as UV irradiation or starvation for carbon or phosphorus sources endogenously generate oxidative stress, and therefore, resistance against these conditions involves a spectrum of σ^S-controlled proteins similar to that which is crucial to combat exogenous oxidative stress (37, 67, 119, 123, 159, 177, 219, 358, 367).

These proteins include (i) Dps, which is the most abundant nucleoid-associated protein in stationary-phase cells and which scavenges iron and thereby prevents the hydroxyl radical-generating Fenton reaction from occuring close to the DNA (5, 8, 118, 167, 312, 347, 454, 525, 539); (ii) exonuclease III (encoded by xthA), which is involved in DNA repair (103, 124); (iii) UspB, which facilitates resolution of Holliday junctions by RuvC in the RuvABC recombination repair pathway (372); (iv) the catalases KatE and KatG (217, 294, 343, 428); (v) glutathione reductase (encoded by gor) (32); (vi) a periplasmic superoxide dismutase (encoded by sodC) (464); (vii) a secondary iron-sulfur cluster assembly system encoded by the sufABCDSE operon (519); (viii) bacterioferritin encoded by bfr, which by sequestering iron may also prevent Fenton chemistry (51, 519); and (ix) the ferrochelatase encoded by hem, which catalyzes the insertion of Fe²⁺ as the last step in heme biosynthesis (99, 519).

Several of the corresponding genes (e.g., dps, katG, gor, hemH, and the sufABCDSE operon) also belong to the OxyR regulon and this overlap further extends to uncharacterized genes that therefore may have functions in oxidative stress protection (e.g., yaiA, ycfR, yjiD) (8, 283, 428, 519, 541). This dual regulation is probably required because σ^S itself does not significantly accumulate upon exposure to H₂O₂, because its stabilization seems compensated for by reduced rpoS translation due to the OxyR-induced small RNA OxyS (322, 537). Another interesting example of dual regulation is aidB, specifying a DNA repair enzyme, which is under control of σ^S and/or Ada, a regulator that responds to alkylation damage and can also cooperate with vegetative RNAP to activate aidB (263).

Acid resistance

σ^S is involved in strong acid resistance that is either associated with the stationary phase or generated in response to a nonlethal "adaptive" acid shift, a condition that also induces σ^S (28, 141, 280, 514). A σ^S-controlled protein with a key role in acid resistance is glutamate decarboxylase, which is expressed as two nearly identical isozymes from the gadA and gadB genes. These genes are under indirect σ^S control via the small regulatory GadY RNA and the regulatory proteins GadX and GadE (see above). In decarboxylating glutamate, this enzyme scavenges a proton and generates γ-aminobutyrate (GABA) that can be excreted via the glutamate/GABA antiporter GadC, which is encoded in the same operon as GadB (73, 141). When external glutamate is not available, the glutamate decarboxylate reaction is part of a bypass from the tricarboxylic acid (TCA) cycle, that also involves GABA transaminase (GabT) and succinic semialdehyde dehydrogenase (GabD), which are expressed from the equally σ^S-dependent gabDTP operon (324).

Another target of the acid stress regulators under σ^S control is the hdeAB operon, and mutants defective in this operon are acid-sensitive (314, 514, 519). HdeA and HdeB are extracytoplasmic chaperones that prevent protein aggregation in the periplasm at acidic pH (206, 241, 305). Additional σ^S-dependent genes that, like gadA, hdeAB, and the regulatory genes mentioned above, are located on the "acid fitness island " at 78.8 min of the E. coli chromosome, are hdeD, slp, and yhiUV (519). HdeD is a membrane protein involved in acid resistance only at high cell densities, whereas the lipoprotein Slp plays a role in resistance against organic acids, such as succinate, lactate, and formate (316). yhiUV (also termed mdtEF) encodes an efflux pump conferring resistance to rhodamin and sodium dodecyl sulfate (SDS) (201). Actually, some 14% of genes in the σ^S regulon specify membrane proteins. These include several known transport systems, but some of them could, like the MdtE/F system, act as efflux carriers that confer resistance to various toxic compounds (519). Right upstream of the "acid fitness island" and surrounded by additional σ^S-controlled genes of unknown function, the gene encoding the "universal stress protein," UspB, is located. This protein confers resistance against ethanol (130) and has recently been shown to be involved in the RuvABC-mediated recombination repair pathway (372) (and see above).

σ^S-controlled stress-protective genes with unknown molecular functions

Specific stress-protective molecular functions have been assigned to many σ^S-controlled proteins. In other cases, however, the molecular details underlying stress protection by particular proteins have remained mysterious. A striking example is the strongly σ^S-dependent YhbO protein (519). This highly conserved protein has similarity to class I glutamine amidotransferases and belongs to the DJ-1/ThiJ/Pfp1 superfamily, which includes chaperones, proteases, and the Parkinson's disease protein DJ-1. A yhbO mutant is highly sensitive against oxidative, thermal, acid, alkaline, osmotic, and UV stress, but a molecular activity of YhbO has yet to be identified (1).

Genes That Redirect Metabolism

σ^S was originally seen as a global regulator that mainly confers multiple stress resistance (192, 195, 293). However, around 20% of the genes in the σ^S regulon turned out to have metabolic functions (112, 371, 519). During slow growth, σ^S can also take over the expression of certain housekeeping genes that in rapidly growing cells are activated by vegetative RNAP. These include ribosomal genes (392) and glycolytic genes (365). In a way, σ^S thus becomes a second housekeeping sigma factor in slowly growing or stationary-phase cells.

σ^S actively redirects energy metabolism when cells enter into the stationary phase. Many genes under positive control of σ^S are involved in glycolysis, fermentation, anaerobic respiration, and alternative electron flow, whereas genes involved in aerobic respiration ( e.g., sdhCDAB encoding succinate dehydrogenase) tend to be negatively affected by σ^S. One of the most strongly σ^S-activated genes is poxB, which encodes pyruvate oxidase (78). This enzyme, which catalyzes the oxidative decarboxylation of pyruvate to acetate, bypasses the NADH-generating pyruvate dehydrogenase reaction (334), which is downregulated by the equally σ^S-activated repressor PdhR (519). These σ^S-mediated changes suggest that energy metabolism in stationary-phase cells is redirected in a way that counteracts the increased generation of reactive oxygen species, for instance, by NADH dehydrogenase II (119, 323, 334, 519).

A variety of ABC-transport systems for amino acids, oligo- or dipeptides, phosphate sources or polyamines (encoded by the art, dpp, opp, ugp, and pot operons, respectively) are under positive σ^S control (112, 371, 519). On the other hand, σ^S seems to collectively downregulate uptake and/or metabolism of certain alternative carbon sources. As a consequence, rpoS mutations improve growth on certain low-concentrated or nonoptimal carbon sources (83, 115, 244) and increase the expression of flagellar genes (113, 371). This is a reflection of the regulatory antagonism between the cAMP-CRP/Eσ⁷⁰-controlled foraging state (corresponding to the postexponential growth phase in LB medium) and the σ^S-controlled stationary phase with its emphasis on maintenance and stress survival (see further below).

σ^S seems to trigger switching between certain isozymes. It strongly activates talA and tktB, which encode transaldolase A and transketolase 2 that are involved in the pentose phosphate pathway, and for which housekeeping isozymes under vegetative control exist (encoded by talB and tktA, respectively). A similar case is the σ^S-dependent gene fbaB, which encodes one of two fructose-bisphosphate aldolases (262, 371, 519). The molecular or physiological reasons underlying isozyme switching are unknown.

Genes Affecting the Structure of the Cell Envelope and Overall Cellular Shape

During entry into stationary-phase cells change to a shorter, more ovoid cell shape. This is due to σ^S-dependent expression of the morphogene bolA (44, 266, 426), which controls transcription of dacA and dacC (specifying penicillin-binding proteins 5 and 6, respectively) and ampC (encoding a β-lactamase) (424). In addition, BolA downregulates the expression of MreB, a cytoskeletal component required for elongated cell shape (146). BolA probably indirectly affects the expression of these cell envelope proteins, because it appears to be a reductase able to interact with glutaredoxin (209). In addition, one of the promoters of the ftsQAZ operon is activated by σ^S, which may contribute to cell volume-reductive cell division that generates the shorter cells typical for the stationary phase (21).

During entry into the stationary phase, phospholipids are modified to contain cyclopropane fatty acids, which contributes to acid tolerance (79, 97). The cfa gene, which encodes cyclopropane fatty acid synthase, is under σ^S control (122, 509). Also, some lipoproteins show σ^S-regulated expression. These include OsmB, which may stimulate cellular aggregation (234), OsmE (176), and Blc (lipocalin), for which a role in membrane maintenance has been proposed (39, 71, 72).

Genes That Modulate Biofilm Formation and Composition

Bacteria can change from the planktonic, unicellular, and usually motile lifestyle to a multicellular biofilm existence, which is characterized by cell-cell and surface adherence and the production of matrix exopolysaccharides (EPS) (33, 458). For E. coli and related bacteria, overall gene expression in biofilms resembles that in planktonic stationary-phase cells, including the induction of σ^S and many σ^S-dependent genes (34, 35, 92, 109, 215, 429). This is not surprising given that biofilms are characterized by high local cell density , nutrient limitation and acid secretion (150, 458, 516).

At first glance, various reports seemed contradictory with respect to σ^S playing a positive or negative role in biofilm formation. In Salmonella enterica Serovar Typhimurium, σ^S is required for the "rdar" (rough, dry and red on Congo red-containing agar plates) or wrinkled colony morphotype, which reflects a structured biofilm at a wet surface/air interface (419, 421). In several studies with submerged E. coli biofilms, rpoS mutants could still adhere to surfaces but did not establish mature thick biofilms (2, 92, 216, 389) ; other researchers observed that a rpoS mutant formed even thicker biofilms (94). However, these variations are probably due to the use of different biofilm models (static growth in microtiter dishes, growth in a flow chamber or in a colony on agar plates), different kinds of surfaces (glass, plastic, agar, or even gallstones), different media (complex or minimal) and different strains of E. coli and Salmonella. While biofilm formation resembles a developmental process in that it occurs in temporally ordered and tightly controlled phases, it is also a very flexible process with the actual components and the final shape and architecture of the biofilm being modulated by a variety of environmental conditions (331). Even a single species can be capable of forming different types of biofilms under different conditions, for instance, inside a host and in the environment.

Below 30°C and further stimulated by low salt, E. coli and Salmonella produce adhesive curli fimbriae and cellulose (363, 420, 546), which generate the rdar morphotype on agar plates (421), and in the environment are involved for instance in attachment to plant leaves (22). Yet, under some conditions and in certain E. coli and Salmonella strains, curli fimbriae are also made at 37°C and can be involved in adhesion to host tissue (45, 364, 418). Via the biofilm regulator CsgD, curli and cellulose formation depend on σ^S (265, 387, 417, 419, 518). In addition, many diguanylate cyclases and phosphodiesterases (featuring GGDEF and EAL domains, respectively) that "make and break " the biofilm-promoting signaling molecule c-di-GMP, are under σ^S control (374, 447, 518). Several of these GGDEF/EAL domain proteins inversely coordinate motility and the expression of CsgD as well as the production of cellulose (238, 374, 440, 441, 518) (see also Fig. 5 and the section on σ^S-controlled downstream networks above).

In addition, the σ^S-regulated morphogene bolA was found to positively modulate biofilm formation (505). The σ^S-dependent ycgZ-ymgABC operon encodes the small proteins YmgA and YmgB that stimulate the activity of the Rcs two-component system, which downregulates CsgD expression and activates the production of the matrix component colanic acid and the expression of the acid resistance genes (282, 484). The ycgZ-ymgABC operon is activated during an intermediate phase of biofilm formation (109), and its expression is further stimulated by environmental signals such as low temperature and blue light (484) or reduced by indole, which generates flat biofilms (281). Thus, this system seems an important modulator of biofilm maturation in the environment. Two additional genes, yliH and yceP, are involved in this modulation by indole (108) and are expressed in a σ^S-dependent manner, especially at low temperatures (369, 521).

Virulence Genes

It is not surprising that σ^S is involved in surviving the harsh conditions that pathogenic E. coli or salmonellae encounter within their hosts, which include the exposure to acidic pH, bile, cationic antimicrobial peptides, or oxidative bursts generated by the innate immune system (104, 131). rpoS mutants exhibit reduced virulence (4, 128, 252, 413, 512, 522) and are good candidates for life vaccines (96, 110, 279, 423). rpoS mutants of Salmonella enterica serovar Typhimurium show reduced colonization of murine Peyer's patches, indicating that σ^S-dependent gene expression is required for the initial stages of systemic infection (352).

σ^S regulates a number of virulence genes. Specifically, in enterohemorrhagic E. coli O157:H7 (whose genome is approximately 20% larger than that of the K-12 strains) these include the genes encoding the type III-secreted proteins EspA, EspB, and EspD (36) ; the regulatory gene ler, which controls some genes located on the LEE pathogenicity island (112, 260) ; and an enterohemorrhagic E. coli-specific hemolysin operon (ehxCABD) (288). Other LEE-associated genes seem negatively regulated by σ^S (112, 476). By transcriptional profiling of an O157:H7 strain, approximately 10% of the σ^S-controlled genes observed were found to be unique to O157:H7 (112).

In S. enterica serovar Typhimurium, σ^S and σ^S-dependent genes are highly expressed after entry into macrophages and epithelial cells (81, 523). The best-studied of these genes are the plasmid-encoded spv genes that are required for intracellular growth in deep lymphoid organs such as spleen and liver (172). σ^S activates the regulatory gene spvR and, together with SpvR, can antagonize the inhibition of the spvABCD operon by H-NS (82, 183, 252, 253, 354, 400, 523). Furthermore, σ^S activates the expression of SEF14 fimbriae in S. enterica serovar Enteritidis (120), the production of the Vi capsular polysaccharide (423), and the expression of the taiA-hlyE virulence operon in S. enterica serovar Typhi (149).

Genes Involved in Stationary Phase and Stress-Induced Mutagenesis

Genetic variability is increased when cells are in stationary phase or exposed to various stresses. This seems a severe stress survival strategy, which operates at the level of the cell population, and a driving force for evolution (142, 473), that for instance can generate antibiotic resistance (378). The products of several σ^S-controlled genes are involved in this stationary-phase or stress-induced mutagenesis, which actually can produce very different types of mutations, suggesting that multiple mechanisms are involved. One is the error-prone translesion repair DNA polymerase IV (Pol IV) encoded by dinB. Not only the expression of Pol IV but also, via the SMC-like protein complex SbcCD, its activity are under σ^S control (273, 381, 449, 459). While Pol IV is responsible for approximately 85% of stress-induced mutations arising from DNA double-strand break repair, the remaining mutations arise from the activity of Pol II, which operates with higher fidelity and was recently found to be also under σ^S control (147). In addition, the mismatch repair proteins MutS and MutH are downregulated in a σ^S-dependent manner (133, 486).

However, there are additional σ^S-affected mutagenic events that do not seem to involve these systems (40, 198, 273, 295). In a search for genes that generated a mutator phenotype when overproduced, rpoS, iraD (yjiD, the product of which stabilizes σ^S, see above), dinB, as well as several other genes were isolated (528). The latter included two genes for which a positive control by σ^S has been observed: (i) appY, which encodes an AraC-like transcriptional regulator that activates the hya and cbdAB-appA operons, encoding hydrogenase I, a third cytochrome oxidase, and an acidic phosphatase (14); and (ii) paaX, encoding a transcriptional repressor of the paa operon, which is involved in the degradation of phenylacetic acid (137, 519). How these factors can stimulate mutagenesis has yet to be clarified. On the other hand, a σ^S-controlled antimutator, the DNA glycosylase Mug, has also been described (330).

Regulatory Genes

A series of regulatory genes that encode DNA-binding transcription factors are under σ^S control. The implications with respect to some regulatory cascades and networks downstream of σ^S have already been illustrated above (involving the σ^S-controlled regulators MlrA, CsgD, GadX, GadW, and GadE). During entry into stationary phase or upon osmotic upshift, σ^S also enhances the expression of four response regulator genes (arcA, basR, cpxR, and ompR) (186, 519). ArcA controls numerous genes in response to the redox state of the respiratory chain (307) and is also involved in a positive feedback loop that further stabilizes σ^S (325) (see above). BasR plays a role in metal resistance (178) and CpxR is a global regulator of an extracytoplasmic stress response and of biofilm formation (116, 368, 411).

Additional σ^S-controlled DNA-binding proteins belong to the families of AraC-, GntR-, LysR-, MerR-, and TetR-like regulators. In addition, the expression of two abundant nucleoid-associated proteins, CbpA (which has a DnaJ-like domain) and the α-subunit of IHF (encoded by himA) is positively regulated by σ^S (519). In many cases, the functions of these regulatory proteins are incompletely understood or even unknown. Functionally characterized regulatory proteins, for which σ^S regulation has been observed (519), include: (i) FhlA, a σ⁵⁴-activating AAA+-ATPase, that stimulates the expression of several operons involved in formate metabolism (430); (ii) PdhR, the GntR-like repressor of the genes for pyruvate dehydrogenase (391); (iii) NsrR, a nitric oxide-sensitive repressor of some 30 genes that include those that generate resistance against nitric oxide but also flagellar operons (42, 370, 461); and (iv) YncC (McbR), a GntR-like repressor of ybiM, which encodes a periplasmic protein that somehow stimulates colanic acid production (538). In a nutshell, these functions again point to the importance of σ^S in metabolic adaptation, multiple stress resistance, and biofilm formation.

In addition to controlling transcriptional regulators, σ^S is also involved in the expression of regulatory factors with a role in posttranscriptional regulation. Thus, YmdB, a partially σ^S-dependent protein induced at low temperature and during entry into stationary phase, acts as a direct inhibitor of RNase III (243). As RNase III is important for the degradation of rpoS mRNA (27, 320), YmdB may be involved in a positive feedback loop that stimulates σ^S expression.

GENETIC VARIABILITY OF σ^S AND THE BALANCE BETWEEN GROWTH AND STRESS SURVIVAL

Different laboratory strains of E. coli K-12 or Salmonella show variations in their σ^S levels and several point and small-deletion mutations were detected in the corresponding rpoS alleles (218, 231, 467). Similar mutations have also been found in enteropathogenic E. coli (38, 115, 514). While these mutants show decreased resistance against exogenous stresses, they seem to have an advantage in competitive colonization of the intestine (257). Interestingly, a similar rpoS polymorphism also occurs in clinical isolates of serovar Typhi, but not in serovar Typhimurium, possibly because the latter uses σ^S for expressing important virulence genes (401) (see also above). Also, commensal E. coli strains isolated from the environment or healthy hosts do not show rpoS polymorphism (212).

When σ^S-proficient E. coli laboratory strains were exposed to long-term starvation, rpoS -related mutations arose and even took over the cultures. These "GASP" mutations ("growth advantage in stationary phase"), which have been localized in rpoS itself, in genes involved in rpoS regulation, as well as other genes, do not eliminate σ^S completely, but result in attenuated σ^S function (139, 533, 534, 545). Sometimes, these mutants can stably coexist with σ^S-proficient strains (138, 410). Because stress resistance even quantitatively correlates with σ^S levels (37), the mutants with reduced σ^S levels or activity are less stress-resistant and also lose other σ^S-dependent phenotypes such as rdar colony morphology, but obviously grow better with the low nutrient levels derived from the debris of lysed cells (100, 139, 410, 533). Null mutations in rpoS arise during continuous growth under conditions of glucose or phosphate limitation in chemostats (356, 510), or when cells are grown on agar plates with succinate, i.e., a carbon source that supports slow growth only (83).

Overall, rpoS mutations arise under conditions of continuous low nutrient supply, but absence of severe stress (besides nutrient limitiation). Phenotypically, the mutants perfectly reflect these selective conditions, because they show a broader nutritional capability and compete better when confronted with low nutrient supply, but they are impaired in multiple stress resistance. Interestingly, even rpoS null mutants came up in glucose-limited chemostat cultures, but additional application of stress (mildly acidic pH) selected for mutants only partially reduced in σ^S activity (135, 244, 356). An additional benefit of attenuated rpoS mutations could be the avoidance of the increased mutation rates associated with high σ^S levels and activity (see above).

All these studies indicate a regulatory trade-off, i.e., that maximal nutritional competence on the one hand and high levels of multiple stress resistance on the other are mutually exclusive. At the molecular level, this reflects sigma factor competition for limiting RNAP core enzyme, since genes for high-affinity nutrient-scavenging systems are usually activated by Eσ⁷⁰ cooperating with cAMP-CRP or other transcription factors, whereas generating multiple stress resistance is the domain of Eσ^S (134, 135, 453). As apparent from a wider perspective, this balance is part of a more general choice between mutually exclusive lifestyles. These are the foraging motile lifestyle of postexponentially and therefore slowly growing cells and the stationary-phase lifestyle characterized by maintenance metabolism, stress resistance, high cell density, and cellular adherence. The molecular basis of this decision is the interplay between sigma factor competition for core RNAP (involving σ⁷⁰, σ²⁸, σ^S, σ^E, and σ⁵⁴) and complex nucleotide second messenger signaling [involving cAMP, (p)ppGpp, and c-di-GMP] (373, 374) (and see above).

But why is this intricately balanced regulation not sufficient for cells to properly adapt to the various conditions mentioned above, and the balance actually reset by mutations in rpoS? The reason may be that this lifestyle transition probably operates as a switch with a relatively sharp threshold (at a distinct low nutrient concentration). This seems so because (i) the major regulatory players, i.e., the sigma factors, show mutual inhibition of activity; (ii) accessory factors such as FliZ, Crl, and Rsd further support this antagonism; and (iii) at least the low-nutrient state dominated by Eσ^S is stabilized by positive feedback loops (see above). Mutations that attenuate σ^S levels and/or its efficiency in sigma factor competition actually readjust the threshold of switching to the "high Eσ^S/stress survival" state to occur at even lower nutrient concentrations. While no selective pressure for threshold resetting seems to exist during the complex life cycles of commensal E. coli (which retain wild-type rpoS alleles, see above), enteropathogenic E. coli seem to experience such a selection in their natural environment, and so do laboratory strains when exposed to the rather artificial condition of continuous long-term growth at very low nutrient concentrations in the absence of any other stressful disturbance.

ORIGIN AND PHYSIOLOGICAL ROLES OF σ^S IN OTHER GRAM-NEGATIVE BACTERIA

σ^S-like sigma factors have also been identified in other gram-negative bacteria that belong to the γ-, β-, and δ-proteobacteria (though not in the α- and ε-clades). As in the paradigmatic E. coli, these σ^S-like sigmas have often been associated with stationary-phase gene regulation, various kinds of stress resistance and/or virulence (16, 17, 62, 86, 179, 182, 210, 274, 286, 290, 296, 394, 446, 455, 498, 502, 530, 547). Even in the spirochete Borrelia burgdorferi, a sigma factor gene, which is itself regulated by multiple environmental signals and is involved in virulence gene expression, has been annotated as rpoS (68, 70). However, as σ^S is closely related to the vegetative σ⁷⁰ (42% overall identity in E. coli), it is not trivial to decide whether genes annotated as ‘rpoS‘ in clades other than the γ-proteobacteria are real homologs of E. coli rpoS and not just additional independent 'evolutionary spin-offs' of the vegetative sigma factor gene(s).

It is obvious that rpoS evolved from a duplication of a rpoD-like ancestral gene (170). In adopting a new function, a sigma factor has to coevolve with a cognate promoter sequence, since selection operates on a phenotype determined by gene expression mediated by this sigma. Thus, the defining criterion of a sigma factor (and its homologs) is its own specific mechanism of promoter recognition. Two features functionally distinguish σ^S from the vegetative σ⁷⁰: (i) the recognition of C(−13) in a promoter specifically by its K173 residue (the corresponding amino acid in σ⁷⁰ is the oppositely charged E458) (29); and (ii) the special role in transcription initiation of a 16-amino-acid σ^S-specific C-terminal extension that is not present in σ⁷⁰ (106, 165, 285, 360, 406, 407). All γ-proteobacteria that do have σ^S (some pathogens such as Xanthomonas and Acinetobacter seem to have lost it), contain this lysine residue as well as the well-conserved C-terminal extension in their σ^S proteins (Table 1). β-Proteobacteria have the lysine replaced by an arginine, and do have the C-terminal extension, although it has limited similarity to the corresponding region in E. coli σ^S. Also among the β-proteobacteria, some pathogens such as Bordetella and Neisseria species do not have an rpoS-like gene. Among the δ-proteobacteria, a gene annotated as rpoS exists in Geobacter sulfurreducens (274), but other representatives, such as Bdellovibrio and Desulfovibrio, lack rpoS-like genes. rpoS-like genes in δ-proteobacteria usually are considered homologs of rpoS in γ/β-proteobacteria (86). Strikingly, however, the Geobacter "σ^S" rather resembles rpoD-encoded vegetative sigmas, because it contains a negatively charged residue (D) at the position that corresponds to E458 in σ⁷⁰ and K173 in σ^S of E. coli, and also does not have the C-terminal extension. Finally, the Borrelia sigma factor annotated as a σ^S has a leucine at the K173 position, no C-terminal extension, and just 34% identity to E. coli σ^S.

TABLE 1.Comparison of functionally important elements in sigma factors annotated as σ^S (RpoS) in γ-, β-, and δ-proteobacteria and spirochetes
Sigma factor from:	Identity to E. coli σ^{S (%)}	Amino acid at K173 position	C-terminal 16-amino-acid extension^a	Presence of RssB
γ-Proteobacteria^b	60–99	K	+	+ ^c
β-Proteobacteria^d	50–53	R	(+)	-
δ-Proteobacteria^e	47	D	-	-
Spirochete^f	34	L	-	-
σ⁷⁰ (RpoD) of E. coli	42	E	-	+ ^g
^aIn E. coli, the σ^S-specific C-terminal extension is EILQTQGLNIEALFRE.
^bExamples of σ^S-like sigmas in γ-proteobacteria and their identities to E. coli σ^S are: Salmonella enterica serovar Typhimurium. 99%; Yersinia pseudotuberculosis, 93%; Shewanella oneidensis , 79%, pseudomonads, about 75% ; Vibrio cholerae , 73%, Legionella pneumophila, 68%; the intracellular pathogen Coxiella burnetii , 58%.
^c Some γ-proteobacteria apparently have lost RssB, e.g., L. pneumophila and C. burnetii.
^d Examples of β-proteobacteria: Ralstonia solanacearum, 53%; Burkholderia pseudomallei , 50% identity to E. coli σ^S.
^eGeobacter sulfurreducens.
^fBorrelia burgdorferi and Borrelia afzelii.
^gBut σ⁷⁰ is not regulated by RssB.

Based on these properties and the phylogenetic distance between spirochetes and proteobacteria (173), it seems highly likely that at least the Borrelia sigma factor gene annotated as rpoS has evolved from an independent duplication of a rpoD-like ancestral gene. Given its closer similarity to the vegetative sigmas than to the σ^S factors of γ- and β-proteobacteria, the same may also apply to the Geobacter σ^S. In fact, this would provide an explanation for the absence of a σ^S-like sigma factor in α-proteobacteria, which phylogenetically are located between the δ- and the γ/β-proteobacteria (173). However, a more thorough analysis of rpoD/rpoS-like genes in various δ-proteobacteria is required before the question of whether they have a true homolog of the σ^S present in γ/β-proteobacteria or not, can be settled.

Within the γ-proteobacteria, the molecular details of σ^S acting as a transcription factor as well as its regulation by RssB appear to be conserved (with notable exceptions such as L. pneumophila, which does not seem to have a homolog of RssB; Table 1). Yet, the overall physiological function of σ^S can vary. Enteric bacteria other than E. coli also use σ^S as a master regulator for a stationary phase and general stress response (18, 313, 321, 452, 451). In other γ-proteobacteria, this function is in principle maintained, but further adapted to the requirements of the specific environmental niches inhabited (86, 182, 207, 210, 456, 455, 502). In Vibrio cholerae, σ^S is induced during the mucosal escape response, activates genes involved in motility and chemotaxis, and in general prepares cells for an outside existence characterized by starvation and multiple environmental stress (353, 530). Moreover, σ^S is involved in environmentally modulated biofilm formation (346). In L. pneumophila, σ^S controls genes required for growth inside ameba and for transmission between host ameba and macrophages (17, 59, 179, 207, 474). In Xenorhabdus nematophilus, rpoS is required for mutualistic colonization of the nematode intestine, but it is dispensable for pathogenizing larval-stage insects (506). Pseudomonads have integrated σ^S-dependent gene regulation into their complex two-component and quorum-sensing network that controls exoenzyme expression and virulence (182, 432, 433) and σ^S seems to affect biofilm formation (200, 526). Overall, this functional diversification also indicates how easily genes are recruited into or lost from the σ^S regulon ; in fact, because of the similarity of σ^S-dependent promoters to vegetative promoters, one or two point mutations in a promoter could be sufficient to recruit a vegetatively controlled gene into the σ^S regulon or vice versa.

PERSPECTIVES

Twenty years after the first description of σ^S as a central regulator of stationary-phase gene expression in E. coli (267), it is now possible to integrate the molecular and physiological functions as well as the amazingly complex regulation of σ^S and stationary-phase gene expression into a wider and whole system-integrating picture, which describes the transitions between entire bacterial lifestyles. However, even for the E. coli paradigm, many important questions remain to be solved. While we now understand reasonably well the molecular function of σ^S in promoter recognition and the core mechanisms of σ^S regulation, there are vast blanks with respect to multiple signal integration into σ^S control. Moreover, the architecture and dynamic behavior of the complex networks directed by σ^S and the molecular and physiological functions of numerous target genes are still a huge territory for further research. Furthermore, the analysis of the roles and regulation of σ^S homologs in other proteobacteria promises new insights into evolutionary adaptation to a variety of environmental and host-associated niches. Finally, stationary-phase gene regulation and general stress responses in gram-negative bacteria that clearly do not have σ^S homologs seem a widely open field for exciting discoveries, as recently demonstrated with α-proteobacteria (143).

ACKNOWLEDGMENTS

Research in my laboratory has been generously funded by the Deutsche Forschungsgemeinschaft, an ERC Advanced Investigator Grant, and the Fonds der Chemischen Industrie.

References

1. Abdallah, J., T. Caldas, F. Kthiri, R. Kern, and G. Richarme. 2007. YhbO protects cells against multiple stresses. J. Bacteriol. 189:9140–9144.[PubMed] [CrossRef]

2. Adams, J. L., and R. J. C. MacLean. 1999. Impact of rpoS deletion on Escherichia coli biofilms. Appl. Environ. Microbiol. 65:4285–4287.[PubMed]

3. Adler, J., and B. Templeton. 1967. The effect of environmental conditions on the motility of Escherichia coli. J. Gen. Microbiol. 46:175–184.[PubMed]

4. Alam, M. S., M. H. Zaki, J. Yoshitake, T. Akuta, T. Ezaki, and T. Akaike. 2006. Involvement of Salmonella enterica serovar Typhi RpoS in resistance to NO-mediated host defense against serovar Typhi infection. Microb. Pathog. 40:116–125.[PubMed] [CrossRef]

5. Almirón, M., A. Link, D. Furlong, and R. Kolter. 1992. A novel DNA binding protein with regulatory and protective roles in starved Escherichia coli. Genes Dev. 6:2646–2654.[PubMed] [CrossRef]

6. Altuvia, S., and E. G. H. Wagner. 2000. Switching on and off with RNA. Proc. Natl. Acad. Sci. USA 97:9824–9826.[PubMed] [CrossRef]

7. Altuvia, S., D. Weinstein-Fischer, A. Zhang, L. Postow, and G. Storz. 1997. A small, stable RNA induced by oxidative stress: roles as a pleiotropic regulator and antimutator. Cell 90:43–53.[PubMed] [CrossRef]

8. Altuvia, S., M. Almirón, G. Huisman, R. Kolter, and G. Storz. 1994. The dps promoter is activated by OxyR during growth and by IHF and σ^S in stationary phase. Mol. Microbiol. 13:265–272.[PubMed] [CrossRef]

9. Amsler, C. D., M. Cho, and P. Matsumura. 1993. Multiple factors underlying the maximum motility of Escherichia coli as cultures enter post-exponential growth. J. Bacteriol. 175:6238–6244.[PubMed]

10. Anderson, K. L., J. E. Whitlock, and V. J. Harwood. 2005. Persistence and differential survival of fecal indicator bacteria in subtropical waters and sediments. Appl. Environ. Microbiol. 71:3041–3048.[PubMed] [CrossRef]

11. Arluison, V., S. Hohng, R. Roy, O. Pellegrini, P. Régnier, and T. Ha. 2007. Spectroscopic observation of RNA chaperone activities of Hfq in post-transcriptional regulation by small non-conding RNA. Nucleic Acids Res. 35:999–1006.[PubMed] [CrossRef]

12. Arnquist, A., A. Olsén, and S. Normark. 1994. σ^S-dependent growth phase induction of the csgBA promoter in Escherichia coli can be achieved in vivo by σ⁷⁰ in the absence of the nucleoid-associated protein H-NS. Mol. Microbiol. 13:1021–1032.[PubMed] [CrossRef]

13. Arnquist, A., A. Olsén, J. Pfeifer, D. G. Russell, and S. Normark. 1992. The Crl protein activates cryptic genes for curli formation and fibronectin binding in Escherichia coli HB101. Mol. Microbiol. 6:2443–2452.[PubMed] [CrossRef]

14. Atlung, T., K. Knudsen, L. Heerfordt, and L. Brøndsted. 1997. Effect of σ^S and the transcriptional activator AppY on induction of the Escherichia coli hya and cbdAB-appA operons in response to carbon and phosphate starvation. J. Bacteriol. 179:2141–2146.[PubMed]

15. Azam, T. A., A. Iwata, A. Nishimura, S. Ueda, and A. Ishihama. 1999. Growth phase-dependent variation in protein composition of the Escherichia coli nucleoid. J. Bacteriol. 181:6361–6370.[PubMed]

16. Bachman, M. A., and M. S. Swanson. 2001. RpoS co-operates with other factors to induce Legionella pneumophila virulence in the stationary phase. Mol. Microbiol. 40:1201–1214.[PubMed] [CrossRef]

17. Bachman, M. A., and M. S. Swanson. 2004. Genetic evidence that Legionella pneumophila RpoS modulates expression of the transmission phenotype in both the exponential and the stationary phase. Infect. Immun. 72:2468–2476.[PubMed] [CrossRef]

18. Badger, J. L., and V. L. Miller. 1995. Role of RpoS in survival of Yersinia enterocolitica to a variety of environmental stresses. J. Bacteriol. 177:5370–5373.[PubMed]

19. Baker, T. A., and R. T. Sauer. 2006. ATP-dependent proteases of bacteria: recognition logic and operating principles. Trends Biochem. Sci. 31:647–653.[PubMed] [CrossRef]

20. Balandina, A., L. Claret, R. Hengge-Aronis, and J. Rouvière-Yaniv. 2001. The Escherichia coli histone-like protein HU regulates rpoS translation. Mol. Microbiol. 39:1069–1079.[PubMed] [CrossRef]

21. Ballesteros, M., S. Kusano, A. Ishihama, and M. Vicente. 1998. The ftsQ1p gearbox promoter of Escherichia coli is a major sigma S-dependent promoter in the ddlB-ftsA region. Mol. Microbiol. 30:419–430.[PubMed] [CrossRef]

22. Barak, J. D., L. Gorski, P. Naraghi-Arani, and A. O. Charkowski. 2005. Salmonella enterica virulence genes are required for bacterial attachment to plant tissue. Appl. Environ. Microbiol. 71:5685–5691.[PubMed] [CrossRef]

23. Barembruch, C., and R. Hengge. 2007. Cellular levels and activity of the flagellar sigma factor FliA of Escherichia coli are controlled by FlgM-modulated proteolysis. Mol. Microbiol. 65:76–89.[PubMed] [CrossRef]

24. Barne, K. A., J. A. Bown, S. J. W. Busby, and S. D. Minchin. 1997. Region 2.5. of the Escherichia coli RNA polymerase σ⁷⁰ subunit is responsible for the recognition of the “extended ” motif at promoters. EMBO J. 16:4034–4040.[PubMed] [CrossRef]

25. Barron, A., G. May, E. Bremer, and M. Villarejo. 1986. Regulation of envelope protein composition during adaptation to osmotic stress in Escherichia coli. J. Bacteriol. 167:433–438.[PubMed]

26. Barth, M., C. Marschall, A. Muffler, D. Fischer, and R. Hengge-Aronis. 1995. A role for the histone-like protein H-NS in growth phase-dependent and osmotic regulation of σ^S and many σ^S-dependent genes in Escherichia coli. J. Bacteriol. 177:3455–3464.[PubMed]

27. Basineni, S. R., R. Madhugiri, T. Kolmsee, R. Hengge, and G. Klug. 2009. The influence of Hfq and ribonucleases on the stability of the small non-coding RNA OxyS and its target rpoS in E. coli is growth phase dependent. RNA Biol. 6:584–594.[PubMed] [CrossRef]

28. Bearson, S. M. D., W. H. Benjamin , Jr., W. E. Swords, and J. W. Foster. 1996. Acid shock induction of RpoS is mediated by the mouse virulence gene mviA of Salmonella typhimurium. J. Bacteriol. 178:2572–2579.[PubMed]

29. Becker, G., and R. Hengge-Aronis. 2001. What makes an Escherichia coli promoter σ^S-dependent? Role of the -13/-14 nucleotide promoter positions and region 2.5 of σ^S. Mol. Microbiol. 39:1153–1165.[PubMed] [CrossRef]

30. Becker, G., E. Klauck, and R. Hengge-Aronis. 1999. Regulation of RpoS proteolysis in Escherichia coli: The response regulator RssB is a recognition factor that interacts with the turnover element in RpoS. Proc. Natl. Acad. Sci. USA 96:6439–6444.[PubMed] [CrossRef]

31. Becker, G., E. Klauck, and R. Hengge-Aronis. 2000. The response regulator RssB, a recognition factor for σ^S proteolysis in Escherichia coli, can act like an anti-σ^S factor. Mol. Microbiol. 35:657–666.[PubMed] [CrossRef]

32. Becker-Hapak, M., and A. Eisenstark. 1995. Role of rpoS in the regulation of glutathione oxidoreductase (gor) in Escherichia coli. FEMS Microbiol. Lett. 134:39–44.[PubMed]

33. Beloin, C., A. Roux, and J.-M. Ghigo. 2008. Escherichia coli biofilms. Curr. Top. Microbiol. Immunol. 322:249–289.[PubMed] [CrossRef]

34. Beloin, C., and J.-M. Ghigo. 2005. Finding gene-expression patterns in bacterial biofilms. Trends Microbiol. 13:16–19.[PubMed] [CrossRef]

35. Beloin, C., J. Valle, P. Latour-Lambert, P. Faure, M. Kzreminski, D. Balestrino, J. A. Haagensen, S. Molin, G. Prensier, B. Arbeille, and J.-M. Ghigo. 2004. Global impact of mature biofilm lifestyle on Escherichia coli K-12 gene expression. Mol. Microbiol. 51:659–674.[PubMed] [CrossRef]

36. Beltrametti, F., A. U. Kresse, and C. A. Guzmán. 1999. Transcriptional regulation of the esp genes of enterohemorrhagic Escherichia coli. J. Bacteriol. 181:3409–3418.[PubMed]

37. Berney, M., H. U. Weilenmann, J. Ihssen, C. Bassin, and T. Egli. 2006. Specific growth rate determines the sensitivity of Escherichia coli to thermal, UVA, and solar disinfection. Appl. Environ. Microbiol. 72:2586–2593.[PubMed] [CrossRef]

38. Bhagwat, A. A., J. Tan, M. Sharma, M. Kothary, S. Low, B. D. Tall, and M. Bhagwat. 2006. Functional heterogeneity of RpoS in stress tolerance of enterohemorrhagic Escherichia coli strains. Appl. Environ. Microbiol. 72:4978–4986. [CrossRef]

39. Bishop, R. E., S. S. Penfold, L. S. Frost, J.-V. Höltje, and J. H. Weiner. 1995. Stationary phase expression of a novel Escherichia coli outer membrane lipoprotein and its relationship with mammalian apolipoprotein-D: implications for the origin of lipocalins. J. Biol. Chem. 270:23097–23103.[PubMed] [CrossRef]

40. Bjedov, I., O. Tenaillon, B. Gerard, V. Souza, E. Denamur, M. Radman, F. Taddei, and I. Matic. 2003. Stress-induced mutagenesis in bacteria. Science 300:1404–1409.[PubMed] [CrossRef]

41. Blot, N., R. Mavathur, M. Geertz, A. Travers, and G. Muskhelisvili. 2006. Homeostatic regulation of supercoiling sensitivity coordinates transcription of the bacterial genome. EMBO Rep. 7:710–715.[PubMed] [CrossRef]

42. Bodenmiller, D. M., and S. Spiro. 2006. The yjeB (nsrR) gene of Escherichia coli encodes a nitric oxide-sensitive transcriptional regulator. J. Bacteriol. 188:874–881.[PubMed] [CrossRef]

43. Boehm, A., M. Kaiser, H. Li, C. Spangler, K. C.A., M. Ackerman, V. Kaever, V. Sourjik, V. Roth, and U. Jenal. 2010. Second messenger-mediated adjustment of bacterial swimming velocity. Cell 141:107–116.[PubMed] [CrossRef]

44. Bohannon, D. E., N. Connell, L. K., A. Tormo, M. Espinosa-Urgel, M. M. Zambrano, and R. Kolter. 1991. Stationary-phase-inducible “gearbox” promoters: Differential effects of katF mutations and role of σ⁷⁰. J. Bacteriol. 173:4482–4492.[PubMed]

45. Bokranz, W., X. Wang, H. Tschape, and U. Römling. 2005. Expression of cellulose and curli fimbriae by Escherichia coli isolated from the gastrointestinal tract. J. Med. Microbiol. 54:1171–1182.[PubMed] [CrossRef]

46. Bonnefoy, E., and J. Rouviere-Yaniv. 1991. HU and IHF, two homologous histone-like proteins of Escherichia coli, form different protein-DNA complexes with short DNA fragments. EMBO J. 10:687–696.[PubMed]

47. Boos, W., U. Ehmann, H. Forkl, W. Klein, M. Rimmele, and P. Postma. 1990. Trehalose transport and metabolism in Escherichia coli. J. Bacteriol. 172:3450–3461.[PubMed]

48. Bordes, P., A. Conter, V. Moales, J. Bouvier, A. Kolb, and C. Gutierrez. 2003. DNA supercoiling contributes to disconnect σ^S accumulation from σ^S-dependent transcription in Escherichia coli. Mol. Microbiol. 48:561–571.[PubMed] [CrossRef]

49. Bordes, P., R. Repoila, A. Kolb, and C. Gutierrez. 2000. Involvement of differential efficiency of transcription by Eσ^S and Eσ⁷⁰ RNA polymerase holenzymes in growth phase regulation of the Escherichia coli osmE promoter. Mol. Microbiol. 35:845–853.[PubMed] [CrossRef]

50. Botsford, J. L., and J. G. Harman. 1992. Cyclic AMP in prokaryotes. Microbiol. Rev. 56:100–122.[PubMed]

51. Bou-Abdallah, F., A. C. Lewin, N. E. Le Brun, G. R. Moore, and N. D. Chasteen. 2002. Iron detoxification properties of Escherichia coli bacterioferritin: attenuation of oxyradical chemistry. J. Biol. Chem. 277:37064–37069.[PubMed] [CrossRef]

52. Bouché, S., E. Klauck, D. Fischer, M. Lucassen, K. Jung, and R. Hengge-Aronis. 1998. Regulation of RssB-dependent proteolysis in Escherichia coli: a role for acetyl phosphate in a response regulator-controlled process. Mol. Microbiol. 27:787–795.[PubMed] [CrossRef]

53. Bougdour, A., C. Cunning, P. J. Baptiste, T. Elliott, and S. Gottesman. 2008. Multiple pathways for regulation of sigmaS (RpoS) stability in Escherichia coli via the action of multiple anti-adaptors. Mol. Microbiol. 68:298–313.[PubMed] [CrossRef]

54. Bougdour, A., C. Lelong, and J. Geiselmann. 2004. Crl, a low temperature-induced protein in Escherichia coli that binds directly to the stationary phase sigma subunit of RNA polymerase. J. Biol. Chem. 279:19540–19550.[PubMed] [CrossRef]

55. Bougdour, A., S. Wickner, and S. Gottesman. 2006. Modulating RssB activity: IraP, a novel regulator of σ^S stability in Escherichia coli. Genes Dev. 20:884–897.[PubMed] [CrossRef]

56. Bouveret, E., and A. Battesti. 2011. The stringent response, p. 231–250. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses, 2nd ed. ASM Press, Washington, DC.

57. Bouvier, J., S. Gordia, G. Kampmann, R. Lange, R. Hengge-Aronis, and C. Gutierrez. 1998. Interplay between global regulators of Escherichia coli: effect of RpoS, H-NS and Lrp on transcription of the gene osmC. Mol. Microbiol. 28:971–980.[PubMed] [CrossRef]

58. Brodolin, K., N. Zenkin, A. Mustaev, D. Mamaeva, and H. Heumann. 2004. The σ⁷⁰ subunit of RNA polymerase induces lacUV5 promoter-proximal pausing of transcription. Nat. Struct. Mol. Biol. 11:551–557.[PubMed] [CrossRef]

59. Broich, M., K. Rydzewski, T. L. McNealy, R. Marre, and A. Flieger. 2006. The global regulatory proteins LetA and RpoS control phospholipase A, lysophospholipase A, acyltransferase, and other hydrolytic activities of Legionella pneumophila JR32. J. Bacteriol. 188:1218–1226.[PubMed] [CrossRef]

60. Brombacher, E., A. Baratto, C. Dorel, and P. Landini. 2006. Gene expression regulation by the curli activator CsgD protein: modulation of cellulose biosynthesis and control of negative determinants for microbial adhesion. J. Bacteriol. 188:2027–2037.[PubMed] [CrossRef]

61. Brombacher, E., C. Dorel, A. J. B. Zehnder, and P. Landini. 2003. The curli biosynthesis regulator CsgD co-ordinates the expression of both positive and negative determinants for biofilm formation in Escherichia coli. Microbiology 149:2847–2857.[PubMed] [CrossRef]

62. Brown, D. G., J. K. Swanson, and C. Allen. 2007. Two host-induced Ralstonia solanacearum genes, acrA and dinF, encode multidrug efflux pumps and contribute to bacterial wilt virulence. Appl. Environ. Microbiol. 73:2777–2786.[PubMed] [CrossRef]

63. Brown, L., and T. Elliott. 1996. Efficient translation of the RpoS sigma factor in Salmonella typhimurium requires host factor I, an RNA-binding protein encoded by the hfq gene. J. Bacteriol. 178:3763–3770.[PubMed]

64. Brown, L., and T. Elliott. 1997. Mutations that increase expression of the rpoS gene and decrease its dependence on hfq function in Salmonella typhimurium. J. Bacteriol. 179:656–662.[PubMed]

65. Brown, N. L., J. V. Stoyanov, S. P. Kidd, and J. L. Hobman. 2003. The MerR family of transcriptional regulators. FEMS Microbiol. Rev. 27:145–163.[PubMed] [CrossRef]

66. Brown, P. K., C. M. Dozois, C. A. Nickerson, A. Zuppardo, J. Terlonge, and R. Curtiss III. 2001. MlrA, a novel regulator of curli (Agf) and extracellular matrix synthesis by Escherichia coli and Salmonella enterica serovar typhimurium. Mol. Microbiol. 41:349–363.[PubMed] [CrossRef]

67. Bucheli-Witschel, M., C. Bassin, and T. Egli. 2010. UV-C inactivation in Escherichia coli is affected by growth conditions preceding irradiation, in particular by the specific growth rate. J. Appl. Microbiol. 109:1733–1744.[PubMed]

68. Burtnick, M. N., J. S. Downey, P. J. Brett, J. A. Boylan, J. G. Frye, T. R. Hoover, and F. C. Gherardini. 2007. Insights into the complex regulation of rpoS in Borrelia burgdorferi. Mol. Microbiol. 65:277–293.[PubMed] [CrossRef]

69. Böhringer, J., D. Fischer, G. Mosler, and R. Hengge-Aronis. 1995. UDP-glucose is a potential intracellular signal molecule in the control of expression of σ^S and σ^S-dependent genes in Escherichia coli. J. Bacteriol. 177:413–422.[PubMed]

70. Caimano, J. J., R. Iver, C. H. Eggers, C. Gonzalez, E. A. Morton, M. A. Gilbert, I. Schwartz, and J. D. Radolf. 2007. Analysis of the RpoS regulon in Borrelia burgdorferi in response to mammalian host signals provides insight into RpoS function during the enzootic cycle. Mol. Microbiol. 65:1193–1217.[PubMed] [CrossRef]

71. Campanacci, V., D. Nurizzo, S. Spinelli, C. Valencia, M. Tegoni, and C. Cambillau. 2004. The crystal structure of the Escherichia coli lipocalin Blc suggests a possible role in phospholipid binding. FEBS Lett. 562:183–188.[PubMed] [CrossRef]

72. Campanacci, V., R. E. Bishop, S. Blangy, M. Tegoni, and C. Cambillau. 2006. The membrane bound bacterial lipocalin Blc is a functional dimer with binding preference for lysophospholipids. FEBS Lett. 580:4877–4883.[PubMed] [CrossRef]

73. Capitani, G., D. de Biase, C. Aurizi, H. Gut, F. Bossa, and M. G. Grutter. 2003. Crystal structure and functional analysis of Escherichia coli glutamate decarboxylase. EMBO J. 22:4027–4037.[PubMed] [CrossRef]

74. Carabetta, V. J., B. K. Mohanty, S. R. Kushner, and T. J. Silhavy. 2009. The response regulator SprE (RssB) modulates polyadenylation and mRNA stability in Escherichia coli. J. Bacteriol. 191:6812–6821.[PubMed] [CrossRef]

75. Carabetta, V. J., T. J. Silhavy, and I. M. Cristea. 2010. Teh response regulator SprE (RssB) is required for maintaining poly(A) polymerase I-degradosome association during stationary phase. J. Bacteriol. 192:3713–3721.[PubMed] [CrossRef]

76. Castanié-Cornet, M.-P., H. Treffandier, A. Francez-Charlot, C. Gutierrez, and K. Cam. 2007. The glutamate-dependent acid resistance system in Escherichia coli: essential and dual role of the His-Asp phosphorelay RcsCDB/AF. Microbiology 153:238–246.[PubMed] [CrossRef]

77. Castanié-Cornet, M.-P., K. Cam, B. Bastiat, A. Cros, P. Bordes, and C. Gutierrez. 2010. Acid stress response in Escherichia coli: mechanism of regulation of gadA transcription by RcsB and GadE. Nucleic Acids Res. 38:3546–3554.[PubMed] [CrossRef]

78. Chang, Y.-Y., A.-Y. Wang, and J. E. Cronan , Jr. 1994. Expression of Escherichia coli pyruvate oxidase (PoxB) depends on the sigma factor encoded by the rpoS (katF) gene. Mol. Microbiol. 11:1019–1028.[PubMed] [CrossRef]

79. Chang, Y.-Y., and J. E. Cronan , Jr. 1999. Membrane cyclopropane fatty acid content is a major factor in acid resistance of Escherichia coli. Mol. Microbiol. 33:249–259.[PubMed] [CrossRef]

80. Checroun, C., and C. Gutierrez. 2004. σ^S-dependent regulation of yehZYXW, which encodes a putative osmoprotectant ABC transporter of Escherichia coli. FEMS Microbiol. Lett. 236:221–226.[PubMed]

81. Chen, C. Y., L. Eckmann, S. J. Libby, F. C. Fang, S. Okamoto, M. F. Kagnoff, J. Fierer, and D. G. Guiney. 1996. Expression of Salmonella typhimurium rpoS and rpoS-dependent genes in the intracellular environment of eukaryotic cells. Infect. Immun. 64:4739–4743.[PubMed]

82. Chen, C. Y., N. A. Buchmeier, S. Libby, F. C. Fang, M. Krause, and D. G. Guiney. 1995. Central regulatory role for the RpoS sigma factor in expression of Salmonella dublin plasmid virulence genes. J. Bacteriol. 177:5303–5309.[PubMed]

83. Chen, G., C. L. Patten, and H. E. Schellhorn. 2004. Positive selection for loss of RpoS function in Escherichia coli. Mutat. Res. 554:193–203.[PubMed]

84. Chevance, F. F. V., and K. T. Hughes. 2008. Coordinating assembly of a bacterial macromolecular machine. Nat. Rev. Microbiol. 6:455–465.[PubMed] [CrossRef]

85. Cheville, A. M., K. W. Arnold, C. Buchrieser, C. M. Cheng, and C. W. Kaspar. 1996. rpoS regulation of acid, heat, and salt tolerance in Escherichia coli O157:H7. Appl. Environ. Microbiol. 62:1822–1824.[PubMed]

86. Chiang, S. M., and H. E. Schellhorn. 2010. Evolution of the RpoS regulon: origin of RpoS and the conservation of RpoS-dependent regulation in bacteria. J. Mol. Evol. 70:557–571.[PubMed] [CrossRef]

87. Chilcott, G. S., and K. T. Hughes. 2000. Coupling of flagellar gene expression to flagellar assembly in Salmonella enterica serovar typhimurium and Escherichia coli. Microbiol. Mol. Biol. Rev. 64:694–708. [CrossRef]

88. Claret, L., and J. Rouvière-Yaniv. 1997. Variation in HU composition during growth of E. coli: the heterodimer is required for long term survival. J. Mol. Biol. 273:93–104.[PubMed] [CrossRef]

89. Cohen-Or, I., Y. Shenhar, D. Biran, and E. Z. Ron. 2010. CspC regulates rpoS transcript levels and complements hfq deletions. Res. Microbiol. 161:694–700.[PubMed] [CrossRef]

90. Colland, F., M. Barth, R. Hengge-Aronis, and A. Kolb. 2000. Sigma factor selectivity of Escherichia coli RNA polymerase: a role for CRP, IHF and Lrp transcription factors. EMBO J. 19:3028–3037.[PubMed] [CrossRef]

91. Colland, F., N. Fujita, D. Kotlarz, A. Ishihama, and A. Kolb. 1999. Positioning of σ^S, the stationary phase σ factor, in Escherichia coli RNA polymerase-promoter open complexes. EMBO J. 18:4049–4059.[PubMed] [CrossRef]

92. Collet, A., P. Cosette, C. Beloin, J.-M. Ghigo, C. Rihouey, P. Lerouge, G. A. Junter, and T. Jouenne. 2008. Impact of rpoS deletion on the proteome of Escherichia coli grown planktonically and as biofilm. J. Proteome Res. 7:4659–4669.[PubMed] [CrossRef]

93. Conter, A., C. Gangneux, M. Suzanne, and C. Gutierrez. 2001. Survival of Escherichia coli during long-term starvation: effects of aeration, NaCl, and the rpoS and osmC gene products. Res. Microbiol. 152:17–26.[PubMed] [CrossRef]

94. Corona-Izquierda, F. P., and J. Membrillo-Hernández. 2002. A mutation in rpoS enhances biofilm formation in Escherichia coli during exponential phase of growth. FEMS Microbiol. Lett. 211:105–110.[PubMed] [CrossRef]

95. Costanzo, A., and S. Ades. 2006. Growth phase-dependent regulation of the extracytoplasmic stress factor, sigmaE by guanosine 3′,5′-bispyrophosphate (ppGpp). J. Bacteriol. 188:4627–4634.[PubMed] [CrossRef]

96. Coynault, C., V. Robbe-Saule, and F. Norel. 1996. Virulence and vaccine potential of Salmonella typhimurium mutants deficient in the expression of the RpoS (σ^S) regulon. Mol. Microbiol. 22:149–160.[PubMed] [CrossRef]

97. Cronan , J. E., Jr. 2002. Phospholipid modifications in bacteria. Curr. Opin. Microbiol. 5:202–205.[PubMed] [CrossRef]

98. Cunning, C., and T. Elliott. 1999. RpoS synthesis is growth rate regulated in Salmonella typhimurium but its turnover is not dependent on acetyl phosphate synthesis or PTS function. J. Bacteriol. 181:4853–4862.[PubMed]

99. Dailey, T. A., and H. A. Dailay. 2002. Identification of [2Fe-2S] clusters in microbial ferrochelatases. J. Bacteriol. 194:2460–2464. [CrossRef]

100. Davidson, C. J., A. P. White, and M. G. Surette. 2008. Evolutionary loss of the radar morphotype in Salmonella as a result of high mutations rates during laboratory passage. ISME J. 2:293–307.[PubMed] [CrossRef]

101. Davies, C. M., J. A. Long, M. Donald, and N. J. Ashbolt. 1995. Survival of fecal microorganisms in marine and freshwater sediments. Appl. Environ. Microbiol. 61:1888–1896.[PubMed]

102. Deighan, P., C. Pukhrambam, B. E. Nickels, and A. Hochschild. 2011. Initial transcribed region sequences influence the composition and functional properties of the bacterial elongation complex. Genes Dev. 25:77–88.[PubMed] [CrossRef]

103. Demple, B., J. Halbrook, and S. Linn. 1983. Escherichia coli xth mutants are hypersensitive to hydrogen peroxide. J. Bacteriol. 153:1079–1082.[PubMed]

104. Dersch, P., and R. Hengge-Aronis. 2003. Survival of environmental and host-associated stress, p. 37–73. In A. R. M. Coates (ed.), Stationary Phase and Disease. Cambridge University Press, Cambridge, UK.

105. Desmarais, T. R., H. M. Solo-Gabriele, and C. J. Palmer. 2002. Influence of soil on fecal indicator organisms in a tidally influenced subtropical environmental. Appl. Environ. Microbiol. 68:1165–1172.[PubMed] [CrossRef]

106. Ding, Q., S. Kusano, M. Villarejo, and A. Ishihama. 1995. Promoter selectivity control of Escherichia coli RNA polymerase by ionic strength: differential recognition of osmoregulated promoters by Eσ^D and Eσ^S holoenzymes. Mol. Microbiol. 16:649–656.[PubMed] [CrossRef]

107. Dodd, C. E., and T. G. Aldsworth. 2002. The importance of RpoS in the survival of bacteria through food processing. Int. J. Food Microbiol. 74:189–194.[PubMed] [CrossRef]

108. Domka, J., J. Lee, and J. M. Wood. 2006. YliH (BssR) and YceP (BssE) regulate Escherichia coli K-12 biofilm formation by influencing cell signaling. Appl. Environ. Microbiol. 72:2449–2459.[PubMed] [CrossRef]

109. Domka, J., J. Lee, T. Bansal, and T. K. Wood. 2007. Temporal gene expression in Escherichia coli K-12 biofilms. Environ. Microbiol. 9:332–346.[PubMed] [CrossRef]

110. Domínguez-Bernal, G., A. Tierrez, A. Bartolomé, S. Martínez-Pulgarin, F. J. Salguero, J. Antonio Orden, and R. de la Fuente. 2008. Salmonella enterica serovar Cholerasuis derivatives harbouring deletions in rpoS and phoP regulatory genes are attenuated in pigs, and survive and multiply in porcine intestinal macrophages and fibroblasts, respectively. Vet. Microbiol. 130:298–311.[PubMed] [CrossRef]

111. Dong, T., and H. E. Schellhorn. 2009. Control of RpoS in global gene expression of Escherichia coli in minimal medium. Mol. Genet. Genomics 281:19–31.[PubMed] [CrossRef]

112. Dong, T., and H. E. Schellhorn. 2009. Global effect of RpoS on gene expression in pathogenic Escherichia coli O157:H7 strain EDL933. BMC Genomics 3:349. [CrossRef]

113. Dong, T., M. G. Kirchhof, and H. E. Schellhorn. 2008. RpoS regulation of gene expression during exponential growth of Escherichia coli K12. Mol. Genet. Genomics 279:267–277.[PubMed] [CrossRef]

114. Dong, T., R. Yu, and H. E. Schellhorn. 2011. Antagonistic regulation of motility and transcriptome expression by RpoN and RpoS in Escherichia coli. Mol. Microbiol. 79:375–386.[PubMed] [CrossRef]

115. Dong, T., S. M. Chiang, C. Joyce, R. Yu, and H. E. Schellhorn. 2009. Polymorphism and selection of rpoS in pathogenic Escherichia coli. BMC Microbiol. 9:118.[PubMed] [CrossRef]

116. Dorel, C., P. Lejeune, and A. Rodrigue. 2006. The Cpx system of Escherichia coli, a strategic signaling pathway for confronting adverse conditions and for settling biofilm communities? Res. Microbiol. 157:306–314.[PubMed] [CrossRef]

117. Dorman, C. J. 2004. H-NS: a universal regulator for a dynamic genome. Nature Rev. Microbiol. 2:391–400.[PubMed] [CrossRef]

118. Dukan, S., and D. Touati. 1996. Hypochlorous acid stress in Escherichia coli: resistance, DNA damage, and comparison with hydrogen peroxide stress. J. Bacteriol. 178:6145–6150.[PubMed]

119. Dukan, S., and T. Nyström. 1998. Bacterial senescence: stasis results in increased and differential oxidation of cytoplasmic proteins leading to developmental induction of the heat shock regulon. Genes Dev. 12:3431–3441.[PubMed] [CrossRef]

120. Edwards, R. A., B. C. Matlock, B. J. Heffernan, and S. R. Maloy. 2001. Genomic analysis and growth-phase-dependent regulation of the SEF14 fimbriae of Salmonella enterica serovar Enteritidis. Microbiology 147:2705–2715.[PubMed]

121. Eguchi, Y., E. Ishii, K. Hata, and R. Utsumi. 2010. Regulation of acid resistance by connectors of two-component signal transduction systems in Escherichia coli. J. Bacteriol. 193:1222–1228.[PubMed] [CrossRef]

122. Eichel, J., Y. Y. Chang, D. Riesenberg, and J. E. Cronan. 1999. Effect of ppGpp on Escherichia coli cyclopropane fatty acid synthesis is mediated through the RpoS sigma factor (σ^S). J. Bacteriol. 181:572–576.[PubMed]

123. Eisenstark, A., M. J. Calcutt, M. Becker-Hapak, and A. Ivanova. 1999. Role of Escherichia coli rpoS and associated genes in defense against oxidative damage. Free Rad. Biol. Med. 21:975–993. [CrossRef]

124. Eisenstark, A. 1989. Bacterial genes involved in response to near-ultraviolet radiation. Adv. Genet. 26:99–147.[PubMed] [CrossRef]

125. England, P., L. F. Westblade, G. Karimova, V. Robbe-Saule, F. Norel, and A. Kolb. 2008. Binding of the unorthodox transcription activator, Crl, to the components of the transcription machinery. J. Biol. Chem. 283:33455–33464.[PubMed] [CrossRef]

126. Estrem, S. T., W. Ross, T. Gaal, Z. W. S. Chen, W. Niu, R. H. Ebright, and R. L. Gourse. 1999. Bacterial promoter architecture: subsite structure of UP elements and interactions with the carboxy-terminal domain of the RNA polymerase a subunit. Genes Dev. 13:2134–2147.[PubMed] [CrossRef]

127. Fang, F. C., and S. Rimsky. 2008. New insights into transcriptional regulation by H-NS. Curr. Opin. Microbiol. 11:113–120.[PubMed] [CrossRef]

128. Fang, F. C., S. J. Libby, N. A. Buchmeier, P. C. Loewen, J. Switala, J. Harwood, and D. G. Guiney. 1992. The alternative σ factor KatF (RpoS) regulates Salmonella virulence. Proc. Natl. Acad. Sci. USA 89:11978–11982.[PubMed] [CrossRef]

129. Farewell, A., K. Kvint, and T. Nyström. 1998. Negative regulation by RpoS: a case of sigma factor competition. Mol. Microbiol. 29:1039–1051.[PubMed] [CrossRef]

130. Farewell, A., K. Kvint, and T. Nyström. 1998. uspB, a new σ^S-regulated gene in Escherichia coli which is required for stationary-phase resistance to ethanol. J. Bacteriol. 180:6140–6147.[PubMed]

131. Felipe-López, A., and M. Hensel. 2011. Bacterial responses to the host cell, p. 385–398. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses. ASM Press, Washington, DC.

132. Fender, A., J. Elf, K. Hampel, B. Zimmermann, and E. G. H. Wagner. 2010. RNAs actively cycle on the Sm-like protein Hfq. Genes Dev. 24:2621–2626.[PubMed] [CrossRef]

133. Feng, G., H. C. T. Tsui, and M. E. Winkler. 1996. Depletion of the cellular amounts of the MutS and MutH methyl-directed mismatch repair proteins in stationary phase Escherichia coli K-12 cells. J. Bacteriol. 178:2388–2396.[PubMed]

134. Ferenci, T. 2001. Hungry bacteria—definition and properties of a nutritional state. Environ. Microbiol. 3:605–611.[PubMed] [CrossRef]

135. Ferenci, T. 2003. What is driving the acquisition of mutS and rpoS polymorphisms in Escherichia coli? Trends Microbiol. 11:457–461.[PubMed] [CrossRef]

136. Ferenci, T. 2008. The spread of a beneficial mutation in experimental bacterial populations: the influence of the environment and genotype on the fixation of rpoS mutations. Heredity 100:446–452.[PubMed] [CrossRef]

137. Ferrandez, A., J. L. García, and E. Díaz. 2000. Transcriptional regulation of the divergent paa catalytic operons for phenylacetic acid degradation in Escherichia coli. J. Biol. Chem. 275:12214–12222.[PubMed] [CrossRef]

138. Finkel, S. E., and R. Kolter. 1999. Evolution of microbial diversity during prolonged starvation. Proc. Natl. Acad. Sci. USA 96:4023–4027.[PubMed] [CrossRef]

139. Finkel, S. E. 2006. Long-term survival during stationary phase: evolution and the GASP phenotype. Nat. Rev. Microbiol. 4:113–120.[PubMed] [CrossRef]

140. Fischer, D., A. Teich, P. Neubauer, and R. Hengge-Aronis. 1998. The general stress sigma factor σ^S of Escherichia coli is induced during diauxic shift from glucose to lactose. J. Bacteriol. 180:6203–6206.[PubMed]

141. Foster, J. W. 2004. Escherichia coli acid resistance: tales of an amateur acidophile. Nat. Rev. Microbiol. 2:898–907.[PubMed] [CrossRef]

142. Foster, P. L. 2007. Stress-induced mutagenesis in bacteria. Crit. Rev. Biochem. Mol. Biol. 42:373–397.[PubMed] [CrossRef]

143. Francez-Charlot, A., J. Frunzke, and J. A. Vorholt. 2011. The general stress response in alpha-proteobacteria, p. 291–300. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses. ASM Press, Washington, DC.

144. Franze de Fernandez, M. T., L. Eoyang, and J. T. August. 1968. Factor fraction required for the synthesis of bacteriophage Qb RNA. Nature (London) 219:588–590.[PubMed] [CrossRef]

145. Frederiksson, A., M. Ballesteros, C. N. Peterson, O. Persson, T. J. Silhavy, and T. Nyström. 2007. Decline in ribosomal fidelity contributes to the accumulation and stabilization of master stress response regulator sigmaS upon carbon starvation. Genes Dev. 21:862–874.[PubMed] [CrossRef]

146. Freire, P., R. N. Moreira, and C. M. Arraiano. 2009. BolA inhibits cell elongation and regulates MreB expression levels. J. Mol. Biol. 385:1345–1351.[PubMed] [CrossRef]

147. Frisch, R. L., Y. Su, P. C. Thornton, J. L. Gibson, S. M. Rosenberg, and P. J. Hastings. 2010. Separate DNA Pol II- and Pol IV-dependent pathways of stress-induced mutation during double-strand-break repair in Escherichia coli are controlled by RpoS. J. Bacteriol. 192:4694–4700.[PubMed] [CrossRef]

148. Frye, J., J. E. Karlinsey, H. R. Felise, B. Marzolf, N. Dowidar, M. McClelland, and K. T. Hughes. 2006. Identification of new flagellar genes of Salmonella enterica serovar typhimurium. J. Bacteriol. 188:2233–2243.[PubMed] [CrossRef]

149. Fuentes, J. A., M. R. Jofré, N. A. Villagra, and G. C. Mora. 2009. RpoS- and Crp-dependent transcriptional control of Salmonella typhi taiA and hlyE genes: role of environmental conditions. Res. Microbiol. 160:800–808.

150. Fux, C. A., J. W. Costerton, P. S. Stewart, and P. Stoodley. 2005. Survival strategies of infectious biofilms. Trends Microbiol. 13:34–40.[PubMed] [CrossRef]

151. Gaal, T., M. J. Mandel, T. J. Silhavy, and R. L. Gourse. 2006. Crl facilitates RNA polymerase holoenzyme formation. J. Bacteriol. 188:7966–7970.[PubMed] [CrossRef]

152. Gaal, T., W. Ross, S. T. Estrem, L. H. Nguyen, R. R. Burgess, and R. L. Gourse. 2001. Promoter recognition and discrimination by Eσ^S RNA polymerase. Mol. Microbiol. 42:939–954.[PubMed] [CrossRef]

153. Gentry, D. R., V. J. Hernandez, L. H. Nguyen, D. B. Jensen, and M. Cashel. 1993. Synthesis of the stationary-phase sigma factor s^S is positively regulated by ppGpp. J. Bacteriol. 175:7982–7989.[PubMed]

154. Georgellis, D., O. Kwon, and E. C. C. Lin. 2001. Quinones as the redox signal for the Arc Two-component system of bacteria. Science 292:2314–2315.[PubMed] [CrossRef]

155. Germer, J., A. Muffler, and R. Hengge-Aronis. 1998. Trehalose is not relevant for in vivo activity of σ^S-containing RNA polymerase in Escherichia coli. J. Bacteriol. 180:1603–1606.[PubMed]

156. Germer, J., G. Becker, M. Metzner, and R. Hengge-Aronis. 2001. Role of activator site position and a distal UP-element half-site for sigma factor selectivity at a CRP/H-NS activated σ^S-dependent promoter in Escherichia coli. Mol. Microbiol. 41:705–716.[PubMed] [CrossRef]

157. Giangrossi, M., S. Zattoni, A. Tramonti, D. De Biase, and M. Falconi. 2005. Antagonistic role of H-NS and GadX in the regulation of the glutamate decarboxylase-dependent acid resistance system in Escherichia coli. J. Biol. Chem. 280:21498–21505.[PubMed] [CrossRef]

158. Girgis, H. S., Y. Liu, W. S. Ryu, and S. Tavazoie. 2007. A comprehensive genetic characterization of bacterial motility. PLoS Genet. 3:e154. [CrossRef]

159. Gong, L., K. Takayama, and S. Kjelleberg. 2002. Role of spoT-dependent ppGpp accumulation in the survival of light-exposed starved bacteria. Microbiology 148:559–570.[PubMed]

160. Gordia, S., and C. Gutierrez. 1996. Growth-phase-dependent expression of the osmotically inducible gene osmC of Escherichia coli K-12. Mol. Microbiol. 19:729–736.[PubMed] [CrossRef]

161. Gottesman, S. 2005. Micros for microbes: non-coding regulatory RNAs in bacteria. Trends Genet. 21:399–404.[PubMed] [CrossRef]

162. Gottesman, S. 2011. Roles of mRNA stability, translational regulation, and small RNAs in stress response regulation, p. 59–74. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses. ASM Press, Washington, DC.

163. Gourse, R. L., W. Ross, and T. Gaal. 2000. UPs and downs in bacterial transcription initiation: the role of the alpha subunit of RNA polymerase in promoter recognition. Mol. Microbiol. 37:687–695.[PubMed] [CrossRef]

164. Grainger, D. C., M. D. Goldberg, D. J. Lee, and S. J. Busby. 2008. Selective repression by Fis and H-NS at the Escherichia coli dps promoter. Mol. Microbiol. 68:1366–1377.[PubMed] [CrossRef]

165. Gralla, J. D., and D. R. Vargas. 2006. Potassium glutamate as a transcriptional inhibitor during bacterial osmoregulation. EMBO J. 25:1515–1521.[PubMed] [CrossRef]

166. Gralla, J. D., and Y. X. Huo. 2008. Remodeling and activation of Escherichia coli RNA polymerase by osmolytes. Biochemistry 47:13189–12196.[PubMed] [CrossRef]

167. Grant, R. A., D. H. Gilman, S. E. Finkel, R. Kolter, and J. M. Hogle. 1998. The crystal structure of Dps, a ferritin homolog that binds and protects DNA. Nat. Struct. Biol. 5:294–303.[PubMed] [CrossRef]

168. Grigorova, I. R., N. J. Phleger, V. K. Mutalik, and C. A. Gross. 2006. Insights into transcriptional regulation and sigma competition from an equilibrium model of RNA polymerase binding to DNA. Proc. Natl. Acad. Sci. USA 103:5332–5337.[PubMed] [CrossRef]

169. Gruber, T., and C. A. Gross. 2003. Multiple sigma subunits and the partitioning of bacterial transcription space. Annu. Rev. Microbiol. 57:441–466.[PubMed] [CrossRef]

170. Gruber, T. M., and D. A. Bryant. 1997. Molecular systematic studies of eubacteria, using σ⁷⁰-type sigma factors of group 1 and 2. J. Bacteriol. 179:1734–1747.[PubMed]

171. Gualdi, L., L. Tagliabue, and P. Landini. 2007. Biofilm formation-gene expression relay system in Escherichia coli: modulation of σ^S-dependent gene expression by the CsgD regulatory protein via σ^S protein stabilization. J. Bacteriol. 189:8034–8043.[PubMed] [CrossRef]

172. Gulig, P. A., H. Danbar, D. G. Guiney, A. J. Lax, F. Norel, and M. Rhen. 1993. Molecular analysis of spv virulence genes of the Salmonella virulence plasmids. Mol. Microbiol. 7:825–830.[PubMed] [CrossRef]

173. Gupta, R. S. 2000. The phylogeny of proteobacteria: relationships to other eubacterial phyla and eukaryotes. FEMS Microbiol. Rev. 24:367–402.[PubMed] [CrossRef]

174. Gutierrez, C., and J. C. Devedjian. 1991. Osmotic induction of gene osmC expression in Escherichia coli. J. Mol. Biol. 220:959–973.[PubMed] [CrossRef]

175. Gutierrez, C., J. Barondess, C. Manoil, and J. Beckwith. 1987. The use of transposon TnphoA to detect genes for cell envelope proteins subject to a common regulatory stimulus. J. Mol. Biol. 195:289–297.[PubMed] [CrossRef]

176. Gutierrez, C., S. Gordia, and S. Bonnassie. 1995. Characterization of the osmotically inducible gene osmE of Escherichia coli K-12. Mol. Microbiol. 16:553–563.[PubMed] [CrossRef]

177. Gérard, F., A. M. Dri, and P. L. Moreau. 1999. Role of Escherichia coli RpoS, LexA, and H-NS global regulators in metabolism and survival under aerobic, phosphate-starvation conditions. Microbiology 145:1547–1562.[PubMed] [CrossRef]

178. Hagiwara, D., T. Yamashino, and T. Mizuno. 2004. A genome-wide view of the Escherichia coli BasS-BasR two-component system implicated in iron-responses. Biosci. Biotechnol. Biochem. 68:1758–1768.[PubMed] [CrossRef]

179. Hales, L. M., and H. A. Shuman. 1999. Legionella pneumophila rpoS is required for growth within Acanthamoeba castellanii. J. Bacteriol. 181:4879–4889.[PubMed]

180. Hammar, M., A. Arnquist, Z. Bian, A. Olsén, and S. Normark. 1995. Expression of two csg operons is required for production of fibronectin- and Congo red-binding curli polymers in Escherichia coli K-12. Mol. Microbiol. 18:661–670.[PubMed] [CrossRef]

181. Haugen, S. P., M. B. Berkmen, W. Ross, and R. L. Gourse. 2006. rRNA promoter regulation by nonoptimal binding of sigma region 1.2: an additional recognition element for RNA polymerase. Cell 125:1069–1082.[PubMed] [CrossRef]

182. Heeb, S., C. Valverde, C. Gigot-Bonnefoy, and D. Haas. 2005. Role of the stress sigma factor RpoS in GacA/RsmA-controlled secondary metabolism and resistance to oxidative stress in Pseudomonas fluorescens CHA0. FEMS Microbiol. Lett. 243:251–258.[PubMed] [CrossRef]

183. Heiskanen, P., S. Taira, and M. Rhen. 1994. Role of rpoS in the regulation of Salmonella plasmid virulence (spv) genes. FEMS Microbiol. Lett. 123:125–130.[PubMed] [CrossRef]

184. Helmann, J. D. 2011. Regulation by alternative sigma factors, p. 31–44. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses, 2nd ed. ASM Press, Washington , DC.

185. Hengge, R., and K. Turgay. 2009. Proteolysis in prokaryotes—from molecular machines to a systems perspective. Res. Microbiol. 160:615–617.[PubMed] [CrossRef]

186. Hengge, R. 2008. The two-component network and the general stress sigma factor RpoS (σ^S) in Escherichia coli. Adv. Exp. Med. Biol. 631:40–53.[PubMed] [CrossRef]

187. Hengge, R. 2009. Principles of cyclic-di-GMP signaling. Nat. Rev. Microbiol. 7:263–273.[PubMed] [CrossRef]

188. Hengge, R. 2009. Proteolysis of σ^S (RpoS) and the general stress response in Escherichia coli. Res. Microbiol. 160:667–676.[PubMed] [CrossRef]

189. Hengge, R. 2010. Role of cyclic Di-GMP in the regulatory networks of Escherichia coli, p. 230--252. In A. J. Wolfe and K. L. Visick (ed.), The Second Messenger Cyclic Di-GMP. ASM Press, Washington, DC.

190. Hengge, R. 2011. The general stress response in Gram-negative bacteria, p. 251–289. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses, 2nd ed. ASM Press, Washington, DC.

191. Hengge-Aronis, R., W. Klein, R. Lange, M. Rimmele, and W. Boos. 1991. Trehalose synthesis genes are controlled by the putative sigma factor encoded by rpoS and are involved in stationary phase thermotolerance in Escherichia coli. J. Bacteriol. 173:7918–7924.[PubMed]

192. Hengge-Aronis, R. 1993. Survival of hunger and stress: the role of rpoS in stationary phase gene regulation in Escherichia coli. Cell 72:165–168.[PubMed] [CrossRef]

193. Hengge-Aronis, R. 1996. Back to log phase: σ^S as a global regulator in the osmotic control of gene expression in Escherichia coli. Mol. Microbiol. 21:887–893.[PubMed] [CrossRef]

194. Hengge-Aronis, R. 1996. Regulation of gene expression during entry into stationary phase, p. 1497–1512. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology. ASM Press, Washington, DC.

195. Hengge-Aronis, R. 2000. The general stress response in Escherichia coli, p. 161–178. In G. Storz and R. Hengge-Aronis (ed.), Bacterial Stress Responses. ASM Press, Washington, DC.

196. Hengge-Aronis, R. 2002. Signal transduction and regulatory mechanisms involved in control of the σ^S subunit of RNA polymerase in Escherichia coli. Microbiol. Mol. Biol. Rev. 66:373–395.[PubMed] [CrossRef]

197. Hengge-Aronis, R. 2002. Stationary phase gene regulation: what makes an Escherichia coli promoter σ^S-dependent? Curr. Opin. Microbiol. 5:591–595.[PubMed] [CrossRef]

198. Hersh, M. N., R. G. Ponder, P. J. Hastings, and S. M. Rosenberg. 2004. Adaptive mutation and amplification in Escherichia coli: two pathways of genome adaptation under stress. Res. Microbiol. 155:352–359.[PubMed] [CrossRef]

199. Heuveling, J., A. Possling, and R. Hengge. 2008. A role for Lon protease in the control of the acid resistance genes of Escherichia coli. Mol. Microbiol. 69:534–547.[PubMed] [CrossRef]

200. Heydorn, A., B. K. Ersbøll, M. Hentzer, M. R. Parsek, M. Givskov, and S. Molin. 2000. Experimental reproducibility in flow-chamber biofilms. Microbiology 146:2409–2415.[PubMed]

201. Hirakawa, H., Y. Inazumi, T. Masaki, T. Hirata, and A. Yamaguchi. 2005. Indole induces the expression of multidrug exporter genes in Escherichia coli. Mol. Microbiol. 55:1113–1126.[PubMed] [CrossRef]

202. Hiratsu, K., H. Shinagawa, and K. Makino. 1995. Mode of promoter recognition by the Escherichia coli RNA polymerase holoenzyme containing the σ^S subunit: identification of the recognition sequence of the fic promoter. Mol. Microbiol. 18:841–850.[PubMed] [CrossRef]

203. Hirsch, M., and T. Elliott. 2002. Role of ppGpp in rpoS stationary phase regulation in Escherichia coli. J. Bacteriol. 184:5077–5087.[PubMed] [CrossRef]

204. Holland, A.-M., and P. N. Rather. 2008. Evidence for extracellular control of RpoS proteolysis in Escherichia coli. FEMS Microbiol. Lett. 286:50–59.[PubMed] [CrossRef]

205. Holland, K., S. J. Busby, and G. S. Lloyd. 2007. New targets for the cyclic AMP receptor protein in the Escherichia coli K-12 genome. FEMS Microbiol. Lett. 274:89–94.[PubMed] [CrossRef]

206. Hong, W., W. Jiao, J. Hu, J. Zhang, C. Liu, X. Fu, D. Shen, B. Xia, and Z. Chang. 2005. Periplasmic protein HdeA exhibits chaperone-like activity exclusively within stomach pH range by transforming into disordered conformation. J. Biol. Chem. 280:27029–27034.[PubMed] [CrossRef]

207. Hovel-Miner, G., S. Pampou, S. P. Faucher, M. Clarke, I. Morozova, P. Morozov, J. J. Russo, H. A. Shuman, and S. Klachikov. 2009. σ^S controls multiple pathways associated with intracellular multiplication of Legionella pneumophila. J. Bacteriol. 191:2461–2473.[PubMed] [CrossRef]

208. Huo, Y. X., A. Z. Rosenthal, and J. D. Gralla. 2008. General stress response signalling: unwrapping transcription complexes by DNA relaxation via the sigma38 C-terminal domain. Mol. Microbiol. 70:369–378.[PubMed] [CrossRef]

209. Huynen, M. A., C. A. Spronk, T. Gabaldon, and B. Snel. 2005. Combining data from genomes, Y2H and 3D structure indicates that BolA is a reductase interacting with a glutaredoxin. FEBS Lett. 579:591–596.[PubMed] [CrossRef]

210. Hülsmann, A., T. M. Rosche, I. S. Kong, H. M. Hassan, D. M. Beam, and J. D. Oliver. 2003. RpoS-dependent stress response and exoenzyme production in Vibrio vulnificus. Appl. Environ. Microbiol. 69:6114–6120.[PubMed] [CrossRef]

211. Ihssen, J., and T. Egli. 2004. Specific growth rate and not cell density controls the general stress response in Escherichia coli. Microbiology 150:1637–1648.[PubMed] [CrossRef]

212. Ihssen, J., E. Grasselli, C. Bassin, P. François, J.-C. Piffaretti, W. Köster, J. Schrenzel, and T. Egli. 2007. Comparative genomic hybridization and physiological characterization of environmental isolates indicate that significant (eco-)physiological properties are highly conserved in the species Escherichia coli. Microbiology 153:2052–2066.[PubMed] [CrossRef]

213. Ilag, L. L., L. F. Westblade, C. Deshayes, A. Kolb, S. J. Busby, and C. V. Robinson. 2004. Mass spectrometry of Escherichia coli RNA polymerase: interactions of the core enzyme with sigma70 and Rsd protein. Structure (Camb.) 12:269–275.[PubMed]

214. Ishihama, A. 2000. Functional modulation of Escherichia coli RNA polymerase. Annu. Rev. Microbiol. 54:499–518.[PubMed] [CrossRef]

215. Ito, A., a. Taniuchi, T. May, K. Kawata, and S. Okabe. 2009. Increased antibiotic resistance of Escherichia coli in mature biofilms. Appl. Environ. Microbiol. 75:4093–4100.[PubMed] [CrossRef]

216. Ito, A., T. May, K. Kawata, and S. Okabe. 2008. Significance of rpoS during maturation of Escherichia coli biofilms. Biotechnol. Bioeng. 99:1462–1471.[PubMed] [CrossRef]

217. Ivanova, A., C. Miller, G. Glinsky, and A. Eisenstark. 1994. Role of rpoS(katF) in oxyR-independent regulation of hydroperoxidase I in Escherichia coli. Mol. Microbiol. 12:571–578.[PubMed] [CrossRef]

218. Ivanova, A., M. Renshaw, R. V. Guntaka, and A. Eisenstark. 1992. DNA base sequence variability in katF (putative sigma factor) gene of Escherichia coli. Nucleic Acids Res. 20:5479–5480.[PubMed] [CrossRef]

219. Ivanova, A. B., G. V. Glinsky, and A. Eisenstark. 1997. Role of RpoS regulon in resistance to oxidative stress and near-UV radiation in Delta-oxyR suppressor mutants of Escherichia coli. Free Radical Biol. Med. 23:627–636.[PubMed] [CrossRef]

220. Jasieki, J., and G. Wegrzyn. 2003. Growth-rate dependent RNA polyadenylation in Escherichia coli. EMBO Rep. 4:172–177.[PubMed] [CrossRef]

221. Jenal, U., and J. Malone. 2006. Mechanisms of cyclic-di-GMP signaling in bacteria. Annu. Rev. Genet. 40:385–407.[PubMed] [CrossRef]

222. Jenal, U., and R. Hengge-Aronis. 2003. Regulation by proteolysis in bacterial cells. Curr. Opin. Microbiol. 6:163–172.[PubMed] [CrossRef]

223. Jenal, U. 2004. Cyclic di-guanosine-monophosphate comes of age: a novel secondary messenger involved in modulating cell surface structures in bacteria? Curr. Opin. Microbiol. 7:185–191.[PubMed] [CrossRef]

224. Jenkins, D. E., J. E. Schultz, and A. Matin. 1988. Starvation-induced cross-protection against heat or H₂O₂ challenge in Escherichia coli. J. Bacteriol. 170:3910–3914.[PubMed]

225. Jenkins, D. E., S. A. Chaisson, and A. Matin. 1990. Starvation-induced cross-protection against osmotic challenge in Escherichia coli. J. Bacteriol. 172:2779–2781.[PubMed]

226. Jin, D. J., and J. E. Cabrera. 2006. Coupling the distribution of RNA polymerase to global gene regulation and the dynamic structure of the bacterial nucleoid in Escherichia coli. J. Struct. Biol. 156:284–291.[PubMed] [CrossRef]

227. Jishage, M., A. Iwata, S. Ueda, and A. Ishihama. 1996. Regulation of RNA polymerase sigma subunit synthesis in Escherichia coli: intracellular levels of four species of sigma subunit under various growth conditions. J. Bacteriol. 178:5447–5451.[PubMed]

228. Jishage, M., and A. Ishihama. 1998. A stationary phase protein in Escherichia coli with binding activity to the major sigma subunit of RNA polymerase. Proc. Natl. Acad. Sci. USA 95:4953–4958.[PubMed] [CrossRef]

229. Jishage, M., and A. Ishihama. 1999. Transcriptional organization and in vivo role of the Escherichia coli rsd gene, encoding the regulator of RNA polymerase sigma D. J. Bacteriol. 181:3768–3776.[PubMed]

230. Jishage, M., K. Kvint, V. Shingler, and T. Nyström. 2002. Regulation of sigma factor competition by the alarmone ppGpp. Genes Dev. 16:1260–1270.[PubMed] [CrossRef]

231. Jishage, M. l., and A. Ishihama. 1997. Variation in RNA polymerase sigma subunit composition within different stocks of Escherichia coli W3110. J. Bacteriol. 179:959–963.[PubMed]

232. Josaitis, C. A., T. Gaal, and R. L. Gourse. 1995. Stringent control and growth-rate-dependent control have nonidentical promoter sequence requirements. Proc. Natl. Acad. Sci. USA 92:1117–1121.[PubMed] [CrossRef]

233. Jubelin, G., A. Vianney, C. Beloin, J. M. Ghigo, J. C. Lazzaroni, P. Lejeune, and C. Dorel. 2005. CpxR/OmpR interplay regulates curli gene expression in response to osmolarity in Escherichia coli. J. Bacteriol. 187:2038–2049.[PubMed] [CrossRef]

234. Jung, J. U., C. Gutierrez, F. Martin, M. Ardourel, and M. Villarejo. 1990. Transcription of osmB, a gene encoding an Escherichia coli lipoprotein, is regulated by dual signals. J. Biol. Chem. 265:10574–10581.[PubMed]

235. Kaasen, I., J. McDougall, and A. R. Strøm. 1994. Analysis of the otsBA operon for osmoregulatory trehalose synthesis in Escherichia coli and homology of the OtsA and OtsB proteins to the yeast trehalose-6-phosphate synthase/phosphatase complex. Gene 145:9–15.[PubMed] [CrossRef]

236. Kaasen, I., P. Falkenberg, O. B. Styrvold, and A. R. Strøm. 1992. Molecular cloning and physical mapping of the otsBA genes, which encode the osmoregulatory trehalose pathway of Escherichia coli: evidence that transcription is activated by KatF (AppR). J. Bacteriol. 174:889–898.[PubMed]

237. Kabir, M. S., T. Sagara, T. Oshima, Y. Kawagoe, H. Mori, R. Tsunedomi, and M. Yamada. 2004. Effects of mutations in the rpoS gene on cell viability and global gene expression under nitrogen starvation in Escherichia coli. Microbiology 150:2543–2553.[PubMed] [CrossRef]

238. Kader, A., R. Simm, U. Gerstel, M. Morr, and U. Römling. 2006. Hierarchical involvement of various GGDEF domain proteins in rdar morphotype development of Salmonella enterica serovar typhimurium. Mol. Microbiol. 60:602–616.[PubMed] [CrossRef]

239. Kalir, S., and U. Alon. 2004. Using a quantitative blueprint to reprogram the dynamics of the flagella gene network. Cell 117:713–720.[PubMed] [CrossRef]

240. Kaper, J. B., J. P. Nataro, and H. L. T. Mobley. 2004. Pathogenic Escherichia coli. Nat. Rev. Microbiol. 2:123–140.[PubMed] [CrossRef]

241. Kern, R., A. Malki, J. Abdallah, J. Tagourti, and G. Richarme. 2007. Escherichia coli HdeB in an acid stress chaperone. J. Bacteriol. 189:603–610.[PubMed] [CrossRef]

242. Kim, E. Y., M. S. Shin, J. H. Rhee, and H. E. Choy. 2004. Factors influencing preferential utilization of RNA polymerase containing sigma-38 in stationary-phase gene expression in Escherichia coli. J. Microbiol. 42:103–110.[PubMed]

243. Kim, K.-S., R. Manasherob, and S. N. Cohen. 2009. YmdB: a stress-responsive ribonuclease-binding regulator of E. coli RNase III activity. Genes Dev. 22:3497–3508. [CrossRef]

244. King, T., A. Ishihama, A. Kori, and T. Ferenci. 2004. A regulatory trade-off as a source of strain variation in the species Escherichia coli. J. Bacteriol. 186:5614–5620.[PubMed] [CrossRef]

245. Klauck, E., J. Böhringer, and R. Hengge-Aronis. 1997. The LysR-like regulator LeuO in Escherichia coli is involved in the translational regulation of rpoS by affecting the expression of the small regulatory DsrA-RNA. Mol. Microbiol. 25:559–569.[PubMed] [CrossRef]

246. Klauck, E., M. Lingnau, and R. Hengge-Aronis. 2001. Role of the response regulator RssB in σ^S recognition and initiation of σ^S proteolysis in Escherichia coli. Mol. Microbiol. 40:1381–1390.[PubMed] [CrossRef]

247. Klein, A. H., A. Shulla, S. A. Reimann, D. H. Keating, and A. J. Wolfe. 2007. The intracellular concentration of acetyl phosphate in Escherichia coli is sufficient for direct phosphorylation of two-component response regulators. J. Bacteriol. 189:5574–5581.[PubMed] [CrossRef]

248. Ko, M., and C. Park. 2000. Two novel flagellar components and H-NS are involved in the motor function of Escherichia coli. J. Mol. Biol. 303:371–382.[PubMed] [CrossRef]

249. Kolb, A., D. Kotlarz, S. Kusano, and A. Ishihama. 1995. Selectivity of the E. coli RNA polymerase Eσ³⁸ for overlapping promoters and ability to support CRP activation. Nucleic Acids Res. 23:819–826.[PubMed] [CrossRef]

250. Kolmsee, T., and R. Hengge. 2011. Rare codons play a positive role in the expression of the stationary phase sigma factor RpoS (σ^S) in Escherichia coli. RNA Biol. 8:913–921.

251. Kolter, R., D. A. Siegele, and A. Tormo. 1993. The stationary phase of the bacterial life cycle. Annu. Rev. Microbiol. 47:855–874.[PubMed] [CrossRef]

252. Kowarz, L., C. Coynault, V. Robbe-Saule, and F. Norel. 1994. The Salmonella typhimurium katF (rpoS) gene: cloning, nucleotide sequence, and regulation of spvR and spvABCD virulence plasmid genes. J. Bacteriol. 176:6852–6860.[PubMed]

253. Kowarz, L., V. Robbe-Saule, and F. Norel. 1996. Identification of cis-acting DNA sequences involved in the transcription of the virulence regulatory gene spvR in Salmonella typhimurium. Mol. Gen. Genet. 251:225–235.[PubMed]

254. Kress, W., Z. Maglica, and E. Weber-Ban. 2009. Clp chaperone-proteases: structure and function. Res. Microbiol. 160:618–628.[PubMed] [CrossRef]

255. Krin, E., A. Danchin, and O. Soutourina. 2010. Decrypting the H-NS-dependent regulatory cascade of acid stress resistance in Escherichia coli. BMC Microbiol. 10:273.[PubMed] [CrossRef]

256. Krin, E., A. Danchin, and O. Soutourina. 2010. RcsB plays a central role in H-NS-dependent regulation of motility and acid stress resistance in Escherichia coli. Res. Microbiol. 161:363–371.[PubMed] [CrossRef]

257. Krogfelt, K. A., M. Hjulgaard, K. Sørensen, P. S. Cohen, and M. Givskov. 2000. rpoS gene function is a disadvantage for Escherichia coli BJ4 during competitive colonization of the mouse large intestine. Infect. Immun. 68:2518–2524.[PubMed] [CrossRef]

258. Kusano, S., Q. Q. Ding, N. Fujita, and A. Ishihama. 1996. Promoter selectivity of Escherichia coli RNA polymerase Eσ⁷⁰ and Eσ³⁸ holoenzymes – effect of DNA supercoiling. J. Biol. Chem. 271:1998–2004.[PubMed] [CrossRef]

259. Kuznedelov, K., L. Minakhin, A. Niedziela-Majka, S. L. Dove, D. Rogulja, B. E. Nickels, A. Hochschild, T. Heyduk, and K. Severinov. 2002. A role for interaction of the RNA polymerase flap domain with the s subunit in promoter recognition. Science 295:855–857.[PubMed] [CrossRef]

260. Laaberki, M. H., N. Janabi, E. Oswald, and F. Repoila. 2006. Concert of regulators to switch on LEE expression in enterohemorrhagic Escherichia coli O157:H7: interplay between Ler, GrlA, H-NS and RpoS. Int. J. Med. Microbiol. 296:197–210.[PubMed] [CrossRef]

261. Lacour, S., A. Kolb, and P. Landini. 2003. Nucleotides from −16 to −12 determine specific promoter recognition by bacterial σ^S-RNA polymerase. J. Biol. Chem. 278:37160–37168.[PubMed] [CrossRef]

262. Lacour, S., and P. Landini. 2004. σ^S-dependent gene expression at the onset of stationary phase in Escherichia coli: function of σ^S-dependent genes and identification of their promoter sequences. J. Bacteriol. 186:7186–7195.[PubMed] [CrossRef]

263. Landini, P., and M. R. Volkert. 2000. Regulatory responses of the adaptive response to alkylation damage: a simple regulon with complex regulatory features. J. Bacteriol. 182:6543–6549.[PubMed] [CrossRef]

264. Landini, P., L. I. Hajec, L. H. Nguyen, R. R. Burgess, and M. R. Volkert. 1996. The leucine-responsive regulatory protein (Lrp) acts as a specific repressor for σ^S-dependent transcription of the Escherichia coli aidB gene. Mol. Microbiol. 20:947–955.[PubMed] [CrossRef]

265. Landini, P. 2009. Cross-talk mechanism in biofilm formation and responses to environmental and physiological stress in Escherichia coli. Res. Microbiol. 160:259–266.[PubMed] [CrossRef]

266. Lange, R., and R. Hengge-Aronis. 1991. Growth phase-regulated expression of bolA and morphology of stationary phase Escherichia coli cells is controlled by the novel sigma factor σ^S (rpoS). J. Bacteriol. 173:4474–4481.[PubMed]

267. Lange, R., and R. Hengge-Aronis. 1991. Identification of a central regulator of stationary-phase gene expression in Escherichia coli. Mol. Microbiol. 5:49–59.[PubMed] [CrossRef]

268. Lange, R., and R. Hengge-Aronis. 1994. The cellular concentration of the σ^S subunit of RNA-polymerase in Escherichia coli is controlled at the levels of transcription, translation and protein stability. Genes Dev. 8:1600–1612.[PubMed] [CrossRef]

269. Lange, R., and R. Hengge-Aronis. 1994. The nlpD gene is located in an operon with rpoS on the Escherichia coli chromosome and encodes a novel lipoprotein with a potential function in cell wall formation. Mol. Microbiol. 13:733–743.[PubMed] [CrossRef]

270. Lange, R., D. Fischer, and R. Hengge-Aronis. 1995. Identification of transcriptional start sites and the role of ppGpp in the expression of rpoS, the structural gene for the s^S subunit of RNA-polymerase in Escherichia coli. J. Bacteriol. 177:4676–4680.[PubMed]

271. Lange, R., M. Barth, and R. Hengge-Aronis. 1993. Complex transcriptional control of the σ^S-dependent stationary phase-induced and osmotically regulated osmY (csi-5) gene suggests novel roles for Lrp, cyclic AMP (cAMP) receptor protein-cAMP complex and integration host factor in the stationary phase response of Escherichia coli. J. Bacteriol. 175:7910–7917.[PubMed]

272. Laurie, A. D., L. M. Bernardo, C. C. Sze, E. Skarfstad, A. Szalewska-Palasz, T. Nyström, and V. Shingler. 2002. The role of the alarmone (p)ppGpp in sigma N competition for core RNA polymerase. J. Biol. Chem. 278:1494–1503.[PubMed] [CrossRef]

273. Layton, J. C., and P. L. Foster. 2003. Error-prone DNA polymerase IV is controlled by the stress-response sigma factor, RpoS, in Escherichia coli. Mol. Microbiol. 50:549–561.[PubMed] [CrossRef]

274. Leang, C., and D. R. Lovley. 2005. Regulation of two highly similar genes, omcB and omcC, in a 10 kb chromosomal duplication in Geobacter sulfurreducens. Microbiology 151:1761–1767.[PubMed] [CrossRef]

275. Lease, R. A., and M. Belfort. 2000. A trans-acting RNA as a control switch in Escherichia coli: DsrA modulates function by forming alternative structures. Proc. Natl. Acad. Sci. USA 97:9919–9924.[PubMed] [CrossRef]

276. Lease, R. A., and M. Belfort. 2000. Riboregulation by DsrA RNA: transactions for global economy. Mol. Microbiol. 38:667–672.[PubMed] [CrossRef]

277. Lease, R. A., M. E. Cusick, and M. Belfort. 1998. Riboregulation in Escherichia coli: DsrA RNA acts by RNA:RNA interaction at multiple loci. Proc. Natl. Acad. Sci. USA 95:12456–12461.[PubMed] [CrossRef]

278. Lee, C. R., S. H. Cho, H. J. Kim, A. Peterkofsky, and Y.-J. Seok. 2010. Potassium mediates Escherichia coli enzyme IIA(Ntr)-dependent regulation of sigma factor selectivity. Mol. Microbiol. 78:1468–1483.[PubMed] [CrossRef]

279. Lee, H. Y., S. A. Cho, I. S. Lee, J. H. Park, S. H. Seok, M. W. Baek, D. J. Kim, S. H. Lee, S. J. Hur, S. H. Ban, Y. K. Lee, Y. K. han, Y. K. Cho, and J. H. Park. 2007. Evaluation of phoP and rpoS mutants of Salmonella enterica serovar Typhi as attenuated typhoid vaccine candidates: virulence and protective immuno responses in intranasally immunized mice. FEMS Immunol. Med. Microbiol. 51:310–318.[PubMed] [CrossRef]

280. Lee, I. S., J. Lin, H. K. Hall, B. Bearson, and J. W. Foster. 1995. The stationary-phase sigma factor σ^S (RpoS) is required for a sustained acid tolerance response in virulent Salmonella typhimurium. Mol. Microbiol. 17:155–167.[PubMed] [CrossRef]

281. Lee, J., A. Jayaraman, and T. K. Wood. 2007. Indole is an inter-species biofilm signal mediated by SdiA. BMC Microbiol. 7:42.[PubMed] [CrossRef]

282. Lee, J., R. Page, R. García-Contreras, J.-M. Palermino, X.-S. Zhang, O. Doshi, T. K. Wood, and W. Peti. 2007. Structure and function of the Escherichia coli protein YmgB: a protein critical for biofilm formation and acid resistance. J. Mol. Biol. 373:11–26.[PubMed] [CrossRef]

283. Lee, J. H., W. S. Yeo, and J. H. Roe. 2004. Induction of the sufA operon encoding Fe-S assembly proteins by superoxide generators and hydrogen peroxide: involvement of OxyR, IHF and an unidentified oxidant-responsive factor. Mol. Microbiol. 51:1745–1755.[PubMed] [CrossRef]

284. Lee, S. J., and J. D. Gralla. 2001. Sigma38 (rpoS) RNA polymerase promoter engagement via −10 region nucleotides. J. Biol. Chem. 276:30064–30071.[PubMed] [CrossRef]

285. Lee, S. J., and J. D. Gralla. 2004. Osmo-regulation of bacterial transcription via poised RNA polymerase. Mol. Cell 14:153–162.[PubMed] [CrossRef]

286. Leophart, A. B., D. K. Thompson, K. Huang, E. Alm, X. F. Wan, A. Arkin, S. D. Brown, L. Wu, T. Yan, X. S. Liu, G. S. Wickham, and J. Zhou. 2006. Transcriptome profiling of Shewanella oneidensis gene expression following exposure to acidic and alkaline pH. J. Bacteriol. 188:1633–1642.[PubMed] [CrossRef]

287. Lesniak, J., W. A. Barton, and D. B. Nikolov. 2003. Structural and functional features of the Escherichia coli hydroperoxide resistance protein OsmC. Protein Sci. 12:2838–2843.[PubMed] [CrossRef]

288. Li, H., A. Granat, V. Stewart, and J. R. Gillespie. 2008. RpoS, H-NS, and DsrA influence EHEC hemolysin operon (ehxCABD) transcription in Escherichia coli O157:H7 strain EDL933. FEMS Microbiol. Lett. 285:257–262.[PubMed] [CrossRef]

289. Lim, B., and C. A. Gross. 2011. Cellular response to heat shock and cold shock, p. 93–114. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses, 2nd ed. ASM Press, Washington, DC.

290. Lin, Y. H., C. Miyamoto, and E. A. Meighen. 2002. Cloning, sequencing, and functional studies of the rpoS gene from Vibrio harveyi. Biochem. Biophys. Res. Commun. 293:456–462.[PubMed] [CrossRef]

291. Liu, M., T. Durfee, J. E. Cabrera, K. Zhao, D. J. Jin, and F. R. Blattner. 2005. Global transcriptional programs reveal a carbon source foraging strategy by Escherichia coli. J. Biol. Chem. 280:15921–15927.[PubMed] [CrossRef]

292. Loewen, P. C., and B. L. Triggs. 1984. Genetic mapping of katF, a locus that with katE affects the synthesis of a second catalase species in Escherichia coli. J. Bacteriol. 160:668–675.[PubMed]

293. Loewen, P. C., and R. Hengge-Aronis. 1994. The role of the sigma factor σ^S (KatF) in bacterial global regulation. Annu. Rev. Microbiol. 48:53–80.[PubMed] [CrossRef]

294. Loewen, P. C. 1992. Regulation of bacterial catalase synthesis, p. 97–115. In J. Scandalios (ed.), Molecular Biology of Free Radical Scavenging Systems. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.

295. Lombardo, M. J., I. Aponyi, and S. M. Rosenberg. 2004. General stress response regulator RpoS in adaptive mutation and amplification in Escherichia coli. Genetics 166:669–680.[PubMed] [CrossRef]

296. Ma, L., J. Chen, R. Liu, X. H. Zhang, and Y. A. Jiang. 2009. Mutation or rpoS gene decreased resistance to environmental stresses, synthesis of extracellular products and virulence of Vibrio anguillarum. FEMS Microbiol. Ecol. 70:130–136.[PubMed] [CrossRef]

297. Ma, Z., H. Richard, D. L. Tucker, T. Conway, and J. W. Foster. 2002. Collaborative regulation of Escherichia coli glutamate-dependent acid resistance by two AraC-like regulators, GadX and GadW (YhiW). J. Bacteriol. 184:7001–7012.[PubMed] [CrossRef]

298. Ma, Z., N. Masuda, and J. W. Foster. 2004. Characterization of the EvgAS-YdeO-GadE branched regulatory circuit governing glutamate-dependent acid resistance in Escherichia coli. J. Bacteriol. 186:7378–7389.[PubMed] [CrossRef]

299. Ma, Z., S. Gong, H. Richard, D. L. Tucker, T. Conway, and J. W. Foster. 2003. GadE (YhiE) activates glutamate decarboxylase-dependent acid resistance in Escherichia coli K-12. Mol. Microbiol. 49:1309–1320.[PubMed] [CrossRef]

300. Maeda, H., N. Fujita, and A. Ishihama. 2000. Competition among seven Escherichia coli sigma subunits: relative binding affinities to the core RNA polymerase. Nucleic Acids Res. 28:3497–3503.[PubMed] [CrossRef]

301. Magnusson, L. U., A. Farewell, and T. Nyström. 2005. ppGpp: a global regulator in Escherichia coli. Trends Microbiol. 13:236–242.[PubMed] [CrossRef]

302. Majdalani, N., C. Cunning, D. Sledjeski, T. Elliott, and S. Gottesman. 1998. DsrA RNA regulates translation of RpoS message by an anti-antisense mechanisms, independent of its action as an antisilencer of transcription. Proc. Natl. Acad. Sci. USA 95:12462–12467.[PubMed] [CrossRef]

303. Majdalani, N., D. Hernandez, and S. Gottesman. 2002. Regulation and mode of action of the second small RNA activator of RpoS translation, RprA. Mol. Microbiol. 46:813–826.[PubMed] [CrossRef]

304. Majdalani, N., S. Chen, J. Murrow, K. St. John, and S. Gottesman. 2001. Regulation of RpoS by a novel small RNA: the characterization of RprA. Mol. Microbiol. 39:1382–1394.[PubMed] [CrossRef]

305. Malki, A., H. T. Le, S. Milles, R. Kern, T. Caldas, J. Abdallah, and G. Richarme. 2008. Solubilization of protein aggregates by the acid stress chaperones HdeA and HdeB. J. Biol. Chem. 283:13679–13687.[PubMed] [CrossRef]

306. Malpica, R., B. Franco, C. Rodriguez, O. Kwon, and D. Georgellis. 2004. Identification of quinone-sensitive redox switch in the ArcB sensor kinase. Proc. Natl. Acad. Sci. USA 101:13318–13323.[PubMed] [CrossRef]

307. Malpica, R., G. R. Sandoval, C. Rodriguez, B. Franco, and D. Georgellis. 2006. Signaling by the arc two-component system provides a link between the redox state of the quinone pool and gene expression. Antioxid. Redox Signal. 8:781–795.[PubMed] [CrossRef]

308. Mandel, M. J., and T. J. Silhavy. 2005. Starvation for different nutrients in Escherichia coli results in differential modulation of RpoS levels and stability. J. Bacteriol. 187:434–442.[PubMed] [CrossRef]

309. Mandin, P., and S. Gottesman. 2010. Integrating anaerobic/aerobic sensing and the general stress response through the ArcZ small RNA. EMBO J. 29:3094–3104.[PubMed] [CrossRef]

310. Mangan, M. W., S. Lucchini, V. Danino, T. O. Cróinín, J. C. Hinton, and C. J. Dorman. 2006. The integration host factor (IHF) integrates stationary-phase and virulence gene expression in Salmonella enterica serovar Typhimurium. Mol. Microbiol. 59:1831–1847.[PubMed] [CrossRef]

311. Martin, A., T. A. Baker, and R. T. Sauer. 2008. Protein unfolding by a AAA+ protease is dependent on ATP hydrolysis rates and substrate energy landscapes. Nat. Struct. Mol. Biol 15:139–145.[PubMed] [CrossRef]

312. Martinez, A., and R. Kolter. 1997. Protection of DNA during oxidative stress by the nonspecific DNA-binding protein Dps. J. Bacteriol. 179:5188–5194.[PubMed]

313. Martínez-García, E., A. Tormo, and J. M. Navarro-Llorens. 2001. Further studies on RpoS in enterobacteria: identification of rpoS in Enterobacter cloacae and Kluyvera cryocrescens. Arch. Microbiol. 175:395–404.[PubMed] [CrossRef]

314. Masuda, N., and G. M. Church. 2002. Escherichia coli gene expression responsive to levels of the response regulator EvgA. J. Bacteriol. 184:6225–6234.[PubMed] [CrossRef]

315. Masuda, N., and G. M. Church. 2003. Regulatory network of acid resistance genes in Escherichia coli. Mol. Microbiol. 48:699–712.[PubMed] [CrossRef]

316. Mates, A. K., A. K. Sayed, and J. W. Foster. 2007. Products of the Escherichia coli acid fitness island attenuate metabolite stress as extremely low pH and mediate a cell density-dependent acid resistance. J. Bacteriol. 189:2759–2768.[PubMed] [CrossRef]

317. McCann, M. P., C. D. Fraley, and A. Matin. 1993. The putative s factor KatF is regulated posttranscriptionally during carbon starvation. J. Bacteriol. 175:2143–2149.[PubMed]

318. McCann, M. P., J. P. Kidwell, and A. Matin. 1991. The putative sigma factor KatF has a central role in development of starvation-mediated general resistance in Escherichia coli. J. Bacteriol. 173:4188–4194.[PubMed]

319. McCleary, W. R., and J. B. Stock. 1994. Acetyl phosphate and the activation of two-component response regulators. J. Biol. Chem. 269:31567–31572.[PubMed]

320. McCullen, C. A., J. N. Benhammou, N. Majdalani, and S. Gottesman. 2010. Mechanism of positive regulation by DsrA and RprA small noncoding RNAs: pairing increases translation and protects rpoS mRNA from degradation. J. Bacteriol. 192:5559–5571.[PubMed] [CrossRef]

321. McMeechan, A., M. Roberts, T. A. Cogan, F. Jørgensen, A. Stevenson, C. Lewis, G. Rowley, and T. J. Humphrey. 2007. Role of the alternative sigma factors sigmaE and sigmaS in survival of Salmonella enterica serovar Typhimurium during starvation, refrigeration and osmotic shock. Microbiology 153:263–269.[PubMed] [CrossRef]

322. Merrikh, H., A. E. Ferrazzoli, A. Bougdour, A. Olivier-Mason, and S. T. Lovett. 2009. A DNA damage response in Escherichia coli involving the alternative sigma factor, RpoS. Proc. Natl. Acad. Sci. USA 106:611–616.[PubMed] [CrossRef]

323. Messner, K. R., and J. A. Imlay. 1999. The identification of primary sites of superoxide and hydrogen peroxide formation in the aerobic respiratory chain and sulfite reductase complex of Escherichia coli. J. Biol. Chem. 274:10119–10128.[PubMed] [CrossRef]

324. Metzner, M., J. Germer, and R. Hengge. 2004. Multiple stress signal integration in the regulation of the complex σ^S-dependent csiD-ygaF-gabDTP operon in Escherichia coli. Mol. Microbiol. 51:799–811.[PubMed] [CrossRef]

325. Mika, F., and R. Hengge. 2005. A two-component phosphotransfer network involving ArcB, ArcA and RssB coordinates synthesis and proteolysis of σ^S in E. coli. Genes Dev. 19:2770–2781.[PubMed] [CrossRef]

326. Mikulecky, P. J., M. K. Kaw, C. C. Brescia, J. C. Takach, D. D. Sledjeski, and A. L. Feig. 2004. Escherichia coli Hfq has distinct interaction surfaces for DsrA, rpoS and poly(A) RNAs. Nat. Struct. Mol. Biol 11:1206–1214.[PubMed] [CrossRef]

327. Milo, R., S. Shen-Orr, S. Itzkovitz, N. Kashtan, D. Chklovskii, and U. Alon. 2002. Network motifs: simple building blocks of complex networks. Science 298:824–827.[PubMed] [CrossRef]

328. Mitchell, J. E., T. Oshima, S. E. Piper, C. L. Webster, L. F. Westblade, G. Karimova, D. Ladant, A. Kolb, J. L. Hobman, S. J. Busby, and D. J. Lee. 2007. The Escherichia coli regulator of σ⁷⁰ protein, Rsd, can up-regulate some stress-dependent promoters by sequestering σ⁷⁰. J. Bacteriol. 189:3489–3495.[PubMed] [CrossRef]

329. Mitrophanov, A. Y., and E. A. Groisman. 2008. Signal integration in bacterial two-component regulatory systems. Genes Dev. 22:2601–2611.[PubMed] [CrossRef]

330. Mokkapati, S. K., F. de Henestrosa, and A. S. Bhagwat. 2001. Escherichia coli DNA glycosylase Mug: a growth-regulated enzyme required for mutation avoidance in stationary phase. Mol. Microbiol. 41:1101–1111.[PubMed] [CrossRef]

331. Monds, R. D., and G. A. O´Toole. 2009. The developmental model of microbial biofilms: ten years of a paradigm up for review. Trends Microbiol. 17:73–87.[PubMed] [CrossRef]

332. Monod, J. 1947. The phenomenon of enzymatic adaptation. Growth 11:223–289.

333. Monteil, V., A. Kolb, C. Mayer, S. Hoos, P. England, and F. Norel. 2010. Crl binds to domain 2 of σ^S and confers a competitive advantage on a natural rpoS mutant of Salmonella enterica serovar Typhi. J. Bacteriol. 192:6401–6410.[PubMed] [CrossRef]

334. Moreau, P. L. 2004. Diversion of the metabolic flux from pyruvate dehydrogenase to pyruvate oxidase decreases oxidative stress during glucose metabolism in nongrowing Escherichia coli cells incubated under aerobic, phosphate starvation conditions. J. Bacteriol. 186:7364–7368.[PubMed] [CrossRef]

335. Moreno, M., J. P. Audia, S. M. D. Bearson, C. Webb, and J. W. Foster. 2000. Regulation of sigma-S degradation in Salmonella enterica serovar typhimurium: in vivo interactions between sigma-S, the response regulator MviA (RssB) and ClpX. J. Mol. Microbiol. Biotechnol. 2:245–254.[PubMed]

336. Muffler, A., D. D. Traulsen, D. Fischer, R. Lange, and R. Hengge-Aronis. 1997. The RNA-binding protein HF-I plays a global regulatory role which is largely, but not exclusively, due to its role in expression of the σ^S subunit of RNA polymerase in Escherichia coli. J. Bacteriol. 179:297–300.[PubMed]

337. Muffler, A., D. D. Traulsen, R. Lange, and R. Hengge-Aronis. 1996. Posttranscriptional osmotic regulation of the σ^S subunit of RNA polymerase in Escherichia coli. J. Bacteriol. 178:1607–1613.[PubMed]

338. Muffler, A., D. Fischer, and R. Hengge-Aronis. 1996. The RNA-binding protein HF-I, known as a host factor for phage Qbeta RNA replication, is essential for the translational regulation of rpoS in Escherichia coli. Genes Dev. 10:1143–1151.[PubMed] [CrossRef]

339. Muffler, A., D. Fischer, S. Altuvia, G. Storz, and R. Hengge-Aronis. 1996. The response regulator RssB controls stability of the σ^S subunit of RNA polymerase in Escherichia coli. EMBO J. 15:1333–1339.[PubMed]

340. Muffler, A., M. Barth, C. Marschall, and R. Hengge-Aronis. 1997. Heat shock regulation of σ^S turnover: a role for DnaK and relationship between stress responses mediated by σ^S and σ³² in Escherichia coli. J. Bacteriol. 179:445–452.[PubMed]

341. Mukhopadhyay, S., J. P. Audia, R. N. Roy, and H. E. Schellhorn. 2000. Transcriptional induction of the conserved alternative sigma factor RpoS in Escherichia coli is dependent on BarA, a probable two-component regulator. Mol. Microbiol. 37:371–381.[PubMed] [CrossRef]

342. Mulvey, M. R., and P. C. Loewen. 1989. Nucleotide sequence of katF of Escherichia coli suggest KatF protein is a novel σ transcription factor. Nucleic Acids Res. 17:9979–9991.[PubMed] [CrossRef]

343. Mulvey, M. R., J. Switala, A. Borys, and P. C. Loewen. 1990. Regulation of transcription of katE and katF in Escherichia coli. J. Bacteriol. 172:6713–6720.[PubMed]

344. Munro, P. M., G. N. Flatau, R. L. Clement, and M. J. Gauthier. 1995. Influence of the RpoS (KatF) sigma factor on maintenance of viability and culturability of Escherichia coli and Salmonella typhimurium in seawater. Appl. Environ. Microbiol. 61:1853–1858.[PubMed]

345. Møller, T., T. Franch, P. Hojrup, D. R. Keene, H. P. Bachinger, R. G. Brennan, and P. Valentin-Hansen. 2002. Hfq, a bacterial Sm-like protein that mediates RNA-RNA interaction. Mol. Cell 9:23–30.[PubMed] [CrossRef]

346. Müller, J., M. C. Miller, A. T. Nielsen, G. K. Schoolnick, and A. M. Spormann. 2007. vpsA- and luxO-independent biofilms of Vibrio cholerae. FEMS Microbiol. Lett. 275:199–206.[PubMed] [CrossRef]

347. Nair, S., and S. E. Finkel. 2004. Dps protects cells against multiple stresses during stationary phase. J. Bacteriol. 186:4192–4198.[PubMed] [CrossRef]

348. Nguyen, L. H., and R. R. Burgess. 1997. Comparative analysis of the interactions of Escherichia coli σ^S and σ⁷⁰ RNA polymerase holoenzyme with the stationary phase-specific bolAp1 promoter. Biochemistry 36:1748–1754.[PubMed] [CrossRef]

349. Nguyen, L. H., D. B. Jensen, N. E. Thompson, D. R. Gentry, and R. R. Burgess. 1993. In vitro functional characterization of overproduced Escherichia coli katF/rpoS gene product. Biochemistry 32:11112–11117.[PubMed] [CrossRef]

350. Nickels, B. E., and A. Hochschild. 2004. Regulation of RNA polymerase through the secondary channel. Cell 118:281–284.[PubMed] [CrossRef]

351. Nickels, B. E., S. L. Dove, K. S. Murakami, S. A. Darst, and A. Hochschild. 2002. Protein-protein and protein-DNA interactions of σ⁷⁰ region 4 involved in transcription activation by lambda cI. J. Mol. Biol. 324:17–34.[PubMed] [CrossRef]

352. Nickerson, C. A., and R. Curtiss III. 1997. Role of sigma factor RpoS in initial stages of Salmonella typhimurium infection. Infect. Immun. 65:1814–1823.[PubMed]

353. Nielsen, A. T., N. A. Dolganov, G. Otto, M. C. Miller, C. Y. Wu, and G. K. Schoolnik. 2006. RpoS controls the Vibrio cholerae mucosal escape response. PLoS Pathog. 2:e109.[PubMed] [CrossRef]

354. Norel, F., V. Robbe-Saule, M. Y. Popoff, and C. Coynault. 1992. The putative sigma factor KatF (RpoS) is required for the transcription of the Salmonella typhimurium virulence gene spvB in Escherichia coli. FEMS Microbiol. Lett. 99:271–276. [CrossRef]

355. Notley, L., and T. Ferenci. 1996. Induction of RpoS-dependent functions in glucose-limited continuous culture: what level of nutrient limitation induces the stationary phase of Escherichia coli. J. Bacteriol. 178:1465–1468.[PubMed]

356. Notley-McRobb, L., T. King, and T. Ferenci. 2002. rpoS mutations and loss of general stress resistance in Escherichia coli populations as a consequence of conflict between competing stress responses. J. Bacteriol. 184:806–811.[PubMed] [CrossRef]

357. Nyström, T., C. Larsson, and L. Gustafsson. 1996. Bacterial defense against aging: role of the Escherichia coli ArcA regulator in gene expression, readjusted energy flux and survival during stasis. EMBO J. 15:3219–3228.[PubMed]

358. Nyström, T. 2004. Stationary-phase physiology. Annu. Rev. Microbiol. 58:161–181.[PubMed] [CrossRef]

359. Ohnishi, K., K. Kutsukake, H. Suzuki, and T. Iino. 1990. Gene fliA encodes an alternative sigma factor specific for flagellar operons in Salmonella typhimurium. Mol. Gen. Genet. 221:139–147.[PubMed] [CrossRef]

360. Ohnuma, M., N. Fujita, A. Ishihama, K. Tanaka, and H. Takahashi. 2000. A carboxy-terminal 16-amino-acid region of σ³⁸ of Escherichia coli is important for transcription under high-salt conditions and sigma activities in vivo. J. Bacteriol. 182:4628–4631.[PubMed] [CrossRef]

361. Ojangu, E.-L., A. Tover, R. Teras, and M. Kivisaar. 2000. Effects of combination of different −10 hexamers and downstream sequences on stationary phase-specific sigma factor σ^S-dependent transcription in Pseudomonas putida. J. Bacteriol. 182:6707–6713.[PubMed] [CrossRef]

362. Olsen, A. S., J. Møller-Jensen, R. G. Brennan, and P. Valentin-Hansen. 2010. C-terminally truncated derivatives of Escherichia coli Hfq are proficient in riboregulation. J. Mol. Biol. 404:173–182.[PubMed] [CrossRef]

363. Olsén, A., A. Jonsson, and S. Normark. 1989. Fibronectin binding mediated by a novel class of surface organelles on Escherichia coli. Nature 338:652–655.[PubMed] [CrossRef]

364. Olsén, A., M. J. Wick, M. Mörgelin, and L. Björck. 1998. Curli, fibrous surface proteins of Escherichia coli interact with major histocompatibility complex class I molecules. Infect. Immun. 66:944–949.[PubMed]

365. Olvera, L., A. Mendoza-Vargas, N. Flores, M. Olvera, J. C. Sigala, G. Gosset, E. Morett, and F. Bolívar. 2009. Transcription analysis of central metabolism genes in Escherichia coli. Possible roles of σ³⁸ in their expression, as a response to carbon limitation. PLoS One 4:e7466.[PubMed] [CrossRef]

366. Opdyke, J. A., J.-G. Kang, and G. Storz. 2004. GadY, a small-RNA regulator of acid response genes in Escherichia coli. J. Bacteriol. 186:6698–6705.[PubMed] [CrossRef]

367. Oppezzo, O. J., C. S. Costa, and R. A. Pizarro. 2011. Influence of rpoS mutations on the response of Salmonella enterica serovar Typhimurium to solar radiation. J. Photochem. Photobiol. B. 102:20–25.[PubMed] [CrossRef]

368. Otto, K., and T. J. Silhavy. 2002. Surface sensing and adhesion of Escherichia coli controlled by the Cpx-signaling pathway. Proc. Natl. Acad. Sci. USA 99:2287–2292.[PubMed] [CrossRef]

369. Park, K. H., M. Song, and H. E. Choy. 2007. Initial characterization of yliH in Salmonella typhimurium. J. Microbiol. 45:558–565.[PubMed]

370. Partridge, J. D., D. M. Bodenmiller, M. S. Humphrys, and S. Spiro. 2009. NsrR targets in the Escherichia coli genome: new insights into DNA sequence requirements for binding and a role for NsrR in the regulation of motility. Mol. Microbiol. 73:680–694.[PubMed] [CrossRef]

371. Patten, C. L., M. G. Kirchhhof, M. R. Schertzberg, R. A. Morton, and H. E. Schellhorn. 2004. Microarray analysis of RpoS-mediated gene expression in Escherichia coli K-12. Mol. Genet. Genomics 272:580–591.[PubMed] [CrossRef]

372. Persson, O., T. Nyström, and A. Farewell. 2010. UspB, a member of the sigma-S regulon, facilitates RuvC resolvase function. DNA Repair (Amst.) 9:1162–1169.[PubMed] [CrossRef]

373. Pesavento, C., and R. Hengge. 2009. Bacterial nucleotide-based second messengers. Curr. Opin. Microbiol. 12:170–176.[PubMed] [CrossRef]

374. Pesavento, C., G. Becker, N. Sommerfeldt, A. Possling, N. Tschowri, A. Mehlis, and R. Hengge. 2008. Inverse regulatory coordination of motility and curli-mediated adhesion in Escherichia coli. Genes Dev. 22:2434–2446.[PubMed] [CrossRef]

375. Peterson, C. N., M. J. Mandel, and T. J. Silhavy. 2005. Escherichia coli starvation diets: essential nutrients weigh in distinctly. J. Bacteriol. 187:7549–7553.[PubMed] [CrossRef]

376. Peterson, C. N., N. Ruiz, and T. J. Silhavy. 2004. RpoS proteolysis is regulated by a mechanism that does not require the SprE (RssB) response regulator phosphorylation site. J. Bacteriol. 186:7403–7410.[PubMed] [CrossRef]

377. Peterson, C. N., V. J. Carabetta, T. Chowdhury, and T. J. Silhavy. 2006. LrhA regulates rpoS translation in response to the Rcs phosphorelay system in Escherichia coli. J. Bacteriol. 188:3175–3181.[PubMed] [CrossRef]

378. Petrosino, J. F., R. S. Galhardo, L. D. Morales, and S. M. Rosenberg. 2009. Stress-induced beta-lactam antibiotic resistance mutation and sequences of stationary phase mutations in the Escherichia coli chromosome. J. Bacteriol. 191:5881–5889.[PubMed] [CrossRef]

379. Phadtare, S., and M. Inouye. 2001. Role CspC and CspE in regulation of expression of RpoS and UspA, the stress response proteins in Escherichia coli. J. Bacteriol. 183:1205–1214.[PubMed] [CrossRef]

380. Piper, S. E., J. E. Mitchell, D. J. Lee, and S. J. Busby. 2009. A global view of Escherichia coli Rsd protein and its interaction. Mol. Biosyst. 5:1934–1947. [CrossRef]

381. Ponder, R. G., N. C. Fonville, and S. M. Rosenberg. 2005. A switch from high-fidelity to error-prone DNA double-strand break repair underlies stress-induced mutation. Mol. Cell 19:791–804.[PubMed] [CrossRef]

382. Potrykus, K., and M. Cashel. 2008. (p)ppGpp: still magical? Annu. Rev. Microbiol. 62:35–51.[PubMed] [CrossRef]

383. Pratt, L. A., and T. J. Silhavy. 1996. The response regulator, SprE, controls the stability of RpoS. Proc. Natl. Acad. Sci. USA 93:2488–2492.[PubMed] [CrossRef]

384. Pratt, L. A., and T. J. Silhavy. 1998. Crl stimulates RpoS activity during stationary phase. Mol. Microbiol. 29:1225–1236.[PubMed] [CrossRef]

385. Price, C. W. 2011. General stress response in Bacillus subtilis and related Gram-positive bacteria, p. 301–318. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses, 2nd ed. ASM Press, Washington, DC.

386. Price, S. B., C.-M. Cheng, C. W. Kaspar, J. C. Wright, F. J. DeGraves, T. A. Penfound, M.-P. Castanié-Cornet, and J. W. Foster. 2000. Role of rpoS in acid resistance and fecal shedding of Escherichia coli O157:H7. Appl. Environ. Microbiol. 66:632–637.[PubMed] [CrossRef]

387. Prigent-Combaret, C., E. Brombacher, O. Vidal, A. Ambert, P. Lejeune, P. Landini, and C. Dorel. 2001. Complex regulatory network controls initial adhesion and biofilm formation in Escherichia coli via regulation of the csgD gene. J. Bacteriol. 2001:7213–7223. [CrossRef]

388. Prigent-Combaret, C., G. Prensier, T. T. Le Thi, Q. Vidal, P. Lejeune, and C. Dorel. 2000. Developmental pathway for biofilm formation in curli-producing Escherichia coli strains: role of flagella, curli and colanic acid. Environ. Microbiol. 2:450–464.[PubMed] [CrossRef]

389. Prouty, A. M., and J. S. Gunn. 2003. Comparative analysis of Salmonella enterica serovar Typhimurium biofilm formation on gallstones and on glass. Infect. Immun. 71:7154–7158.[PubMed] [CrossRef]

390. Pruteanu, M., and R. Hengge-Aronis. 2002. The cellular level of the recognition factor RssB is rate-limiting for σ^S proteolysis: Implications for RssB regulation and signal transduction in σ^S turnover in Escherichia coli. Mol. Microbiol. 45:1701–1714.[PubMed] [CrossRef]

391. Quail, M. A., and J. Guest. 1995. Purification, characterization and mode of action of PdhR, the transcriptional repressor of the pdhR-aceEF-lpd operon of Escherichia coli. Mol. Microbiol. 15:519–529.[PubMed] [CrossRef]

392. Raffaelle, M., E. I. Kanin, J. Vogt, R. R. Burgess, and A. Z. Ansari. 2005. Holoenzyme switching and stochastic release of sigma factor from RNA polymerase in vivo. Mol. Cell 20:357–366.[PubMed] [CrossRef]

393. Rajkowitsch, L., and R. Schroeder. 2007. Dissecting RNA chaperone activity. RNA 13:2053–2060.[PubMed] [CrossRef]

394. Ramos-González, M. I., and S. Molin. 1998. Cloning, sequencing, and phenotypic characterization of the rpoS gene from Pseudomonas putida KT2440. J. Bacteriol. 180:3421–3431.[PubMed]

395. Ranquet, C., and S. Gottesman. 2007. Translational regulation of the Escherichia coli stress factor RpoS: a role for SsrA and Lon. J. Bacteriol. 189:4872–4879.[PubMed] [CrossRef]

396. Repoila, F., N. Majdalani, and S. Gottesman. 2003. Small non-coding RNAs, co-ordinators of adaptation processes in Escherichia coli: the RpoS paradigm. Mol. Microbiol. 48:855–861.[PubMed] [CrossRef]

397. Resch, A., B. Vecerek, K. Palavra, and U. Bläsi. 2010. Requirement of the CsdA DEAD-box helicase for low temperature riboregulation of rpoS mRNA. RNA Biol. 7:96–102. [CrossRef]

398. Richard, H., and J. W. Foster. 2007. Sodium regulates Escherichia coli acid resistance, and influences GadX- and GadW-dependent activation of gadE. Microbiology 153:3154–3161.[PubMed] [CrossRef]

399. Ring, B. Z., W. S. Yarnell, and J. W. Roberts. 1996. Function of E. coli RNA polymerase sigma factor σ⁷⁰ in promoter-proximal pausing. Cell 86:485–493.[PubMed] [CrossRef]

400. Robbe-Saule, V., F. Schaeffer, L. Kowarz, and F. Norel. 1997. Relationships between H-NS, σ^S, SpvR and growth phase in the control of spvR, the regulatory gene of the Salmonella plasmid virulence operon. Mol. Gen. Genet. 256:333–347.[PubMed] [CrossRef]

401. Robbe-Saule, V., G. Algorta, I. Rouilhac, and F. Norel. 2003. Characterization of the RpoS status of clinical isolates of Salmonella enterica. Appl. Environ. Microbiol. 69:4352–4358.[PubMed] [CrossRef]

402. Robbe-Saule, V., M. D. Lopes, A. Kolb, and F. Norel. 2007. Physiological effects of Crl in Salmonella are modulated by σ^S level and promoter specificity. J. Bacteriol. 189:2976–2987.[PubMed] [CrossRef]

403. Robbe-Saule, V., V. Jaumouille, M. C. Prevost, S. Guadagnini, C. Talhouarne, H. Mathout, A. Kolb, and F. Norel. 2006. Crl activates transcription initiation of RpoS-regulated genes involved in the multicellular behavior of Salmonella enterica serovar typhimurium. J. Bacteriol. 188:3983–3994.[PubMed] [CrossRef]

404. Rockabrand, D., K. Livers, T. Austin, R. Kaiser, D. Jensen, R. Burgess, and P. Blum. 1998. Roles of DnaK and RpoS in starvation-induced thermotolerance of Escherichia coli. J. Bacteriol. 180:846–854.[PubMed]

405. Rockabrand, D., T. Arthur, G. Korinek, K. Livers, and P. Blum. 1995. An essential role for the Escherichia coli DnaK protein in starvation-induced thermotolerance, H₂O₂ resistance, and reductive division. J. Bacteriol. 177:3695–3703.[PubMed]

406. Rosenthal, A. Z., M. Hu, and J. D. Gralla. 2006. Osmolyte-induced transcription: −35 region elements and recognition by σ³⁸. Mol. Microbiol. 59:1052–1061.[PubMed] [CrossRef]

407. Rosenthal, A. Z., Y. Kim, and J. D. Gralla. 2008. Poising of Escherichia coli RNA polymerase and its release from the sigma38 C-terminal tail for osmY transcription. J. Mol. Biol. 376:938–949.[PubMed] [CrossRef]

408. Rosenthal, A. Z., Y. Kim, and J. D. Gralla. 2008. Regulation of transcription by acetate in Escherichia coli: in vivo and in vitro comparisons. Mol. Microbiol. 68.[PubMed] [CrossRef]

409. Ross, W., K. K. Gosink, J. Salomon, K. Igarashi, C. Zou, A. Ishihama, K. Severinov, and R. L. Gourse. 1993. A third recognition element in bacterial promoters—DNA binding by the alpha-subunit of RNA polymerase. Science 262:1407–1413.[PubMed] [CrossRef]

410. Rozen, D. E., N. Philippe, A. de Visser, R. E. Lenski, and D. Schneider. 2009. Death and cannibalism in a seasonal environment facilitate bacterial coexistence. Ecol. Lett. 12:34–44.[PubMed] [CrossRef]

411. Ruiz, N., and T. J. Silhavy. 2005. Sensing external stress: watchdogs of the Escherichia coli envelope. Curr. Opin. Microbiol. 8:122–126.[PubMed] [CrossRef]

412. Ruiz, N., C. N. Peterson, and T. J. Silhavy. 2001. RpoS-dependent transcriptional control of sprE: regulatory feedback loop. J. Bacteriol. 183:5974–5981.[PubMed] [CrossRef]

413. Rychlik, I., and P. A. Barrow. 2005. Salmonella stress management and its relevance to behaviour during intestinal colonisation and infection. FEMS Microbiol. Rev. 29:1021–1040.[PubMed] [CrossRef]

414. Rychlik, I., G. Martin, U. Methner, M. Lovell, L. Cardova, A. Sebkova, M. Sevcik, J. Damborsky, and P. A. Barrow. 2002. Identification of Salmonella enterica serovar Typhimurium genes associated with growth suppression in stationary-phase nutrient broth cultures and in the chicken intestine. Arch. Microbiol. 178:411–420.[PubMed] [CrossRef]

415. Römling, U., and D. Amikam. 2006. Cyclic di-GMP as a second messenger. Curr. Opin. Microbiol. 9:218–228.[PubMed] [CrossRef]

416. Römling, U., M. Gomelsky, and M. Y. Galperin. 2005. C-di-GMP: the dawning of a novel bacterial signalling system. Mol. Microbiol. 57:629–639.[PubMed] [CrossRef]

417. Römling, U., M. Rohde, A. Olsén, S. Normark, and J. Reinköster. 2000. AgfD, the checkpoint of multicellular and aggregative behaviour in Salmonella typhimurium regulates at least two independent pathways. Mol. Microbiol. 36:10–23.[PubMed] [CrossRef]

418. Römling, U., W. Bokranz, W. Rabsch, X. Zogaj, M. Nimtz, and H. Tschäpe. 2003. Occurrence and regulation of the multicellular morphotype in Salmonella serovars important in human disease. Int. J. Med. Microbiol. 293:273–285.[PubMed] [CrossRef]

419. Römling, U., W. D. Sierralta, K. Eriksson, and S. Normark. 1998. Multicellular and aggregative behaviour of Salmonella typhimurium strains is controlled by mutations in the agfD promoter. Mol. Microbiol. 28:249–264.[PubMed] [CrossRef]

420. Römling, U., Z. Bian, M. Hammar, W. D. Sierralta, and S. Normark. 1998. Curli fibers are highly conserved between Salmonella typhimurium and Escherichia coli with respect to operon structure and regulation. J. Bacteriol. 180:722–731.[PubMed]

421. Römling, U. 2005. Characterization of the rdar morphotype, a multicellular behaviour in Enterobacteriaceae. Cell. Mol. Life Sci. 62:1234–1246.[PubMed] [CrossRef]

422. Sak, B. D., A. Eisenstark, and D. Touati. 1989. Exonuclease III and the catalase hydroperoxidase II in Escherichia coli are both regulated by the katF product. Proc. Natl. Acad. Sci. USA 86:3271–3275.[PubMed] [CrossRef]

423. Santander, J., K. L. Roland, and R. Curtiss III. 2008. Regulation of Vi capsular polysaccharide synthesis in Salmonella enterica serotype Typhi. J. Infect. Dev. Ctries. 2:412–420.[PubMed]

424. Santos, J. M., M. Lobo, A. P. Matos, M. A. De Pedro, and C. M. Arraiano. 2002. The gene bolA regulates dacA (PBP5), dacC (PBP6) and ampC (AmpC), promoting normal morphology in Escherichia coli. Mol. Microbiol. 45:1729–1740.[PubMed] [CrossRef]

425. Santos, J. M., P. Freire, F. Mika, R. Hengge, and C. M. Arraiano. 2006. Bacterial polyadenylation links transcription with mRNA degradation via σ^S proteolysis. Mol. Microbiol. 60:177–188.[PubMed] [CrossRef]

426. Santos, J. M., P. Freire, M. Vicente, and C. M. Arraiano. 1999. The stationary-phase morphogene bolA from Escherichia coli is induced by stress during early stages of growth. Mol. Microbiol. 32:789–798.[PubMed] [CrossRef]

427. Sauter, C., J. Basquin, and D. Suck. 2003. Sm-like protein in eubacteria: the crystal structure of the Hfq protein from Escherichia coli. Nucleic Acids Res. 31:4091–4098.[PubMed] [CrossRef]

428. Schellhorn, H. E. 1995. Regulation of hydroperoxidase (catalase) expression in Escherichia coli. FEMS Microbiol. Lett. 131:113–119.[PubMed] [CrossRef]

429. Schembri, M. A., K. Kjaergaard, and P. Klemm. 2003. Global gene expression in Escherichia coli biofilms. Mol. Microbiol. 48:253–267.[PubMed] [CrossRef]

430. Schlensog, V., S. Lutz, and A. Böck. 1994. Purification and DNA-binding properties of FHLA, the transcriptional activator of the formate hydrogenlyase system from Escherichia coli. J. Biol. Chem. 269:19590–19596.[PubMed]

431. Schumacher, M. A., R. F. Pearson, T. Møller, P. Valentin-Hansen, and R. G. Brennan. 2002. Structures of the pleiotropic translational regulator Hfq and an Hfq-RNA complex: a bacterial Sm-like protein. EMBO J. 21:3546–3556.[PubMed] [CrossRef]

432. Schuster, M., A. C. Hawkins, C. S. Harwood, and E. P. Greenberg. 2004. The Pseudomonas aeruginosa RpoS regulon and its relationship to quorum sensing. Mol. Microbiol. 51:973–985.[PubMed] [CrossRef]

433. Schuster, M., and E. P. Greenberg. 2007. Early activation of quorum sensing in Pseudomonas aeruginosa reveals the architecture of a complex regulon. BMC Genomics 8:287.[PubMed] [CrossRef]

434. Schweder, T., K.-H. Lee, O. Lomovskaya, and A. Matin. 1996. Regulation of Escherichia coli starvation sigma factor (σ³⁸) by ClpXP protease. J. Bacteriol. 178:470–476.[PubMed]

435. Sevcik, M., A. Sebková, J. Volf, and I. Rychlík. 2001. Transcription of arcA and rpoS during growth of Salmonella typhimurium under aerobic and microaerobic conditions. Microbiology 147:701–708.[PubMed]

436. Sezonov, G., D. Joseleau-Petit, and R. D’Ari. 2007. Escherichia coli physiology in Luria-Bertani broth. J. Bacteriol. 189:8746–8749.[PubMed] [CrossRef]

437. Shen-Orr, S. S., R. Milo, S. Mangan, and U. Alon. 2002. Network motifs in the transcriptional regulation network of Escherichia coli. Nat. Genet. 31:64–68.[PubMed] [CrossRef]

438. Shin, M., M. Song, J. H. Rhee, Y. Hong, Y. J. Kim, Y.-J. Seok, K. S. Ha, S. H. Jung, and H. E. Choy. 2005. DNA looping-mediated repression by histone-like protein H-NS: specific requirement of E σ⁷⁰ as a cofactor for looping. Genes Dev. 19:2388–2398.[PubMed] [CrossRef]

439. Shin, S., M.-P. Castanie-Cornet, J. W. Foster, J. A. Crawford, C. Brinkley, and J. B. Kaper. 2001. An activator glutamate decarboxylate genes regulates the expression of enteropathogenic Escherichia coli virulence genes through control of the plasmid-encoded regulator, Per. Mol. Microbiol. 41:1133–1150.[PubMed] [CrossRef]

440. Simm, R., A. Lusch, A. Kader, M. Andersson, and U. Römling. 2007. Role of EAL-containing proteins in multicellular behavior of Salmonella enterica serovar typhimurium. J. Bacteriol. 189:3613–3623.[PubMed] [CrossRef]

441. Simm, R., M. Morr, A. Kader, M. Nimtz, and U. Römling. 2004. GGDEF and EAL domains inversely regulate cyclic di-GMP levels and transition from sessility to motility. Mol. Microbiol. 53:1123–1134.[PubMed] [CrossRef]

442. Sledjeski, D., and S. Gottesman. 1995. A small RNA acts as an antisilencer of the H-NS-silenced rcsA gene of Escherichia coli. Proc. Natl. Acad. Sci. USA 92:2003–2007.[PubMed] [CrossRef]

443. Sledjeski, D. D., A. Gupta, and S. Gottesman. 1996. The small RNA, DsrA, is essential for the low temperature expression of RpoS during exponential growth in E. coli. EMBO J. 15:3993–4000.[PubMed]

444. Sledjeski, D. D., C. Whitman, and A. Zhang. 2001. Hfq is necessary for regulation by the untranslated RNA DsrA. J. Bacteriol. 183:1997–2005.[PubMed] [CrossRef]

445. Small, P., D. Blankenhorn, D. Welty, E. Zinser, and J. L. Slonczewski. 1994. Acid and base resistance in Escherichia coli and Shigella flexneri: role of rpoS and growth pH. J. Bacteriol. 176:1729–1737.[PubMed]

446. Solis, R., I. Bertani, G. Degrassi, G. Devescovi, and V. Venturi. 2006. Involvement of quorum sensing and RpoS in rice seedling blight caused by Burkholderia plantarii. FEMS Microbiol. Lett. 259:106–112.[PubMed] [CrossRef]

447. Sommerfeldt, N., A. Possling, G. Becker, C. Pesavento, N. Tschowri, and R. Hengge. 2009. Gene expression patterns and differential input into curli fimbriae regulation of all GGDEF/EAL domain proteins in Escherichia coli. Microbiology 155:1318–1331.[PubMed] [CrossRef]

448. Soper, T., P. Mandin, N. Majdalani, S. Gottesman, and S. A. Woodson. 2010. Positive regulation by small RNAs and the role of Hfq. Proc. Natl. Acad. Sci. USA 107:9602–9607.[PubMed] [CrossRef]

449. Sorvik, K. A., and P. L. Foster. 2011. The SMC-like protein complex SbcCD enhances DNA polymerase IV-dependent spontaneous mutation in Escherichia coli. J. Bacteriol. 193:660–669.[PubMed] [CrossRef]

450. Soutourina, O., A. Kolb, E. Krin, C. Laurent-Winter, S. Rimsky, A. Danchin, and P. Bertin. 1999. Multiple control of flagellum biosynthesis in Escherichia coli: role of H-NS protein and the cyclic AMP-catabolite activator protein complex in transcription of the flhDC master operon. J. Bacteriol. 181:7500–7508.[PubMed]

451. Spector, M. P., F. Garcia del Portillo, S. M. D. Bearson, A. Mahmud, M. Magut, B. B. Finlay, G. Dougan, J. W. Foster, and M. J. Pallen. 1999. The rpoS-dependent starvation-stress response locus stiA encodes a nitrate reductase (NarZYWV) required for carbon-starvation-inducible thermotolerance and acid tolerance in Salmonella typhimurium. Microbiology 145:3035–3045.[PubMed]

452. Spector, M. P. 1998. The starvation-stress response (SSR) of Salmonella. Adv. Microb. Physiol. 40:233–279.[PubMed] [CrossRef]

453. Spira, B., X. Hu, and T. Ferenci. 2008. Strain variation in ppGpp concentration and RpoS levels in laboratory strains of Escherichia coli. Microbiology 154:2887–2895.[PubMed] [CrossRef]

454. Stephani, K., D. Weichart, and R. Hengge. 2003. Dynamic control of Dps protein levels by ClpXP and ClpAP proteases in Escherichia coli. Mol. Microbiol. 49:1605–1614.[PubMed] [CrossRef]

455. Stockwell, V. O., and J. E. Loper. 2005. The sigma factor RpoS is required for stress tolerance and environmental fitness of Pseudomonas fluorescens Pf-5. Microbiology 151:3001–3009.[PubMed] [CrossRef]

456. Stockwell, V. O., K. Hockett, and J. E. Loper. 2009. Role of RpoS in stress tolerance and environmental fitness of the phyllosphere bacterium Pseudomonas fluorescens strain 122. Phytopathology 99:689–695.[PubMed] [CrossRef]

457. Stokes, N. R., H. D. Murray, C. Subramaniam, R. L. Gourse, P. Louis, W. Bartlett, S. Miller, and I. R. Booth. 2003. A role for mechanosensitive channels in survival of stationary phase: regulation of channel expression by RpoS. Proc. Natl. Acad. Sci. USA 100:15959–15964.[PubMed] [CrossRef]

458. Stoodley, P., K. Sauer, D. G. Davies, and J. W. Costerton. 2002. Biofilms as complex differentiated communities. Annu. Rev. Microbiol. 56:187–209.[PubMed] [CrossRef]

459. Storvik, K. A., and P. L. Foster. 2010. RpoS, the stress response sigma factor, plays a dual role in the regulation of Escherichia coli ´s error-prone DNA polymerase IV. J. Bacteriol. 192:3639–3644.[PubMed] [CrossRef]

460. Storz, G., and R. Hengge (ed.). 2011. Bacterial Stress Responses, 2nd ed. ASM Press, Washington, DC.

461. Storz, G., and S. Spiro. 2011. Sensing and responding to reactive oxygen and nitrogen species, p. 157–174. In G. Storz and R. Hengge (ed.), Bacterial Stress Responses, 2nd ed. ASM Press, Washington , DC.

462. Storz, G., J. A. Opdyke, and A. Zhang. 2004. Controlling mRNA stability and translation with small, non-coding RNAs. Curr. Opin. Microbiol. 7:140–144.[PubMed] [CrossRef]

463. Storz, G., S. Altuvia, and K. M. Wassarman. 2005. An abundance of RNA regulators. Annu. Rev. Biochem. 74:199–217.[PubMed] [CrossRef]

464. Strohmeier-Gort, A., D. M. Ferber, and J. A. Imlay. 1999. The regulation and role of the periplasmic copper, zinc superoxide dismutase of Escherichia coli. Mol. Microbiol. 32:179–191.[PubMed] [CrossRef]

465. Strøm, A. R., and I. Kaasen. 1993. Trehalose metabolism in Escherichia coli: stress protection and stress regulation of gene expression. Mol. Microbiol. 8:205–210.[PubMed] [CrossRef]

466. Stüdemann, A., M. Noirclerc-Savoye, E. Klauck, G. Becker, D. Schneider, and R. Hengge. 2003. Sequential recognition of two distinct sites in σ^S by the proteolytic targeting factor RssB and ClpX. EMBO J. 22:4111–4120.[PubMed] [CrossRef]

467. Sutton, A., R. Buencamino, and A. Eisenstark. 2000. rpoS mutants in archival cultures of Salmonella enterica serovar Typhimurium. J. Bacteriol. 182:4375–4379.[PubMed] [CrossRef]

468. Takayanagi, Y., K. Tanaka, and H. Takahashi. 1994. Structure of the 5 ′ upstream region and the regulation of the rpoS gene of Escherichia coli. Mol. Gen. Genet. 243:525–531.[PubMed] [CrossRef]

469. Tanaka, K., S. Kusano, N. Fujita, A. Ishihama, and H. Takahashi. 1995. Promoter determinants for Escherichia coli RNA polymerase holoenzyme containing σ³⁸ (the rpoS gene product). Nucleic Acids Res. 23:827–834.[PubMed] [CrossRef]

470. Tanaka, K., Y. Takayanagi, N. Fujita, A. Ishihama, and H. Takahashi. 1993. Heterogeneity of the principal sigma factor in Escherichia coli: the rpoS gene product, σ³⁸, is a second principal sigma factor of RNA polymerase in stationary phase Escherichia coli. Proc. Natl. Acad. Sci. USA 90:3511–3515.[PubMed] [CrossRef]

471. Tani, T. H., A. Khodursky, R. M. Blumenthal, P. O. Brown, and R. G. Matthews. 2002. Adaptation to famine: a family of stationary-phase genes revealed by microarray analysis. Proc. Natl. Acad. Sci. USA 99:13471–13476.[PubMed] [CrossRef]

472. Teich, A., S. Meyer, H. Y. Lin, L. Andersson, S. O. Enfors, and P. Neubauer. 1999. Growth rate related concentration changes of the starvation response regulators σ^S and ppGpp in glucose-limited fed-batch and continuous cultures of Escherichia coli. Biotechnol. Progr. 15:123–129.[PubMed] [CrossRef]

473. Tenaillon, O., E. Denamur, and I. Matic. 2004. Evolutionary significance of stress-induced mutagenesis in bacteria. Trends Microbiol. 12:264–270.[PubMed] [CrossRef]

474. Tiaden, A., T. Spirig, S. S. Weber, H. Brüggemann, R. Bosshard, C. Buchrieser, and H. Hilbi. 2007. The Legionella pneumophila response regulator LqsR promotes host cell interactions as an element of the virulence regulatory network controlled by Rpos and LetA. Cell. Microbiol. 9:2903–2930.[PubMed] [CrossRef]

475. Tomoyasu, T., A. Takaya, E. Isogai, and T. Yamamoto. 2003. Turnover of FlhD and FlhC, master regulator proteins for Salmonella flagellum biogenesis, by the ATP-dependent ClpXP protease. Mol. Microbiol. 48:443–452.[PubMed] [CrossRef]

476. Tomoyasu, T., A. Takaya, Y. Handa, K. Karata, and T. Yamamoto. 2005. ClpXP controls the expression of LEE genes in enterohaemorrhagic Escherichia coli. FEMS Microbiol. Lett. 253:59–66.[PubMed] [CrossRef]

477. Tomoyasu, T., T. Ohkishi, Y. Ukyo, A. Tokumitsu, A. Takaya, M. Suzuki, K. Sekiya, H. Matsui, K. Kutsukake, and T. Yamamoto. 2002. The ClpXP ATP-dependent protease regulates flagellum synthesis in Salmonella enterica serovar typhimurium. J. Bacteriol. 184:645–653.[PubMed] [CrossRef]

478. Touati, E., E. Dassa, and P. L. Boquet. 1986. Pleiotropic mutations in appR reduce pH 2.5 acid phosphatase expression and restore succinate utilization in CRP-deficient strains of Escherichia coli. Mol. Gen. Genet. 202:257–264.[PubMed] [CrossRef]

479. Tramonti, A., M. De Canio, and D. De Biase. 2008. GadX/GadW-dependent regulation of the Escherichia coli acid fitness island: transcriptional control at the gadY-gadW divergent promoters and identification of four novel 42 bp GadX/GadW-specific binding sites. Mol. Microbiol. 70:965–982.[PubMed]

480. Tramonti, A., M. De Canio, I. Delany, V. Scarlato, and D. De Biase. 2006. Mechanisms of transcription activation exerted by GadX and GadW at the gadA and gadBC gene promoters of the glutamate-based acid resistance system in Escherichia coli. J. Bacteriol. 188:8118–8127.[PubMed] [CrossRef]

481. Tramonti, A., P. Visca, M. De Canio, M. Falconi, and D. De Biase. 2002. Functional characterization and regulation of gadX, a gene encoding an AraC/XylS-like transcriptional activator of the Escherichia coli glutamic acid decarboxylase system. J. Bacteriol. 184:2603–2613.[PubMed] [CrossRef]

482. Traxler, M. F., D. E. Chang, and T. Conway. 2006. Guanosine 3′,5′-bispyrophosphate coordinates global gene expression during glucose-lactose diauxie in Escherichia coli. Proc. Natl. Acad. Sci. USA 103:2374–2379.[PubMed] [CrossRef]

483. Traxler, M. F., V. M. Zacharia, S. Marquardt, S. M. Summers, H. T. Nguyen, S. E. Stark, and T. Conway. 2011. Discretely calibrated regulatory loops controlled by ppGpp partition gene induction across the "feast to famine" gradient in Escherichia coli. Mol. Microbiol. 79:830–845.[PubMed] [CrossRef]

484. Tschowri, N., S. Busse, and R. Hengge. 2009. The BLUF-EAL protein YcgF acts as a direct anti-repressor in a blue light response of E. coli. Genes Dev. 23:522–534.[PubMed] [CrossRef]

485. Tsui, H.-C. T., H.-C. L. Leung, and M. E. Winkler. 1994. Characterization of broadly pleiotropic phenotypes caused by an hfq insertion mutation in Escherichia coli K-12. Mol. Microbiol. 13:35–49.[PubMed] [CrossRef]

486. Tsui, H. C. T., G. Feng, and M. E. Winkler. 1997. Negative regulation of mutS and mutH repair gene expression by the Hfq and RpoS global regulators of Escherichia coli K-12. J. Bacteriol. 179:7476–7487.[PubMed]

487. Tu, X., T. Latifi, A. Bougdour, S. Gottesman, and E. A. Groisman. 2006. The PhoP/PhoQ two-component system stabilizes the alternative sigma factor RpoS in Salmonella enterica. Proc. Natl. Acad. Sci. USA 103:13503–13508.[PubMed] [CrossRef]

488. Tucker, D. L., N. Tucker, Z. Ma, J. W. Foster, R. L. Miranda, P. S. Cohen, and T. Conway. 2003. Genes of the GadX-GadW regulon in Escherichia coli. J. Bacteriol. 185:3190–3201.[PubMed] [CrossRef]

489. Tuveson, R. W., and R. B. Jonas. 1979. Genetic control of near-UV (300–400 nm) sensitivity independent of the recA gene in strains of Escherichia coli K-12. Photochem. Photobiol. 30:667–676.[PubMed] [CrossRef]

490. Typas, A., and R. Hengge. 2005. Differential ability of σ^S and σ⁷⁰ of Escherichia coli to utilize promoters containing half or full UP-element sites. Mol. Microbiol. 55:250–260.[PubMed] [CrossRef]

491. Typas, A., and R. Hengge. 2006. Role of the spacer between the -35 and -10 region in σ^S promoter selectivity in Escherichia coli. Mol. Microbiol. 59:1037–1051.[PubMed] [CrossRef]

492. Typas, A., C. Barembruch, and R. Hengge. 2007. Stationary phase reorganisation of the E.coli transcription machinery by Crl protein, a fine-tuner of σ^S activity and levels. EMBO J. 26:1569–1578.[PubMed] [CrossRef]

493. Typas, A., G. Becker, and R. Hengge. 2007. The molecular basis of selective promoter activation by the σ^S subunit of RNA polymerase. Mol. Microbiol. 63:1296–1306.[PubMed] [CrossRef]

494. Typas, A., S. Stella, R. C. Johnson, and R. Hengge. 2007. The -35 sequence location and the Fis-sigma factor interface determine σ^S selectivity of the proP (p2) promoter in Escherichia coli. Mol. Microbiol. 63:780–796.[PubMed]

495. Ueguchi, C., N. Misonou, and T. Mizuno. 2001. Negative control of rpoS expression by phosphoenolpyruvate: carbohydrate phosphotransferase system in Escherichia coli. J. Bacteriol. 183:520–527.[PubMed] [CrossRef]

496. Uehara, T., K. R. Parzych, T. Dinh, and T. G. Bernhardt. 2010. Daughter cell separation is controlled by cytokinetic ring-activated cell wall hydrolysis. EMBO J. 29:1412–1422.[PubMed] [CrossRef]

497. Updegrove, T., N. Wilf, X. Sun, and R. M. Wartell. 2008. Effect of Hfq on RprA-rpoS mRNA pairing: Hfq-RNA binding and the influence of the 5´rpoS mRNA leader region. Biochemistry 47:11184–11195.[PubMed] [CrossRef]

498. Utaisincharoen, P., S. Aricharoen, K. Limposuwan, S. Tungpradabkul, and S. Sirisinha. 2006. Burkholderia pseudomallei RpoS regulates multinucleated giant cell formation and inducible nitric oxide synthase expression in mouse macrophage cell line (RWA 264.7). Microb. Pathog. 40:184–189.[PubMed] [CrossRef]

499. Valentin-Hansen, P., M. Eriksen, and C. Udesen. 2004. The bacterial Sm-like protein Hfq: a key player in RNA transactions. Mol. Microbiol. 51:1525–1533.[PubMed] [CrossRef]

500. Vanaja, S. K., T. M. Bergholz, and T. S. Whittam. 2009. Characterization of the Escherichia coli O157:H7 Sakai GadE regulon. J. Bacteriol. 191:1868–1877.[PubMed] [CrossRef]

501. Vassylyev, D. G., S.-I. Sekine, O. Laptenko, L. J., M. N. Vassylyeva, S. Borukhov, and S. Yokoyama. 2002. Crystal structure of a bacterial RNA polymerase holoenzyme at 2.6 Å resolution. Nature 417:712–719.[PubMed] [CrossRef]

502. Vasudevan, P., and K. Venkitanarayanan. 2006. Role of the rpoS gene in the survival of Vibrio parahaemolyticus in artificial seawater and fish homogenate. J. Food Prot. 69:1438–1442.[PubMed]

503. Vecerek, B., M. Beich-Frandsen, A. Resch, and U. Bläsi. 2010. Translational activation of rpoS mRNA by the non-coding RNA DsrA and Hfq does not require ribosome binding. Nucleic Acids Res. 38:1284–1293.[PubMed] [CrossRef]

504. Vidal, O., R. Longin, C. Prigent-Combaret, C. Dorel, M. Heooreman, and P. Lejeune. 1998. Isolation of an Escherichia coli K-12 mutant strain able to form biofilms on inert surfaces: involvement of a new ompR allele that increases curli expression. J. Bacteriol. 180:2442–2449.[PubMed]

505. Vieira, H. L., P. Freire, and C. M. Arraiano. 2004. Effect of Escherichia coli morphogene bolA on biofilms. Appl. Environ. Microbiol. 70:5682–5684.[PubMed] [CrossRef]

506. Vivas, E. I., and H. Goodrich-Blair. 2001. Xenorhabdus nematophilus as a model for host-bacterium interactions: rpoS is necessary for mutualism with nematodes. J. Bacteriol. 183:4687–4693.[PubMed] [CrossRef]

507. Vytvytska, O., J. S. Jakobsen, G. Balcunatie, J. S. Andersen, M. Baccarini, and A. von Gabain. 1998. Host factor I, Hfq, binds to Escherichia coli ompA mRNA in a growth rate-dependent fashion and regulates its stability. Proc. Natl. Acad. Sci. USA 95:14118–14123.[PubMed] [CrossRef]

508. Wade, J. T., D. C. Roa, D. C. Grainger, D. Hurd, S. J. Busby, K. Struhl, and E. Nudler. 2006. Extensive functional overlap between sigma factors in Escherichia coli. Nat. Struct. Mol. Biol 13:806–814.[PubMed] [CrossRef]

509. Wang, A.-Y., and J. E. Cronan , Jr. 1994. The growth phase-dependent synthesis of cyclopropane fatty acids in Escherichia coli is the result of an RpoS (KatF)-dependent promoter plus enzyme instability. Mol. Microbiol. 11:1009–1017.[PubMed] [CrossRef]

510. Wang, L., b. Spira, Z. Zhou, L. Feng, R. P. Maharjan, X. Li, F. Li, C. McKenzie, P. R. Reeves, and T. Ferenci. 2010. Divergence involving global regulatory gene mutations in an Escherichia coli population evolving under phosphate limitation. Genome Biol. Evol. 2:478–487.[PubMed] [CrossRef]

511. Wang, S., R. T. Fleming, E. M. Westbrook, P. Matsumura, and D. B. McKay. 2006. Structure of the Escherichia coli FlhDC complex, a prokaryotic heteromeric regulator of transcription. J. Mol. Biol. 355:798–808.[PubMed] [CrossRef]

512. Wang, Y., and K. S. Kim. 2000. Effect or rpoS mutations on stress-resistance and invasion of brain microvascular endothelial cells in Escherichia coli K1. FEMS Microbiol. Lett. 182:241–247.[PubMed] [CrossRef]

513. Wassarman, K. M., and G. Storz. 2000. 6S RNA regulates E. coli RNA polymerase activity. Cell 101:613–623.[PubMed] [CrossRef]

514. Waterman, S. R., and P. L. C. Small. 1996. Identification of σ^S-dependent genes associated with the stationary-phase acid resistance phenotype of Shigella flexneri. Mol. Microbiol. 21:925–940.[PubMed] [CrossRef]

515. Waterman, S. R., and P. L. Small. 2003. Transcriptional expression of Escherichia coli glutamate-dependent acid resistance genes gadA and gadBC in an hns rpoS mutant. J. Bacteriol. 185:4644–4647.[PubMed] [CrossRef]

516. Watnick, P., and R. Kolter. 2000. Biofilm, city of microbes. J. Bacteriol. 182:2675–2679.[PubMed] [CrossRef]

517. Weber, A., S. A. Kögl, and K. Jung. 2006. Time-dependent proteome alterations under osmotic stress during aerobic and anaerobic growth in Escherichia coli. J. Bacteriol. 188:7165–7175.[PubMed] [CrossRef]

518. Weber, H., C. Pesavento, A. Possling, G. Tischendorf, and R. Hengge. 2006. Cyclic-di-GMP-mediated signaling within the σ^S network of Escherichia coli. Mol. Microbiol. 62:1014–1034.[PubMed] [CrossRef]

519. Weber, H., T. Polen, J. Heuveling, V. Wendisch, and R. Hengge. 2005. Genome-wide analysis of the general stress response network in Escherichia coli: σ^S-dependent genes, promoters and sigma factor selectivity. J. Bacteriol. 187:1591–1603.[PubMed] [CrossRef]

520. Weichart, D., R. Lange, N. Henneberg, and R. Hengge-Aronis. 1993. Identification and characterization of stationary phase-inducible genes in Escherichia coli. Mol. Microbiol. 10:407–420.[PubMed] [CrossRef]

521. White-Ziegler, C. A., S. Um, N. M. Pérez, A. L. Berns, A. J. Malhowski, and S. Young. 2008. Low temperature (23°C) increases expression of biofilm-, cold-shock- and RpoS-dependent genes in Escherichia coli K-12. Microbiology 154:148–166.[PubMed] [CrossRef]

522. Wilmes-Riesenberg, M. R., J. W. Foster, and R. Curtiss III. 1997. An altered rpoS allele contributes to the avirulence of Salmonella typhimurium LT2. Infect. Immun. 65:203–210.[PubMed]

523. Wilson, J. A., T. J. Doyle, and P. A. Gulig. 1997. Exponential-phase expression of spvA of the Salmonella typhimurium virulence plasmid: induction in intracellular salts medium and intracellularly in mice and cultured mammalian cells. Microbiology 143:3827–3839.[PubMed] [CrossRef]

524. Wise, A., R. Brems, V. Ramakrishnan, and M. Villarejo. 1996. Sequences in the -35 region of Escherichia coli rpoS-dependent genes promote transcription by E σ^S. J. Bacteriol. 178:2785–2793.[PubMed]

525. Wolf, S. G., D. Frenkiel, T. Arad, S. E. Finkel, R. Kolter, and A. Minsky. 1999. DNA protection by stress-induced biocrystallization. Nature 400:83–85.[PubMed] [CrossRef]

526. Xu, K. D., M. J. Franklin, C.-H. Park, G. A. McFeters, and P. S. Stewart. 2001. Gene expression and protein levels of the stationary phase sigma factor, RpoS, in continuously-fed Pseudomonas aeruginosa biofilms. FEMS Microbiol. Lett. 199:67–71.[PubMed] [CrossRef]

527. Yamashino, T., C. Ueguchi, and T. Mizuno. 1995. Quantitative control of the stationary phase-specific sigma factor, σ^S, in Escherichia coli: involvement of the nucleoid protein H-NS. EMBO J. 14:594–602.[PubMed]

528. Yang, H., E. Wolff, M. Kim, A. Diep, and J. H. Miller. 2004. Identification of mutator genes and mutational pathways in Escherichia coli using a multicopy cloning approach. Mol. Microbiol. 53:283–295.[PubMed] [CrossRef]

529. Yang, S., C. R. Lopez, and E. L. Zechiedrich. 2006. Quorum sensing and multidrug transporters in Escherichia coli. Proc. Natl. Acad. Sci. USA 103:2386–2391.[PubMed] [CrossRef]

530. Yildiz, F. H., and G. K. Schoolnik. 1998. Role of rpoS in stress survival and virulence of Vibrio cholerae. J. Bacteriol. 180:773–784.[PubMed]

531. Yim, H. H., and M. Villarejo. 1992. osmY, a new hyperosmotically inducible gene, encodes a periplasmic protein in Escherichia coli. J. Bacteriol. 174:3637–3644.[PubMed]

532. Yim, H. H., R. L. Brems, and M. Villarejo. 1994. Molecular characterization of the promoter of osmY, an rpoS dependent gene. J. Bacteriol. 176:100–107.[PubMed]

533. Zambrano, M. M., and R. Kolter. 1996. GASPing for life in stationary phase. Cell 86:181–184.[PubMed] [CrossRef]

534. Zambrano, M. M., D. A. Siegele, M. Almirón, A. Tormo, and R. Kolter. 1993. Microbial competition: Escherichia coli mutants that take over stationary phase cultures. Science 259:1757–1760.[PubMed] [CrossRef]

535. Zgurskaya, H. I., M. Keyhan, and A. Matin. 1997. The σ^S level in starving Escherichia coli cells increases solely as a result of its increased stability, despite decreased synthesis. Mol. Microbiol. 24:643–651.[PubMed] [CrossRef]

536. Zhang, A., K. M. Wassarman, C. Rosenow, B. C. Tjaden, G. Storz, and S. Gottesman. 2003. Global analysis of small RNA and mRNA targets of Hfq. Mol. Microbiol. 50:1111–1124.[PubMed] [CrossRef]

537. Zhang, A., S. Altuvia, A. Tiwari, L. Argaman, R. Hengge-Aronis, and G. Storz. 1998. The OxyS regulatory RNA represses rpoS translation and binds the Hfq (HF-I) protein. EMBO J. 17:6061–6068.[PubMed] [CrossRef]

538. Zhang, X. S., R. García-Contreras, and T. K. Wood. 2008. Escherichia coli transcription factor YncC (McbR) regulates colanic acid and biofilm formation by repressing expression of periplasmic protein YbiM (McbA). ISME J. 2:615–631.[PubMed] [CrossRef]

539. Zhao, G., P. Ceci, A. Ilari, L. Giangiacomo, T. M. Laue, E. Chiancone, and N. D. Chasteen. 2002. Iron and hydrogen peroxide detoxification properties of DNA-binding protein form starved cells: a ferritin-like DNA-binding protein of Escherichia coli. J. Biol. Chem. 277:27689–27696.[PubMed] [CrossRef]

540. Zhao, K., M. Liu, and R. R. Burgess. 2007. Adaptation in bacterial flagellar and motility systems: from regulon members to “foraging”-like behavior in E. coli. Nucleic Acids Res. 35:4441–4452.[PubMed] [CrossRef]

541. Zheng, M., X. Wang, L. J. Templeton, D. R. Smulski, R. A. LaRossa, and G. Storz. 2001. DNA microarray-mediated transcriptional profiling of the Escherichia coli response to hydrogen peroxide. J. Bacteriol. 183:4562–4570.[PubMed] [CrossRef]

542. Zhou, A. N., and S. Gottesman. 1998. Regulation of proteolysis of the stationary-phase sigma factor RpoS. J. Bacteriol. 180:1154–1158.[PubMed]

543. Zhou, Y., and S. Gottesman. 2006. Modes of regulation of RpoS by H-NS. J. Bacteriol. 188:7022–7025.[PubMed] [CrossRef]

544. Zhou, Y., S. Gottesman, J. R. Hoskins, M. R. Maurizi, and S. Wickner. 2001. The RssB response regulator directly targets σ^S for degradation by ClpXP. Genes Dev. 15:627–637.[PubMed] [CrossRef]

545. Zinser, E. R., and R. Kolter. 2004. Escherichia coli evolution during stationary phase. Res. Microbiol. 155:328–336.[PubMed] [CrossRef]

546. Zogaj, X., M. Nimtz, Rohde, M., W. Bokranz, and U. Römling. 2001. The multicellular morphotypes of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix. Mol. Microbiol. 39:1452–1463.[PubMed] [CrossRef]

547. Zusman, T., O. Gal-Mor, and G. Segal. 2002. Characterization of a Legionella pneumophila relA insertion mutant and roles of RelA and RpoS in virulence gene expression. J. Bacteriol. 184:67–75.[PubMed] [CrossRef]