Stationary-Phase Gene Regulation in <i>Escherichia coli</i><sup>?</sup>
Module
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:
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§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.
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 σB, 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 σS (RpoS) as a master regulator of the stationary phase and general stress response, with σS being under complex transcriptional and posttranscriptional control. In β-proteobacteria, σS 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.
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.
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, σFliA, σS, σE, 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).
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 σFliA 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, σS does not yet seem to gain significant access to RNAP core , because the large majority of σS-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 σFliA (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 σS-dependent genes by binding to and blocking their −10 promoter regions. FliZ thereby gives priority to FlhDC-driven motility during the initial phase of σS 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, σS 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 σS-containing RNAP holoenzyme (EσS) formation. This results in the activation of σS-dependent genes in a fine-tuned temporal order that probably reflects the affinity of their promoters for EσS (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 σE (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 σFliA 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, σS-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.
During transition into stationary-phase induction of σS 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 σS and activate σS-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 σS accumulation and therefore generate "cross-protection" (224, 225, 267, 318) (Fig. 2).
In addition, σS 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, σS 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 discovery of σS (RpoS) as a master regulator for stationary-phase gene expression (267) was based on the observations that σS 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 "σS " were proposed because of the function in stationary phase and stress responses (267).
Basic structural properties of σS
In vitro, σS 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, σS 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, EσS 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 σS-dependent gene expression. Again in contrast to Eσ70, the activity of EσS is positively modulated by these molecules by a mechanism that involves the C-terminal region of σS (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 σS-β-flap interaction have been implicated in the formation of a "poised" state of promoter-bound EσS from which EσS can productively escape in response to certain stress conditions that result in DNA relaxation and/or accumulation of potassium glutamate (166, 208, 407).
The σS promoter consensus sequence and the "σS promoter selectivity paradox"
Initial sequence comparisons of σS-controlled promoters left researchers puzzled with the finding that EσS 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 EσS revealed −35 and −10 hexamers basically identical to the σ70 consensus promoter (TTGACA and TATACT, respectively) (152). Nevertheless, EσS 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 "σS 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 EσS binding (152), natural σS-controlled promoters have been selected during evolution for a complex combination of properties: (i) an optimal balance of EσS binding, open complex formation, and promoter clearance ; (ii) a distinct degree of σS selectivity; and (iii) a certain expression level, depending on the physiological roles of the respective genes. In vivo many genes are exclusively σS-dependent, but there are also promoters that can be activated by both holoenzymes such that Eσ70 is responsible for expression during growth, and EσS "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 σS-selective promoters thus follows a complex mixed strategy that relies upon a series of special properties of EσS and its interactions with promoter sequences. As a consequence, σS-dependent promoters have modular structures with each promoter using a specific combination of these properties. Features that can contribute to σS selectivity of a promoter are summarized in the following (for a more detailed account, the reader is referred to reference 493).
EσS is more tolerant than Eσ70 with respect to deviations from the "optimal" consensus promoter
EσS-mediated transcriptional activity is less compromized by partially degenerate −35 regions (49, 152, 261, 284, 491, 524). Moreover, EσS 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 EσS than by Eσ70, this "deviation from the optimum" strategy can increase σS 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 EσS is indifferent to the −18 nucleotide (446). Because of its reduced stringency with respect to the −35 sequence and spacer length and sequence, EσS 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 EσS with respect to the −35 position is probably related to the domains 4 of σS and σ70 interacting differently with the β-flap domain of the RNAP core enzyme (259, 491).
EσS makes additional contacts to DNA sites outside the core −10 hexamer in a manner distinct from Eσ70
A hallmark of σS-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 σS (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 σS selectivity, the presence of C(−13) also enhances absolute promoter strength with EσS in comparison with all three other bases at this position. T(−14), however, provides higher σS 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 EσS-mediated transcription initiation. Thus, σS-dependent promoters identified by analyzing runoff transcripts on microarrays ( ROMA), which were obtained with EσS and linearized chromosomal DNA fragments in vitro, represented a smaller subset of the σS regulon in which promoters with a C(−13) were strongly underrepresented (301).
Moreover, σS-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 EσS-mediated transcription (361, 390), and probably optimizes promoter melting at these promoters, because EσS binds untwisted but not melted promoters more weakly than Eσ70 (284). Taking together all these promoter elements, the consensus sequence for a σS-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 σS 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" σS-dependent promoter also has an information content similar to that of standard promoters with typical −35 and −10 hexamers (493).
EσS uses UP-elements in a manner distinct from Eσ70
There is another class of σS-dependent promoters that do show recognizable (though usually not perfect) −35 regions, but, despite this similarity to vegetative promoters, they are σS-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 EσS-mediated expression, but interferes with Eσ70 activity, whereas either a proximal half-site or a full UP-element strongly favors Eσ70-dependent expression. EσS-mediated promoter activation by a distal half-site obligatorily requires the presence of a −35 element, i.e., fixation of domain 4 of σS at this position of the promoter (490). The striking discrimination between the two sigma factors is probably based on the failure of σS 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 σS 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 EσS (493). In conclusion, a second type of σS-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).
EσS 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 σS-controlled promoters, and preliminary data indicate that transcription by EσS may be less affected than by Eσ70 at these sites (493). In addition, σS seems more prone to leave the early elongation complex than σ70 (392), which not only increases the cellular pool of free σS available for sigma factor competition for RNAP core, but can make EσS-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 σS selectivity of a promoter.
In contrast to "simple" alternative RNAP holoenzymes, as, for instance, Eσ32, which is responsible for the heat shock response (289), EσS resembles the vegetative Eσ70 because its activity can be modulated by additional regulatory proteins that bind to promoter regions. As a consequence, expression of σS-dependent genes does not necessarily just follow σS accumulation in the cell. Rather, subsets of the σS 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 σS accumulation (519). The remaining genes exhibited σS-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 σS-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 σS 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 σS 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 EσS 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 EσS and Eσ70, and thereby activates the two target promoters yaiCp and csgBp, respectively (518).
Abundant histone-like proteins
In addition to "classical" regulators, major nucleoid-associated or "histone-like" proteins can differentially affect promoter activation by EσS and Eσ70 and thereby contribute to σS selectivity. The regulons controlled by the leucine-responsive regulatory protein, Lrp, and σS show a large overlap (471), and, in an lrp mutant background, numerous genes exhibit altered ratios of σS-dependence (519). Similarly, integration host factor, IHF, modulates the expression of many σS-controlled genes (310). Depending on the specific promoter, activation or repression by these factors is more strongly imposed upon EσS 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 σS-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 EσS-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 EσS (438). However, EσS 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 σS control, thus making gadA expression indirectly dependent on σS (157).
In apparently rare cases, H-NS can exert a direct positive effect on σS-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 σS selectivity, which, in the csiD promoter, depends on the position of cAMP-CRP binding (156, 324).
The primary requirement for inducing the general stress response in E. coli is σS accumulation, with expression patterns of many σS-dependent genes then being fine-tuned by additional signal input. Because σS 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 σS proteolysis (Fig. 3; reviewed in reference 196) . Moreover, the efficiency of σS in sigma factor competition for RNAP core, and therefore σS activity, is subject to regulation (reviewed in reference 493).
The highly fine-tuned gradual accumulation of σS during the transition from exponential to postexponential and finally to stationary phase mostly relies on the control of σS expression and combines mechanisms of transcriptional and translational regulation (268, 270). However, rapid "emergency" accumulation of σS in response to potentially life-threatening stress conditions is mainly dependent on a rapid inhibition of σS proteolysis (28, 53, 188, 222, 268, 308, 340). In addition, σS stabilization against proteolytic degradation also allows σS to accumulate further in starved cells, when expression rates for σS (as for most proteins) are reduced again (268, 535).
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 (rpoSp1) 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 rpoSp1 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 σS 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 σS 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 rpoSp1 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 rpoSp1 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 σS levels under all conditions, but still exhibit an induction of σS 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 σS closely correlate both during steady-state growth and upon sudden growth rate transitions (472). While (p)ppGpp seems to also contribute to stabilization of σS protein during entry into the stationary phase (230) (and see below), a major part of its positive effect on σS is due to a role in rpoS transcription, as shown with transcriptional rpoS::lacZ fusions. When in these fusion constructs the original rpoSp1 was replaced by tacp or lacUV5p, expression was still clearly reduced in relA spoT mutants (albeit somewhat less than when rpoSp1 was present) (203, 270). Thus, it appears that (p)ppGpp affects both transcription initiation at rpoSp1 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.
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 σS 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 σS 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 σS 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 σS and high σS 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 σS protein (322) (and see below), with the net result that σS 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 σS 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.
Although rapidly growing cells do express σS 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, σS 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, σS 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 σS proteolysis and its control by stress signals have recently been reviewed in detail (188).
Role of the proteolytic targeting factor RssB in ClpXP-mediated σS turnover
σS is degraded by the ATP-dependent complex ClpXP protease (434). However, ClpXP alone cannot bind to σS, 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 σS to ClpXP (28, 339, 383, 542). RssB directly binds to σS with a 1:1 stoichiometry (30, 31, 246, 335, 544). Within σS, 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 σS that makes the direct contact to the C(−13) position that is highly conserved in σS-dependent promoters (29) (and see above). By contrast, both domains of RssB are required to bind σS in vitro and to downregulate σS 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 σS (for a detailed discussion, see reference 188). Thus, in vitro binding and turnover of σS are stimulated by phosphorylation (30, 544). Interestingly, RssBD58P or RssBD58A 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 σS 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 σS 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 σS. 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 σS 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 σS (188, 466). In contrast to σS, RssB does not or only weakly interact with ClpX6 in the absence of its substrate σS (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-σS 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 σS 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 σS, which is consistent with σS 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 σS. RssB then inhibits activity of σS, as it interferes with σS binding to RNAP core; when overproduced in stoichiometric amounts with σS, it can act like an anti-sigma factor (31, 246, 335, 542).
Role of ArcB-mediated RssB phosphorylation in the control of σS proteolysis
Proteolysis of σS 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 σS and RssB in one way or another. This allows degradation of other ClpXP substrates to continue, while σS proteolysis becomes inhibited and σS 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 σS 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 σS proteolysis and relieves rpoS transcriptional repression, respectively, with both processes integrating into a gradual accumulation of σS (325).
Role of the σS-RssB feedback cycle in the control of σS proteolysis
Interestingly, the slow and fine-tuned regulation of RssB phosphorylation does not contribute to the rapid "emergency" stabilization of σS triggered by many suddenly arising acute stress conditions (325, 376). Rather, sudden inhibition of σS proteolysis depends on rapid alterations of the ratio between cellular σS and RssB available for interaction. This requires that (i) RssB is the rate-limiting factor for σS degradation in vivo, which was experimentally demonstrated (390); and (ii) an appropriate σS:RssB ratio is maintained under nonstress conditions, i.e., during growth when σS 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 σS (390, 412). Thus, σS activates the expression of its own "destruction factor" (Fig. 4). This homeostatic negative feedback allows maintenance of the same high σS degradation rates under conditions of changing σS expression rates (up to a threshold where the relatively weak rssAp promoter becomes saturated) (18). In addition, this homeostatic feedback loop sets the stage for controlling the rate of σS proteolyis by rapidly acting competition, titration, and sequestration mechanisms that alter the ratio between σS and RssB available for interaction. Moreover, the more slowly operating homeostatic feedback loop can confer adaptation, i.e., can result in rapid σS stabilization being more or less transient (188).
Role of RssB sequestration for the control of σS proteolysis
Many mechanisms can affect the σS:RssB ratio, which provides a basis for multiple-signal integration in the control of σS degradation (Fig. 4). On the one hand, RssB can be titrated by sudden strong σS synthesis, which means that σS stabilization can actually be a passive consequence of increased σS 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 σS. 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 σS or even with σS 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 σS (311). Inactivation of a fraction of the (now stable) σS by RssB and ClpXP has indeed been observed in starved cells (31, 542). Because of the excess of σS over RssB, only a fraction of σS can be "co-trapped" and therefore inactivated, and the large remainder would be stable and free to interact with RNAP core enzyme.
Role of EσS holoenzyme formation in the control of σS proteolysis
Finally, RssB competes with RNAP core enzyme for σS binding (544) (Fig. 4). As a consequence, the cellular level of free σS available to interact with RssB, and thus the rate of σS degradation, is also modulated by processes that affect binding of σS to RNAP core enzyme. Therefore, Crl, a protein that binds to σS and stimulates its association with RNAP, has a stabilizing effect on σS (54, 492). The anti-σ70 factor Rsd, which reduces the level of σ70 available for competition with σS for RNAP core (228), could have a similar effect on σS turnover. Overall, a complex protein-protein interaction network connects the σS degradation and σS activity cycles, such that stress-induced changes in many of these interactions can indirectly affect σS proteolysis. Moreover, this network can exert several homeostatic functions: increased EσS formation will be compensated for by increased RssB expression in the longer run; also, increased EσS formation at the expense of Eσ70 formation will result in decreased σS expression, since rpoS is transcribed from Eσ70- controlled promoters.
Open questions in the regulation of σS proteolysis
In conclusion, we now begin to understand the molecular mechanisms by which σS 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, σS 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 σS 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 σS turnover, has yet to be identified (204).
Finally, σS 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 σS 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 σS, 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 σS 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 σS proteolysis (74, 75), but further work is clearly required here.
Components affecting sigma factor competition for RNAP core enzyme
In order to become active, σS 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 σS, σFliA, σ 32, and/or σE 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 σS (see above) and σE (95), (iii) may contribute to σS 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 EσS formation and activity
While all these mechanisms promote alternative RNAP holoenzyme formation, they do not favor EσS formation specifically. Nevertheless, EσS 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, σS levels reach only about one-third of those of σ70 (227) and the in vitro affinity of purified σS for RNAP core is the lowest of all sigmas in E. coli (300). Therefore, additional factors are required for σS 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 σS activity at certain target genes (384). Crl binds to domain 2 of free and RNAP-associated σS (54, 125, 333) and stimulates the expression of a large set of σS-dependent genes independently of any specific promoter motif (492). In vivo, where σS has to compete with other sigmas for RNAP core, Crl is a dedicated σS-auxiliary factor, in particular, when σS levels are low (402, 403, 492). In vitro, Crl stimulates gene expression by EσS more than by Eσ70 (151), and promotes EσS formation and activation under conditions where σS and σ70 compete for RNAP core (492). It does so by forming a 1:1 complex with σS, 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 EσS holoenzyme (125).
Role of K+ in stimulating σS activity
High intracellular levels of K+ (as found in a mutant devoid of enzyme IIA(Ntr) of the nitrogen PTS) increase σS-dependent gene expression without affecting σS levels. In vitro, K+ stimulates σS binding to RNAP core at the expense of Eσ70 formation (278). Moreover, K+ also directly stimulates EσS activity on certain promoters in a process that requires the C-terminal 16-amino-acid extension of σS 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 EσS may add to enhanced synthesis and stabilization of σS, thus explaining the very strong osmoregulation of many σS-dependent genes (519).
Role of FliZ protein as an antagonist of σS activity
Finally, FliZ was recently identified as a factor that negatively affects the expression of many σS-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 σFliA (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 σS by recognizing an operator sequence that resembles the extended −10 region of σS-dependent promoters. FliZ does so with a structural element that mimics the first α-helix (α3.0) in domain 3 of σS, which makes contact to the extended−promoter element, and, in fact, the C(−13) promoter nucleotide is of key importance for interaction with both EσS and FliZ (377).
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 σS-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 σS-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).
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 σS (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 σS control, with σS being the top regulator in regulatory cascades. Moreover, many target genes are under direct as well as indirect σS 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 σS 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, σS 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.
The currently best studied example is the network by which σS 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). EσS directly controls csgD transcription, but does so in conjunction with a multitude of regulatory factors, many of which are themselves under σS 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 σS (374, 447, 518), whereas yhjH is a class III gene in the flagellar hierarchy (148, 158, 248, 374, 414). Since σS, 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, σS 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, EσS 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.
Another σS-controlled network consisting of nested feedforward loops is responsible for acid resistance (141). EσS, 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 σS-controlled YmgB protein (282, 484).
Taken together, a similar complex pattern as in curli/cellulose control appears in the acid resistance network: σS 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 σS-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).
Finally, some σS-controlled genes are involved in regulatory feedback loops that affect σS itself. The most important one is certainly σS-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 σS proteolysis by numerous stress signals (as described above). In addition, positive feedback loops probably assist robust accumulation of σS in stationary phase or upon stress exposure: (i) via sigma factor competition, σS 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 σS-dependent (229); (iii) σS 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 σS stabilization (accordingly, an arcA mutant stabilizes σS to a lesser extent during entry into the stationary phase and shows reduced survival and stress resistance) (325, 357); and (iv) the σS-dependent biofilm regulator CsgD may be involved in further stabilizing σS, 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 σS-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.
The σS-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, σS 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).
σS 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 σS 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 σS-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, σS-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 σS, but high levels of trehalose accumulate only in growing cells shifted to high osmolarity (155, 191, 236). Another example is σS-controlled expression of mechanosensitive channels, which become active only in response to osmotic downshift (457).
Because of the large size of the σS regulon, it is often difficult to pinpoint the specific genes required for particular σS-controlled phenotypes. Consequently, complex phenotypes, as for instance σS-dependent thermotolerance in stationary phase, are still not understood on a mechanistic basis, whereas, on the other hand, many σS-controlled genes have remained functionally uncharacterized. This review can just give a cursory overview of a limited number of σS-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.
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, σS is also induced upon osmotic upshift in growing cells (193, 337), and σS-dependent expression was observed for 326 genes under these conditions. These included 186 genes not found under other conditions where σS accumulates, indicating that σS also serves as a major osmoregulator (519). Proteomic analysis corroborated the induction of many σS-controlled proteins upon osmotic upshift (517). σS 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 σS-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
σS 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 σS-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 σS 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 σS and/or Ada, a regulator that responds to alkylation damage and can also cooperate with vegetative RNAP to activate aidB (263).
Acid resistance
σS 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 σS (28, 141, 280, 514). A σS-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 σS 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 σS-dependent gabDTP operon (324).
Another target of the acid stress regulators under σS 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 σS-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 σS 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 σS-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).
σS-controlled stress-protective genes with unknown molecular functions
Specific stress-protective molecular functions have been assigned to many σS-controlled proteins. In other cases, however, the molecular details underlying stress protection by particular proteins have remained mysterious. A striking example is the strongly σS-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).
σS was originally seen as a global regulator that mainly confers multiple stress resistance (192, 195, 293). However, around 20% of the genes in the σS regulon turned out to have metabolic functions (112, 371, 519). During slow growth, σS 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, σS thus becomes a second housekeeping sigma factor in slowly growing or stationary-phase cells.
σS actively redirects energy metabolism when cells enter into the stationary phase. Many genes under positive control of σS 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 σS. One of the most strongly σS-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 σS-activated repressor PdhR (519). These σS-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 σS control (112, 371, 519). On the other hand, σS 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 σS-controlled stationary phase with its emphasis on maintenance and stress survival (see further below).
σS 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 σS-dependent gene fbaB, which encodes one of two fructose-bisphosphate aldolases (262, 371, 519). The molecular or physiological reasons underlying isozyme switching are unknown.
During entry into stationary-phase cells change to a shorter, more ovoid cell shape. This is due to σS-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 σS, 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 σS control (122, 509). Also, some lipoproteins show σS-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).
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 σS and many σS-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 σS playing a positive or negative role in biofilm formation. In Salmonella enterica Serovar Typhimurium, σS 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 σS (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 σS 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 σS-controlled downstream networks above).
In addition, the σS-regulated morphogene bolA was found to positively modulate biofilm formation (505). The σS-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 σS-dependent manner, especially at low temperatures (369, 521).
It is not surprising that σS 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 σS-dependent gene expression is required for the initial stages of systemic infection (352).
σS 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 σS (112, 476). By transcriptional profiling of an O157:H7 strain, approximately 10% of the σS-controlled genes observed were found to be unique to O157:H7 (112).
In S. enterica serovar Typhimurium, σS and σS-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). σS 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, σS 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).
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 σS-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 σS 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 σS control (147). In addition, the mismatch repair proteins MutS and MutH are downregulated in a σS-dependent manner (133, 486).
However, there are additional σS-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 σS, see above), dinB, as well as several other genes were isolated (528). The latter included two genes for which a positive control by σS 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 σS-controlled antimutator, the DNA glycosylase Mug, has also been described (330).
A series of regulatory genes that encode DNA-binding transcription factors are under σS control. The implications with respect to some regulatory cascades and networks downstream of σS have already been illustrated above (involving the σS-controlled regulators MlrA, CsgD, GadX, GadW, and GadE). During entry into stationary phase or upon osmotic upshift, σS 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 σS (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 σS-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 σS (519). In many cases, the functions of these regulatory proteins are incompletely understood or even unknown. Functionally characterized regulatory proteins, for which σS 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 σS in metabolic adaptation, multiple stress resistance, and biofilm formation.
In addition to controlling transcriptional regulators, σS is also involved in the expression of regulatory factors with a role in posttranscriptional regulation. Thus, YmdB, a partially σS-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 σS expression.
Different laboratory strains of E. coli K-12 or Salmonella show variations in their σS 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 σS 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 σS-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 σS completely, but result in attenuated σS function (139, 533, 534, 545). Sometimes, these mutants can stably coexist with σS-proficient strains (138, 410). Because stress resistance even quantitatively correlates with σS levels (37), the mutants with reduced σS levels or activity are less stress-resistant and also lose other σS-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 σS activity (135, 244, 356). An additional benefit of attenuated rpoS mutations could be the avoidance of the increased mutation rates associated with high σS 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 EσS (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, σS, σE, 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 EσS is stabilized by positive feedback loops (see above). Mutations that attenuate σS levels and/or its efficiency in sigma factor competition actually readjust the threshold of switching to the "high EσS/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.
σS-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 σS-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 σS 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 σS 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 σS-specific C-terminal extension that is not present in σ70 (106, 165, 285, 360, 406, 407). All γ-proteobacteria that do have σS (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 σS 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 σS. 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 "σS" 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 σS of E. coli, and also does not have the C-terminal extension. Finally, the Borrelia sigma factor annotated as a σS has a leucine at the K173 position, no C-terminal extension, and just 34% identity to E. coli σS.
TABLE 1.Comparison of functionally important elements in sigma factors annotated as σS (RpoS) in γ-, β-, and δ-proteobacteria and spirochetes| Sigma factor from: | Identity to E. coli σS (%) | Amino acid at K173 position | C-terminal 16-amino-acid extensiona | Presence of RssB |
| γ-Proteobacteriab | 60–99 | K | + | + c |
| β-Proteobacteriad | 50–53 | R | (+) | - |
| δ-Proteobacteriae | 47 | D | - | - |
| Spirochetef | 34 | L | - | - |
| σ70 (RpoD) of E. coli | 42 | E | - | + g |
|
|
|
|
|
|
|
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 σS factors of γ- and β-proteobacteria, the same may also apply to the Geobacter σS. In fact, this would provide an explanation for the absence of a σS-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 σS present in γ/β-proteobacteria or not, can be settled.
Within the γ-proteobacteria, the molecular details of σS 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 σS can vary. Enteric bacteria other than E. coli also use σS 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, σS 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, σS is involved in environmentally modulated biofilm formation (346). In L. pneumophila, σS 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 σS-dependent gene regulation into their complex two-component and quorum-sensing network that controls exoenzyme expression and virulence (182, 432, 433) and σS seems to affect biofilm formation (200, 526). Overall, this functional diversification also indicates how easily genes are recruited into or lost from the σS regulon ; in fact, because of the similarity of σS-dependent promoters to vegetative promoters, one or two point mutations in a promoter could be sufficient to recruit a vegetatively controlled gene into the σS regulon or vice versa.
Twenty years after the first description of σS 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 σS 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 σS in promoter recognition and the core mechanisms of σS regulation, there are vast blanks with respect to multiple signal integration into σS control. Moreover, the architecture and dynamic behavior of the complex networks directed by σS 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 σS 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 σS homologs seem a widely open field for exciting discoveries, as recently demonstrated with α-proteobacteria (143).
Research in my laboratory has been generously funded by the Deutsche Forschungsgemeinschaft, an ERC Advanced Investigator Grant, and the Fonds der Chemischen Industrie.
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